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GOLIKOV Marine Biological Station Zoological Institute H АРУ ARD Millport, United Kingdom St. Petersburg, Russia VERSITY jallen O uacf gla.ac.uk Ц Ni А. V. GROSSU Е. Е. BINDER Universitatea Bucuresti Museum d'Histoire Naturelle Romania Geneve, Switzerland T. HABE P. BOUCHET Muséum National d'Histoire Naturelle Paris, France bouchet@cimrs1.mnhn.fr P. CALOW University of Sheffield United Kingdom R. CAMERON Sheffield United Kingdom R.Cameron @sheffield.ac.uk J. G. CARTER University of North Carolina Chapel Hill, U.S.A. MARYVONNE CHARRIER Universite de Rennes France Maryvonne.Charrier @univ-rennes1.fr В. Н. COWIE University of Hawaii Honolulu, HI., U.S.A. A. H. CLARKE, Jr. Portland, Texas, U.S.A. B. C. CLARKE University of Nottingham United Kingdom R. DILLON College of Charleston SC, U.S.A. С. J. DUNCAN University of Liverpool United Kingdom D. J. EERNISSE California State University Fullerton, U.S.A. E. GITTENBERGER Rijksmuseum van Natuurlijke Historie Leiden, Netherlands sbu2eg @ rulsfb.leidenuniv.de F. GIUSTI Universita di Siena, Italy giustif @ unisi.it Tokai University Shimizu, Japan R. HANLON Marine Biological Laboratory Woods Hole, Mass., U.S.A. G. HASZPRUNAR Zoologische Staatssammlung Muenchen Muenchen, Germany haszi @zi. biologie. uni-muenchen.de J. M. HEALY University of Queensland Australia jhealy @zoology.uq.edu.au D. M. HILLIS University of Texas Austin, U.S.A. К.Е. HOAGLAND Council for Undergraduate Research Washington, DC, U.S.A. Elaine @ cur.org В. HUBENDICK Naturhistoriska Museet Goteborg, Sweden S. HUNT Lancashire United Kingdom R. JANSSEN Forschungsinstitut Senckenberg, Frankfurt am Main, Germany М. $. JOHNSON University of Western Australia Nedlands, WA, Australia msj @cyllene.uwa.edu.au В. М. KILBURN Natal Museum Pietermaritzburg, South Africa M. À. KLAPPENBACH Museo Nacional de Historia Natural Montevideo, Uruguay J. KNUDSEN Zoologisk Institut Museum Kobenhavn, Denmark С. LYDEARD University of Alabama Tuscaloosa, U.S.A. clydeard @ biology.as.ua.edu C. MEIER-BROOK Tropenmedizinisches Institut Tubingen, Germany Н. К. MIENIS Hebrew University of Jerusalem Israel J. Е. MORTON The University Auckland, New Zealand J. J. MURRAY, Jr. University of Virginia Charlottesville, U.S.A. R. NATARAJAN Marine Biological Station Porto Novo, India DIARMAID O'FOIGHIL University of Michigan Ann Arbor, U.S.A. J. OKLAND University of Oslo Norway T. OKUTANI University of Fisheries Tokyo, Japan W. L. PARAENSE Instituto Oswaldo Cruz, Rio de Janeiro Brazil J. J. PARODIZ Carnegie Museum Pittsburgh, U.S.A. R. PIPE Plymouth Marine Laboratory Devon, United Kingdom RKPI@wpo.nerc.ac.uk J. P. POINTIER Ecole Pratique des Hautes Etudes Perpignan Cedex, France pointier @ gala. univ-perp. fr W. F. PONDER Australian Museum Sydney QUAY: Academia Sinica Qingdao, People’s Republic of China D. G. REID The Natural History Museum London, United Kingdom S. G. SEGERSTRALE Institute of Marine Research Helsinki, Finland А. STANCZYKOWSKA Siedlce, Poland Е STARMUHLNER Zoologisches Institut der Universitat Wien, Austria У. 1. STAROBOGATOV Zoological Institute St. Petersburg, Russia J. STUARDO Universidad de Chile Valparaiso C. THIRIOT University P et M. Curie Villefranche-sur-Mer, France thiriot @obs-vifr.fr 5. TILLIER Museum National d'Histoire Naturelle Paris, France J.A.M. VAN DEN BIGGELAAR University of Utrecht The Netherlands М. Н. VERDONK Rijksuniversiteit Utrecht, Netherlands H. WÂGELE Ruhr-Universitát Bochum Germany Heike. Waegele @ ruhr-uni-bochum.de ANDERS WAREN Swedish Museum of Natural History Stockholm, Sweden В. В. WILSON Dept. Conservation and Land Management Kallaroo, Western Australia H. ZEISSLER Leipzig, Germany A. ZILCH Forschungsinstitut Senckenberg Frankfurt am Main, Germany MALACOLOGIA, 2004, 46(1): 1-17 ANATOMY OF A SMALL CLAM, ALVEINUS OJIANUS (BIVALVIA: KELLIELLIDAE), WITH A DISCUSSION ON THE TAXONOMIC STATUS OF THE FAMILY George A. Evseev, Natalya K. Kolotukhina & Olga Ya. Semenikhina Institute of Marine Biology, Russian Academy of Sciences, Vladivostok 690041, Russia; inmarbio@mail.primorye.ru ABSTRACT The anatomy of the small bivalve Alveinus ojianus (Yokoyama, 1927) of 1.8-2.0 mm shell length from shallow bays of the northwestern Pacific was studied. The morphological fea- tures, in which A. ojianus differs from other bivalve species occurring in the shelf waters include: a reduced gill consisting of the inner demibranchs, the posterior filaments of which are replaced by a membrane lining the cloaca; one siphon, the walls of which consist of the cuticular epithelium lining the mantle cavity; the inner mantle fold of the adult is thickened and, in addition to the mantle gland, contains extracellular granules; the labial palps have no sorting ridges, and the gill flexure is without a food groove; the stomach has a thickened cuticular lining and a helicoid structure, which are homologues of the gastric shield of other bivalves; and there are no sorting areas, digestive pouch, caecum or dorsal hood in the stomach. Comparison of its internal morphological features with those of other bivalve taxa demonstrates the close similarity of the Kelliellidae to the deep-water Verticordiidae but not Veneroida, to which Kelliellidae are usually referred. However, comparison of Kelliellidae and Verticordiidae based on shell morphology shows that differences between them makes their relationship improbable. One of the reasons for these contradictions appears to be the paedomorphic development of A. ojianus and a mixed nature of the morphological features used for the taxonomy. Key words: anatomy, mantle, extracellular granules, gill, stomach, paedomorphic features. INTRODUCTION The mollusks with a shell length no more than 3—5 mm hold a special position in the taxonomic systems of the Bivalvia. In their morphological features, they can be divided into two groups. One group is the miniature adult bivalves — many species of Propeamus- siidae, Carditinae, Circinae and others — in which features are homologous to those of larger taxa, and which are usually included in a subfamily with those larger taxa (Habe, 1977; Dijkstra & Kastoro, 1997; Oliver & Zuschin, 2001). Small bivalves of the second group belong to families or superfamilies — Crenellidae, Montacutidae, Lasaeidae, Philo- bryidae, Condylocardiidae and others — in which the macroforms are usually absent (Habe, 1977; Morton, 1978; Morton & Scott, 1989; Middelfart, 2000). Their shell morpho- logical features are usually characterized by hypoplasia, and among their internal organs there are structures that are unknown in macroforms (Oldfield, 1961; Morton, 1981; Mikkelsen & Bieler, 1989, 1992; Allen, 2000). As a result of this, in most cases the taxo- nomic features of the adult small forms are difficult to compare with those of the larger taxa, on which one of the most prevalent sys- tems of Heterodonta is based (Cox, 1969; Keen, 1969). The taxonomic status and phylo- genetic relationships some of them are still being discussed (Ockelmann, 1964; Cosel & Salas, 2001). Alveinus ojianus (Yokoyama, 1927) belongs to the otherwise deep-water family Kelliellidae, a member of the second group of the small bivalves. In addition to A. ojianus and another species of this genus recently found in the Red and Arabian seas (Oliver & Zuschin, 2001), in the family there are 12-14 species of Kelliella mainly inhabiting the waters of the eastern and southern Pacific (Knudsen, 1970; Bernard, 1989). Allen (2001) has provided a detailed description of this genus, including internal morphology, as well as the taxonomic history of the Kelliellidae and its relationships with other deep-water taxa. In contrast to deep-water Kelliella, species of the genus Alveinus occur both in shallow bays and open parts of the coastal zone (Miyadi & Habe, 1957; Oliver & Zuschin, 2001). Some data on biology of A. ojianus are to be found in general studies on the ecology and 2 EVSEEV ET AL. taxonomy of bivalves (Habe, 1950, 1973, 1977; Scarlato, 1981; Evseev, 2000). Accord- ing to these works, А. ojianus is widely distrib- uted in Peter the Great Bay and eastern part of the Sea of Japan, as well as off the Pacific coast of Japan and the South Kuril Islands. It is common in sandy mud at a depth of 2-22 m, where its density may exceed 1,000 speci- mens per m2. The bivalves attach to the sand grains with 1—2 byssus threads. The shell length of A. ojianus rarely exceeds 2 mm. The mollusks are easily identified by their triangu- lar, lustrous brownish shell. In this study, we examined the anatomy of A. ojianus, no data on which were found in the literature. There is also no information on the phylogenetic relationships of the Kelliellidae with its sister families. In this connection, we also attempted to estimate the taxonomic and phylogenetic significance of the internal fea- tures of A. ojianus for their use as additional taxonomic characters for the family Kellielli- dae, for which the taxonomy is almost com- pletely based on shell morphology. MATERIALS AND METHODS The adult and juvenile specimens of A. ojianus from the Amursky Bay and Vostok Bay, as well as the open part of Peter the Great Bay, the Sea of Japan, were used in this study. The bivalves were collected with a dredge from the research vessel “Lugovoye” in September 1999 and sampled by SCUBA-diving to 12-14 т depth in July-September 2000-2001 and in March 2002. Most of the specimens investigated measured from 350-1,000 um in length. The anatomy was studied by means of serial histological sections. The mollusks were pre- liminarily fixed in 96% ethanol. The shell valves were removed with a fine needle. The speci- mens were embedded in paraffin using routine methods, but the holding time in alcohol, chlo- roform and paraffin was greatly reduced. The sections were made from 7-10 um with a slid- ing microtome, mounted on glass slides and stained with Erlich and Boemer’s haemo- toxylin. A light microscope was used to exam- ine the sections. ABBREVIATIONS IN THE FIGURES 1 first cardinal tooth 3a third anterior cardinal tooth 3b Шиа posterior cardinal tooth aa anterior adductor abf ascending branchial filaments ag apical gland of the foot al line of attachment of the inner demibranch to the visceral mass an anus ar anterior retractor ba branchial axis bg byssal gland с cloaca cg terminal cuticula of the gill cls cutucular lining of the stomach cpg cerebral-pleural ganglion cso crystalline style opening ct loose connective tissue dd digestive diverticula ea exhalant aperture ec excretory cells eg extracellular granules of the mantle lobe fi foot g mantle gland gg extracellular granules of the apical gland gh _ gastric helicoid structure gr gastric ridges as “traffic circles” of the crystalline style ia inhalant aperture ici intracapillaceous inclusions ice inner ciliated epithelium icg intracellular granules id inner demibranch ifc interfilamentary connective ig intestinal groove ip — inhalant sensory papillae ipe inner pavement epithelium iue inner unciliated epithelium k kidney Ic branchial lateral cilia If longitudinal muscle fibres № branchial latero-frontal cilia lg marginal groove of the lunule li pit of the internal ligament Iml left mantle lobe le external ligament lp labial palps mg mid-gut mm mucous masses mmf middle mantle fold n nimpha O oesophagus obc opening of the byssogenous canal omf outer mantle fold ose outer stratified epithelium оу ovary p pericardium pa posterior adductor АМАТОМУ OF ALVEINUS OJIANUS 3 pd post-apertural dilatation of the mid-gut pg pedal ganglion pj posterior mantle junction pp posterior sensory papillae pr posterior retractor ps passage from the stomach to the post- apertural dilatation of mid-gut pss passage from the style sac to the post- apertural dilatation of mid-gut pt1 posterior lateral tooth pvj postero-ventral mantle junction r rectum rc rotary cilia of the crystalline style sac rml right mantle lobe rsw right stomach wall $ stomach sc statocyst capsule si exhalant siphon skr branchial skeletal rods sla slit-like aperture of the crystalline style sac ss crystalline style sac st statocyst st! statolith t testis th tooth of the gastric helicoid structure ty typhlosole ucf branchial unciliated filaments vj ventral mantle fusion vsw ventral stomach wall RESULTS А general view, details of the right shell, and the internal topography of A. ojianus are shown in Figures 1-4. Mantle The mantle edge has three folds. The thin outer fold is located along the posterior, ven- tral and anterior shell margin. The thin middle fold fuses near the anterior adductor and ven- trally (Fig. 1, vj) and forms a broad pedal- byssal gape occupying most of the ventral mantle edge. The inhalant aperture of the mantle cavity is separated from the exhalant aperture by the postero-ventral fusion of the fold (Fig. 1, ру). The thickened edges of the inhalant aperture bear laterally three marginal sensory papillae each. There are one to three short guard cilia at the papillae tips. Each side of the thickened, fused part of the middle mantle fold has four similar lateral pa- pillae between the inhalant and exhalant aper- tures. The exhalant aperture terminates in a conic siphon with a smooth, tapering opening. Cuticular tissue forms very thin, semi-trans- parent walls of the siphon. Like the base of the inhalant aperture, the base of the siphon is thickened and its posterior edge bears three papillae. The total number of sensory papillae on both sides of the inhalant and exhalant apertures amounts to 17. In juveniles, there may be fewer papillae; for example, in a speci- men of approximately 500 um shell length, two pairs of papillae were found at the lateral edges of the inhalant aperture, two pairs of common papillae and one pair of short papil- lae were located between the inhalant and exhalant apertures. As in adults, three papillae were present dorsal to the siphon. FIGS. 1, 2. Alveinus ojianus. FIG. 1. Lateral and ventral views of living specimen. Bar = 500 um. FIG. 2. Internal view of right shell valve showing hinge teeth. Bar = 300 um. 4 EVSEEV ET AL. FIG. 3. Topography of internal organs as seen from left side with left shell valve and left mantle lobe removed. Bar = 200 um. FIG. 4. Internal morphology and configuration as seen from left side (sagittal section). Bar = 100 um. АМАТОМУ OF ALVEINUS OJIANUS 5 The inner mantle fold of the adult is usually thickened (20-40 um), whereas the thickness of the other proximal mantle part does not exceed 6-8 um. In width, the thickened fold may vary from relatively narrow ridge to the crescent belt, which starts ventral to the ante- rior adductor, continues on either side of the pedal-byssal gape and ends close to the inhal- ant aperture. Sections made near the posterior edge of the pedal-byssal gape (Fig. 5) and through the central part of the inner thickened fold (Fig. 6) show that the thickening and the proximal di- latation of this fold are formed by connective and glandular tissue, as well as by yellowish extracellular granules. The surface of the inner wall of the mantle represents a smooth transparent epithelium consisting of polygonal pavement cells with FIGS. 5, 6. Extracellular granules of mantle. Bar = 50 um. FIG. 5. Transverse section through posterior part of layer of granules. Fig 6. Diagonal section through central part of their layer. marked nuclei and nucleoli. The wrinkled outer layer of the fold consists of a stratified semi- transparent epithelium with large cells and clearly distinguishable nuclei. Within the thick- ened fold, there 1$ a layer of deeply stained, vesicular floccular glandular tissue, under which the oval or oblong-angular grayish or yellowish granules are located. The granules usually form one, sometimes two layers. They are enveloped in the homoge- neous milk-white masses, which fuse into a single substance resembling the mucous se- cretion of the labial palps. Where granules form two layers, their size ranges from 20-30 um in the upper layer and from 10-12 to 15-20 um in the lower layer. If granules are in one layer, their length amounts to 35—40 um, and they are large and roundly elongate. The granules are joined to the outer or inner epithelium by their short sides. At the same time, contacting sides of the granules are often free of the milk-white masses. The glandular tissue sometimes sepa- rates the granule layers. The granules occur inside the mantle lobes in both males and females. ш adults, the total number of granules is 60—80. On left and right lobes of the same individual, the arrangement may be asymmetrical and varying in shape, size and quantity of the granules. These ap- pear to be different stages in their formation. In juveniles of 350—400 um shell length, gran- ules were not found. Adults 500-700 um long collected in October-November have from 20-25 to 40-55 granules in the anterior part of the mantle cavity, where they were located in a single layer. No granules were found in bivalves sampled at the end of March. Muscular System The anterior adductor is elongate and nar- rowed towards the retractor. The posterior adductor is larger and more rounded in shape. As in Turtonia minuta (Fabricius, 1780) and Lasaea rubra (Montagu, 1803) (Oldfield, 1955), both adductors consist of smooth, bundled muscle fibres (Figs. 3, 4). In other bivalves, muscle fibres of this type usually form the outer portion of adductor, which is responsible for maintaining valve closure (Yonge, 1936). An inner adductor component of “quick” cross-striated muscle fibres is lack- ing in A. ojianus. Asystem of circular, longitudinal and diago- nal muscle fibres is located within the foot. The longitudinal and diagonal fibres continue 6 EVSEEW ETAL dorsally and form a pair of anterior and a pair of posterior pedal retractors, which do not dif- fer from adductors in their color and structure. The other muscles are represented by smaller bundles and fine fibres scattered within the mantle and the visceral mass. Gills The subquadrate gills cover practically all the visceral mass. They consist of inner left and right demibranchs, each with long de- scending and short ascending lamellae. The descending lamellae extend from the bran- chial axes. The latter run almost parallel to the external ligament between the subumbonal enlargement of the visceral mass and the pos- terior adductor (Fig. 3, ba). The ascending lamella terminates dorsally in arcuate chiti- nous bridges attached to the visceral mass along a line between the adductors. The de- scending and ascending lamellae join ven- trally in a flexure with one or two interlamellar connectives. Thin, rounded, chitinous inter- — — FIGS. 7, 8. Branchial filaments. Bar = 30 um. FIG. 7. Descending filaments. FIG. 8. Ascending filaments. filamentar connectives that join skeletal rods of adjacent filaments are rare and irregular. The homorhabdic gill filaments are oval in a transverse section, with two skeletal rods join- ing abfrontally (Figs. 7, 8, skr). The filament walls consist of a one-layer ciliary epithelium with indistinct cell borders. The blood vessels are without muscle septa. Rare blood cells and deeply stained organic inclusions, which appear to be bacterial in a nature, occur within some vessels. The interfilamentar space 1$ filled with long cilia, of which the latero-frontal cilia are the most pronounced. Laterally, each filament bears symmetrical thick cilia (?) of unclear function, the orientation of which 1$ opposite that of the remaining cilia. The former are similar to the “anomalous” latero-frontal cilia delimiting the water fluxes (Atkins, 1938) but differ from them in direction and location. The diameter of the filament is about 30—40 ит; the filament number varies from 17 to 22 in an adult demibranch. Anteriorly, the demibranch is located be- tween the labial palps. However, the ventral marginal food groove is indistinct. The height of the ascending lamella decreases posteri- orly, and it is lacking behind the visceral mass. In its place, the descending lamella of the left demibranch joins the descending lamella of the right demibranch to form the reno-anal cavity (cloaca or suprabranchial cavity) (Pelseneer, 1906). Anteriorly, the lateral walls of the cloaca are formed by the distal limbs of the kidney. The posterior region of the lateral walls and the posterior wall of the cavity are lined internally with an elastic cuticular tissue (Figs. 3, 4, cg). The anus opens into the cloaca through the posterior wall; the cloaca commu- nicates ventrally with the siphon through the exhalant aperture. The postero-ventral wall of the cloaca joins the inner fold of the mantle between the inhalant and exhalant apertures. Foot The elongate foot is comparatively large. Its middle part is cylindrical; the apical part is pointed and ciliated. At its base, the foot usu- ally expands abruptly (Fig. 4). The postero- ventral part of the foot forms a well-marked heel. /n vivo, the foot may greatly expand, becoming about twice or three times as long as the shell. In fixed specimens, the foot is usually directed towards the labial palps. The outer wrinkled surface of the foot is covered with a stratified epithelium. АМАТОМУ OF ALVEINUS ОЛАМИ$ 7 There are two glands in the foot. The byssal gland (Fig. 4, bg) lies in the posterior part of the foot and consists of large, deeply stained cells forming the secreting lamellae. The latter are separated by narrow passages, which are filled with a homogeneous, poorly staining secretion. This system of the lamellae and passages converges ventrally to a broad byssogenous canal that opens anterior to the heel. The byssogenous canal is lined with one layer of similarly staining cells. The byssus threads are very thin (5-7 ит) and semi-trans- parent. Their proximal part is club-shaped; the distal tip bears a small terminal disc, which 1$ attached to large grains of sand or gravel. There are usually one or two byssus threads. The second apical gland (Fig. 4, ag) lies in the central and distal, or only in the distal parts of the foot. It is separated from the byssal gland by a layer of loose connective tissue and by a system of the blood lacunae, within which connective tissue islets and rare radial muscle fibres are scattered. The gland cells also form secreting lamellae, but these are narrower, more compact and less intensely stained in some places than the byssal gland cells. In the intercellular space of the apical gland, isometric or elongate homogenous, faintly stained granules sometimes occur. These are similar to the mantle cavity gran- ules in shape and dimensions. Unlike the byssal gland cells, the cells of the apical gland are small and rare in some individuals. In oth- ers, the secreting cells may be absent, and only the axial part of the foot consists of rela- tively well-stained glandular tissue surrounded by numerous lacunae. The byssal groove is not marked on the ven- tral side of the foot, and no openings are found in the distal and middle parts of the foot. The canal of the apical gland appears to open FIG. 9. Pedal ganglion and statocysts. Bar = 30 ит. either in the byssogenous canal or in the short byssal slit. Statocysts These are symmetrical sense organs repre- sented by spherical capsules approximately 30 um diameter. A spherical yellow or brown- ish statolith measuring up to 16-17 ит in di- ameter lies within each statocyst (Fig. 9). Cilia were not observed on the inner capsule walls. The statolith structure appears to be radial. A dark dot is sometimes seen in the center of statolith. The capsules are located in the anterior part of the foot lateral to the pedal ganglion. They belong to the type В, occurring in the Anomalodesmata (Morton, 1985). Labial Palps These are parallel anterior and posterior lamellae attached dorsal to the visceral mass (Fig. 4, Ip). The lamellae are ventrally elon- gated. Near their base is a broad funnel- shaped mouth that opens into a long oesophagus. The smooth inner surface of the palps and walls of the oesophagus are cov- ered by long, dense cilia, which are inclined proximally. Sorting ridges on the palps are lacking. The ciliary area is usually covered with dense mucus. Under the ciliated epithe- lium there are scattered, intensely staining mucous gland cells, as well as loose connec- tive tissue with numerous lacunae. The outer surface of the palps is unciliated. In fixed adults, the length of the anterior palps, including bases, is 250-300 um, whereas the length of the free ends does not usually exceed 100-150 ит. However, in liv- ing specimens the anterior palps are capable of expanding posteriorly up to 400-500 um. Alimentary Canal A long oesophagus enters the stomach antero-ventrally (Fig. 4, 0). The oesophageal walls consist of one layer of columnar ciliated epithelial cells, the long, dense cilia of which are directed towards the stomach. Mucous gland cells are scattered among the epithelial cells. Deep folds, three to four in number, may be seen in a transverse section of the oe- sophagus (Fig. 10). The stomach is hemispherical (Fig. 4, s), consisting of anterior and posterior sections. FIG. 10. Oesophagus (transverse section). Bar = 30 um. The lateral walls and antero-ventral section are thin and lined with the columnar digestive (?) epithelial cells containing numerous inclu- sions of different shape and density. The cell borders on the inside of stomach cavity are usually indistinct. Ciliated cells were not found. EVSEEV ET AL. The dorsal and right-dorsal walls of the ante- rior section are composed of a thickened cu- ticular tissue (Fig. 4, c/s) that is devoid of the inner covering of the digestive cells. In the center of the anterior section of the stomach there is a cup-shaped organ 100 um in diameter joined to the right-dorsal wall of the stomach (Fig. 11, gh). It is lined by the same thick cuticular tissue as the wall. In a transverse section, the organ is shaped like a convoluted or helicoid lamina. The initial bifur- cated part appears to function as the erosive tooth of the gastric shield. The distal height of the wall of the helicoid structure is 30-40 um. The antero-ventral wall of the helicoid struc- ture is absent, and ventrally there are the curved ridges (Fig. 11, gr) running towards the intestinal groove. The contents of the anterior section of the stomach consist of a mucous secretion and algae, among which the diatoms Thalassiosira, Pyxidicula, Odantella predominate. The dia- toms are represented both by the whole FIG. 11. Antero-dorsal wall of stomach and food contents (left view). Bar = 50 пт. АМАТОМУ OF ALVEINUS OJIANUS iue sla FIG. 13. Crystalline style sac. Transverse section through middle part and ventral wall of stomach. Bar = 50 um. 10 EVSEEV ЕТ AL. FIG. 14. Crystalline style sac. Transverse section through ventral wall and mid-gut opening. Bar = 50 um. thecas (25-30 um length) and large fragments forming assemblages near the oesophagial opening. The small fragments forming two to three diffuse spots usually occur anterior to the helicoid structure, where one of the ducts of the digestive diverticula appears to be located. FIG. 15. Mid-gut (transverse section). Bar = 20 ит. The relatively large crystalline style sac forms the posterior section of the stomach. The single layer of epithelial cells forming the sac walls has large nuclei and numerous in- clusions (Figs. 12-14). The inner surface of the sac is densely lined with deeply staining cilia, which rotate the short thickened style clockwise. A thin membrane coat continuous with the similar coat of the stomach wall cov- ers the outer surface of the sac. The sac communicates with the stomach by means of a longitudinal slit-like aperture (Fig. 12, sla). The dorsal part of the aperture, where the style projects from the sac, is broad, whereas the middle part, to which the intesti- nal groove leads from the stomach, is nar- rower (Fig. 13, s/a). In the middle part of the sac, the slit-like aperture is formed from the edges of the anterior sac wall, which are turned inward. Ventrally, the slit-like aperture extends to the bottom of the sac. In a trans- verse section through this area (Fig. 14), the АМАТОМУ OF ALVEINUS OJIANUS 11 left inward turned side of the aperture looks like the typhlosole, which is curved like a small tongue. It runs in a spiral along the posterior edge of the opening and the right wall of the sac into the post-apertural dilatation of the mid-gut. In other words, there are two pas- sages that lead into the post-apertural dilata- tion of the gut. One of them (Fig. 14, ps) communicates gut with stomach via the intes- tinal groove. The other (Fig. 14, pss) leads from the sac cavity into the gut through the slit-like aperture. The mid-gut runs from the bottom of the style sac towards the oesophagus (Fig. 4). At the anterior part of the visceral mass, it imme- diately loops and runs back to the base of the foot. Then the gut ascends dorsally along the posterior wall of the visceral mass to the peri- cardial cavity. The hindgut rounds the adduc- tor not only on its dorsal side, as in all bivalves, but also to the ventral side. The rec- tum turns anteriorly and dorsally. There are no differences in transverse sec- tions of the gut immediately posterior to the stomach (Fig. 15) and across the ascending part (Figs. 12-14). The gut wall consists of a single layer of large cylindrical cells with dis- tinct nuclei and transparent or stained cyto- plasm. In diagonal and longitudinal sections, the cell aggregations look like twisted, in- tensely staining fibrous bands with the dense cytoplasmic inclusions. The outer wall of the gut is covered by a thin membrane. A similar membrane appears to line the inner gut wall. No cilia and no typhlosole were found in the lumen of the intestine. The food wastes in the beginning of the mid- gut represent a thickened grayish mass con- sisting of fragments of algal thecas. Contents of the ascending section are brownish in color. Gaps outlining the borders of the pellets ap- pear in the contents. There are isolated oval dark or brownish pellets in the hindgut. The digestive diverticula consist of two lobes, which unite under the umbo and sur- round the stomach laterally. The lobes taper ventrally and extend almost to the base of the foot. In some individuals, large triangular- rounded cells of the diverticular gland together form oval rosettes. In sections transverse to the stomach wall, a branching net of canals located among the cell groups and converging towards the stomach wall can be observed. There are vacuoles, faintly staining nuclei and small dark cytoplasmic granules within the cells. In other individuals (Fig. 11), the thin gland has short broad canals and large cells, the borders of which are indistinct. This ap- pears to be caused by autolysis. In this case, the gland cells contain numerous transparent brownish granules, among which the deeply stained granules form aggregations. Digestive ducts and tubules typical of most bivalves were not found. Pericardium and Kidneys The pericardial cavity lies anterior and dor- sal to the posterior adductor (Figs. 3, 4). Within the cavity, there is a transparent ven- tricle, through which the rectum passes, and two thin-walled postero-ventral auricles. The kidney consists of two elongated distal limbs and lies posterior to the pericardium. Within the limbs, faintly staining floccular tissue and large excretory cells with the brownish or dark granules occur (Fig. 16). Each distal limb ex- tends laterally to the terminal cuticula of the gill forming the cloaca and anteriorly to the posterior wall of the visceral mass. In the ven- FIG. 16. Kidney. Transverse section through distal part of left limb and terminal cuticula of gill. Bar = 50 um. 2 EVSEEV ET AL. tral part of the cloaca, the kidney limbs only join the terminal cuticula of the gill, and the renal openings are located near the base of the exhalant siphon. Reproductive System The species is dioecious; it becomes mature at а shell length of approximately 0.8-1.0 mm. The ovary lies in the visceral mass and sur- rounds the digestive diverticula and the stom- ach laterally and posteriorly. The anterior part of the ovary is located in the subumbonal en- largement of the visceral mass. Ventrally, the ovary borders the base of the foot. The testis is similarly located in the visceral mass. The walls of the follicle of the ovary or testis are very thin. In the ovary, the oval and oval- angular oocytes were predominant among the detached oocytes. The large growing oocytes of 30-40 um, sometimes 45 ит, diameter with well-marked nuclei and nucleoli were attached to the follicle wall. No пре oocytes were found in sections. Unlike the ovary, the testis was filled by both spermatocytes | and II and by nutritional cells. Ripe spermatozoa with the large heads occurred in center of the follicle. The largest spermatocytes were 6-7 ит in diameter. Their large nuclei were lightly stained, with small nucleoli and granules of chromatin. Gonial and nutritional cells were attached to the wall. DISCUSSION Alveinus ojianus is a member of the family Kelliellidae, which appears to consist of 3—4 genera and 23-25 species, including fossil taxa (Habe, 1953, 1977; Keen, 1969; Bernard, 1989; Hayami & Kase, 1993; Allen, 2001; Oliver & Zuschin, 2001). The taxonomic status and phylogenetic relationships of this family still remain insufficiently well-founded and this could be the result of poor data on the internal comparative and functional morphology both of this species and the family as a whole. As a result, the taxonomic significance of some internal features of the Kelliellidae that could also be useful phylogenetic characters is not yet determined. The mantle and apical glands as well as the extracellular mantle granules of A. ojianus are examples of such features of unclear taxo- nomic significance. A mantle gland that is structurally and topologically similar to that of A. ojianus is known not only in such sister families as the Vesicomyidae (Morton, 1986; Allen, 2001), but also in the more distant Hiatellidae, Crassatellidae, Carditidae, Thyasi- ridae and Verticordiidae (Pelseneer, 1906; Allen, 1968; Yonge, 1969, 1971; Allen & Turner, 1974). In these taxa, mucus secreted by the gland is used for pseudofaeces forma- tion, for attaching sand grains to a shell and, probably in the Verticordiidae, for encapsula- tion of motile prey caught in the mantle cavity. There are also species (for example, Turtonia minuta), in which only the female mantle gland takes part in the formation of brooding capsules (Oldfield, 1955, 1963). This species also has an apical gland. None of the above taxa, including Т. minuta, contain extra- cellular granules, such as are found in the thickened inner mantle fold of A. ojianus and possibly in other species of Kelliellidae (Clausen, 1958). These granules begin to form at the end of September or the beginning of October when the shell length of the juvenile A. ojianus ex- ceeds 350-400 um. In sections through the mantle of adults, the granules are located both on and under the mantle fold and on the epi- thelium. In some sections of the gland, there are “empty places”, which are similar to the granules in shape, size and location. Brood- ing, in which these granules might be used for nutrition of the young, as for instance in the eggs of some gastropods (Thorson, 1936), is absent in A. ojianus. In this species, there is no hypobranchial gland that can be used for nutrition of the brooded young (Owen, 1961; Morton, 1977, 1982). The pelagic larvae of A. ojianus occur in Peter the Great Bay in August and settle at a shell length about 230-240 um. Formation of the granules at the beginning of the thermal minimum and their absence in spring indicate that the granules can be used as a food resource not for the young, but for the adult mollusk itself during the winter game- togenesis. Their presence is also independent of sex. The stomach is another organ that is impor- tant in the taxonomy of A. ojianus. Its morphol- ogy noticeably differs from those in related families Arcticidae, Glossidae, Trapezidae and Veneridae (Purchon, 1960; Reid, 1965). There is no cuticular lining or helicoid structure in the stomachs of members of these families, but they possess digestive pouch, dorsal hood, caecum, gastric shield, and sorting areas con- sisting of ridges and grooves. The stomach of АМАТОМУ OF ALVEINUS OJIANUS 13 Vesicomyidae has not been studied. But in a transverse section, the digestive tubules of large vesicomyids, for example Calyptogena (Morton, 1986), are similar to the tubules of most bivalves. The mid-gut, hindgut and rec- tum of Calyptogena differ from the gut of A. ojianus in having a folded, ciliated epithelium on the inner wall and in the presence of a typhlosole in the rectum or by absence of the anterior loop of the gut as, for instance, in Isorropodon (Cosel & Salas, 2001). The slit- like aperture of the style sac opening is not in the mid-gut, as in many bivalves, but in the posterior section of the stomach. It is a re- markable distinguishing feature of the alimen- tary canal of A. ojianus and, possibly, other members of Kelliellidae. Taking into account the general configuration and composition of functionally important sections of the stomach, which have been used as taxonomic charac- ters of suborders or orders in the Bivalvia (Purchon, 1978, 1987; Starobogatov, 1992), as well as other internal features — siphons, papillae, mantle apertures, labial palps, termi- nal cuticula of the gill, foot and its glands (Table 1) — the inclusion of Kelliellidae in the same subdivision of the Veneroida as the above families may be considered as insuffi- ciently founded. On the other hand, some morphological fea- tures of A. ojianus that seem to be taxonomi- cally important in comparison to veneroid families, occur in more distant phylogenetic lines. For instance, the cuticular lining of the stomach wall of A. ojianus and, possibly, other species of Kelliellidae is most similar either to that of more primitive Nucinellidae, Sole- myiidae and Nuculidae, or of the specialized Verticordiidae, Poromyidae and Cuspidariidae (Starobogatov, 1992). A more detailed com- parison shows that, like A. ojianus, the stom- ach of primitive taxa has no caecum, no digestive pouch, and no large ciliary fields, but has sorting ridges, large pouch-like digestive tubules, and small embayments in the right- dorsal wall resembling the dorsal hood (Purchon, 1956; Allen & Sanders, 1969). In addition to the cuticular lining occupying a small part of the stomach wall, a common gas- tric shield joined with the underlying columnar cells by means of pseudociliary cuticular connectives may also occur in stomach of the primitive mollusks (Halton & Owen, 1968). However, the cup-shaped helicoid structure is lacking, and the general configuration of the stomach of the primitive bivalve differs mark- edly from that of A. ojianus and other taxa, such that the former could be separated as a special subclass based on their morphology and digestion features alone (Purchon, 1987). As in A. ojianus, the stomachs of specialized taxa (e.g., Verticordiidae) have no dorsal hood, caecum, sorting areas and digestive tubules similar to those of other bivalves (Allen & Turner, 1974). The cuticular (scleroprotein) lining may bear the irregular, almost parallel ridges covering the most part of the anterior section of the stomach or fan-like divergent curved ridges resembling those located at the bottom of the A. ojianus helicoid structure. The cuticular lining of the verticordiid stomach also has spiral structures (Allen & Turner, 1974: fig. 63), which are comparable with the helicoid structure of A. ojianus in shape and location. In addition to the dorsal opening, the style sac seems to have an anterior slit-like aperture that opens into the stomach, as in A. ojianus (Allen & Turner, 1974: figs. 19, 77). Other simi- larities to A. ojianus include a mid-gut consist- ing of columnar cells lacking cilia, although in some verticordiids a typhlosole may occur in- side the gut. Also, the labial palps of the Verticordiidae are usually funnel-shaped and without the sorting ridges, but possess wing- shaped processes, buccal cavities or special glands used for capture and partial digestion of motile prey. In addition to the above features, as well as valves and additional tentacles in the mantle apertures that are features concerned with nutritional specialization at specific or generic levels (Allen & Turner, 1974), the Verticor- diidae have important characters at the famil- ial level that are absent in A. ojianus. These are a thickened muscular wall surrounding the stomach and oesophagus, dilatation of the hindgut — an analogue of the “masticatory stomach’ of Cephalaspidea (Ivanov, 1985) — а radial mantle gland consisting of separate is- lets, and a shell, which morphologically and structurally differs from the Kelliellidae, such that inclusion of these families within the same order is not possible. Therefore, although there is no relationship between the Kelliellidae and the Verticordii- dae, a similar evolutionary route can cause a high degree of likeness in the internal mor- phological features of these taxa (Table 1). One of the features of this evolutionary path is their small body size as, for instance, in many species of Kelliellidae, Verticordiidae, and Vesicomyidae, in which the average shell 14 EVSEEV EWAE TABLE 1. The internal morphological features of the Kelliellidae, Veneroida and Verticordiidae and their significance in the taxonomy and evolutionary development of Kelliellidae. (+) — feature present; (+) — feature may be absent; (-) — feature absent; (*) — feature has significance; (+) — feature has no significance; (?) — significance of feature not determined. Taxa Significance Paedo- Morphological Features Kelliellidae Veneroida Verticordiidae Taxonomy morphosis Mantle Organs Mantle gland + E E $ = Extracellular granules + - ® % Only exhalant siphon + - + er = Apical siphonal papillae - + - = = Ваза! siphonal papillae os 2 + ii ы Ciliated mantle lobes - + - + 2 Mantle Cavity Organs Inner and outer branchial lamellae differ in size - + + Ё is Only inner branchial lamellae + - = si e Ventral marginal food groove + + + 2 ? Free posterior gill end 2 + = 2 ? Homorhabdic branchial filaments + - + A h Adductor consisting of one portion + - E Е x Visceral Mass Organs Byssal gland + - + 2 2 Apical gland + - iS É Ventral foot groove - - + 3 # Ciliated foot er + - Y # Statocysts of B2-type ar ? ? Alimentary Canal Organs Funnel-shaped labial palps 7 - а = A Sorting ridges of labial palps m: + + 5 = Labial muscle rim - + - = = Cuticular lining of stomach + - + 5 ? Gastric helicoid structure Fe - 3 ? Digestive pouch - + - = ei Dorsal hood + Eb + = Caecum of stomach - + + 2 Sorting areas of stomach - + - + E Typhlosoles of stomach Sa + + = ? Digestive tubules a + + ? + Crystalline style sac combined with gut - = 2 E z Crystalline style sac combined with stomach + = Y 4 Typhlosole of gut + > + 2 Dilatation of gut - - E 2 ANATOMY OF ALVEINUS OJIANUS 15 length does not exceed 4-5 mm (Knudsen, 1970; Allen & Turner, 1974; Bernard, 1989; Warén, 1989; Hayami & Kase, 1993; Allen, 2001; Cosel & Salas, 2001). Mollusks of such a size usually have either only an inner demibranch or an inner and underdeveloped outer demibranch. The number of the gill bran- chial filaments rarely exceeds 22-23. At the same time, the branchial axis is located dor- sally, and the posterior filaments are replaced by a membrane or differ from other filaments in their shape and ciliary system (Pelseneer, 1906). The adductors of these bivalves some- times consist only of the outer portion of smooth muscle fibres, which in ontogenesis were previously the internal portion of cross- striated fibres (Oldfield, 1955). In these mol- lusks, the pedal gape of the mantle cavity may be not separated from the inhalant aperture (Clausen, 1958). In cases in which the pedal даре 1$ separate, the inhalant siphon 1$ usu- ally absent, and the exhalant siphon repre- sents a simple tube without the apical papillae, as in juvenile Veneridae (Ansell, 1962). The alimentary canal is also characterized by such “juvenilization” (Table), that is, paedo- morphosis (De Beer, 1958). The paedomor- phic features include the funnel-shaped labial palps, the total or partial absence of the sort- ing ridges on the palps, the morphologically indistinct mouth without the labial muscle rim, the epithelium of the oesophagus covered with long cilia, which occurs in macroforms in the veliger or pediveliger stages, the absence of the digestive pouch, caecae and sorting areas as, for example, in Erycinidae and other bivalve families (Oldfield, 1955, 1963; Chanley 8 Andrews, 1971; Alatalo et al., 1984). These features as well as the underdeveloped gill and adductors, the primitive inhalant aperture without protection of the mantle cavity by valves; the unciliated epithelium of the mantle cavity and intestinal tract and cloaca, and the accelerated reproductive development, which in A. ojianus matures at a shell length of about 1 mm, suggest incomplete somatic develop- ment of not only the internal organ, but also the outer skeletal organ, the shell. Thus, the Verticordiidae and the Kelliellidae are of different phylogenetic lines, but with a similar pattern of evolutionary development - paedomorphosis, in which the post-juvenile stages prove to be “cut” (De Beer, 1958; Gould, 1977) in taxa of both families. There- fore, based on internal morphological charac- ters, an attempt to determine the place of Kelliellidae in the taxonomic system rested on the conchological features of adults, meets with failure. This is caused by insufficiently studied internal organs of bivalves as a whole. In conchological features, the family Kellielli- dae appears to be among the most primitive Heterodonta, but it is impossible to determine its place more distinctly because of poorly studied juvenile stages of Heterodonta with morphological characters similar to those of adult kelliellids. ACNOWLEDGEMENTS We greatly appreciate Prof. John A. Allen (University Marine Biological Station, Millport) for critical reading the manuscript and very useful comments. We also thank him for send- ing reprints of papers. We are very grateful to Dr. Paula M. Mikkelsen (American Museum of Natural History, New York), Dr. Eugene V. Coan (California Academy of Sciences, San- Francisco) and anonymous reviewer for their helpful comments, beneficial criticism and cor- rections. We thank Dr. Konstantin A. Lutaenko (Institute of Marine Biology, Vladivostok), who provided us with benthic samples from the research vessel “Lugovoye”. We are grateful to Dr. Vladimir V. Malakhov (Moscow State University, Moscow) for his consultation and support of our work. We thank Dr. Alexander |. Kafanov (Institute of Marine Biology, Vladi- vostok), Dr. P. Graham Oliver (National Mu- seum of Wales, Cardiff), and Dr. Rudo von Cosel (Museum National D'Histoire Naturelle, Paris) who provided us with reprints of papers. 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Revised ms. accepted 4 July 2003 + ya | Ñ e VU ‘ | Zr: О OG A = > = . o | e > pe BEER u rod o = В ia | oh y 7 y : A м @ ui fes: must JAN Y DS A PPD ste) u | | | О cue EMM Mie? qe “4 av D =a oF | >) E tf Lu MALACOLOGIA, 2004, 46(1): 19-35 MORPHOLOGICAL AND MOLECULAR ANALYSIS OF THE STATUS AND RELATIONSHIPS OF OXYCHILUS PAULUCCIAE (DE STEFANI, 1883) (GASTROPODA: PULMONATA: ZONITIDAE) Giuseppe Manganelli', Simone Cianfanelli?, Nicola Salomone* & Folco Giusti! ABSTRACT Morphological data shows that О. paulucciae (De Stefani, 1883) belongs to Oxychilus ($. str.), sensu Giusti & Manganelli (1999), and is distinguished from sympatric, similarly shelled species, such as О. draparnaudi (Beck, 1837) and О. meridionalis (Paulucci, 1881), by its larger shell (diameter: 13.9-17.4 mm), smaller umbilicus (about 1/8 of shell diameter), narrow mid-penial region, internal ornamentation of proximal penis consisting of longitudinal pleats, and a less developed vaginal gland, often forming an incomplete ring around proximal vagina. DNA sequence data, analysing the ITS-1 region in two specimens of О. paulucciae and representatives of several other species occurring in Tuscany, О. draparnaudi, О. majori (Paulucci, 1886), O. meridionalis, O. pilula (Paulucci, 1886) and O. uzielli (Issel, 1873), indicates that O. paulucciae represents a well-differentiated evolutionary lineage and suggests it has close relationships with O. meridionalis and O. uziellii. Finally, analysis of morphological characters and DNA sequencing data demonstrates that Oxychilus lanzai Forcart, 1967, is a junior synonym of O. paulucciae. Key words: Zonitidae, Oxychilus paulucciae, Oxychilus lanzai, systematics. INTRODUCTION During the 1960s, the malacologist L. Forcart received many specimens of Tuscan Oxychilus, most collected by Prof. B. Lanza (Museo Zoologico de “La Specola”, Universita di Firenze) in NW Tuscany (provinces of Massa Carrara, Lucca and Florence). Based on these specimens, Forcart (1967) produced a first revision of many taxa of the species group described from Tuscany in the nineteenth century (Table 1), and described a new species: O. (Ortizius) lanzai Forcart, 1967. In the late 1960s, one of us (FG), together with M. Mazzini, was involved in the study of the malacofauna of the Apuan Alps (NW Tuscany) as part of a project promoted by the Società Italiana di Biogeografia (Giusti & Mazzini, 1971). The study of the Oxychilus material collected on this occasion led to revision of that studied by L. Forcart (1967, 1968). It became evident that Forcart, using diaphanized preparations of the whole distal genitalia mounted on glass slides, sometimes leading Swiss misinterpreted the internal structure of the penis, which was then considered very important for the diagnosis of subgenera: parallel rows of papillae for Oxychilus s. str.; parallel uninterrupted pleats for Ortizius Forcart, 1957. This happened for specimens from Tana di Magnano, Garfagnana, Province of Lucca, which he assigned to O. (Oxychilus) paulucciae (De Stefani, 1883), but when re- examined turned out to be a species of Ortizius, anatomically identical to that from the Grotta della Risvolta, Apuan Alps, which he assigned to O. lanzai. Giusti 8 Mazzini (1971) declined to express formal synonymy between O. /anzai and O. paulucciae, because they realized that since Forcart did not have topotypical specimens of the classic Tuscan taxa for anatomical inves- tigation, he based his study on spirit material with shells similar to those of the types, thus misinterpreting some classic species (shell shape is very rarely diagnostic in species of Oxychilus). They merely stated that the entire group of nominal species described for NW Tuscany required revision before the problem ‘Dipartimento di Scienze Ambientali, Universita di Siena, Via Mattioli 4, 53100 Siena, Italy; manganelli@unisi.it ?Museo Zoologico de “La Specola”, Sezione del Museo di Storia Naturale dell'Università di Firenze, Via Romana 17, 1-50125 Firenze, Italy 3Dipartimento di Biologia Evolutiva, Universita di Siena, Via Mattioli 4, 53100 Siena, Italy 20 MANGANELLI ЕТ AL. TABLE 1. Nominal taxa of the species group introduced for Tuscan Oxychilus (excluding those established for species living in the Tuscan Archipelago) (for syntypes kept in the Museo di Zoologia “La Specola”, the collection number is followed by the number of specimens). Nominal taxon Status Zonites Uziellii Issel, 1872: 60-61. Oxychilus uziellii (\ssel, 1872) Type material: lectotype (MZUF 689) and one paralectotype (Manganelli & Giusti, 1985, (MZUF 11521) in Paulucci collection. 1993, 2001) Type locality: “Fra i detriti del Gombo, presso Pisa”, but see Manganelli & Giusti (2000). Hyalina scotophila De Stefani, 1879: 38-39. probably junior synonym of Type material: 4 syntypes; one in Paulucci collection (MZUF Oxychilus draparnaudi (Beck, 738) and three in Museo di Storia Naturale dell Accademia dei 1837) (Manganelli & Giusti, Fisiocritici in Siena. 2001) Type locality: “Siena, in un profondo condotto sotterraneo” Hyalinia meridionalis Paulucci, 1881: 78-79, pl. 1, fig. 6. Oxychilus meridionalis Type material: lectotype (MZUF 13187) and 30 paralectotypes (Paulucci, 1881) (Manganelli in Paulucci collection (781/12, 828/7, 829/1, 830/3, 832/2, 8 Giusti, 2001) 13188/5). MZUF 781, 829, 830, 832, 833 belong to other species (Manganelli 8 Giusti, 2001). Type locality: “Fabbriche presso i Bagni di Lucca”. Hyalinia Isseliana Paulucci, 1882: 165-168, pl. 9, fig. 13. junior synonym of Oxychilus Type material: lectotype (MZUF 687) and 4 paralectotypes meridionalis (Paulucci, 1881 (MZUF 688/3, 13346/1) in Paulucci collection (Manganelli & (Manganelli & Giusti, 2001) Giusti, 2001). Type locality: “Fabbriche presso i Bagni di Lucca (Lucca; Toscana). Hyalinia Guidoni De Stefani, 1883: 35, 1888: fig. 3. nominal taxon in need of Type material: unknown. revision Type locality: “Forno Volasco, 480 [т]”. Hyalinia Paulucciae De Stefani, 1883: 35-36, 1888: fig. 1. Oxychilus paulucciae (De Type material: no syntype being known, a neotype was Stefani, 1883) (this paper) designated (Fig. 1). The neotype (a spirit specimen) 1$ in the Museo Zoologico de “La Specola”, Sezione del Museo di Storia Naturale dell’Universitä di Firenze (Italy) (MZUF 17597) Type locality: “Alp. E. Vagli 850” (p. 36) and “Strada nazionale presso il Ponte di Ceserana” (caption of un-numbered plate). “Alp. E” is for “Pendici orientali delle Alpi Apuane dall'alveo del fiume Serchio e dell'Aulella fino alla crina” (p. 17). Following the designation of the neotype, the type locality becomes “Vagli di sopra, Valle Arnetola, 930 т as! (Vagli di sotto, Lucca), 32TPP0084”. Hyalina scotophila var. пота Paulucci, 1886: 12-13, pl. 1, fig. 2. junior synonym of Oxychilus Type material: 27 syntypes (MZUF 788/4, 789/9, 790/5, 791/7, draparnaudi (Beck, 1837) 792/1, 13347/1) in Paulucci collection. The shell MZUF 792 is (Giusti, 1968; Manganelli et that illustred by Paulucci (1886). al., 1995) Type locality: “alla Fonte dell’Appetito presso Porto Santo Stefano, presso la vetta del Telegrafo, sopra al Convento de’ Passionisti, in vicinanza delle scogliere di Calagrande ... Allisola del Giglio in una localita denominata «Franco». (Continues) REVISION OF OXYCHILUS PAULUCCIAE 2A (Continued) Nominal taxon Hyalinia nitidula var. amiatae Westerlund, 1886: 57. Type material: 33 syntypes in Paulucci collection (MZUF 807/12, 19290/10, 19291/11). Status junior synonym of Oxychilus draparnaudi (Beck, 1837) (Manganelli et al., 1995) Type locality: “Italien, М. Amiata” [in localita La Scarpa]. Hyalinia sylvicola Westerlund, 1886: 59. Type material: 8 syntypes in Paulucci collection (MZUF 805). nominal taxon in need of revision Type locality: “Italien, Bosco di San Vittore in Toskana”. [= Pozza delle Monache, Bosco di San Vettore]. Hyalinia blauneri var. cloacarum Westerlund, 1886: 61. Type material: 8 syntypes in Paulucci collection (MZUF 804) nominal taxon in need of revision Type locality: “Italien b. Volterra” [= Fogna della Chiesa di Camporbiano]. Hyalina scotophila var. dilatata Westerlund, 1886: 61. Type material: 8 syntypes in Paulucci collection (MZUF 19289). nominal taxon in need of revision Type locality: “Ital., San Martino b. Palma” [= San Martino alla Palma] Oxychilus (Ortizius) lanzai Forcart, 1967: 114—115, fig. 1, pl. 1, fig.1. Type material: The holotype (spirit specimen, MZUF 462), seven paratypes (spirit specimens, MZUF 454/2, 463/5 from junior synonym of Oxychilus paulucciae (De Stefani, 1883) (this paper) “Grotta del Buggine” and six paratypes (four spirit specimens, MZUF 683; two shells, MZUF 691) from “Grotta della Risvolta” are in the Museo Zoologico de “La Specola”, Sezione del Museo di Storia Naturale dell’Universita di Firenze (Italy). Four other paratypes from “Grotta del Buggine” (two spirit specimens, NMB 6562: two spirit specimens, MFP) are in the Naturhistorisches Museum Basel (Switzerland) and in the Museo “Felice Роеу”, La Habana (Cuba) respectively. Type locality: “Toskana, Prov. Lucca, Apuaner Alpen, Grotta del Buggine 315 т (N. 166 T.) bei Cardoso Stazzemese”. Oxychilus (Ortizius) tongiorgii Giusti, 1969: 367-369, figs. 1-2, 5A, pl. 1, figs. 1, 2. Type material: holotype and 9 paratypes in Giusti collection. junior synonym of Oxychilus meridionalis (Paulucci, 1881) (Manganelli & Giusti, 2001) Type locality: “Grotta dei Ladri (п. 262 T. Pi) Monti Pisani nei pressi di Asciano”. Oxychilus (Ortizius) forcartianus Giusti, 1969a: 369-371, figs 3, 4, 5B, pl. 1, figs. 3, 4. Type material: holotype and 3 paratypes in Giusti collection. junior synonym of Oxychilus meridionalis (Paulucci, 1881) (Manganelli & Giusti, 2001) Type locality: “Grotta dei Fiorentini ргеззо Pomarance (Grosseto)”. of the relationships between O. paulucciae and O. lanzai could be tackled. The oldest established Tuscan Oxychilus species — Zonites uziellii Issel, 1872; Hyalinia meridionalis Paulucci, 1881; and Hyalinia isseliana Paulucci, 1881 — have now been re- vised (Manganelli & Giusti, 1985, 1993, 2000, 2001), and it is therefore possible to revise O. paulucciae, to clarify its relationships with other Tuscan Oxychilus and to resolve the problem of its synonymy with O. /anzai. The first problem with Hyalinia paulucciae is that its type-material has not been traced. The malacological collection of De Stefani, which remained in Pisa when he moved to Siena and then Florence, was irreparably damaged during the Second World War. In order to de- fine this nominal taxon objectively, we have selected a neotype, because the “qualifying conditions”, required for the designation of a neotype, exist (ICZN, 1999: Art. 75.3). The neotype, the shell and genitalia of which are 22 MANGANELLI ЕТ AL. shown in Figure 1 (shell) and Figures 3-4 (genitalia), is deposited in the Museo Zoologico de “La Specola”, Sezione del Museo di Storia Naturale dell’Universitä di Firenze, Italy (catalogue no. 17597). It is a spirit speci- men collected near Vagli, one of the two locali- ties where De Stefani reported his species. lts shell matches the original description perfectly. MATERIAL AND METHODS Morphological Analysis Whole shells were photographed under the light microscope (Wild M5A). All dimensions — NW number of whorls (Ehrmann, 1933: fig. 12), SD shell diameter, SH shell height and UD umbilicus diameter — were measured us- ing a micrometer. Live specimens were drowned in water, then fixed and preserved in 75% ethanol buffered with sodium carbonate. The bodies were iso- lated after crushing the shells and dissected under the light microscope (Wild M5A) using thin-pointed watchmaker's tweezers. Anatomi- cal details were drawn using a Wild camera lucida. Some parts of the genital organs — duct of bursa copulatrix, distal vagina, epiphallus, flagellum, proximal portion of penis, “bottle- neck”, distal penis and penial sheath — were measured by micrometer. Radulae were extracted manually from buc- cal bulbs, washed in 75% ethanol, mounted on copper stubs with electronconductive glue, sputter-coated with gold and photographed using a Philips 505 SEM. All specimens listed in material examined belong to anatomically determined popula- tions. The material examined 1$ listed as fol- lows: locality, municipality and province names in parenthesis, UTM reference, collector(s), date, number of specimens in parenthesis (sp spirit preserved specimen/s, sh shell/s) and bibliographical reference, in parenthesis, if they are voucher specimens. Locality names and UTM references are according to the of- ficial 1:25,000 scale map of Italy (series М 891). Key to museum and collection acronyms: FGC, collection F. Giusti, Dipartimento di Scienze Ambientali, University of Siena, Italy; NMB, Naturhistorisches Museum Basel, Swit- zerland; MFP, Museo “Felice Poey”, La Habana, Cuba; MZUF, Museo Zoologico “La Specola”, Sezione del Museo di Storia Natu- rale dell'Universitá di Firenze, Italy; SCC, S. Cianfanelli collection, Firenze, Italy. Key to acronyms in figures: В, “bottle-neck”; BC, bursa copulatrix; BS, “bottle-neck” sheath; BW, body wall; DBC, duct of bursa copulatrix; DP, distal portion of penis; E, epiphallus; EO, epiphallus opening; F, flagellum; FO, free ovi- duct; POS, prostatic portion of ovispermiduct; PP, proximal portion of penis; PR, penial re- tractor; PS, penial sheath; UOS, uterine por- tion of ovispermiduct; V, vagina; VD, vas deferens; VG, vaginal gland. Molecular Analysis DNA Extraction, PCR and Sequencing Two specimens of O. paulucciae and others of several species occurring in Tuscany — O. draparnaudi (Beck, 1837), O. majori (Paulucci, 1886), O. meridionalis (Paulucci, 1881), O. pilula (Paulucci, 1886), and O. uzielli (Issel, 1873) — were used for molecular analysis. One specimen of O. paulucciae (O. paulucciae 1) was collected in the type locality of this species and the other (O. paulucciae 2) in a cave where part of the type material of O. lanzai was col- lected. Collection sites and codes of the samples used for molecular analysis are indi- cated in Table 2. Total DNA was extracted from fresh foot muscle using standard phenol/chlo- roform and ethanol precipitation methods as described in Salomone et al. (2002). We ampli- fied the ITS-1 region by PCR using the primer pair CS249 (5 TCGTAACAAGGTTTCCG3’) and DT421 (5 GCTGCGTTCTTCATCG3') (Schlôtterer et al., 1994). PCR amplification was performed in a reaction volume of 50 ul following a profile consisting of 25 cycles with temperatures of 95°C for 20”, 55°C for 30” and 72°C for 30”, plus a final extension step at 72°C for 5’. The products obtained using these con- ditions were very clean single bands, showing no evidence of double or ambiguous bands. After elimination of excess nucleotides and primers by gel separation and purification with Nucleospin Extract (Genenco) columns, both strands of the final products were sequenced using the two amplification primers. Sequenc- ing reactions were performed at the core facil- ity of MWG-BIOTECH, Ebersberg, Germany. All sequences were checked manually for se- quencing errors and submitted to GenBank (Accession Nos. AY373635-AY373645). REVISION OF OXYCHILUS PAULUCCIAE 23 TABLE 2. Material examined for DNA sequencing. Taxon Locality Oxychilus draparnaudi 1 Castello di Brolio (Gaiole in Chianti, Siena), 32TPP9909, G. Manganelli & L. Manganelli leg. 01.10.2000 Oxychilus draparnaudi 2 Giglio Island: Giglio Castello (Isola del Giglio, Grosseto), 32TPM5692, V. Vignoli leg. 30.06.2000 Oxychilus та/оп Monte Argentario: Grotta di Punta degli Stretti 250 T/GR (Monte Argentario, Grosseto), 32TPN7800, $. Cianfanelli 8 С. Manganelli leg. 07.10.2001 Oxychilus meridionalis 1 Сараппо (Castelnuovo Berardenga, Siena), 32TPP9308, С. Manganelli 8 L. Manganelli leg. 28.05.2000 Oxychilus meridionalis 2 Passo della Calla (Santa Sofia, Forli), 32TQP2060, $. Cianfanelli & С. Manganelli leg. 16.09.2001 Oxychilus meridionalis 3 Fabbriche di Bagni di Lucca (Bagni di Lucca, Lucca), 32TPP3175, $. Cianfanelli & E. Lori leg. 15.04.2003 Oxychilus paulucciae 1 Apuane Alps: Vagli di sopra, Valle Arnetola (Vagli di sotto, Lucca), 32TPP0084, $. Cianfanelli & М. Calcagno leg. 24.06.2000 Oxychilus paulucciae 2 Apuane Alps, Grotta della Risvolta 158T/LU (Stazzema, Lucca), 32TPP0372, $. Cianfanelli М. Calcagno leg. 14.10.2001 Oxychilus pilula Capraia Island: Il Laghetto (Capraia Isola, Livorno), 32TNN6665, F. Barbagli & S. Lotti leg. 22.06.2001 Oxychilus uziellii Fosso delle Filicaie, San Giusto in Salcio (Gaiole in Chianti - Radda in Chianti, Siena), 32TPP9115, G. Manganelli & L. Manganelli leg. 28.05.2000 Retinella olivetorum Fosso delle Filicaie, San Giusto in Salcio (Gaiole in Chianti - Radda in Chianti, Siena), 32TPP9115, G. Manganelli & L. Manganelli leg. 28.05.2000 Phylogenetic analysis Sequences were aligned using Clustal W (Thompson et al., 1994) and slightly modified by eye. Boundaries of the ITS-1 region were estimated by comparison with those deter- mined for Albinaria caerulea (Deshayes, 1835) (Genbank Acc. No. AF136012). Phylogenetic analyses were performed with PAUP* (version 4.0610; Swofford, 2001) with Retinella olivetorum (Gmelin, 1791) as outgroup, using maximum parsimony (MP) and maximum like- lihood (ML). MP reconstruction was performed by an exhaustive search of the most parsimo- nious tree(s) with equal weighting of all charac- ters. Gaps were treated as missing data. For ML analysis, the appropriate substitution model of DNA evolution that best fitted the data set was determined by the likelihood ratio test and the Akaike information criterion (AIC; Akaike, 1974) with Modeltest 3.04 (Posada & Crandall, 1998). A ML heuristic search (Step-Add ran- dom; TBR branch swapping) was then run un- der the likelihood setting estimated by Modeltest. Support for individual nodes was evaluated by bootstrap analysis (heuristic search) with 1000 replications. RESULTS Redescription of O. paulucciae (De Stefani, 1883) Identification A medium-sized species of Oxychilus (s. str.), sensu Giusti & Manganelli (1999), a “sub- genus” of Oxychilus characterized by: penis with flagellum; penial retractor inserted at apex of flagellum; internal ornamentation of proximal penis consisting of pleats or pleats and rows of papillae without apical thorns; epiphallus usually longer than proximal penis, its internal wall with slender longitudinal pleats; mucous gland mainly vaginal; long mesocone of central tooth). Oxychilus paulucciae is identified with respect to similar- shelled sympatric species (O. draparnaudi 24 MANGANELLI ET AL. and ©. meridionalis) by a larger shell (shell diameter: 13.9-17.4 mm) with small umbilicus (about 1/8 of shell diameter), narrow mid-pe- nial region, internal ornamentation of proximal penis consisting of longitudinal pleats, and vaginal gland often forming incomplete ring around proximal vagina. Description Body pale gray in colour; neck and upper part of sides with variably wide areas with pits (with phylacites); foot slender, of aulacopod type, with sole longitudinally tripartite (central part whitish, lateral parts pale gray); eyes FIGS. 1, 2. Two shells of Oxychilus paulucciae (De Stefani, 1883) from Vagli di sopra, Valle Arnetola, 930 т asl (Vagli di sotto, Lucca), 32TPP0084, $. Cianfanelli & M. Calcagno leg. 8.10.2000 (MZUF 17597, neotype) (FIG. 1) and Grotta della Risvolta, 220 т asl, no. 158T/LU (Stazzema, Lucca), 32TPP0372, М. Bodon 8 $. Cianfanelli leg. 9.11.1997 (FIG. 2). REVISION OF OXYCHILUS PAULUCCIAE 25 № PR 3 4 mm fil: 4-5 2mm ; à 4 M , à À Я 3 FIGS. 3-5. Distal genitalia (FIG. 3) and internal ornamentation of flagellum and proximal penis (FIGS. 4, 5) in specimens of Oxychilus paulucciae (De Stefani, 1883) from Vagli di sopra, Valle Arnetola, 930 т asl (Vagli di sotto, Lucca), 32TPP0084, $. Cianfanelli 8 М. Calcagno leg. 8.10.2000 (MZUF 17597, neotype) (FIGS. 3-4) and $. Cianfanelli & М. Calcagno leg. 24.6.2000 (FIG. 5). 26 MANGANELLI ET AL. present, normal in size; kidney sigmurethrous; jaw oxygnathous. Shell (Figs. 1, 2; Forcart, 1967: pl. 1, fig.1, figs. 1, 1a-1c [as O. lanzai], figs. 4, 4a—4c) dextral, medium in size, discoidal, de- pressed, thin and fragile, subtransparent, glossy when fresh, whitish-yellow or pale greenish, sometimes opalescent below; sur- face smooth, with variably evident growth lines and microsculpture consisting of very fine wavy spiral lines; spire usually tectiform, 5 1/12-5 1/2 whorls, gradually increasing in size, last whorl dilated near aperture, its last quarter descending slightly or not at all, rarely slightly angled at periphery; sutures shallow; umbilicus small, about 1.4-2.6 mm wide (usually 1/8-1/9, rarely 1/7 and in only one case 1/5 of maximum shell diameter), sometimes eccentric; aperture oval, oblique; peristome interrupted, simple, not thickened or reflected, its superior vertex starting at, or slightly above, periphery of last whorl. Di- mensions (30 shells measured). Number of whorls: 5 1/4 + 1/6 (5-5 5/6); shell diameter: 15.5 + 1.0 mm (13.9-17.4); height: 6.1 + 0.6 mm (5.3—7.2); umbilicus diameter: 1.9 + 0.2 mm (1.4—2.6). Genitalia (Figs. 3-12; Forcart, 1967: fig. 1 [аз O. lanzai], fig. 3). General scheme of geni- talia as in Oxychilus (s. str.), sensu Giusti & Manganelli (1999). Only distal genitalia are described here (a total of 21 adult specimens were dissected for study of genital structure during the various phases of the research). Female genitalia include free-oviduct, bursa copulatrix and its duct, and vagina. Distal free oviduct and most proximal vagina enveloped by muff of spongy glandular tissue forming vaginal gland; vaginal gland relatively unde- veloped, sometimes enough to form continu- ous ring around wall of most proximal part of vagina, distal part of free oviduct and of duct of bursa copulatrix, sometimes reduced to cover only one side (that facing free-oviduct) of proximal vagina and of distal duct of bursa copulatrix (large portion of wall on opposite side is uncovered); in both cases, vaginal gland often envelopes one side (that facing free oviduct) of distal canal of bursa copulatrix; duct of bursa copulatrix long (7.5 mm; п: 2), initially moderately flared, narrowing before entering oval or pyriform bursa copulatrix; dis- tal vagina (that without glandular muff) vari- ably long (2.7-4.9 тт; п: 2) and wide, reducing in calibre slightly or not at all near genital atrium. Male distal genitalia include vas deferens, epiphallus and penial complex (flagellum and penis). Epiphallus variably long (7.3-10.3 mm; п: 2) and slender, internal walls bearing series of very slender longitudinal pleats. Fla- gellum usually very long (3.7-4.7 тт; п: 2), with penial retractor muscle ending at apex (sometimes thin muscular branch extends on one side to end at about half flagellum length). Penis variably long (9.9-14.3 тт; п: 2) with clear distinction into proximal and dis- tal parts due to “bottle-neck” (terminal, slen- der part of proximal penis: minimum caliber recorded 0.5-0.62 mm: п: 4), enveloped by thin, distinct, translucent sheath. Proximal penis rather short (5.4-5.9 mm; п: 2). Distal penis variable in length (4.5-8.4 тт; п: 2), enveloped by variably long (2.5-4.1 тт; п: 2) penial sheath, proximally very thin, tra- versed on one side by vas deferens, then slightly thicker for rest of length. Internal sur- face of flagellum and proximal penis sur- rounding opening of epiphallus into penis with many small radially disposed pleats, sometimes fragmented into rows of variably large papillae; lateral surface, and that oppo- site opening of epiphallus into penis, having slender longitudinal pleats with jagged sides, frequently fragmented into rows of variably large papillae. A variable number (9-12) of these pleats continues on rest of proximal penis, converging, fusing and reducing in number before continuing, with a more or less marked interruption at “bottle-neck”, in- side distal penis, where they are usually wider with jagged sides. Very short, thin- walled duct connects distal penis (level with where penial sheath originates) to genital atrium in which vagina also ends. Radula consisting of many rows of about 31-35 teeth, according to formula: 11-13 M/1 + 0-1 LM/2 + 3-4 L/3 + C/3 + 3-4 L/3 + 0-1 LM/2 + 11-13 МЛ (6 specimens examined). Central teeth with well-developed basal plate, apical portion of which V-like, with pointed vertices; body of tooth wide, providing base for long, slender, pointed mesocone flanked by two very short ectocones. On both sides of each central tooth, three-four lateral tricuspid teeth, sometimes one latero-marginal bicuspid tooth and series of monocuspid marginal teeth in decreasing order of size. REVISION OF ОХУСНШИ$ PAULUCCIAE 27 FIGS. 6, 7. Distal genitalia (FIG. 6) and internal ornamentation of flagellum and proximal penis (FIG. 7) in a specimen of Oxychilus paulucciae (De Stefani, 1883) from Grotta del Buggine Stazzemese, 315 m asl, no. 166 T/LU (Stazzema, Lucca), 32TPP0573, B. Lanza & P. Lanza leg. 1960 (NMB 6562-a, paratype). 28 MANGANELLI ET AL. 9-10 2mm FIGS. 8-10. Distal genitalia (FIG. 8), internal ornamentation of flagellum and proximal penis (FIG. 9) and mid-penis region (FIG. 10) in a specimen of Oxychilus paulucciae (De Stefani, 1883) from Tana del Pollone di Magnano, 565 m asl, no. 1017 T/LU (Villa Collemandina, Lucca), 32TPP1592, B. Lanza leg. 30.4.72 (MZUF 15856). REVISION OF OXYCHILUS PAULUCCIAE 29 11 4 mm FIGS. 11, 12. Distal genitalia (FIG. 11) and mid-penis region (FIG. 12) in a specimen of Oxychilus paulucciae (De Stefani, 1883) from Grotta della Risvolta, 220 m asl, no. 158 T/LU (Stazzema, Lucca), 32TPP0372, М. Bodon & $. Cianfanelli leg. 9.11.1997. Material Examined PPO7 Grotta del Buggine Stazzemese, 315 т asl, no. 166 T/LU (Stazzema, Lucca), NP98 Buca della Freddana, 650 т asl, no. 32TPP0573, no collector and date (2 230 T/MS (Massa, Massa Carrara), sp, FGC), B. Lanza leg. 18.10.59 (2 sp 32TNP9783, G. Comotti leg. 12.7.85 [paratypes of Oxychilus lanzai], MZUF (1 sp, 2 sh, FGC). 454); В. Lanza & P. Lanza leg. 1960 (1 sp 30 PPO3 PPT PP19 PP28 МАМСАМЕН ET AL. [holotype of Oxychilus lanzai], MZUF 462; 5 sp [paratypes], MZUF 463; 2 sp [paratypes of Oxychilus lanzai], ММВ 6562-a). Grotta della Risvolta, 220 m asl, no. 158 T/LU (Stazzema, Lucca), 32TPP0372, В. Lanza leg. 23.7.61 (4 sp [paratypes of Oxychilus lanzai], MZUF 683; 2 sh [paratypes of Oxychilus lanzai], MZUF 691); В. Lanza leg. 15.3.64 (3 sp, FGC); B. Lanza leg. 21.12.69 (1 sp, FGC); M. Bodon & S. Cianfanelli leg. 9.11.97 (5 sp, 3 sh, SCC 8097/1742); S. Cianfanelli leg. 14.10.01 (1 sp, 29 sh, SCC 11647/2819). Vagli di sopra, Valle Arnetola, 930 т as! (Vagli di sotto, Lucca), 32TPP0084, $. Cianfanelli & M. Calcagno leg. 24.06. 2000 (2 sp, 2 sh, FGC; 22 sh, SCC 9328/2291; 7 sh, SCC 9330/2290; 1 sh, SCC 9485/2400), S. Cianfanelli & M. Calcagno leg. 8.10.2000 (1 sp, MZUF 17597 [neotype of O. paulucciae]; 7 sh SCC 9484/2399). Buca delle Fate di San Rocco, 635 m asl, no. 362 T/LU (Pescaglia, Lucca), 32TPP1170, P. Magrini leg. 22.8.79 (1 sp, 3 sh, FGC). Grotta della Faglia, Pania di Corfino (Villa Collemandina, Lucca), F. Utili leg. 24.11.63 (2 sp, FGC). Tana dei Gracchi di Sasso Rosso, 755 т asl, no. 289 Т/ LU (Villa Collemandina, Lucca), 32TPP1293, Е. Utili leg. 22.11.63 (1 sp, FGC). Tana del Pollone di Magnano, 565 т asl, no. 1017 T/LU (Villa Collemandina, Lucca), 32TPP1592, В. Lanza leg. 30.4.72 (8 sp, MZUF 15856; 1 sp, MZUF 15858). Tana di Magnano, 635 т asl, по. 162 ИО (Villa Collemandina, Lucca), 32TPP1592, В. Lanza & В. Malkin leg. 30.11.59 (1 sp det. O. paulucciae by Forcart, 1967, 1968, NMB 6561-a; 1 sp det O. paulucciae by Forcart, 1968, MZUF 445), F. Utili leg. 24.11.63 (5 sp, 1 sh, ЕСС), В. Lanza leg. 15.3.64 (2 sp, 2 sh, FGC), B. Lanza leg. 24.10.65 (1 sp, FGO), F. Utili leg. 5.12.65 (2 sp, 1 sh, FGC); P. Brignoli 8 A. Vigna Taglianti leg. 3.11.67 (3-sp, 3 sh, FGC): В, Lanza leg. 17.3.68 (11 sp, 20 sh, FGC). Grotta dell’Iseretta, 650 m asl, по. 823 T/LU (Bagni di Lucca, Lucca), 32TPP2882, P. Magrini leg. 6.74 (2 sp, FGC). Etymology De Stefani (1883) named this species after the famous Italian malacologist, Marquise Marianna Paulucci (1835-1919) and Forcart (1967) after Prof. Benedetto Lanza, former di- rector of the Museo Zoologico de “La Specola” (Florence), who collected the specimens used for the description. Habitat All the specimens of O. paulucciae were collected inside caves. The area inhabited by the species 1$ karstic. The species 1$ presum- ably adapted to subterranean life and may be defined as troglobie. Geographical Distribution Species with reduced distribution, limited to northwestern Tuscany (Fig. 13). Status and Conservation Not globally threatened. Despite its limited distribution, O. paulucciae does not seem to be under any particular threat at present. Molecular Data Sequence Data Analysis The ITS-1 region ranged from 544 (O. uziellii) to 593 bp (O. meridionalis 1 and O. meridionalis 2) in length. After deletion of am- biguously aligned positions, the data-set in- cluded a total of 660 nucleotide positions, 61 of which were phylogenetically informative under the parsimony criterion. Uncorrected percentage sequence divergence (p-distance) between the two O. paulucciae specimens (1, Vagli di sopra; 2, Grotta della Risvolta) was 1.1%. Oxychilus meridionalis 1 (Capanno) and O. meridionalis 2 (Passo della Calla) are separated by a sequence divergence 0.3%, whereas a divergence of 2.3% distinguished them from O. meridionalis 3 (Bagni di Lucca). Sequence difference between О. draparnaudi 1 (Castello di Brolio) and O. draparnaudi 2 (Giglio Castello) was 1.3%. Genetic distances between morphologically defined species ranged from 2.5% (O. meridionalis 2 and O. paulucciae 2) to 7.9% (O. draparnaudi 2 and O. majori). REVISION OF OXYCHILUS PAULUCCIAE 31 à к № Q т й À = À Nb FIG. 13. The distribution of Oxychilus paulucciae (De Stefani, 1883) on UTM map of central-northern Italy. Phylogenetic Analysis Maximum parsimony analysis supported a single best tree shown in Figure 14 (tree length: 297, CI: 0.926, RI: 0.788). The likeli- hood ratio test and AIC from Modeltest sup- ported the HKY+G model (Hasegawa et al., 1985, including among-site rate heterogene- Ку) аз the best fit substitution model for the data. Parameters estimated for this model were: Ti: Tv ratio = 0.789, gamma shape pa- rameter = 1.02 and base frequencies А = 0.1994, Т = 0.2592, С = 0.26248, С = 0.2790. A ML analysis incorporating these parameters generated a tree with a likelihood score (-InL) of 2193.43 (Fig. 15). Parsimony and likelihood analyses pro- duced essentially the same topologies. The clade grouping, in MP topology, the two O. draparnaudi specimens with O. pilula repre- sents the only difference between the two reconstructions. In both topologies, all con- specific specimens grouped together in clades strongly supported by bootstrap values. The two reconstructions also suggest close phylo- genetic relationships between O. majori, O. paulucciae, O. uziellii and O. meridionalis. DISCUSSION Hyalinia paulucciae and Oxychilus lanzai Re-examination of the specimens from Grotta della Risvolta (Figs. 2, 11, 12) and Grotta del Buggine (Figs. 6, 7), which Forcart (1967) as- signed to O. (Ortizius) lanzai, and those from Tana di Magnano, which Forcart (1967) as- signed to O. (Oxychilus) paulucciae, confirmed that they belong to a single species. These specimens have the same characters as those collected near Vagli, the type locality of O. paulucciae. Similar conclusions were provided by molecular data. The two specimens (one from Grotta della Risvolta and one from Vagli) analysed were genetically very similar, forming a well differentiated evolutionary lineage with respect to all the other Oxychilus examined. The level of genetic divergence observed was corre- lated with geographic sampling and fall within the range observed for the other conspecific Oxychilus. The observed congruence between morphological and molecular data definitively demonstrates that O. /anzai is a junior synonym of O. paulucciae, as hypothesized by Giusti & Mazzini (1971) and Riedel (1980, 1997, 1998). 32 MANGANELLI ET AL. 10 changes O. paulucciae 1 O. paulucciae 2 O. meridionalis 1 a meridionalis 2 O. meridionalis 3 O. uziellii O. majori O. pilula O. draparnaudi 1 O. draparnaudi 2 Retinella olivetorum FIG. 14. Most parsimonious tree calculated from ITS-1 sequence data. Bootstrap values are indicated at nodes (1,000 replications). Morphological Analysis Oxychilus paulucciae belongs to Oxychilus ($. str.), sensu Giusti & Manganelli (1999), being characterized by: penis with flagellum; penial retractor inserted at apex of flagellum; internal ornamentation of penis consisting of pleats or rows of papillae without apical thorns; epiphallus long, usually longer than proximal penis; internal wall of epiphallus with slender longitudinal pleats; mucous gland mainly vaginal; long mesocone of cen- tral tooth. Among Oxychilus (s. str.) species it holds an intermediate position, sharing a narrow mid- penial portion (“bottle-neck”) enveloped by a thin sheath with O. diductus (Westerlund, 1886), О. draparnaudi, О. majori, О. mortilleti (Pfeiffer, 1859), О. oglasicola Giusti, 1968, and О. oppressus (Shuttleworth, 1878), but unlike them it has internal ornamentation of proximal penis consisting of longitudinal pleats very similar to that of O. meridionalis (O. diductus: Manganelli et al., 2002: figs. 7-11; O. draparnaudi: Giusti & Manganelli, 1997: figs. 15-30; Manganelli 8 Giusti, 1998: figs. 19-22; O. majori: figs. 4-8; O. meridionalis: Manganelli 8 Giusti, 2001: figs. 9-31; O. mortilleti: Manganelli & Giusti, 1998: figs. 5— 17; O. oglasicola: Manganelli et al., 1999: figs 12-14; O. oppressus: Riedel, 1967: figs. 1, 2; personal unpublished data). REVISION OF OXYCHILUS PAULUCCIAE 33 0.05 substitutions/site O. paulucciae 1 71 O. paulucciae 2 82 O. meridionalis 1 95 96 | Г O. meridionalis 2 94 O. meridionalis 3 85 O. идей O. majori O. pilula O. draparnaudi 1 70 O. draparnaudi 2 Retinella olivetorum FIG. 15. Maximum likelihood tree calculated from ITS-1 sequence data. Bootstrap values are indicated at nodes (1,000 replications). Consequently, internal ornamentation of the proximal penis consisting of longitudinal pleats readily distinguishes O. paulucciae from O. diductus, O. draparnaudi, O. majori, O. mortilleti, O. oglasicola and O. oppressus, and the narrow mid-penial region (“bottle- neck”) distinguishes it from O. meridionalis. Besides the structure of the mid-penial region, O. paulucciae can be distinguished from O. meridionalis by its pale grey body, larger shell (shell diameter: 15.5 + 1.0 mm) with small umbilicus (about 1/8 of shell maximum diam- eter), and vaginal gland that often forms in- complete ring around proximal vagina (body slate blue in colour; smaller shell: shell diam- eter: 13.1 + 2.0 mm, with larger umbilicus, about 1/6-1/7 of shell diameter; vaginal gland always forming complete ring around proximal vagina; for detailed description of O. meridionalis: Manganelli 8 Giusti, 2001). Oxychilus paulucciae also shares the inter- nal ornamentation of the proximal penis, con- sisting of longitudinal pleats, with the species traditionally assigned to Ortizius Forcart, 1957 (type species: Hyalina (Polita) helvetica Blum, 1881). Only one of the 28 species assigned by Riedel (1980, 1998) to this subgenus (Giusti & Manganelli, 2002: table 1) occurs within the area inhabited by O. paulucciae: O. clarus (Held, 1838). It is impossible to confuse the two species: O. clarus has a very small whit- ish shell (Kerney et al., 1983: pl. 10). 34 МАМСАМЕН ET AL. Phylogenetic Relationships Our phylogenetic reconstructions indicate that the species analysed in this study do in- deed represent well-differentiated evolutionary lineages. In particular, the three species for which multiple specimens were available al- ways formed monophyletic clades supported by bootstrap analysis, suggesting good overall resolution of the data set. Molecular data also indicated that О. paulucciae is genetically close to О. meridionalis, the two taxa separated by a sequence divergence ranging from 2.5 to 3.7%. This result is congruent with morphological evi- dence, if it is admitted that the internal structure of proximal penis, not the narrow mid-penis region, supports taxonomic relationships within the genus. Another interesting finding of this study was the close relationship between O. paulucciae, O. meridionalis, О. uziellii, and O. majori, which supports the existence of a Tuscan radiation of the genus. The last two taxa, O. uziellii, and O. majori, are character- ized by some highly derived features in the penial complex. This makes it difficult to unam- biguously infer their systematic affinities based on morphological evidence. Morphologically derived taxa represent a challenge for system- atics that can only be addressed by a molecu- lar approach. Finally, molecular data indicated that O. pilula and O. draparnaudi are distantly related taxa with respect to the above species. CONCLUSION Both morphological and ITS-1 sequence data indicates that O. paulucciae 1$ close, but distinct from O. meridionalis, a widespread Tuscan species. The molecular results also showed that O. uziellii and O. majori, two other morphologically highly derived Tuscan species, are closely related to O. meridionalis and O. paulucciae and distinct from the most common O. draparnaudi. Combined morphological and molecular analysis of wider taxonomic sample, espe- cially type species of the many subgenera of Oxychilus, will further clarify the taxonomy of the genus and the relationships within oxychiline zonitids. ACKNOWLEDGMENTS We thank Antonella Daviddi and Leonardo Gamberucci for technical assistance, Helen Ampt for revising the English, Marco Bodon (Genoa, Italy), Micaela Calcagno (Florence, Italy), Gianni Comotti (Nembro, Bergamo, Italy), Elisabetta Lori (Pistoia, Italy), Paolo Magrini (Florence, Italy), Augusto Vigna Taglianti (Rome, Italy), and Franco Utili (Florence, Italy) for field collection and Benedetto Lanza (Flo- rence, Italy), Ambros Hánggi and Jolanda Ineichen-Riedi (Basel, Switzerland) for informa- tion about or loan of material from the Naturhistorisches Museum Basel (Switzerland). This research was funded by grants from “Bioitaly Tuscany”, “EEC Regulation 2081/93 — Objective 5/b” projects and by Universita di Siena (PAR 2001, project “| molluschi поп marini della fauna italiana: filogenesi, sistematica, faunistica, zoogeografia, conser- vazione) and Museo Zoologico de “а Specola”, Sezione del Museo di Storia Naturale dell’Universita di Firenze (Italy). 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Nucleic Acids Research, 22: 4673—4680. WESTERLUND, C. A., 1886, Fauna der in der paláarktischen Region (Europa, Kaukasien, Sibirien, Turan, Persien, Kurdistan, Armenien, Mesopotamien, Kleinasien, Syrien, Arabien, Egypten, Tripolis, Tunesien, Algerien und Marocco) lebenden Binnenconchylien. |. Fam. Testacellidae, Glandinidae, Vitrinidae & Leucochroidae. Hakan Ohlsson’s Buch- druckerei, Lund, 88, 7 pp. Revised ms. accepted 4 July 2003 o AU y _ + . ' o Er ' o р 0 = . =; = : 7 | -« 7 Ss = 7 + 7 DEF be 2 is ¿ai Mi da — Ñ PE O A Иры >= 7 oe | $ bs Ö Er | | = tan | | pA O à et NAS MA + 1 ° BS y 3 | ET” wi Ne: : (2 ' u on u р | $ . ko y = = | u à = | y ка u o a ATAN a 20 , ase TE gere und vt ма o > у o > é > SV AN MEN т | u i т, > yO "HN pit Ñ y ie o > AL | У? р u u de 7 y EN ” | y = 7 o LEE LE o a4 mi ee ve та я кА PER a Ree os PS A u я rc aye cer aise Y e Lo a 7 | u = Ñ | MAS > o 710 u | en: o [ a ig Suh 4 ‘à u o ER a u | CPR CARE. u Lie, 5 fan 7 7 LA р | в 2 7 р > Fe Be aa | к | hal | м 7 y À a й - & sE : Zu р 7 ое 7 : нами & y | ' ES 7 a n 7 О . 1 ON | o | 7 o Ty a #3 o e : K | 4 o bl Ñ Ex om"; a+ de Г | à "14004 a u i | a o = ae 10 “= AA | р . | ый р Вх vo 7 En LC u р sa u SE y IN su cda A и nite 7 . | | = . Y u LAS у О > 7 = es Ri = snes E. u = A 7 o in y > te . o +) Le ¡ESA ri et и D LE >it où № y e e 2 10 | ma LOT ый ‘i ie $ MATE ми 4 Фей 22 do A Fo г | ria yon Qui pro | - UN Au > > oni: | md o ¡DAI a ù Las 10 OA. nr Di MALACOLOGIA, 2004, 46(1): 37-55 THE BIOLOGY AND FUNCTIONAL MORPHOLOGY OF FOEGIA NOVAEZELANDIAE (BIVALVIA: ANOMALODESMATA: CLAVAGELLOIDEA) FROM WESTERN AUSTRALIA Brian Morton Western Australian Museum, Francis Street, Perth, Western Australia 6000, Australia; prof_bsmorton@hotmail.com ABSTRACT As more representatives of the adventitious, tube-building anomalodesmatan Clavagelloidea are examined, a pattern of extraordinary adaptive radiation is being re- vealed. Despite its name, Foegia novaezelandiae is known only from the Holocene and Recent of Western Australia and is thus possibly very modern. A few tubes are held in the collections of the Western Australian Museum, Perth, and a single living individual has been collected from a hypoxic beach at Dampier, Western Australia. Like other clavagelloids, using a muscular pedal disc, Г. novaezelandiae pumps interstitial water into its mantle cavity via the pedal gape, and hence the pedal slit and tubules of its anterior “watering pot” com- ponent of the adventitious tube. Foegia novaezelandiae is similar to Brechites vaginiferus in being amyarian, except for minute anterior pedal retractor muscles in the latter. As with B. vaginiferus also, pallial re- tractor muscles effect siphonal and pedal disc retraction. The adventitious tube of Е. novaezelandiae 1$ more complex in that the shell valves are recessed and largely hidden externally by additional bulbous concretions of tube material secreted from anterior and posterior pallial crests. Also like B. vaginiferus, F. novaezelandiae pumps interstitial water into the mantle cavity, probably collecting interstitial bacteria and dissolved organic mate- rial as nutritional supplements. Unlike В. vaginiferus, however, Е. novaezelandiae has ап agglomeration of organic material and bacteria adhering to its highly convoluted periostracum anteriorly, particularly that of the pedal disc and thus inside the adventitious tube. Such bacteria may help detoxify the hydrogen sulphide contained in the interstitial water of the hypoxic sediment that F novaezelandiae inhabits. However, F novaezelandiae has a full complement of mantle cavity and intestinal organs for the processing of food fil- tered from the seawater above. Key words: Foegia novaezelandiae, Clavagelloidea, adventitious tube formation, anatomy, tube function, watering-pot shell. INTRODUCTION The adventitious tubes of the diverse repre- sentatives of the Clavagelloidea d’Orbigny, 1843, constitute some of the weirdest and rar- est bivalve structures. The most recent cla- distic analysis of the Anomalodesmata by Harper et al. (2000) did not identify sister groups but noted that Clavagella and its allies first appeared in the Cretaceous, whereas Brechites and its allies are known from the Oligocene onwards. Savazzi (2000) also noted that representatives of the Clavagelloidea seem to fall into two groups comprising those that (1) have their left valve united into the fab- ric of an adventitious tube in the case of 37 endobenthic (Stirpulina) and epibenthic (Dianadema) genera, or a crypt in the case of nestling and boring species (Clavagella, Bryopa), with the right valve free inside it, and (ii) those in which both valves are incorporated into the structure of an adventitious tube, again in the case of endobenthic (Brechites, Foegia) and epibenthic (Humphreyia) genera. The anatomies of species of Clavagella, Bryopa and Dianadema have been described by Soliman (1971), Savazzi (1999, 2000) and Morton (1984a, 2003), respectively, and those of representatives of Brechites and Humphreyia by Morton (1984b, 2002a, b). Two of the above genera, that is, Dianadema and Humphreyia, are known only from Australia, 38 MORTON and Smith (1971, 1976, 1998) and Lamprell & Healy (1998) catalogue the species recorded from that continent. These authors consider that in Australia the genus Brechites comprises three subgenera, that is, Brechites, s.s., plus Penicillus and Foegia, the second subgenus being represented by В. (P.) philippinensis (Chenu, 1843) and the third by В. (F.) novaezelandiae (Bruguiére, 1789) and В. (F.) veitchi Smith, 1971. Brechites (F.) novae- zelandiae is the type species of Foegia but, as noted by Smith (1971), other than for a description of its adventitious tube, virtually nothing else is known about it and there are no extant specimens with tissues available for study. During January 2000, a research trip was made to Western Australia and a single living individual of Foegia novaezelandiae was col- lected. On this and subsequent visits, the small collection of tubes of this species in the West- ern Australian Museum was examined. Obser- vations on the living animal and the collection of examined tubes are herein reported upon to provide an insight into the biology and anatomy of one of the strangest species, of one of the strangest superfamilies (Clavagelloidea) within the Bivalvia (Morton, 1981a, 19854). MATERIALS AND METHODS The specimen of Foegia novaezelandiae was collected from intertidal mud on the beach adjacent to the leased property of Dampier Salt Co. Ltd., Karratha, Western Australia. It was buried anterior end down, that is, the watering pot, with the posterior tube projecting just above the mud surface. As described for Brechites vaginiferus (Chenu, 1843) by Morton (2002a), the ante- rior end only of the adventitious tube of Foegia novaezelandiae was placed within a transpar- ent tub with a lid that had a central hole in it to hold the tube in place and containing a sus- pension of Ehrlich’s haematoxylin in seawa- ter. The whole animal and tub was then placed in a much larger, also transparent, container of filtered seawater and left overnight. The liv- ing animal was subsequently dissected and the ciliary currents of the organs of the mantle cavity studied by application, again, of a sea- water suspension of Ehrlich’s haematoxylin. The specimen was fixed in 5% formalin even- tually and, following routine histological pro- cedures, sectioned transversely at 6 um and every tenth section retained. Alternate slides were stained in either Ehrlich’s haematoxylin and eosin or Masson’s trichrome. The nine specimens of Foegia novae- zelandiae contained in the collections of the Western Australian Museum, Perth, were ex- amined and the dimensions of all intact tubes measured to the nearest 1 mm. These were: greatest width, total length and length to the first growth (or possibly repair) increment. ABBREVIATIONS USED IN FIGURES AC Anterior concretion AN Anus APC Anterior pallial crest APCC Anterior pallial crest cavity AU Auricle CA Ctenidial axis CE Cuticular fusion CM Circular muscle CP Ctenidial plica C-P-V-CONN Cerebro-pleural visceral connective DD Digestive diverticula DK Distal limb of the kidney ES Exhalant siphon F Foot FPA Fourth pallial aperture H Heart HA Haemocoel HG Hypobranchial gland IBC Infra-branchial chamber ID Inner demibranch IE Inner epithelium IER Inner labial palp IP Inner layer of periostracum IS Inhalant siphon K Kidney KC Kidney concretion KT Kidney tubule LM Longitudinal muscle LV Left shell valve MG Mid gut N Nerve O Oesophagus OA Organic agglomeration OD Outer demibranch OE Outer epithelium OEP Outer labial palp OP Outer layer of periostracum OS Osphradium OV Ovary E Periostracum PC Posterior concretion BIOLOGY OF FOEGIA 39 PD Pedal disc PE Pericardium REG Periostracal groove PG Pedal gape PK Proximal limb of the kidney PL Pallial line PPC Posterior pallial crest PPCC Posterior pallial crest cavity PRM Pallial retractor muscle R Rectum RA Renal aperture RV Right shell valve S Siphons SA Saddle SBC Supra-branchial chamber SC Sensory cell SN Siphonal nerve SVI Shell valve impression ME Testes ТМЕ Transverse muscle fibres \/ Ventricle VM Visceral mass VMG Ventral marginal food groove TAXONOMIC CONSIDERATIONS Smith (1971) discussed the taxonomy of Brechites (Foegia) novaezelandiae (Bruguière, 1789). He regarded Aspergillum agglutinans Lamarck, 1818 (р. 430), and А. novae- hollandiae Chenu, 1843 (p. 3, pl. 4, fig. 8), to be synonyms. Penicillus novae Zelandiae Bruguière, 1789 (p. 129-130), was based on an ambiguous illustration in Favanne de Montcervelle & Favanne de Montcervelle (1780: 642, plate 79, fig. E), and misattributed to New Zealand. А neotype may be needed to stabilize the concept, because the original material has not come to light. No type mate- rial of А. agglutinans has been found. Two syntypes of Aspergillum novaehollandiae Chenu, 1843, are held in the collections of the Natural History Museum, London (1968668), and these are figured here (Fig. 1). Gray (1858a: 313) differentiated Foegia Gray, 1847 (p. 188), from other genera in his Aspergillidae Gray, 1858, a junior synonym of the Clavagellidae Orbigny, 1844, in several important respects: “Umbo more or less cov- ered with a swollen prominence in front; the whole of the valves except the umbo or nucleus enclosed in the tube; fringe indistinct, formed like the hole in the disk, of short thick separate tubes”. The above description is gen- erally correct, and because of other anatomi- cal differences, | agree with Gray (1858a) that the genus Foegia is valid. It is possible that Foegia might date from Gray (1842: 77), where there is a definition but no named species. In any event, the type species of Foegia is Peni- cillus novae Zelandiae Bruguiére, 1789, by monotypy in Gray (1847). The species under consideration is, therefore, Foegia novae- zelandiae (Bruguiére, 1789). FIG. 1. Foegia novaezelandiae. The two syn- types of Aspergillum novaehollandiae (NHM London 1968668). 40 MORTON DISTRIBUTION In the collection of the Department of Earth and Planetary Sciences of the Western Aus- tralian Museum are a number of local Ho- locene subfossils of Foegia novaezelandiae: 1.Kwinana (south of Perth). Dredged from Cockburn Sound, 3 specimens (WAM 69.1070a, b, c). 2.Fremantle. Dredged from a fishing anchor- age, 1 specimen (WAM 70.2034). In the collection of Recent Mollusca in the Western Australian Museum are 16 specimens of Foegia novaezelandiae collected from ei- ther Cockburn Sound, South Fremantle, Woodman's Point or Leighton Beach, all loca- tions again just south of Perth. One was dredged from 1-2 fathoms (2-4 m), and all were dead when collected. Only nine tubes are intact. Smith (1971) records that Foegia novae- zelandiae occurs along “The central and south west coast of Western Australia and two speci- mens from the north coast of Queensland” (p. 152). Cotton (1961) does not record the spe- cies from South Australia, nor do Wells 8 Bryce (2000) from Western Australia, presumably because of its rarity. Smith (1976) illustrates (р. 201, map 3) the range of F novaezelandiae. Lamprell 8 Healy (1998) agree with this distri- bution pattern and report that the species oc- curs from depths of 3-22 m in sand. The record herein, from Dampier, though intertidal, 1$ within the distribution range described, and therefore F. novaezelandiae is a Southern Hemisphere, warm temperate-tropical species. BIOLOGY The single specimen from the Dampier Salt Co. Ltd. lease at Karratha, Western Australia, was collected from the intertidal of an un- named muddy beach, the landward drainage onto which has been restricted by construc- tion of a bund to create solar salt pond “0”. The seaward remnant of the original creek which drained onto the beach, lies opposite and is divided into two outlets by West Inter- course Island. Mangroves fringe the beach: an Avicennia forest to the seaward is followed landward, in succession, by Rhizophora scrub, Avicennia scrub, and (locally) Ceriops- Avicennia heath grading into a salt flat. The main water influence here is the tides because the hypersaline (salinities > 40%), drainage from the land, as reported upon by Morton (2002a) for this part of Western Australia, has been halted by construction of the bund and causeway for pond “0”. This has thus in turn adversely impacted not only beach dynamics but also interstitial water character. Whether FIG. 2. Foegia novaezelandiae. A. The adventitious tube; B. the siphonal tube as seen from the posterior aspect; C. a closer view of the tube showing the calcareous tube and periostracum beneath the adhering detritus and D, the watering pot as seen from the anterior aspect. Note the dorso-ventrally aligned pedal slit (for abbreviations see рр. 38-39). BIOLOGY OF FOEGIA 41 natural or perturbed, the substratum of sandy- mud in this Foegia novaezelandiae habitat is hypoxic, and the specimen was oriented ver- tically in it with the posterior end of the tube projecting above the sediment surface by some 10 mm. Semeniuk & Wurm (1987) de- scribe in broad terms the characteristics of the shore seaward of pond “0” and provide basic maps (figs. 21, 22) of the area. ANATOMY Adventitious Tube The nine tubes of Foegia novaezelandiae in the collections of the Western Australian Mu- seum range in total length from 69-98 mm and in maximum width from 13-16 mm. The living individual from Dampier was 130 mm long and 15 mm wide. Some tubes in the collection have either a single growth increment or a repair at a length ranging from 80-94 mm. The Dampier individual has two (Fig. 2A). The relationships between tube width and total tube length and length to the first growth increment or repair are illustrated in Figure 3. Where there is no growth increment, the two measurements are 16 y = -0.0156x + 15.315 =. R? = 0.0355 Е = 3 > 8 E y = 0.0039x + 13.79 13 В? = 0.0029 $ 12 Y the same. Although the correlations are poor, the lines of best fit are similar. Four individu- als, each with one growth or repair mark on the tube, lie on the right side of the plot sug- gesting that any such increment occurs at a length of between - 85-100 mm. The above implies that the adventitious tube 1$ secreted but once when the contained animal becomes an adult, but that it can be subsequently ex- tended or repaired posteriorly, as in Brechites vaginiferus (Morton, 2002a). The tube о the living Foegía novaezelandiae is illustrated in Figure 2. The main shaft of the tube (Fig. 2A) is covered in sand grains and other hard detritus, except posteriorly and anteriorly at the watering pot disc. Posteriorly, there are two growth (or repair) increments, both secreted internal to the preceding one. These are covered sparsely in detritus and raised above the sediment surface. Viewed from the posterior aspect (Fig. 2B), the tube aperture is 8-shaped in cross-section match- ing the configuration of the siphons, which project up into it. In places, the shell debris is worn away from the tube beneath exposing the calcareous tube with a thin adhering film of periostracum (Fig. 2C). Seen from the an- terior end (Fig. 2D), the watering pot disc has + e © © On 14 = OS $ $ &---- Tube width versus total tube length @——® Tube width versus to first growth increment/repair Tubes with one growth increment/repair 50 55 60 65 70 75 80 85 90 95 100 Tube length (mm) FIG. 3. Foegia novaezelandiae. The relationship between adventitious tube width and (1), total length and (ii), length to the first growth (or repair) increment. 42 MORTON PC AC FIG. 4. Foegia novaezelandiae. A view of the dorsal surface of the adventitious tube showing the true shell valves and enclosing anterior and posterior bulbous projections (for abbreviations see pp. 38-39). a dorso-ventrally aligned pedal slit and an ar- ray of open tubules which, as shown by Gray (1858a), do not have a distinct “fringe” sepa- rating it from the tube's shaft, as is the case in Brechites vaginiferus and where it is identi- fied as a distinct “line” (Morton, 2002a: fig. 1). Tube Function When the watering pot of the living individual of Foegia novaezelandiae was placed in a suspension of Ehrlich's haematoxylin in sea- water, the animal clarified it within 12 hours. Thus, as with Brechites vaginiferus (Morton, 2002a), Е. novaezelandiae pumps interstitial water into the mantle cavity through the pedal slit and tubules that constitute the watering pot. Shell As noted by Gray (1858a), the shell of Foegia novaezelandiae is covered by two, anterior and posterior, bulbous secretions and is gen- erally hidden within the fabric of the adventi- tious tube. However in one specimen in the Western Australian Museum collection (from Cockburn Sound, (i) of (iv) specimens col- lected in 1965; broken base only; S 14232), the shell valves are partly visible. This speci- men was cleaned carefully with dilute nitric acid, to remove sand grains and other debris and is illustrated in Figure 4. The two shell valves have parted and are ~ 3 mm long. They are equivalve and inequilateral, that is, anteri- orly foreshortened and posteriorly elongate, and thus of the same general form as in all clavagelloids hitherto described, for example, Brechites vaginiferus, Humphreyia strangei and Dianadema multangularis (Morton, 2002a, b, 2003). The umbones are slightly pointed, and there is a trace of a radial sculpture of periostracal spinules, similar to those de- scribed for Lyonsia hyalina by Prezant (1979a) and for the clavagelloids listed above. Around BIOLOGY OF FOEGIA 43 PPCC УМ РЕ АРСС FIG. 5. Foegia novaezelandiae. An internal view of the adventitious tube showing the positions of the true shell valves and pallial line lying below the saddle (for abbreviations see pp. 38-39). the two shell valves and uniting them, is a “saddle” of secondarily secreted shell which has fine concentric growth lines also seen in other clavagelloids (see above). Shell and saddle are sunk into the general fabric of the adventitious tube. A thick, bulbous concretion covers the antero-dorsal region of the right valve, and a second, similarly bulbous con- cretion is present posteriorly. Internally, the shell, saddle and adventi- tious tube of the Dampier specimen (Fig. 5) are united and covered by a smooth calcar- eous concretion. The positions of the valves appear as depressions surrounded by raised borders of secondarily and internally se- creted calcium carbonate. Pockets where anterior and posterior pallial crests are in- serted above the valves to create the bul- bous secretions covering them are also evident. Two crescentric pallial-line scars encircle the antero-lateral sides of the shell valve impressions. The Pilbarra region of Western Australia is mineral rich and the in- ternal surface of the anterior watering pot was stained brown with iron oxide. Internal Anatomy The living animal of Foegia novaezelandiae was removed from its tube and is illustrated in Figure 6A-C. The siphons have contracted. The entire body is enclosed in periostracum secreted by the general mantle epithelium. Covering the mantle immediately beneath the true shell and, therefore, approximately en- compassing the pericardium, the periostracum is a transparent skin (this is illustrated as a light stippling in Figure 6C). Elsewhere, cov- ering siphons, pedal disc and the general mantle surface, the light brown periostracum is thick and wrinkled. From the dorsal view (Fig. 6A), the pericardium contains a heart, which comprises a central ventricle, pen- etrated by the rectum, and lateral auricles. Posteriorly, there are paired kidneys, over which the rectum passes. Anteriorly, the vis- ceral mass contains the digestive diverticula and the paired ovaries. From what is the crescentric remnant of a pallial line, pallial re- tractor muscles pass into the mantle in ante- rior, ventral and posterior directions to effect 44 MORTON РАМ В y к АРЕН ppc АРС N PL | de E OV FIG. 6. Foegia novaezelandiae. А generalized picture of the anatomy, as seen from А, dorsal; B, ventral and C, right lateral aspects. Note that in C the periostracum surrounding the pericardium is illustrated with a light stippling as in Figure 2C: elsewhere the periostracum is brown, thick and wrinkled (for abbreviations see pp. 38-39). BIOLOGY OF FOEGIA 45 Srey EEE у 1 <=. <«— 10 mm к fo 1 Pee Ape «a CF ее An р =a = AN PL : ; у»: > ILP PD FIG. 7. Foegia novaezelandiae. An interval view of the organs and ciliary currents of the mantle cavity as seen from the right side (for abbreviations see pp. 38-39). contraction of the body within its adventitious tube. There are no other muscles. Also seen dorsally, above the visceral mass, are ante- rior and posterior pallial crests. From the ventral view (Fig. 6B), the periostracum-covered pedal disc lies antero- ventrally, and in its centre is a dorso-ventrally aligned pedal gape. Where the siphons meet the remainder of the mantle, there is a mid- ventral fourth pallial aperture. The animal, as seen from the right side (Fig. 6C), shows the heart within the pericardium and the rectum passing over the kidneys, the pallial retractor muscles and the anterior pedal disc and gape. Also seen are the fourth pallial aperture and the anterior and posterior pallial crests. Organs and Ciliary Currents of the Mantle Cavity The extended body of Foegia novaezelandiae is shown in Figure 7 after being opened on the right side. The most obvious feature is the long paired ctenidia, each of which consists of a com- plete inner demibranch and the dorsally directed descending lamella only of the outer. The ctenidia extend into the apex of the siphons and thus separate supra- from infra-branchial cham- bers. The ciliary currents of the ctenidia are of Type E (Atkins, 1937a) and pass collected par- ticles anteriorly towards the mouth in the ctenidial axis and in the ventral marginal food groove of the inner demibranch via small labial palps. The visceral mass is small with a little foot antero-ventrally. No statocysts have been iden- tified, although they occur in most anomalo- desmatans (Morton, 1985b), but were similarly not seen in Dianadema multangularis (Morton, 2003). Their absence in this specimen may be because only every 10" transverse sec- tion was kept but this would mean any missed statocysts would be very small, that is, < 60 um in length. Within the visceral mass, dorsal ovaries are separate from ventral testes. The ciliary currents of the visceral mass are directed towards its postero-ventral edge where unwanted particles fall onto the mantle mid- ventrally. As in Brechites vaginiferus (Morton, 2002a), the ciliary currents on the internal sur- face of the pedal disc radiate outwards and downwards from the pedal gape. The ciliary currents on the internal surface of the mantle are downward, complementing those о the vis- ceral mass but, mid-ventrally, strong ciliary cur- rents transfer unwanted material posteriorly, where it is ejected from the inhalant siphon as pseudofaeces There are also posteriorly di- rected ciliary currents in the supra-branchial chamber and which presumably help to trans- fer faeces to the exhalant aperture because the anus is located deep inside the siphons on the posterior surface of the paired kidneys. Musculature Foegia novaezelandiae has no adductor and pedal retractor muscles. The pallial line 1$ short, ~ 3 mm, on each side of the body and from it arise pallial retractor muscles that ex- tend anteriorly, ventrally and posteriorly. The attachment of the pallial retractor muscles to the adventitious tube, at the pallial line, 1$ shown in transverse section in Figure 8. 46 MORTON DD O PRM C-P-V-CONN OEP CM LM FIG. 8. Foegia novaezelandiae. A transverse section through the visceral mass and mantle (for abbreviations see pp. 38-39). FIG. 9. Foegia novaezelandiae. A transverse section through the outer mantle epithelium of the pedal disc showing the periostracum and agglomeration of adhering organic material and bacterial cells (for abbreviations see pp. 38-39). BIOLOGY OF FOEGIA 47 FIG. 10. Foegía novaezelandiae. А ЗЕМ micrograph of the outer surface of the pedal disc, that is, inside the adventitious tube, showing attached inorganic and organic detritus and rod-shaped bacteria. Mantle The mantle margin of Foegía novaezelandiae is shown in transverse section in Figure 8. Mantle fusion is of Туре С (Yonge, 1982), that is, inner, middle and inner surfaces of the outer mantle folds, so that virtually everywhere the outer surface of the general mantle is enclosed in thick periostracum. The pallial retractor muscles extend into the mantle (Fig. 7) and posteriorly form longitudinal fibres that retract the siphons. Laterally, the mantle has a capa- cious haemocoel and circular muscles from both the left and right assist in pallial contrac- tion. The mantle of the pedal disc is shown in transverse section in Figure 9. The outer epi- thelium is thrown into many folds and at the apex of each pleat there is a swollen cell ~ 8 um in diameter which is innervated by tiny subepithelial nerves. The epithelium also se- cretes the periostracum, which comprises two layers. The inner is thick, up to 50 um and stains blue in Masson’s trichrome. It is prob- ably mucoid. The outer layer is thin (2 um), stains red in Masson’s trichrome and is thrown into complex fibrous folds and strands. Around the pedal disc but diminishing towards the si- phons, the outer surface of the periostracum is covered in an agglomeration of organic material. Within this are slightly curved, rod- shaped bacteria, ~ 1.5-2 um in length, and which do not stain in either Masson’s trichrome or Ehrlich’s haematoxylin, but shine a bright yellow-green. This agglomeration of organic material and bacteria attached to the pedal disc, being inside the adventitious tube, is in darkness. It is not present in the similarly endobenthic Brechites vaginiferus (Morton, 1984a: fig 16a). The agglomeration of inor- ganic and organic detritus with the bacteria attached to the pedal disc, as seen under the SEM, is illustrated in Figure 10. Siphons As is typical of all clavagelloids studied hith- erto (Morton, 1984a, b, 2002a, b, 2003), and for other anomalodesmatans (Prezant, 1979b; Morton, 1981b), radial mantle glands at the apices of the siphons of F novaezelandiae produce a secretion which attaches sand grains and other detritus to the thick periostracum of their outer surfaces to cam- ouflage them. The siphons are shown in trans- verse section in Figure 11A. Internally, there are 16 pallial nerves that, in other clavagelloids, for example, Brechites vaginiferus (Morton, 2002a) relate to the number of sensory papil- lae, which surround the siphonal orifices. The siphonal wall is illustrated in greater detail in Figure 11B. Externally, are outer and inner layers of the periostracum. Internal to the outer epithelium is a haemocoel and inter- nal to this are successive layers of longitudi- nal, circular, longitudinal and circular muscles. Criss-crossing the longitudinal muscle blocks are transverse and oblique fibres that must create the tonus which extends and contracts the siphons, in cooperation with the other 48 MORTON 1mm 2.5mm CM SN FIG. 11. Foegia novaezelandiae. Transverse sections through A, the siphons showing the thick perio- stracum and pallial nerves and В, the siphonal wall in greater detail (for abbreviations see pp. 38-39). muscles and blood-filled haemocoels of the mantle. In terms of its muscular complexity, the siphons of Foegia novaezelandiae are very similar to those of Brechites vaginiferus (Morton, 1984a: fig.14) and Humphreyia strangei (Morton, 2002b: fig.12). Ctenidia The long, homorhabdic ctenidia (Fig. 7) are also illustrated diagrammatically in transverse section in Figure 12. Approximately five pli- cae make up the descending lamella of the outer demibranch and about eight both lamel- lae of the inner. There is a ventral marginal food groove in the latter. Each plica comprises a maximum of 20 filaments anteriorly, but only two as the ctenidia decline in size posteriorly (Big: 7). As in other clavagelloids, for example, Brechites vaginiferus (Morton, 1984a, 2002a), the epithelium ventral to the kidneys and which forms the dorsal surface of the supra-branchial chamber of the outer demibranch is modified into a hypobranchial gland. The descending lamella of the outer demibranch attaches to the visceral mass by a cuticular junction, as does the ascending lamella of the inner (Atkins, 1937b). This was first described for ап anomalodesmatan, that 1$, Laternula truncata, by Morton (1973) and is considered characteristic of all representatives. Medially, adjacent to the cuticular junction 1$ an osphradium that has not hitherto been de- scribed for any anomalodesmatan, although it has been reported in other bivalves, for ex- ample, Corbicula fluminea (Kraemer, 1981). Left and right osphradia (Fig. 12) extend from the labial palps to the posterior end of the vis- ceral mass. In transverse section (Fig. 13), each osphradium lies between the cuticular junction of the outer demibranch with the vis- ceral mass and the hypobranchial gland. It comprises a central core of cells between which nerve fibres pass towards the periph- ery. The ощег epithelium is thin (4 um) but periodically along its margin there are swollen sensory cells ~ 8 um tall and towards which the nerves are oriented. Pericardium and Kidneys The pericardium and kidneys are illustrated in Figure 6A and C and in transverse section in Figure 12. The rectum is enclosed by the ventricle of the heart (in turn surrounded by the pericardium) but lies dorsal to the paired kidneys. Each kidney comprises a capacious distal limb and a bag-like proximal limb that opens into the supra-branchial chamber of the inner demibranch at ciliated renal apertures (Fig. 12). There are no pericardial proprio- receptors such as occur in Humphreyia strangei and Dianadema multangularis BIOLOGY OF FOEGIA д) RS a) 7 = VMG OV 2.5 mm ТЕ FIG. 12. Foegia novaezelandiae. A transverse section through the paired kidneys Showing the renal apertures, the ctenidia and the position of the paired hypobranchial glands and osphradia within the supra-branchial chamber of the outer demibranch (for abbreviations see pp. 38-39). CF IBC FIG. 13. Foegia novaezelandiae. A transverse section through the hypobranchial gland and osphradium in the supra-branchial chamber of the outer demibranch (for abbreviations see pp. 38-39). 49 50 MORTON KT KC 20 um FIG. 14. Foegia novaezelandiae. À transverse section through two distal limb tubules of the kidney showing the contained concretions (for abbreviations see рр. 38-39). (Morton, 2002b, 2003) probably because there are no remnants of the posterior pedal retrac- tor muscles as in Brechites vaginiferus, which similarly does not have such sense organs (Morton, 2002а). Distal kidney tubules are illustrated in trans- verse section in Figure 14. The cells are some 10 pm tall, largely vacuolated, and contain approximately spherical concretions, between 6-8 ит in diameter and which stain blue in Masson’s trichrome but with a lighter staining core. Such concretions also occur in the lu- mina of the distal limb tubules. DISCUSSION The first, detailed description of a tube-dwell- ing clavagelloid (Aspergillum dichotomum) was by Lacaze-Duthiers (1883). Three-quar- ters of a century later, Purchon (1956, 1960) described Brechites penis and, later, Smith (1971, 1976, 1998) produced simple illustra- tions of Australian species, but not Foegia novaezelandiae. Subsequently, Morton (1984a, 2002a, b) described Brechites vaginiferus and the cemented Humphreyia strangei. Clavagelloids that unite only the left valve into the fabric of a crypt (Clavagella and Bryopa) have been described by Owen (1835), Soliman (1971) and Morton (1984b). The strange, cemented species, Dianadema multangularis, with tubules that form a crown over the dorsal part of the shell and adventi- tious tube, was described by Morton (2003) and suggested to be similar functionally to the North American, Late Cretaceous Ascaulo- cardium armatum (Pojeta & Sohl, 1987). Savazzi (1982, 1999) described adaptations of clavagelloids to a tube-dwelling mode of life, and Carter (1978) described how the tubes of gastrochaenids are formed. The gastro- chaenids Cucurbitula and Eufistulana (Morton, 1982, 1983) are convergently very similar to Dianadema and Brechites, respectively, in forming adventitious tubes. However, the shell valves of gastrochaenids do not unite with the tubes. Also, there is no anterior pedal slit nor are there tubules giving access to interstitial waters. Morton (1984a, 2002a) speculated on the process of tube formation in Brechites vaginiferus as, earlier, had Gray (1858b) and Smith (1978). These authors agree that the adventitious tube is secreted but once and that posterior extension is possible either as the animal grows or has to extend itself either to keep pace with an accreting habitat or to ef- fect repair. Because the whole body internal to the tube is covered in thick periostracum, Morton (1984a, 2002a) believed erroneously that the tube of B. vaginiferus was created by a secretion produced from glands in the apex of the siphons pouring down the outside of the periostracum-covered adult, between it and the burrow, to form a structure that matched the configuration and surface structure of the burrow wall. Subsequently, Morton (2002b, 2003) showed that the tubes of Humphreyia strangei and Dianadema multangularis could not be secreted in this way, since both are cemented epibenthically with no burrow tem- plate. Formation probably results from the mantle epithelium secreting sequentially either periostracum or adventitious tube, in a man- ner similar to that described by Savazzi (2000) for the ligament of Bryopa. BIOLOGY OF FOEGIA АЕ Se a NE Burrow wall Mantle Burrow wall Mantle Burrow wall Adventitious tube Mantle Burrow wall SR SE Adventitious tube Mantle Burrow wall Adventitious tube * № «+ > en $ ES ЛЕ a e Mantle Ca its FIG. 15. Foegia novaezelandiae. Generalized illustrations of longitudinal sections through the shell, saddle and adventitous tube showing the postulated method of construction (for abbreviations see рр. 38-39). Dil 52 MORTON In Foegia novaezelandiae the process of tube formation is more complicated than that of other clavagelloids and is illustrated in Fig- ure 15А-Е. Initially, the tube is secreted the same way as in Brechites vaginiferus (Morton, 1984a, 2002a), in that the juvenile shell is cov- ered by periostracum: (Fig. 15A, arrow 1). The animal expands hydrodynamically enlarging its burrow to full adult size, and a second layer of periostracum is then secreted by the mantle and covers the whole body. The anterior and posterior pallial crests secrete this too overthe tiny shell valves (Fig. 15B, arrow 2). Secre- tion of periostracum 2 having halted, the ad- ventitious tube is then produced by the mantle (Fig. 15C). Extra secretions of the tube by the pallial crests produce the bulbous protuber- ances above the true shell valves largely hid- ing them. Internally too, further secretions by the dorsal mantle unite shell valves, saddle and tube, creating the situation whereby the former are effectively incorporated into the total structure of the adventitious tube (Fig. 15D). Finally (Fig. 15E, arrow 3), a further layer of periostracum is produced by the mantle so that the whole animal, within its tube, is now cov- ered in periostracum which is thin and trans- parent dorsally (small arrow 3), and thick and wrinkled all over the rest of the mantle (large arrow 3). The secretion of the adventitious tube of Foegia novaezelandiae is thus highly complex involving the mantle in a sequence of secre- tions of different properties to produce: (i) shell and saddle (covered by periostracum), (ii) a second layer of periostracum, (Ш) the main component of the adventitious tube and, finally, (iv) a third layer of periostracum. This results in the peculiar situation wherein the animal is encased within periostracum, within a tube, within periostracum and within a burrow. The hydrodynamic forces within the mantle and siphons of Foegia novaezelandiae which pump the animal up to its full size before pro- duction of the adventitous tube and subse- quently extend the siphons following contraction, must, as postulated for Brechites vaginiferus (Morton, 2002a), in the absence of any adductor muscles, be created by con- tractions of the pedal disc. In F. novae- zelandiae, the pedal disc must also create the hydrodynamic forces in the haemocoels of the mantle and siphons, acting agonisti- cally with the circular, longitudinal and trans- verse muscles within the latter, to effect siphonal extension. The paired supra-bran- chial osphradia of F novaezelandiae are of interest in this respect. Bivalve osphradia are usually simple structures and generally be- lieved to monitor water flow through the ctenidia (Kraemer, 1981). However, in the case of F. novaezelandiae, perhaps they monitor the complex hydrodynamic forces in the mantle cavity and assist in their synchronisation. Foegia novaezelandiae is also of interest in another respect. The thick anterior cover- ing of periostracum, especially around the pedal disc, is thrown into complex folds not seen in other tube-dwelling clavagelloids, for example, Brechites vaginiferus (Morton, 1984a: fig. 12). It also possesses an external covering of an agglomeration of organic ma- terial and anucleate “cells”. Sand grains and other inorganic detritus covering the siphonal apices exposed to light are not present in the pedal disc agglomeration in the dark. The “cells” are bacteria: might they be sulphide oxidizing? Foegia novaezelandiae is unusual in that it occupies hypoxic mud. Is it possible that it has within the base of its tube and into which interstitial water is pumped, a collec- tion of symbiotic bacteria that help to detoxify the sulphide in the incoming water? Might such bacteria also provide it with a supple- mentary source of nutrition in the form of re- duced carbon and amino acids fixed and produced by them, respectively? This may reduce dependence on short-term inputs of organic matter from the tropical, nutrient de- ficient waters above (Rochford, 1980). This study cannot answer these questions until more intact specimens are available for study. Reid (1990) surveyed the occurrence of chemoautotrophic sulphide oxidizing bacteria in the Bivalvia and showed that they occur within the ctenidial filaments in specialized bacteriocytes and are characteristic of hydro- thermal vent species, for example, Calyptogena and Bathymodiolus, shallow water represen- tatives of the Lucinoidea (Taylor & Glover, 2000) and Solemyoidea, many of which inhabit sulphur-rich sediments (Dando et al., 1986). Foegia novaezelandiae does not have intra- cellular, ctenidial bacteria, but the record of free-living bacteria with characteristics of sul- phide-oxidizing ones on the pedal disc periostracum is of interest and deserves fur- ther study. It is now known that the adventitious tubes of clavagelloids fulfil a number of functions. These are: BIOLOGY OF FOEGIA 93 (i) Creating the rigid external skeleton against which the pedal disc can pump interstitial water into and out of the mantle cavity to generate the hydrodynamic pres- sures necessary in the pallial haemocoels to extend the siphons following retraction. (ii) The same pumping action may supply the animal with interstitial bacteria and dis- solved organic material and mineral salts, which probably act as sources of nutrients accessory to the material collected by suspension feeding from the tropical, nu- trient poor overlying water (Rochford, 1980). (iii) Aeration of the interstitial water may be achieved by pumping mantle cavity water obtained from the sea above via the si- phons into the burrow heading. (iv) Possible detoxification of interstitial wa- ter, by burrow aeration. (v) Possible detoxification of hydrogen sul- phide in the incoming interstitial water by (loosely symbiotic?) chemoautotrophic bacteria and the supply of reduced car- bon and amino acids to the host. Our understanding of the adaptive radiation of the Clavagelloidea increases with each new species studied. It stems from initial, but sepa- rate, adaptations in the Cretaceous (Clavagellidae: Clavagella and Dianadema) and Oligocene (Penicillidae: Brechites and Foegia) to life within a tube but how such ad- aptations arose and from what ancestor(s) are unknown (Harper et al., 2000). ACKNOWLEDGEMENTS | am grateful to F. E. Wells, D. $. Jones and M. Hewitt of the Western Australian Museum, Perth, Western Australia, for organizing field research at Dampier, Western Australia, in 2000, to S. Slack-Smith and G. W. Kendrick, also of the Western Australian Museum, for assistance in accessing the collections of Re- cent and fossil Mollusca, respectively. E. V. Coan of Palo Alto, California, and J. D. Taylor, Natural History Museum, London, are thanked for assistance with the complex taxonomy of Foegia novaezelandiae, and the latter is fur- ther thanked for providing the SEM photograph (Fig. 10) of the pedal disc. The Director and staff of the Western Aus- tralian Museum are thanked for the provision of facilities and many kinds of help and hospi- tality, respectively. LITERATURE CITED ATKINS, D., 1937a, On the ciliary mechanisms and interrelationships of lamellibranchs. Part 3. Types of lamellibranch gills and their food currents. Quarterly Journal of Microscopical Science, 79: 375—421. ATKINS, D., 1937b, On the ciliary mechanisms and interrelationships of lamellibranchs. Part IV. Cuticular fusion with special reference to the fourth aperture in certain lamellibranchs. Quarterly Journal of Microscopical Science, 79: 423-445. BRUGUIERE, M., 1789, Encyclopedie methodique; histoire naturelle des vers, Vol.1 (XV), 126-130, genus 33. Paris: Pankouche. CARTER, J. 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The Nautilus, 93: 93-95. PREZANT, R. S., 1979b, The structure and func- tion of the radial mantle glands of Lyonsia hyalina (Bivalvia: Anomalodesmata). Journal of Zoology, London, 187: 505-516. PURCHON, К. D., 1956, A note on the biology of Brechites penis (L.). Lamellibranchia. Zoologi- cal Journal of the Linnean Society, 43: 43-54. PURCHON, БК. D., 1960, A further note on the biology of Brechites penis (L.). Lamellibranchia. Proceedings of the Malacological Society of London, 34: 19-23. REID, R. G. B., 1990, Evolutionary implications of sulphide-oxidizing symbioses in bivalves. Pp.127-140, т: в. MORTON, ed., The Bivalvia — Proceedings of a Memorial Symposium in Honour of Sir Charles Maurice Yonge, Edinburgh 1986. Hong Kong University Press, Hong Kong, viii + 355 pp. ROCHFORD, D. J., 1980, Nutrient status of the oceans around Australia. CS/RO Division of Fisheries and Oceanography Annual Report, 1977-1979: 9-20. SAVAZZI, E., 1982, Adaptations to tube dwelling in the Bivalvia. Lethaia, 15: 275-297. SAVAZZI, E., 1999, Boring, nestling and tube- dwelling bivalves. Pp. 205-237, in: Е. SAVAZZI, ed., Functional morphology of the invertebrate skeleton. Chichester: Wiley & Sons, x + 706 pp. SAVAZZI, E., 2000, Morphodynamics of Bryopa and the evolution of clavagellids. Pp. 313-327, in: Е. М. HARPER, J. О. TAYLOR & J. A. CRAME, eds., The Evolutionary Biology of the Bivalvia. Lon- don: Geological Society Special Publications, 177: vii + 494 pp. SEMENIUK, V. 8 P.A. S. WURM, 1987, The man- groves of the Dampier Archipelago, Western Australia. Journal of the Royal Society of West- ern Australia, 69: 29-87. SMITH, В. J., 1971, A revision of the family Clavagellidae (Pelecypoda Mollusca) from Australia with descriptions of two new species. Journal of the Malacological Society of Aus- tralia, 2: 135-161. SMITH, В. J., 1976, Revision of the Recent spe- cies of the family Clavagellidae (Mollusca: Bivalvia). Journal of the Malacological Society of Australia, 3: 187-209. SMITH, В. J., 1978, Further notes on the Clavagellidae, with speculation on the process of tube growth. Journal of the Malacological Society of Australia, 4: 77-79. SMITH, B. J., 1998, Superfamily Clavagelloidea. Pp. 412-415, in: P. L. BEESLEY, G. J. В. ROSS 8 А. WELLS, eds., Mollusca: the southern synthe- sis. Fauna of Australia. Volume 5, Part A. Melbourne, Australia: CSIRO Publishing, xvi + 563 pp. SOLIMAN, С. М., 1971, On a new clavagellid bi- valve from the Red Sea. Proceedings of the Malacological Society of London, 39: 389-397. BIOLOGY OF FOEGIA TAYLOR, J. D. & GLOVER, E. A., 2000, The anatomy, chemosymbiosis and evolution of the Lucinidae. Pp. 207-225, in: E. M. HARPER, J. D. TAYLOR & J. A. CRAME, eds., The evolutionary biology of the Bivalvia: London: Geological So- ciety, Special Publications, 177: vii + 494 pp. WELLS, F. E. & C. W. BRYCE, 2000, Seashells of Western Australia. Perth: Western Austra- lian Museum, 207 pp. YONGE, C. M., 1982, Mantle margins with a re- vision of siphonal types in the Bivalvia. Jour- nal of Molluscan Studies, 48: 102-103. Revised ms. accepted 18 August 2003 т FULL : cpa! 5 tot 7 PTT | pit м Are MALACOLOGIA, 2004, 46(1): 57-71 NEW SPECIES OF THE GENUS KELLIA (BIVALVIA: KELLIIDAE) FROM THE COMMANDER ISLANDS, WITH NOTES ON KELLIA COMANDORICA SCARLATO, 1981 Gennady M. Kamenev The Institute of Marine Biology, Russian Academy of Science, Vladivostok 690041, Russia; kamenev@mail333.com, inmarbio@mail.primorye.ru ABSTRACT Anew species, Kellia kussakini, is described from the Commander Islands. This species has a small (to 4.8 mm), translucent, pear-shaped, very inflated, almost globular shell (shell length, height, and width almost equal), with a slightly polished, yellowish-gray periostracum and posteriorly placed beaks. It was found in the subtidal zone (depth 5-20 т) of Bering and Medny islands, on a rocky platform, with population density to 1,190 speci- mens/m?. Scarlato (1981) described Kellia comandorica Scarlato, 1981, from the Com- mander Islands after study of a small amount of material (10 specimens). Later, Coan et al. (2000) synonymized K. comandorica with K. suborbicularis (Montagu, 1803). A study of extensive material (146 specimens) has shown that K. comandorica is a separate species having characters that distinguish it from other species of Kellia. An expanded description of K. comandorica is given. Key words: Kellia, Kelliidae, Bivalvia, Commander Islands, morphology, distribution. INTRODUCTION The bivalve mollusk fauna of the Com- mander Islands shelf has been poorly stud- ied. The most complete species list of bivalve mollusks of the Commander Islands was pub- lished after examination of the extensive ma- terial collected by two joint expeditions of IMB-PRIFO (the sealer “Krylatka”, 1972; RV “Rakitnoye”, 1973) to these islands, as well as an analysis of previous investigations (Kamenev, 1995). However, there still were a number species requiring additional investiga- tion and more accurate identification. Subse- quently, a few papers devoted to the study of these species were published (Kamenev, 1996, 2002; Kamenev & Nadtochy, 2000). Further examination of bivalve mollusks col- lected in the shelf zone of the Commander Islands revealed one new species of the ge- nus Kellia which was erroneously identified as Kellia suborbicularis (Montagu, 1803) (Kamenev, 1995). In addition, another species of this genus, Kellia comandorica Scarlato, 1981, described by Scarlato (1981) based on a small amount of material, is abundant in the intertidal and subtidal zones of the Com- mander Islands. Scarlato (1981) described K. ЭЙ comandorica in detail and provided а compara- tive diagnosis with distinguishing characters of this species, and photos of the holotype. Coan et al. (2000) considered this species as a зупопут of К. suborbicularis. À study of а large quantity of К. comandorica, which is а common mollusk in the Commander Islands, has clearly shown that it is a well-identifiable, separate species of Kellia. The goal of this paper is to describe the new species and ex- pand the description of К. comandorica, with new data on its morphology, ecology, and geo- graphical distribution. MATERIAL AND METHODS In this study | used the material collected by the joint expeditions of IMB-PRIFO in the subtidal zone of the Kuril Islands (the sealer “Krylatka”, September-October 1969) and Commander Islands (the sealer “Krylatka”, July 1972; RV “Rakitnoe”, August-October 1973) and the expedition of IMB in the inter- tidal zone of the Commander Islands (June- August 1972). The material ofthe new species and of K. comandorica from the subtidal zone of the Commander Islands was fixed and stored in 70% ethanol in IMB. Material of K. 58 KAMENEV comandorica from the Kuril Islands and the intertidal zone of the Commander Islands was fixed in 70% ethanol and stored dry in IMB. For comparison purposes, collections of К. suborbicularis — 88 specimens from the North Atlantic (CAS, NHM, NMW) and more 300 specimens from the northeastern Pacific (CAS, UW); of Kellia japonica Pilsbry, 1895 — 2 specimens from Japan (NSMT Mo 73530) and 16 specimens from the Pacific seas of Russia (MIMB); of Kellia porculus Pilsbry, 1904 — 1 specimen from Japan (NSMT Mo 73531); and of Kellia subrotundata (Dunker, 1882) — 1 specimen from (NSMT Mo 73532) were used. All material of these species was stored dry. Shell Measurements Figure 1 shows the shell morphology mea- surements. Shell length (L), anterior end length (A), height (H), width (W) (not shown) were measured for each valve. The ratios of these parameters to shell length (A/L, H/L, W/L, re- spectively) were determined. Shell measure- ments were made using a caliper and an ocular micrometer with an accuracy of 0.1 mm. The following material was measured: (1) 85 specimens, 1 right and 5 left valves of К. comandorica from Urup Island, Kuril Is- lands, (MIMB, 15 specimens, 1 right, 5 left valves) and the Commander Islands (MIMB, IMB, 70 specimens). (2) 97 specimens of the new species from the Commander Islands (IMB). (3) 44 specimens of K. suborbicularis from the North Atlantic: Weymouth, Dorset, Sea area 16, United Kingdom (NMW 1953.183, 24 specimens); Tenby, Pembrokeshire, Sea area 21, United Kingdom (NMW 1953.183, 9 specimens); Guernsey, Channel Is., Sea area 17, United Kingdom (NMW 1953.183, 3 specimens); Plymouth, United Kingdom (NHM 20030382, 2 specimens); Isle of Herm, Guernsey, United Kingdom (NHM 20030383, 2 specimens); England (CAS 165845, 2 specimens); England (CAS 165846, 2 specimens). (4) 21 specimens of K. suborbicularis from the northeastern Pacific: Monterey Bay, Cali- fornia (CAS 161254, 8 specimens); Orcas Island, San Juan Islands, San Juan County, Puget Sound, Washington (CAS 161256, 7 specimens); Alaska (CAS 161255, 6 specimens). Statistics Statistical analysis of the material used a package of statistical programs STATISTICA (Borovikov & Borovikov, 1997) and Data Analy- sis Module of MS Excel 97. The calculated indices (A/L; H/L; W/L) are less susceptible to change compared with other measured parameters. Therefore, the statistical analysis was performed using only these characteristics. All data was tested with a Kolmogorov test for their fit to a normal dis- tribution. The distribution of some indices was different from the norm. Therefore all analy- ses were performed on log,, transformations of the original variables. All indices for pairs of different valves of Kellia species were com- pared using the Student (T) parametric test and one-way analysis of variance (ANOVA). Throughout this study, statistical significance was defined as P < 0.05. Abbreviations The following abbreviations are used in the paper: CAS — California Academy of Sciences, San Francisco; IMB — Institute of Marine Biol- ogy, Russian Academy of Sciences, Vladivostok; MIMB — Museum of the Institute of Marine Biol- ogy, Vladivostok; NHM — The Natural History Museum, London; NMW — National Museums & Galleries of Wales, Cardiff; NSMT — National FIG. 1. Placement of shell measurements: L — shell length; Н — height; А — anterior end length. KELLIA IN THE COMMANDER ISLANDS 59 Science Museum, Tokyo; PRIFO — Pacific Re- search Institute of Fisheries and Oceanography, Vladivostok; UW — University of Washington, Seattle; ZIN — Zoological Institute, Russian Acad- emy of Sciences, St.-Petersburg. SYSTEMATICS Family Lasaeidae Gray, 1842 Genus Kellia Turton, 1822 Type species: Mya suborbicularis Montagu, 1803 Diagnosis Shell small (< 30 mm), thin, ovate to globu- lar, inflated, inequilateral, equivalve. Surface with growth lines. Periostracum thin, adher- ent, colorless, gray, green to yellow. Beaks prosogyrate, almost central. Hinge plate nar- row. Right valve with one cardinal tooth and posterior lateral tooth; left valve with two car- dinal teeth and posterior lateral tooth. Liga- ment internal, partly lodged in a lanceolate resilifer, situated between cardinal and lateral teeth. Pallial line without pallial sinus. Kellia comandorica Scarlato, 1981 (Figs. 2-20, Table 1) Kellia comandorica Scarlato, 1981: 321, pls. 284 (holotype), 285. Kellia suborbicularis (Montagu, 1803), Coan et al., 2000: 323 (partim). Type Material and Locality Holotype (ZIN 9372), Commander Islands, Coll. E. F. Gurjanova, 1930 (Scarlato, 1981). Material Examined 16 lots (MIMB 2989, 2990, 2992, 2994, 2996-3000, 3047-3050, 3052, 3053, 3055) from the tidal zone of Urup Island, Kuril Is- lands (15 specimens, 1 right, 5 left valves); 63 lots (MIMB 2993, 3001, 3002, 3052, 3054, IMB) from the intertidal and tidal zones of the Commander Islands (131 specimens). Total of 146 specimens, 1 right, and 5 left valves. Description (expanded from that of Scarlato,1981) Exterior. Shell small (to 16.8 mm), ovate-an- gular, high (H/L = 0.735-0.976), equivalve, in- flated (W/L of valve 0.198-0.397), inequilateral, thin, solid. Surface with conspicuous, often rather rough growth lines. Periostracum thin, adherent, non-polished, colorless or gray, ex- tending into inner surface. Beaks small, mod- erately projecting above dorsal margin, slightly anterior to midline (sometimes central) (A/L = 0.314-0.5), rounded, prosogyrate. Anterior and posterior ends rounded. Anterodorsal margin slightly convex, gently descending ventrally, smoothly transitioning to slightly curved ante- rior margin. Ventral margin slightly curved. Posterodorsal margin slightly convex, rather steeply descending to rounded posterior mar- gin. TABLE 1. Kellia comandorica Scarlato, 1981. Summary statistics of the shell measurements (mm) and indices: L — shell length; A — anterior end length; H — height; W — width. Numerator indicates the summary statistics for the right valve, denominator — for the left valve. Statistics L A H Mean 9.31 4.26 7.91 9.21 4.21 7.82 SD 0.33 0.15 0.29 0.32 0.15 0.28 SE 3.04 1.41 DAD 2.99 1.39 2.69 Min 3.4 HO) 2.5 3.4 1.6 2.5 Max 16.8 7.8 15.2 16.8 7.8 15.2 n 86 86 86 W A/L H/L W/L 2.47 0.458 0.846 0.264 2.47 0.458 0.846 0.265 0.10 0.003 0.005 0.003 0.10 0.003 0.005 0.003 0.9 0.027 0.049 0.031 0.93 0.027 0.048 0.030 0.9 0.314 0.735 0.198 0.9 0.314 0.735 0.198 Sail 0.500 0.976 0.389 0.500 0.976 0.397 86 86 86 86 90 90 90 90 60 KAMENEV Interior. Right valve with one cardinal tooth and posterior lateral tooth; left valve with two cardinal teeth and posterior lateral tooth. In right valve, cardinal tooth large, elongate, flat- tened, with a flat top, anteroventrally directed, situated at edge of inner part of anterodorsal shell margin; posterior lateral tooth large, long, extending along posterodorsal shell margin. In left valve, anterior cardinal tooth large, elon- gate, flattened, often triangular, anteroventrally directed, situated at edge of inner part of anterodorsal shell margin; posterior cardinal tooth small, rounded, isolated, fingerlike, with rounded top, situated exactly under beak; pos- terior lateral tooth large, long, extending along posterodorsal shell margin. Internal ligament FIGS. 2-10. Kellia comandorica Scarlato, 1981. 2-4: MIMB (3002), Gladky Cape, Medny Island, Commander Islands, intertidal zone, shell length 16.4 mm. 5: ММВ (2994), Lidina Cape, Urup Is- land, Kuril Islands, 20 m, shell length 16.0 mm. 6: Poludennaya Bight, Medny Island, Commander Islands, 20 m, shell length 11.8 mm. 7: Poludennaya Bight, Medny Island, Commander Islands, 20 m, shell length 12.2 mm. 8: Peschany Cape, Medny Island, Commander Islands, shell length 13.1 mm. 9: MIMB (2998), Van-der-Linda Cape, Urup Island, Kuril Islands, 10 m, shell length 14.7 mm. 10: Polovina Cape, Bering Island, Commander Islands, 5 m, dorsal view of both valves of a young speci- men. Bar = 1 mm. KELLIA IN THE COMMANDER ISLANDS 61 FIGS. 11-20. Kellia comandorica Scarlato, 1981. 11-14: Peschany Cape, Medny Island, Commander Islands, 15 m. 11, 12: Right and left valves of an adult specimen. 13, 14: Hinge of right and left valves. 15-20. Phedoskina Cape, Bering Island, Commander Islands, 5 m. 15, 16: Right and left valves of a young specimen. 17, 18: Hinge of right and left valves. 19, 20: Ventral view of hinge of right and left valves showing resilifer. Bar = 500 um. 62 well-developed, large, situated between car- dinal and lateral teeth, posteriorly directed, partly lodged in lanceolate resilifer extending obliquely posterior to beaks. Anterior adduc- tor muscle scar large, rounded; posterior muscle scar large, ovate-angular, longer and wider than anterior scar. Pallial line without pallial sinus. Shell interior with conspicuous radial rows of fossae extending to pallial line. Variability Shell shape and proportions, as well as width of the valves vary markedly (Table 1, Figs. 5— 7). The shell shape varies from ovate-elongate with relatively small shell height to rounded with height almost equal to shell length. The shell is most often slightly angular but sometimes it is regularly ovate without angles. The specimens frequently have a deformed shell because of living in small holes and crevices of boulders and rocky platforms, preventing normal growth. The position of the beaks is also variable. Usu- 140° RUSSIA KAMENEV ally the beaks are anteriorly placed but some- times they occupy the central position. The sizes and shape of cardinal and lateral teeth in both valves vary little. All investigated speci- mens, independent of the age, habitat, and geographic area, had conspicuous radial rows of fossae on the inner shell wall. Distribution and Habitat (Fig. 21) Kellia comandorica occurs near the Com- mander Islands and Urup Island (Kuril Islands). Near Bering Island and Medny Island (Com- mander Islands), K. comandorica is a com- mon species of the bottom fauna. lt was recorded from the intertidal zone to 20 m depth, on boulders and rocky platforms, at a bottom temperature from 4.0 to 10.2*C, with population density to 170 specimens/m?. Near Urup Island (Kuril Islands) this species was found at depth from 5 to 20 m, on boulders and rocky platforms, with population density to 40 specimens/m?. Sea of Okhotsk No 2 Urup Is. Pacific Ocean 160° FIG. 21. Distribution of Kellia comandorica, Scarlato 1981. KELLIA IN THE COMMANDER ISLANDS 63 FIGS. 22-39. Shells of Kellia species. 22-27. Kellia suborbicularis (Montagu, 1803) from the north- eastern Pacific. 22-25: CAS (161254), Monterey Bay, California, 18-22 m, shell length 26.1 mm. 26-27: CAS (161255), Alaska, left valve, shell length 13.8 mm. 28-33. Kellia suborbicularis (Montagu, 1803) from the North Atlantic. 28-29: NMW (1953.183), Tenby, Pembrokeshire, Sea area 21, United Kingdom, right valve, shell length 11.7 mm. 30-31: NHM (20030382), Plymouth, United Kingdom, right valve, shell length 9.0 mm. 32-33: CAS (165845), England, right valve, shell length 8.9 mm. 34-35: Kellia japonica Pilsbry, 1895, NSMT (Mo 73530), Nagashima, Mie Prefecture, Japan, right valve, shell length 11.5 mm. 36-37: Kellia subrotundata (Dunker, 1882), NSMT (Mo 73532), Nosappu Cape, Hokkaido, Japan, right valve, shell length 11.4 тт. 38-39: Kellia porculus Pilsbry, 1904, NSMT (Mo 73531), Ushimado, Okayamo Prefecture, Seto Inland Sea, Japan, right valve, shell length 8.3 mm. эе$$0; OLIS [EIDE ие; — эещз elpel ие} seus |EIPEJ Jule) jo SMOJ |егрел эещз|ерел Jule) seus jeipes Jule) soeuns U}IM sauneuos UJIM saunauos Чум SOWNSWOS yjoous snonoidsuos yjim UJIM saunauos UJIM sauneuos ||э4$ ¡eusaju] seul] saul| JMOIB seul] Soul saul Soul umouß чбпол saul| snon9IdsU09 UMOJÓ }UIE} YM UMOIP уме} ym UMOIB ше} M UMOIB уме} UM = ‘snonoidsuoo yum UMOIP jure, JIM JO JUIEJ UM S9EUNS Jays MOJ9Á-ysIAeJb AesB-ysimoyjaA — Aes ло $$э0]09 Mol|eÂ-usiAe16 Kei6-ysimol|e mole ‘paysiod JO moe ‘paysijod moJjaÁ ‘paysijod 'paysijod Ajybiys 'paysijod-uou JO mojjaAk ‘paysijod ‘peysod AjuBiys wnoejsousd (6£t'0) > au1pru 0} (1970) эмир (2150) aumpiu (8gy'0) эмир (£rt'0) эр LL Jouaque Авуби$ (98£'0) эцчирии 0} Jouajue 0} 1012}504 0} Jouajue (FLr0) эчирии 0} лоизие a ‘ualjoms ‘чб 0} Jouajue ‘чбч А!буо$ 'mo| Anubis ‘ч@ч Anubis ‘Mol 0} лоиэие mo] Anubis ‘мо! (71/v) syeag 2 (055`0) (705`0) (ove'0) (v0€'0) SE pajeyul Алэл (68z'0) payeyu! pajeyur Алэл pajeyyul Алэл (voz'0) разеци! (09z'0) payeyu! pajeyur Алэл (T/M) эмел uuu ‘fuel р O'€L SEL gr 8'91 L'9c Lbb ‘хеш |945 (zv8'0) (088`0) (8/8`0) — (096`0) seINGo\6 (978`0) (608`0) (168`0) (7H) aJeJpenbans-ajeno ¡eouaydsgns ajespenb рэрипол 'padeys-1esd летбие-эело эзебио|э-э}ело лепбие-эело odeus eus pljos pljos pijos зизопзиед ‘эбел pros pros pros 1945 EJEPUNJOIGNS y snjnaiod y eoruodef y IUIMESSNY Y вэиориешоэ y (эшэеа IN) (шорбим рэлип) siajoeleyo suejnoiquogns ‘M sueyjnoiquogns ‘У ‘u}fue] pus Jolajue — y ‘рим эмел — М ‘juBieu — H ‘биз! ¡Jays — 7 'взэюэа$ е/эу jo злэоелецо Ббицециалашиа ‘2 3798vL 64 KELLIA IN THE COMMANDER ISLANDS 65 Comparisons Kellia comandorica is easily distinguished from other species of this genus by its shell with radial rows of fossae on the inner wall (Table 2). Moreover, К. comandorica differs from Kellia kussakini by its larger, lower, and less inflated ovate-angular shell with anteri- orly placed beaks. In shell shape and proportion К. coman- dorica is similar to К. suborbicularis from the North Atlantic. However, with the exception of the above-mentioned distinguishing character, in contrast to X. suborbicularis from the North Atlantic, this species has a less inflated shell with the non-polished periostracum and less anteriorly placed beaks (Table 1, 3). Mean values and variances of the indices charac- terizing the position of beaks (A/L) and the relative width (W/L) were significantly differ- ent in these species (Table 4, Figs. 28-33). Unlike К. comandorica, the shell of К. suborbicularis from the northeastern Pacific is non-angular, rounded-ovate or ovate-elongate with faint growth lines and the polished yellow or grayish-yellow periostracum, interiorly smooth, sometimes with the faint radial striae especially noticeable along the ventral shell margin (Tables 1, 2, 4, 5, Figs. 22-27). In contrast to К. japonica, К. porculus and К. subrotundata, K. comandorica has a less inflated shell with a non-polished gray periostracum, conspicuous growth lines, and less anteriorly placed beaks (Table 2, Figs. 34-39). Remarks In the northwestern Pacific, the Russian and Japanese malacologists recognize another species К. japonica (Scarlato, 1981; Kafanov, 1991; Okutani, 2000), which Coan et al. (2000) also considered a synonym of K. suborbi- cularis. However, the very wide geographic distribution of K. suborbicularis may suggest that it is a composite that includes several species (Scarlato, 1981). The placement of K. japonica in synonymy with К. suborbicularis further extends the range of K. suborbicularis. The study of different-age individuals (from 0.3 mm to 11.7 mm) of К. suborbicularis from the North Atlantic showed that the shell shape varies from ovate-elongate to globular (Figs. 28-33). However, the proportion of speci- mens with the ovate-elongate shell was small. On the whole, in comparison with K. sub- orbicularis from the northeastern Pacific, in- dividuals of this species from the North Atlantic have a relatively small (< 12 mm), markedly more rounded, higher, and more in- flated shell with less anteriorly placed beaks and a slightly polished yellowish-gray periostracum (Tables 3, 5). Mean values and variances of the indices characterizing the position of beaks (A/L), the relative height (H/ L) and width (W/L) in K. suborbicularis from the North Atlantic were significantly different from the mean values and variances of the same indices of this species from the north- eastern Pacific (Table 4). TABLE 3. Kellia suborbicularis (Montagu, 1803) from the North Atlantic (NMW 1953.183; NHM 20030382, 20030383; CAS 165845, 165846). Summary statistics of the shell measurements (mm) and indices: L — shell length; A — anterior end length; H — height; W — width. Numerator indicates the summary statistics for the right valve, denominator - for the left valve. Statistics L A H W AIL H/L WIL Mean 6.91 3.02 5.92 242 0.443 0.857 0.303 6.92 3.02 5.92 2.13 0.442 0.857 0.304 SD 0.32 0.12 0.27 0.11 0.005 0.006 0.005 0.32 0.12 0.27 0.12 0.005 0.006 0.005 SE Dail 0.81 1.80 0.75 0.033 0.037 0.035 al 0.81 1.80 0.77 0.034 0.037 0.036 Min Dun AP Dal 0.6 0.346 0.776 0.211 Dir 12 2x 0.6 0.337 0.776 0.211 Max ТИ 4.6 9.4 3.8 0.500 0.944 0.367 Her 46 9.4 3.8 0.500 0.944 0.367 п 44 44 44 44 44 44 44 44 44 44 44 44 44 44 66 KAMENEV TABLE 4. Results of comparison by pairs of mean values (Student (T) test) and variances (ANOVA) of indices of the right and left valves of Kellia comandorica, К. kussakini, and К. suborbicularis: L - shell length; А - anterior end length; H - height; W — width; P — probability that index values in К. comandorica, К. kussakini, and К. suborbicularis are drawn from the same population; п — number of valves of compared species, respectively; * — significant difference. Right valves Left valves Indices il; E F B n т Р Е E n K. comandorica and K. suborbicularis (North Atlantic) A/L* -2.60 0.006 766 0.006 86/44 -2.73 0.004 8.75 0.004 90/44 H/L 1.45 0.080 1.77 0.187 86/44 1.44 0.008 1.76 0.187 90/44 WIL* 6.24 <0.001 42.69 <0.001 86/44 6.23 <0.001 43.38 <0.001 90/44 K. comandorica and K. suborbicularis (Northeastern Pacific) A/L* 8.23 < 0.001 48.43 < 0.001 86/21 8.43 < 0.001 51.50 <0.001 90/21 H/L* 3.78 < 0.001 10.62 0.002 86/21 3.87 < 0.001 11.24 0.001 90/21 W/L 0.83 0.205 0.47 0.492 86/21 0.87 0.196 0.49 0.486 90/21 К. kussakini and К. comandorica A/L* 1346 <0.001 178.21 <0.001 86/97 13.59 <0.001 179.79 <0.001 86/97 H/L* 11.02 <0.001 125.47 <0.001 86/97 11.25 < 0.001 128.38 < 0.001 86/97 W/L* 15.99 <0.001 252.99 <0.001 86/97 16.04 <0.001 253.16 <0.001 86/97 K. kussakini and K. suborbicularis (North Atlantic) A/L* -12.39 <0.001 160.41 <0.001 97/44 -12.26 < 0.001 159.50 <0.001 97/44 H/L* -8.88 <0.001 76.69 <0.001 97/44 -8.85 <0.001 79.36 <0.001 97/44 WIL* -5.79 < 0.001 35.06 < 0.001 97/44 -5.65 <0.001 33.44 <0.001 97/44 K. kussakini and K. suborbicularis (Northeastern Pacific) A/L* 18.60 <0.001 205.68 <0.001 97/21 18.60 <0.001 205.68 <0.001 97/21 H/L* 11.88 <0.001 147.89 <0.001 97/21 11.89 <0.001 148.39 <0.001 97/21 WI/L* 13.55 <0.001 111.36 <0.001 97/21 13.80 <0.001 109.01 <0.001 97/21 К. suborbicularis (North Atlantic) and К. suborbicularis (Northeastern Pacific) A/L* 4.34 <0.001 13.74 <0.001 44/21 4.24 <0.001 12.95 <0.001 44/21 H/L* 4.82 <0.001 23.61 <0.001 44/21 4.84 <0.001 23.76 <0.001 44/21 WIL* 6.09 <0.001 27.30 <0.001 44/21 6.16 <0001 27:03 0100s 44/241 Se ee. ———unururs m hhuur en Most specimens of K. suborbicularis from the pared. However, a detailed study of this ques- northeastern Pacific had a ovate-elongate, markedly less inflated shell, reaching more than 26 mm in length, with more anteriorly placed beaks and yellow, strongly polished periostracum (Figs. 22-27, Table 5). The specimens with an analogous shell shape (more 20 mm in length) were also recorded off the Kuril Islands (materials from the expe- dition with the R/V “Akademik Oparin”, 1 July — 4 August 2003). Allthese morphological char- acteristics are observed in Kellia laperousii (Deshayes, 1839) (Oldroyd, 1925; Scarlato, 1981), which Coan et al. (2000) also consid- ered to be a synonym of K. suborbicularis. They stated that no consistent difference from the North Atlantic K. suborbicularis can be found when similar-sized specimens are com- tion is needed. Taking into account the results of comparison of specimens of K. suborbicula- ris from the North Atlantic and the northeast- ern Pacific, | think that most likely a separate species К. laperousii occurs in the North Pa- cific. The species of the genus Kellia - K. Japonica, К. porculus and К. subrotundata — living off the coast of Japan (Okutani, 2000), are most similar to K. suborbicularis in shape and shell proportions (Tables 2, 3, Figs. 28- 39). In addition, the species from Japan are also very similar to each other. Some differ- ences between these species exist in the de- gree of inflation of the shell, the position and form of the beaks, and the color of the periostracum (Table 2). However, the differ- KELLIA IN THE COMMANDER ISLANDS 67 TABLE 5. Kellia suborbicularis (Montagu, 1803) from the northeastern Pacific (CAS 1612564, 161255, 161256). Summary statistics of the shell measurements (mm) and indices: L — shell length; A — anterior end length; H — height; W — width. Numerator indicates the summary statistics for the right valve, denominator — for the left valve. Statistics L A H Mean 15.56 6.40 12.55 15.56 6.40 12.55 SD 0.97 0.37 0.75 0.97 0.37 0.75 SE 4.47 1.70 3.44 4.47 1170) 3.44 Min 9.4 42 7.3 9.4 4.2 YES Max 26.1 10.4 19.4 26.1 10.4 19.4 n 21 РИ РИ 21 21 21 ence in shell proportions can be attributed to the individual variability of one species. Be- cause only a very limited material on Kellia species from Japan was at my disposal, no final conclusion can be made. It is not improb- able that after a detailed study of more exten- sive material, these species prove to be a synonym of one species different from K. suborbicularis or K. laperousii. Kellia kussakini Kamenev, new species Figs. 40—53, Table 6 Type Material and Locality Holotype (MIMB 7770), Phedoskina Cape, Bering Island, Commander Islands, Pacific Ocean, 5 m, rocky platform, bottom water tem- perature of 10.0°C, Coll. М. |. Lukin, 23-1X-1973 (RV “Rakitnoye”); paratypes (96) (ММВ 7771) from holotype locality. Other Material Examined 38 specimens with slightly damaged shells from type locality; 1 specimen from Najushka Bight, Bering Island, Commander Islands, Pacific Ocean, 20 m, rocky platform, bottom water temperature of 9.8°C, Coll. G. T. Beloko- nev, 24-1Х-1973 (RV “Rakitnoye”); 1 specimen from Nerpichy Cape, Bering Island, Com- mander Islands, Pacific Ocean, 10 m, rocky platform, bottom water temperature of 7.4°C, Coll. V. I. Lukin, 21-VII-1972 (the sealer “Krylatka”); 2 specimens from Peschany Cape, W A/L H/L W/L 4.06 0.414 0.809 0.259 4.08 0.414 0.809 0.260 0.30 0.004 0.008 0.005 0.30 0.004 0.008 0.005 1.36 0.020 0.038 0.023 1.37 0.020 0.038 0.021 2.0 0.380 0.743 0.213 2.0 0.380 0.743 0.213 8.2 0.463 0.873 0.314 8.2 0.463 0.873 0.314 21 21. 21 21 21 21 21 21 Medny Island, Commander Island, Bering Sea, 10 m, boulders, bottom water temperature of 5.0°C, Coll. V. P. Kashenko, 10-VII-1972 (the sealer “Krylatka”); 5 specimens from Poludennaya Bight, Medny Island, Com- mander Island, Pacific Ocean, 15 m, rocky platform, bottom water temperature of 4.2°C, Coll. V. I. Lukin, 17-VII-1972 (the sealer “Krylatka”); 1 right valve from Vodopadsky Cape, Medny Island, Commander Islands, Pacific Ocean, 20 m, rocky platform, bottom water temperature of 7.6°C, Coll. G. T. Belokonev, 11-IX-1973 (RV “Rakitnoye”). To- tal of 47 specimens and 1 right valve. Description Exterior. Shell very small (to 4.8 mm), pear- shaped, with slightly narrowing anterior end, almost globular, very high (H/L = 0.833-1.036, shell height almost equal to or sometimes greater than length), equivalve, very inflated (W/L of valve = 0.278-0.429, shell width al- most equal to length), inequilateral, thin, frag- ile, translucent. Surface with faint growth lines. Periostracum thin, adherent, slightly polished, yellowish-gray, extending into inner surface. Beaks small, high, strongly projecting above dorsal margin, slightly posterior, sometimes central or slightly anterior (A/L = 0.371-0.579), rounded, prosogyrate. Anterior end slightly narrowed, rounded, lower than posterior shell end. Posterior end rounded. Anterodorsal margin slightly convex, rather steeply de- scending ventrally, smoothly transitioning to 68 KAMENEV strongly curved anterior margin. Ventral mar- gin strongly curved. Posterodorsal margin slightly convex, steeply descending ventrally, smoothly transitioning to curved posterior margin. Interior: Right valve with one cardinal tooth and posterior lateral tooth; left valve with two cardinal teeth and posterior lateral tooth. In right valve, cardinal tooth large, elongate, flat- tened, with rounded or flat top, anteroventrally directed, situated at edge of inner part of anterodorsal shell margin; posterior lateral tooth large, long, extending along postero- dorsal shell margin. In left valve, anterior car- dinal tooth larger, than posterior, elongate, flattened, with rounded top, anteroventrally directed, situated at edge of inner part of anterodorsal shell margin; posterior cardinal tooth smaller, rounded, isolated, fingerlike, with rounded top, situated exactly under beak; posterior lateral tooth large, long, ex- KELLIA IN THE COMMANDER ISLANDS 69 tending along posterodorsal shell margin. Internal ligament well developed, large, situ- ated between cardinal and lateral teeth, pos- teriorly directed, partly lodged in lanceolate resilifer extending obliquely posterior to beaks. Anterior adductor muscle scar large, rounded; posterior muscle scar large, ovate- angular, longer and wider than anterior scar. Pallial line without pallial sinus. Shell interior smooth, without radial rows of fossae or striae. Variability Shell shape and proportions change little with age. In young specimens (< 2.5 mm), the shell is less high, less inflated, and beaks are more pos- teriorly placed. In adults, shell height and width, the position of beaks vary slightly (Table 6, Figs. 43—46). As a whole, shell remains pear-shaped, with more narrow anterior end, and almost globu- lar because of almost absolute equality of the length, height, and width. The beaks are mostly posteriorly placed (the beaks of 8 out of 97 mea- sured specimens are anteriorly placed). The sizes and shape of cardinal and lateral teeth in both valves vary little. Afew specimens had three car- dinal teeth in the left valve (Fig. 53). Distribution and Habitat (Fig. 54) This species was recorded near Bering and Medny islands, Commander Islands, at a depth from 5 to 20 m, on boulders and rocky platforms, at a bottom water temperature from 4.2°C to 10.0°C, with population density to 1,190 specimens/m? (type locality). Comparisons This species is easily distinguished from other species of this genus by its small, al- most globular, very high, and inflated shell with posteriorly placed high beaks (Table 2). Mean values and variances of the indices charac- terizing the position of beaks (A/L), the rela- FIGS. 49-53. Kellia kussakini Kamenev, new species, paratypes (MIMB 7771) from holotype locality. 49, 50: Hinge of right and left valves. 51, 52: Ventral view of hinge of right and left valves showing resilifer. 53: Hinge of left valves with three cardinal teeth. Bar = 300um. 70 KAMENEV Bering Sea Pacific Ocean 166° FIG. 54. Distribution of Kellia kussakini ( № - type locality). TABLE 6. Kellia kussakini Kamenev, new species. Shell measurements (mm), indices of holotype (MIMB 7770), and summary statistics of holotype and paratypes (ММВ 7771) characters: L — shell length; A — anterior end length; H — height; W — width. Numerator indicates shell measurements, indices, and summary statistics for the right valve, denominator — for the left valve. Statistics E A H W A/L H/L W/L Holotype 4.0 2 3.8 1.4 05525 0.950 0.350 4.0 21 3.8 1.4 01525 0.950 0.350 Holotype and paratypes Mean 2.95 1252 214 1.01 0:5 0.917 0.340 2.95 52 271 1.01 051% 0.917 0.340 D 0.06 0.03 0.06 0.03 0.003 0.004 0.003 0.06 0.03 0.06 0.03 0.003 0.004 0.003 SE 0.60 0.29 0.59 0.27 0.027 0.037 0.034 0.60 0.29 0.59 0.27 0.027 0.037 0.034 Min mle 0.8 12 0.4 0.444 0.833 0.278 1.4 0.8 1.2 0.4 0.444 0.833 0.278 Мах 4.8 2.6 4.6 Re) 0.579 1.036 0.429 4.8 2.6 4.6 1.8 0.579 1.036 0.429 n 97 97 97 97 97 97 97 97 97 97 97 97 97 97 KELLIA IN THE COMMANDER ISLANDS TA tive height (H/L) and width (W/L) in K. kussakini were significantly different from the mean val- ues and variances of the same indices of K. comandorica and K. suborbicularis (Table 4). Besides, unlike K. comandorica, it has a smooth interior shell wall without the radial rows of fossae. In shell shape, K. kussakini is close to K. porculus (Figs. 38, 39), living off the coast of Japan, from which it is distinguished in having a more high, very inflated, fragile and translucent shell with posteriorly placed beaks and a slightly polished, yellowish-gray periostracum (Table 2). Etymology The specific name honors Oleg G. Kussakin, Academician of the Russian Academy of Sci- ences, a famous Russian researcher of the marine fauna of the intertidal zone of Russian Pacific seas and world isopod fauna, who de- voted all his life to the study of the northwest- ern Pacific fauna. ACKNOWLEDGMENTS | am very grateful to Dr. V. A. Nadtochy (PRIFO, Vladivostok) and Mrs. N. V. Kameneva (MIMB, Vladivostok) for great help during work on this manuscript; to Professor T. W. Pietsch and Dr. K. Stiles (UW, Seattle) for arrangement of my visit to the UW and work with the bivalve mollusk collection, for all-round, very kind, and friendly help during my life and work in Seattle; to Professor A. J. Kohn and Dr. G. Jensen (UW, Seattle) for great help during work with the bi- valve mollusks collection at the UW; to Mr. Gary Cook (Berkeley) for all-round, very kind and friendly help during my life and work in San Fran- cisco; to Dr. P. D. Roopnarine and Miss E. Kools (Department of Invertebrate Zoology, CAS, San Francisco) for arrangement of my work with the bivalve mollusks collection at the CAS and great help during this work; to Dr. H. Saito (NSMT, Tokyo) for sending the specimens of K. japonica, K. porculus, and K. subrotundata; to Dr. Gra- ham Oliver, Ms. Harriet Wood (NMW, Cardiff), Dr. David G. Reid, and Mrs. Amelia MacLellan (NHM, London) for sending the specimens of K. suborbicularis from the North Atlantic; to Dr. E. V. Coan (Department of Invertebrate Zool- ogy, CAS, San Francisco) for consultations, help in communication with other specialists, and comments on the manuscript; to Professor G. J. Vermeij (University of California at Davis, Davis) for help in communication with other spe- cialists; to Mr. D. V. Fomin (IMB, Vladivostok) for help in work with the scanning microscope; to Ms. T. М. Kaznova (IMB, Vladivostok) for help with translating of the manuscript into English; to Dr. George M. Davis for help in the publica- tion of the manuscript; to Dr. J. A. Allen for com- ments on the manuscript. This research was supported by Grant 01- 04-48010 from the Russian Foundation for Basic Research. LITERATURE CITED BOROVIKOV, V. P. & |. Р. BOROVIKOV, 1997, “STATISTICA”. Statistical analysis and processing of data using “WINDOWS”. Filin Press, Moscow, 608 pp. [in Russian]. COAN, Е. V., Р. H. SCOTT & Е. R. BERNARD, 2000, Bivalve seashells of western North America. Marine bivalve mollusks from Arctic Alaska to Baja California. Santa Barbara Museum of Natural History, 764 pp. KAFANOV, A. 1., 1991, Shelf and continental slope bivalve molluscs of the northern Pacific Ocean: a check-list. Far Eastern Branch of the Academy of Sciences of the USSR, Vladivostok, 200 pp. [in Russian, with English summary]. KAMENEV, С. M., 1995, Species composition and distribution of bivalve mollusks on the Commander Islands shelf. Malacological Review, 28: 1-23. KAMENEV, С. M., 1996, Additional data on morphology and geographic distribution of Adontorhina cyclia Berry, 1947 (Bivalvia: Thyasiridae), newly reported from the northwestern Pacific. The Veliger, 39(3): 213- 219. KAMENEV, G. M., 2002, Genus Parvithracia (Bivalvia: Thraciidae) with descriptions of a new subgenus and two new species from the northwestern Pacific. Malacologia, 44(1): 107- 134. KAMENEV, G. M. & V. A. NADTOCHY, 2000, Mendicula ferruginosa (Forbes, 1844) (Bivalvia, Thyasiridae) from the Far Eastern Seas of Russia. Ruthenica, 10(2): 147-152. [in Russian, with English abstract]. OKUTANI, T., 2000, Marine mollusks in Japan. Tokai University, Tokyo, Japan. xlvii + 1175 pp., incl. 542 pls. OLDROYD, |. S., 1925, The marine shells of the west coast of North America, Vol. 1 [Bivalvia]. Stanford University, Publications, University Series, Geological Sciences, 1(1): 247 pp., 57 pls. SCARLATO, O. A., 1981, Bivalve mollusks of temperate waters of the northwestern Pacific. “Nauka” Publ. House, Leningrad, 480 pp. [in Russian]. Revised ms. accepted 28 August 2003 MALACOLOGIA, 2004, 46(1): 73-78 REVISION OF THE REPRODUCTIVE MORPHOLOGY OF THREE LEPTAXIS SPECIES (GASTROPODA, PULMONATA, HYGROMIIDAE) AND ITS IMPLICATION ON DART EVOLUTION Joris М. Koene* & Igor V. Muratov? ABSTRACT Many species of land snails have one or more sharp, calcareous “love darts” that are used to stab the partner during mating. These darts are produced and stored in special- ized organs called stylophores. Because their number and position varies among species, stylophores are often used for identification and classification, especially in the family Hygromiidae. Having several stylophores, and thus several darts, is presumably the an- cestral state from which species with one stylophore evolved. Species with small acces- sory sacs or rudimentary stylophores located above the functional stylophore are therefore thought to represent intermediate forms between species with double and single stylophores. We investigated the stylophores, darts, and associated reproductive organs of three spe- cies of the hygromiid genus Leptaxis — L. erubescens, L. nivosa and L. undata. In all the specimens of the investigated species, a small sac located just above the stylophore was found to be present. We conclude that this previously overlooked organ represents a rudi- ment of a stylophore, leading us to conclude that Leptaxis should be considered as an intermediate form in the evolution towards a single stylophore in the Hygromiidae. Keywords: love dart, dart sac, stylophore, snail, stylommatophora, Helicoidea, rudiment. INTRODUCTION In the reproductive system of many her- maphroditic land snail species, one or more sharp, calcareous structures are present. When these are produced in a specialized or- gan, the stylophore (also referred to as dart sac), they are called darts. In many species, these “love darts” are stabbed through the partner’s skin during mating (Adamo & Chase, 1990; Reyes Tur et al., 2000). In Helix aspersa (Muller, 1774) — often called Cornu aspersum, Cryptomphalus aspersus, or Cantareus aspersus — this “dart shooting” results in the transfer of an allohormone that inhibits sperm digestion and thereby increases sperm storage and fertilization success (Koene & Ter Maat 2001; Koene & Chase, 1998a; Rogers & Chase, 2001, 2002; Landolfa et al., 2001; Landolfa, 2002). Recently, a comparative study demonstrated that the evolution of darts may be driven by sexual conflict (Koene & Schulenburg, submit- ted), thus explaining the diversity in number and shape of darts. For example, Trichia has two conical darts without blades (Schileyko, 1978a); Leptaxis and Hygromia each have one dart with two (differently arranged) blades (re- spectively: Spence, 1911; Giusti & Manganelli, 1987); Helix has a dart with four blades (Hasse et al., 2002); and Monachoides has one dart with seven blades (Koene & Schulenburg, submitted). Some species, such as Cepaea nemoralis and C. hortensis, which are other- wise remarkably similar, can most easily be distinguished by the shape of their darts (Kerney et al., 1983). Despite the large diver- sity in shapes, darts are rarely used for taxo- nomic purposes. Conversely, stylophores are traditionally used for identification and classi- fication of land snails within the superfamily Helicoidea (Nordsieck, 1987; Schileyko, 1989). Species with one stylophore are thought to have evolved from ancestral species bear- ing several stylophores (Schileyko, 1989). When more than one stylophore is present, different arrangements are possible. Several stylophores can be arranged around the vagi- nal duct (e.g., Humboldtiana: Thompson & Brewer, 2000). Two pairs of stylophores can be present on opposing sites of the vaginal ‘Faculty of Earth and Life Sciences, Vrije Universiteit, De Boelelaan 1085, 1081 HV Amsterdam, The Netherlands; joris.koene@falw.vu.nl 2Department of Malacology, The Academy of Natural Sciences, Philadelphia, Pennsylvania, USA 74 КОЕМЕ & MURATOV duct (e.g., Trichia: Schileyko, 1978а). Alterna- tively, only one pair of stylophores can be present (Hygromia: Giusti 8 Мапдапей, 1987). In these latter two cases, within each stylophore pair, the stylophore that is furthest away from the genital opening normally does not contain a dart, but see Taniushkin et al. (1999) for a possible case of atavistic devel- opment of darts in the upper stylophores of Xeropicta krynickii (Krynicki, 1833). There are also species with morphologies that clearly represent intermediate stages be- tween the above-mentioned forms. In such cases, a stylophore has become reduced in size and no longer produces a dart; such a rudimentary organ is then often referred to as an accessory sac (Nordsieck, 1993), internal dart sac (Giusti & Manganelli, 1987), or upper stylophore (Schileyko, 1989). Because these terms all describe the same organ, probably at different stages of reduction, we have cho- sen to use the term upper stylophore through- out the rest of this paper. Obviously, these intermediates provide important information about the course of evolution of the stylo- phore(s). Because rudimentary organs can be greatly reduced in size, they have sometimes been overlooked in previous studies. This is the case for the genus Leptaxis, which is why we redescribe the morphology of the stylophores, darts and associated reproduc- tive organs of three species of this genus. MATERIAL AND METHODS Species of the genus Leptaxis inhabit Macaronesia, which includes 32 islands grouped in five major archipelagos: Azores, Canaries, Cape Verde, Madeira, and Selvagens Leptaxis erubescens Leptaxis undata Leptaxis nivosa FIG. 1. Shells of the investigated Leptaxis species. LEPTAXIS DART EVOLUTION O (Mitchell-Thomé, 1976). We have focused on the reproductive morphology of species en- demic to the Madeiran archipelago. Among these, Leptaxis erubescens (Lowe, 1831) is the only species of this genus that occurs on all the different island groups — Madeira, Porto Santo, and The Desertas — of this volcanic archipelago (Cook, 1996). The other Leptaxis species are confined to one of the island groups (Cook, 1996). Of these species we in- vestigated, L. undata (Lowe, 1831) from Ma- deira and L. nivosa (Sowerby, 1824) from Porto Santo. Figure 1 shows the shells of the inves- tigated species. Dry and alcohol preserved specimens of L. erubescens (N = 2) were obtained from the malacological collection of the Academy of Natural Sciences of Philadelphia (ANSP 128459 A9427H). Several specimens (frozen at -80°C) of this species (N = 4), as well as of L. undata (N = 4) and L. nivosa (N = 5), were generously made available to us by P. Van Riel (Royal Belgian Institute of Natural Sciences, Brussels). The specimens of each species, which were all adult, were dissected to remove the repro- ductive tract. Subsequently, the reproductive organs were drawn using a camera lucida. To Leptaxis erubescens L. nivosa AG KPSC y HD $0 0.2 ст Br L. undata \ DG LS de 22 22 EL } А VD US ) > x Р PRM 0.2 ст | °С\ 0.2 ст FIG. 2. Comparison of the position and relative size of the upper and lower stylophores of Leptaxis erubescens, L. undata , and L. nivosa. The reproductive system of L. erubescens is shown to depict the other reproductive structures that are mentioned т the text. Abbreviations: A, appendage; AG, albumen gland; BC, bursa copulatrix; BT, bursa tract; DG, digitiform gland; FL, flagellum; FPSC, fertilization pouch-spermathecal complex; G, genital pore; HD, hermaphroditic duct; LS, lower stylophore; P, penis; PRM, penis retractor muscle, SO, spermoviduct; US, upper stylophore; V, vaginal duct; VD, vas deferens. 76 КОЕМЕ & MURATOV avoid damage of the darts, the stylophores were carefully cut out of the reproductive tracts and placed overnight in 1N МаОН, which dis- solved all the tissue and mucus but left the dart intact. For cross-sections darts were care- fully broken in two. The intact and broken darts were consecutively prepared for electron mi- croscopy by placing them on small aluminium plates with an electrically conducting adhesive (Leit-Tab, Plano). Subsequently, they were coated with gold using a Metalloplan (Leitz). The darts were then placed under a scanning electron microscope (S-530 SEM, Hitachi) and photos were taken. RESULTS In all the mature specimens of each species, one large stylophore, containing one dart, was present. This stylophore was positioned in such L. erubescens L. undata а way that it curved slightly around the vaginal duct. Besides this stylophore, we also found a small sac situated between the larger stylophore and the vaginal duct in all species. The position of this organ suggests that we are dealing with the rudiment of an upper stylophore. Addition- ally, a flattened, non-hollow appendage at the base of the vagina is present. Figure 2 gives an overview of the morphology of the investigated Leptaxis species showing the positions and rela- tive sizes of the stylophores, the small sac (i.e., upper stylophore) and the appendage. The two mucus glands of each of the species are situ- ated above the stylophore around the vaginal duct. Each of these digitiform glands has sev- eral branches that join а the base. These glands, as well as the rest of the reproductive system, are only depicted for L. erubescens. The darts of all three species have a round base and a broad corona by which they are at- tached to a tubercle in the stylophore (Fig. 3). L. nivosa 250 um | 250 um FIG. 3. Electron microscopic pictures of the darts of Leptaxis erubescens, L. undata, and L. пмоза. LEPTAXIS DART EVOLUTION TT Approximately halfway towards the tip of the dart the curved shaft broadens and flattens out, thus forming two large blades (Fig. 3). The dart is curved and lightly contorted, which is illus- trated by the electron microscopic picture of the side view of a dart of L. erubescens (Fig. 3), also reflecting the shape of the stylophore. DISCUSSION It has long been thought, based on morpho- logical data, that the genus Leptaxis fully con- forms to the European Hygromiidae with one stylophore (Mandahl-Barth, 1943; Backhuys, 1975; Schileyko, 1989). Interestingly, Pilsbry (1894: 292-293) stated after having investi- gated several Leptaxis species: “| had ex- pected to find in Leptaxis some archaic characters preserved; for its geographic posi- tion and the shell-peculiarities argue for the group an ancient origin; but the evidence shows that however remote in the past the type was derived from the continental fauna, the main anatomical features of modern European Helices were then well established”. Our find- ing of the small organ just above the stylophore in the investigated species of Leptaxis sug- gests that Pilsbry was correct in expecting some ancient characters. The position of the previously overlooked organ is consistent with the position of the upper stylophore in the genus Trichia (Schileyko, 1978a) and the internal or acces- sory dart sac in the genus Hygromia (Giusti & Manganelli, 1987; Nordsieck, 1987). There- fore, we conclude that the investigated spe- cies of the Leptaxis genus possess a rudiment of an upper stylophore. This rudiment has probably been overlooked for so long because the small organ is well hidden in connective tissue between the vaginal duct and the much larger functional stylophore that contains the contorted dart. Nevertheless, the presence of the upper stylophore in Leptaxis has impor- tant implications for the phylogenetic position of this genus within the Hygromiidae. Much of the molluscan phylogeny is heavily based on traits of the reproductive morphology and, es- pecially within the Hygromiidae, the presence and number of (reduced) stylophores play an important role in the classification within the family (Nordsieck, 1987, 1993; Schileyko, 1989). Several observations can be made with re- spect to the reproductive morphology of the family Hygromiidae. There are clear morpho- logical differences between the phylogeneti- cally older subfamily Trichiinae and the younger subfamily Hygromiinae. All Trichiinae have two pairs of stylophores, that is, two up- per and two lower stylophores (e.g., Trichia: Schileyko, 1978a). Most of the differences in the stylophore morphology between genera of Trichiinae are relatively small, while important morphological changes are found within the Hygromiinae. In this subfamily, one pair of stylophores has been lost, consequently many species have one upper and one lower stylophore (e.g., Hygromia: Giusti & Manganelli, 1987). Additionally, a further reduction of the upper stylophore and an enlargement of the lower stylophore occurred (e.g., Leptaxis: this paper; Lindholmomneme: Schileyko, 1978b), culminating with total loss of the upper stylophore (e.g., Monachoides: Schileyko, 1978b, 1989). Simultaneously with this evolution towards a single stylophore, the dart seems to be- come more elaborate. Perpendicular blades on the dart occur in several genera of Hygromiinae, resulting in different dart shapes, and increasing the dart’s surface area. Presumably, this allows the dart to transfer larger amounts of the product from the mucus glands (Fedoseeva, 1994; Adamo & Chase, 1996; Koene & Schulenburg, sub- mitted). However, it is still unclear whether the hygromiid dart is used in a similar way as the helicid dart (Koene & Chase, 1998a, b; Rogers & Chase, 2001; Landolfa et al., 2001) to transfer an allohormone (Koene & Ter Maat, 2001, 2002). Hence, behavioural data are required to determine how the Leptaxis dart is used. Observations of the mating behaviour of Leptaxis may also shed light on the function of the appendage at the base of the genital system (see also Mandahl-Barth, 1943). To conclude, we found the rudiment of an upper stylophore in three species of Leptaxis, which has previously gone unnoticed (Mandahl-Barth, 1943; Backhuys, 1975, Schileyko, 1989). The presence of this small organ is of importance because it indicates that Leptaxis links Hygromiinae with two (upper and lower) stylophores (e.g., Lindholm- omneme) and Hygromiinae with single stylophores (e.g., Monachoides). Therefore, our findings lead us to conclude that this ge- nus is an intermediate form in the evolution towards a single stylophore. 78 КОЕМЕ & MURATOV ACKNOWLEDGEMENTS We are grateful to P. Van Riel for generously providing some of the specimens. We thank H. Reise, P. Van Riel, and H. Schulenburg for valu- able comments and discussion, and J. Lange and C. Levesque for their technical assistance. JMK was supported by a Jessup-McHenry Award of the Academy of Natural Sciences of Philadelphia and a Casimir-Ziegler Stipend of the Royal Netherlands Academy of Arts and Sciences. LITERATURE CITED ADAMO, S.A. & В. CHASE, 1990, The “love dart” of the snail Helix aspersa injects a pheromone that decreases courtship duration. Journal of Experimental Zoology, 255: 80-87. ADAMO, S. A. & R. CHASE, 1996, Dart shoot- ing in helicid snails: an “honest” signal or an instrument of manipulation? Journal of Theo- retical Biology, 180: 77-80. BACKHUYS, W., 1975, Zoogeography and tax- onomy of the land and freshwater molluscs of the Azores. Backhuys & Meesters, Amsterdam, xii + 350 pp., 32 pls. COOK, L. M., 1996, Habitat, isolation and the evolution of Madeiran landsnails. Biological Journal of the Linnean Society, 59: 457—470. FEDOSEEVA, E.A., 1994, Comparative morphol- ogy of darts in the superfamily Helicoidea (Gas- tropoda, Pulmonata). Ruthenica, 4: 103-110. GIUSTI, Е. & С. MANGANELLI, 1987, Notulae Malacologicae, XXXVI. On some Hygromiidae (Gastropoda: Helicoidea) living in Sardinia and in Corsica. (Studies on the Sardinian and Corsican Malacofauna VI). Bollettino Malacologico, 23: 123-206. HASSE, B., J. C.MARXEN, W. BECKER, H. EHRENBERG 4 M. EPPLE, 2002, A crystallo- graphic study of the love dart (gypsobelum) of the land snail Helix pomatia (L.). Journal of Molluscan Studies, 68: 249-254. KERNEY, М. P., R. A. D. CAMERON & J. H. JUNGBLUTH, 1983, Die Landschnecken Nord- und Mitteleuropas. Verlag Paul Parey, Ham- burg 8 Berlin. KOENE, J. M. 8 R. CHASE, 1998a, Changes in the reproductive system of the snail Helix aspersa caused by mucus from the love dart. Journal of Experimental Biology, 201: 2313-2319. KOENE, J. M. & R. CHASE, 1998b, The love dart of Helix aspersa Miller is not a gift of calcium. Journal of Molluscan Studies, 64: 75-80. KOENE, J. M. & H. SCHULENBURG, Shooting darts in hermaphroditic land snails: counter- adaptive co-evolution. Submitted. KOENE, J. M. & A. TER MAAT, 2001, “Allo- hormones”: a class of bioactive substances favoured by sexual selection. Journal of Com- parative Physiology A, 187: 323-326. KOENE, J. M. & A. TER MAAT, 2002, The dis- tinction between pheromones and allohor- mones. Journal of Comparative Physiology A, 188: 163-164. LANDOLFA, M. A., 2002, On the adaptive func- tion of the love dart of Helix aspersa. Veliger, 45: 231-249. LANDOLFA, М. A., О. М. GREEN & К. CHASE, 2001, Dart shooting influences paternal repro- ductive success in the snail Helix aspersa (Pulmonata, Stylommatophora). Behavioural Ecology, 12: 773-777. MANDAHL-BARTH, G., 1943, Systematische Untersuchungen úber die Heliciden-Fauna von Madeira. Abhandlungen der Senckenbergischen Naturforschenden Gesellschaft, 469: 1-93. MITCHELL-THOME, R. C., 1976, Geology of the Middle Atlantic Islands. Beitrage zur regionalen geologie de Erde, 12. Gebrüder Borntraeger, Berlin, ix + 282 pp., 6 pls. NORDSIECK, H., 1987, Revision des Systems der Helicoidea (Gastropoda: Stylommatophora). Archiv ftir Molluskenkunde, 118: 9-50. NORDSIECK, H., 1993, Das System der palaarktischen Hygromiidae (Gastropoda: Stylommatophora: Helicoidea). Archiv fur Molluskenkunde, 122: 1-23. PILSBRY, H. A., 1894, Manual of Conchology. Second Series: Pulmonata, vol. IX. Philadelphia. REYES TUR, B., A: Е. VEEAZQUES <& YO: CABRERA, 2000, Conducta de apareamiento y aspectos de la relación des sistema reproductor en Polymita muscarum Lea 1834 (Gastropoda: Pulmonata). (Mating behaviour and some aspects of the structure function re- lationship in reproductive system of Polymita muscarum Lea 1834 (Gastropoda: Pulmonata)). Revista Biología, 14: 160-166. ROGERS, D. & К. CHASE, 2001, Dart receipt promotes sperm storage in the garden snail Helix aspersa. Behavioural Ecology and So- ciobiology, 50: 122-127. ROGERS, D. 8 R. CHASE, 2002, Determinants of paternity in the garden snail Helix aspersa. Behavioural Ecology and Sociobiology, 52: 280-295. SCHILEYKO, A. A., 1978a, On the systematics of Trichia s. lat. (Pulmonata: Helicoidea: Hygromiidae). Malacologia, 17: 1-56. SCHILEYKO, A. A., 1978b, Molljuski (Helicoi- dea). Fauna SSSR, 3(6): 1-384, Leningrad. SCHILEYKO, A. A., 1989, Taxonomic status, phylogenetic relations and system of the Helicoidea sensu lato (Pulmonata). Archiv fúr Molluskenkunde, 120: 187-236. SPENCE, G. C., 1911, On the dart of Helix undata Love. Journal of Conchology, 13: 210. TANIUSHKIN, А.1., $. S. ZHILTSOV & A. М. SUVOROV, 1999, A cause of occurrence of two darts in the upper stylophore in Xeropicta krynickii (Pulmonata Hygromiidae). Ruthenica, 9: 163-164. THOMPSON, Е. С. & С. P. BREWER, 2000, Landsnails of the genus Humboldtiana from northern Mexico (Gastropoda, Pulmonata, Helicoidea, Humboldtianidae). Bulletin of the Florida Museum of Natural History, 43: 49-77. Revised ms. accepted 15 December 2003 MALACOLOGIA, 2004, 46(1): 79-125 PRELIMINARY PHYLOGENETIC STUDY OF BRADYBAENIDAE (GASTROPODA: STYLOMMATOPHORA: HELICOIDEA) Min Wu Hebei University, Hezuolu 1, Baoding 071002, China; minwu@mail.hbu.edu.cn ABSTRACT Morphological variation in the terminal genitalia of genera of Bradybaeninae is compared and discussed. This is the first attempt to study the anatomy of the endemic Chinese bradybaenids Cathaica (Pliocathaica), Pseudiberus (Platypetasus), and Metodontia. A pre- liminary phylogenetic analysis of bradybaenids was performed based on the character matrix from the present study. The focus was primarily on the terminal genitalia. Helix (Helicidae) and Camaena (Camaenidae) were used as outgroups. The results suggest that several previous taxonomic arrangements for the subdivision of this family, based on the analyses using shell features and/or superficial anatomy of genital system, are unsuitable. The cladistic analysis suggests that the use of the subfamily Helicostylinae, sensu lato, might not be suitable for use as the sister group of the known Bradybaeninae. Two new endemic genera from western China are described based on the comparison of the terminal genitalia: Aegistohadra п. gen. and Eueuhadra п. gen. They are monophyl- etic and are readily distinguished from other bradybaenids by a synapomorphy, the pres- ence of penial caecum. Nanina delavayana Heude, 1885, is designated as the type species of Aegistohadra. The type species of Eueuhadra 1$ a new species, E. gonggashanensis. Key words: Stylommatophora, Helicoidea, Bradybaenidae, China, terminal genitalia, phylogeny, phylogenetic analysis, new taxa. INTRODUCTION The Bradybaenidae (= Bradybaenidae + Helicostylidae, sensu Schileyko 1991) are a large group of terrestrial snails widely distrib- uted in eastern Asia, with one species in Eu- rope. Historically, more than 150 authors (Richardson, 1983; Wu, unpublished cata- logue) have published on Chinese bradybaenids. However, most work on the classification of higher taxa of China was based on shell, not anatomical characters (Pilsbry, 1888-1894; Möllendorff, 1899; Dautzenberg, 1914-1915; Bavay & Dautzenberg, 1900, 1915; Blume, 1925; Ping 8 Yen, 1932; Yen, 1939; Zilch, 1940; almost all previous work). Therefore, knowledge on the bradybaenid systematics has remained unsatisfactory. The monograph by Wiegmann (1900), in which species from 12 genera and subgenera are described, was the first study dealing spe- cifically with the anatomy of bradybaenid geni- talia. More recently, some malacologists have made comparative studies of the genital mor- 79 phology, mainly based on their native bradybaenid taxa (Schileyko, 1978; Azuma, 1982; literature of Japanese workers, cited by Nordsieck, 2002; Lee & Kwon, 1993, 1994; Wu, 2001; Wu & Guo, 2003). Many authors have focused on the general structures, such as the size of dart sac, the presence/absence of a flagellum, and the number of mucous glands. Schileyko (1978) gave a much more precise, detailed description of the terminal genitalia of Russian bradybaenids that in- cludes the above traditionally described char- acter and internal dissections of the penis and dart apparatus. More recently, Nordsieck (1987) stated that the bradybaenid groups are characterized by apomorphies of the genital organs. However, similar work covering most en- demic Chinese bradybaenid taxa, which is essential for understanding the general anatomy of bradybaenids and construction of a sound taxonomic framework, has been lack- ing. The present work compares the structure of the terminal genitalia of some genera of the 80 Bradybaenidae based on dissection of their type species or non-type congeners. Two new bradybaenid genera are proposed based on anatomical and shell characters. А preliminary phylogenetic analysis is performed based on the data obtained from these dissection re- sults. This phylogeny is compared to the three bradybaenid subdivision plans comprehen- sively reviewed recently by Nordsieck (2002), widely used thus far in China (e.g., Yen, 1939; Zilch, 1960), Russia (e.g., Schileyko, 1978), and Japan (e.g., Kuroda & Habe, 1949; Minato, 1988). MATERIALS AND METHODS This study is based on specimens from the collections of the Zoological Museum, Insti- tute of Zoology, Chinese Academy of Sciences (IZCAS), and from those belonging to Forschungsinstitut und Naturmuseum Senckenberg (SMF). Many genera are repre- sented by the non-type congeners rather than by the type species, because of the paucity of alcohol-preserved specimens in museums and the absence of specimens from type localities. All examined specimens (except specimens of IZCAS00067, which were first fixed in for- malin before being placed in 70% ethanol) are preserved in 70% ethanol. For preparing the dissections, a tiny hole was carefully drilled into the shell apex to assist removal of the soft parts of the snail using water pressure. All the illustrations were drawn using a stereo micro- scope and camera lucida. Shell and genital measurements were taken with 0.01 mm and 0.1 mm accuracy respectively for the new taxa described. Whorl number was counted as de- scribed by Kerney 8 Cameron (1979) and was taken with 1/8 whorl accuracy. Both color and length of soft parts in the descriptions refer to those observed and measured after alcohol preservation. Type specimens of the new spe- cies are deposited in IZCAS, Beijing. Таха studied are listed in Appendix | along with locality data and museum accession num- bers. Descriptions of new taxa are given in Appendix Il. Abbreviations The abbreviations used in the text and in the illustrations are explained as: ADC — channel connecting accessory sac and dart sac; AG — albumen gland; App — vaginal empty appen- dicula; AS — accessory sac (= inner stylophore in Giusti et al., 1992); ASC — accessory sac chamber; At — atrium; BC — bursa copulatrix; BCD - bursa copulatrix duct; C23 — chamber produced by V2 and V3 in dart sac; DS — dart sac (= outer stylophore in Giusti et al., 1992); Dt — love dart; DtC — chamber containing the dart = dart sac chamber; DVM — membranous sac surrounding dart sac and/or distal region of vagina near atrium (= basal genital sheath in Cuezzo, 1998). When preparing the genita- lia for observation, the structures were care- fully preserved for future examination. Ep — epiphallus, the region between the pe- nis and the insertion of the vas deferens. The delimitation is esily recognized when the ephiphallic papilla (= verge in Cuezzo, 1998: 102) is present. When the epiphallic papilla is lacking, the continuous ridge structure can help to distinguish the epiphallic region from that of penis (Cuezzo, 1998). It is notable that the concept used by Cuezzo (1998) differs from that used by Giusti et al. (1992), who defined the epiphallus as “from end of vas deferens to point of attachment of penial re- tractor”. The term epiphallus of Cuezzo (1998) is used here, because the point of attachment of penial retractor varies among different bradybaenid groups, and in most cases it is not level with the ephiphallic papilla. EpP - epiphallic papilla (= “penial” verge in Schileyko, 1991); MAC — mucous gland-acces- sory sac channel; MG — mucous gland (= dart gland in Nordsieck, 2002); OD — oviduct; Ov — ovotestis; Р — penis; according to the epiphallus concept used by Cuezzo (1998), the term pe- nis used in this study refers to the region be- tween the epiphallic papilla and the atrium, or when the epiphallic papilla is absent, it refers to the region close to the atrium and internally possesses the similar and continuous pilaster/ ridge structure. PLs — polylayered structure in dart sac and/ or accessory sac, produced by wavy and spongy connective tissue. PLs is not separate, but connected tightly with neighboring tissue, and if present, is visible when the dart appa- ratus is dissected sagittally. This structure oc- curs as occupying most part of dart sac (e.g., in Fig. 14C) or a small and limited region (e.g., in Fig. 11B) in the dart apparatus; PR — penial retractor muscle; PS — penial sheath; PP — penial pilaster(s)/ridge(s); SPC — simple pe- nial caecum; T — talon; UV — free oviduct; Va — vagina; VD — vas deferens; V1 — a valvule op- posite the opening of mucous glands, in sag- TABLE 1. Data set containing 28 characters (characters 0-27) and 26 taxa used т the phylogenetic analysis. 1021177127137 1415 16 7 18 19 2012122125: 24_25 26127 (ALS Е бет 59 9 002020250 0 ZO! NC LAZO: 2 2 m2 2 252 92760 1 1 20 0 1 1 1 0820720700 Camaena Albers, 1850 Helix Linnaeus, 1758 Oe) OW 2 OO 2 WO 0 о 1 2 0 1 1 090 0-0 0) ооо © 1 1 Mastigeulota Pilsbry, 1895 PHYLOGENY OF BRADYBAENIDAE ооо ооо ооо ооо ооо оо ооо о ооо ото ооо ооо оо ооо ооо ооо ооо ооо ооо ооо оо о о ооо ооо ооо ооо о ооо ооо отт ооо оо ооо Or KT ооо ооо оо оо NNNNOOONNNOOOONO oocoooooooooooowN oorcoooooooooooonx< Se) OO e Ser? оо OO OO OO m en ON Sr le) nsoorcorcomnmntToooooo ee ehe) in e (©) (=) (©) (>) (©) OO 00060 ss © © oO 0 0 = ооо ооо оо о ооо ооо ооо о о о о ооо OONONONNNNOOONON SONO SS ee (=) (>) (>) OL (S) ONS (SO) SS BS (> (©) OCT OTK мот оо ооо от KT NOOO > < охтомямтоооо ооо ооо омотт мо оо оо о ооо охот ооо ооо = oO O © (o>) O 00 5 = со 0 So © т ES N N © 60 © © LO) E 0 Oo — 1) Es = © © © = - Oo 21S Зоо 0NQO— EQU Ее = Goo -2D6 D C= £& DO" O AS D © © © © = ant oO _ боб E=002:0 0% = n © = [er (ol 5 rn Oo” oO — (= - © фо= O So sma2 ооо о += Обо < < Ф ооо бо обес Oso 235259 00530 DD Oo 3< ZS3>06++0D9D20522000 Фо оо о Фобос 5цхаяхоа часа я чан 24010 0100 0009 PI ARA 1 1 ооо 1 1 020720 1 1 ZOO E AO) 1 1 1 1 1 1 1 1 0 1 00 Euhadra Pilsbry, 1890 00 1 Nesiohelix Kuroda & Emura, 1943 0 0 0 0 0 Aegistohadra, п. gen. 0 1 0020700 1 1 1 1 1 0207207070 007500510 1 0207207205 022020 ооо 0.0 0 Eueuhadra, п. gen. 1 1 0 0 0502400009 022 05200 1 1 1 1 1 1 1 1 Calocochlea Hartmann, 1842 Pfeifferia Gray, 1853 1 00277007 00 2020,20 ll VA оо ООО 0 1 Trichobradybaena Wu, 2003 81 82 Ща! plane of dart sac (as т Fig. 6D); V2 — а valvule opposite V1 and closest to atrium, to- gether with V1 forming a muscular tube con- taining dart(s) (as in Fig. 60); V3 — a valvule between V2 and V4, in sagittal plane of дай sac (as in Fig. 60); V4 — most inner/proximal valvule in DC, together with V1 forming a chamber containing love dart(s) (as in Fig. 6D). Terms V1-V4 are employed, for convenience only, to show the sagittal plane of dart sac. PR FIG. 1. Fruticicola fruticum (O. Е. Müller, 1774), IZCAS01009-2. A, general view of genitalia; В, dart sac and part of vagina, sagittal section, with cross-section of accessory sac. Structured DVM indicated by a thick solid arrow; C, penis, opened, with cross-section, showing a fold formed by the penial pilasters. A & В showing the elongated vagina section between dart sac and atrium. Bars equal 1 mm. PHYLOGENY OF BRADYBAENIDAE 83 Each view of the three dimensional portrayal represents only one part of the boundary of the chamber near it. Cladistic Analysis Cladistic analyses were performed using the computer program Hennig86 Version 1.5 (Farris, 1988) and program Winclada Version 1.00.08 (Nixon, 2002). The analysis of the character distribution on the cladograms was carried out using the program Winclada. All the 28 characters used, observed from terminal genitalia except Character 27 from mantle, are based on a selection made after my study of the representatives for terminal groups. Of the characaters (0-27), seven bi- nary characters and the remaining multistate characters were coded as non-additive. To avoid artificial judgement, character polarity is obtained as one of the results of the analysis rather than as an apriori assumption (Nixon 8 Carpenter, 1993). Therefore, all characters involved are treated as undirected and unor- dered. No missing character state occurred in the examined terminals (Table 1). Consider- ing that fused coding involves a loss of phylo- genetic information (Lee & Bryant, 1999), the inapplicable characters (e.g., coding of char- acter 1 was separated from that fused with character 0) were separately coded when the character-variable 1$ inapplicable in some taxa. Selection of the Ingroup and Outgroup Taxa Besides including two newly proposed gen- era, the ingroup bradybaenid taxa considered were those included in the subfamily Bradybaeninae by Richardson (1983), except Armandiella Ancey, Tricheulota Pilsbry, Plecteulota Môllendorff, Neseulota Ehrmann, Archaeoxesta Kobelt, Coccoglypta Pilsbry, Coneulota Pfeffer, Dolicheulota Pilsbry, and Ponsadenia Schileyko, because alcohol-pre- DVM FIG. 2. Bradybaena similaris (Rang, 1831), IZCAS01072-1. A, lateral view of дай apparatus and penial complex; B, basal view of dart apparatus; C, penis, opened; D, dart sac, sagittal section. DVM indicated by thick solid arrows in A & D. Bar equals 1 mm. 84 served material was unavailable. Semi- buliminus Möllendorff was excluded because it was recently grouped into Metodontia (Wu, in review). Halolimnohelix Germain, Haplohelix Pilsbry, Urguessella Preston, and Vicarihelix Pilsbry listed in Richardson’s bradybaenid catalog (1983; also in Thiele, 1931) were ex- cluded because they are, on the basis of anatomy, non-bradybaenid helicoids (Nordsieck, 1986, 1987; Schileyko, 1991). Results of previous cladistic analyses for Helicoidea were used as the departure point for outgroup selection. According to the cla- dogram based on a molecular database (Wade et al., 2001: fig. 3c), the Camaenidae— Helicidae—Polygridae group forms the sister group of Bradybaenidae. In another anatomy- based cladistic analysis of Xanthonychidae (= Helminthoglypidae; Cuezzo, 1998), the sister relationship of the Bradybaenidae and Xanthonychidae-Helicidae groups are sup- ported by four synapomorphies. Therefore, in this study, the Helicidae and Camaenidae were chosen as outgroups. All the 23 genera and subgenera were treated as separated terminal taxa. The type species was available for only 11 ingroup and one outgroup genera. These are: Acusta, FIG. 3. Karaftohelix weyrichii (Schrenck, 1867), IZCAS01080-2. A, В, lateral views of dart sac, DVM indicated by thick solid arrows; C, dart sac, sagittal section, with cross-section showing DVM, DVM indicated by thick solid arrows, neck-structure indicated by thick hollow arrows; D, penis, opened, with cross-sections, fold formed by the penial pilasters indicated by thick solid arrows. Bar equals 1 mm. PHYLOGENY OF BRADYBAENIDAE 85 Bradybaena, Cathaica, Fruticicola, Mastigeu- lota, Pseudaspasita, Nesiohelix, Aegistohadra п. gen., Eueuhadra n. gen., Trichobradybaena, Pfeifferia, and Helix. Otherwise, only those spe- cies commonly accepted in a group were used as the representatives for their generic group. RESULTS Character Descriptions Character 0: Presence of the membranate sac surrounding the dart sac and/or the distal region of the vagina near to the atrium (DVM). (0) absent; (1) present (Figs. 1B, 2A, 2D, 9A, indicated by thick solid arrows). Remarks: The dart sac 15 inserted on the va- gina. In very few cases, the dart sac is ba- sally wrapped by a layer of membrane, which sometimes appears to be sac-like (Fig. 3A— C, indicated by thick solid arrows) near the atrium, completely or partially. Character 1: The DVM inernally simple or structured: (0) not applicable because DVM absent; (1) DVM present, internally simple (Figs. 2A, 2D, 9A, indicated by thick solid arrows); (2) DVM present, internally structured, with numerous cells (Figs. 1B, 3C, indicated by thick solid arrows). Character 2: The proximal dart sac and/or the distal region of the vagina are/is wholy en- circled by the DVM or not: (0) not applicable because DVM absent; (1) DVM present, proximal dart sac partially encircled by DVM (Figs. 1B, 3B, C, indicated by thick solid arrows); (2) DVM present, proximal dart sac wholly wrapped by DVM (Fig. 2A, indicated by thick solid arrows). Character 3: Presence of the penial sheath: (0) absent; (1) present. Remarks: п Nesiohelix swinhoei, Aegisto- hadra n. gen., Eueuhadra n. gen., Pfeifferia micans, and Calocochlea coccomelos, the penial sheath is lacking (Figs. 5D, 6A, С, 7A, B, 8A, B, 9A, D). In the other genera, the penial sheath 1$ always present (e.g., Azuma, 1982). In the outgroup Helix pomatia, the penial sheath is present and developed, wrap- ping the whole penis and the basal part of penial retractor (Fig. 10D, E, indicated by thick solid arrows). In bradybaenid genera, the penial sheath, if present, cannot be morpho- logically distinguished from that of Helix. Character 4: Differentiation status of the pe- nial pilasters: (0) penial pilasters not differentiated; (1) pe- nial pilasters differentiated near epiphallus; (2) penial pilasters differentiated near atrium. Remarks: Differentiated penial pilasters are those thickened, deep, and/or morphologi- cally distinguishable from the neighboring Zig-zag ones of moderate thickness. In most species examined, the penial pilasters are somewhat thickened near the epiphallus, becoming thinner near the atrium (e.g., Figs. 2C, 11C, 12C, 13C, 14E, 15D). It is charac- teristic that the pilasters on the penial inner wall differentiate towards the atrium or to- FIG. 4. Metodontia yantaiensis (Crosse & Debeaux, 1863), IZCAS00131-1. A, general view of genitalia, with cross-sections of penial sheath and penis; B, dart sac, sagittal section. Bar equals 1 mm. 86 wards the epiphallus. In Fruticicola fruticum and Karaftohelix weyrichii, the penial pilas- ters differentiate near the epiphallus and form an asymmetrically projecting fold (Figs. 1C, 3D), which is similar to the asymmetri- cal epiphallic papilla of Nesiohelix swinhoei (Fig. 5E), and assumed to serve as the epiphallic papilla. In Stilonodiscus moellen- dorffi (Fig. 16B), S. yeni, S. entochilus, and Laeocathaica (Laeocathaica) subsimilis (Fig. 17В), the penial pilasters become thickened and differentiated near the atrium. Especially FIG. 5. Nesiohelix swinhoei (L. Pfeiffer, 1865), IZCAS00055-2. A, dart sac, sagittal section; В, sagittal section of accessory sac; C, cross section of mucous glands insertion on accessory sac, mucous tube entrance indicated by a thick solid arrow; D, penis and epiphallus, sagittal section, with cross-section of epiphallus; E, sagittal section of penis-epiphallus region, diagrammatic, with cross-section; F, epiphallus and flagellum, opened, with cross-sections; G, cross-section of dart sac, showing two pieces of dart; H, a piece of dart, with cross-sections. Bars equal 1 mm. PHYLOGENY OF BRADYBAENIDAE 87 in Stilonodiscus, such differentiated pilasters Character 6: Symmetry of the epiphallic pa- are high and valvule-shaped. pilla (EpP): Character 5: Presence of the epiphallic papilla: (0) not applicable because epiphallic papilla (0) epiphallic papilla absent (e.g., Fig. 11C); absent; (1) epiphallic papilla present and (1) present (e.g., Fig. 12C). symmetric (е:9., 710$. 12С, 18): (2) FIG. 6. Aegistohadra delavayana п. gen. & comb., IZCAS00132-3. A, general view of terminal genitalia, the sac of vagina opposite to dart sac indicated by a thick solid arrow (left), penial caecum indicated by a thick solid arrow (right); B, basal view of dart sac, diagrammatic; C, penial complex, opened, with cross-sections, penial caecum indicated by a thick solid arrow; D, dart sac, sagittal section, the sac of vagina opposite to dart sac indicated by a thick solid arrow. Bar equals 1 mm. 88 WU PR Ep EoP FI P FIG. 7. Eueuhadra gonggashanensis, п. gen. & sp., IZCAS00067-13, Paratype. А, general view of terminal genitalia; В, penial complex, penis and partially epiphallus opened, with cross-sections; C, epiphallus and flagellum, opened, with cross-sections; D, dart sac, sagittal section; E, section of penis- epiphallus region, diagrammatic. Bar equals 1 mm. PHYLOGENY ОЕ BRADYBAENIDAE 89 epiphallic papilla present and asymmetric (Figs. 5E, 8D, 9D). Remarks: In Nesiohelix swinhoei, Aegisto- hadra п. gen., Eueuhadra п. gen., Pfeifferia micans, Calocochlea coccomelos, Cathaica (Pliocathaica) gansuica, Aegista (Aegista) accrescens, Aegista (Plectotropis) gerlachi, A SÍ ji NI ZA Laeocathaica (Laeocathaica) subsimilis, Acusta ravida, Trishoplita dacostae, and Euhadra herklotsi (Figs. 5D, 6C, 7B, 8D, 9D, 12C, 13F, 14E, 17B, 18D, 19D, 20C), amore or less protruding epiphallic papilla is present. In the remaining bradybaenid genera the epiphallic papilla is depressed or missing. FIG. 8. Calocochlea coccomelos (Sowerby, 1840), SMF 323619. A, general view of genitalia; B, dart sac, sagittal section; C, mucous glands, sagittal section; D, above: Penis-epiphallus region, opened; below: cross-section of penis-epiphallus transition, diagrammatic; valve-shaped epiphallic papilla indicated by thick solid arrows. Bars equal 1 mm. 90 WU Character 7: The epiphallic papilla valve- shaped or papilla-shaped: (0) not applicable because epiphallic papilla absent; (1) epiphallic papilla present, valve- shaped (Figs. 8D & 9D, indicated by thick solid arrows); (2) epiphallic papilla present, papilla-shaped (e.g., Figs. 5D, 10D). Character 8: Presence of the penial caecum: (0) absent; (1) present (e.g., Figs. 6A, 6C, indicated by thick solid arrows). Remarks: This structure can be easily dis- tinguished from the following simple penial caecum (SPC) by the PC pilasters, which are differentiated from those of the caecum. In the simple penial caecum (SPC), which characterizes the genera Trichobradybaena and Mastigeulota, the penial pilasters form- ing the inner wall of caecum are just the ex- tended parts from its outer/entering pilasters. Character 9: The simple penial caecum: (0) absent; (1) present (Figs. 11B, 23A, indi- cated by thick solid arrows). Character 10: Presence of the flagellum: (0) absent; (1) present. FIG. 9. Pfeifferia micans Pfeiffer, 1845, SMF 323620. A, general view of genitalia; B, dart sac, sagittal section except mucous glands; C, mucous glands, sagittal section; D, penis-epiphallus region, opened, valve-shaped epiphallic papilla indicated by a thick solid arrow. Bars equal 1 mm. PHYLOGENY OF BRADYBAENIDAE 91 Ep MG FIG. 10. Helix pomatia (Linné, 1758), IZCAS00188-1. A, general view of terminal genitalia, bar equals 2 mm; B, dart sac, sagittal section with mucous glands removed; C, section of partial dart sac, with cross-section; D, penial complex, opened, with cross-sections, middle two thick solid arrows indicating penis-epiphallus chambers; E, section of penis-epiphallus region, diagrammatic, penial sheath indicated by a thick solid arrow; F, distal penis near atrium opening, opened; G, basal view of dart sac, diagrammatic. 92 Remarks: The flagellum and the vas defer- ens insertion structure are almost the same in the species examined. Flagellum, if present, with inner ridges simple or somewhat complexly arranged. Insertion of vas defer- ens on flagellum inwardly forms a more or less distinct C-shaped (in cross-section) fold towards the tip of flagellum (Figs. 5F, 7C, 10D, 130, 14F, 19E, 20A). The only exception 1$ Aegistohadra delavayana, n. comb., in which a depressed pilaster instead of the distinct C-shaped fold is present (Fig. 6C). These structures are the same in bradybaenid gen- era and in Helix pomatia. Therefore, if present, the flagellum of the various groups examined might be considered homologous. Character 11: Presence of the polylayered structure (PLs) in accessory sac: (0) PLs absent (e.g., Figs. 1B, 5A, 8B); (1) PLs present (e.g., Figs. 2D, 10C, 11B, 13C, 14C, 22D); (2) not applicable because dart sac absent. Remarks: In Metodontia yantaiensis, the ac- cessory sac has some wavy and spongy connective tissue (polylayered structure, PLs) (Fig. 4B). In Pseudaspasita binodata, such structure seems to be weakly devel- oped (Fig. 21C). This kind of structure can be easily distinguished from the folds/pilas- ters on the inner wall of the accessory sac (Fig. 16E) by the compactness and paral- lelism in the arrangement of its filaments/ layers. In Bradybaena similaris and Cathaica (Cathaica) fasciola, the structure is much developed and situated between insertion of mucous glands and vagina (Figs. 2D, 22D). In Aegista (Aegista) it is highly developed and uppermost, and it wraps the dart chamber (DtC) (Figs. 13C, 14C). In Aegista (Plectotropis), PLs occupies the whole accessory sac that is externally vis- ible and the region between dart sac and the vagina. Interestingly, in Helix pomatia, the polylayered structure is also present, at the pit formed by both dart the sac and each of the accessory sacs/trunk of basal mucous stalks (Fig. 10C). The observed PLs of the taxa studied are provisionally assumed to v2 V3 V4 Pls DIC FIG. 11. Mastigeulota kiangsinensis (E. Martens, 1875), IZCAS00003-1. A, lateral view of dart apparatus; B, dart sac, sagittal section; C, penis, opened, simple penial caecum (SPC) indicated by a thick solid arrow. Bars equal 1 mm. PHYLOGENY OF BRADYBAENIDAE 93 be homologous in origin, because they oc- and vagina (region |) (e.g., Fig. 22D); (2) PLs cur only in the specific region in dart appa- present, distributed between mucous glands ratus, and assumed to be related to dart insertion and дай caecum (DtC; region II) shooting or pumping the mucus out during (e.g., Figs. 4B, 10C, 11B); (3) PLs present copulation. at region | & Il (e.g., Fig. 14С). Character 12: Distribution of polylayered struc- Character 13: The common entrance of mu- ture (PLs) in accessory sac: cous glands: (0) not applicable because PLs absent; (1) (0) mucous glands without common entrance distributed between mucous glands insertion (e.g., Figs. 19B, 20D); (1) with common en- VD Va FIG. 12. Cathaica (Pliocathaica) gansuica (Möllendorff, 1899), IZCAS00210-1. A, basal view of dart sac and penial complex; B, lateral view of dart sac and penial complex; C, penis, opened; D, dart sac, sagittal section, neck-structure region of dart sac indicated by four thick hollow arrows. Bar equals 1 mm. 94 trance; (2) not applicable because dart sac absent. Remarks: There are two ways by which the mucous glands are inserted on the acces- sory sac, which can only be observed when the accessory sac is cut open sagittally. Usu- ally, the mucous glands open into the ac- cessory sac through a common duct (Figs. 18; 29-48160, 7D, IB 12D) MSC MAC, 15€, 166.176, 18B..216; 220, 23E: 24B): Another situation was found in Karaftohelix weyrichii, Trishoplita dacostae, Euhadra DS herklotsi, and Nesiohelix swinhoei (Figs. ЗС, 5C, 19B, 20D), with two to numerous sepa- rate tubes rather than a common tube open- ing into the accessory sac. Character 14: The distinguishability of the ac- cessory sac from outside of the dart sac: (0) indistinct from outside of the dart sac; (1) distinct from outside of the dart sac; (2) not applicable because dart sac absent. Remarks: The accessory sac cannot always be distinguished externally by an apparent external boundary from the dart sac (e.g., Fig. —— MG — Va AS At Va B FIG. 13. Aegista (Aegista) accrescens (Heude, 1882), IZCAS00027-4. A, B, lateral views of dart sac; C, dart sac, sagittal section; D, flagellum, opened, with cross-sections; E, penial complex, penis opened, with cross-sections; F, section of penis-epiphallus region, diagrammatic. Bar equals 1 mm. PHYLOGENY ОЕ BRADYBAENIDAE 95 Р$ Va PLs OGC an FIG. 14. Aegista (Plectotropis) gerlachi (E. Martens, 1881), IZCAS00044-2. A, basal view of dart sac; B, lateral view of dart sac, outer tissue partially removed to show the polylayered structure inside dart sac; C, dart sac, sagittal section; D, polylayered structure in accessory sac, magnified; E, penis and epiphallus, opened; F, penial complex, with cross-sections. Bar equals 1 mm. 96 WU 2B). Various genera show different patterns of the accessory sac, which has little relation- ship with its size from external view. The ac- cessory sac is situated usually on the bottom of dart sac, except in Acusta, where it is situ- ated near the top of dart sac (Fig. 18B). The structurally simplest accessory sac is an empty sac, only with a few depressed folds (= pilas- ters) on its inner wall (e.g., Figs. 1B, 7D 8 16С). Character 15: The accessory sac 1$ bipartite (e.g., Figs. 14A, 22C) or undivided (e.g., Figs. 5A, 7A): (0) accessory sac divided into two parts; (1) accessory sac undivided; (2) not applicable because dart sac absent. Character 16: Presence of V1-V4 in the dart apparatus (= V2 is present): (0) V1-VA indistinct (= V2 is indistinct/ab- sent) (e.g., Figs. 8B, 9B); (1) V1-V4 distinct (= V2 is distinct) (e.g., Figs. 7D, 23E); (2) not applicable because dart sac absent. Remarks: Inside the dart sac, several valvules (V1-V4) form a tube that contains one love dart (or two in Nesiohelix) serving as mating-related organ (e.g., Figs. 7D, 23E). According to this study, the position and the number of the valvules are intraspecifically stable but vary among the genera studied. The term valvule is used here for the first time in land snail anatomy. It is a small valve- like structure that describes the nature of the chamber boundry (dart sac, accessory sac chanmber), visible in sagittal section. How- ever, the position of VI-V4 can easily be determind even when V2 is absent, because: (1) V1 and V4 always form the opening of a muscular tube containing the love dart(s); (2) the space between V4 and V3 is usually the opening of the accessory sac (the only ex- ception is in Acusta, in which the accessory sac is situated on the top of dart sac); (3) the space between V3 and V2 is C23, which varies from presence as a pronounced chamber to totally absence. Such absence means V2 is lacking morphologically. For this reason, the complexity of the development FIG. 15. Pseudobuliminus (Pseudobuliminus) piligerus (Möllendorff, 1899), IZCAS00085-21. A, В lateral views of terminal genitalia; C, dart sac, sagittal section; D, penial complex, penis opened, with cross-sections. Bar equals 1 mm. PHYLOGENY OF BRADYBAENIDAE 97 DS FIG. 16. Stilpnodiscus. À, В, С Stilpnodiscus moellendorffi Wu, 2001, IZCAS00081 -4, Paratype. A, lateral view of terminal genitalia; В, penis, opened, with cross-sections: С, dart sac, sagittal section; О, E Stilpnodiscus entochilus Möllendorff, 1899, IZCAS00076-2. D, basal view of dart sac; E, dart sac, sagittal section, in detail, with cross-sections of penis. Bar equals 1 mm. 98 of the dart sac inner structure is considered, described, and employed for the first time as an important and necesssary character for the dart sac in the Bradybaenidae. Character 17: Presence of a papilla within accessory sac formed by the mucous glands insertion: (0) without papilla; (1) with a papilla; (2) not applicable because dart sac absent. Remarks: А papilla with a tiny роге or sev- eral tiny pores for the entrance of mucous from mucous glands into the accessory sac is sometimes present. If the mucous gland ducts merge into one common tube, the pa- pilla also has one pore, as in Acusta ravida (Fig. 18B, G). When the mucous glands en- ter the accessory sac separately, two papil- lae are present, as in Trishoplita dacostae (Fig. 19B, indicated by two lower thick solid arrows) or a somewhat complex structure with numerous pores as in Nesiohelix swinhoei and Euhadra herklotsi (Figs. 5C, 20D). In most genera, such a structure is absent (other Figs.). Character 18: Presence of the structure de- rived from mucous glands entering papilla leading to DtC: (0) not applicable because mucous glands entrance papilla absent; (1) mucous glands entrance papilla present, its derived part does not lead to DtC; (2) mucous glands entrance papilla present, its derived part leads to DtC (Figs. 19B, 20D, respectively indicated by a upper thick solid arrow). Character 19: Number of branches of mucous glands: (0) numerous mucous branches; (1) one spherical mucous gland (Figs. 8, 9); (2) two branches of mucous glands; (3) not appli- cable because dart sac absent. С FIG. 17. Laeocathaica (Laeocathaica) filippina (Heude, 1882), IZCASO0006. A, В IZCAS00006-5. A, lateral view of terminal genitalia; В, penial complex, penis opened; С, 17СА$00006-6, dart sac, sagittal section. Bar equals 1 mm. PHYLOGENY ОЕ BRADYBAENIDAE 99 Character 20: The length of vaginal region between dart sac and atrium: (0) region short; (1) region pronouncedly elongated (e.g., Figs. 9A, 18B); (3) not ap- plicable because dart sac absent. Character 21: Proximal part of dart sac elon- gated, forming a neck-structure. (0) neck-structure absent; (1) neck-structure present (Figs. 3A, 12D); (2) not applicable because dart sac absent. Character 22: Presence of penis-epiphallus chamber(s): (0) absent; (1) a simple chamber present (Fig. 7B, E); (2) more chambers present (Fig. ТОВ, Е) Remarks: The penis-epiphallus chamber oc- curs in the wall of penis-epiphallus junction. Dissection shows that there are three cases of differentiation. (1) № is solid (i.e., without any chamber within) between the epiphallic papilla and its wall. (2) There is only a simple chamber between the epiphallic papilla and its wall. (3) As seen in Helix pomatia (Helicidae) (Fig. 10D, E), more than one chamber is developed in this area, and some of them extend into the penial wall. All FIG. 18. Acusta ravida (Benson, 1842), IZCAS00944-2. A, general view of genitalia; B, dart sac and part of vagina, sagittal section, with cross-section of vagina; C, region near penial sheath; D, proximal region of penis, opened, showing epiphallic papilla; E, section of penis-epiphallus region, diagrammatic; F, distal region of penis, opened; G, section of accessory sac, papilla of entrance for mucous tubes indicated by a thick solid arrow, diagrammatic. Bar equals 1 mm. 100 WU FIG. 19. Trishoplita dacostae Gude, 1900. IZCAS00174-2. A, lateral view of dart apparatus; B, dart sac, sagittal section, upper thick solid arrow indicating the structure derived from the mucous glands entering papilla leading to dart chamber, two lower thick solid arrows indicating the mucous glands entering papilla; C, the mucous glands entering papillae and the derived structure; D, penial complex, with penis opened; E, epiphallus and flagellum, opened, with cross-sections. Bar equals 1 mm. PHYLOGENY OF BRADYBAENIDAE 101 FIG. 20. Euhadra herklotsi (E. Martens, 1861), IZCAS01076-1. A, epiphallus and flagellum, with cross- sections; B, section of penis-epiphallus region, diagrammatic; C, penis-epiphallus region, penis opened; D, dart sac, sagittal section, upper thick solid arrow indicating the structure derived from the mucous glands entering papilla leading to dart chamber, lower thick solid arrow indicating the structure of mucous glands entering papilla. Bar equals 1 mm. 102 WU > FIG. 21. Pseudaspasita binodata (Möllendorff, 1886), IZCAS01075-2. A, general view of terminal genitalia; B, basal view of dart sac, diagrammatic; C, dart sac, sagittal section. Bar equals 1 mm. © ERAS At = LA N Г. = Sr À С = N PS de MG Dt DtC PLs va v1 v2 FIG. 22. Cathaica (Cathaica) fasciola (Draparnaud, 1801), 1ZCAS01074-6. A, lateral view of dart sac; B, lateral view of dart sac, with cross-sections of penial sheath and penis; C, basal view of terminal genitalia; D, dart sac, sagittal section; E, polylayered structure in accessory sac. Bar equals 1 mm. PHYLOGENY ОЕ BRADYBAENIDAE 103 bradybaenid genera examined, except Eueuhadra, п. gen., fall into the first case. Character 23: Number of darts per dart sac: (0) dart sac containing 1 dart; (1) dart sac containing 2 darts (Fig. 5G, Н); (2) not ap- plicable because dart sac absent. Remarks: In Nesiohelix swinhoei (type spe- cies of the genus Nesiohelix; Richardson, 1983, the type species mistakenly given as Nesiohelix caspari; see original introduction of genus by Kuroda 8 Emura, 1943), the dart sac contains two darts, each of which is wrapped by a muscular tube. These two muscular tubes are attached closely but dis- tinctly divided. In this study, the two darts are the same length rather than “one larger, the other smaller” (Kuroda 8 Emura, 1943: text-fig. 1), semi-circled in cross-sections, and blunt apically. In some other congeneric species of Nesiohelix, such as N. samarangae (Kuroda 8 Miyanaga, 1942) and N. moreletiana (Heude, 1882), the dart sac contains two darts (Habe, 1945, not fig- ured), which is confirmed in this study and is an important synapomorphy characteriz- ing Nesiohelix. Character 24: Internal pilaster of accessory sac differentiated or not: (0) not differentiated; (1) differentiated (Fig. 1B); (2) not applicable because dart sac absent. Character 25: Position where the accessory sac is inserted on dart sac: (0) accessory sac inserted on the bottom of dart sac; (1) accessory sac on the upper side of dart sac (Fig. 18В); (2) not applicable be- cause dart sac absent. Remarks: The accessory sac 1$ usually situ- ated on the bottom of dart sac, except in Acusta, where it is situated at/near the top of dart sac (Fig. 18B, compared to its nor- mal position, e.g., that shown in Fig. 9A). The abnormal position of accessory sac can be observed in all anatomically known FIG. 23. Trichobradybaena submissa (Deshayes, 1873), IZCAS00010-3. A, general view of genitalia, simple penial caecum (SPC) indicated by a thick solid arrow; B, C, lateral views of dart sac; D, penis, partially opened; E, dart sac, sagittal section. Bars equal 1 mm (after Wu 8 Guo, 2003). 104 WU Acusta species and is preliminarily consid- by thick solid arrows); (2) not applicable be- ered as the inverse of accessory sac in po- cause dart sac absent. sition (Wu, unpublished paper on Acusta). Character 27: Relation of the mantle to the shell. Character 26: Presence of sacs inserted on (0) shell is not partially enclosed by mantle; vagina oppsite to dart sac: (1) the mantle partially enclosing shell (ob- (0) absent; (1) present (Fig. 6A, D, indicated served in Pfeifferia micans, SMF323620). FIG. 24. Pseudiberus (Platypetasus) chentingensis Yen, 1935, IZCAS00163. A, lateral view of terminal genitalia, with cross-sections of epiphallus and vas deferens; B, dart sac, sagittal section; C, penial complex, with penis opened. Bar equals 1 mm. PHYLOGENY ОЕ BRADYBAENIDAE 105 FIG. 25. Camaena platyodon (L. Pfeiffer, 1846), IZCAS00833. Male section of terminal gentalia, left, showing задана! section of penial wall; right, showing cross-sections of penis, epiphallus and flagellum. Cladistic Analysis The data matrix (Table 1) was submitted to HENNIG86 and Winclada. All the observed apomorphies were included among the char- acters because they are useful for the char- acterization of certain terminals, although they are not informative for the construction of the phylogenies. The removal of apomorphies in the analysis will decrease the steps in con- 106 structing a cladogram, but will not influence its reliability. Two types of analyses were per- formed. In the first type of analysis, all char- acters were weighted equally. Data sets were calculated with an exact algorithm (implicit enumeration). Another type of analysis used the successive weighting function provided by the Hennig86 program, which is considered by Carpenter (1988) to be the best method for weighting characters and choosing among equally parsimonious cladograms. The first analysis produced 11 equally parsimonious trees (EPTs) with length of 96, consistency index (Cl) 0.54, and retention index (RI) 0.67. Extended branch swapping was then applied to the initial tree using the branch-breaking WU (bb*) command, producing 3,502 bb trees. A strict consensus tree (SCT) (Fig. 26) was then summarized from these 3,502 trees with Winclada in order to find the most unambigu- ous monophylies. In the second analysis, after two iterations of successive approximations weighting and branch-breaking, 87 trees were retained, each with length 274, Cl = 0.73, RI = 0.87. The cla- dograms obtained by the first and the second type of analyses were then introduced to Winclada for rerooting and mapping the dis- tribution of characters. Based on the trees re- sulting from the second type of analysis, the rooted SCT was produced using Winclada (Fig. 27): Camaena Helix Acusta Karaftohelix Pliocathaica Laeocathaica Aegistohadra n. gen. Eueuhadra n. gen. Nesiohelix Trishoplita Euhadra Aegista Plectotropis Pseudaspasita Mastigeulota Metodontia Cathaica Platypetasus Stilpnodiscus spa Stilpnodiscus spb Pseudobuliminus Bradybaena Trichobradybaena Fruticicola Calocochlea Pfeifferia FIG. 26. Rooted SCT resulted from 3,502 EPTs (| = 96, Cl = 54, RI = 67) based on equally weighted characters. PHYLOGENY OF BRADYBAENIDAE In the 3,502 ЕРТ$ constructed based on the equally weighted characters, 51 trees were found to be exactly equal respectively, in to- pology, to those obtained by philosophy of successive approximations weighting. In other 3 11131415161718192021 23242526 Pseudobuliminus — _—_—————_—— В O Camaena 022222233222222 1112 | Helix 4 1017182025 O-0-0-0-0 Acusta 201111 3 1823 и 4 O-O Nesiohelix 131719 011 O-0-0 5 В 151622 010 18 Trishoplita 110 ; Euhadra 26 y 38 Aegistohadra n. gen. O 1 01 1922 о Eueuhadra п. gen. Aegista 111216 15 à ово о Plectotropis 130 0 12 Pseudaspasita 4 Laeocathaica 012 4 1316 21 [700000-0— Karaftohelix 121100 1 . ы (ois Pliocathaica O0 111216 } = o— Cathaica 4 Stilonodiscus spa 567 9 1112 . o-0-0— Mastigeulota 000 112 Platypetasus 14 Ho— Sf/pnodiscus spb 0 111216 , 0-00— Metodontia 120 19 O 111216 2 0-0-O 110 290 107 words, 51 out of 87 trees based on weighted characters had the exact topology with the cla- dograms from the first analysis. When the rooted SCT (Fig. 27) was summarized from the 87 trees from the second analysis, only R JN B—B—B- B—E—E- АА B—E—E- A—A—A- A—A—A- A- Bien B—B—B- ВВ Bradybaena 9 12 Trichobradybaena : oe Fruticicola B = H — Н = 27 11 Pfeifferia a H = Calocochlea FIG. 27. Left: Rooted SCT resulted from 87 EPTs (| = 274, CI = 73, RI = 87) based on weighted characters, showing the distribution of character states. Solid circle — nonhomoplasious change; empty circle - homoplasious change. Right: Showing suprageneric classification by different authors: A — Aegistinae/Aegistini, В — Bradybaeninae/Bradybaenini, Е — Euhadrinae/Euhadrini; H — Helicostylidae/Helicostylinae; R — Russian authors, J — Japanese authors, N — Nordsieck (Nordsieck, 2002). 108 one node collapsed, so this rooted SCT was thought to be informative and proper for being used both to interpret the present bradybaenid phylogeny and to indicate the reliability of the monophyletic groups. As shown in the rooted SCT obtained by us- ing the second type of analysis (Fig. 27), the ingroup was well defined by three nonhomo- plasious synapomorphies characters 15(1), 16(1), and 22(0) (Fig. 27). Eight clearly distin- guished monophylies supported by nonhomo- plasious synapomorphy/synapomorphies are as follows: (a) The clade (Trishoplita, Euhadra), was sup- ported by character state 18(2). The mono- phyly of this clade was also confirmed by the SCT; (b) The clade composed of Aegistohadra and Eueuhadra, supported by character state 8(1); (c) The monophyly (Aegista, Plectotropis, Pseudaspasita), supported by character state 12(3). The monophyly of this clade was also confirmed by the SCT; (d) The clade composed of Karaftohelix and Pliocathaica, supported by synapomorphic character state 21(1); (e) The clade composed of Cathaica, Stilonodiscus, Mastigeulota, Platypetasus, Metodontia, Pseudobuliminus, Brady- baena, Trichobradybaena, Fruticicola, Calocochlea, and Pfeifferia, supported by synapomorphic character states 5(0), 6(0) and 7(0); (f) The clade embedded in (e), Mastigeulota, Platypetasus, Stilpnodiscus spb, Metodontia, Pseudobuliminus, Brady- baena, Trichobradybaena, Fruticicola, Calocochlea, and Pfeifferia, supported by synapomorphic character state 14(0). The monophyly of this clade was also confirmed by the SCT; (g) The clade composed by Bradybaena, Trichobradybaena, Fruticicola, Calococh- lea, and Pfeifferia, supported by synapomorphic character states 1(1) and 2(2). The monophyly of this clade was also confirmed by the SCT; (п) The clade Calocochlea and Pfeifferia, sup- ported by synapomorphic character states 6(2), 7(1), and 19(1). The monophyly of this clade was also confirmed by the SCT. Based on the characters extracted from ter- minal genitalia, only part of the examined bradybaenid genera could be characterized by their autapomorphies. The terminal taxa Trishoplita, Euhadra, Aegista, Pliocathaica, Calocochlea, Pseudobuliminus, and Platy- petasus had the opposite situation, that is, considering the anatomy of terminal genitalia, they were not characterized by derived char- acters. As indicated in Figure 27, they seemed to be defined by the “loss of character state(s)” rather than autapomorphies that could be di- rectly observed. DISCUSSION The proposed phylogeny of bradybaenid genera has almost no similarity with the previ- ous systems reviewed by Nordsieck (2002). Previously, knowledge of bradybaenid system- atics came from the shell and very few genital features, and resulted from methodologically subjective analyses. The present cladograms, with too many branches, are not strongly sup- ported, indicating that this is a preliminary re- sult, providing a testable hypothesis of realtionships among bradybaenid genera. The hypothesis reflected by the cladogram in Fig- ure 27 is preferred, because it represents the best testable systematic hypothesis explain- ing the present data set. While the hypothesis presented is limited and requires the addition of data from many un- studied taxa, focus on some monophyletic branches with relatively strong support shows convincing results. The monophyletic clade of Calocochlea and Pfeifferia, representatives of the Helicostylinae Ihering, 1909, is well nested in the ingroup, suggesting that Helicostylinae are a bradybaenid group rather than a sepa- rate family (Helicostylidae sensu Schileyko, 1991). In the definition of the family Helicostylidae sensu Schileyko (Schileyko, 1991: 221), “the flagellum is variously devel- oped but is always present” is a dubious char- acter, because the present work shows that both Pfeifferia micans (type species of Pfeifferia) and Calocochlea coccomelos have no flagellum (Figs. 8, 9). The present phylogeny, by artificially exclud- ing helicostyline groups, is more or less com- patible with the tripartite plan of bradybaenid genera, that is, tripartitite classification of (1) modified Aegistini (= subfamily Aegistinae sensu Kuroda & Habe, 1949, listed as a tribe by Nordsieck, 2002; partial Aegistinae sensu Schileyko, 1991), (2) Euhadrini (= Euhadrinae Minato, 1988, listed as a tribe by Nordsieck, 2002: including both Nesiohelix and Euhadra), PHYLOGENY OF BRADYBAENIDAE 109 and (3) Bradybaenini (= Bradybaeninae sensu Kuroda & Habe, 1949, listed as a tribe by Nordsieck, 2002) (Fig. 27). The Aegistini was distinguished from Bradybaenini by the pres- ence of the flagellum. The present phyloge- netic hypothesis suggests the flagellum has been at least convergently lost in Brady- baenidae twice (Fig. 27). Accordingly, this character should not be employed as the proper character defining Aegistinae as used in the original designation (Kuroda & Habe, 1949). The present hypothesis shows the reli- ability for the monophyly of (Trishoplita, Euhadra), which are distributed in both Aegistini (including genus Tishoplita: Kuroda 8 Habe, 1949; Minato, 1988; Schileyko, 1991; Nordsieck, 2002) and Euhadrini. Therefore, Aegistini should be condisered a paraphyletic group as indicated by the evidence that Trishoplita is embedded in the clade of Euhadrini. As clearly indicated by the all ETPs (3,502 ETPs, not figured) from the first analysis and cladogramed based on the weighted charac- ters (in all 87 ETPs, not figured), Acusta oc- curred most basally in the cladograms. Also indicated by the rooted SCTs (Figs. 26, 27), Acusta, which was placed in the Bradybaenini (= Bradybaeninae sensu Russian and Japa- nese authors), was confirmed as the sister group to all the remaining bradybaenids examinated. Thus, the Bradybaeninae is a paraphyletic group, and Acusta should not be placed in Bradybaenini (sensu Nordsieck, 2002). In summary, the result obtained dem- onstrates the Bradybaenini (sensu Nordsieck, 2002) is not monophyletic. Cathaica was divided into several subgen- era by Andreae (1911) based on shell charac- ters. This study examined two of them, Cathaica (s. str.) and Cathaica (Pliocathaica). The results here show that Cathaica (s. str.) has a much closer relationship to the termi- nals in Clade (e), than to Pliocathaica, which is closest to Karaftohelix (Fig. 27). Accordingly, subgroups of Cathaica may be polyphyletic. Some characters used by other authors are thought to be unreliable after careful dissec- tions and thus are omitted from the present study. The widening of the basal bursa copulatrix duct, which was used by Schileyko (1991) as a diagnostic character of Brady- baenidae sensu Schileyko, is not included in the present data set, because this part varies in thickness according to physiological state, for example in Aegista accresens (Heude, 1882), as observed by the author. Some char- acters once used to describe the genitalia are ambiguous and thus should be avoid being used. For example, the development status of the accessory sac, which is an autapo- morphy in the diagnostic definition for Helico- stylidae (sensu Schileyko, 1991) as “ап accessory sac is weakly developed or lack- ing”, seems not to be so definite and conse- quently is less informative or misleading. In the Bradybaeninae (sensu Nordsieck, 2002), the accessory sac shows a variety of devel- opment states, such as size range, differen- tiation of the internal pilasters, and occurrence of the mucous glands entering papilla. There- fore, the accessory sac comprises a series of characters instead of a character with several character states. In Helicostylinae (= Helico- stylidae sensu Schileyko, 1991), the genera Calocochlea and Pfeifferia (Figs. 8B, 9B) have an accessory sac with similar structure as those seen in the bradybaenine, for example, in Trichobradybaena (Fig. 23E) and in Pliocathaica (Fig. 12D). This suggests some characters, such as seen in the non-homo- plasious characters (both synapomorphies and autapomorphies) in this study, should be given special attention as to whether they are shared by or transformed into certain states in any other bradybaenid genus not covered in this work. Careful consideration of this prob- lem will enhance the reliability of the phylog- eny obtained. Cuezzo (1998) points out that there are three different problems seen in the published lit- erature of the Xanthonychidae (= Helmintho- glyptidae). | see the same problems in the current the study of the Bradybaenidae. Virtu- ally in all the published literature, the system- atics of Bradybaenidae is established on “arbitrary narrative character transformations”. Any effort to make a predictive classification of the Bradybaenidae (or any other group), as Nordsieck (2002) suggests, should be based on testable hypotheses, and after as many species as possible are examined. The present work does not aim to provide a definitive clas- sification of the Bradybaenidae, as many gen- era and and many other important characters, for example, anatomical (besides terminal genitalia), molecular, and chromosomal, are not included in the data set. However, it does suggest that the phylogeny of the Brady- baenidae is complex and considerable further work on the systematics for this group 1$ needed. 110 ACKNOWLEDGEMENTS The author is in debt to Mr. Hartmut Nord- sieck and Dr. Ronald Janssen (Natural His- tory Museum of Senckenberg, Germany) for helpful opinions on this the manuscript and for lending important specimens. Prof. Folco Giusti (Dipartimento di Scienze Ambientali, Italy) and Prof. Anatoly А. Schileyko (А. М. Severtzov Institute of Problems of Evolution, Russia) reviewed the manuscript and provided valued and helpful comments. Dr. Larisa A. Prozorova (Russian Academy of Sciences) provided some critical specimens. Dr. Bernhard Hausdorf (Universität Hamburg, Germany) shared the author his experience in the construction of phylogenies. Prof. Edmund Gittenberger (National Museum of Natural History, Netherlands) has provided lit- erature. 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KWON, 1994, Studies on the genital organs of fourteen species of family Bradybaenidae in Korea. Korean Journal of Malacology, 10(1): 9-18. MINATO, H., 1988, A systematic and biblio- graphic list of the Japanese land snails. Shirahama. x + 294 pp. MOLLENDORFF, О. F., 1899, Binnen-Mollusken aus Westchina und Centralasien. |. Annuaire du Musée Zoologique de L’Academie Impériale des Sciences de St. — Pétersbourg, 4: 46-144, 7 pls. NIXON, K. C. & J. M. CARPENTER, 1993, On outgroups. Cladistics, 9: 413—426. NIXON, K. C., 2002, WinClada ver. 1.00.08. Published by the author, Ithaca, New York. NORDSIECK, H., 1986, The system of the Stylommatophora (Gastropoda), with special regard to the systematic position of the Clausiliidae, Il. Archiv für Molluskenkunde, 117 (1/3): 93-116. NORDSIECK, H., 1987, Revision des Systems der Helicoidea (Gastropoda: Stylommato- phora). Archiv für Molluskenkunde, 118: 9-50. NORDSIECK, H., 2002, The systematics of the Bradybaeninae (Gastropoda: Stylommato- phora: Bradybaenidae), an example for the PHYLOGENY ОЕ BRADYBAENIDAE da work of divergent systematic schools. Mittei- lungen der Deutschen Malakozoologischen Gesellschaft, 67: 41-47. PILSBRY, H. A., 1888-1894, In: С. W. TRYON & Н. A. PILSBRY, Manual of Conchology, (2)4: 121- 193 [1888]; (2)4: 194-296 [1889]; (2)5: 179- 216 [1889]; (2)6: 193-324 [1891]; (2)7: 129-225 [1892]; (2)8: 113-314; (2)9: 49-338 [1894]. PILSBRY, H. A., 1934, Zoological results of the DOLAN West China expedition of 1931. Part Il, molluscs. Proceedings of the Academy of Natu- ral Sciences of Philadelphia, 86: 5-28, 6 pls. PING, С & T.-C. YEN, 1932, Some gastropods from Sin-Kiang. The Science Quarterly of the National University of Peking, 3(3): 127-148. RICHARDSON, L., 1983, Bradybaenidae: cata- log of species. Tryonia, 12: 1-479. SCHILEYKO, A. A., 1978, Land mollusks of the superfamily Helicoidea. Fauna of the Ц. $. $. R., Mollusks, 3(6): 1-384. SCHILEYKO, A. A., 1991, Taxonomic status, phylogenetic relations and system of the Helicoidea sensu lato (Pulmonata). Archiv fúr Molluskenkunde, 4/6: 187-236. THIELE, J., 1931, Handbuch de systematischen Weichtierkunde. (translated by J. S. Bhatti, general editor: V. $. Kothekar) 2: 688-702. WADE, С. M., Р. В. MORDAN & В. CLARKE, 2001, А phylogeny of the land snails (Gas- tropoda: Pulmonata). Proceedings of the Royal Society of London, B, 268: 413-422. WIEGMANN, F., 1900, Binnen-Mollusken aus Westchina und Centralasien. Annuaire du Musée Zoologique de L'Academie Impériale des Sciences de St.-Pétersbourg, 5: 1-131. WIKTOR, A., D.-N. CHEN & М. WU, 2000, Stylommatophoran slugs of China (Gas- tropoda: Pulmonata) (A prodromus). Folia Malacologica, 8(1): 3-35. WU, M., 2001, Contribution to the knowledge of Chinese endemic genus Stilpnodiscus (Gastropoda: Pulmonata: Bradybaenidae), with descriptions of two new species. Folia Malacologica, 9(2): 81-89. WU, М. & J.-Y. GUO, 2003, Contribution to the knowledge of the Chinese terrestrial malaco- fauna (Helicoidea): description of a new bradybaenid genus with three species. The Veliger, 46(3): 239-251. YEN, Т. С., 1935, The non-marine gastropods of north China. Part | Publications du Musée Hoangho Paiho de Tien Tsin, 34: 1-57, 5 pls. YEN, T. C., 1939, Die chinesischen Land- und Süßwasser- Gastropoden des Natur-Museums Senckenberg. Abhandlungen der Senckenber- gischen Naturforschenden Gesellschaft, 444: 129-157, pls. 13-16. ZILCH, A., 1940, Landschnecken aus Fukien (China). Archiv für Molluskenkunde, 72: 113-118. ZILCH, A., 1959-1960, Gastropoda, Teil 2 Eu- thyneura. — Handb. Paláozool., 6 (2), ХИ + 834 pp. Berlin-Nikolassee (Borntraeger). Revised ms. accepted 11 January 2004 APPENDIX |: Taxa studied Acusta ravida (Benson, 1842), type species of the genus: IZCAS00944, Jiangning County, Jiangsu Province, coll. unknown. Four adult specimens (two dissected) and two young specimens. Aegista (Aegista) accrescens (Heude, 1882): IZCAS00027, Xiushan County (28.4°N, 108.9°E), Sichuan Province, 1986-VII-21, coll. De-Niu Chen 4 Jia-Xiang Gao. Six adult specimens (three dissected). Aegista (Plectotropis) gerlachi (E. Martens, 1881): IZCAS00044, Guangdong Province, other collection data unknown. Nine adult (two dissected) and three young specimens. Bradybaena similaris (Rang, 1831) (not A. Férussac 1821; see Nordsieck, 2002), type species of the genus: IZCAS01072, Fuzhou, Fujian Province, 1975-X-16, coll. unknown. Forty adult (two dissected) and 17 young specimens. Cathaica (Cathaica) fasciola (Draparnaud, 1801), type species of the genus: 12CAS01074, Pi County, Xuzhou, Jiangsu Province, 2000-V-2, coll. Qi-Lian Qin. Nu- merous specimens (three dissected). Cathaica (Pliocathaica) gansuica (Möllendorff, 1899): IZCAS00210, Dachuanxiang, Zhouqu County, Gansu Province, 1200 m alt., 1998- V-9, coll. De-niu Chen & Guo-Qing Zhang. 166 specimens (two dissected). Fruticicola fruticum (O. F. Müller, 1774), type species of the genus: IZCAS01009, lime- stone quarry near Klodzko, Wapniarka Mt., Lower Silesia, Poland, 1999-VI-26, coll. Min Wu 8 Andrzej Wiktor. Seven adult speci- mens (two dissected) and one young speci- men. Karaftohelix weyrichii (Schrenck, 1867): 1ZCAS01080, near Yushno-Sakhalinsk City, Sakhalin Island, Russia, 2001-VII-29, coll. Larisa A. Prozorova. One adult (dissected) and four young specimens. Mastigeulota kiangsinensis (E. Martens, 1875), type species of the genus: 12CAS00003, Huangnipo, Badong County (31.0°М, 110.3°E), Hubei Province, coll. De- Niu Chen, 1984-VI-29. Six adult (two dis- sected) and one young specimen. Metodontia yantaiensis (Crosse 8 Debeaux, 1863): IZCAS00131, Quyang County, Hubei Province, coll. Min Wu. Fifteen adult speci- mens (two dissected) and 19 young speci- mens. 112 Pseudaspasita binodata (Möllendorff, 1886), type species of the genus: IZCAS01075, Beiquan Park, Beipei, Chonggin, 1964-V-12. Twenty-seven adults (two dissected) and 12 young specimens. Pseudiberus (Platypetasus) chentingensis Yen, 1935: IZCAS00163, Jiaozuo, Henan Province, 1999-VII-22, coll. Guang-Wen Chen. Six adults (two dissected) and 17 young specimens. Stilonodiscus moellendorffi Wu, 2001: IZCAS00081, type specimens, Shanggou, Shawanxiang, Dangchang County, Gansu Province, 1998-VI-6, coll. De-Niu Chen & Guo-Qing Zhang. Stilonodiscus entochilus Móllendorff, 1899: IZCAS00076, Guoyuanxiang, Nanping County (now Jiuzhaigou County) (33.2°N, 104.2°E), Sichuan Province, 1000 m alt., coll. De-Niu Chen & Guo-Qing Zhang, 1998- V-18. 25 adult (four dissected) and 17 young specimens. Laeocathaica (Laeocathaica) subsimilis (Deshayes, 1873): 17СА$00006, Xingjian- xiang, Nanchong (30.8%N, 106.1°E), Sichuan Province, coll. unknown, 1964-V-20. Eleven adult specimens (four dissected). Pseudobuliminus (Pseudobuliminus) piligerus (Möllendorff, 1899): 17СА$00085, Anchang- hexiang, Wen County (33.0%N, 104.6°E), Gansu Province, 1200 alt., coll. De-Niu Chen & Guo-Qing Zhang, 1998-V-19. 287 specimens (three dissected). Trishoplita dacostae Gude, 1900: IZCAS00174, Kobayashi Hiyazaki, Japan, coll. unknown, 1998-X. Six adult (three dis- sected) and six young specimens. Euhadra herklotsi (E. Martens, 1861): IZCAS01076, Ishigakijima, 1931-VII, coll. Shikanu (?). Two adult specimens (one dis- sected). Nesiohelix swinhoel (L. Pfeiffer, 1865), type species of the genus (in Richardson, 1983, the type species mistakenly given as Nesiohelix сазрап; Kuroda & Emura, 1943): IZCAS00055, Yilan County (24.7°N, 121.7°E), Taiwan Province, coll. unknown, 1896-X. Two adult specimens dissected. Aegistohadra delavayana (Heude 1885) n. gen. and comb.: IZCAS00132, Zhibenshan Mt., Baoshan (26.3°N, 104.4°E), Yunnan Province, coll. De-Niu Chen, 1981-VI-23. Four adult specimens (two dissected) and three young specimens. Paratypes, IZCAS- type-2902-1 and IZCAS-type-2902-2, Fa Kouan Tchen, coll. Unknown. Aegistohadra seraphinica (Heude, 1889): paratypes, IZCAS-type-3071-1 and IZCAS- type-3071-2, Si-lin, Guangxi, coll. Unknown. Eueuhadra gonggashanensis, п. gen. € sp., type species of the genus: IZCAS00067, west slope of Gonggashan Mt., Kangding County (30.0°N, 101.9°E), Sichuan Prov- ince; coll. De-Niu Chen 8 Jia-Xiang Gao, 1982-IX-9. Fifteen adult (four dissected) and seven young specimens; IZCAS01061, bor- der of Jiuzaigou County and Songpan County (33%02'14.4"N, 103°42’32.1"Е), Sichuan Province, coll. Min Wu. One adult specimen dissected. Trichobradybaena submissa (Deshayes, 1873), type species of the genus: IZCAS00010, Hanzhong, Shaanxi Province, 1992-IV-15, coll. De-Niu Chen. Numerous specimens (three dissected). Helix pomatia (Linné, 1758) (Helicidae), type species of the genus: IZCAS00188, lime- stone quarry near Klodzko, Wapniarka Mt., Lower Silesia, Poland, 1999-VI-26, coll. Min Wu & Andrzej Wiktor. One adult specimen dissected. Calocochlea coccomelos (Sowerby, 1840): SMF323619, Philippines: Sibuyan, ex Moellendorff. One specimen dissected. Pfeifferia micans Pfeiffer, 1845, type species of the genus: SMF323620, Philippinen: Cagayan, Pamplona, O. v. Moellendorff. One adult specimen dissected. Camaena platyodon (L. Pfeiffer, 1846) (Camaenidae): IZCAS00833, Hainan, other collection data lacking. Eleven adult (three dissected) and one young specimens. APPENDIX Il: New taxa Aegistohadra, п. gen. Type species: Nanina delavayana Heude, 1885: 102, pl. xxvi, fig. 8. Aegistohadra delavayana (Heude, 1885), n. gen. & comb. (Figs. 6, 28-31; Table 2) Material Four adults (1ZCAS00132-1-4) of which two are full grown but broken and three young shells were examined, Zhibenshan Mt., Baoshan (26.3°N, 104.4°E) (“Yunlong County” in original label is a printing error), Yunnan Province; coll. De-Niu Chen, 1981- VI-23. 113 PHYLOGENY OF BRADYBAENIDAE OC 0 910 Y}PIM 194$ - A gç'ol €O'2L A "IS - Lae y-ZEL00SVOZI €9'0 18'9 - - 9/1 3/6 yl ze €1'0Z €-ZELOOSWOZI - - 96 SL < 8/1 8/6 - 86'0z Z-ZELO0SVOZI 95`0 EL + L8 zi Le'zl 9/1 р 0,67 65-91 L-ZELOOSVIZI UJPIM Jajaweıp ypIm биг spoymyouos = SHOUM UPI juBieu AuBleH Snoliqun ¡elmuedy jeınnedy -ojoJd “ON ‘ON TELS lleys иазеше!р snollIquu ‘qwos 9 ‘чэб ‘и euefenejap елрецо}з!6бау ‘TE LOOSVOZI зиэшюэа$ jo sjunoo pue ззиэшелпзеэш |еэ!бо|оцэцоЭ ‘с 3719VL 114 Two paratype specimens of Мапта delavayana H., IZCAS-type-2902-1 and IZCAS- type-2902-2 (Fig. 32), Fa Kouan Tchen, coll. unknown. Two paratype specimens of Helix seraphinica Heude, 1889 (Fig. 33). Etymology The genus name is derived from the names of two bradybaenid genera Aegista and Euhadra. Diagnosis Female part of genitalia with sac-shaped structure on vagina opposite to dart sac. Short Description Shell strongly depressed, sinistral, thick and solid. Umbilicus broad. Protoconch with radial wrinkles. Penial sheath absent; penis with a penial caecum near penial retractor; epiphallus with a flagellum; penis-epiphallus chamber absent; accessory sac undivided; in dart ap- paratus polylayered structure absent; \/1-\/4 in the dart apparatus present; two sacs in- serted on vagina opposite to dart sac. Full Description Shell sinistral, thin but solid. Apex distinct. Whorls convex. Suture strongly impressed. Umbilicus narrow to moderately wide. Col- umella oblique; columellar lip dilated, slightly covering umbilicus. Adult shell and young shell with smooth surface, spiral furrows irregularly and sparsely present, ribs absent; growth lines not accompanied by irregular thickenings, background microscopic ripples absent. Protoconch with radial wrinkles. Immature shells unkeeled and unangulated. Body whorl large, unkeeled, weakly descending in front, with convex bottom. Aperture rounded, ob- lique. Lip toothless, equally expanded, thin within. Peristome reflexed equally. Parietal callus indistinct. Shell dull, opaque; yellowish brown with two brown bands, one above and one beneath periphery, the lower sometimes not as distinct as the upper. Bottom of body whorl yellowish brown (Figs. 29, 30). Animal uniformly gray. Jaw arcuate with 7- 8 ribs dentating the concave margin, ribs con- tiguous, wide. In a paratype (IZCAS00132-2), radula with 169 rows of teeth, each with one FIG. 28. Distribution map. Square: Aegistohadra delavayana п. gen. & comb.; dots: Eueuhadra gonggashanensis n. gen. & sp. PHYLOGENY OF BRADYBAENIDAE IS central tooth and 53 lateral teeth at each side; central tooth and lateral teeth L,-L,, unicus- pid; L,,-L,, each with an endocone and ап ectocone (Fig. 31E). Genitalia: Penial sheath absent (Figs. 6A, 31A), except for some basal connective tis- sue present near atrium covering penis. Pe- nis short, slender, with a finger-shaped penial caecum near penial retractor (Figs. 6A, 6C, 31A, 31B). Retractor simple, thin or thick, short FIG. 29. Aegistohadra delavayana n. gen. & comb., shell near mature, IZCAS00132-1. A, apical view; B, basal view; C, apertural view. Bar equals 5 mm. or in moderate length. Epiphallus thick, short, with more or less protruding symmetrical epiphallic papilla (Figs. 6C, 31C). Penis- epiphallus chamber absent. Flagellum roundly blunt at end, with fairly smooth surface, thick, short (Figs. 6A, 6C, 31A, 31B); innerly folds not forming a C-shaped open tube towards fla- gellum or epiphallus (Figs. 6C). Pore of penial papilla located near the pore leading to penial caecum, mainly built by two pilasters derived from four thicker ones longitudinally arranged along penial inner wall (Figs. 6C, 31C). Dart sac developed, with an accessory sac below. Accessory sac large in size, slightly elongated (Figs. 6A, 6D, 31A). Dart sac containing one dart. Dart about 7.0 mm in length, almost straight, slightly expanding basally; cross sec- tion of dart throughout rounded or ovate at lower part, upper 1/4 with 2 opposite sharp ridges. (Fig. 31F). Inside dart sac, ADC shar- ing same entrance with DtC; V1-V4 present, V2 merged into a pilaster towards vagina; V1, V3 and V4 forming DtC; C23 present, but opened to vagina (Fig. 6D). Two sacs on va- gina opposite dart apparatus, one with two highly ridged pilasters, and another just be- neath the first one and with connective tissue inside, of unknown function (Figs. 6A, В & D); DVM absent. Mucous glands with two lobules, each as long as dart sac, stalks distinct, sepa- rated from dart sac and tied tightly to the trunk of vagina, inserting near base of dart sac (Fig. 31A). Lobules simply branched, distally sac- shaped. Bursa copulatrix slightly elongated, FIG. 30. Aegistohadra delavayana (Heude, 1885), п. gen. & comb., shell, IZCAS00132-3. А broken but adult shell, showing aperture structure. Bar equals 5 mm. 116 WU FIG. 31. Aegistohadra delavayana (Heude, 1885), п. gen. 8 comb., IZCAS00132-2. A, general view . of genitalia; B, penial complex; C, penis and epiphallus, opened; D, penial caecum (PC), opened; E, teeth of radula, bar equals 25 um; Е, dart, with cross-sections; С, a leaf of ovotestis. А-О, Е, С, bars equal 1 mm. PHYLOGENY OF BRADYBAENIDAE ПЕЙ well differentiated from its duct (Fig. 31A). Bursa copulatrix duct moderately long, insert- ing low on vagina (Fig. 31A). Ovotestis palm- shaped, with single stalk (Fig. 31G). Holotype: dart sac 10.0 mm in length, 2.5 mm in width, ratio of width to length 0.3; mucous duct length 9.3 mm; vagina length 13.8 тт; bursa copulatrix duct 18.0 mm long, basal width 1.3 mm; transverse diameter (maj.) of bursa copulatrix 1.8 mm, sagittal diameter (maj.) of bursa copulatrix 3.5 mm; vas deferens length 16.3 тт; penis length 12.0 тт; flagellum length 5.8 тт; epiphallus 3.8 mm in length; penial retractor 3.8 mm long (Fig. 31G). FIG. 32. Aegistohadra delavayana (Heude, 1885), п. gen. 8 comb., shell, paratypes, А-С, IZCAS-type- 2902-1; D-F, IZCAS-type-2902-2. Bar equals 10 mm. 118 Range Southwestern China (Fig. 28). Remarks This species can be distinguished from all known bradybaenid species in that the female part of the genitalia has a sac-shaped struc- ture on the vagina opposite to the dart sac. It also differs from all bradybaenids, except Eueuhadra gonggashanensis, п. gen. & sp., in having a pronounced penial caecum. Based on shell features, although distinctly larger (diam. maj.: 55 mm; min.: 48 тт; alt.: 30 mm), it is possible that Helix seraphinica Heude, 1889, from Silin (as “Xilin” in today's spelling, Guangxi Province) should be placed in Aegistohadra because of their similar shell shape (Fig. 33), as suggested by H. Nordsieck FIG. 33. Helix seraphinica Heude, 1889, shell, paratypes, А-С, IZCAS-type-3071-1; D-F, IZCAS-type- 3071-2. Bars equal 5 mm. PHYLOGENY OF BRADYBAENIDAE 119 (pers. comm.). However, Helix seraphinica we cannot be certain until its anatomy is known, considering the great morphological diversity shown in helicoid shells. Eueuhadra п. gen. Type species: Eueuhadra gonggashanensis, n. sp. Eueuhadra gonggashanensis, п. sp. (Figs. 7, 28, 34-39; Table 3) Material Holotype (IZCAS00067-1), West slope of Gonggashan Mt., Kangding County (30.0°N, 101.9°E), Sichuan Province; coll. De-Niu Chen & Jia-Xiang Gao, 1982-IX-9. Paratypes 14 (IZCAS00067-2-15), the same data as holo- type; seven young specimens (IZCAS00067- 16-22) were also examined; paratype 1 (17СА$01061), border of Jiuzaigou County and Songpan County (33°02'14.4’N, 10342'32.1”E), Sichuan Province; 3311 та. Ss. |.; coll. Min Wu, 2001-X-4. Etymology The genus name comes from “eu-” (real) and the bradybaenid genus Euhadra. The species is named after the holotype locality: Gonggashan Mountains. Diagnosis A simple penis-epiphallus chamber present; dart sac with multiple mucous branches. Short Description Shell depressed, dextral, thin but solid. Um- bilicus very narrow and more or less covered by columellar margin of the peristome. Protoconch shell granulose. Penial sheath absent; penis distally with an outstanding tube-shaped penial caecum; epiphallus with a flagellum; a simple penis- epiphallus chamber present; dart sac with a distinct and relatively large accessory sac on the end on which a bundle of mucous glands is inserted per one common duct; accessory sac undivided; in dart apparatus, polylayered structure absent, V1-V4 present. Range W China. Full Description Shell dextral, depressed, thin but solid. Apex distinct. Whorls convex. Suture impressed. Umbilicus very narrow and more or less cov- ered by columellar margin of peristome. Col- umella very oblique. Spiral furrows absent, without ribs, growth lines not accompanied by irregular thickenings, microscopic ripples ab- sent. Protoconch finely granulose, granulation regularly arranged. Teleoconch finely and un- evenly granulose on upper spire. Immature shells bluntly angulated. Whorls increasing rapidly; body whorl fairly large, unkeeled, FIG. 34. Eueuhadra gonggashanensis, n. gen. & sp., shell, IZCAS00067-1, Holotype, A, apical view; B, basal view; C, apertural view. Bar equals 5mm. WU 120 $0`0 690 650 616 10'6 mE ge 99°81 GO LL L90LOSVOZI 70`0 vo 0 v6 0 OL OL G8 OL nl "ev 96 LC 6/ LL y1-29000SVIZI 70`0 990 78`0 gr OL vv LL С G 8177 Gv ZL €l-Z9000SV9ZI GO 0 870 st! OL LL IL ZI G nV 19 ez GL'EL 7L-L9000SVOZI 70`0 9G 0 £6 0 £96 GL'OL С EV 87 02 6r LL LL-L9000SVOZI 70`0 vo 0 c8 0 Z9 OL vy LL hl EV G9 cc vec cl 01-29000SvV97Z1 90 0 79'0 OP L LC bb ÿ8 LL С SLY 28 cc 65 CL 6-/9000SvV971 GO 0 9G 0 6C | £9 LL evel A he L292 897, 8-29000SVIZI GO 0 £G'O OC L 08 OL 99 LL al! ev Gr ez ve cl 1-19000SWIZI 70`0 590 ÿ6 0 08 6 Lb bb SL AZ басс СИ ар 9-19000SVIZI $0'0 550 0) 08 OL ÿ9' LL int! 8/,t 8c ez 18 CL G-/9000SvV971 900 950 99| LO'LL 10`51 SL Y 96 rc 80 VL y-L9000SV9OZI 90 0 cS 0 931 cS OL 69 LL Tt ev 0677 L6 LL €-L9000SVOZI 70`0 190 860 90 LE 19° 2! 81, VLE? oL el c-L9000SVOZI sadfjeied 70`0 09'0 £O EL ZG bb 9ÿ CL YA G ¡AA voy! L-Z9000SVOZI adAjojoH YIPIM [19ys U}PIM Jayaweip U}PIM y¡Bue] SHOUM Y9UO9 SHOUM U}pIM yBieu eyeweip зпоашй AUBISH sha quin jeınyady jeınyady -0}01d ‘ON ‘ON ЭЧ$ BUS ‘ds ‘u 9 ‘uab ‘и 'sisugueysebbuob eipeyneng jo sadAyesed pue adÁjojoy jo ззипоэ pue sjuawalnseau ¡esibojoy9uo) ‘€ FJIGVL PHYLOGENY OF BRADYBAENIDAE 1241 slightly descended in front, with convex base. Aperture rather broadly lunate, more or less oblique. Lip toothless, uniformly thickened within, forming a ring-like thickening. Peris- tome thin, uniformly reflexed. Parietal callus distinct. Periostracum uniformly in greenish brown, bandless. Bottom of body whorl with same or lighter colour (Figs. 34, 35 8 39; Table 3). Animal with numerous brown spots on the anterior half. Jaw arcuate with 10-12 ribs FIG. 35. Eueuhadra gonggashanensis, п. gen. & sp., shell, 17СА$00067-5, Paratype, a shell with periostracum on. A, apical view; B, basal view; C, apertural view. Bar equals 5 mm. dentating the concave margin, ribs wide and almost contiguous. Radula of holotype with 133 rows of teeth, each with one central tooth and 44 lateral teeth on both sides; central tooth with 1 tiny cusp at each side; lateral teeth L,-L,, each with an ectocone; L,,—L,, each with a tiny endocone and an ectocone; main cones and ectocones of L,,—L,, bicuspid respectively, two cusps of ectocone roundly blunt (Fig. 36D). Genitalia: Penial sheath absent (Figs. 7A, 36A, 37A). Penis of moderate length, swollen, with a tube-like penial caecum (PC) near pe- nial retractor. Penial retractor short. Epiphallus thick, short. Epiphallic papilla depressed, sym- metrical. Penis internally with three thick pe- nial pilasters and two thinner ones among them (Figs. 36B, 37C). Near to the pore lead- ing to penial caecum, a papilla, built partially by above-mentioned pilasters present (Fig. 7B). Flagellum thick, short, smooth, abruptly tapering and forming a vermiform appendix (Fig. 36A). Penis-epiphallus chamber present, small, simple (Fig. 7E). Vas deferens inserted on flagellum, with inner folds forming a C- shaped open tube towards flagellum (Fig. 7C). Dart sac containing one dart. Dart approxi- mately 2 mm, medially rounded, apically trap- ezoid in cross section (Fig. 37E, only seen from the spirit material of IZCAS01061; in all dissected specimens of 17СА$00067, the darts are completely eroded, because of hav- ing been first fixed in formalin before being preserved in alcohol). Mucous glands longer than dart sac, inserted at the end of accessory sac; with 11-13 (in two specimens of IZCAS00067) mucous lobules radially ar- ranged, stalk of lobule indistinct; each lobule simply branched and consisting of slightly ex- panded vesicles, not expanded distally (Fig. 36A). Accessory sac developed, тпепу simple except for some narrow pilasters; a bundle of mucous glands inserting on the end of AS, from a common entering tube, its inner entrance without papilla (Fig. 7D); ADC share the same entrance with DtC; V1-V4 present; V1, V2 and V4 forming DtC; C23 present and tiny, with entrance leading to ADC (Figure 7D). DVM absent. Polylayered structure (PLs) ab- sent. Bursa copulatrix ovate, not well differen- tiated from bursa copulatrix duct (Fig. 36A). Bursa copulatrix duct of moderate length, wide, inserting high on vagina (Figs. 36A, 37A). Ovotestis palm-shaped, distinctly branched and two stalks closely arranged and having a common duct; in holotype, ovotestis embed- 122 WU FIG. 36. Eueuhadra gonggashanensis, п. gen. & sp., IZCAS00067-1, Holotype, A, general view of genitalia, bar equals 2 mm; В, penis, opened; С, penial caecum, opened; D, teeth of radula, bar equals 25 um; E, a leaf of ovotestis, bar equals 0.5 тт; Е, ovotestis matrix, magnified. B, С, bars equal 1 тт. PHYLOGENY OF BRADYBAENIDAE 123 Е Ер РВ С FIG. 37. Eueuhadra gonggashanensis, п. gen. & sp., genitalia, IZCAS01061, paratype, A, general view of genitalia; В, a branch of mucous glands; С, penis, penial caecum (PC), and flagellum, opened; D, ovotestis; E, dart with cross sections. Bars equal 1 mm. 124 ded in matrix composed of disordered fibers (Fig. 36F; in the other examined specimens the matrix normal); stalks fairly long (Figs. 36E, 37D). Holotype: dart sac 3.3 mm in length, 1.8 mm in width, ratio of width to length 0.5; mucous duct length 7.3 mm; vagina length 4.6 mm; free oviduct 12.2 mm; bursa copulatrix duct length 10.3 mm, bursa copulatrix duct basal width 1.5 mm; transverse diameter (maj.) of bursa copulatrix 1.5 mm, sagittal diameter (maj.) 2.2 mm; vas deferens length 13.7 mm; penis length 7.0 mm; epiphallus 6.4 mm; flagellum length 3.0 mm; PR length 2.2 mm. Range Western Sichuan, the species was known only from two localities where type and holo- type material were collected (Fig. 28). Remarks This species differs from all known bradybaenids by having а simple penis- epiphallus chamber. The sisterhood of this species and Aegistohadra delavayana (Heude, 1885), n. gen. & comb., is suggested by their common derived character the penial caecum. Ecology This species (17СА$01061) inhabits high mountains (Fig. 38B), in very low density. The environment is extremely wet, inside a dark fir forest, where the stones and fallen trunks are covered by a thick layer of lichen and moss which sometimes reaches the thickness of approximately 50 cm (Fig. 38C, D). The speci- men (IZCAS01061), the only collection after careful search of about 500 т? in the forest was found inactive under moss. The popula- tion in Jiuzaigou seems to be isolated from another known population in the Gonggashan Mountains. Based on 30-days field work cov- ering the area from Dujiangyan to Jiuzaigou along Minjiang River, it has been confirmed that these two populations are fairly sepa- FIG. 38. Habitat of Eueuhadra gonggashanensis, п. gen. & sp. A, Paratype, IZCAS01061, in its habitat; B-D, natural environment conditions of locality for Paratype IZCAS01061. PHYLOGENY OF BRADYBAENIDAE 125 rated. In the same area, по helicoid snails were found and only the non-helicoid snail Deroceras (Deroceras) altaicum (Simroth, 1886) (Wiktor et al., 2000), which is widely dis- tributed in the whole vally of Minjiang River and its neighboring mountains. It is also inter- esting that no conchologically similar species was recorded in this region before (e.g., Pilsbry, 1934). FIG. 39. Eueuhadra gonggashanensis, п. gen. & sp., shell, 17СА$01061, paratype, A, apical view; B, basal view; C, apertural view. Bar equals 5 mm. MALACOLOGIA, 2004, 46(1): 127-156 TOWARD COMPREHENSIVENESS: INCREASED MOLECULAR SAMPLING WITHIN CYPRAEIDAE AND ITS PHYLOGENETIC IMPLICATIONS Christopher P. Meyer Florida Museum of Natural History, University of Florida, Gainesville, Florida 32611 USA; cmeyer@flmnh.ufl.edu ABSTRACT This paper introduces 73 additional taxa to the existing mitochondrial molecular data- base of 202 taxa for the Cypraeidae and addresses the systematic implications of their inclusion. Five outgroup members from the Ovulidae are also added. Sequence data are included from all previously missing extant named genera (Propustularia, Barycypraea and Schilderia), completing the overall “generic-level” framework for living cowries. Newly added taxa include 47 recognized species, 25 subspecies, and six undescribed taxa. Phy- logenetic results generally are consistent with previous arrangements, with few minor ad- justments. The most significant findings are that: (1) currently recognized Nesiocypraea 1$ broken into two disparate clades, a deeply rooting Nesiocypraea sensu stricto group and the more derived Austrasiatica (Lorenz, 1989). (2) Two newly included Barycypraea taxa are sister to Zoila, reaffirming the validity of the subfamilial clade Bernayinae. (3) The inclusion of a significant number of added Erroneini taxa (N = 24) creates a phylogenetic challenge because of poor support and recovered relationships inconsistent at first glance with traditionally recognized affinities. In order to maintain nomenclatural consistency, Erronea is maintained at a generic level, whereas Adusta is dropped to subgeneric status within Erronea. Greater than 90% of currently recognized species are included, and 93% of these are supported by molecular criteria. Moreover, more than 70% of the tested, recognized subspecies are distinct. The phylogeny provides one of the most comprehen- sive, species-level frameworks to date for testing diversification theories in the marine tropics. Key words: Cypraeidae, molecular systematics, taxon sampling, Cypraea. INTRODUCTION Cowries (Gastropoda: Cypraeidae) are taxo- nomically one of the best known of all mollus- can groups, and have been used frequently to examine speciation and biogeographic pat- terns in the marine tropics (Schilder, 1965, 1969; Foin, 1976; Kay, 1984, 1990; Meyer, 2003). A wealth of taxonomic (Schilder & Schilder, 1938, 1971; Schilder, 1939; Lorenz & Hubert, 1993; Groves, 1994; Lorenz, 2002), anatomical (Troschel, 1863; Vayssiére, 1923, 1927; Riese, 1931; Risbec, 1937; Schilder, 1936; Kay, 1957, 1960, 1963, 1985, 1996; Bradner & Kay, 1996; Lorenz, 2000), biogeo- graphic (Schilder, 1965, 1969; Foin, 1976; Burgess, 1985; Liltved, 1989; Lorenz & Hubert, 1993; Lorenz, 2002) and fossil data (Schilder & Schilder, 1971; Kay, 1990, 1996; Groves, 1994) is available for the group; how- ever, what has been lacking is a well-resolved, comprehensive species-level phylogeny. 127 These phylogenetic hypotheses of relationship establish sister pairs at the appropriate taxo- nomic level and provide the framework to test diversification theories. Meyer (2003) introduced molecular data for 234 taxa in Cypraeidae and generated phylogenetic hypotheses for most major clades as well as sister-group relation- ships for most species. Systematics for Cypraeidae were reviewed in light of the results and diversification patterns within the tropics were addressed. The study presented herein significantly increases the comprehensiveness of taxon sampling in the group by introducing 73 Cypraeidae and five Ovulidae taxa to the exist- ing molecular dataset and discusses their sys- tematic implications. In addition to broader taxonomic sampling, this paper presents the re- sults of broader geographic sampling. The ap- pendix lists 147 localities added across the various taxa. Five outgroup taxa from six locali- ties are included, and 67 recognized cypraeid species or subspecies are added from 75 locali- 128 MEYER ties. The remaining 66 localities were added to supposedly known taxa, but revealed six previ- ously unrecognized taxa, some of which may correspond to names currently in synonymy upon review of type localities. MATERIALS AND METHODS Recognition Criteria: ESU versus OTU The ultimate goal of this project is to con- struct a comprehensive phylogeny of cypraeid gastropods at the appropriate level for diversi- fication studies. As such, the operational taxo- nomic unit (OTU) chosen for phylogenetic analyses generally represents an evolution- arily significant unit (ESU) that must fulfill some minimal criteria established through genetic scrutiny. First, mtDNA haplotypes of sampled individuals must represent a mono- phyletic clade; yet this alone is not sufficient, because any phylogeny has a plethora of monophyletic groups, because a clade re- quires only two individuals. Thus, auxiliary cri- teria are required to delineate significant units. Within cowries, these additional criteria are (1) geographic distinction or allopatry, (2) signifi- cant genetic distance from the sister group such that pairwise distance comparisons yield a bimodal distribution, and/or (3) taxonomic recognition by previous workers. An OTU is included in analyses only if at least two of these three criteria are met. Most OTUs fulfill all three criteria and are considered evolution- arily significant units (ESUs) (sensu Moritz, 1994). These criteria are erected in order to delineate independent evolutionary trajecto- ries, but do not guarantee that the units are reproductively isolated. In a few instances, two of the three criteria (genetic separation and taxonomic recognition) are not supported by the third (exclusive geographic signatures). While the genetic differences (monophyly) between populations indicate some indepen- dent period of evolutionary history between geographic regions, it appears that, on occa- sion, haplotypes from outlying regions can mix back into the sister gene pool. The few cases where all three criteria are not fulfilled always occur on the periphery of regions (e.g., Marquesas, Hawaii) and show asymmetrical, “downstream”, dispersal events (Fig. 1). As circumscribed, all ESUs discussed indicate independent evolutionary histories, but alter- native criteria, such as either nuclear markers or breeding experiments, are needed to verify reproductive isolation. Molecular Methods Most methods follow protocols detailed in Meyer (2003) for all aspects of preservation, extraction, amplification, and sequencing. Tis- sue samples were acquired from a variety of catholicorum 964 catholicorum 908 TAITAE 2131 X catholicorum 897 gaskoinii 655 gaskoini 972 gaskoini 975 gaskoini 971 gaskoini 2362 gaskoini gaskoini 973 gaskoini 974 gaskoini 977 gaskoini 654 gaskoini 976 astaryi 2136 ситтой ТК 2138 astaryi 1472 astaryi 2134 astaryi 2135 astaryi 2137 cumingii TIK 1970 astaryi 1105 astaryi 1471 astaryi 2133 GARCIAI 2413 ситтой TIK 2140 cumingii RR 1966 cumingii RR 2144 cumingii RR 2143 cumingii RR 1968 cumingii TIK 1969 cumingii TIK 2139 cumingii RR 2141 cumingii RR 2142 cumingii RR 1967 cumingii HUA 700 100 catholicorum gaskoini astaryi 1% cumingii FIG. 1. ESU vs. OTU criteria. Phylogram show- ing the relationships among members of the Pacific Cribrarula subclade, with bootstrap val- ues for major groups. Four distinct clades are evident, and the names presented on the right: Cribrarula catholicorum, С. gaskoini, С. astaryi, and C. cumingii. Note that single individuals of two newly included taxa, С. taitae and С. garciai (white stars), nest within two of the major clades and show little variation (a single mutation). These two new taxa are introduced as OTUs, because of their distinct morphology and geog- raphy (American Samoa and Easter Island, re- spectively), but are currently not considered ESUs by molecular criteria. All individuals from the Marquesas are С. astaryi; however, two in- dividuals of С. cumingii possess haplotypes be- longing to the С. astaryi clade as well (dark stars). While the two haplotype clusters are distinct, the pattern indicates uni-directional exchange of lar- vae downstream from the Marquesas (С. astaryi). Molecular criteria recognize these two clades as ESUs with historically limited exchange. (TIK = Tikehau, RR = Rangiroa, HUA = Huahine, all C. astaryi from Marquesas, all C. gaskoini from Hawaii, and all C. catholicorum from Solomon Islands) INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE 129 sources and locations (listed in the acknowledgements and appendix). Most samples were preserved т 95% ethanol. DNA extraction was performed using DNAzol (Chomczynski et al., 1997) using one-half vol- umes and following the manufacturer's proto- col (Molecular Research Center, Inc.) with the exception that the digestion step was in- creased by an additional 24 or 48 h. PCR was performed as described in Meyer (2003). COI primers were as follow (from Folmer et al., 1994): LCO-1490 (5’—3’) GGT CAA CAA ATC АТА ААС ATA TTG С, and HCO-2198 (5'-3”) TAA ACT TCA GGG TGA CCA AAA ATC A. For problematic taxa, these primers were de- generated as follows: dgLCO-1490 (5’—3’) GGT CAA CAAATC АТАААС AYA TYG С, and dgHCO-2198 (5’—3’) TAAACT TCA GGG TGA CCA AAR AAY CA. Two internal primers were designed for small amplifications of degraded DNA: InCypLCO (5’-3’) CGT YTA AAT AAT ATAAGY ТТУ TG, and InCypHCO (5'-3”) CGT ATA TTA ATA ATT СТТ GTA AT. Palumbi's (1996) 16Sar and 16Sbr primers were used for 16S: 16Sar (5’—3’) CGC СТС TTT ATC AAA ААС АТ, and 16Sbr (5'-3”) CCG GTC TGAACT CAG ATC АСС T. Two internal primers were designed for small amplifications of degraded DNA: In16Sar (5’—3’) GGG CTA GTATGAATG GTT TGA, and In16Sbr (5’—3’) ATG СТС TTA ТСС CTATGG TAA СТ. The polymerase chain reaction was carried out in 50 ul volumes, us- ing 1 ul of template. Each reaction included 5 ul 10X PCR buffer, 5 ul dNTPs (10mM stock), 2 ul of each primer (10uM stock), 3 м MgCl, solution (25 mM stock), 0.2 ul Taq (5 Units/pl stock) and 31.8 ul ddH,O. Reactions were run for 35-40 cycles with the following parameters: an initial one min denaturation at 95°C; then cycled at 95°C for 40 sec (denaturation), 40°C to 44°C (CO!) or 50°C to 54°C (16S) for 40 sec (annealing), and 72°C for 60 sec (extension). Successfully amplified products were cleaned for cycle sequencing using Wizard® PCR Preps (Promega). Sequencing also followed Meyer (2003) with all new sequences generated using ABI chemistry and sequencers. Sequences were generated from the resulting electrophenograms using Sequencher (Gene Codes). All primer sequences, aligned СО! and 16$ sequences and Nexus files are available at the archived data web pages of the Florida Museum of Natural History Malacology Depart- ment (http://www.flmnh.ufl.edu/malacology/ archdata/Meyer2004), and new sequences are deposited in Genbank under accession num- bers AY534351 through AY534503. Phylogenetic Analyses The 297 operational taxonomic units (OTUs) presented in this paper were selected from an extensive database comprised of over 2,000 sequenced individuals. In general, taxa are included if they exhibit distinctive geographic and/or genetic signatures. In most instances, new OTUs are recognized in the literature as either species (N = 47) or subspecies (N = 25). This paper introduces six previously un- recognized taxa. The increasing size of this dataset presents computational and heuristic challenges for phylogenetic analyses. Two weighted trans- version bias parsimony searches (3:1 and 5:1) were performed on the complete dataset us- ing PAUP* (Swofford, 1998). At first, 250 ran- dom-addition replicate searches were performed, but with a tree limit of ten imposed to minimize search time on suboptimal islands. After 250 replicates, the most parsimonious topologies were used as starting trees for ex- haustive searches without tree limits. This strategy was employed for both weighted analyses, and the most parsimonious topolo- gies were pooled and evaluated using likeli- hood criteria. ModelTest v. 3.06 (Posada & Crandall, 1998) was used to select the most appropriate model for likelihood parameters. The most likely weighted parsimonious trees were then compared using consensus meth- ods. A two-tiered, compartmentalized strategy was adopted that followed Meyer (2003) for levels of topological support. The strict con- sensus topology derived from the most likely overall analyses was divided into four subequal components called basal, midi, mid2, and derived. Because the basal, mid1 and mid2 cohorts are necessarily paraphyletic groups that include the common ancestor and some, but not all, of its descendants, repre- sentative derived clades were included in the paraphyletic analyses. In this way multiple derived member clades overlapped between more basal and derived analyses, and the overall topology could be “scaffolded” together by linking clades shared in both basal and derived compartments. Within each of the four subanalyses, parsi- mony searches were performed using a 5:1 transversion bias. Both bootstrap (Felsenstein, 1985) analyses (1,000 replicates) and decay (Bremer, 1994) analyses (TreeRot v2; Sorenson, 1999) were performed to establish levels of support. Results from Bayesian meth- 130 MEYER ods (Mr. Bayes v3.04b) are not reported in this paper, but were generated for the four sub- groups and compared to the combined parsi- mony/likelihood methods utilized in PAUP*. Overwhelmingly, they were consistent with the results presented here, but on few occasions differed in hypotheses of relationship. The scaffolded parsimony global topologies were compared to the scaffolded Bayesian topol- оду using likelihood criteria in PAUP*. The combined topology derived from the compart- mentalized Bayesian subsets was less likely than the overall topologies found using the combined parsimony/likelihood criteria. It ap- pears that Bayesian results depended on taxon sampling and outgroup inclusion. While this finding may be of interest to the general sys- tematic community, it is not a point specifically addressed in this paper. RESULTS The final culled dataset contained 297 OTUs and 1,107 characters, 493 base pairs from 16$ and 614 bases from COI. For 16$, alignment followed those presented in Meyer (2003) based on secondary structure. Weighted par- simony searches resulted in 512 equally most parsimonious trees (MPTs) for 3:1 Ti: Tv and 480 trees for 5:1 searches. Derived portions of the comprehensive topology were consistent. Thus, all named clades (subfamilies, tribes and gen- era) presented in Figure 2 are found in all to- pologies, except one mentioned below. However, the topologies recovered from alter- nate weightings differed in five deeper regions, all of which are poorly supported regardless of methodology. First, 5:1 topologies placed the clade consisting of Propustularia/Nesiocypraeal Ipsa basal as sister to all other cowries. In 3:1 topologies this clade moves up one node and is sister to Erosariinae. Second, the pustulose clade consisting of Nucleolaria/Cryptocypraeal Staphylaea is monophyletic in 5:1 trees, while in 3:1 topologies these genera are a basal paraphyletic grade leading to the clade includ- ing Monetaria/Perisserosa/Erosaria. Third, in 5:1 topologies Perisserosa is sister to Erosaria, whereas in 3:1 trees, Perisserosa is sister to Monetaria. Fourth, the arrangement of major groups along the backbone from Umbiliini to Cypraeovulinae conflicts. Results from 5:1 searches are shown in Figure 2, whereas in 3:1 topologies, Notocypraea and Cypraeovula (Cypraeovulinae) are a basal sister grade lead- ing to more derived member groups. Finally, the basal arrangement within Erroneini is dif- ferent. In 3:1 topologies Purpuradusta is more basal, while in 5:1 trees, Erronea is more basal. When alternative topologies were evaluated using ModelTest, the GTR+I+G model was se- lected as the best-fit model. When both the 3:1 MPTs and 5:1 MPTs were evaluated using the selected likelihood criteria [lset base = (05315128 05136452 0.111915) INstz=s6; Rmat = (0.99559 41.36057 1.0461 1.68935 22.78834), rates = gamma, shape = 0.562423, Pinvar = 0.48426], the 5:1 subset was signifi- cantly more likely (ANOVA: p < 0.001, average -In likelihood = 49513.8). Therefore, results from the 5:1 searches are presented herein. The overall relationships among major sub- groups recovered in the 5:1 MPTs are more consistent with both morphological and fossil evidence in addition to being more likely based on molecular data. In particular, a monophyl- etic pustulose clade is more parsimonious for conchological and anatomical features, be- cause it is more likely that a bumpy shell was derived a single time, rather than being derived either twice independently, or derived once then lost. Also, the basal, paraphyletic status of Notocypraea and Cypraeovula within the 3:1 topologies is inconsistent with the fossil record for both groups relative to more derived mem- bers of the 3:1 МРТ$ (i.e., Umbilia, Barycypraea, and Zoila), which appear earlier in the record and root more deeply in the 5:1 topologies. Also, the sister-group relationship of the two genera is more consistent with paleobiogeography (the breakup of Gondwanaland) and recognized affinities based on both conchological and de- velopmental criteria. The other major discrep- ancies between the 3:1 and 5:1 MPTs (most basal cowries, Perisserosa affinities, and posi- tion of Purpuradusta) are more ambiguous based on alternate criteria (morphological or paleontological). Suprageneric Relationships (Fig. 2) Overall, suprageneric results were consistent with previous systematic findings (Meyer, 2003), with two exceptions. First, /psa falls out- side Erosariinae and is no longer sister to Erosariini, but instead is allied with newly in- cluded Propustularia and Nesiocypraea sensu stricto. New sequence data from Nesiocypraea teramachii neocaledonica did not result in an affinity with other recognized “Nesiocypraea” species (N. hirasei, N. sakurai and N. langfordi). Instead, Nesiocypraea teramachii roots more deeply in the phylogeny as a distant sister to lpsa childreni, within a clade that includes both Ipsa and Propustularia. Thus, the inclusion of INCREASED MOLECULAR ЗАМРИМС IN CYPRAEIDAE 1331 Ovulidae PROPUSTULARIA NESIOCYPRAEA Ipsa Cryptocypraea Nucleolaria Staphylaea Monetaria Perisserosa Erosaria | pe Umbilia | Umbiliini i BASAL LA. Cypraeidae Erosariini Erosariinae Muracypraea Cypraea Macrocypraea Cypraeinae Leporicypraea Mauritiini Mauritia BARYCYPRAEA 7 Zoila Bernayinae Talparia Luria Trona Annepona Chelycypraea Austrocypraein Austrocypraea Arestoides NT Lyncina! Pustularia Neobernaya Pseudozonaria SCHILDERIA PESO Zonaria MID 2 Bistolidini Cypraeini MID 1 Luriini Luriinae Pustulariinae Notocypraea Cypraeovula Palmadusta Bistolida Ovatipsa Talostolida Cribrarula Austrasiatica Pamulacypraea Erronea? Purpuradusta Contradusta Erroneini DERIVED Erroneinae Notadusta Eclogavena Melicerona Blasicrura FIG. 2. Strict suprageneric consensus topology of 480 most parsimonious trees derived from a 5:1 Ti: Tv weighted search strategy of all 297 OTUs. Subfamilies are indicated with arrows and tribes are listed to the right. The four compartments for further subanalyses are bracketed to the right. The four newly added genera are capitalized and bolded. ‘Lyncina includes the subclades Callistocypraea, Miolyncina and Lyncina as reported in Meyer (2003). 2Austrasiatica replaces the prior use of Nesiocypraea for the same clade. *Erronea now includes Adusta, formerly rec- ognized as the sister taxon. 132 MEYER two new ancient lineages (Propustularia and Nesiocypraea) affects the relative position of lpsa. Moreover, the finding that Nesiocypraea teramachii is not related to other previously rec- ognized Nesiocypraea, compels me to recog- nize the clade Austrasiatica proposed by Lorenz (1989) at the generic level for the group includ- ing Austrasiatica hirasei, A. sakurai, and A. langfordi. There are some conchological and anatomical features that support this separa- tion. The left posterior terminal ridge in Nesiocypraea is more produced and separate from the body of the shell, whereas in Aus'rasiatica, the ridge is continuous with the boay. Lorenz (pers. comm.) also states that (1) Nesiocypraea lacks a distinct embryonic band- ing, having instead only a darker middorsal zone, (2) Nesiocypraea have a proportionally larger spire, and (3) the darker pattern of the shell is absent in juvenile Austrasiatica, only gained after the deflection of the labral margin; whereas, the darker pattern can be part of ju- venile Nesiocypraea shells. Additionally, the rachidian tooth of Nesiocypraea lacks the prominent paired basal denticles present in the three Austrasiatica taxa, and the tooth shape is less elongated and squared, whereas the rachidian in Austrasiatica narrows toward the cusps (Bradner & Kay, 1996). The fact that Austrasiatica was erected to differentiate the three species (albeit incorrectly aligned with Schilderia) is also an indication that the two lin- eages possess independent histories. The deep position of Propustularia within the cowrie phy- logeny is not surprising because it is one ofthe oldest of extant taxa, extending back to the Lower Eocene (Kay, 1996). The second suprageneric difference concerns the relative position of Zoila in the overall phy- logeny and is caused by the inclusion of se- quence data for two taxa from the ancient lineage Barycypraea. These new data indicate that Barycypraea teulerei and Barycypraea fultoni are sister taxa, and they are sister to Zoila. This BarycypraealZoila clade is recog- nized as the extant members of the subfamily Bernayinae, a group that includes many extinct fossil members and extends back into the Me- sozoic (Kay, 1996). These new data change the relative position of Zoila to Cypraeinae (Meyer, 2003); however, the topology in this region of the phylogeny is poorly supported. The final suprageneric addition to the molecu- lar database is the inclusion of sequence data from Schilderia achatidea, the single, living rep- resentative from an older, more diverse genus of European affinities. Previously, the paraphyletic arrangement of the genera Pseudozonaria and Zonaria was a surprising result (Meyer, 2003). These new data for Schilderia place the genus as sister to Zonaria to the exclusion of Pseudozonaria (and Neobernaya), and phylogenetic results main- tain their independent, paraphyletic status. These finding are more consistent with geo- graphic affinities than recognized taxonomic affinities (Pseudozonaria is often considered a subgenus of Zonaria), as both Neobernaya and Pseudozonaria are currently restricted to the eastern Pacific whereas Schilderia and Zonaria are restricted to the western Atlantic. Basal Compartment (Fig. 3) Five Ovulidae taxa are added in these analy- ses: Pseudocypraea exquisita, Volva volva, Primovula concinna, Dentiovula takeoi, and Prosimnia semperi. Within Ovulidae, only a few major clades are well supported and may be the results of poor taxon sampling. First, the clade Eocypraeinae appears well supported and includes Pedicularia, Jenneria and Pseudocypraea. Eocypraeinae is sister to a strongly supported clade (Ovulinae) that in- cludes the remaining Ovulidae. Within the Ovulinae, two subgroups are well supported and represent the major clades Volvini and Ovulini. Of the added Ovulidae, Volva falls into Volvini, but Prosimnia unexpectedly falls into Ovulini as do Primovula and Dentiovula. These results are generally consistent with Cate’s (1974) arrangement of higher-level relation- ships within the Ovulidae. Cyphoma gibbosum falls basal to these two sisters in the strict con- sensus topology; however, its position is poorly supported, and it is expected to move within the Volvini with the inclusion of more taxa. Monophyly of Ovulidae is not addressed herein and would require the inclusion of more distant representatives from Lamellaridae, Triviidae and Eratoidae. The Cypraeidae basal group includes the genera Propustularia, Nesiocypraea, Ipsa, Cryptocypraea, Nucleolaria, Staphylaea, Monetaria, Perisserosa, and Erosaria. Propustularia, Nesiocypraea, and /psa form a clade that roots deeply within the phylogeny and is sister to all other cowries. Each of the three genera is represented by only a single taxon, and only Nesiocypraea contains additional rec- ognized species missing from the dataset (Nesiocypraea midwayensis, N. lisetae and N. aenigma). While sharing a most recent com- mon ancestor, the three genera are highly di- vergent from each other, representing significant periods of independent history. Two INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE a Strict 93 17 1 1100] a 66 100 1 90 10 79 10 Pedicularia pacifica Jenneria pustulata Pseudocypraea adamsonii Pseudocypraea exquisita (1 Cyphoma gibbosum Neosimnia aequalis Volva volva (2) Phenacovolva weaveri Phenacovolva tokioi Ovula ovum Serratovolva dondani Calpurnus verrucosus Crenavolva rosewateri Primovula concinna (3) Procalpurnus lacteus Crenavolva tokuoi Crenavolva cf rosewateri Dentiovula takeoi (4) Prionovolva brevis Adamantia florida Prosimnia semperi (5) [Surinamensis (6) | Nesiocypraea Ipsa Cryptocypraea nucleus i Nucieolaria limacina interstincta limacina limacina staphylaea laevigata staphylaea staphylaea semiplota Eocypraeinae eeuINAO Propustularia Staphylaea annulus obvelata* moneta caputophidii caputserpentis caputdraconis Monetaria Perisserosa marginalis (8) citrina (9) helvola helvola helvola cf. callista (10) helvola hawaiiensis turdus irrorata (11) Erosaria erosa (Pacific) erosa (Indian) tigris (Indian) tigris (Pacific) pantherina Cypraea Talparia Luria isabellamexicana 133 Pedicularia pacifica Jenneria pustulata Pseudocypraea adamsonii Pseudocypraea exquisita Cyphoma gibbosum Neosimnia aequalis Volva volva Phenacovolva weaveri Phenacovolva tokioi Ovula ovum Serratovolva dondani Calpurnus verrucosus Crenavolva rosewateri Primovula concinna Procalpurnus lacteus Crenavolva tokuoi Crenavolva cf rosewateri Dentiovula takeoi Prionovolva brevis Adamantia florida Prosimnia semperi surinamensis teramachii neocaledonica childreni dillwyni nucleus granulata* limacina interstincta limacina limacina staphylaea laevigata staphylaea staphylaea semiplota annulus obvelata* moneta caputophidii caputserpentis caputdraconis guttata marginalis citrina helvola helvola helvola cf. callista helvola hawaiiensis turdus irrorata albuginosa poraria beckii macandrewi englerti kingae thomasi cernica spurca acicularis nebrites erosa (IO) erosa (PO) labrolineata boivinii ocellata gangranosa miliaris eburnea* lamarckii cf. redimita lamarckii lamarckii 0.1 substitutions/site =) — 0.05 substitutions/site armeniaca hesitata capricornica cf. petilirostris mus tigris (10) tigris (PO) pantherina talpa exusta pulchra cinerea lurida tessellata isabella isabellamexicana FIG. 3. Basal Compartment cladogram and phylogram. Bootstrap values are presented above branches in the cladogram and rescaled decay values below. Bolded taxa are new additions to the data set. Their identity number shown in parentheses follows the listing in the Appendix. Generic or suprageneric groupings are indicated to the right of the cladogram. OTUs with an asterisk (*) are not ESUs based on molecular criteria. Phylogram to the right is based on likelihood distances using a GTR+I+G model of sequence evolution. Note that the scaling for branch lengths changes between Ovulidae and Cypraeidae. 134 МЕУЕК are known exclusively from the Indo-Pacific (Nesiocypraea and Ipsa) and one (Propustularia) from the western Atlantic, but has a fossil record from North America, the Caribbean, and Europe (Kay, 1996). The splits among these ancient groups are among the earliest of all extant species and may have ос- curred in the Mesozoic. While reasonably sup- ported as a clade, this basal group is not strongly supported as the most basal sister, and in other analyses (3:1) moves up to become sister of the remaining basal taxa (Erosariinae). The final six genera from the basal compart- ment form the strongly supported clade Erosariinae and is the sister group to all remain- ing extant species. Membership and relation- ships within the Erosariinae are consistent with previous findings (Meyer, 2003). Five taxa from Erosaria are added: Erosaria marginalis, E. citrina, E. helvola cf. callista, E. macandrewi, and Е. englerti. Ten independent lineages are strongly supported (bootstraps > 90/decays > 6) within Erosaria, but interrelationships among them are not (< 50/< 4). Erosaria marginalis and E. citrina, both from the western Indian Ocean, are strongly supported as sister taxa. This саде 1$ poorly supported as sister to the E. helvola complex. Within Erosaria helvola, three ESUs are identifiable: E. helvola hawaliensis from Hawaii, E. helvola cf. callista from the Marquesas, and E. helvola helvola from the remainder of the IndoPacific. The newly included ESU, E. helvola cf. callista, may need a new name, because the type locality of E. helvola callista is Tahiti (Shaw, 1909), not the Marquesas. These five taxa are sister to the remaining Erosaria; however, the basal po- sition is poorly supported. Erosaria turdus is a monotypic, deeply divergent lineage. Newly added Erosaria irrorata, a species restricted to the oceanic islands of the Pacific, is poorly sup- ported as sister to a strongly supported clade (97/12) including E. albuginosa and E. poraria. These three taxa are sister to a well-supported lineage (92/6) of eight taxa that | tentatively rec- ognize as Paulonaria at the subgeneric level. New sequence data from Erosaria macandrewi, a Red Sea taxon, closely ally that species with Е. beckii. These two species are sister to the remaining Paulonaria taxa. The final additional taxon within Paulonaria is Erosaria englerti, a species endemic to Easter Island and Sala y Gomez. Erosaria englerti shares a more recent common ancestor with the remaining five Paulonaria taxa. All other relationships within Erosaria are the same as those presented in Meyer (2003) and are indicated in Figure 3. Newly added haplotypes from E. lamarckii lamarckii populations of the western Indian Ocean exhibit a recent divergence from the previously recorded Е. lamarckii cf. redimita of the Andaman Sea. One final finding from addi- tional Erosaria sequence data is that haplotypes from Erosaria miliaris and E. eburnea individu- als interfinger, indicating that either the diver- gence between these two taxa is very recent and lineage sorting has not occurred, or that these two taxa represent a cline across the western Pacific from a colored dorsum in the west to white shells in the east. Mid1 Compartment (Fig. 4) The second paraphyletic compartment con- tains mostly large-shelled taxa from the follow- ing tribes: Umbiliini, Cypraeini, Mauritiini, Luriini, Austrocypraeini, and the genus Pustularia. All six clades are well supported (> 70/> 5) except for Austrocypraeini. As in Meyer (2003), inter- relationships among these major suprageneric clades are resolved in the consensus, but poorly supported. Austrocypraeini and Luriini are sis- ters and recognized as the subfamily Luriinae. Barycypraea and Zoila are sisters and recog- nized as the subfamily Bernayinae. Cypraeini and Mauritiini are sisters and recognized as the subfamily Cypraeinae. In the current topology, Pustularia and all remaining cowries share a more recent common ancestor. This large clade is sister to Luriinae, which in turn is sister to Bernayinae, and this inclusive clade is sister to Cypraeinae. As in Meyer (2003), Umbiliini is sis- ter to all remaining mid1, mid2 and derived taxa. Within the mid1 compartment, 13 taxa are added to the sequence database. The first addi- tion falls within the genus Umbilia and is tenta- tively recognized as Umbilia cf. petilirostris. A single divergent sequence was generated from tissue samples collected from the deep waters in the Capricorn Channel off Queensland, Aus- tralia. Seven sequenced individuals were com- pletely identical, while an eighth sample from a subadult shell was significantly divergent. This single sample may represent the newly de- scribed Umbilia petilirostris Darragh, 2002; how- ever, authors disagree on its taxonomic status (Wilson & Clarkson, in press). Until more com- prehensive sampling is done in the region, | present the divergent sequence as a different ESU, which does not preclude it from being lumped within U. capricornica at a later date with more exhaustive sampling. The relation- ships within Umbilia remain as in previous analyses (Meyer, 2003). The second taxon added to mid1 is [ероп- cypraea mappa aliwalensis from Natal, South ats FIG. 4. Mid 1 Compartment cladogram and phylogram. All other information as in Fig. INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE Strict 93 15 99 13 > 97 = 51 24 nucleus ranulata* limacina interstincta limacina limacina staphylaea laevigata staphylaea staphylaea semiplota armeniaca hesitata capricornica tigris (Indian) tigris (Pacific) cervinetta cervus zebra valentia mappa mappa mappa aliwalensis (16) mappa rosea geographica mappa viridis scurra indica scurra scurra mauritiana depressa dispersa depressa depressa grayana eglantina histrio maculifera martybealsi maculifera maculifera maculifera scindata arabica (AmSamoa) (17 arabica arabica arabica asiatica Larabicaimmanis | teulerei (18) 5 marginata ketyana (20) marginata marginata venusta rosselli eludens decipiens mariellae (21) friendii jeaniana friendii thersites* talpa exusta (22) pulchra cinerea lurida tessellata isabella stercoraria mariae testudinaria reevei argus argus argus contrastriata (23 nivosa broderipii (24) leucodon aurantium porteri lynx vitellus ventriculus (Xmas) (25) ventriculus schilderorum sulcidentata kuroharai (26) leviathan carneola propinqua [globulusgoblus | globulus brevirostris cicercula margarita mauiensis bist. keelingensis (27) bistrinotata bistrinotata bist. sublaevis (28) spadicea annettae robertsi arabicula nigropunctata achatides picta sanguinolenta zonaria pyrum ) angelicae* Cryptocypraea Nucleolaria Staphylaea Umbilia Muracypraea Cypraea Macrocypraea Leporicypraea Mauritia Barycypraea Zoila Talparia Luria Trona Annepona Chelycypraea Austrocypraea Arestoides Lyncina Pustularia Neobernaya Pseudozonaria Schilderia Zonaria 0.05 substitutions/site 135 dillwyni nucleus granulata* limacina interstincta limacina limacina staph. laevigata staph. staphylaea semiplota armeniaca hesitata capricornica cf. petilirostris mus tigris (10) tigris (PO) pantherina cervinetta cervus zebra valentia mappa mappa mappa aliwalensis mappa rosea geographica mappa viridis mappa admirabilis scurra indica scurra scurra mauritiana depressa dispersa depressa depressa grayana eglantina histrio maculifera martybealsi maculifera maculifera maculifera scindata arabica (AmSamoa) arabica arabica arabica asiatica arabica immanis teulerei fultoni massieri marginata ketyana marginata marginata venusta rosselli eludens decipiens mariallae friendii jeaniana friendii thersites* talpa exusta pulchra cinerea lurida tessellata isabella isabellamexicana stercoraria mariae testudinaria reevei argus argus argus contrastriata nivosa broderipii leucodon aurantium porteri lynx vitellus ventriculus (XmaslO) ventriculus (PO) schilderorum sulcidentata kuroharai leviathan carneola propinqua globulus globulus globulus brevirostris cicercula margarita mauiensis bistrinotata keelingensis bistrinotata bistrinotata bistrinotata sublaevis spadicea annettae robertsi arabicula nigropunctata achatidea zonaria pyrum angelicae* sanguinolenta picta 136 MEYER Africa, and falls as sister to Leporicypraea mappa rosea. Lorenz (2002) has recently re- vised the taxonomy of the mappa group in light of molecular findings. Importantly, the names | associated previously with ESUs have changed, and those changes are reflected in the Appendix and also discussed herein. The taxon | previously recognized as Leporicypraea тарра viridis from SE Polynesia 1$ now recog- nized as Leporicypraea admirabilis. The taxon | previously recognized as Leporicypraea mappa panerythra from the non-continental portions of the western Pacific 1$ now recog- nized as Leporicypraea mappa viridis. The other taxon names remain the same. Sequences of L. mappa “rewa” from Pacific localities (Fiji, Vanuatu, Palau, and South China Sea) interfinger with haplotypes of L. mappa geographica individuals from Indian Ocean lo- calities (NW Australia, Phuket, Seychelles, and Zanzibar). Therefore, | recognize only a single taxon, L. mappa geographica, for this clade. Because of its conchological distinctiveness and sympatry with conspecifics, Lorenz (2002) elevated L. mappa geographica to specific sta- tus with Indian and Pacific subspecies. Based on the genetic difference between mappa-com- plex conspecifics and geographic overlap, spe- cific status is certainly acceptable. However, the remaining L. mappa subspecies are para- phyletic. The phylogeny Lorenz (2002: 27) pre- sents is correct and reflects this arrangement. Certainly, other recognized cowrie species are derived from paraphyletic parent species (e.g., Eclogavena coxeni and others; see Meyer, 2003: table 4, and cases herein), and L. geographica would have to be added to this list. These results suggest a third species sis- terto L. geographica should be recognized that would include both L. mappa viridis and L. mappa admirabilis. L. mappa geographica in- dividuals have been found sympatrically with both L. mappa mappa and L. mappa viridis in- dividuals in the Pacific Ocean. However, as yet, L. mappa mappa and L. mappa viridis haplo- types have not been found together. One new undescribed taxon is added to Mauritia. Haplotypes of M. arabica individuals from American Samoa cluster independently from haplotypes of M. arabica individuals from other Pacific localities. Shells from Samoan in- dividuals tend to be smaller, more heavily mar- gined and more circular than individuals from other Pacific localities. Results from increased sampling in both М. depressa depressa (М = 10) and M. depressa dispersa (N = 10) main- tain their independent, reciprocally monophyl- etic status, albeit recently diverged. As in previous findings, the interrelationships among major lineages in Mauritia are poorly supported. Consensus methods and poor support result in two polytomies (Fig. 4). Further genetic data will be needed to address this region of the phy- logeny as all extant taxa have been sampled. New sequence data from Barycypraea teulerei and B. fultoni place them as sister taxa and align them with the genus Zoila to form the group Bernayinae. Sequence data presented for Barycypraea fultoni are of B. fultoni amorimi from Mozambique. The Australian Zoila marginata complex is split into two ESUs as increased sampling indicates fixed molecular differences between populations separated by the Southwest Cape region between capes Naturaliste and Leeuwin. Further sampling di- rectly within this region may uncover interme- diate haplotypes that would link the two ESUs and suggest a cline instead of two independent lineages. Such a finding is the case in the Zoila friendii complex. However, as none have been discovered yet, | present the data as two tenta- tive ESUs: Zoila marginata marginata to the south and Z. marginata ketyana to the west. Other described Z. marginata taxa (Lorenz, 2001; 2002) within each ESU interfinger, and do not fulfill molecular criteria for recognition. Sequence data from Zoila mariellae are the fi- nal addition to the Bernayinae clade. While the exact provenance of the animal sequenced is unknown, it is likely from the northwestern shelf of Australia. Molecular results place Z. mariellae as a distinct sister to Z. decipiens, also from the northwestern shelf, as expected. Following along the phylogeny, the clade Luriinae comes next. Talparia and Luria are strongly supported as the clade Luriini. A small fragment from 16S was amplified from a de- graded Talparia exusta specimen, and as ex- pected, the taxon is sister to the more widespread Talparia talpa. Surprisingly, se- quence divergence between the two species appears to be relative small, indicating a more recent divergence than expected. Better-pre- served material from 7. exusta is needed be- fore these relative results can be confidently assessed. The inclusion of four new taxa to the Austrocypraeini (Arestoides argus contrastriata, Lyncina broderipii, L. ventriculus from the In- dian Ocean, and L. kuroharai) does not help in resolving interrelationships among member taxa. Arestoides argus is broken into a Pacific clade, A. argus argus, and a western Indian Ocean clade, A. argus contrastriata, based on additional sequence data from the Indian Ocean. Lyncina broderipii appears as sister to L. nivosa within the Callistocypraea clade, as INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE 137 predicted in Meyer (2003). A single sampled individual of L. ventriculus from Christmas Is- land in the Indian Ocean falls significantly out- side the haplotype cluster of individuals (N = 6) from various regions of the Pacific basin. Lyncina ventriculus is an oceanic taxon, and because of the geographic gap between sites across continental Southeast Asia, | choose to present the Christmas Island form as new, undescribed, distinct ESU. Further sampling of individuals from Christmas Island may change this interpretation, but they are currently lack- ing. A single sample of Lyncina kuroharai was sequenced and the results place it closely re- lated to L. sulcidentata, an endemic Hawaiian taxon. The shallow split between these two taxa indicates a relatively recent common ancestor. Faunal ties have been documented in other cowrie species between Hawaii and Japan, most notably in Luria isabella, and the close affinities between L. kuroharai and L. sulcidentata represent another example of this biogeographic link. The final two ESUs added within the mid1 compartment are members of the genus Pustularia, and more specifically are recognized subspecies of Pustularia bistrinotata. A single Р bistrinotata keelingensis individual was se- quenced, is distinct, and appears as sister to the remaining P. bistrinotata complex. Further- more, P. bistrinotata sublaevis individuals (N = 5) from southeast Polynesia (Tuamotu and Societies) cluster together, forming a third ESU within Р bistrinotata. Mid2 Compartment (Fig. 5) The third phylogenetic compartment, mid2, contains members from the genera Neobernaya, Pseudozonaria, Schilderia, Zonaria, the subfamily Cypraeovulinae, and the tribe Bistolidini of the subfamily Erroneinae. Interrelationships among member clades are consistent with previous findings (Meyer, 2003). Neobernaya and Pseudozonaria are sisters, and that clade is sister to the remaining cow- ries. The inclusion of sequence data from the genus Schilderia (S. achatidea), place the group as sister to Zonaria, and together this clade shares a more recent ancestor with the remaining taxa. The subfamily Cypraeovulinae includes both the South African Cypraeovula and South Australian Notocypraea and is sis- ter to the western IndoPacific Erroneinae, which is composed of two tribes: Bistolidini and Erroneini. Within the mid2 compartment, 25 taxa are added to the existing sequence database; at least one ESU is added within each genus ex- cept the monotypic Neobernaya. Pseudo- zonaria nigropunctata, a Galapagos endemic, falls into the eastern Pacific clade as a diver- gent sister to P arabicula, although not strongly supported. The position of Schilderia achatidea has been mentioned previously as sister to Zonaria, now found exclusively in the eastern Atlantic. Two taxa are added from Zonaria. Zonaria picta from the Cape Verde Islands falls near the base of Zonaria, and its relationship with other Zonarid taxa is ambiguous, resulting in a polytomy at the base of the group. Alterna- tive phylogenetic reconstructions at the base of the group show small internodes, indicative of a short radiative burst, with little divergence since. New sequence data from Pseudozonaria angelicae are extremely similar to haplotypes from Р pyrum (both Р pyrum angolensis and Р pyrum senegalensis). | include Р angelicae as a taxon in the phylogeny, but prefer to con- sider it at most a subspecies until further se- quence data are available within the Р. pyrum complex, as | have reservations concerning di- vergences along the mostly continuous West African/Mediterranean coastline. Sequence data from six additional taxa are included within Cypraeovulinae, two from Notocypraea and four from Cypraeovula. In Notocypraea, | tentatively recognize two ESUs within Notocypraea angustata, with a phyloge- netic break somewhere between Port Lincoln and Port Macdonnel, South Australia. Two di- vergent haplotype clusters exist without inter- mediate states. Again, further data may change this interpretation, but at present | chose to rep- resent these as different ESUs indicating dis- tinct evolutionary trajectories. Sequence data from a single specimen of Notocypraea hartsmithi, a rare species from southeastern Australia, indicate that the species is sister to all remaining Notocypraea taxa. Within Cypraeovula, four taxa are added, but their in- clusion does not change previous interpreta- tions that the group is composed of predominately four divergent lineages with mi- nor differences within each. New sequence data from both Cypraeovula fuscorubra and C. fuscodentata closely align these taxa with C. capensis. New sequence data from C. mikeharti and C. algoensis closely align those taxa with C. edentula and C. alfredensis. Noting the shal- low divergences among recognized species in Figure 5, | am doubtful that many of the de- scribed subspecies within Cypraeovula (sum- marized in Lorenz, 2002) will fulfill my molecular criteria for ESU status. As some species are differentiated currently by only a 138 Strict 67 3 95 14 98 25 55 6 72 6 79 8 3 79 15 73 6 1 3 3 3 76 7 00 21 80 5 3 00 38 99 10 100 21 100 20 96 12 5 1100 80 1100 MEYER globulus globulus globulus brevirostris cicercula margarita mauiensis bistrinotata keelingensis bistrinotata bistrinotata bistrinotata sublaevis _ spadicea annettae robertsi arabicula nigropunctata (29) lachatidea (30) | sanguinolenta picta (31) zonaria pyrum angelicae* (32, hartsmithi (33) piperita pulicaria comptoni declivis angustata (P. Lincoln) (34) angustata connelli castanea iutsui fuscorubra (35) capensis fuscodentata (36) coronata edentula alfredensis mikeharti* (37) lutea saulae lentiginosa исгас ziczac ziczac misella contaminata distans contaminata contaminata asellus bitaeniata asellus vespacea asellus asellus diluculum artuffeli clandestina candida cland. clandestina (39) clandestina passerina stolida stolida stolida clavicola stolida diagues (40) stolida rubiginosa owenii (41) erythraeensis goodallii hirundo ursellus (WP) ursellus (Andaman) (42) kieneri kieneri kien depriesteri A kien depriesteri B coloba chinensis amiges (43) chinensis chinensis teres pellucens subteres (44) Uatior (45) | gaskoini taitae* (46) catholicorum cumingii pellisserpentis (48) cribraria comma esontropia francescoi (49) esontropia esontropia fallax gaspardi cribraria cf. australiensis cribraria cf. abaliena (50) exmouthensis exmouthensis exmouth. magnifica (51) cribraria rottnestensis cribraria melwardi (52) cribraria cribraria cribraria abrolhensis (53) langfordi cavatoensis hirasei katsuae [MUSUMEA | serrulifera minoridens oryzaeformis hammondae microdon microdon microdon chrysalis gracilis gracilis gracilis notata fimbriata fimbriata fimbriata cf. unifasciata fimbriata marquesana Lfimbriata waikikiensis | Pustularia Neobernaya Pseudozonaria Schilderia Zonaria Notocypraea Cypraeovula Palmadusta Bistolida Ovatipsa Talostolida Cribrarula Austrasiatica Palmulacypraea Purpuradusta globulus globulus globulus brevirostria cicercula margarita mauiensis bistrinotata keelingensis bistrinotata bistrinotata bistrinotata sublaevis ra spadicea annettae robertsi arabicula nigropunctata achatidea zonaria pyrum angelicae* sanguinolenta picta hartsmithi piperita pulicaria comptoni declivis angustata (Port Lincoln) angustata angustata connelli castanea iutsui fuscorubra capensis fuscodentata coronata edentula alfredensis mikeharti* algoensis contaminata distans contaminata contaminata asellus bitaeniata asellus vespacea asellus asellus saulae lutea lentiginosa ziczac ziczac ziczac misella diluculum artuffeli clandestina candida clandestina clandestina clandestina passerina stolida stolida stolida clavicola stolida diagues stolida rubiginosa owenii vasta erythraeensis goodallii hirundo ursellus (WP) ursellus (Andaman) kieneri kieneri Кеп depriesteri А kien depriesteri В coloba chinensis amiges chinensis chinensis teres pellucens subteres latior gaskoini taitae* catholicorum astaryi garciai* cumingii pellisserpentis cribraria comma esontropia francescoi esontropia esontropia fallax cribraria australiensis cribraria cf. abaliena gaspardi exmouthensis exmouthensis exmouthensis magnifica cribraria rottnestensis melwardi cribraria cribraria cribraria abrolhensis langfordi cavatoensis hirasei sakuraii katsuae musumea serrulifera minoridens oryzaeformis hammondae microdon microdon microdon chrysalis gracilis gracilis gracilis notata fimbriata fimbriata fimbriata cf. unifasciata fimbriata marquesana fimbriata waikikiensis 0.05 substitutions/site FIG. 5. Mid 2 Compartment cladogram and phylogram. All other information as in Fig. 3. INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE 139 single mutation (e.g., Cypraeovula mikeharti/ С. algoensis or С. castanea/C. iutsui), there simply is not enough room for differences to have accumulated between taxa. This is not to say that described entities are not indepen- dent. Indeed, because Cypraeovula taxa are direct developers with limited dispersal and gene flow, regional differences are expected on small geographic scales, much like the South Australian endemic clades Umbilia, Zoila, and Notocypraea. However, based on the genetic similarity among sampled member Cypraeovula, much of this variation has to be very recently derived. This pattern is borne out in the South Australian direct developers that have been more extensively sampled. The tribe Bistolidini within Erroneinae is com- posed of members from five genera: Palmadusta, Bistolida, Ovatipsa, Talostolida and Cribrarula. As in Meyer (2003), the basal root of Bistolidini is poorly resolved. Overall analyses place either Palmadusta as sister to the other four genera or Palmadusta and Bistolida as a clade, sister to the remaining three. Compartmentalized analyses place Palmadusta at the base, although poorly sup- ported. The addition of 15 ESUs did not help in resolving this issue. Only one taxon is added to the Palmadusta clade, but it alters the sub- specific designations previously ascribed (Meyer, 2003). New haplotypes from Andaman Sea P clandestina individuals form a distinct monophyletic clade. This new ESU is sister to the western Indian Ocean P. clandestina passerina, and the two of them are sister to the Pacific P clandestina clade and the Japanese endemic P. artuffeli. Based on a review of P clandestina subspecies and type localities, the Pacific clade that | had formerly (Meyer, 2003) recognized as P clandestina clandestina should be P clandestina candida, and the new Р. clandestina clade from the Andaman Sea now bears the name P. clandestina clandestina. | also reviewed the subspecies and type locali- ties for the three Р asellus ESUs previously unnamed (Meyer, 2003). Based on increased sampling and conchological comparisons, | ten- tatively ascribe the following subspecific des- ignations for the three clades: P asellus asellus for the western Indian Ocean clade, P. asellus vespacea for the Seychelles to western Pacific clade, and P. asellus bitaeniata for the Melanesian and Pacific clade (Fig. 5, Appen- dix). Unfortunately, the addition of P clandestina clandestina does not help in resolving the basal nodes of Palmadusta. As shown in Figure 5, the base of Palmadusta is poorly resolved and sister group assignments are ambiguous. Afew lineages remain strongly supported (Р asellus, P. clandestina/diluculum, P. ziczac and P. contaminata), but confident hypotheses of other interrelationships require further data. Three taxa are added to Bistolida: B. stolida diagues, В. owenii and an undescribed, dis- tinct eastern Indian Ocean clade of B. ursellus. Individuals of B. stolida diagues from the Seychelles fall as sister to B. stolida rubiginosa. Bistolida owenii, a western Indian Ocean taxon, is sister to the Red Sea endemic В. erythraeensis. À new Bistolida ursellus se- quence from the Andaman Sea is poorly sup- ported as sister to the remaining B. ursellus taxon from the Pacific basin. Its placement is equally parsimonious as either sister to B. ursellus (Pacific) or forming a B. ursellus grade leading to the B. kieneri lineage. The topology of the two В. ursellus taxa as sisters is more likely and consistent with morphology. One taxon is added to Ovatipsa and two taxa to Talostolida. Within Ovatipsa, the subspe- cies O. chinensis amiges from the Pacific ba- sin and Western Australia is distinct from O. chinensis chinensis from the Philippines west- ward through the Indian Ocean to the east coast of Africa. Various other O. chinensis subspecies have been described within the Indian Ocean (e.g., Lorenz & Hubert, 1993), and preliminary data indicate that these Indian Ocean subspecies may represent very recent divergences within what | am currently recog- nizing as O. chinensis chinensis. However, until more individuals are sampled, | maintain them all under the taxon Ovatipsa chinensis chinensis. Within Talostolida, two taxa are added that appear as sisters to each other: Т. subteres from southeastern Polynesia and 7. latior from Hawaii. These two taxa are sister to Talostolida pellucens. All four taxa currently included within Talostolida are deeply divergent independent ESUs. A single haplotype of Talostolida teres “alveolus” (sensu Lorenz, 2002) is completely identical to haplotypes of T. teres teres individuals from both the Society Islands and the Tuamotu. Moreover, 7. teres individuals from SE Polynesia have been de- scribed by Lorenz (2002) as a distinct subspe- cies T. teres “апае”; however sampled individuals of Т. teres from SE Polynesia interfinger with individuals sampled from the Western Pacific (Papua New Guinea and Guam). Therefore, the data do not support Т. teres “janae” as a valid taxon, based on my criteria. All Marquesan individuals sequenced possess T. pellucens haplotypes, whereas all T. teres-like individuals from the remainder of SE Polynesia possess T. teres haplotypes. 140 MEYER The Cribrarula clade includes eight additional taxa, making it the most diverse genus within Bistolidini. Two taxa, Cribrarula taitae from American Samoa and C. garciai from Easter Island, are added to the deeply divergent Pa- cific subclade. Both taxa are recently divergent members from their respective sister taxon. Cribrarula taitae appears as a closely related sister to С. catholicorum, and С. garciai is closely related to С. cumingii. Only a single in- dividual from each of the two taxa was included in these analyses, and the results would be better addressed with multiple samples. Two members are added to the Western Indian Ocean subclade: Cribrarula pellisserpentis and С. esontropia francescoi, both from Madagas- car. Cribrarula esontropia francescoi is a closely related sister to С. esontropia esontropia, which includes С. esontropia cribellum (Meyer, 2003). Cribrarula pellisserpentis is a deeply divergent member within the western Indian Ocean subclade and is sister to the other three ESUs. Four taxa are added to the remaining Cribrarula member clade. A single individual of C. cribraria from Masirah, Oman, appears significantly di- vergent from population samples of the previ- ously unnamed C. cribraria ESU from the Andaman Sea. Conchologically, this individual approximates the western Indian Ocean taxon С. cribraria abaliena and 1$ tentatively recog- nized as such. A single individual of C. cribraria australiensis from Western Australia falls within the Andaman С. cribraria cluster; therefore, | tentatively adopt the name C. cribraria cf. “australiensis” for a taxon that extends from the Andaman Sea southward to Western Austra- lia. More exhaustive sampling is required to confirm these geographic patterns. A single in- dividual of C. exmouthensis magnifica from Broome 1$ significantly different from samples of C. exmouthensis exmouthensis from the Exmouth Gulf region, therefore validating the status of that taxon. Additional samples of C. cribraria rottnestensis (N = 3) further validate the taxon's uniqueness. Eight individuals of C. melwardi from northeastern Australia all share a common ancestor and are reciprocally mono- phyletic with respect to the remaining C. cribraria individuals. Moreover, a single C. cribraria cribraria individual from the same reef (Lamont Reef in the Bunker Group) clusters as expected with other Pacific C. cribraria cribraria individuals. The final taxon included is C. cribraria abrolhensis (N = 3), and haplotypes are shallowly divergent but reciprocally mono- phyletic with respect to samples of C. cribraria cribraria (N = 30) from predominately western Pacific localities (Appendix). More thorough analyses and discussion of this fascinating, species-rich group is in preparation (Meyer et al., in prep.). Derived Compartment (Fig. 6) The final compartment analyzed is the derived monophyletic clade recognized as the tribe Erroneini. This clade includes the following nine genera: Austrasiatica, Palmulacypraea, Erronea, Purpuradusta, Contradusta, Nota- dusta, Eclogavena, Melicerona and Blasicrura. Many (25) taxa are added within the tribe, and phylogenetic analyses result in some surpris- ing affinities. For the most part, major genera are well supported, but their interrelationships are not. Three taxa currently ascribed to Austrasiatica were included in previous analy- ses (Meyer, 2003); however, they were consid- ered as representatives of the genus Nesiocypraea. As discussed earlier, the find- ing that Nesiocypraea teramachii is distantly related raises the subgenus Austrasiatica to generic status for the clade that includes Austrasiatica langfordi, А. hirasei and А. sakurai. As in Meyer (2003), Austrasiatica 1$ sister to all other Erroneini taxa, followed by Pamulacypraea as sister to the remainder. As predicted in Meyer (2003), the newly added Pamulacypraea musumea falls as sister to P katsuae. Even with the addition of 24 taxa (a 67% increase), the topology among the rest of the major Erroneini lineages 1$ ambiguous. Six added “Erronea” species form a basal grade leading to the Adusta/Erronea split previously recognized in Meyer (2003). | take a conserva- tive approach and redefine Erronea to include all these taxa and subsume Adusta to a well- supported subclade within the group, as the new data demonstrate that Adusta and Erronea (including the more recent additions) are not equivalent (sisters). If Adusta were to be main- tained at equivalent generic status, Erronea would represent a paraphyletic group. Purpuradusta, Eclogavena, Melicerona and Blasicrura are all well-supported monophyletic lineages. As in Meyer (2003), Notadusta is well supported only if restricted to members of the Notadusta punctata complex. However, be- cause Notadusta martini is often considered a member of Notadusta, | include it within Notadusta here, although poorly supported. In a similarly conservative manner, | include two of the added taxa within Contradusta, although again poorly supported. Support for relation- ships among these seven genera is poor and is likely because of the short internode length between divergent lineages. INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE Strict 100 51 7 73 3 176 3 1 59 7 95 9 78 2 199 99 17 98 13 99 17 100 84 23 71 1 3 80 8 51 3 100 log 5 6 | зв 1 5 [97 59 1 100 81 14 9 6 5 193 9 91 4 98 64 78 2 1 1 75 62 = 100 66 2 95 94 100 a 7 97 98 = 21 | 100 5] 84 2 192 100 = 16 1 ! 73 7 100 99 6 8 | 63 100 1 0 82 100 8 | 99 1 100 100 0 1 94 5 1181 4 152 1 goodallii hirundo stolida stolida stolida clavicola stolida diagues stolida rubiginosa owenii vasta erythraeensis ursellus ursellus (Andaman) kieneri kieneri kien depriesteri A kien depriesteri B langfordi cavatoensis hirasei sakuraii 2] katsuae musumea (54) xanthodon (55) pallida (56) vredenburgi (57) rabaulensis (58) fernandoi (59) onyx adusta subviridis subviridis onyx melanesiae subviridis dorsalis pyriformis (60) cylindrica lenella (61) cylindrica cylindrica ovum ovum (62) ovum palauensis caurica elongata (63) caurica draceana caurica spp. 1 (64) caurica quinquefasciata caurica spp. 2 (65) caurica derosa errones ovum chrysostoma caurica samoensis (66) caurica caurica serrulifera minoridens oryzaeformis (67) hammondae microdon microdon microdon chrysalis (68) gracilis gracilis gracilis notata fimbriata fimbriata fimbriata cf. unifasciata fimbriata marquesana (69) Lfimbriata waikikiensis (70) | walkeri bregeriana barclayi (71) | pulchella (72) martini hungerfordi (73) punctata berinii B punctata berinii A punctata (Andaman) punctata trizonata (74) RES punctata dayritiana quadrimaculata thielei coxeni quad. quadrimaculata listeri melvilli (75) listeri felina (76) interrupta pallidula pallidula pallidula rhinoceros pallidula cf. vivia (77) summersi (78) Bistolida Austrasiatica Palmulacypraea Erronea Purpuradusta Contradusta Notadusta Eclogavena Melicerona Blasicrura 0.05 substitutions/site 141 stolida stolida stolida clavicola stolida diagues stolida rubiginosa owenii vasta erythraeensis goodallii hirundo ursellus ursellus (Andaman) kieneri kieneri kien depriesteri A kien depriesteri B langfordi cavatoensis hirasei sakuraii katsuae musumea xanthodon pallida vredenburgi rabaulensis fernandoi onyx adusta subviridis subviridis onyx melanesiae subviridis dorsalis pyriformis cylindrica lenella cylindrica cylindrica ovum ovum ovum palauensis caurica elongata caurica draceana caurica spp. 1 caurica quinquefasciata caurica spp. 2 caurica derosa errones ovum chrysostoma caurica samoensis caurica caurica serrulifera minoridens oryzaeformis hammondae micr. microdon micr. chrysalis gracilis gracilis gracilis notata fimbriata fimbriata fimbriata cf. unifasciata fimbriata marquesana fimbriata waikikiensis walkeri bregeriana barclayi pulchella martini hungerfordi punctata berinii A punctata berinii B punctata (Andaman) punctata trizonata punctata punctata dayritiana quadrimaculata thielei coxeni quadrimaculata quadrimaculata listeri melvilli listeri listeri felina interrupta pallidula pallidula pallidula rhinoceros pallidula cf. vivia summersi FIG. 6. Derived Compartment cladogram and phylogram. All other information as in Fig. 3. 142 MEYER Twelve additional taxa are added to Етопеа. Six of the additions are traditionally recognized as distinct species, four have been recognized as subspecies, and two are newly discovered, but may have names associated with them that have been placed into synonymy. Of the new species, three form a relatively well-supported clade: Erronea rabaulensis shares a more re- cent common ancestor with E. fernandoi (80/1), and those two are sister to Е. vredenburgi (84/ 3). The three additional Erronea species all nest deeply within the clade, and their relationships are not well supported. Erronea pallida appears as sister to the clade of the previously described three species and Adusta. Erronea pyriformis is relatively well supported (81/6) as the sister to the clade previously recognized as Erronea (Meyer, 2003). Finally, Erronea xanthodon falls at the base of Erronea and is sister to all other Erronea taxa. Within the crown Erronea subclade, six taxa are added that are all tradi- tionally recognized at the subspecific level. In- dividuals of Erronea cylindrica lenella (N = 8, all from New Caledonia) form a monophyletic group strongly supported (91/6) as sister to the clade including the remaining E. cylindrica in- dividuals plus two subspecies of Е. ovum. These results imply that E. cylindrica at the specific level is a paraphyletic taxon. Newly added individuals of Erronea ovum ovum from both Singapore and the Philippines (N = 15) form a monophyletic group sister to Е. ovum palauensis (N = 7). The four remaining, newly added taxa are all members of the Erronea caurica complex. First, individuals (N = 7) of the newly described E. caurica samoensis ap- pear as a distinct lineage sister to individuals (N = 15) from the remainder of the Pacific and Western Australia (E. caurica caurica). Four geographically structured haplotype clades are found exclusively in the Western Indian Ocean. Erronea caurica dracaena is Currently restricted to the Seychelles based on sampling. Newly added individuals from East Africa and Mada- gascar form a haplotype clade that | recognize as Erronea caurica elongata. Individuals of E. caurica quinquefasciata from the Red Sea, East Africa and Oman form the third monophyletic group. Finally, newly sequenced individuals from Masirah (N = 7) form a private haplotype clade (E. caurica ssp. #1) sister to E. caurica quinquefasciata. The final, newly added taxon (E. caurica ssp. #2) within the E. caurica com- plex is a clade (N = 18) that includes individu- als primarily from India, but with a few individuals from Masirah, Oman. This haplo- type саде 1$ sister to the clade recognized pre- viously as E. caurica cf. derosa from the Andaman Sea (Meyer, 2003). The Erronea caurica complex and the associated E. cylindrica, E. ovum and E. errones species will be more thoroughly addressed in another pa- per (Meyer, in prep.) as the group exhibits re- markable geographic structuring, polyphyly of recognized species (E. ovum), and evidence of introgression based on nuclear markers. Purpuradusta is well supported and contains four newly added taxa that fall in expected re- lationships. The southeastern Polynesian en- demic species Purpuradusta oryzaeformis is distinct and sister to P. minoridens that ranges throughout the remainder of the western IndoPacific. A single specimen of P. microdon from East Africa falls outside the haplotype clade of other sampled individuals from the Pacific basin (N = 5). This East African popula- tion is recognized as Purpuradusta microdon chrysalis. Two peripheral populations of Purpuradusta fimbriata in the Pacific Basin are introduced. First, Hawaiian populations of P. fimbriata are distinct (N = 7) and were previ- ously recognized as P. fimbriata waikikiensis; thus this name 1$ resurrected as a valid entity. Second, individuals from the Marquesas are also distinct genetically, consistent with the subspecies designation of Lorenz (2002), P fimbriata marquesana (N = 14). Both of these Pacific P. fimbriata subclades share a more recent history with the widespread Pacific sub- species Р fimbriata unifasciata, as expected. Two newly added species, “Erronea” barclayi and “Erronea” pulchella, come out as sister species in phylogenetic analyses. Moreover, these two taxa appear as sister to Contradusta in the most likely topology. Because of these results, and the poorly supported nature о their relationships, | tentatively place the two taxa in the genus Contradusta, with the caveat that they may be removed with future data. These results are somewhat surprising, particularly because “Contradusta” pulchella is thought to be closely related to Erronea pyriformis be- cause of the darkly stained columellar denti- tion and overall conchological similarities. The sister relationship between Contradusta pulchella and С. barclayi is more acceptable as their divergence is deep, and the phyloge- netic affiliations of С. barclayi were more diffi- cult to predict based on morphological criteria. Another surprising result is the sister relation- ship between Notadusta martini and “Erronea” hungerfordi. Given these phylogenetic results, | tentatively place “Erronea” hungerfordi within Notadusta, but with little confidence, although it is reasonably supported (73/4), and suspect that it may be removed with more samples and INCREASED MOLECULAR SAMPLING IN CYPRAEIDAE 143 sequence data. Within the remaining Notadusta complex, individuals of N. punctata trizonata (N = 9) form a monophyletic group sister to the Pacific N. punctata punctata clade. Finally, in regards to Notadusta, “Notadusta” rabaulensis was mentioned previously as a member of Erronea and “Notadusta” musumea as Palmulacypraea, further reducing the member- ship of Notadusta (Meyer, 2003). The final four additions to the dataset fall into Melicerona and Blasicrura. First, two taxa are added to Melicerona. Samples of Melicerona listeri melvilli (N = 5) from Queensland, Austra- lia, form a monophyletic group sister to the re- maining Melicerona taxa. (Two rostrate and melanistic individuals interfinger among the other three haplotypes indicating that the tera- tology is likely driven by phenotypic responses to environmental conditions rather than having a genetic basis.) Samples of Melicerona felina from both Oman and East Africa form a mono- phyletic group, and because the haplotypes from the two regions interfinger, there is no evidence for a distinction between the subspe- cies M. felina felina and M. felina fabula. Within Blasicrura, two taxa are added, based on the sequencing results. First, samples of Blasicrura summersi, a Fijian and Tonga endemic, appear as a recently divergent sister to the also newly included B. pallidula cf. vivia from American Samoa. This clade is sister to the Melanesian subspecies Blasicrura pallidula rhinoceros, as expected based on geography. This resulting topology indicates that the Blasicrura pallidula complex is paraphyletic. DISCUSSION The ultimate goal of this project is to construct a comprehensive phylogeny of cypraeid gas- tropods at the appropriate level for diversifica- tion studies. From a molecular perspective, all ESUS presented are effectively equal units of diversity, whether they are currently recognized as species, subspecies or some other level. There are some noted exceptions as OTUs were used on occasion that represented un- sorted or clinal variation within an ESU (e.g., Erosaria miliaris/eburnea). However, on a gen- eral scale, each taxon shown in the phylogenies (Figs. 3-6) represents an independent evolu- tionary trajectory. Because so much taxonomic information is available for cowries, it is informative to see how molecular criteria compare with recog- nized taxonomic entities. The most recent com- pilation of the cowries is that of Lorenz (2002), and | will use his checklist (рр. 250-291) аз a benchmark for comparisons. Lorenz recog- nizes 232 species, of which | have sequenced 210 (> 90%), and they are presented herein. The missing species are as follows: Nesiocypraea aenigma, N. lisetae, N. midway- ensis, Austrasiatica alexhuberti, Erosaria ostergaardi, Zoila perlae, Lyncina camelopar- dis, L. joycae, Pustularia chiapponii, Cypra- eovula colligata, C. cruickshanki, C. immelmani, Palmadusta androyensis, P. johnsonorum, Austrasiatica deforgesi, Palmulacypraea boucheti, Р omii, Есодауепа luchuana, Erronea (?) angioyorum, and E. nymphae. Se- quences from samples of both Purpuradusta barbieri and “Talostolida” rashleighana have been obtained, but were too late for inclusion in these analyses. All missing species are rare, with small ranges located generally at the pe- riphery of their putative sister species based on conchological and anatomical characters. Of the 210 sequenced species, phylogenetic com- parisons and molecular criteria support all but 15 (93%) as ESUs. The 15 recognized species not supported by my criteria are discussed be- low. For Nucleolaria granulata, Monetaria obvelata, Erosaria eburnea, Zoila orientalis, 2. thersites, Luria controversa, L. gilvella, Notocypraea occidentalis, and Palmadusta humphreysii, multiple individuals were se- quenced and the haplotypes interfingered within their closest relative. For the next six species that | do not support, only a single in- dividual was sequenced, thus they may indeed represent a very young independent trajectory. However, when compared to the genetic diver- sity within their closest relative, the genetic dif- ference is unremarkable, and in some instances, only a single mutation different from putative conspecifics: Zonaria angelicae, Z. petitiana, Cypraeovula mikeharti, Bistolida brevidentata, Cribrarula garciai, and C. taitae. While genetic data are overall broadly con- sistent with taxa recognized at the specific level, the results are even more remarkable when compared among taxa recognized at subspe- cific levels. Lorenz recognizes 260 taxa at the зибзресйс level. Of those 260 subspecies, | have sequenced at least two individuals from 160 in order to assess their validity. Molecular criteria support 113 (> 70%) of these taxa as legitimate ESUs. Moreover, sequence results indicate an additional 20 distinct ESUs not recognized as subspecies by Lorenz (but sometimes mentioned as important varieties or forms). A full listing of sampled taxa and their current ESU status as indicated by the prior criteria can be found at the Cowrie Ge- 144 МЕУЕК netic Database Project Website (http:// www.flmnh.ufl.edu/cowries). The website in- cludes other information, such as localities sampled, numbers of individuals for each taxon, and photographs of the specimens sequenced. Overwhelming molecular support for tradition- ally recognized taxa, both at specific and sub- specific levels, is extremely encouraging. First, from a taxonomic standpoint, these molecular results corroborate the excellent work done by centuries of malacological researchers, at both professional and amateur levels. Similar mo- lecular surveys of other diverse groups will pro- vide valuable comparisons in order to assess taxonomic congruence (e.g., Jackson & Cheetham, 1990) and address concordant di- versification patterns. Second, from a molecu- lar perspective, sequence data provide a suitable, objective, relative metric for circum- scribing appropriate evolutionary units. Assum- ing rate constancy in the molecules (COI only, in prep.), molecular divergences can constrain the tempo of diversification and assess the dis- tinctiveness of purported taxa. A growing body of molecular data across the diversity of life undoubtedly will provide insight to some of our most fundamental evolutionary questions. ACKNOWLEDGEMENTS An ever-growing number of individual and institutions have contributed and supported this ongoing research. Without their assistance, the work would not be possible. The following per- sons are recognized: Nonoy Alonzo, Vicente Azurin, Paul Barber, Don Barclay, Marty Beals, Victor Bonito, Philippe Bouchet, Michel Boutet, Roy Caldwell, Carlos Carvalho, Hank Chaney, John Chester, Peter Clarkson, Lori Bell Colin, Pat Colin, Allen Collins, Harry Conley, Vince Crayssac, Carolyn Cruz, Donald Dan, Martyn Day, Bruno de Bruin, Helen deJode, John Earle, Andrew Edinger, Mark Erdmann, Melissa Frey, Michel Garcia, Bill Gibbs, Serge Gofas, Terry Gosliner, Jeroen Goud, Robert Gourguet, Fabien Goutal, Paulo Granja, Kibata Mussa Haji, Jerry Harasewych, ltaru Hayami, Brian Hayes, Claus Hedegaard, Ed Heiman, Bert Hoeksema, John Hoover, John Jackson, Maurice Jay, Scott Johnson, Paul Kanner, Yasunori Kano, Tomoki Kase, Norbert Kayombo, Shigemitsu Kinjo, Lisa Kirkendale, Kitona Kombo Kitona, Utih Kukun, Senthil Kumar, Jean Paul Lefort, Bill Liltved, Hung-Chang Liu, Charlotte Lloyd, Felix Lorenz, Jr., Felix Lorenz, Sr., Larry Madrigal, Marlene Martinez, Gerald McCormack, Mohammed Mohammed, Hugh Morrison, Gowele Mtoka, Mtumwa Mwadini, Peter Ng, Steve Norby, Shuichi Ohashi, Yoshihiro Omi, Ina Park, Marcel Pin, Cory Pittman, Xavier Pochon, Matt Rich- mond, Raphael Ritson-Williams, Goncalo Rosa, Gary Rosenberg, Teina Rongo, Fred Schroeder, Mike Severns, Pauline Severns, Hung-Long Shi, Brian Simison, Michael Small, John Starmer, Steve Tettlebach, David Touitou, Martin Wallace, Chia-Hsiang Wang, Dave Watts, Barry Wilson, Woody Woodman, Shu-Ho Wu. The following institutions are acknowledged: Florida Museum of Natural History; University of California Mu- seum of Paleontology; Academy of Natural Sci- ences of Philadelphia; Bernice P. Bishop Museum, Honolulu, Hawaii; California Academy of Sciences; Institute of Marine Sciences, Zan- zibar; University of Dar es Salaam; Jackson- ville Shell Club; Musée National d'Histoire Naturelle, Paris, France; National Museum of Natural History Naturalis, Leiden, The Nether- lands; Santa Barbara Museum of Natural His- tory; National Museum of Natural History; and Suganthi Devadason Marine Research Institute. | also would like to thank Felix Lorenz, Jr., for his thoughtful comments, as well as the reviews of four anonymous reviewers. Final decisions and opinions are wholly mine. This research has been financially supported by the following NSF grants: DEB-9807316, DEB 0196049, and OCE-0221382. LITERATURE CITED BRADNER, H. 8 E. A. 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Kamenev The Institute of Marine Biology, Russian Academy of Science, Vladivostok 690041, Russia; kamenev@mail333.com, inmarbio@mail.primorye.ru ABSTRACT A new species, Abrina scarlatoi, is described from the Commander and Kuril islands. This species has a small (to 11.2 mm), ovate-trigonal, high, almost equilateral shell with a non-polished, gray or light brown periostracum and conspicuous growth lines. The external ligament is attached to a short, wide nymph. The internal ligament is lodged in an ovate- elongate resilifer, which extends obliquely posterior to the beaks. Abrina scarlatoi was found in shelf zones of the Commander Islands (depth 3-100 m) and Kuril Islands (intertidal zone to 120 m), on rocky platforms and boulders, covered by a thick layer of lime red algae, brown algae, and sponges, with a population density up to 30 specimens/m?. The taxonomic status of Abrina magna Scarlato, 1965, and A. hainanensis Scarlato, 1965, is also discussed. Key words: Abrina, Semelidae, Bivalvia, Commander and Kuril islands. INTRODUCTION Previously, four species of the genus Abrina — A. сиперуда Scarlato, 1981; A. sachalinica Scarlato, 1981; A. shiashkotanika Scarlato, 1981; and A. tatarica Scarlato, 1981 — have been listed in Russian fauna (Scarlato, 1981). However, examination has shown that they are species of Macoma Leach, 1819 (Tellinidae) (Kamenev & Nadtochy, 1999). Study of the bivalve fauna of the Commander Islands shelf revealed an unknown species tentatively assigned to the genus Abrina Habe, 1952 (Kamenev, 1995; Bujanovsky, 1997). Detailed examination of the material from the Commander Islands and additional specimens from the Kurils has led me to regard it as a new species of Abrina. MATERIAL AND METHODS In this study, | have used the material col- lected by the IMB intertidal expedition to the Kuril Islands (June—July, 1967), joint IMB-PRIFO expeditions to the Commander Islands (8-28 July 1972, sealer “Krylatka”; 30 August-6 Octo- ber 1973, R/V “Rakytnoe”) and the Kuril Islands (July-November 1987, R/V “Tikhookeansky”), 157 and joint IMB - PIBOC expedition to Sakhalin Island and the Kuril Islands (1 July-4 August 2003, R/V “Akademik Орапп”). For comparison purposes, collections of the following taxa were used: Abrina lunella (Gould, 1861) (NSMT); A. kinoshitai (Kuroda & Habe, 1958) (NSMT, NSMI); A. declivis (Sowerby, 1868) (SBMNH); A. magna Scarlato, 1965, and A. hainanensis Scarlato, 1965 (both ZIN), and different species of other genera of the Semelidae (UW, CAS, USNM). Abrina declivis was stored in 70% ethanol. All other materials were stored dry. Shell Measurements Figure 1 shows the position of the shell mor- phology measurements. Shell length (L), height (H), width of each valve (W) not shown, anterior end length (A), maximal distance from posterior shell margin to top of pallial sinus (L1), and minimal distance from top pallial si- nus (L2) to anterior adductor muscle scar (L2) were measured for each valve. The ratios of these parameters to shell length (H/L, W/L, A/ L, L1/L, L2/L, respectively) were determined. Shell measurements were made using an ocu- lar micrometer with an accuracy of 0.1 mm. | made measurements of 34 specimens of the new species. 158 KAMENEV FIG. 1. Placement of shell measurements: L - shell length; Н - height; А - anterior end length; L1 - maximal distance from posterior shell margin to top of pallial sinus; L2 - minimal distance from top of pallial sinus to anterior adductor muscle scar. Abbreviations The following abbreviations are used in the paper: CAS — California Academy of Sciences, San Francisco; IMB - Institute of Marine Biol- ogy, Russian Academy of Sciences, Vladivostok; MIMB — Museum of the Institute of Marine Biology, Vladivostok; NHMI — Natu- ral History Museum and Institute, Chiba; NSMT — National Science Museum, Tokyo; PIBOC - Pacific Institute of Bioorganic Chem- istry, Russian Academy of Sciences, Vladivostok; PRIFO — Pacific Research Insti- tute of Fisheries and Oceanography, Vladivostok; SBMNH - Santa Barbara Mu- seum of Natural History, Santa Barbara; USNM - United States National Museum of Natural History, Smithsonian Institute, Wash- ington, D.C.; UW — University of Washington, Seattle; ZIN — Zoological Institute, Russian Academy of Sciences, St. Petersburg. SYSTEMATICS Family Semelidae Stoliczka, 1870 Genus Abrina Habe, 1952 Type species: Abra kanamarui Kuroda, 1951; = Macoma lunella Gould, 1861 Diagnosis Shell small (< 20 mm), thin to medium in thickness, moderately inflated, subtrigonal, ovate-trigonal or ovate, white, equivalve or with right valve sometimes more inflated, equilateral to longer anteriorly. Posterior end attenuate, with radial ridge along postero- dorsal margin, sometimes flexed to right. Periostracum thin, adherent or dehiscent, silky to dull, colorless, tan, gray, light brown. Surface with faint or conspicuous growth lines. Beaks orthogyrate, central or posterior. Hinge weak, two cardinal teeth in each valve; lateral teeth absent. Ligament opisthodetic, parivincular, both external and internal; ex- ternal seated on a nymph not projecting above dorsal margin; internal lodged in ob- lique resilifer posterior to cardinal teeth. Pal- lial sinus long, sometimes slightly different length and form in each valve, partly confluent with pallial line. Abrina scarlatoi Kamenev, new species Figs. 2-19, Table 1 Type Material and Locality Holotype (MIMB 9529), Polovina Bight, Bering Island, Commander Islands, Bering Sea, 3 m, rocky platform, bottom water tem- perature of 8.0°C, Coll. V. N. Romanov, 26- VII-1972 (sealer “Krylatka”); paratypes (30): paratypes (2) (MIMB 9530) from the holotype locality; paratypes (5) (MIMB 9531), Tonky Cape, Bering Island, Commander Islands, Bering Sea, 10 m, rocky platform, bottom wa- ter temperature of 9.1°C, Coll. $. D. Vavilin, 13-1Х-1973 (R/V “Rakitnoye”); paratype (ММВ 9532), Kamni Bobrovye — Kitolovnaya Bed, Medny Island, Commander Islands, Bering Sea (54`58.0’М, 167'21.5'E), 100 m, rocky plat- form, Coll. V. I. Lukin, 18-1X-1973 (R/V “Rakitnoye”); paratypes (2) (MIMB 9533), Cherny Cape, Medny Island, Commander Is- lands, Bering Sea, 15 m, rocky platform, bot- tom water temperature of 9.4°C, Coll. V. 1. Lukin, 17-1X-1973 (R/V “Rakitnoye”); paratypes (2) (MIMB 9534), Palata Cape, Medny Island, Commander Islands, Pacific Ocean, 20 m, rocky platform, bottom water temperature of 5.0°C, Coll. V. I. Lukin, 16-VII- 1972 (sealer “Krylatka”); paratype (MIMB 9535), Sivuchy Kamen, Medny Island, Bering Sea, 10 m, boulders, bottom water tempera- NEW SPECIES OF THE BIVALVE GENUS ABRINA 159 ture of 8.6 C, Coll. V. I. Lukin, 2-X-1973 (R/V ter temperature of 9.0 C, Coll. С. T. Belokonev, “Rakytnoe”); paratypes (3) (MIMB 9536), Tonky 13-1Х-1973 (R/V “Rakytnoye”); paratypes (2) Cape, Bering Island, Commander Islands, (ММВ 9537), Vodopadskogo Cape, Medny Is- Bering Sea, 20 m, rocky platform, bottom wa- land, Commander Islands, Pacific Ocean FIGS. 2-13. Abrina scarlatoi Kamenev, new species. FIGS. 2-5: Holotype (ММВ 9529), Polovina Bight, Bering Island, Commander Islands, Bering Sea, 3 m, shell length 9.8 mm. FIGS. 6, 7: Paratype (MIMB 9538), Phedoskina Cape, Bering Island, Commander Islands, Pacific Ocean, 5 m, right and left valves of a young specimen. FIGS. 8, 9: Paratype (MIMB 9531), Tonky Cape, Bering Island, Commander Islands, Bering Sea, 10 m, right and left valves with ligament. FIG. 10: Paratype (MIMB 9530), from holotype locality, right valve without ligament. FIG. 11: Paratype (MIMB 9534), Palata Cape, Medny Island, Commander Islands, Pacific Ocean, 20 m, left valve without ligament. FIG. 12: Paratype (MIMB 9533), Cherny Cape, 15 m, Medny Island, Commander Islands, Bering Sea, right valve without ligament. FIG. 13: MIMB 9549, Nadezda Strait (Rashua Island - Matua Island), Kuril Islands, 48°00’N, 153°15’E, 50 m. Bar = 1 mm. 160 KAMENEV (54'38.6’М, 167 43.5'E), 40 т, rocky platform, Coll. V. |. Lukin, 3-X-1973 (R/V “Rakytnoe”); paratypes (3) (MIMB 9538), Phedoskina Cape, Bering Island, Commander Islands, Pacific Ocean, 5-15 m, rocky platform, bottom water temperature of 9.8-10.0 C, Coll. V. I. Lukin, 23- IX-1973 (R/V “Rakytnoe”); paratypes (3) (ММВ 9539), Peregrebnogo Cape, Bering Island, Commander Islands, Bering Sea, 15-20 m, rocky platform, bottom water temperature of 10.0°C, Coll. B.l. Sirenko, 5-1Х-1973 (R/V “Rakytnoe”); paratype (ММВ 9540), Bujan Bight, Bering Island, Commander Islands, Bering Sea, 5 m, rocky platform, bottom water temperature of 7.6 C, Coll. V. I. Lukin, 28-VII- 1972 (sealer “Krylatka”); paratype (ММВ 9541), Poloviny Bay, Bering Island, Commander Is- lands, Bering Sea, 10 m, rocky platform, bot- tom water temperature of 9.8°C, Coll. V. I. Lukin, 27-1Х-1973 (R/V “Rakytnoe”); paratypes (4) (MIMB 9542) Ushishir Islands, Kuril Islands (42°30.2’N, 152`51.0’Е), 87-120 m, boulders covered Бу Spongia, Coll. С. М. Kamenev, 19- VII-2003 (R/V “Akademik Oparin”). Other Material Examined One slightly damaged specimen (MIMB 9543), Korabelnaya Bight, Medny Island, Com- mander Islands, Bering Sea, 5 m, rocky plat- form, bottom water temperature of 6.8°C, Coll. V. |. Lukin, 14-VII-1972 (sealer “Krylatka”); one slightly damaged specimen (MIMB 9544), Kamny Bobrovye, Medny Island, Commander Island, Bering Sea, 5 m, bottom water tem- perature of 5.6 C, Coll. V. 1. Lukin, 13-VII-1972 (sealer “Krylatka”); one left valve (ММВ 9545), Palata Cape, Medny Island, Commander Is- lands, Pacific Ocean, 15 m, rocky platform, bottom water temperature of 5.2°C, Coll. V. 1. Lukin, 16-\1-1972 (sealer “Krylatka”); one left valve (MIMB 9546), Phedoskina Cape, Bering Island, Commander Islands, 20 m, rocky plat- form, bottom water temperature of 9.9°C, Coll. V. |. Lukin, 23-IX-1973 (R/V “Rakitnoye”); one slightly damaged specimen (MIMB 9547), Utesnaya Bight, Second Kuril Strait, Paramushir Island, Kuril Islands, intertidal zone, boulders with brown algae of the gen- era Fucus and Alaria, Coll. M. В. Ivanova, 7- V11-1967; one specimen (ММВ 9548), Burevestnik Village, Iturup Island, Kuril Islands, Sea of Okhotsk, intertidal zone, boulders with brown algae of the genus А/апа, Coll. O. С. Kusakin, 24-VII-1967; one specimen (ММВ 9549), Nadezda Strait (Rashua Island - Matua Island), Kuril Islands (48°00’N, 153°15’E), 50 m, rocky platform, Coll. V. I. Lukin, 19-VIII-1987 (R/V “Tikhookeansky”). Total of 5 specimens and 2 left valves. Description Exterior. Shell small (to 11.2 mm), ovate- trigonal, high (H/L = 0.740-0.827), equivalve, moderately inflated (W/L = 0.181-0.235), al- most equilateral (slightly longer anteriorly, sometimes equilateral or longer posteriorly), thin, solid, white under periostracum. Surface with conspicuous growth lines. Periostracum non-polished, gray, sometimes light brown, dehiscent, easily peeled off near beaks, ex- tending into inner surface, thrown into small wrinkles, more conspicuous at shell margins. Beaks orthogyrate, small, slightly rounded, moderately projecting above dorsal margin, slightly posterior or anterior to midline, some- times central (A/L = 0.474-0.583). Anterior end rounded. Posterior end narrow, obliquely subtruncate, with faint radial ridge from beaks to ventral limit of posterior end. Anterodorsal margin slightly convex, gently descending ven- trally, smoothly transiting to rounded anterior end. Ventral margin slightly curved. Posterodorsal margin short, straight, gently descending ventrally, forming noticeable angle at transition to posterior margin. Posterior margin straight, rather steeply descending ventrally, forming rounded angle at transition to ventral margin. External ligament short (1/2 posterodorsal margin length), attached to short, wide nymph not projecting above dor- sal margin. Interior. Hinge plate wide, sometimes pro- jecting into shell cavity in area of cardinal teeth. Hinge weak, with two cardinal teeth in each valve. In left valve, anterior tooth wide, long, reaching edge of hinge plate; posterior tooth very narrow, lamellate, shorter, not reaching edge of hinge plate, almost parallel to anterior tooth. In right valve, anterior and posterior teeth almost same length and width (anterior tooth slightly shorter and wider). Internal liga- ment well developed, reaching edge of hinge plate, lodged in ovate-trigonal or ovate-elon- gate resilifer, which extends obliquely poste- rior to beaks. Anterior adductor muscle scar large, ovate, vertically extended; posterior adductor scar large, rounded, shorter and wider than anterior scar. Pallial sinus distinct, moderate, reaching past midline (L1/L = 0.603-0.698), broad, rounded anteriorly, of NEW SPECIES OF THE BIVALVE GENUS ABRINA 161 FIGS. 14-19. The hinge of the different age specimens of Abrina scarlatoi Kamenev, new species. FIGS. 14-16. Hinge of right valve. FIG. 14: Paratype (MIMB 9538), Phedoskina Cape, Bering Island, Commander Islands, Pacific Ocean, 10 m, shell length 6.2 mm. FIG. 15: Paratype (MIMB 9536), Tonky Cape, Medny Island, Commander Islands, Bering Sea, 20 m, shell length 6.8 mm. FIG. 16: Paratype (MIMB 9533), Cherny Cape, Medny Island, Commander Islands, Bering Sea, shell length 7.6 mm. FIGS. 17-19. Hinge of left valve. FIG. 17: Paratype (MIMB 9530), from holotype locality, shell length 6.2 mm. FIG. 18: Paratype (MIMB 9534), Palata Cape, Medny Island, Commander Islands, Pacific Ocean, 20 m, shell length 7.5 mm. FIG. 19: Paratype (MIMB 9531), Tonky Cape, Bering Island, Commander Islands, Bering Sea, 10 m, shell length 8.5 mm. Bar = 500 um. same shape and size in both valves (L1/L and L2/L of left valve 0.655 and 0.184; L1/L and L2/L of right valve 0.654 and 0.187), substan- tially detached, confluent with pallial line for more than 1/2 of its length. Shell interior often with faint radial striae. Variability Shell shape and proportions change with age. п young specimens (< 4 mm), in con- trast to adults, the shell is more elongate and angular; the posterodorsal margin atthe tran- sition to the posterior margin forms a distinct angle; the posterior margin more steeply de- scends ventrally, forming a pointed acute angle at the transition to ventral margin; the ventral margin is almost straight; the beaks are placed more posteriorly (АЛ. = 0.54- 0.583). The periostracum of young speci- mens has very fine, short, discontinuous radial lines in the area of the beaks. In young and adult specimens, the relative length, shape, and degree of confluence of the pal- lial sinus with the pallial line vary slightly. Sometimes, the length and shape of pallial sinus of right and left valves are slightly dif- ferent (Table 1). Distribution and Habitat (Fig. 20) Commander Islands: Bering Island and Medny Island; Kuril Islands: Paramushir Is- land; Nadezda Strait (Rashua Island - Matua Island) (48°00’N, 153°15’E); Ushishir Islands (42°30.2’N, 152'51.0'E); Iturup Island. 162 Near the Commander Islands, this species was found at depths from 3 m (Polovina Bight, Bering Island) to 100 т (Kamni Bobrovye - Kitolovnaya Bed, Medny Island, 54'58'N, 167'21'5E) on a rocky platform and boulders covered by a thick layer of lime red algae, with a population den- sity up to 30 specimens/m?; near the Kuril Is- lands — from the intertidal zone (Paramushir Island, Кигир Island) to 120 т (Ushishir Islands) on boulders covered by brown algae of the gen- era Fucus and Alaria or sponges. Comparisons In contrast to other species of Abrina, A. scarlatoi has the shell with rough, conspicu- ous growth lines, gray, a non-polished, dehis- cent periostracum, wide hinge plate, and a short, wide nymph (Table 2). Moreover, A. scarlatoi differs from А. lunella (Figs. 21-28) in its smaller, higher shell with less posteriorly placed beaks and in having the hinge with non- bifid cardinal teeth and a very narrow, lamel- RUSSIA KAMENEV late posterior cardinal tooth in the left valve (Gould, 1861; Kuroda, 1951; Habe, 1952, 1977, 1981; Kuroda et al., 1971; Ito et al., 1986; Kamenev & Nadtochy, 1999; Okutani, 2000); from A. kinoshitai, in a smaller, higher, more inflated, ovate-trigonal shell without a flexure of the posterior end, with less posteriorly placed beaks, a shorter pallial sinus of the same shape and size in both valves, and in having the hinge with non-bifid cardinal teeth and a very narrow, lamellate posterior cardinal tooth in the left valve (Ito, 1967, 1989; Kuroda et al., 1971; Habe, 1977; Tsuchida & Kurozumi, 1995; Kamenev & Nadtochy, 1999); from A. declivis, in the more elongate shell with a much less attenuate pos- terior end and in having the hinge with non- bifid cardinal teeth and a very narrow, lamellate posterior cardinal tooth in the left valve (Scott, 1994); from A. sibogai (Prashad, 1932), A inanis (Prashad, 1932), and A. weberi (Prashad, 1932), in the shell with less posteriorly placed beaks and lacking lunule and a escutcheon (Prashard, 1932). Bering Sea Sea of Okhotsk e Commander Islands ~ fe Paramushir Is. 2? o. Nadezda Strait en |" Ushishir Is. Is. ро Pacific Ocean 150° FIG. 20. Distribution of Abrina scarlatoi. 160° NEW SPECIES OF THE BIVALVE GENUS ABRINA 163 TABLE 1. Abrina scarlatoi Kamenev, new species. Shell measurements (mm), indices and summary statistics of all characteristics: L - shell length; H - height; W - width; A - anterior end length; L1 - maximal distance from the posterior shell margin to the top of pallial sinus; L2 - minimal distance from the top of pallial sinus to the anterior adductor muscle scar. Numerator indicates shell measurements and indices for the left valve, denominator - for the right valve. Depository № H W A L1 AT ENVIE ЗАЛЕ ТЕ ЕЕ Holotype MIMB 9529 DSC 22277470 78.475 2a) (0827 0.224! 0.480`0:653 0214 Siow 9 22 47 64 21 (0827 0.224 0480) 01653 02414 Paratype MIMB 9530 ¡ARAS ALO 3391 5:0 ALA (0789: 0-21: 01513 0658207184 ORIO 16 39 50 14 10:789.0.211 0:513 0658: 041184 Paratype MIMB 9530 62: 47 12 34 41 12 0758 0.194 0.548 0:661 0194 625547 12 54 41 1.2 0.758 0.194 0.548 0.661 0.194 Paratype MIMB 9531 CSG OO 4:3" Sr 17 0776 0224 01506, 0671 0200 56:6 19143 157 (17 0776 20722450506 0:6 7102200 Paratype ММВ 9531 80104 ESA 40 53 16 0500 0225 0500 0663 0200 8.0 6.4 18 40 53 1.6 0.800 0.225 0.500 0.663 0.200 Paratype MIMB 9531 ROMEO OMIS OM SIMS 0692022410 5180/6716 Oxia о © Ш 9 So 1.3 ,0.78970,2247 0818370. 8712 02171 Paratype MIMB 9531 68 54 16 7347 47 10 0194 0235 05 0691 0447 691154 kG 3445 12 0794 0255 915101662 0176 Paratype MIMB 9531 DOS MIE SOS CD ON 0765. 0: 196 0 5680/6415 R 02161 9.025.475 ll 3.0 36 09 0.768 0.196 0.536 0.6437 0.161 Paratype MIMB 9532 ИУ © Von 4 39 49 16 07497 0:152 01506101623) 0.208 TT STORIES OS ON 1 04001620 5060/6719 07195 Paratype MIMB 9533 LAA als) NES GS 1.2 0.803 0.211 0.474 0.671 0.158 (COROS 16 36 50 12 09803 021110474 0658 9.155 Paratype MIMB 9533 66 25:07 14 32 244 12 0158 0212 0455 0667 0.152 60504 32143 13 07591 0.21210455 09652 02197 Paratype MIMB 9534 TOR OT SAS 2 1570778702007 0:52070.69370 200 wa 99 9 99 91 0870/2008 01520) 01667 9 0:227 Paratype MIMB 9534 DA TO 27 35 0) 0141 0204 0500 101648 01685 54 40 1.1 2.7 3.5 1.0 0.741 0.204 0.500 0.648 0.185 Paratype MIMB 9535 MOST EC Зи 4 MES 01753: 02190.50 O644 0178 TS 3757821675735 475 1400775290 21990/279%0/6 1680192 Paratype MIMB 9536 12 56 1А 96 49 15 0779 0194 0.500 0.681 07208 Е 316-28 15 07780.19 050080667 70208 Paratype MIMB 9536 6:02 Sur а 136 43 16 07501 0:206; 01529 0.632 0235 u 5:1 14 36 4.4 14 0.750 0.206 0.529 0.647 0.206 Paratype MIMB 9536 LOS SN D OA 310) 0:9 0778101200) 101533 06670200 25723152 2.095 214310. 20/00/7786 02000 555206670200 Paratype MIMB 9537 ELISA WA 5 46 15 01500194 0486 0639} 0208 #2 54 (14 935 46 15 09750 0194 04865 01639 01208 Paratype MIMB 9537 AA 121 30137 12 0163 0203 050801627 0.203 DORE MIS OS 21076350203 ©:50810:62710:203 Paratype MIMB 9538 ИВО 2 0025 O OMS 0 189 0541106271 069 SNMP O 7227372 OOO SOMO OA 0162205189 Paratype MIMB 9538 65 249 14 93 42.12 01754 0215 0508 0646 0.155 65 24914 33 42 12 (0752 0215 10/5080 64620185 Paratype MIMB 9538 02 TS ds Sl es LS OE. 0210 0150001618 NO 240 62 4813 SA 3.9 1.3 0.774 0.210 0.500 0.629 0.210 Paratype MIMB 9539 а OS EN 09 0102 01211 0526101632 0140 5 244 12 30 98 08 0772 0211 0:526 0:66 0.190 Paratype MIMB 9539 8 0’ 20 -24 096 75’ 0199 0:51 0:629.10:162 И 28 07 2221072747206 09757 0139 0:54 10164910. 162 Paratype MIMB 9539 24 1905 14 16 04 0792 0208 0.583 01667 07167 О 14 10 04 0792 0.208 01583" 0661 0167 Paratype MIMB9540 ОВ SET ES 09 Oren 0208} 02526) 0,695 9151 a (99) > © — — N 00 (99) ©) o co о 00 — — = М о 00 o on N 00 o (o>) | [Ce] o _ — о (continues) 164 (continued) Depository L H W A Paratype MIMB 9541 3911191110822 3.3 1731 0.85 22 Paratype ММВ 9542 11.2 90 24 5.5 112, 290: 24% 62575 Paratype MIMB 9542 10.5 85 2.3 5.3 10.5. 8.5. 423-553 Paratype MIMB 9542 112 91 24 541 11220291 2,41 . 5.1 Paratype ММВ 9542 41 33 10 22 4.1 88 105722 MIMB 9547 60 48 13 353 6:0 4.8 13 33 MIMB 9548 7.8 MEM 16. 40 1.8. 61 1.6 4.0 ММВ 9549 Ar Tel. 2, 1 dur 4 74 Nhe, SALT Statistics L H W Mean 5.81 531 142 5:46 6.81 5.31 1.42 3.45 SD 2.10 1.72 0.48 0.96 2.10 1.72 048 0.96 SE 0.36 0.30 0.08 0.16 0.36 0.30 0.08 0.1 Min ES OS ET 4 1.9 0.5 1.4 Мах E SEO NS 11269" 24 55 N BET ET Eu ee 4 34 34 3 Etymology The specific name honors Orest A. Scarlato, Academician of the Russian Academy of Sci- ences, a famous Russian researcher of the marine bivalve fauna of Russia. Remarks The genus Abrina also includes species A. magna Scarlato, 1965, and A. hainanensis Scarlato, 1965, described by Scarlato (1965) from Hainan Island, South China Sea, China. All the material of A. magna (4 specimens and 103 shells) was collected from assemblages of empty shells on sandy beaches of Hainan Is- land and northern Vietnam. The material of A. hainanensis (the holotype and 10 additional specimens) is much smaller, but with the ex- ception ofthe holotype, was also sampled from assemblages of empty shells on sandy shores Hainan Island and the Gulf of Thailand (Bangkok). The holotype was collected in the KAMENEV L1 E27 AE ММ AM Е 12 2.5 0.6 0.795 0.205 0.564 0.641 0.154 2.5 0.6 0.795 0.205 0.564 0.641 0.154 7.2 1.7 0.782 0.214 0.491 0.643 0.152 7.2 1.8 0.782 0.214 0.491 0.643 0.160 7:0 22 «02604 0.219 0:505 0 6670210 7.9 2.2 0804 0.219 0.505 (01667 702110 78 17 0813 0.214 01455 0696 01152 7.8 1.8 0.813 0.214 0.455 0.696 0.161 2.7 0.8 0.791 0.244 0.537 0.659 0.195 2.7 0.8 0.791 0.244 0.537 0.659 0.195 40 11. 0.800 0.217 0:550: 0.667 0'183 4.0 1.1 0.800 0.217 0.550 0.667 0.183 47 1.3 0.782 0.205 0.513 0.603 0.167 47 1.3 0.782 0.205 0.513. 01603 10167 63 19 0755 01681 0500 0.670 0202 6.3 2.0 0755 0.181 0.500. 0:670107213 L1 [2 НМ АХ ЕЕ ЕЕ 4.47 1.26 0.778 0.208 0.514 0.655 0.184 4.46 1.28 0.778 0.208 0.514 0.654 0.187 1.43 0.43 0.023 0.014 0.026 0.024 0.024 1.42 0.44 0.021 0.014 0.027 0.019 0.021 0.24 0.07 0.004 0.002 0.005 0.003 0.004 0.24 0.07 0.004 0.002 0.005 0.003 0.004 16 0.4 0.740 0.181 0.455 0.603 0.140 16 0.4 0.740 0.181 0.455 0.603 0.140 718 2.2 0.827 0.244 0.583 0.698 0.235 7.8 34 34 34 34 34 34 34 3 34 34 34 34 34 34 intertidal zone of Hainan Island in the estuary of river, on silty sand among the mangroves. Having studied all materials relating to these species in the ZIN collection, | think that Scarlato (1965) erroneously assigned these species to Abrina. The hinge plate in these spe- cies is very wide, projects into the shell cavity in the area of resilifer. The hinge is weak, with two cardinal teeth in the right valve and two cardinal teeth in the left valve of A. hainanensis, and one cardinal tooth in the left valve of A. magna (Figs. 29-34). External and internal liga- ments are very large. The external ligament 1$ deeply sunken, almost internal, separated from the resilium by a slight ridge. The resilium is lodged in a large, trigonal resilifer behind the cardinal teeth. Lateral teeth are absent. Thus, the hinge of both species is identical to the hinge of the genus Psammotreta (Tellinidae) (Keen, 1969), except that the left valve of A. magna bears one cardinal tooth instead of two. To all appearances, the posterior cardinal tooth on the left valve of A. magna 1$ partly or com- NEW SPECIES OF THE BIVALVE GENUS ABRINA 165 FIGS. 21-34. Shells of Abrina species. FIGS. 21-28. Abrina lunella (Gould, 1861), NSMT (Mo 73503), Shiroko, Suzuka-shi, Mie Prefecture, Japan. FIGS. 21-24: Shell length 12.8 mm. FIGS. 25, 26: Shell length 10.2 mm. FIGS. 27, 28: Hinge of left and right valves. Bar = 1 mm. FIGS. 29-34. Abrina magna Scarlato, 1965. FIGS. 29, 30: ZIN (17), Tonkin Bay, North Vietnam, South China Sea, right valve, length 63.0 mm. FIGS. 31-34. ZIN (20), North Vietnam, South China Sea. FIGS. 31, 32: Right valve, length 49.6 mm. 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А|э}елэроци рэзеции А|эуелэрош Ajayesapou (YN эмел uu ‘y}fuel] У, 002 vel 00! all ep! $9 ‘хеш ||3US (712.0) ybiy (067`0) MO] (958`0) чбч (821.0) (669`0) mo] (15/`0) uBiy (1/H) ‘euoßin-s}eno чецобц}-э]ело 'ajeBuoja-ajeno jeuoBujgns чбщ‘1еицобц-э}ело ‘эзебио|э-э]ело 2чоб-э}ело adeus Jleys иэдэм y sıueul “y ıebogis “y SIAI[28P Y 10JEUE9S “y I¡EYYSOUN] “Y ejjaun] “y SJ9J0e.1ey) "y¡Bua] pus Jolajue - y “yjpim эмел - М uBiey - н ¿yyBua] les - 7 ‘dds euugy Jo siajoeJeyo Bunenualama ‘г FIGVL 166 NEW SPECIES OF THE BIVALVE GENUS ABRINA 167 pletely reduced with age. Unlike A. hainanensis, all valves of A. magna were very large, 42 to 73 mm long (valves of A. hainanensis are 9.5 to 22.3 mm in length). Moreover, since all ma- terial on this species was collected in assem- blages of empty shells, almost on all left and right valves the cardinal teeth were partly or completely destroyed, and the ligament and periostracum were lacking. Therefore, it is not inconceivable that the thin and weak posterior cardinal tooth was broken in all left valves. Examination of the descriptions and figures of members of the genus Psammotreta (Keen, 1969, 1971; Habe, 1977; Lamprell & White- head, 1992; Okutani, 2000) shows that A. ma- gna is most likely a synonym of Psammotreta (Tellinimactra) edentula (Spengler, 1798), in- habiting the intertidal and upper subtidal zones of Japan, South China, North Vietnam and Aus- tralia. Abrina magna is identical to P. (T.) edentula in hinge structure and morphology of the external and internal ligaments. Moreover, it has similar shape, proportions and size of the shell, a very deep pallial sinus in both valves, and scars of the anterior and posterior adduc- tors differing in shape and size (Figs. 29-34). It is possible that the material of A. hainanensis comprises young specimens of P (Т.) edentula. However, it is not unlikely that A. hainanensis is a separate species of the same subgenus. A more thorough study of specimens of different species of Psammotreta is needed to make a correct identification of A. magna and A. hainanensis. DISCUSSION Scarlato (1981) described new species of Abrina on the basis of a study of young speci- mens of Macoma (Kamenev 8 Nadtochy, 1999). The main morphological characteristic on the basis of which these species were pre- viously included in Abrina, was the presence of an internal ligament in an oblique resilifer posterior to the cardinal teeth. The genera Abrina and Macoma are similar in most mor- phological characteristics. The main distin- guishing characteristic of Abrina is the presence of a well-developed internal ligament in the resilifer, a narrow groove posterior to the cardinal teeth. In Macoma, an internal liga- ment is absent. However, studies of the com- mon northwestern Pacific Macoma species — M. loveni (Jensen, 1905), M. calcarea (Gmelin, 1791), M. balthica (Linne, 1758), M. crassula (Deshayes, 1855), M. lama Bartsch, 1921, M. incongrua (Martens, 1865) — show the pres- ence of an internal ligament in young speci- mens (Kamenev 8 Nadtochy, 1999). Thus, a well-developed internal ligament lodged in oblique resilifer in representatives of the ge- nus Macoma 1$ a juvenile characteristic that is preserved in Abrina during its entire life. Morphological similarity of the genera Abrina and Macoma, and the presence of an internal ligament in young specimens of species of Macoma, at first leads one to suggest that the present species is a juvenile of species of Масота. п Macoma, a well-developed resilium is found only in individuals up to 5-6 mm in shell length, whereas in specimens with a shell length more than 10 mm, it is lacking (Kamenev & Nadtochy, 1999). A study of A. scarlatoi of different ages showed that both young and adult specimens of this species have a well-developed resilium, lodged in the oblique resilifer posterior to the cardinal teeth. The shape of the resilifer changes with age, but its position and relative size remain un- changed. Furthermore, A. scarlatoi differs from most species of Macoma (Scarlato, 1981; Coan et al., 2000) in the lack of a flexure to the right of the posterior shell margin and by having pallial sinuses of similar shape and size in both valves. Therefore, | think that the spe- cies described herein belongs to the genus Abrina, not to Macoma. ACKNOWLEDGMENTS | am very grateful to Mrs. М. V. Kameneva (MIMB, Vladivostok) for great help during work on this manuscript; to Professor T. W. Pietsch and Dr. K. Stiles (UW, Seattle) for arrangement of my visit to the UW and work with the bivalve mollusks collection, for all-round, very kind, and friendly help during my stay and work in Se- attle; to Professor A. J. Kohn and Dr. G. Jensen (UW, Seattle) for great help during work with the bivalve mollusks collection at the UW; to Dr. P. D. Roopnarine and Miss. E. Kools (De- partment of Invertebrate Zoology, CAS, San Francisco) for arrangement of my work with the bivalve mollusks collection at the CAS and great help during this work; to Mr. Gary Cook (Ber- keley) for all-round, very kind and friendly help during my stay and work in San Francisco; to Drs. В. |. Sirenko and A. V. Martynov and all collaborators of Marine Research Laboratory (ZIN, St. Petersburg) for sending of the speci- mens of Abrina species and help during work with collection of bivalve mollusks of ZIN; to Dr. H. Saito (NSMT, Tokyo) for sending of the specimens of A. /unella and A. kinoshitaï; to Drs. 168 KAMENEV T. Kurozumi and E. Tsuchida (NHMI, Chiba) for sending of the specimens of А. kinoshitai and reprints of necessary papers; to Dr. K. Amano (Joetsu University of Education, Joetsu) for sending of the reprints of scientific papers nec- essary for work; Ms. R. N. Germon (USNM, Washington) for sending the specimens of dif- ferent species from the Semelidae; to Dr. E. V. Coan (Department of Invertebrate Zoology, CAS, San Francisco) and Paul Valentich Scott (Department of Invertebrate Zoology, SBMNH, Santa Barbara) for consultations and sending of the copies of scientific papers necessary for work; to Mr. D. V. Fomin (IMB, Vladivostok) for help in work with the scanning microscope; to Ms. T. N. Kaznova (IMB, Vladivostok) for help with translating of the manuscript into English; to Professor George M. Davis for help in the publication ofthe manuscript; and to two anony- mous reviewers for comments on the manu- script. This research was supported by Grant 01- 04-48010 from the Russian Foundation for Basic Research. LITERATURE CITED BUJANOVSKY, A. 1., 1997, On the fauna and ecology of the bivalves of the shallow water shelf zone of the Commander Islands. Pp. 242-253, in: A. V. RZHAVSKY, ed., Benthic flora and fauna of the shelf zone of the Commander Islands. Vladivostok: Dalnauka Press. 270 pp. [in Russian, with English abstract]. COAN, Е. V., Р. H. SCOTT & Е. К. BERNARD, 2000, Bivalve seashells of western North America. Marine bivalve mollusks from Arctic Alaska to Baja California. Santa Barbara Mu- seum of Natural History. viii + 764 pp. GOULD, A. A., 1861, Descriptions of shells col- lected by the North Pacific Exploring Expedi- tion. Proceeding Boston Society Natural History, 8: 14-40. HABE, T., 1952, Genera of Japanese shells. N 3. Pelecypoda, 187-280. [in Japanese]. HABE, T., 1977, Systematics of Mollusca in Ja- pan. Bivalvia and Scaphopoda. Tokyo. 372 pp. [in Japanese]. HABE, T., 1981, Bivalvia. A catalogue of molluscs of Wakayma Prefecture, the Province of КИ. |. Bivalvia, Scaphopoda and Cephalopoda. Pub- lications of the Seto Marine Biological Labora- tory, Special Publication Series, 7(1): 25-224. ITO, K., 1967, A catalogue of the marine mollus- can shell-fish collected on the coast of and off Tajima, Hyogo Prefecture. Bulletin of the Ja- pan Sea Regional Fisheries Research Labo- ratory, 18: 39-91 [in Japanese, with English abstract]. ITO, K., 1989, Distribution of molluscan shells in the coastal areas of Chuetsu, Kaetsu and Sado Island, Niigata Prefecture, Japan. Bulletin of the Japan Sea Regional Fisheries Research Laboratory, 39: 37-133 [in Japanese, with En- glish abstract]. ITO, K., Y. МАТАМО, У. YAMADA & $. IGARASHI, 1986, Shell species cught S/S Rokko-Maru off the coast Ishikawa Prefecture. Bulletin of the Ishikawa Prefectural Fisheries Experimental Station, 4: 1-179 [in Japanese, with English abstract]. KAMENEV, С. M., 1995, Species composition and distribution of bivalve mollusks on the Commander Islands shelf. Malacological Re- view, 28: 1-23. KAMENEV, G. M. 8 V. A. NADTOCHY, 1999, Species of Macoma (Bivalvia: Tellinidae) from the Pacific coast of Russia, previously de- scribed as Abrina (Bivalvia: Semelidae). Malacologia, 41(1): 209-230. KEEN, A. M., 1969, Superfamily Tellinacea. Pp. 613—643, in: R. C. MOORE, ed., Treatise on In- vertebrate Paleontology: Mollusca 6, Part N (Bivalvia). Lawrence, Kansas: University of Kansas Press. xxxviii + 952 pp. KEEN, А. М., 1971, Sea shells of tropical west America; marine mollusks from Baja Califor- та to Peru. Stanford, California: Stanford Uni- versity Press. xvi + 1064 pp. KURODA, T., 1951, Descriptions of a new ge- nus of a marine gastropod, Kanamurua, gen. n., and a new species of a bivalve, Abra kanamarui, sp. n., dedicated to Mr. T. Kanamaru on his 60" birthday. Venus, 16: 68— 72 [in Japanese]. KURODA, T., T. HABE & К. OYAMA, 1971, Sea shells of Sagami Bay. Tokyo: Maruzeh. xiv + 741 [in Japanese] + 489 pp. [in English]. LAMPRELL, К. & T. WHITEHEAD, 1992, Bivalves of Australia. Vol. 1. Bathurst: Crawford House Press. 182 pp. OKUTANI, T., 2000, Marine mollusks in Japan. Tokyo: Tokai University. xlvii + 1175 pp. PRASHAD, B., 1932, The Lamellibranchia of the Siboga Expedition. Systematic part II: Pelecypoda (exclusive of the Pectinidae), in: M. WEBER, ed., Siboga-Expeditie 34(53c). Leiden. 353 pp. SCARLATO, O.A., 1965, Bivalve mollusks of the superfamily Tellinacea of the Chinese seas. Studia Marina Sinica, 8: 27-114 [in Chinese, with Russian summary]. SCARLATO, O. A., 1981, Bivalve mollusks of temperate waters of the northwestern Pacific. Leningrad: NAUKA Press. 480 pp. [in Russian]. SCOTT, P. H., 1994, Bivalve molluscs from the southeastern waters of Hong Kong. Pp. 55-100, in: B. MORTON, ed., The malacofauna of Hong Kong and southern China Ill: Proceedings of the Third International Workshop on the Malac- ofauna of Hong Kong and Southern China, Hong Kong, 13 April-1 May 1992. Hong Kong: Hong Kong University Press. xxii + 504 pp. TSUCHIDA, Е. 8 Т. KUROZUMI, 1995, Fauna of marine mollusks of sea around Otsuchi Bay, lwate Prefecture (5) BIVALVIA-2. Otsuchi Ma- rine Research Center Report, 20: 13-42 [in Japanese]. Revised ms. accepted 8 February 2004 MALACOLOGIA, 2004, 46(1): 169-183 SMELE STRUCTURES OF SELECTED GASTROPODS FROM HYDROTHERMAL VENTS AND SEEPS Steffen Kiel Freie Universität Berlin, Institut für Geologische Wissenschaften, Fachrichtung Paläontologie, Malteserstrasse 74-100, 12249 Berlin, Germany; steffen.kiel@gmx.de ABSTRACT Shell structures of 24 gastropod species from hydrothermal vents and seeps are elec- tron microscopically investigated, and the ecological and phylogenetic implications of their shell structures are discussed. The presence of prismatic complex crossed lamellar, and regularly foliated structure in the Neolepetopsidae provides further evidence for their posi- tion as sister group of the Acmaeidae. The Lepetodriloidea are considered to be derived from, or to have a common ancestor with the Fissurellidae based on their complex crossed lamellar structure and on the presence of shell pores. The earlier hypothesis that Pelto- spiridae derived from Neomphalidae by reduction of complex crossed lamellar structure cannot be supported; both groups show the same array of shell structures. It is shown that shell pores are a frequent feature in Neomphalidae and Peltospiridae. Dissolution of the inner shell walls is documented for Bathynerita naticoides. The trend that small and thin- shelled gastropod groups tend to reduce their shell structure to intersected crossed platy, can also be observed in the vent/seep gastropods. Generally, their shell structures appear to reflect those of the phylogenetic group to which they belong, rather than being influ- enced by the peculiarities of the extreme environment they inhabit. Keywords: Gastropoda, shell structure, deep-sea, hydrothermal vent, cold seep, phylogeny. INTRODUCTION Chemosynthetic ecosystems in the deep-sea harbor highly endemic faunas (Tunnicliffe et al., 1996). The gastropods that live there are no exception to this: 95-98% of the species and 70% of the genera are endemic to vents and seeps, and five families are found exclu- sively here (Warén 8 Bouchet, 2001). Origin and phylogenetic relationships of many of the endemic taxa are still debated. Shell structures have only been described for three out of the about 125 gastropods spe- cies known from chemosynthetic ecosystems: Neomphalus fretterae (Batten, 1984), Melano- drymia aurantiaca (Hickman, 1984), and Lepetodrilus elevatus (Hunt, 1993). The scope of the present study is to provide an overview over the shell structures of the gastropod fami- lies present at chemosynthetic ecosystems, and to discuss their ecological and phyloge- netic implications. Additionally, these data can help to clarify the identity of fossil vent and seep gastropods. 169 MATERIALS AND METHODS The majority of the material used here 1$ from the study of Warén 8 Bouchet (2001), and was provided by the Muséum National d’ Histoire Naturelle in Paris (MNHN). Three ad- ditional species were provided by the Natural History Museum of Los Angeles County (LACM). All investigated specimens had the size of adult specimens as reported in the literature. The shell structure of protoconchs and onto- genetic changes in shell structures were not the subject of this study. Shell mineralogy was not studied, and is only inferred from the known mineralogy in related groups or 1$ noted in cases when the structures have an unequivocal mineralogy. To observe the shell structures, pieces of shell were broken off the apertural region to obtain fresh fracture zones. The material was then mounted on stubs, coated with gold, and observed with several scanning electron microscopes in Paris and Hamburg. 170 KIEL The different types of shell structures were determined following the scheme of Carter & Clark (1985) and Hedegaard (1990, 1997). All figures in this study are oriented in a way that the outer side of the shell is up. The shell structures present in each species are listed from the outer side of the shell towards the inside. The taxonomic framework is that of Warén & Bouchet (2001). Abbrevations in Figures ccl complex crossed lamellar hom homogenous ica intersected crossed acicular icp intersected crossed platy nac nacre pcc prismatic complex crossed lamellar per organic periostracum Мо regularly foliated rsp regular spherulitic prismatic scl simple crossed lamellar spr simple prismatic FIGS. 1, 2. Neolepetopsis cf. gordensis. FIG. 1: Upper side of shell with the outer complex crossed lamellar layer and the prismatic complex crossed lamellar layer below (bar = 100 um). FIG. 2: Detail of the prismatic complex crossed lamellar layer (bar = 10 pm). RESULTS Subclass Patellogastropoda Family Neolepetopsidae The prismatic complex crossed lamellar and regularly foliated structures are always com- posed entirely of calcite (Hedegaard, 1990). Neolepetopsis cf. gordensis McLean, 1990 - complex crossed lamellar (Fig. 1) - prismatic complex crossed lamellar (Figs. 122) Mid-America Trench, Jalisco Block, 18°22’N-104°23’W; seep т 3,000-3,300 m (MNHN). Eulepetopsis vitrea McLean, 1990 - prismatic complex crossed lamellar (Figs. 3,4) - regularly foliated (Fig. 5) - simple prismatic East Pacific Rise, NE of Ге de Paques, site Rehu, 17°24’S-113°12’W; vent in 2,578 m (MNHN). The prismatic complex crossed lamellar layer in this species has a very similar appearance as the “outer calcitic crossed lamellar” layer of Patella crenata described by Bandel & Geldmacher (1996). These authors com- pared their terminology only to those of Boggild (1930) and MacClintock (1967), but not to those of the more recent works of Lindberg (1986, 1988) and Hedegaard (1990). However, due to their similarity the “outer calcitic crossed lamellar” layer of Pa- tella crenata is here considered the same structure as prismatic complex crossed lamellar. Paralepetopsis ferrugivora Warén & Bouchet, 2001 - prismatic complex crossed lamellar (Figs. 6, 7) Mid-Atlantic Ridge, Lucky Strike; vent т about 1,650 m (MNHN). This specimen lacked any further details on its label, the depth is derived from the description of the Lucky Strike vent field (Van Dover et al., 1996). Hedegaard (1990) noted that the prisms of the prismatic complex crossed lamellar struc- ture are always convex towards the outer side of the shell. This is also observed in the three neolepetopsids investigated here. Hedegaard (1990) also pointed out that these prisms show ribbed surfaces, which SHELL STRUCTURES OF VENT AND SEEP GASTROPODS 171 he interpreted as the edges of the second order lamellae. This is also observed here, and these ribs have quite different appear- ances: in Neolepetopsis cf. gordensis they are fine tubercles (Fig. 2), in Paralepetopsis ferrugivora they are coarse and irregular (Fig. 7), and in Eulepetopsis vitrea they form a distinct grid-like pattern with tuberculate intersections (Fig. 4). FIGS. 3-5. Eulepetopsis vitrea. FIG. 3: Outer prismatic complex crossed lamellar layer (bar = 10 um). FIG. 4: Close-up on the grid-like, ribbed surface of the prisms (bar = 1 um). FIG. 5: Regularly foliated layer (Баг = 10 um). Subclass Cocculiniformia Family Pyropeltidae Pyropelta musaica McLean 8 Haszprunar, 1987 - simple prismatic (Figs. 8, 9) - simple crossed lamellar (Figs. 8, 9) - simple prismatic (Fig. 8) California, Santa Catalina Basin, between San Clemente and Santa Santa Catalina, 33°12’N, 118°30’W; whale bone from 1,240 m; (LACM 146909). The shell consists of at least five alternating layers of simple crossed lamellar and simple prismatic structure, with the simple crossed lamellar layer becoming progressively thicker towards the outer side of the shell. The microcrystals of the simple crossed lamellar layers are not very densely packed. Subclass Vetigastropoda Family uncertain Sahlingia xandaros Warén & Bouchet, 2001 - simple prismatic (Fig. 10) - intersected crossed acicular or platy (Fig. 10) - homogenous (Fig. 10) FIGS. 6, 7. Paralepetopsis ferrugivora. FIG. 6: Overview showing that the entire shell is composed of prismatic complex crossed lamellar structure (bar = 10 um). FIG. 7: Close-up on the prisms, showing their ribbed surface (bar = 10 um). 172 KIEL FIGS. 8, 9. Pyropelta musaica. FIG. 8: Cross- section showing five layers with simple crossed lamellar structure, with four thin layers with simple prismatic structure between them, marked by thin white bars (bar = 10 um). FIG. 9: Close-up on three simple crossed lamellar layers (bar = 10 pm). Alaska, Aleutian Trench, Kodiak Seep, 56°55.65’М, 149°32.90’W (LACM 1999-45); seep in 4,430 m. The crossed layer has a granular appearance making it difficult to distinguish between in- tersected crossed acicular or platy structure. FIG. 10. Sahlingia xandaros, showing the thin outer layer of simple prismatic structure, the remaining shell is composed of intersected crossed acicular or platy structure (bar = 10 um). FIGS. 11, 12. Protolira valvatoides. FIG. 11: Organic periostracum, simple prismatic, and intersected crossed platy structure (bar = 10 um). FIG. 12: Outer side of shell with organic periostracum and the intersected crossed platy layer (bar = 10 um). FIG. 13. Bruceiella athlia, showing the intersected crossed platy and simple prismatic layers, and the intersected crossed acicular layer below (bar = 10 um). SHELL STRUCTURES OF VENT AND SEEP GASTROPODS 173 Family Skeneidae Protolira valvatoides Warén & Bouchet, 1993 - simple prismatic (Fig. 11) - intersected crossed platy (Figs. 11, 12) - Simple prismatic (Fig. 11) Mid-Atlantic Ridge, Lucky Strike, site Pago- das, 54”18.32'N-32*16.51W; vent in 1,685 m (MNHN). Many of the microcrystals have a granular appearance, and show cavities between each other. Bruceiella athlia Warén & Bouchet, 1993 - intersected crossed platy (Fig. 13) - simple prismatic (Fig. 13) - intersected crossed acicular (Fig. 13) Aleutian Trench, site Shumagin, 54°18.06’N— 157°12.11’W; seep in 2,524 т (ММНМ). Many of the microcrystals have a granular appearance, and show cavities between each other. Family Sutilizonidae Sutilizona theca McLean, 1989 - simple prismatic (Fig. 14) - intersected crossed platy (Fig. 14) East Pacific Rise, 11°46’N, 103°47’W; vent in 2,715 m (Paratype LACM 2355). Family Lepetodrilidae Hunt (1992) used powder diffraction to show that the shell of Lepetodrilus elevatus is com- posed entirely of aragonite. It is therefore as- sumed that the shells of the lepetodrilids investigated here are also composed of ara- gonite. FIG. 14. Sutilizona theca, showing the thin outer layer of simple prismatic structure, the remaining shell is composed of intersected crossed platy structure (bar = 3 um). Lepetodrilus pustulosus McLean, 1988 - simple prismatic (Fig. 15) - complex crossed lamellar (Figs. 15, 16) East Pacific Rise, sites Parigo, Genesis, Elsa, 12°48.52’N-103°56.48’W; vent in 4,808 m (MNHN). There are occasionally fine pores perpen- dicular to the shell’s surface, with an aver- age diameter of 1 um (Fig. 16). Pseudorimula midatlantica McLean, 1992 - homogenous - complex crossed lamellar Mid-Atlantic Ridge, Snake Pit, site Elan, 23°23’ N—44°56’W; vent in 3,520 т (MNHN). There are occasionally fine pores perpen- dicular to the shell’s surface, with an aver- age diameter of 1-2 um. Family Trochidae Bathymargarites symplector Warén 8 Bouchet, 1989 FIGS. 15, 16. Lepetodrilus pustulosus. FIG. 15: Cross section showing the outer simple prismatic layer, and the inner complex crossed lamellar layer (bar = 100 um). FIG. 16: Close-up on the crossed lamellar layer, arrows indicate the fine, vertical pores (bar = 10 um). 174 KIEL FIGS. 17, 18. Bathymargarites symplector. FIG. 17: Overview showing the regular spherulitic prismatic upper layer, and the nacreous inner layer (Баг = 50 um). FIG. 18: Close-up on the outer side of the shell showing the thin homogenous layer and the upper part of the regular simple prismatic layer (bar = 10 um). Subclass uncertain Family Neomphalidae FIG. 20. Melanodrymia aurantiaca, view on the simple crossed lamellar structure (bar = 10 um). - homogenous (Fig. 18) Retiskenea diploura Warén & Bouchet, 2001 - regular spherulitic prismatic (Figs. 17, 18) - homogenous (Fig. 19) - columnar nacre (Fig. 17) - simple prismatic (Fig. 19) East Pacific Rise 13°М; the label in the box - intersected crossed platy (Fig. 19) indicates “same as Warén & Bouchet, 1993: - simple prismatic 11-13, figs. 10A-E, 11А-В”; it is thus likely Aleutian Trench, site Shumargin, to be from a vent in 2,616-2,635 т (MNHN). 54°18.17’N-157°11.82’W,; seep in 4,808 т (Paratype, MNHN). The microcrystals of the crossed platy layer are not very densely packed and have a granular appearance. Melanodrymia aurantiaca Hickman, 1984 - simple prismatic (Fig. 20) - simple crossed lamellar (Fig. 20) - simple prismatic (Fig. 20) East Pacific Rise, sites Parigo, Pogosud, Genesis, 12°48.52’N-103°56.48’W; vent in 2,630 m (ММНМ). Hickman (1984) reported a thick layer with complex prismatic structure in this species. Hedegaard (1990) pointed out that Mac Clintock’s (1967) “complex prismatic” struc- FIG. 19. Retiskenea diploura, the homogenous, ture is identical with the "simple crossed simple prismatic, and intersected crossed platy lamellar” structure of Carter & Clark (1985). structures merge into each other (bar = 10 um). Thus, the superficial differences between SHELL STRUCTURES OF VENT AND SEEP GASTROPODS 175 FIG. 21. Cyathermia naticoides, upper part of a cross-section with simple prismatic and complex crossed lamellar structure, arrow indicates a shell роге (bar = 10 um). Hickman’s (1984) and my descriptions of the shell structures of Melanodrymia aurantiaca is likely to be only a difference in terminology. Cyathermia naticoides Warén & Bouchet, 1989 - simple prismatic (Fig. 21) - complex cross lamellar (Fig. 21) FIG. 22. Pachydermia laevis, homogenous and complex crossed lamellar structure, arrow indicates a shell pore (bar = 100 рт). FIG. 23. Peltospira smaragdina has an outer layer with simple prismatic structure, and an inner layer with complex crossed lamellar structure, arrows indicate the broad shell pores (bar = 10 pm). East Pacific Rise, sites Julie, Genesis, Parigo, 12°48.96’N-103°46.62’W; vent in 2,630 m (MNHN). Pachydermia laevis Warén & Bouchet, 1989 - homogenous (Fig. 22) - complex crossed lamellar (Fig. 22) East Pacific Rise, site Genesis, 12°48.56’N— 103°46.58’W; vent in 2,630 т (MNHN). There are occasionally fine pores perpen- dicular to the shell's surface with an aver- age diameter of 3 um (Fig. 22). Planorbidella planispira (Warén 8 Bouchet, 1989) - homogenous - complex crossed lamellar - simple prismatic East Pacific Rise, site Elsa, 12°48.09’N- 103°46.34’W; vent in 2,630 т (MNHN). Family Peltospiridae Peltospira smaragdina Warén & Bouchet, 2001 - simple prismatic (Fig. 23) - complex cross lamellar (Fig. 23) Mid-Atlantic Ridge, Lucky Strike, site Sintra, 37°17.50’N-32°16.47’W; vent in 1,622 т (MNHN). There are occasionally fine pores perpen- dicular to the shell’s surface, with an aver- age diameter of 4 um (Fig. 23). Ctenopelta porifera Warén & Bouchet, 1993 - homogenous, with traces of unidentified shell structures (Figs. 24, 25) - simple prismatic (Fig. 25) East Pacific Rise, sites Totem, Genesis, Elsa, 12°48.71'N-103°56.53’W; vent in 2,630 m (MNHN). 176 KIEL The shell is perforated by fine pores with an average diameter of 4 ит (Figs. 24, 25); these pores have not been observed in the internal septum. Lirapex costellata Warén & Bouchet, 2001 - simple prismatic - complex crossed lamellar - homogenous - simple prismatic Mid-Atlantic Ridge, Lucky Strike, site Tour Eiffel, 37°17.32 N-32°16.51’W; vent in 1,685 т (MNHN). Недедаага (1990) presented shell structure data for three species he considered peltospirids. Among these, Hyalogyrina glabra has subsequently been assigned to the heterobranch family Hyalogyrinidae (Warén 8 Bouchet, 2001). The other two — Xyloskenea costulifera and Bathyxylophila excelsa — were placed in the Skeneidae (Marshall, 1988), and FIGS. 24, 25. Ctenopelta porifera. FIG. 24: Outer layer with homogenous, and remnants of an unidentified structure, arrow indicates a shell pore (bar = 10 um). FIG. 25: Lower side of shell with homogenous?, and simple prismatic structure, arrow indicates a shell pore (bar = 10 um). no subsequent work has been done on them. However, the Skeneidae are a heterogenous group of small-shelled trochoids, and the as- signment of Xyloskenea costulifera and Bathyxylophila excelsa to either the Skeneidae or the Peltospiridae is still uncertain (Marshall, pers. comm., 2003). Subclass Neritimorpha Family Neritidae Bathynerita naticoidea Clarke, 1989 - homogenous (Fig. 26) - simple cross lamellar (Fig. 26) - simple prismatic Louisiana Slope, Bush Hill Seep, 27°46.91’N—91°30.34’W; seep in 540-580 m (MNHN). FIGS. 26, 27. Bathynerita naticoidea. FIG. 26: Mainly complex crossed lamellar structure, and thin, homogenous outer layer (bar = 100 um). FIG. 27: View on the shell’s interior showing that the inner walls are dissolved (bar = 100 um). SHELL STRUCTURES ОЕ VENT AND SEEP GASTROPODS A Hedegaard (1990) found the outer, homog- enous layers of the five Neritidae investigated by him to be composed of calcite. It is thus likely that the thin homogenous outer layer of Bathynerita naticoidea is also composed of calcite. The inner shell walls of Bathynerita naticoidea are dissolved (Fig. 27). Dissolution of the inner shell walls is characteristic for the Neritidae, but has apparently never been docu- mented for Bathynerita naticoidea. Family Phenacolepidae Shinkailepas briandi Warén & Bouchet, 2001 - homogenous (dense) (Fig. 29) - homogenous (granular) (Fig. 29) - complex crossed lamellar (Figs. 28, 29) - intersected crossed platy (Fig. 28) - simple prismatic Mid-Atlantic Ridge, Lucky Strike, site Sintra, 37°17.50’N-32°16.47’W; vent in 1,622 m (MNHN). FIGS. 28, 29. Shinkailepas briandi. FIG. 28: inner side of the shell, showing the transition from complex crossed lamellar to intersected crossed platy structure (bar = 10 um). FIG. 29: Outer side of the shell showing the homogenous outer layer; the homogenous layer is very dense in the rib, and more granular away from the rib (bar = 100 um). The homogenous outer layer that builds the ridges on the shell surface is similar to the outer layer described for Phenacolepas pulchellus by Hedegaard (1990). Hedegaard (1990) assumed that this layer has calcitic shell mineralogy, which is also likely in the species investigated here. FIGS. 30-32. Provanna variabilis. FIG. 30: Overview showing the inner and outer, simple prismatic layers, and a central layer with complex crossed lamellar structure (bar = 10 um). FIG. 31: Close-up on the upper side of the shell, showing the slightly detached organic periostracum, and the transition from the outer simple prismatic to the complex crossed lamellar layer (bar = 10 um). FIG. 32: Inner simple prismatic layer is absent in this part of the shell (bar = 50 um). 178 KIEL vie HT IAE LT TS ARRET Pa НЕ: FIG. 33. Alviniconcha hessleri, the organic periostracum is about two and a half times thicker than the shell (bar = 100 pm). Subclass Caenogastropoda Family Provannidae Provanna variabilis Warén & Bouchet, 1986 - simple prismatic (Figs. 30-32) - complex crossed lamellar (Figs. 30-32) - Simple prismatic (Fig. 30) Juan de Fuca Ridge, 47°57’N-129°04’W; vents in 2,212 m (MNHN). In the innermost portion of the complex crossed lamellar layer the microcrystals are sometimes only loosely packed, although they are densely packed in the remaining part of the layer (Fig. 30). The inner simple prismatic layer may be present or absent at different parts of the shell (compare Figs. 30 and 32). Alviniconcha hessleri Okutani 8 Ohta, 1988 - simple prismatic - complex crossed lamellar (Fig. 33) - simple prismatic Mariana Back Arc Basin, site Alice Springs, 1812.59 N-144*42.43'E; vent in 3,630- 3,655 m (MNHN). Subclass Heterobranchia Family Xylodisculidae Xylodiscula analoga Warén & Bouchet, 2001 - intersected crossed platy (Fig. 34) - simple prismatic (Fig. 34) Mid-Atlantic Ridge, Lucky Strike, site Tour FIG. 34. Xylodiscula analoga has mainly intersected crossed platy structure, and a thin layer with simple prismatic structure at the inner side of the shell. The “smeared” area in the center of the Due and the loose packing of microcrystals probably indicate a high content of organic material bar = 10 um). SHELL STRUCTURES OF VENT AND ЗЕЕР GASTROPODS 179 Eiffel, 37°17.32’N-32°16.51’W; vent in 1,685m (MNHN). The microcrystals are not very densely packed. Family Hyalogyrinidae Hyalogyrina umbellifera Warén & Bouchet, 2001 - simple prismatic (Figs. 35, 36) - intersected crossed platy (Figs. 35, 36) Aleutian Trench, site Shumagin, 54°18.17’N- 157°11.82’W; seep in 4,808 m (Paratype, MNHN). This composition of shell structure is similar to that described for Hyalogyrina glabra (Hedegaard, 1990). The microcrystals are not very densely packed. FIGS. 35, 36. Hyalogyrina umbellifera. FIG. 35: Thin outer layer with simple prismatic structure, and intersected crossed platy structure below (bar = 10 um). FIG. 36: Close-up on the transition from simple prismatic to intersected crossed platy structure, note the loose packing of the microcrystals (bar = 10 pm). DISCUSSION Among the purposes of this study was to investigate whether the peculiarities of the vent/seep environment influence the shell structures of the gastropods groups living there. The microcrystals that build the shell structures are not very densely packed in sev- eral species (e.g., in Retiskenea diploura, Hyalogyrina umbellifera, Xylodiscula analoga, and to a lesser extend also in Pyropelta musaica, Protolira valvatoides, and Bruceiella athlia). Such loose packing is rarely observed in gastropods from shallow-marine environ- ments, even in very thin-shelled species (Bandel, pers. comm. 2003; pers. observa- tions). This loose packing is most probably the result of a high organic content in the shell. Loose packing occurs most frequently in small, thin shells with intersected crossed platy struc- ture. This makes it at present impossible to distinguish whether it is related to the extreme vent/seep habitat, or to shell thickness and structure, or both. An obvious correlation, although not related to the vent/seep habitat, is that between the presence of intersected crossed platy struc- ture and shell thickness. This structure occurs more frequently in small, thin-shelled species than in larger, thicker-shelled ones. Hede- gaard (1990) noted that among the Archaeo- gastropoda, intersected crossed platy structure dominates in species from small- shelled groups. In case of the vent/seep gas- tropods investigated here, this tendency can not only be observed among archaeogastro- pods, but also among the Heterobranchia, in the families Hyalogyrinidae and Xylodisculidae. There are no apparent correlations between shell structures and depth or habitat. Neolepetopsidae Anatomical and molecular data indicate a sister group relationship of Neolepetopsidae and Acmaeidae (Lindberg, 1998; Harasewych & McArthur, 2000). The three neolepetopsids investigated here have prismatic complex crossed lamellar shell structure, which Hede- gaard (1990) considered as apomorphy of the Acmaeidae. Hedegaard (1990) also pointed out that regularly foliated structure is present only in few acmaeids, and considered the re- duction of this structure as an apomorphy of the Acmaeidae sensu stricto. Regularly foli- 180 KIEL ated structure is present in Eulepetopsis vitrea, but absent in Neolepetopsis cf. gordensis and Paralepetopsis ferrugivora. The position ofthe Neolepetopsidae as sister group of the Acmaeidae within the Acmaeoidea can thus be supported. However, it should be noted that the patellid Patella crenata also has prismatic complex crossed lamellar structure (Bandel & Geldmacher, 1996), raising some doubt whether this shell structure can actually be considered as an apomorphy of the Acmaeidae. Pyropeltidae Pyropelta musaica has simple crossed lamellar structure like the three cocculinids investigated by Hedegaard (1990), but a multi- layered occurrence of this structure separated by thin layers of a different structure as in Pyropelta musaica was not described. Neither does a fossil cocculinid from the Cretaceous show such a pattern (my data). A total of five investigated Cocculiniformia are far too few to propose this alternation of shell structures as an apomorphy of the Pyropeltidae. How- ever, when future research confirms that this pattern does not occur in other cocculiniforms, it could be used for phylogenetic purposes, and also to identify members of the Pyropel- tidae in the fossil record. Sahlingia Sahlingia xandaros has only simple prismatic and intersected crossed platy structures, which are not very conclusive for phylogenetic pur- poses. Skeneidae The two skeneids have the same shell struc- tures as the three skeneids investigated by Hedegaard (1990). Lepetodrilidae and Sutilizonidae The Lepetodrilidae are considered here to be derived from, or to have a common ances- tor with the Fissurellidae. The two decisive factors are their complex crossed lamellar structure, and their fine shell pores. Among the slit-bearing Vetigastropoda, the Pleuro- tomariidae, Haliotidae, and Seguenziidae have a nacreous shells (Baggild, 1930; Erben & Krampiz, 1972; Bandel, 1979; Hedegaard, 1990; Harasewych, 2002) and are thus less likely to be related. This also pertains to the Palaeozoic slit-bearing Porcellidae, for which nacre is inferred from the presence of nacre in its Mesozoic sister group, the Cirridae (Kiel 8 Fryda, 2004). The two remaining slit-bear- ing groups, Fissurellidae and Scissurellidae, have crossed lamellar structure (Batten, 1975; Bandel, 1998), have shell morphologies simi- lar to those of the lepetodrilids, and are ana- tomically similar (Warén, pers. comm., 2003). Among these, only the fissurellids show shell pores as found in the lepetodrilids. Shell pores evolved independently in several groups of mollusks (Reindl 8 Haszprunar, 1996), can be present or absent in genera of the same fam- ily (e.g., Peltospiridae or Neomphalidae as shown herein), and may even be present or absent in species of the same genus — for example, Shinkailepas briandi without pores (herein) and Shinkailepas myojinensis with pores (Sasaki et al., 2003). However, shell pores have never been reported in scissurellids but frequently in fissurellids. Thus, the coinci- dence of similar shell shape, shell structure, and the presence of shell pores in both groups allows me to propose a close phylogenetic relationship between Fissurellidae and Lepetodrilidae. The sutilizonid Sutilizona theca has inter- sected crossed platy structure, which 1$ nei- ther very conclusive for phylogenetic analysis, nor does it contradict previously suggested relationships (Haszprunar, 1988; Ponder 8 Lindberg, 1997; Warén 8 Bouchet, 2001). Trochidae Bathymargarites symplector has columnar nacre which is a common shell structure among the Trochidae (Wise, 1970; Erben, 1974; Hedegaard, 1990; Hickman & McLean, 1990). It is, however, the only gastropod with nacreous shell investigated here. Among other trochids from vents and seeps, nacre was re- ported from Cataegis meroglypta (McLean & Quinn, 1987; Warén & Bouchet, 1993), and from species of Falsimargarita (Warén & Bouchet, 2001). Neomphalidae and Peltospiridae Hedegaard (1990) proposed that the Pelto- spiridae is derived from the Neomphalidae by reduction of crossed lamellar structure. How- ever, this was based on the incorrect higher taxonomic placement of his species — none of SHELL STRUCTURES OF VENT AND ЗЕЕР GASTROPODS 181 them appears to belong to the Peltospiridae. Two of the three peltospirids investigated here show complex crossed lamellar structure, whereas two of the six Neomphalidae with known shell structure lack complex crossed lamellar structure. These new observations negate derivation of the Peltospiridae from the Neomphalidae, but do provide additional evi- dence that both families are related, as indi- cated by anatomical and molecular studies (Israelsson, 1998; McArthur & Tunnicliffe, 1998; Warén & Bouchet, 2001). Shell pores in the Neomphalidae were first reported from Neomphalus fretterae, which has two types of pores, averaging 0.1 um and 1.0 um in diameter (Batten, 1984). Four out of the nine species of the Neomphalidae and Peltospiridae investigated here (including Neomphalus fretterae) have pores in their shells. In Peltospira smaragdina, Pachydermia laevis, and Ctenopelta porifera, the pores have an average diameter of 1.0-4.0 um. The latter species has additional macropores 30.0-70.0 um in diameter (Warén & Bouchet, 1993). The function of such shell pores is still controver- sial (Reindl & Haszprunar, 1996). Batten (1984) found the highest concentration of pores in Neomphalus fretterae around muscle insertion fields, and therefore interpreted them as muscle insertions. In the case of Ctenopelta porifera, Warén & Bouchet (1993, 2001) suggested the macropores to be related to chemosymbiosis. Although the shell structure of the proto- conch is not the scope of this study, Batten’s (1984) interpretation of the multi-layered protoconch of Neomphalus fretterae deserves comment. Batten (1984) speculated that the three shell layers of the protoconch “may indi- cate that the veliger larval stage may have an extended planktonic mode.” Calcification of the protoconch in archaeogastropods, however, takes place at the beginning of their benthic life, after the velum has been discarded (Bandel, 1982), and thus after the free-swim- ming larval stage. The additional inner layers have thus been built by the benthic juvenile or adult, possibly to strengthen the apical por- tion of the sub-limpet shell. Neritidae and Phenacolepidae Both investigated species, Bathynerita naticoidea and Shinkailepas briandi, have crossed lamellar structures like their shallow- marine relatives. Likewise, both species have a homogenous outer layer with presumably calcitic mineralogy. In this respect, both spe- cies differ from the neritilliid Pisulina, in which the thin outer layer has simple prismatic struc- ture (Kano & Kase, 2000). In contrast to the classification of Warén & Bouchet (2001), Bathynerita has recently been considered to be more closely related to the Phenacolepidae than to the Neritidae (Hodgson et al., 1998; Kano et al., 2002). Unfortunately, shell struc- tures are too uniform among the two groups to provide further evidence to this hypothesis. The inner shell walls of Bathynerita naticoidea are dissolved, a feature that is characteristic for all known neritoids, but had not yet been demonstrated for Bathynerita. Provannidae The observed shells structures in the two provannids are similar to those of other caenogastropods (Bandel, 1990). Hyalogyrinidae and Xylodisculidae Both investigated species have homogenous and intersected crossed platy structure, whereas all other Known heterobranchs have crossed lamellar structure (Bandel, 1990). This devia- tion might result from their small and thin shells, rather than being of phylogenetic importance. In sum, the shell structures of the vent and seep gastropods appear to reflect those of the phylogenetic group to which they belong, rather than being influenced by the peculiari- ties of the extreme environment they inhabit. ACKNOWLEDGMENTS | would like to thank P. Bouchet, Paris, who made my visit to the MNHN in Paris possible; V. Heros and R. von Cosel, Paris, for their help with the collection in Paris; J. McLean and L. Groves, Los Angeles, for making shell material from the NHM available to me; C. Chancogne, and G. Mascarell, Paris, for their help with the SEM; A. Warén, Stockholm, for discussion and for sharing unpublished data; C. T. S. Little, Leeds, for linguistic improvements; K. Bandel, Hamburg, for discussion of shell structures and shell formation in gastropods, and two anonymous reviewers for their critical reading of the manuscript. 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TAYLOR, ed., Origin and evolutionary radiation of the Mol- lusca. London: Oxford University Press. SASAKI, T., T. OKUTANI & К. FUJIKURA, 2003, New taxa and new records of patelliform gas- tropods associated with chemoautosynthesis- based communities in Japanese waters. The Veliger, 46: 189-210. TUNNICLIFFE, V., M. R. FOWLER £ A. G. MCARTHUR, 1996, Plate tectonic history and hot vent biogeography. Pp. 225-238, in: к. J. MACLEOD, ed., Tectonic, magmatic, hydrother- mal and biological segmentation of Mid-Oce- anic ridges. Geological Society, Special Publi- cation. | VAN РО\УЕР. С. LD DESBRUYERES; М. SEGONZAC, Т. СОМТЕТ, А. Е.-М. SALDANHA & С. LANGMUIR, 1996, Biology of the Lucky Strike hydrothermal field. Deep-Sea Research |, 43: 1509-1529. WAREN, A. € P. BOUCHET, 1993, New records, species, genera, and a new family of gastro- pods from hydrothermal vents and hydrocar- bon seeps. Zoologica Scripta, 22: 1-90. WAREN, A. 8 P. BOUCHET, 2001, Gastropoda and Monoplacophora from hydrothermal vents and seeps; new taxa and records. The Veliger, 44: 116-231. WISE, S. W., 1970, Microarchitecture and mode of formation of nacre (mother-of-pearl) in pele- cypods, gastropods, and cephalopods. Eclogae Geologicae Helvetiae, 63: 775-797. Revised ms. accepted 2 April 2004 MALACOLOGIA, 2004, 46(1): 185-202 CREPIDULA CACHIMILLA (MOLLUSCA: GASTROPODA), À NEW SPECIES FROM PATAGONIA, ARGENTINA Maximiliano Cledén' ”, Luiz Ricardo L. Simone? & Pablo Е. Penchaszadeh! ABSTRACT А new species, Crepidula cachimilla, is described based on a population from 15 т depth in San Antonio Oeste, Argentina. Shell length ranged from 5.4 to 28.5 mm for males and from 9.6 to 52.2 mm for females. The minimum shell length recorded for a brooding female was 23.5 mm, and the maximum shell length was 49.3 mm. A detailed anatomical description is given, showing as main characters of the species a relative thick columellar muscle, a greater closure of the pallial cavity aperture by a fusion of the mantle border, a very small osphradium, with about 16 broad filaments, endostyle divided by a middle lon- gitudinal furrow, very large salivary glands, duplication of both gastric ducts to the diges- tive gland, male seminal vesicle very long and with irregular walls, pallial oviduct with a broad vaginal duct and a tall papilla originating both from pallial floor and roof. Brood egg masses of mature females contained from 15 to 65 egg capsules. The triangular-shaped egg capsules measured between 2.2 and 3.4 mm in length and between 2.3 and 3.8 mm in width. Each egg capsule contained between 129 and 563 eggs. The number of eggs per capsule and the egg diameter did not correlate with female shell length. Uncleaved eggs measured between 180 and 200 ит in diameter. They all developed synchronously within the egg capsules. Prehatching veliger shells measured between 260 and 300 ит in length. After hatching at the veliger stage, protoconch length during metamorphosis ranged be- tween 700 and 800 um. These parameters neither coincide with those reported by Hoagland (1977) for the similar Californian Crepidula onyx, nor with the reproductive characters reported by Miloslavich 8 Penchaszadeh (2001) for Crepidula aplysioides, which suppos- edly occurs in the region. Key words: Crepidula cachimilla, new species, Calyptraeoidea, anatomy, reproduction, southwestern Atlantic, Patagonia, hermaphroditism. INTRODUCTION According to Dall (1909: 234), Crepidula onyx (С. В. Sowerby |, 1824) occurs along the Pacific coast from North America to Chile. Based on shell and radular morphology, Parodiz (1939) reported this species on the Atlantic coast of Argentina, from San Matías Gulf to Punta Norte, and Aguirre & Farinati (2000) recorded fossils of this species from the Quaternary period in northeastern Argen- tina. Hoagland (1977) suggested that the At- lantic material studied by Parodiz (1939) should be attributed to C. aplysioides. Crepidula aplysioides has been defined both anatomically (Simone, 2002) and by reproduc- tive patterns (Miloslavich 4 Penchaszadeh, 2001). Based on the differences with the stud- ied sample, we conclude that our material from San Antonio Oeste, Argentina, belongs to an undescribed species. In this paper, we de- scribe this new species, which 1$ restricted to an area of Patagonia, southwestern Atlantic. The study on the calyptraeids has grown considerately in the last few years with the addition of knowledge on the anatomy (e.g., Simone, 2002), molecular biology (e.g., Collin, 2000), and reproductive strategies (e.g., Miloslavich & Penchaszadeh, 2001). From the eastern coast of Americas, knowledge of the informally defined “Crepidula plana complex” is of particular importance (Collin, 2000; Simone, submitted; Simone et al., 2000), of which this paper is a part. Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Buenos Aires, Argentina Alfred Wegener Institut für Polar und Meeresforschung, Bremerhaven, Germany ¿Museu de Zoologia da Universidade de Sao Paulo,Caixa Postal 42594, 04299-970 Sao Paulo, SP, Brazil *To whom correspondence should Бе addressed; mcledon@bg.fcen.uba.ar 186 CLEDÓN ЕТ AL. MATERIALS AND METHODS Three samples were collected in March, May, and August 2001 at 15 m depth at Playa Orengo, San Antonio Oeste (40%53'S, 64*36'W), Argentina, by SCUBA diving. The animals were attached to the bivalves Atrina seminuda (d'Orbigny, 1846) and Aulacomya atra (Molina, 1782) and to stones. Approximately 370 speci- mens were collected. Live individuals were carried to the labora- tory, carefully detached from their substratum, measured to the nearest 0.1 mm precision with a digital vernier calliper, and some specimens dissected for anatomical description in vivo. Shell parameters were measured following Hoagland's (1977) definitions. “О” refers to the length of the shell arc, whereas convexity 1$ the relation between shell arc and shell length; “SL” refers to shell length. The sexual characteristics of the population were determined by the presence or absence of a penis. A total number of 47 egg masses was found and fixed in 5% seawater-formalin. Four ran- domly chosen egg capsules per egg mass were detached, and their length and width were measured under a stereomicroscope. Eggs and embryos contained within these egg capsules were counted and measured, and the presence or absence of cannibalism or nurse eggs was analyzed with a Kruskal-Wallis test. Settlement size was estimated by measur- ing the protoconch length under SEM. Simple linear regression type 2 following natural logarithmic (In) transformations was carried out to identify the parameters of taxo- nomic value. Radular characteristics of six individuals of different sizes were also studied with SEM. The anatomical study was performed using standard methodology, with non-narcotized specimens fixed in 70% ETOH. Dissections were performed under a stereomicroscope, with the specimens immersed in fixative. All draw- ings were done with the aid of a camera lucida. Abbreviations of anatomical structures are as follows: aa, anterior aorta; ab, auricle region beyond ventricle connection; ac, anterior ex- tremity of gill on mantle border; ad, adrectal sinus; af, afferent gill vessel; ag, albumen gland; an, anus; ap, aperture of visceral vas deferens into pallial cavity; au, auricle; bg, buc- cal ganglion; ce, cerebro-pleural ganglia; cg, capsule gland; cm, columellar muscle; cv, ctenidial vein; dd, duct to digestive gland; dg, digestive gland; di, septum separating haemocoel from visceral mass; dm, dorsal shell muscle; dp, posterior duct to digestive gland; en, endostyle; es, esophagus; ey, eye; fd, foot dorsal surface; ff, female folds of genital pa- pilla; fg, food groove; fl, female papilla; fp, fe- male pore; gd, gonopericardial duct; gf, gastric fold; gi, gill; gp, pedal ganglion; gs, gastric shield; ig, probable ingesting gland; in, intes- tine; iu, “U”-shaped loop of intestine on pallial roof; ki, kidney; Il, left lateral expansion (flap) of neck; Im, lateral shell muscle; m1-m14, odontophore muscles; mb, mantle border; ml, mantle region restricting pallial cavity; mo, mouth; ne, nephrostome; ng, nephridial gland; nr, nerve ring; od, odontophore; os, osphra- dium; ov, pallial oviduct; oy, ovary; pb, probos- cis; pc, pericardium; pd, penis sperm groove; pe, penis; pp, penis papilla; pr, propodium; ру, pallial cavity; rg, repugnatorial gland; rl, right lateral expansion (flap) of neck; rn, radular nucleus; rs, radular sac; rt, rectum; sa, sali- vary gland duct; sd, pallial sperm groove; se, subesophageal ganglion; sg, salivary gland; si, siphon-like fold; sr, seminal receptacles; ss, style sac; st, stomach; su, supraesophageal glangion; sv, seminal vesicle; sy, statocyst; te, cephalic tentacle; tg, integument; tm, net of transversal muscles of haemocoel; ts, testis, ve, ventricle; vg, vaginal duct; vm, visceral mass; vo, visceral oviduct. Abbreviations of institutions: AMNH, Ameri- can Museum of Natural History, New York, New York, USA; FMNH, Field Museum of Natural History, Chicago, Illinois, USA; MACN, Museo Argentino de Ciencias Naturales “B. Rivadavia”, Buenos Aires, Argentina; MZSP, Museu de Zoologia da Universidade de Sao Paulo, Sao Paulo, Brazil. RESULTS Crepidula cachimilla, new species (Figs. 1-44) Crepidula onyx Sowerby: Parodiz, 1939: 701, pl. 1, fig. 1; Scarabino, 1977: 185, pl. 3, fig. 5 (non G. B. Sowerby |, 1824). Crepidula aplysioides Reeve: Hoagland, 1977: 369 (Argentinean material only) (non Reeve, 1859). Type Material Holotype: AMNH 306947. Paratypes: AMNH 306957 to 306961, 14 paratypes (5 dry speci- mens); AMNH 306948 to 306956, 9 paratypes CREPIDULA CACHIMILLA М. SP. 187 (4 females, 5 males preserved in ethanol): MZSP 41427 (15 paratypes); FMNH (10 para- types). Type Locality Rio Negro, San Antonio Oeste, Playa Orengo, Argentina (40°53’S, 64°36’W), 15 т depth, on shells of Atrina seminuda and Aulacomya atra and on stones (Figs. 1-3). Etymology The name of the species alludes to the mapuche word meaning great friend and is dedi- cated to our colleagues at the Invertebrates | Laboratory of the Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires. FIGS. 1-3. Crepidula cachimilla on different substrata. FIG. 1: Aulacomya atra. FIG. 2: Atrina seminuda. FIG. 3: Rock. Scale bar = 3 cm. Е female; m: male. Diagnosis Shell outer surface smooth, lacking periostracum; apex projecting posteriorly, slightly away from posterior shell edge. Col- umellar muscle somewhat thick. Pallial cavity aperture restricted at right by a closure of mantle edge. Osphradium small, approximatgely 1/8 of mantle aperture length, with about 16-17 broad, closely spaced filaments. Endostyle divided by a middle longitudinal furrow. Hypo- branchial gland greatly reduced. Transversal fold of kidney at level of nephrostome. Sali- vary glands very large, slightly larger than haemocoel. Both gastric ducts to digestive gland duplicated. Male seminal vesicle very large, coiled, wall markedly irregular. Female seminal receptacles reunited in a same region, mostly 4—5; vaginal duct long, broad; genital papilla tall, with a pair of separate longitudinal folds, ending subterminally. Description Shell (Figs. 1-15): To 50 mm in length and 38 mm in width; walls with 0.46-0.60 mm thick; slightly to strongly convex (convexity = 1.095- 1.350) (Table 1, including other measure- ments). Growth lines covering entire shell and septum. Color opaque-brown, internally bright- chocolate brown. In males, always opaque- brown externally and bright-brown internally. Few individuals (about 1%) with a white shell. Periostracum totally deciduous. Male speci- mens with very thin, brittle, flattened shells (Figs. 4-15). Protoconch smooth, with 1.4 whorls; transition to teleoconch not clearly defined. Aperture elliptical or subcircular. Apex solid, generally prominent, turned to right in females, almost central in males, slightly above margin, never reaching margin in males, extending beyond itin females. Flattened sep- tum never convex, ridge central, margin with a clear central notch, covering less than half of ventral surface. Septal edge translucent, sinuous, slight turned towards right. Muscle scars inconspicuous. Head-Foot (Figs. 16-18): Head differenti- ated, on long, dorsoventrally flattened neck, about half length of foot. Proboscis short, cy- lindrical. Tentacles long, stubby, apex some- what bifid. Eyes dark, small, located on obsolete ommatophores in basal region of lat- eral margin of tentacles. Neck with pair of lat- eral, flattened lappets (nuchal lobes); left 188 CLEDÓN ЕТ AL. expansion narrower than right; right expansion bringing low food groove along its dorsal limit with head (Fig. 17: fg). Foot very ample, oc- cupying about 3/4 of shell concavity, dorsoven- trally greatly flattened, thin; clear longitudinal inner sinus running in median line; shell sep- tum as dorsal foot limit. Mantle fusing with dorsal surface of foot, protruding beyond its borders. Furrow of pedal glands transverse, in anterior margin of foot; anterior margin of foot covered dorsally by posterior region of neck ventral surface. Columellar muscle somewhat reduced, small, but somewhat thickened, contouring whole anterior border of shell septum, slightly taller at right (Figs. 17, 40: cm). Inner haemocoel cavity narrow, run- ning approximately in center of neck region. Inner space almost filled by great quantity of transverse, very slender muscular fibers; these fibers connecting ventral surface of dorsal haemocoel wall with dorsal surface of its ven- tral wall, contouring salivary glands and esophagus (Fig. 18: tm). No vestiges of oper- culum except in very young specimens, being circular, paucispiral, thin, semi-transparent, flexible. Mantle Organs (Figs. 16, 19-22): Mantle border thick, slightly hollow due to broad collar sinuses (Fig. 21). Mantle border surrounding entire shell ventral margin, free in anterior third, attaching to foot borders in posterior 2/3, situ- FIGS. 4-15. Crepidula cachimilla. FIGS. 4-6: Female holotype, AMNH 306947. FIGS. 7-9: Female paratype 1, AMNH 306949. FIGS. 10-12: Female paratype 2, AMNH 306950. FIGS. 13-15: Male paratype 5, AMNH 306955. Scale bar for FIGS. 4-12 = 4 cm. Scale bar for FIGS. 13-15 = 2 cm. CREPIDULA CACHIMILLA М. SP. 189 ated slightly away from foot edge, connecting to it by a thin, semi-transparent portion. Mantle border without appendages, but entirely edged by series of minute repugnatorial glands, im- mersed in central region of mantle edge (Fig. 21: rg). Mantle border with special arrangement of folds in middle region of pallial cavity aper- ture, a Somewhat narrow fold located from gill anterior end running towards left, decreasing and disappearing abruptly at level of osphradium, its broader region with a broad central furrow, its posterior edge expanding weakly beyond mantle border covering ven- trally anterior region of gill, its anterior edge slightly projecting, but not extending beyond mantle edge (Figs. 19, 20, 22). Dorsal shell muscle well developed (Fig. 16: dm), origin small, in about middle-right region of shell, just anterior to septum, its fibers run- ning anteriorly, spraying like fan, inserting in adjacent anterior region of dorsal surface of pallial cavity. Lateral shell muscle (Figs. 16, 19, 20: Im) small, fan-like, located close to right side of mantle border, in region where pallial cavity penetrates shell septum chamber, with a differentiated muscular branch running to- wards mantle border, thickness restricting pal- lial aperture (Fig. 20). Pallial cavity aperture occupying about 2/3 of right-anterior half of shell border (compared to a clock in dorsal view, with head at 12 o’clock, pallial aperture from 11 to 2 o'clock) (Fig. 19); right region of pallial cavity aperture restricted by a broad clo- sure of mantle border, forming a transverse septum (Fig. 20). Pallial cavity deep, broad, triangular, arched, dorsoventrally flattened. Anterior extremity of pallial cavity a little larger than its aperture because of closure in left and right extremities produced by fusion of mantle and foot (Figs. 19, 20: ml). Pallial cavity nar- rowing gradually towards posterior, penetrat- ing at left of visceral mass; cavity length about 2/3 length of animal (Figs. 16, 19). Osphradium small, monopectinate, located between anterior half of gill and mantle bor- der, at some distance from gill anterior end, located about in left region of pallial aperture somewhat perpendicular to longitudinal axis of body (Figs. 19, 20). Osphradium length little more than 1/8 of pallial aperture length, in form of a small fold, attached to mantle, separated from gill structures. Osphradium leaflets cy- TABLE 1. Measurements in mm of the holotype and paratypes. Total Specimen length (L) Height Width Holotype AMNH 52:5 8.5 36.4 306947 (female) Paratype 1 AMNH 30.9 8.4 21.8 306948 (female) Paratype 2 AMNH 42.8 9.2 2985 306949 (female) Paratype 3AMNH 38.7 7.8 27.4 306950 (female) Paratype 4 АММН 31.9 12.6 19.8 306951 (female) Paratype 5 АММН 20.1 6.4 14.7 306952 (male) Paratype 6 AMNH 23.3 St ИИ 306953 (male) Paratype 7 АММН 15.3 Sl 11.6 306954 (male) Paratype 8 АММН 12.3 3.9 10.2 306955 (male) Paratype 9 АММН 26.6 8.2 20.2 306956 (male) Septum Septum free Convexity D length shell length (D/L) 59.3 24.9 18.35 1:19 35.4 11222 16.1 1.14 47.5 28.8 ES kl 41.3 ES 19.1 1.07 42.1 13:3 15:8 1.34 23.8 8.8 9.3 116 25 10.9 10.3 1510 1722 5.8 8.3 1612 14.2 3.9 6.4 als 30.9 128 133 1.16 190 CLEDÓN ЕТ AL. lindrical, close from each other, somewhat thick, low, about 16-17 in number (Fig. 22: 0$) in females. Osphradium ganglion narrow. Gill very large, its base narrow, edging ante- rior and left margin of pallial cavity almost the entirety of its length; anterior gill extremity in right-anterior region of pallial cavity aperture, near its right limit, on thick mantle border; gill posterior extremity in posterior end of pallial cavity (Fig. 20). Gill filaments triangular at their base and with very long, almost straight, nar- row, stiff rod turned to right (Fig. 21: gi); rods FIGS. 16-21. Crepidula cachimilla female anatomy. FIG. 16: Whole dorsal view, specimen extracted from shell. FIG. 17: Same, head-foot, dorsal view, visceral mass and pallial structures removed. FIG. 18: Head and haemocoel, ventral view, foot and neck “sole” removed. FIG. 19: Pallial cavity and visceral mass extracted, ventral view. FIG. 20: Same, left pallial connection sectioned, ventral portion of visceral mass deflected, most gill filaments artificially sectioned. FIG. 21: pallial cavity roof, transversal section in region tangent to rectum. Scales = 2 mm. CREPIDULA CACHIMILLA М. SP. 191 extending about three times longer than their triangular, membranous base; rods beginning in ctenidial vein region, in left margin of cavity roof, and touching food groove of head-foot, in right margin of cavity floor; rod apex rounded, preceded by thicker region. Gill fila- ments connected to each other by cilia, mainly in their thicker apical region, holding them in a somewhat firm position. Gill filaments longer in central gill region, shorting gradually in both extremities; gill anterior extremity with short filaments, abruptly turning forwards, located on mantle border (Fig. 22). Ctenidial vein nar- row, with uniform width along its entire length. Endostyle well developed (Figs. 20, 21, 22: en), yellowish, in form of broad, flat glandular ridge located in middle level of ventral surface of ctenidial vein along its entire length. Endo- style divided longitudinally by a shallow middle furrow. Hypobranchial gland extremely thin, practically absent. About 1/3 of visceral mass encroaching on pallial cavity roof (Fig. 20), occupying about 1/3 of this cavity in poste- rior-right region; pericardium and kidney lo- cated posteriorly; a long intestinal loop, anus and pallial oviduct located anteriorly. Visceral Mass (Figs. 16, 19, 20): Form of a dorsoventrally flattened cone, housed in shell chamber produced by septum, which sepa- rates visceral mass from dorsal surface of foot. Left and anterior region of visceral mass oc- cupied by pallial cavity (Figs. 16, 19, 20). Re- maining regions of visceral mass with stomach as central structure, immediately surrounded by greenish-beige digestive gland (except in some ventral and dorsal areas). Gonad sur- rounding digestive gland, more concentrated anteriorly and at left. Visceral mass encroach- ing on right-posterior region of pallial cavity roof, possessing another ventral flap as pal- lial cavity floor (Fig. 20: vm). Anterior extrem- ity of visceral mass ventral flap ending at anterior border of shell septum, covering col- umellar muscle (Fig. 17). Circulatory and Excretory Systems (Figs. 20, 25): Pericardium somewhat triangular, broad, oblique to longitudinal axis of animal (Fig. 16: pc). Pericardium left region very narrow, in form of a vein connecting gill with auricle, be- ginning at posterior extremity of gill in poste- rior-left end of pallial cavity, running to surround area where visceral mass encroaches into pallial cavity, gradually increasing towards anterior and right (Fig. 25). Remaining peri- cardium limits: (1) anterior and ventral — pal- lial cavity; (2) posterior — visceral mass (go- nad generally); (3) dorsal — mantle; (4) right — kidney. Auricle thin walled, long, narrow, run- ning all along pericardium length, attached to its anterior and dorsal inner surfaces (Fig. 25), connecting with ventricle approximately in its middle portion; auricle having a broad portion beyond ventricle as blind sac (Fig. 25: ab), bearing orifice to nephridial gland. Ventricle elliptical, very muscular; its connection with auricle located about in middle region of its anterior surface, on opposite side bearing ori- gin of aortas. Anterior aorta broad, running towards opposite side from posterior aorta. Anterior aorta running towards right, surround- ing posterior inner pericardium surface, then penetrating head haemocoel. Kidney occupying about half of area of vis- ceral mass within pallial cavity (Fig. 20). Kid- ney limits: (1) dorsal — mantle; (2) ventral — pallial cavity; (3) posterior-right — visceral mass (gonad generally); (4) posterior-left — pericar- dium; (5) anterior — an intestinal loop; (6) lat- eral-right — intestine and oviduct (when present). Kidney central region hollow, with single anterior lobe (Fig. 25). Kidney lobe slightly uniform, covering dorsal surface, in- testinal region passing through kidney cham- ber, and about 1/4 of inner space of kidney adjacent to intestine. Renal lobe having longi- tudinal, branching, narrow folds; a larger fold located at left of nephrostome having a series of anterior branches situated somewhat uni- formly; another transversal, somewhat tall fold located at level of nephrostome. Nephridial gland in renal limit with pericardium, very small, having a series of triangular, transversal, nar- row folds connected with dorsal renal lobe (Fig. 25: ng). Nephrostome a very small slit located in central region of ventral wall (Figs. 20, 25: ne), in anterior region of hollow portion of kid- ney; no inner glandular folds close to it. Adrectal sinus very broad, edging externally intestine loop exposed in pallial cavity, con- nected to main kidney chamber but separated by a thin septum (Figs. 20, 25: ad). Digestive System (Figs. 16, 26-28): Probos- cis short, broad (Figs. 16-18: pb). Pair of nar- row ventral proboscis retractor muscles very thin, immersed in proboscis wall. Mouth lon- gitudinal, in center of anterior proboscis sur- face. Buccal mass very large, occupying most of proboscis inner space and short portion of haemocoel posterior to it. Jaw plates in dorsal 192 CLEDÓN ЕТ AL. wall of buccal mass thin, almost vestigial, broader laterally, short longitudinally. Pair of broad, low dorsal folds beginning well poste- rior to jaws; dorsal chamber between these folds shallow. Odontophore large, occupying most of buccal mass. Odontophore muscles similar to other spe- cies of Crepidula (Simone, 2002) (Figs. 26, 27: m1); several very narrow jugal muscles connecting buccal mass with adjacent wall of snout, more concentrated anteriorly around mouth: m1b pair of dorsal protractor muscles narrow, thin, superficial, originating in anterodorsal region of mouth, close to median line, inserting in posterodorsal-lateral region of odontophore; m1v similar to m1b but located in ventral surface; m2 pair of retractor muscles of buccal mass (retractor of pharynx) broad, FIGS. 22-28. Crepidula cachimilla anatomy. FIG. 22: Anterior portion of pallial cavity, close to mantle border, mantle border slightly deflected for showing anterior gill region. FIG. 23: Central nervous system (nerve ring), ventral view. FIG. 24: Same, dorsal view. FIG. 25: Middle and posterior-right region of pallial roof, ventral view, ventral wall of pericardium and kidney partially removed. FIG. 26: Buccal mass, ventral view. FIG. 27: Same, dorsal view, salivary gland (sg) fully shown. FIG. 28: Middle and distal digestive tubes shown as in situ, ventral view, some adjacent structures also represented. Scales = 1 mm. CREPIDULA CACHIMILLA М. SP. 193 originating in lateral-ventral region of haemocoel just posterior to snout (Fig. 18), running towards anterior, inserting in lateral- posterodorsal region of odontophore cartilages; m2a pair of dorsal tensor muscles of radula, continuation of m2 after insertion in cartilages, running towards anterior, inserting in subradular cartilage in middle region of its dorsal inner surface; mt dorsal transversal muscle, or approximator muscle of cartilages, connecting dorsally both posterodorsal-lateral surfaces of cartilages, lying between superfi- cial membrane that covers odontophore and tissue on middle region of radula; m4 pair of median dorsal tensor muscle of radula very large, thick, originating in ventral-middle-pos- terior region of odontophore cartilages, run- ning towards medial, contouring medial-ventral surface of cartilages, running on their dorsal surface, inserting in subradular cartilage dor- sal-posterior-medial extremities; m5 pair of median radular tensor muscle thick, originat- ing in median-posterodorsal region of odontophore cartilages, near side of m2 in- sertion and m2a origin, covering perpendicu- larly m4 middle region, running medially, inserting along both sides of radular sac (each m5 branch covering a side of radular sac, medially and dorsally); m6 horizontal muscle very thin, uniting anterior half of odontophore cartilages, inserting on their dorsal margin; m7 pair of ventral tensor muscle of radula thin, narrow, originating inside radular sac ventral surface close to each other, running anteriorly, separating gradually from each other, insert- ing in radula ventral border; m8 pair of strong muscles originating in posterodorsal-lateral regions of odontophore cartilages near inser- tion of m2, running attached to dorsal margin of odontophore cartilages, inserting in their anterodorsal region close to horizontal muscle (m6); m9 pair of dorsal-medial tensor muscle of radula broad, thin, originating along dorsal- median surface of radular sac (in its region internal to odontophore), crossing to dorsal surface, inserting in dorsal-ventral border of subradular cartilage; mj jaws and peribuccal muscles somewhat thick, surrounding lateral and dorsal wall of buccal mass, originating around mouth, inserting in middle level of lat- eral and dorsal wall of odontophore; m11 pair of ventral tensor muscles of radula weakly present; m14 pair broad, thin, originating in posterodorsal region of odontophore, close to m2 and m5 origins, running towards ventral and anterior, inserting in snout inner ventral surface in about middle level of odontophore; tissue covering middle region of radula within odontophore, on its dorsal surface. Radula short, little more than odontophore length (Figs. 29, 30); rachidian tooth narrow, strongly curved inwards, central cusp large, sharp, secondary cusps 2-4 similar-sized pairs (formula 2-1-2/0-0 to 4-1-4/0-0), weak pair of lateral reinforcements on its borders; lateral tooth broad (about three times broader than rachidian), curved inward, with about 7-10 short, triangular cusps, along edge on mar- ginal side and 1-3 very weak cusps on edge on rachidian side, cusps decreasing laterally, disappearing about in middle region of tooth, with thick, arched border (formula from 1-1-7/ 0-0 to 3-1-10/0-0); inner marginal tooth long, curved, tall, tip sharply pointed (cusp formula 0-1-5/0-0 to 2-1-7/0-0); outer marginal tooth narrower than inner marginal tooth, thin, and with two small cusps along its inner margin only (cup formula 0-1-2/0-0 to 0-1-3/0-0). Pair of buccal ganglia large, close to each other near median line (Fig. 26: bg), located between buccal mass and adjacent esopha- FIGS. 29, 30. Radula of Crepidula cachimilla. FIG. 29: General view of the radula. Scale bar = 100 um. FIG. 30: Detail of the central and lateral tooth. Scale bar = 64 um. 194 CLEDÓN ЕТ AL. gus. Salivary glands not passing through nerve ring, longer than haemocoel, fitting inside it, bent (Figs. 18, 27: sg); distal end rounded, of about 1/3 of haemocoel width, running towards anterior possessing approximately same width along its length, narrowing close to buccal mass. Ducts of salivary glands broad, sinuous (Fig. 27: sa), running in dorsal surface of buc- cal mass, penetrating adjacent buccal mass wall a short distance, apertures small, in ante- rior region of dorsal folds of buccal mass. Esophagus (Figs. 18, 28: es) narrow, long; anterior esophagus inner surface with pair of broad folds, running straight posteriorly, be- coming gradually slender. Stomach (Fig. 28) somewhat conical, large, occupying about half of visceral mass size; esophagus inserting in left side of its posterior-left region, close to shell apex. Anterior duct to digestive gland located in region of stomach ventral surface preced- ing style sac, separated into two similar-sized, well-spaced ducts, each running in opposite directions, highly dichotomic. Posterior duct to digestive gland also duplicated (distance be- tween this pair greater than that of anterior ducts), each one running in opposite direc- FIGS. 31-35. Crepidula cachimilla anatomy. FIG. 31: Head-foot, male, dorsal view, pallial structures and visceral mass removed. FIG. 32: Visceral mass and adjacent part of pallial cavity, male, ventral view. FIG. 33: Penis and adjacent structures, dorsal view, penis deflected. FIG. 34: Visceral vas deferens extracted, seminal vesicle (sv) uncoiled. FIG. 35: Pallial oviduct, ventral view as in situ, most integument and pallial cover removed (except close to papilla). Scale bars = 1 mm. CREPIDULA CACHIMILLA М. SP. 195 tions, both very narrow, located in ventral re- gion of stomach almost at its posterior end, one of them turned posteriorly. Stomach gradually narrowing towards an- terior and left, arriving close to left-posterior extremity of pallial cavity. Gastric shield oc- cupying about 1/3 of stomach inner surface, located in its right side (Fig. 28: gs). Pair of longitudinal folds separating intestine from style sac running at left (Fig. 28: gf), in re- gion anterior to anterior ducts to digestive glands, abruptly separating one another per- pendicularly, in a T-fashion, surrounding en- tire stomach circumference in this region, forming a low, narrow fold separating style sac from main gastric chamber. À weak con- striction marking region between style sac and main gastric chamber, clearer at right. Digestive gland pale brown in color, surround- ing stomach except some areas on dorsal and ventral surfaces (Figs. 16, 20, 24). Intestine narrow, sinuous (Fig. 28: in), running on an- terior border of visceral mass from left to right, initially in its ventral region, slightly near me- dian line cross to its dorsal region and run- ning up to right-anterior extremity of visceral mass (Fig. 28); running towards left in this region, becoming broader and exposed in pallial cavity, surrounding right and anterior border of kidney, abruptly running towards right in a U-shape, parallel to preceding loop (Figs. 16, 21, 28, 25: iu). Anus small, si- phoned, located in right region of pallial cav- ity close to mantle border (Figs. 21, 28, 25). Final intestine loops filled with several small, elliptical fecal pellets. Male Genital System (Figs. 31-34): Mature males up to 28 mm in shell length. Testis white, located mostly in anterior region of visceral mass (Fig. 32: ts). Sperm duct differentiable in region of testis just at right of esophagus penetration into visceral mass. Seminal vesicle intensely coiled, locally accumulated in ante- rior-right region of visceral mass (Fig. 32: sv); if uncoiled, presenting about same length as visceral mass; wall glandular, greatly irregu- lar, varying from broad to very narrow along its length (Fig. 34). Seminal vesicle abruptly narrowing near pallial cavity, having a very nar- row aperture located in right-posterior end of this cavity (Figs. 32, 34: ap). Pallial sperm groove starting immediately below this aper- ture, running as a relatively deep, narrow fur- row with elevated edges. Pallial sperm groove running along right neck lobe close to its edge (Fig. 31: sd), slightly dorsal; abruptly curving towards left close to penis base, connecting to its posterior base region (Fig. 33). Penis lo- cated behind right cephalic tentacle, curved in same direction, of about 3-4 times its size (Fig. 31). Distal papilla long, about 1/4 of length of remaining penis region, about 1/5 of its width (Fig. 33). Penis groove deep, central, running along ventral surface up to penis papilla tip (Ею: 33). Female Genital System: Ovary cream yellow, surrounding digestive gland, more concen- trated in anterior region of visceral mass (Fig. 19: оу); when mature, oocytes distinguishable by their transparency. Visceral oviduct formed by gradual decrease from right-anterior end of ovary. Gonopericardial duct narrow, relatively short, originating in right-ventral extremity of pericardium, running ventral to visceral glands in area in which visceral mass encroaches to- ward pallial roof, inserting in posterior extrem- ity of pallial oviduct, joined with insertion of visceral oviduct (Fig. 35: gd). Pallial oviduct narrow, located in right-anterior end of pallial cavity (Figs. 16, 20: ov). Seminal receptacles (sr) located in right side of last portion of vis- ceral oviduct, four to five in number, with three always significantly larger (Fig. 35: sr); each a small sac; duct very narrow, long; their inser- tion preceding albumen gland, on right surface. Albumen gland long, narrow, whitish, its walls thick, glandular; located in anterior-right ex- tremity of visceral mass, about half size of cap- sule gland (Fig. 35: ag). Separating albumen from capsule glands a narrow differentiable, paler colored tissue, most probably an ingest- ing gland (Fig. 35: ig). Capsule gland a con- tinuation of albumen gland, but situated perpendicular and slightly dorsal to it, broad, spherical (Fig. 35: cg); walls thick glandular, pale brown; inner duct narrow, U-shaped, length about 1/8 of pallial cavity aperture. Vagi- nal duct (vg) relatively broad, equal in size to albumen gland. Genital pore preceded by tall, long papilla close to mantle border, at right and slightly removed from anus (Fig. 20: fl). Geni- tal papilla with broader base and somewhat conical form; pair of well-spaced low folds run- ning along its posterior-left side; both start gradually in papilla base and terminate at some distance from pore (Fig. 35: ff); posterior fold originating on surface of pallial cavity floor; anterior fold originating from pallial roof. Geni- tal pore a transverse apical slit, perpendicular to papilla folds (Fig. 35: fp). 196 CLEDÓN ЕТ AL. Reproduction Animals categorized into four sexual phases: (1) undifferentiated juveniles, (2) males, (3) transitional individuals, and (4) females. These are easily recognizable under a microscope by observation of the external development of the reproductive organs. Juveniles are without vis- ible sexual organs. Males have a well-devel- oped penis. Transitional individuals have a penis in retraction phase and a developing genital papilla. Females lack a penis and have an easily distinguishable papilla. Undifferentiated juveniles were between 3.7 and 5.1 mm SL (mean: 4.3 SD: 0.4 N: 11). Males were 5.4-28.5 mm SL (mean: 14.1 SD: 0.8 М: 103), always attached to larger individu- als. Females were 9.6-52.2 mm SL (mean: March Relative Frequency (%) 16 22 | | | | | ых м 34 40 46 52 28 Shell length (mm) November 15 - 12 Relative Frequency (%) (o 4 10 16 22 6 = | | | 0 aI T T T T T T | la T T T la I 1 1 28 34 40 46 52 Shell length (mm) FIG. 36. Crepidula cachimilla sex proportion in March (N: 270) and November (N: 286). White: males; black: females. CREPIDULA CACHIMILLA М. SP. 197 37.3 SD: 0.9 М: 252), forming stacks of 2-5 individuals. The smallest brooding female was 23.5 mm SL; the largest was 49.5 mm SL. Peak of female development from August to April (observed in 47 brooding females). No- table period of reproductive rest between March and November. No juveniles were en- countered in the field during winter, being re- flected in the diminution in the proportion of males in the population and their larger shell length in comparison with the summer (Fig. 36). Egg masses with 15-65 capsules (Fig. 37) (mean: 35, SD: 14, N: 47). Egg capsules (Fig. 38) 2.2-3.3 mm in length (mean: 2.8, SD: 0.5, N: 148) and 2.3-3.4 mm in width (mean: 2.6, SD: 0.4, N: 148). Each egg capsule containing 129-441 uncleaved eggs (Fig. 39) (mean: 226, SD: 57, М: 148) in a whitish viscous liquid. А! eggs developing into veliger larvae (Fig. 40) and hatching. It was not possible to measure hatching time. No nurse eggs or cannibalism was observed. No differences in number of embryos between initial and late brood stages in females of same size found. Neither In-transformed mean capsule size (r? = 0.01) nor In-transformed capsule number per mass (г? = 0.13) correlated with shell length of brooding females. There was no positive corre- lation between In-transformed egg number per capsule and In-transformed capsule size (г? = 0.33) or In-transformed female size (r? = 0.02). Uncleaved egg diameter 180-200 um (mean: 191.7; SD: 7.2, N: 20). Protoconch length of juvenile shells (Figs. 41, 42) 700-800 um (mean: 760 SD: 65 N: 11). Habitat Between 10 and 20 m depth, attached to hard substrata. Distribution Known only from northeastern Patagonia including the records from Golfo San Matias to Punta Norte of Parodiz (1939). FIGS. 37-42. Crepidula cachimilla egg mass and protoconch. FIG. 37: Egg mass. Scale bar = 7 mm. FIG. 38: First and late stage egg capsule. Scale bar = 3.5 mm. FIG. 39: Eggs in first division stages. Scale bar = 500 um. FIG. 40: Prehatching stage. Scale bar = 350 um. FIG. 41: Protoconch on an adult shell. Scale bar = 800um. FIG. 42: Detail with SEM of protoconch of an adult shell. Scale bar = 1,100 um. CLEDÓN ЕТ AL. 198 (запициоэ) (€9Z :N ‘ev F LLP (2G ‘N ‘LL F8€ (LS :N ZF6 (6/1 ‘ujewo) sah ‘ueaw) 089-008 :ue9W) G9-ZL ‘UBSU) 6-7 $002 ‘Je 19 yaıneisojy\ snjeginoe snındea/nsog 216) ‘орлейео ou 8£c-voc Суб чеэш = 1161 ‘opielles ерипэа! I 116} ‘орлейео sok - Ler-Srl ve-9L 1161 ‘орлееэ eueiddyyd à LOOZ 'yapezseyouag 0002 'yapezseyouag 8 ou LOL vr 8vbv-18 901 ® UOpaj) oullo3seg ‘эцоцис eunuabie ‘7 Ly8L ‘Либо ou 011 051-08 879 $861 ‘ривбеон esjold 9 1181 MoJeuwe7 Sok = 910 1-80€ GS 1161 ‘орлееэ EJEJEIIP I 9661 UMOIg = $9961 GEL-Z9 - 9661 'UMOJY sisuaquinboo ‘7 LOOZ ‘yepezseyoudd 6581 ‘элээч ou 00$ CITE G+EZ © yoıneisol\ sapıoıs/lde I (02. IN T'LFLL6L (8PL:N ‘vl + 182 (Lv N ‘VL FSE ou ‚uesul) 002—081 (UBB) E9G-6Z1 ues) 9-5) Ápnys $14} eywiy9es I (ansdes/9) (ZZ) ueeuu) (0zz :чеэш) (бу :ueau) yz81 ‘| Âgemos ‘g 9 sof 081—091 005—001 09-61 9861 ‘pue|beoy xAuo I eg ‘| Agemos y 9 UOISSNOSIP 995 ejep ou 005—001 001-09 Суб! 209 XAUO empidalo $бБэ asınN (url) э}эшер ginsdeo Jad sseu 66e sed uononpoidei sarmads 66a panespun $бба jo лэашйм aınsdeo jo Jaquiny бирлебэ/ adualajay ‘в|ииицоеэ ‘D чим рэледшоэ еэмэил\у Lou Seloods e/npide1 ло; зэцзиэоелецо элцопролаэч ‘2 3719VWL 199 CREPIDULA CACHIMILLA N. SP. иблеш ayy Bulyoea, (G:N '9FEL (1621 'uawo) Janau хэае 'padeys-S - 95—61 - ‘иеэц/) бицме. 0Z-+ smean2e snındesAnsog иблеш Bulyoeas 1161 ‘oprees Janau xade ‘pedeus-s - - - - epunag 9 иблеш бицоеэл 1161 ‘орленея Janau хэае 'padeys-S - - - BulmeJo eueidaiiyd 9 un O€Z—-061 0002 'yapezseyouay $ = 95 95“, 98'8—96'5 эелле| sabian очио}$е4‘эцоцис еициэбие I Ly81 ‘Либо A = 07€ 2 0'8-9'p oene] JSBISA eajold ‘9 иблеш Bulyoeas 1181 ‘youewe 7] Janau хэае 'padeys-S - - - эпиэли! Buimeo вер‘ иблеш Buiyoeel 9661 ‘UMOIg Janau хэае 'padeys-S - - - эниэлиГ BuimeJo sisuaquiinboo ‘7 url 009 6581 ‘198 3 z c8l-v6 = ajlueanf Bulmeso sapıoıs/lde 9 зэллбу pue шп 005-092 иоцанозер 24$ sas t'es 6b-S' EZ 9'85-7'5 эелие| лэбцэл вииицоео I vegı ‘| Áquamos ‘g ‘9 3 09 09-17 - эелле| лэбиэл XAUO 9 Sene] vz8! ‘| Áqiamos 'g 9 Е 09 2 0$ лэбиэл эцаоомиеа XAUO ejprdalo adeys unidas (uu) yyBuay Jays (uw) чбиа| ¡Jays (wu) эбе}$ бичэзен зэюэа$ эеша; wnwıxey\ эеше; Buipooig чубиз! 118ys ele (репициоэ) 200 CLEDÓN ЕТ AL. DISCUSSION The shell of Crepidula cachimilla is similar to species occurring in the western Atlantic belonging to the “Crepidula plana complex” (Collin, 2000; Simone, submitted). lts most distinctive characters are the projecting apex, located somewhat away from the posterior shell base, and the absence of periostracum. The characters of Crepidula cachimilla pre- sented in the Diagnosis, mostly morphologi- cal, as well as those summarized in the Table 2, are the main basis differentiating this spe- cies. That set of characters easily separates the new species from the remaining South American taxa. From the Atlantic species with known anatomy, C. cachimilla has a thicker columellar muscle, a condition found only in other species in early stages of the develop- ment, after which the columellar muscle be- comes reduced. As stated by Simone (2002), based on comparison of the ontogeny and phylogeny, the lateral and dorsal shell muscles are also derived from the columellar muscle, and are both thick in C. cachimilla; however, the respective scars in the shell are incon- spicuous. The restriction of the pallial cavity aperture is one of the synapomorphies of the family Calyptraeidae (Simone, 2002); how- ever, in С. cachimilla this state 1$ still more developed, as itis greatly restricted on the right side by a broad fusion of the mantle border. The hypobranchial gland is normally reduced in Crepidula, being a thin glandular layer sur- rounding the visceral structures encroached into pallial cavity roof (Simone et al., 2000; Simone, 2002); however, C. cachimilla has practically no developed hypobranchial gland, the region where it would occur being thin and transparent. The contrary happens with the salivary glands, which are normally small; in C. cachimilla, these glands are longer than the haemocoel, being folded inside this cavity. This state is comparable with that of Bostrycapulus aculeatus (Gmelin, 1791) (also known as Crepidula aculeata); however, in that species, these glands are still larger (Simone, 2002). Other notable feature of С. cachimilla 1$ the duplication of both ducts to digestive gland in the stomach. Despite the conchological peculiarities of C. cachimilla, shell characters alone do not clearly distinguish it from C. onyx, which it resembles in shape, color, and size. This similarity led Parodiz (1939) to assume that the studied species was C. onyx. Such misidentifications are common in this family, with C. argentina (Simone et al., 2000) having been confused with C. protea in Argentina. There are subtle differences in shell shape between C. cachimilla and C. onyx. Crepidula cachimilla tends to have a more pointed apex, and the shell also seems to be less convex, but these features can be strongly affected by the substratum. Anatomical differences between С. cachimilla and C. onyx are not yet known, because there has not been a detailed ana- tomical study of the latter. However, the radu- lar morphology of C. onyx (Hoagland, 1977) is markedly different from that of the studied species, in which the central tooth has 2-4 cusps (formula 2-1-2/0-0 to 4-1-4/0-0); the lat- eral tooth 7-10 cusps, (formula 7-1-0/0-0 to 10-1-0/0-0); and the inner marginal tooth has 1-3 cusps (1-1-0/0-0 to 3-1-0/0-0). In addition, the uncleaved eggs of the Argentinean mate- rial are larger than those of the Californian C. onyx population described by Hoagland (1986). The main difference between the spe- cies is the occurrence of six “malformed”, ог nurse eggs per sac (Hoagland, 1986) and the fact that “frequently fully half the entire num- ber of embryos disintegrate within the capsules and are used as food by the survivors” (Coe, 1942). Although we are unable to assess the frequency of this phenomenon in California, such malformed eggs or disintegrating em- bryos were absent in the studied Argentinean material. Aguirre & Farinati (2000) reported the pres- ence of C. onyx among other Crepidula spe- cies from Quaternary sediments in Argentina. Because of the shell of the species described here is very similar to that of C. onyx, it is rea- sonable to assume that these fossil records belong to the species described here. The occurrence of these fossils proves that this is not an exotic species recently introduced to the area. Additional differences between the reproductive parameters reported by Hoagland (1986) and Coe (1942) for C. onyx and C. cachimilla are: the larger egg diameters and the complete lack of nurse eggs or canni- balism in C. cachimilla, and the different radu- lar morphology. On this basis, the material studied by Parodiz (1939) should be assigned to the new species described here instead of being assigned to C. aplysioides, as proposed by Hoagland (1977). Crepidula cachimilla also differs from other species in many reproductive strategy char- CREPIDULA CACHIMILLA М. SP. 201 acteristics. Crepidula aplysioides Reeve, 1859, is a small (up to 2.0 cm SL; brooding female between 9.4 and 18.2 mm SL) tropical and subtropical species with egg capsules contain- ing fewer eggs than C. cachimilla (Hoagland, 1977). Further reproductive characteristics are given by Miloslavich & Penchaszadeh (2001). The number of eggs per capsule separates C. cachimilla from C. coquimbensis Brown & Olivares, 1996; С. dilatata Lamarck, 1822; and C. protea Orbigny, 1841 (Table 2). In C. cachimilla (Table 2), the egg diameter clearly differs from that of С. argentina (Cledón & Penchaszadeh, 2001), C. philippiana (Gallardo, 1977, 1996), C. fecunda (Gallardo, 1979) and C. dilatata (Gallardo, 1977; Chaparro & Paschke, 1990) (Table 2). In C. cachimilla (Table 2), eggs per capsule are more numerous, and both males and females are larger than those of С. protea (Hoagland, 1983). The larval shell at hatching and the protoconch of juveniles are larger in C. cachimilla (Table 2) than in C. argentina (Cledón & Penchaszadeh, 2001) (Table 2). According to our observations on C. cachimilla, broods containing a large number of capsules (more than 40) always belong to females larger than 31 mm SL. Because of the number of eggs per capsule does not de- pend on the female size, we used this param- eter as species representative. А more extensive comparison of the mor- phology of С. cachimilla with other species of the “Crepidula plana complex” is being pub- lished elsewhere (Simone, submitted), with a phylogenetic analysis of all known species occurring from Florida to Patagonia. Crepidula cachimilla is separated from the remaining species by such plesiomorphies as the thick- ness of the shell muscles (columellar, lateral and dorsal muscles), which are very thin in the other species; the nephridial gland having clearly transverse septa, whereas in the re- maining species this gland has irregular lon- gitudinal folds; the larger size of the salivary glands, which are normally reduced; and re- tention of the ventral tensor muscle of the radula (m11), mostly lost in other species. ACKNOWLEDGEMENTS This study was financially supported by SECyT-Argentina Pict 98-01-04321 (P. E. Pen- chaszadeh р. 1.), UBACyT-X917 (P. E. Pen- chaszadeh р. 1.), Comisión de Investigación Científica de la Provincia de Buenos Aires (CIC), SECyT-Argentina Pict 01-7222 (A. Rumi р. i.), Deutscher Akademischer Austausch- dienst (DAAD) and Fundación Antorchas (M. Scelzo р. i.). We thank Guido Pastorino and Fabrizio Scarabino for their useful comments to an ear- lier MS version. We are especially grateful to Paula Mikkelsen and Sonja Guetz for improv- ing the present version. The morphological study was also part supported by the Fundaçäo de Amparo à Pesquisa do Estado de Säo Paulo, process # 00/11074-5 and 00/11357-7. LITERATURE CITED AGUIRRE, М. L. & Е. А. FARINATI, 2000, Molus- cos del cuaternario marino de la Argentina. Boletin de la Academia Nacional de Ciencias, Córdoba, Rep. Argentina, 64: 235-333. BROWN, D. 1. & С. А. OLIVARES, 1996, А new species of Crepidula (Mollusca: Meso- gastropoda: Calyptraeidae) from Chile: addi- tional characters for the identification of eastern Pacific planar Crepidula group. Journal of Natu- ral History, 30: 1443-1458. CHAPARRO, О. R. & К. A. PASCHKE, 1990, Nurse egg feeding and energy balance in em- bryos of Crepidula dilatata (Gastropoda: Calyptraeidae) during intracapsular develop- ment. Marine Ecology Progress Series, 65: 183-191. CLEDON, M. 8 P. E. PENCHASZADEH, 2001, Reproduction and brooding of Crepidula argentina Simone, Pastorino & Penchaszadeh, 2000 (Gastropoda: Calyptraeidae). The Nau- tilus, 115: 15-21. COE, W. R., 1942, The reproductive organs of the prosobranch mollusk Crepidula onyx and their transformation during the change from male to female phase. Journal of Morphology 70: 501-512. COLLIN, R., 2000, Phylogeny of the Crepidula plana (Gastropoda: Calyptraeidae) cryptic spe- cies complex in North America. Canadian Jour- nal of Zoology, 78: 1500-1514. DALL, W. H., 1909, Report on a collection of shells from Peru, with a summary of the littoral marine Mollusca of the Peruvian zoological province. Proceedings of the U. S. National Museum, 37(1704): 147-294, pls. 20-28. GALLARDO, C., 1977, Crepidula philippiana n. sp. nuevo gastrópodo Calyptraeida de Chile con especial referencia al patrón de desarrollo. Studies on Neotropical Fauna and Environ- ment, 12: 177-185. GALLARDO, C., 1979, Especies gemelas del género Crepidula (Gastropoda, Calyptraeidae) en la costa de Chile; una redescripción de C. dilatata Lamarck y descripción de C. fecunda n. sp. Studies on Neotropical Fauna and Environment 14: 215-226. 202 CLEDÓN ЕТ AL. GALLARDO, C., 1996, Reproduction in Cre- pidula philippiana (Gastropoda, Calyptraeidae) from southern Chile. Studies on Neotropical Fauna and Environment, 31: 1-6. HOAGLAND, K. E., 1977, Systematic review of fossil and recent Crepidula and discussion of evolution of the Calyptraeidae. Malacologia, 16: 353-420. HOAGLAND, K. E., 1983, Ecology and larval development of Crepidula protea (Proso- branchia: Crepidulidae) from southern Brazil: a new type of egg capsule for the genus. The Nautilus, 97: 105-109. HOAGLAND, К. E., 1986, Patterns of encapsu- lation and brooding in Calyptraeidae (Prosobranchia: Gastropoda). American Mala- cological Bulletin, 4: 173-183. MILOSLAVICH, Р. & P. Е. PENCHASZADEH, 2001, Reproduction of Crepidula aplysioides Reeve (Caenogastropoda) from La Restinga Lagoon, Venezuela. P. 224, in: L. SALVINI, J. VOLTZOW, H. SATTMANN & G. STEINER, eds., Ab- stracts, World Congress of Malacology 2001, Vienna, Austria (Unitas Malacologica). MILOSLAVICH, Р., Р. Е. PENCHASZADEH & А. К. CARBONINI, 2003, Reproduction of Crepidula aculeata (Gastropoda, Calyp- traeidae) from the southern Caribbean (Ven- ezuela). Veliger, 46(3): 269-274. PARODIZ, J. J., 1939, Las especies de Crepidula de las costas argentinas. Physis, 17: 685-709. SIMONE, L. R. L., 2002, Comparative morpho- logical study and phylogeny of representatives of the Superfamilies Calyptraeoidea and Hipponicoidea, (Mollusca, Caenogastropoda). Biota Neotropica, 2(2): 1-102. SIMONE, L. R. L., submitted, Morphologic and phylogenetic study of the western Atlantic Crepidula plana complex (Caenogastropoda, Calyptraeidae), with description of three new species from Brazil. Zootaxa. SIMONE, L. К. L., С. PASTORINO & P. E. PEN- CHASZADEH, 2000, Crepidula argentina (Gastropoda: Calyptraeidae), a new species from the littoral of Argentina. The Nautilus, 114: 127-141. Revised ms. accepted 16 February 2004 RESEARCH NOTES MALACOLOGIA, 2004, 46(1): 205-209 CHROMOSOMES OF THE CHINESE MUSSEL ANODONTA WOODIANA (LEA 1834) (BIVALVIA, UNIONIDAE) FROM THE HEATED KONIN LAKES SYSTEM IN POLAND Pawel Woznicki Department of Evolutionary Genetics, University in Olsztyn, Oczapowskiego 5, 10-718 Olsztyn, Poland; pwozn@uwm.edu.pl ABSTRACT The chromosome complement of freshwater mussel Anodonta woodiana was investigated using Giemsa, Ag-NOR and chromomycin A; staining. The diploid chromosome number of this species is 2n = 38, and the arm number (FN) = 76. Nucleolar organizer region (NOR) was found on one chromosome pair, and it was connected to GC-rich chromatin, as visualized by СМАз staining. Key words: Anodonta woodiana, chromosomes, freshwater bivalve, karyotype, NOR. INTRODUCTION The freshwater bivalve mollusk Anodonta woodiana is native to eastern Asia. In recent years, it has been discovered in Europe (Kiss & Pekli, 1988; Beran, 1997) and on several Indonesian islands (Watters, 1997). It has also been collected in the wild in the Dominican Republic and Costa Rica (Watters, 1997). In the heated Konin Lakes of Poland, Anodonta woodiana appeared in the mid- 1980$ following the introduction of silver carp, Hypophthalmichthys molitrix (Val.), from Hun- gary (Afanasjev et al. 2001; Kraszewski 8 Zdanowski, 2001). Anodonta woodiana was observed as a dominant species in some parts of this system of lakes (Protasov et al., 1994). Within Unionidae, the chromosome number is known for 26 species, and most of them have 38 (reviewed in Nakamura, 1985; Barsiene, 1994; Thiriot-Quiévreux, 2002). Five species of Anodonta have been studied cyto- genetically, but only diploid chromosome num- ber (2n = 38) and fundamental arm number (FN = 76) have been established (Nakamura, 1985; Barsiene, 1994). The present report describes the karyotype and location of nucleolar organizer regions (NORs) of Anodonta woodiana from Poland. 205 MATERIALS AND METHODS Nineteen specimens of Anodonta woodiana from the Konin Lakes in central Poland were studied for chromosome complement. A 0.4% solution of cobalt chloride was injected in vivo (0.05-0.1 ml per specimen, depending on shell length, which ranged from 10 to 17 cm). Cobalt chloride blocks two major steps of cel- lular respiration. As the result of tissue hypoxia it stimulates cell proliferation (Webb, 1962, cited by Cucchi 8 Baruffaldi, 1989). After 60 h, 0.1% colchicine solution was in- jected to the mussel's foot in vivo for 6 h. From 0.5 to 1.0 ml of colchicine solution were used (depending on the mussel’s size). Gills were dissected, homogenized in distilled water, and hypotonized for 60 min in distilled water. Cell suspensions were fixed by 3:1 methanol/ace- tic acid and centrifuged three times at 1,000 rpm. Each slide preparation was made using air-drying technique (Thiriot-Quiévreux & Ayraud, 1982). For conventional karyotypes, chromosome preparations were stained with 5% Giemsa in distilled water for 20 min. CMA, staining was done according to Sola et al. (1992) and Ag- NOR staining as described by Howell & Black (1980). 206 WOZNICKI TABLE 1. Relative lengths (RL) and centromeric indices (Cl) of Anodonta woodiana chromosomes. Chromosome pair no. RL SD CI SD Classification 1 3 + 0.06 46.50 + 4.39 т 2 3:39 + 0.08 49.09 + 0.50 m 3 2.98 + 0.06 43.34 + 2.83 m 4 2.60 + 0.03 43.93 + 3.20 m 5 2.53 + 0.07 43.06 + 2.21 m 6 2.41 + 0.04 45.65 + 0.99 m 7 2.42 + 0.01 45.19 + 5.49 m 8 2.31 + 0.13 45.38 + 1.19 m 9 2.21 + 0:07 43.39 + 2.46 m 10 2415 + 0.09 43.76 + 1.96 m 11 3.20 + 0.02 35.78 + 2.34 sm 12 2.96 + 0.14 3126 +.1.45 m-sm 13 2:82 2003 33.47 + 107 sm 14 2.60 + 0.08 36.02 + 2.51 sm 15 2.51 + 0.09 29.65 + 4.91 sm 16 2.50 OM 31.31 IO m-sm 17 2.22 2:00:11 38.11 + 0.68 m-sm 18 2.16 + 0.01 38.27 +227 m-sm 19 2.28 + 0.04 31.96 2:01 sm 1 1 2 EL sa $8 ЦА RA 8 9 10 aT 5 85 a se И 13 т u — 17 FIG. 1. Karyotype of Chinese mussel (Anodonta woodiana). m — metacentric chromosomes, m-sm — meta-submetacentric and sm — submetacentric chro- mosomes. NOR-bearing chromosome pair is framed. Scale bar equals 5 um. ANODONTA WOODIANA CHROMOSOMES 207 Chromosome spreads were analyzed under a Nikon Optiphot 2 fluorescent microscope equipped with UV filters for identification of fluorescent signals and photographed by Coolpix 995 camera. Ten metaphase plates were karyotyped. Mor- phometric measurements of chromosomes were made using the freeware computer appli- cation MicroMeasure version 3.3 available on the Internet at: http://www.colostate.edu/Depts/ Biology/MicroMeasure. The relative length (RL) (100x chromosome length/total haploid length) and the centromeric index (Cl) (100х length of the short arm/total chromosome length) were calculated. Chromosomes were classified according to Levan et al. (1964). In case of six animals sequential staining CMA3/ Ag-NOR was done, and at least three metaphase plates from each specimen were analysed. About 50 interphase nuclei were observed from the same six individuals after silver staining. RESULTS From 19 individuals of Anodonta woodiana, 211 Giemsa-stained metaphase plates were analysed, showing that the diploid chromo- some number was 2n = 38 (Fig. 1). Relative length ranged from 3.74 to 2.15 (Table 1), and the karyotype consisted of 10 pairs of meta- centric, five pairs of meta-submetacentric, and four pairs of submetacentric chromosomes (FN = 76) (Fig. 1, Table 1). Staining with fluorochrome CMA3 revealed bright positive bands at terminal position on the short arm of one chromosome pair of Anodonta woodiana (Fig. 2). The same results were obtained using silver staining (Ag-NOR). Sequential CMA3/Ag-NOR staining procedure of the same metaphases showed that the CMA3 and silver positive signals appeared at the same chromosome site of metacentric chromosome pair no. 6 (Fig. 3). The number of silver-stained interphase nucleoli in A. woodiana cells never exceeded two nucleoli per cell (Fig. 4). DISCUSSION The karyotype of the Chinese mussel has been described for the first time in the present paper. The chromosome number of Anodonta woodiana, 2n = 38 (Fig. 1), is coincident with FIGS. 2-4. Anodonta woodiana. FIG. 2: Metaphase chromosomes of after CMA,-stain- ing. Arrows indicate NOR chromosomes. Scale bar equals 5 um. FIG. 3: Metaphase chromo- somes of after Ag-staining. Arrows indicate NOR chromosomes. FIG. 4: Silver stained interphase nuclei with two active nucleoli. 208 WOZNICKI that reported for other Anodonta spp. - А. anatina, А. grandis, А. piscinalis, А. судпеа, А. subcircularis (Nakamura,1985; Barsiene, 1994). Such a diploid chromosome number is the most frequent one among the bivalve spe- cies previously studied. About 47% of species within the class Bivalvia, particularly from Palaeoheterodonta and Heterodonta possess 38 chromosomes (Thiriot-Quiévreux, 1994). Fundamental chromosome arm number (FN) reported for four Unionidae species equaled 76 (Nakamura, 1985). The same value of FN was observed in Anodonta woodiana, because only bi-armed chromosomes (meta- and sub- metacentrics) were found (Fig. 1; Table 1). In eukaryotes, the 18S, 5.8S, and 28$ ribo- somal RNA genes (called major rDNA) are present in high copy number and are clustered as tandem repeats at one or more chromo- somal sites, termed nucleolar organizer re- gions (NORs) (Long & David, 1980). These clusters can be visualized indirectly by stain- ing complex of residual acidic protein associ- ated with the fibril center of the nucleolus (Ag-NORs) (Jordan, 1987) or using chromomycin Аз (СМАз) staining, which binds to GC rich chromatin (Amemiya & Gold, 1986). These methods do not detect the regions con- taining 5S (minor) rDNA, another multicopy ri- bosomal gene not involved in the formation of the nucleolus (Little & Braaten, 1989). The single NOR locus in Chinese mussel (Fig. 3) represents one of the NOR patterns observed in bivalves. The number of NOR-bearing chro- mosome pairs in these mollusks varies from one in Mya arenaria (Thiriot-Quiévreux et al., 1998), Donax trunculus (Martinez et al., 2002) and Brachidontes pharaonis (Vitturi et al., 2000) to three in Mytilus californianus (Martinez-Lage et al., 1997; Gonzalez-Tizon et al., 2000) and М. trossulus (Martinez-Lage et al., 1997). The chromosomal location of NORs in most species was terminal, as was found in A. woodiana (Figs. 2, 3). It has been suggested that a single pair of chromosomal NORs located terminally may represent a plesiomorphic character (Amemiya & Gold, 1990; Thiriot-Quiévreux, 1994). GC-rich CMA3 positive heterochromatin con- nected to NORs is typical of fish and amphib- ians (Amemiya & Gold, 1986), although it has also been observed in bivalve mollusks (Martinez-Exposito et al., 1997; Martinez-Lage et al., 1994). Staining with fluorochrome CMA3 has revealed the existence of GC bands on one chromosome pair, at the same location as Ag-NOR in Anodonta woodiana (Figs. 2, 3). Other bivalve species show СМАз positive bands on two or more chromosome pairs. In mytilids, some CMA3 bands were present at the NOR sites but also CMA3-negative NORs were present and CMA3 bands not connected to NORs were found (Martinez-Lage et al., 1995; Vitturi et al., 2000). The interstitial loca- tions of CMA3 bands were observed on Donax trunculus (Martinez et al., 2002) and Dreissena polymorpha chromosomes (Woznicki & Boron, 2003). Apart from the single Ag-NOR site СМАз positive signals were found in zebra mussel on almost all chromosomes except pairs 1, 5 and 16 (Woznicki & Boron 2003). The association of NORs with CMA3-bright bands shows that the use of combined, se- quential CMA3/Ag-NOR staining proved to be practicable and reliable for the detection of ri- bosomal regions of the bivalve mollusks chro- mosomes. Present findings provide an initial step in the cytogenetic characterization of invasive aquatic species, Anodonta woodiana and the first case of NORs description in the species from genus Anodonta. ACKNOWLEDGEMENTS The study was supported by project No. 0804.0205 financed by the University of Warmia and Mazury in Olsztyn, Poland. LITERATURE CITED AFANASJEV, S. A., В. ZDANOWSKI & А. KRASZEWSKI, 2001, Growth and population structure of the mussel Anodonta woodiana (Lea, 1834) (Bivalvia, Unionidae) in the heated Konin Lakes system. Archives of Polish Fish- eries, 9: 123-131. AMEMIYA, C. T. & J. R. GOLD, 1986, Chromo- mycin A, stains Nucleolar Organizer Regions of fish chromosomes. Copeia 1: 226-231. AMEMIYA, С. T. 8 J. К. GOLD, 1990, Cytoge- netic studies in North American minnows (Cyp- rinidae). XVII. Chromosomal NOR phenotypes of 12 species, with comments on cyto- systematic relationships among 50 species. Hereditas, 112: 231-247. BARSIENE, J., 1994, Chromosome set changes in molluscs from highly polluted habitats. Pp. 434-446, in: А. В. BEAUMONT, ed., Genetics and evolution of aquatic organisms. London, Chapman & Hall. BERAN, L., 1997, First record of Sinanodonta woodiana (Mollusca: Bivalvia) in the Czech ANODONTA WOODIANA CHROMOSOMES 209 Republic. Acta Societatis Zoologicae Bohemo- slovenicae, 61: 1-2. CUCCHI, С. & А. BARUFFALDI, 1989, А simple in vivo method for increasing mitoses in teleost fish. Cytobios, 60: 165-169. GONZALEZ-TIZON, A., А. MARTINEZ-LAGE, I. REGO, J. AUSIO 8 J. MENDEZ, 2000, DNA contents, karyotypes and chromosomal loca- tion of 18S-5.8S-28S ribosomal loci in some species of bivalve molluscs from the Pacific Canadian coast. Genome, 43: 1065-1072. HOWELL, W. M. & D. A. BLACK, 1980, Con- trolled silver staining of nucleolar organizer regions with a protective colloidal developer: a 1-step method. Experientia 36: 1014-1015. JORDAN, G., 1987, At the heart of the nucleo- lus. Nature 329: 489-499. KISS, A. & J. PEKLI, 1988, On the growth rate of Anodonta woodiana (Lea 1834) (Bivalvia: Unionacea). Bulletin of Agricultural Society Godollo, 1: 119-124. KRASZEWSKI, A. & B. ZDANOWSKI, 2001, The distribution and abundance of the Chinese mussel Anodonta woodiana (Lea 1834) in the heated Konin Lakes. Archives of Polish Fish- eries, 9: 253-265. LEVAN, A., K. FREDGA & A. A. SANDBERG, 1964, Nomenclature for centromeric position on chromosomes. Hereditas, 52: 201-220. LITTLE, К. D. & D. С. BRAATEN, 1989, Genomic organization of human 5$ rDNA and sequence of one tandem repeat. Genomics, 4: 376-383. LONG, Е. O. & I. D. DAVID, 1980, Repeated genes in eucaryotes. Annual Reviews Bio- chemistry, 49: 727-764. MARTINEZ, A., L. MARINAS, A. GONZALEZ- TIZON & J. MENDEZ, 2002, Cytogenetic char- acterization of Donax trunculus (Bivalvia: Donacidae) by means of karyotyping, fluoro- chrome banding and fluorescent in situ hybrid- ization. Journal of Molluscan Studies, 68: 393-396. MARTINEZ-EXPOSITO, М. J., J. MENDEZ & J. PASANTES, 1997, Analysis of NORs and NOR-associated heterochromatin in the mus- sel Mytilus galloprovincialis Lmk. Chromosome Research, 5: 268-273. MARTINEZ-LAGE, A., A. GONZALEZ-TIZON, J. AUSIO & J. MENDEZ, 1997, Karyotypes and Ag-NORs of the mussels Mytilus californianus and M. trossulus from the Pacific Canadian coast. Aquaculture, 153: 239-249. MARTINEZ-LAGE, A., A. GONZALEZ-TIZON & J. MENDEZ, 1994, Characterization of differ- ent chromatin types in Mytilus galloprovincialis L. after C-banding, fluorochrome and restric- tion endonuclease treatments. Heredity, 72: 242-249. MARTINEZ-LAGE, A., A. GONZALEZ-TIZON & J. MENDEZ, 1995, Chromosomal markers in three species of the genus Mytilus (Mollusca: Bivalvia). Heredity, 74: 369-375. NAKAMURA, Н. K.,1985, A review of molluscan cytogenetic information based on the CISMOCH-computerized index system for molluscan chromosomes. Bivalvia, Polyplaco- phora and Cephalopoda. Venus, Japanese Journal of Malacology, 44: 193-225. PROTASOV, А. A., $. A. AFANASJEV, O. O. SINICYNA & В. ZDANOWSKI, 1994, Сотро- sition and functioning of benthic communities. Archives of Polish Fisheries, 2: 257-284. SOLA, L., А. К. ROSSI, V. IASELLI, Е. M. RASCH & P. J. MONACO, 1992, Cytogenetics of bi- sexual/unisexual species of Poecilia. Il. Analy- sis of heterochromatin and nucleolar organizer regions in Poecilia mexicana mexicana by C- banding and DAPI, quinacrine, chromomycin A, and silver staining. Cytogenetics and Cell Genetics, 60: 229-235. THIRIOT-QUIEVREUX, C., 1994, Chromosomal genetics. Pp. 369-388, in: A.R. BEAUMONT, ed., Genetics and evolution of aquatic organisms. London, Chapman & Hall. THIRIOT-QUIEVREUX, C., 2002. Review of the literature on bivalve cytogenetics in the last ten years. Cahiers de Biologie Marine, 43: 17-26. THIRIOT-QUIEVREUX, C. & N. AYRAUD, 1982, Les caryotypes de quelques especes de bivalves et de gasteropodes marins. Marine Biology, 70: 165-172. THIRIOT-QUIEVREUX, C., К. BLICHARSKA 4 M. WOLOWICZ, 1998, Karyotype of Mya arenaria L. (Bivalvia) from the Gulf of Gdansk (Baltic Sea). Polish Archives of Hydrobiology, 45: 523-530. VITTURI, R., P. GIANGUZZA, M.S. COLOMBA 8 S. RIGGIO, 2000, Cytogenetic characteriza- tion of Brachidontes pharaonis (Fisher P., 1870): karyotype, banding and fluorescent in situ hybridization (FISH) (Mollusca: Bivalvia: Mytillidae). Ophelia, 52: 213-220. WATTERS, G. T., 1997, A synthesis and review of the expanding range of the Asian freshwa- ter mussel Anodonta woodiana (Bivalvia: Unionidae). Veliger, 40: 152-156. WOZNICKI, P. 8 A. BORON, 2003, Banding chro- mosome patterns of zebra mussel Dreissena polymorpha (Pallas) from the heated Konin lakes system (Poland). Caryologia, 56: 427-430. Revised ms. accepted 23 January 2004 MALACOLOGIA, 2004, 46(1): 211-216 LOCOMOTION IN HELIX ASPERSA Norbert Buyssens Morphology Unit, Laboratory of Pharmacology, University of Antwerp — UIA, Wilrijk, Belgium; liliane.vandeneynde@ua.ac.be ABSTRACT The pedal waves in Helix aspersa move faster than the foot of the animal. On the other hand, a histological study of the foot could not identify an organized muscular structure expected to be capable of wave construction. Instead, a non-organized tissue with rela- tively few muscle cells and many collagen fibers mixed with vessels and empty cavities was found. We could demonstrate that the movement of the waves was uncoupled from the move- ment of the foot and that the forward displacement of the snail is due to rhythmic fluid accumulation under pressure. This pressure generates a force in backward direction on the substratum, which in turn is used as push off by the animal to move in a forward direction. The snail does not crawl, the waves move independently from the foot sole, and the animal glides smoothly without changing the length or the shape of the foot. The problem how waves are constructed starting from the available muscular material is not solved. We advance a cautious hypothesis that it happens by a cyclic reversible re- cruitment of cells at the moving front and a corresponding dropping off at the rear. Key words: Helix, locomotion, haemolymph. INTRODUCTION During a study of the changes in the shape of smooth muscle cells when undergoing con- traction, our attention was drawn to the foot of the snail as a possible suitable model. Com- mon garden snails were caught, fixed and pro- cessed for histological examination. This showed that the foot did not contain one or more large muscles orientated in one direc- tion, but many discrete muscle cells and fi- bers distributed in an irregular three- dimensional pattern. Moreover, collagen fibers and empty clefts or holes made up the larger part of the space. When watching the charac- teristic waves on the foot sole, the question arose how an apparently unorganized group of individual muscle cells and fibers could manage to assemble ordered unidirectional waves. Closer examination showed that the waves moved faster than the foot, which led to the conclusion that the movement of the waves was uncoupled from the advancing movement of the foot. Atransmission by a fluid interphase, in this case the haemolymph, was considered as a likely possibility and became the aim of this study. Because we do no not know exactly how these waves are structured, 211 the use of the term is purely descriptive. What we see are narrow, dark, transverse stripes separated by wider light segments, which we will call “junctions” for convenience. MATERIALS AND METHODS Snails (n = 150) of the species Helix aspersa were collected in spring and early summer. They were housed in plastic boxes and fed lettuce. Their average weight was 5.2 g (3.9-7.2 д). After dissection, tissues of 20 animals were fixed in a 4% aqueous solution of formalin. Bouin’s, methacarn, and isopropanol fixatives were used when appropriate. Sections of paraffin- embedded material were stained with Sirius red haematoxylin. Additional stains were Masson's trichrome, PAS, PAS with amylase digestion, haematoxylin and eosin, alcian blue at pH 4.2, mucicarmin, Von Kossa’s for calcium, Kernechtrot and Fontana’s silverstaining for melanin. A contracted foot of 21 mm length and an expanded one of 32 mm length were cut in uninterrupted serial sections. Speeds of the animals were recorded while glid- ing on perplex plates over a distance of 50 mm, and for each animal five consecutive displace- 212 BUYSSENS ments were timed. Videotapes were made from animals in different positions, on different substrata and under different angles of illumi- nation. Standard and scanning radiographs of resting and moving animals were taken. Injec- tions of Indian ink droplets in the foot of anaes- thetized animals were studied by videotape and after killing by histology in order to estab- lish the exact location in the foot. Repeated attempts to record electrical ac- tivity were unsuccessful. RESULTS Histology The foot contains from the tip to the tail a moderate number of slender discrete smooth muscle cells or fibers embedded in a loose network of many collagen fibers. Fibers lie in longitudinal, circular, and vertical directions without any preferential pattern. They make many short contacts with each other (Fig. 1). Many vacuolated interstitial cells and many empty spaces occupy the interstitium. Thin- walled muscular vessels (arteries) and spaces lined by flattened endothelial type cells, prob- ably representing veins, can be identified. Many empty spaces not lined by cells are dis- tributed throughout and are particularly con- centrated at the margins of the foot sole. The existence of a ramified communicating system of channels can be demonstrated convincingly by the injection of Evans blue in the tail of a fixed foot. The dye spreads diffusely in the core of the foot and then moves to the margins where it is easily recognized because of their thinness. If sufficient pressure is applied, the dye fills the head and induces the eversion of the antennae. Few nerve fibers can be detected by routine stains. We used a Mab against acetylated tubu- lin (Sigma T 6793, clone 6-11B-1) and DAB as chromogen to demonstrate a nervous network making contact with the smooth muscles. We only found thin ramifications extending to the base of the epithelial cells of the sole that had all the characteristics of sensitive nerve endings. In addition, the sole shows transverse lines regu- larly spaced at 1.0-1.5 mm intervals in a con- tracted foot of 25 mm length. Histology reveals that they are condensations of collagen fibers closely apposed to the sole epithelium. We did not find mention of these structures in the litera- ture. Findings on mucus cells are not reported because they are not relevant to this study. Displacement of the Waves The snail Helix aspersa moves according to a monotaxic anterograde “wave” pattern. The waves move faster than the animal. The junc- tion length is + 4 mm, the thickness of the wave is 1.0-1.5 mm. The wave is composed of two layers, a cranial one which is lighter and a darker caudal one. Generally 10 to 12 waves can be counted at one point in time. Table 1 shows figures that allow calculation of the ra- tio of the speed of the waves to the speed of the animal. The waves move 2.3 times faster than the animal. Prior 8 Gelperin (1974) found in Limax maximus a ratio of 2.2. Jones & Trueman (1970) detected in Patella vulgata a ratio of 3.5. Bonse (1935) reports ratios be- tween 0.93 and 1.48 in Helix pomatia. Unfor- tunately, these authors did not elaborate on this phenomenon in later studies. TABLE 1. Speed of animals versus speed of waves. Speed of animals (n = 90 individuals) Distance covered in 30 sec Speed per second Speed of waves (n = 60 individuals) Length of junction Number of waves in 30 sec Distance covered in 30 sec Speed per second 5 = 50 mm 1.6 mm S=4mm ЗЕ 4 mm x 27.5 = 110 тт 3.7 mm Ratio speed of waves/speed of animals 28 HELIX ASPERSA LOCOMOTION 215 FIG. 1. Section of foot. A. Irregular distribution of muscle fibers in red and collagen fibers in green. Masson's trichrome stain. Scale bar = 200 ит. В. Collagen fibers are red and are closely apposed to the muscle fibers which show many contacts. Sirius Red Hematoxylin. Scale bar = 50 um. Because the progression of waves is not matched by a corresponding progression о the foot, the waves cannot act as “toes” on which the snail could lean to move ahead. Hence the problem of how the waves command foot move- ment must be addressed. Independent from the waves, other irregular undulations are seen а the very margins of the foot over a width less than 1 mm. They move in a caudal direction at a pace of 1 mm every 1-3 FIG. 2. Scanning radiograph showing the tight apposition of the foot on the substratum. A. Frontal view. B. Lateral view. 214 BUYSSENS TABLE 2. Time to pull different weights over a distance of 50 mm in three snails (A-C) of com- parable body weight (body weight is given in parentheses). Time in sec Weight A (5.9 g) В (5.49) С (5.3 9) 99 29 35 27 159 40 41 42 214 90 65 70 sec and reach the tail. They were mentioned by Bonse (1935), and their function is unknown. Before a wave appears, the tip of the foot is dilated by fluid accumulation. In this dilated portion, the first wave is formed. Successive waves develop in caudal direction, but at the very moment they are formed they start to move in cranial direction. It is clear that this is not comparable to a peristaltic wave system originating in the tail and rushing to the tip. When the animals stops, the waves also stop, the most caudally situated first. There is no jamming at the tail. Accidental amputation of the tail does not stop emergence of waves. When the moving animal is grasped by the observer and turned upside down waves con- tinue for a few minutes. Visual observation confirmed by videotaping show that the waves do not cause a retraction of the foot surface but that the junctions bulge as they are filled with fluid. It is clear that the junctions by their expansion push on the substratum and that they are the propulsive elements. Bonse (1935) and Lissmann (1945) have demon- strated that the pressure of the foot on the substratum decreased when the wave passed and resumed to normal when the junction moved over the recorder plate. Waves can persist in feet severed from the body. They can last for 15 min and thereafter continue as irregular slow movements for two hours. These contractions come to an end in the tip of the foot where they first originated: primum movens, ultimum moriens! The mar- ginal undulations were very resistant and could be observed up to six hours after sec- tioning. In the slug Limax maximus. Prior & Gelperin (1974) detected waves after decapi- tation and demonstrated that the presence of the central nervous system was necessary to initiate them. However, once started, the waves could form independently. The impor- tant conclusion of these findings is that waves can develop and function without a pulsating heart. In our material, the snail does not lift its foot in a detectable amplitude during forward move- ment. We could substantiate this in three ways. First, by visual observation of the moving ani- mals with an amputated tail. Second, by al- lowing the snail to glide on black paper and studying the mucus trail. Over distances from 50 cm to 1 m this is a straight ribbon without any irregularities either in the centre or at the margins. Third, by scanning radiography of resting and moving animals, which demon- strates that the foot really sticks over its whole length to the substratum (Fig. 2). The snail glides on its own slime. The bulg- ing junction exerts a backward pressure par- allel to the substratum. This can convincingly be demonstrated by videotaping: when the snail rests on a thin plastic strip and is held by an observer, the marker lines on the strip move backwards. With the same method, it is pos- sible to measure the energy that the snail uses for forward displacement. When we attach to the strip a string with a weight that we let hang over the border of a table, we can measure the time in which the animal can push back- wards the plastic strip with its attached weight. Table 2 shows the figures for three snails of comparable weight. It appears that animal A, for instance, can pull a weight of 9 g over a distance of 50 mm in 29 sec, which is the same speed as a free moving animal. When the weight increases, the speed decreases ac- cordingly. Calculating the energy in our first example, we arrive at 0.0015 milliwatt. The displacement of haemolymph in the foot as demonstrated by the bulging of the junc- tions is not visible when the foot rests on the substratum, because it is a pressure mecha- nism and not a crawling mechanism. The pres- sure generates a force that is parallel to the substratum and directed backwards. These are the conditions for developing shear stress. However, due to the fact that the mass of the substratum is too big to be displaced, the dis- placement or shear strain, occurs in the op- posite direction. This may be an explanation why a backward moving force induces a for- ward displacement. Finally, the behavior of injected Indian ink should be reported. Histological examination showed that the droplets were generally present in pre-existing clefts in the core of the foot. Sometimes they accumulated in the nu- merous spaces close to the epithelium. A lin- ear deposit of a few mm in longitudinal HELIX ASPERSA LOCOMOTION 215 direction is particularly suited for examination. When the arriving wave hits the posterior end of the ink deposit, it is slightly pushed forward and during the passage of the wave slightly stretched. Once the wave has passed, the de- posit resumes its original shape and position. The image looks like the wobbling of a leaf on the waves in a pond. During these small changes in the shape and position of the ink deposits special attention was paid to the shape of the foot and to the continuity of the forward movement. Analysis of the videotapes could not disclose any change, leading to the conclusion that the behavior of the ink mate- rial is an internal event and is not coupled to the foot sole. DISCUSSION Our study demonstrates that in Helix aspersa the pedal waves move faster than the foot. The shape of the foot and the continuity of the for- ward movement do not show any temporal or topographic relationship with the wave move- ment. The displacement and deformation of Indian ink droplets is not coupled to similar changes in the foot sole. These findings allow the conclusion that the waves act indirectly by the intermediary of the haemolymph. This con- clusion is also corroborated by the observa- tion of the filling with fluid of the segments (“junctions”) between the waves, resulting in the pressure on the substratum, leading in turn to the forward movement. In addition, the tem- poral persistence of waves after severing the foot from the body indicates that a pulsating heart is not necessary to maintain waves. The question of how waves are built up from non-organized muscles is not solved. Authors who describe muscle bundles in different fixed directions are fortunate and use these to ex- plain the locomotion of the foot in snails and slugs. Jones (1975) published an exhaustive report on locomotion in Pulmonata. However, he did not address the problems of our present study. To the best of our knowledge, he is the first author who succeeded in catching mov- ing waves. He described in Agrolimax reticulatus the fixation of waves by immersing the moving animals in liquid nitrogen (Jones, 1973). In cryostat sections, he reported a com- pression of oblique muscles and an almost com- plete occlusion of the haemocoel. He presented a micrograph showing muscle fibers that are reduced to strings, lying in many directions and occupying a very small fraction of the tissue area. For a morphologist, it is difficult to corre- late this tissular arrangement with waves. In Helix pomatia, he described the foot as highly muscular. We did not study Helix pomatia, but we can confirm that in Helix aspersa and in several other snails this is not the case. In his review on locomotion of molluscs, Trueman (1983) described for Helix a model of locomotion based on a crawling mechanism. Crawling is a biphasic activity. A transient sta- tionary point or zone serves as an anchor for the contracting or elongating free moving and uplifting segment of the animal. When waves are present, it is assumed that the wave 1$ the anchor and that the animal advances because of the successive anchoring of the waves. The author refers to the work of Denny (1980), who studied the physicochemical properties of the mucus of Agrolimax columbianus. This author proposed a dual reaction of mucus changing from a solid phase in the stationary state to a liquid phase in the moving state, acting like a material ratchet. These properties of the mu- cus may facilitate the crawling mechanism proposed by Trueman. Several other students of snail locomotion also propose the crawling mechanism for for- ward displacement: Trappman (1916), Miller (1974), Gainey (1976), and Moffett (1979). In a study on the histology of the foot and the locomotion of Gastropoda, Elves (1961) mentioned that in Discus rotundatus the mus- culature of the foot is not well developed, and muscle fibers are of small size and few in num- ber. He devoted a short description to Helix aspersa and described in the foot a reticulum of connective tissue fibers and large muscles which run both longitudinally and dorsoven- trally. However, in the accompanying diagram, he depicted the muscular component as a few scattered small bundles occupying a small fraction of the total transverse section area. In his discussion, he mentioned that the locomo- tion may be influenced by an interplay between haemocoel turgor and muscular waves. An interesting type of forward displacement in terrestrial gastropods was discussed by Pearce (1989). It was termed “loping” (derived from galloping) and differs from the gliding pro- gression. п the loping motion, the gastropod lifts its head from the substratum and thrusts it forward, then replaces it on the substratum, forming a low arch in the sole behind the head through which the rest of the body flows to the new stationary point of contact. The mucus trail left consists of more or less elongated dots, in contrast to the continuous mucus trail during gliding progression. Hence, loping is a perfect example of crawling and a strong argument 216 BUYSSENS for the existence of a different “ordinary” (ter- minology of Pearce) gliding mechanism. Inter- estingly, the waves of ordinary gliding are present with loping, but there is no interference. We can only speculate about what happens in Helix aspersa. The signal for wave forma- tion starts at the tip of the foot, resulting in the successive appearance of waves in the cau- dal direction. At the very moment the waves are induced, they start moving in cranial di- rection back to their initial inductive signal. The waves are not at all sinusoidal peristaltic con- tractions or pressure waves like in blood ves- sels. They resemble slice-like condensations of tissue (muscle, collagen, and interstitial cells), which move in an upright position along a horizontal plane parallel to the foot sole. A possible explanation how to visualize the for- mation of such a structure could be that the slices of condensed tissue while moving ac- quire cells at the advancing front and release them again at the trailing front. This phenom- enon of recruitment is known in the forward movement of cells in culture. Bretcher & Aguado-Velasco (1988) describe how lamellipodia, which are formed by cells when they start moving, recruit plasma membrane material at the expense of the trailing end of the cell. In fact, these cells do not advance by moving but by growing. The concept of recruit- ment can perhaps explain why waves are bilayered, the frontal directed half being the recruitment front and the rear half being the propulsion machine. Concerning the muscular structure of the foot an interesting micrograph is published by Ber- пага (1968). It shows in the foot of the large marine snail Polinices lewisi muscle and col- lagen fibers in a three-dimensional pattern reminiscent of the foot of Helix aspersa. Waves were not mentioned, and the foot aspirates and expels ambient water. Analyses of movie films of waves in Helix pomatia have been reported by Bonse (1935) and Lissman (1945) and in Patella vulgata by Jones & Trueman (1970). However, these ex- cellent studies do not address the basic ques- tion how we can understand the formation of waves or why they move faster than the ani- mals. ACKNOWLEDGMENTS The author wishes to thank Rita Van den Bossche for the histology work and Liliane Van den Eynde for logistical and secretarial work. Prof. A. Herman offered graciously his labora- tory facilities. Prof. A. De Schepper and his staff provided generously top quality radiographs. LITERATURE CITED BERNARD, F. R., 1968, The aquiferous system of Polinices lewisi. Journal of the Fisheries Research Board of Canada, 25(3): 541-546. BONSE, H., 1935, Ein Beitrag zum Problem der Schneckenbewegung. Zoologische Jahr- búcher, Abteilung Allgemeine Zoologie und Physiologie der Tiere, 54: 349-384. BRETCHER, М. S. & С. AGUADO-VELASCO, 1998, Membrane traffic during cell locomotion. Current Opinion in Cell Biology, 10: 537-541. DENNY, M. L., 1980, The role of gastropod pedal mucus in locomotion. Nature, 285: 160-161. ELVES, М. W., 1961, The histology of the foot of Discus rotundatus and the locomotion of gas- tropod Mollusca. Proceedings of the Malaco- logical Society London, 34: 346-355. GAINEY, L. F., Jr., 1976, Locomotion in the Gas- tropoda: functional morphology of the foot in Neretina reclivata and Thais rustica. Malacologia, 15(2): 411-431. JONES, H. D., 1973, The mechanism of loco- motion of Agrolimax reticulatus (Mollusca: Gastropoda). Journal of Zoology, London, 171: 489-498. JONES, H. D., 1975, Locomotion. Pp. 1-32, in: V. FRETTER 8 J. PEAKE, eds., Pulmonates, |, Functional anatomy and physiology. London: Academic Press. JONES, H. D. 8 Е. КВ. TRUEMAN, 1970, Loco- motion of the limpet, Patella vulgata L. Journal of Experimental Biology, 52: 201-216. LISSMANN, H. W., 1945, The mechanism of lo- comotion in gastropod molluscs. 1. Kinemat- ics. Journal of Experimental Biology, 21: 58-69. MILLER, 5. L., 1974, Adaptive design of loco- motion and foot form in prosobranch gastro- pods. Journal of Experimental Marine Biology and Ecology, 14: 99-156. MOFFETT, $., 1979, Locomotion in the primi- tive pulmonate snail Melampus bidentatus: foot structure and function. Biological Bulletin, 57: 306-319. PEARCE, T. A., 1989, Loping locomotion in ter- restrial gastropods. Walkerana, 3(10): 229- 237 PRIOR, D. J. & А. GELPERIN, 1974, Behavioral and physiological studies on locomotion in the giant garden slug Limax maximus. Malacologi- cal Review, 7: 50-51. TRAPPMAN, W., 1916, Die Muskulatur von He- Их pomatia L. Zeitschrift für Wissenschaftliche Zoologie, 115: 490-585. TRUEMAN, E. R., 1983, Locomotion in molluscs. Pp. 155-198, in: A. S. M. SALEUDDIN & K. M. WILBUR, eds., The Mollusca, Volume 4, Physi- ology. London: Academic Press. Revised ms. accepted 4 May 2004 MALACOLOGIA, 2004, 46(1): 217-224 COLORATION IN HELICINIDAE (MOLLUSCA: GASTROPODA: NERITOPSINA) Ira Richling Zoologisches Institut, Christian-Albrechts-Universitát zu Kiel Olshausenstraße 40, 24098 Kiel, Germany; ira@richling.de ABSTRACT The coloration of Costa Rican Helicinidae has been studied, with special attention paid to the arboreal species. It is shown that either shell color or mantle pigmentation contribute to the coloration visible in the living animals. Ecological and systematic implications are given. This paper is supplementary to Richling (2004). Keywords: Helicinidae, Costa Rica, Central America, classification, coloration. INTRODUCTION During a recently published revision of the systematics and species differentiation in Costa Rican Helicinidae (Richling, 2004), the role of coloration was studied. As the plates were printed in black and white, | provide here the same plates in color and give additional notes on species in relationship to color pat- terns. Further details on the species, especially those in Figure 6, material and methods and literature are given in Richling (2004). RESULTS AND DISCUSSION Coloration in land snails is mainly determined by the need for camouflage. Thus, it depends strongly on the habitat of the respective spe- cies. The Costa Rican species inhabit tropical rain forests and can be split into two groups: arboreal species and ground dwellers crawl- ing in leaf litter. The arboreal species show a variable, bright coloration, as can be found in other arboreal land snails, for example, spe- cies of Liguus (Orthalicidae), Amphidromus (Camaenidae), and Cepaea (Helicidae), with yellow and red prevailing. They are seldomly greenish in adaptation to leaves. All Costa Rican species of the genus Helicina exhibit this pattern (Figs. 3, 4, 5A-D), with a greenish color developed in certain specimens of Helicina funcki L. Pfeiffer, 1849 (Fig. ЗА), and Helicina escondida Richling, 2004 (Fig. 4H). The ground dwellers, Lucidella lirata (L. Pfeiffer, 1847), Alcadia hojarasca (Richling, 2001), and A/cadia boeckeleri (Richling, 2001), are uniformly brownish, and in addition exhibit a rough surface, that 1$, periostracal hairs or ridges on the shell (Figs. 20-Q, 5F-H). Pyrgodomus microdinus (Morelet, 1851) is the only exception in its strong association to sur- faces of calcareous rocks. In living individu- als, the bright yellow empty shell (Fig. 2R) becomes greenish-grayish because of the underlying dark pigmentation of the mantle. Furthermore, P. microdinus glues particles of detritus on its shell, thus perfectly resembling the rock surface (Fig. 5E). The same applies to the Jamaican Eutrochatella pulchella (Gray, 1825), in which the active camouflage is func- tionally replaced by the white-yellowish mot- tling of the shell (Figs. 6M, P). When comparing the coloration of living ani- mals with empty shells, it becomes obvious that in the Costa Rican arboreal species, two different ways are utilized to produce the vari- able and bright coloration. On one hand, the species have a variable shell color combined with rather thick shells and a uniform mantle pigmentation, for example, Helicina funcki, which can even be nearly reddish; H. pitalensis Wagner, 1910; H. beatrix Angas, 1879; H. talamancensis (Richling, 2001); and H. punctisulcata cuericiensis Richling, 2004 (Figs. 1A—E, L-M, 2А-Е). On the other hand, the shells are thin and more or less transparent with exception of the outer lip (Figs. 1F-K, 2F- N), but the mantle is variously mottled and causes the visible coloration, for example, Helicina tenuis L. Pfeiffer, 1849; H. gemma Preston, 1903; H. monteverdensis Richling, 2004; H. escondida Richling, 2004; and H. chiquitica (Richling, 2001) (Figs. 3D-E, 4C- H, 5A—D). As an artifact in empty shells, the transparency becomes less, for example, com- 218 RICHLING pare Figure 21 and Figure 4F. The latter way seems to have evolved in connection with the very limited availability of calcium carbonate in Costa Rica. It seems that the optimal coloration of small- sized arboreal Helicinidae, about 3-4 mm, is dark. Evidence is given by the very small spe- cies H. chiquitica, in which most individuals are dark (Fig. 5C), populations of Helicina monteverdensis of a reduced average body size (Fig. 4G), and juveniles of Helicina funcki (Fig. 3B). Due to the high adaptability of the colora- tion, its value for systematics is limited, al- though the present study shows that in arboreal species the way to achieve the final coloration is typical for each species. For thin- shelled species, the varying and patterned mantle pigmentation is characteristic. When looking at a number of individuals, this pat- tern shows at certain specificity for different species, but single specimens might show exceptions. In some cases, it is even typical at population level, for example, in Helicina tenuis from Cabo Blanco on the Pacific plain (Fig. 3D) and La Selva on the Caribbean plain (Fig. 3E), or in Helicina monteverdensis in populations about 5 km from each other (Figs. 4F, G). The coloration of head and foot seldom shows species specificity: the upper side is usually dark, especially towards the head and tentacles, whereas the lower side is light. Among the Costa Rican species, Helicina talamancensis represents the only exception. In all specimens studied, the whole body 1$ whitish except for the sharply separated black tentacles (Fig. 4B). LITERATURE CITED RICHLING, I., 2004, Classification of the Heli- cinidae: review of morphological characteris- tics based on a revision of the Costa Rican species and application to the arrangement of the Central American mainland taxa (Mollusca: Gastropoda: Neritopsina). Malacologia, 45(2): 195-440. COLORATION IN HELICINIDAE FIG. 1. Shell coloration of Costa Rican species. A-C. Helicina funcki. À. Rio Barbilla. B. Manzanillo. С. Santa Elena. 0-Е. H. pitalensis. D. Bajo Bonito. E. Península de Osa. Е-1. H. tenuis. F-H. Cabo Blanco. |. La Selva. J-K. H. echandiensis Richling, 2004, campamento Echandi. L-M. H. punctisulcata cuericiensis, Estación Cuerici; scale bars = 4 mm (A-E), 3 mm (F-M). RICHLING FIG. 2. Shell coloration of Costa Rican species. A. Helicina beatrix beatrix, Guayacán. В-С. Н. b. confusa (Wagner, 1908). В. Uatsi. С. Shiroles. D. H. b. riopejensis Richling, 2004, Río Peje. E. H. talamancensis, Bajo Bonito. F-H. H. gemma. Е. Cacao. С. Las Pavas. H. Siquirres. 1-4. H. monteverdensis, Monteverde. K-M. H. escondida, Río Barbilla. N. H. chiquitica, Río Barbilla. O. Alcadia hojarasca, Mirador Gerardo. P. A. boeckeleri, Pitilla. Q. Lucidella lirata, Cahuita. R. Pyrgodomus microdinus, Fila de Cal; scale bars = 3 mm (А-М), 2 тт (М-Р), 1.2 mm (Q-R). COLORATION IN HELICINIDAE FIG. 3. Living animals of Costa Rican species. A. Helicina funcki, Cahuita. В. H. funcki, juvenile, Uatsi. C. H. pitalensis, Bajo Bonito. D. H. tenuis, Cabo Blanco. E. H. tenuis, La Selva. F. H. beatrix confusa, Uatsi. с. H. beatrix confusa, Shiroles (photograph: Vollrath Wiese). Н. H. beatrix riopejensis, Río Peje. RICHLING FIG. 4. Living animals of Costa Rican species. А. Helicina beatrix beatrix, Счауасап. В. H. talamancensis, Bajo Bonito. С. H. gemma, Cacao. D. H. gemma, Las Pavas. Е. H. gemma, Siquirres. F. H. monteverdensis, Monteverde. G. H. monteverdensis, Mirador Gerardo. H. H. escondida, Shiroles COLORATION IN HELICINIDAE FIG. 5. Living animals of Costa Rican species. А. Helicina escondida, Shiroles. В. H. escondida, Rio Barbilla. C. H. chiquitica, Rio Barbilla. D. H. chiquitica, Rio Pacuarito. E. Pyrgodomus microdinus, Fila de Cal (photograph: Vollrath Wiese). Е. Alcadia hojarasca, Mirador Gerardo. С. A. boeckeleri, Pitilla. H. Lucidella lirata, Cahuita. RICHLING FIG. 6. Shell coloration. A. Helicina neritella Lamarck, 1799, Jamaica. В. H. platychila (Mühlfeldt, 1816), Dominica. С. H. orbiculata (Say, 1818), Florida. D. H. turbinata Wiegmann, 1831, Mexico. Е. H. amoena L. Pfeiffer, 1849, Guatemala. F. H. dysoni L. Pfeiffer, 1849, Trinidad & Tobago. G. H. sericea Drouet, 1859, Suriname. H. Angulata brasiliensis (Gray, 1825), Brazil. |. Alcadia major (Gray, 1824), Jamaica. J. A. hollandi (C. В. Adams, 1849), Jamaica. К. A. jamaicensis (Sowerby, 1841), Jamaica. L. A. rotunda (Orbigny, 1841), Cuba. M. Eutrochatella pulchella, Jamaica. N. Lucidella aureola (Férussac, 1822), Jamaica. O. Schasicheila alata (L. Pfeiffer, 1848), Mexico. P. Eutrochatella pulchella, Jamaica; scale bar = 5 mm (А-О). LETTER FROM THE EDITOR MALACOLOGIA, 2004, 46(1): 227-231 SPECIES CHECK-LISTS: DEATH OR REVIVAL OF THE NOUVELLE ÉCOLE? George M. Davis Department of Microbiology and Tropical Medicine George Washington University Medical Center Ross Hall 731, 2300 Eye Street NW, Washington DC 20037, U.S.A.; georgedavis99@hotmail.com With more than 100,000 described species, mollusks belong to the second largest phylum after the Arthropoda. Mollusks have attracted a large number of shell collectors, amateur malacologists, field biologists, conservation- ists, as well as evolutionary biologists, taxono- mists and systematists. As a result, there are huge amounts of та- lacological publications available for most re- gions of our planet, and our knowledge of the group is increasing day by day. However, de- spite the extensive work that has been done, nomenclature and taxonomy of many groups are still in a confused state, and the systemat- ics of numerous taxa is embroiled in contro- versy. Moreover, the problems increase with the recent advent of new anatomical and mo- lecular methods, where the new types of data are often in conflict with the traditional, that is, shell-based taxonomy. This confusion affects most taxonomic lev- els from subspecies to higher taxa and makes it difficult for many non-biologists and even professional biologists to apply or understand the correct name for a taxon. The conse- quences are not-trivial, as an incorrect deter- mination or classification can be the deciding factor in many diverse scientific and non-sci- entific activities. To maintain order in the sometimes chaotic system of publications and taxa, species check-lists are often generated for individual biogeographic regional and/or systematic groups. For the uninitiated, it is difficult to imag- ше how much work 1$ involved to generate a widely acceptable check-list. Often interna- tional groups of scientists have to sort through hundreds or even thousands of primary publi- cations, look at many voucher specimens and work through quantities of field records. They have to carefully consider frequently contra- dicting information, make educated decisions about the “correct” nomenclature and tax- onomy, and ensure compliance with the Inter- national Code of Zoological Nomenclature. 221 And finally, these check-lists have to be up- dated on a regular basis in order to keep pace with the malacological research. Given the nature of species check-lists there are, however, some critical points | would like to discuss. This could possibly help to further improve the quality of those check-lists and/ or to point out some possible pitfalls the users should be made aware of. Although the fol- lowing points might apply to many species- check lists, | will focus my attention on the two most recent European lists, the “Mollusques continentaux de France: liste de référence annotée et bibliographie” by Falkner et al. (2002), and the “Check-list of the non-marine Molluscan species-group taxa of the states of Northern, Atlantic and Central Europe” (CLECOM 1) by Falkner et al. (2001). (1) Naming Species Identifying a species, a “good species”, has often been a very difficult task, due, in part, to the large variety of species concepts and in part to difficulties in the objective selection and interpretation of the characters used, often resulting in a personal view of what a species is. The famous malacologist W. Kobelt (1881) once wrote [translated from German]: “/ obey a simple, practical rule, no matter how unsci- entific it may be. I call a good species what | can diagnose without long and careful com- parisons and measurements ...”. More than half a century ago, matters changed due to the introduction of the biological species con- cept (BSC) (Mayr, 1940) and of other, pro- gressively more refined concepts (review by Hull, 1997). Many traditional taxonomists, nev- ertheless, continue working as in the past. They use an often arbitrary chosen level of morphological difference, frequently calcu- lated “at a glance” to decide what a new “good species” is. However, these good species are actually “morphospecies” (see Giusti & Man- ganelli, 1992), and it is usually tacitly taken 228 DAVIS for granted that they correspond to biological species. Obviously, this practice is extremely subjec- tive. Extremism gave rise to “the lumpers”, who require robust differences to segregate taxa, and in doing so, tend to accept very polymor- phic species. Others, “the splitters”, base their taxonomic decisions on low levels of differen- tiation. More recently, subjective decisions were brought into clear view when molecular research revealed that both splitters and lumpers are frequently wrong, because the levels of morphological difference are often independent of genetic divergence (organisms with a low level of morphological difference may have high levels of genetic divergence, and vice versa). А quick glance at many papers shows that the authors continue to use superficial shell morphological/typological procedures. These are sometimes based on simple differences in shell height or diameter, or on the relation- ship height/diameter, differences that repeat- edly have been shown to be unsuitable for species differentiation. A first conclusion is that creating taxa simply on the basis of shell char- acters (qualitative and quantitative) should al- ways be done with extreme caution (many cases of convergence in shell shape in not related groups are known, convergent evolu- tion is rampant; Davis, 1979), unless there are very compelling reasons (fossils, rare deep sea taxa, taxa of family-level groups notori- ous for being over-named and for having high intraspecific variation, etc.). The situation may not substantially change even when other characters, anatomical, eco-ethological, are added (anatomical characters are not always highly revealing; peculiar ecology, parasitol- ogy or ethology may be the source of certain shell differences). The situation is rendered more problematic by the so-called “fanatisme du nobis” (Dance, 1970), that is, the introduction of new names to give “eternity” to one's own name. The more new names one introduces, the greater 1$ the possibility one of them survives as that of a “good species” or “good subspecies”. The International Code on Zoological No- menclature (ICZN) was created to manage, in a legalistic manner, the naming of species. But it has been abused and indirectly supports proliferation of names that cannot be sup- pressed and therefore must be considered valid and listed in check-lists until a future re- vision is made. The last edition of the ICZN (1999) called for adequate and rigorous spe- cies definitions (see also Hawskworth & Bisby, 1988: 12-13), but since then, the literature shows that this call remains unheaded in many instances. Recent creation of new species based on dubious characters is the survival/revival of the school called “Nouvelle École”, founded by the French malacologist Jules-René Bourguignat in the second half of the 19" century, accord- ing to which a species should be determined on arbitrarily chosen characters, which nearly always meant shell characters. If an individual was found to differ from all others by three characters or more, it should be considered to belong to a species new to science (Dance, 1986). This school has been unanimously con- demned, but never completely abandoned. As a matter of fact, many taxonomists officially criticize the Nouvelle Ecole, but actually fol- low а very similar if not identical practice. Con- tinuation of this practice is fueled by the fact that at least some of the species described by the followers of the Nouvelle Ecole subse- quently turn out to be “good species” (see above). Moreover, recently the method of the Nouvelle Ecole is spreading again due to the strategy of the “valeur patrimoniale” of local faunas to support conservation programs (Falkner et al., 2001: 6; 2002: 19; Bouchet, 2002: 8-12). According to such strategy, “It is extremely difficult to convince engineers or politicians of the value and need for protec- tion of a special, even unique, unnamed form, but from the moment that a name can be pro- vided there exists a recognizable unit which can be referred to” (Falkner et al., 2001: 6). Though these intentions are honorable, taxa of the species group should never be de- scribed for “political” reasons! Apart from the fact that that strategy opens doors to irrespon- sible students and amateurs, it risks to reduce taxonomy to a mere artifice possibly with grim consequences: “if the philosophy of the Nouvelle Ecole had become widely popular its effect on systematic conchology, as may be imagined, could have been catastrophic” (Dance, 1986). While | reject the subjective and unethical practice just described, | am concerned about how one manages the many names given to taxa that have uncertain value. Each taxon correctly (legally) described has value unless the contrary is demonstrated. At the same time, it is a considerable disservice to science and society to inflate check-lists with numerous SPECIES CHECK-LISTS 229 dubious nominal species/subspecies! | am convinced that these taxa must be evaluated critically and that, when it is eventually decided they must be listed as potentially endangered or as a part of a special ecosystem, their un- certain taxonomic status should be made clear. (2) The Use of Subspecies Subspecies are even more difficult to define objectively than species as was well known from the late 19" century (Kobelt, 1881, continued the sentence given above as follows: “... That which | can distinguish only by precise mea- surements | call a variety’). Alas, some mala- cologists still name subspecies as it was done by Kobelt for varieties (minor variations), just to distinguish a little characterized local form, and as a tool to satisfy their “fanatisme du nobis’. Difficulties in using subspecies come, first of all, from the fact that not all species con- cepts recognize the subspecies status. One species concept that explicitly does is the BSC. However, although often claimed by malacolo- gists, the BSC, in its true meaning, is rarely applied in malacology, because it places the taxonomy of natural species within the scheme of population genetics. Within the framework of the BSC, the concept of subspecies gradu- ally evolved from a simple “unit of conve- nience” (Blackwelder, 1967; Dobzhansky et al., 1977; Mayr, 1963, 1982) to became applicable to populations that are kept isolated usually by geographical barriers and that exhibit rec- ognizable phylogenetic partitioning due to the time-dependent accumulation of genetic dif- ferences (O’Brien & Mayr, 1991). Therefore, subspecies should presently never be used without an immense amount of rigorous data that clearly demonstrate that the concept can be applied legitimately, that is, allopatric popu- lations that have diverged sufficiently geneti- cally (based on real genetic data) where they would be elevated to full species were it not for the complete sameness of the mate rec- ognition system and the full capacity to pro- duce a F1 and F2 generation if given the opportunity (Davis, 1994). It is obvious that most check-lists that use subspecies are not based on such rigorous work. To give an example from CLECOM, for Germany there are 36 (!) species and subspe- cies listed for the minute rissooidean genus Bythiospeum. | do not know of any molecular or detailed anatomical study that has looked at variability within and between populations to infer possible genetic breaks in taxa of Bythiospeum that, in turn, could be used to deduce reproductive isolation. Therefore, | would consider the splitting of Bythiospeum a simple matter of pure subjectivity. In fact, pre- liminary genetic data produced by one of my collaborators indicate that the number of Bythiospeum species in Europe is much lower than hitherto believed and that the genus ap- pears to be paraphyletic, demonstrating the diagnostic inadequacy of the morphological methods used up-to-day. Unfortunately, CLECOM gives contradicting information as to legitimacy of the subspecies listed. Whereas on page 3 of the introduction to CLECOM | it is said that “... the CLECOM check-list will contain the nomenclaturally cor- rect names of the species and subspecies that are considered to be valid’, further down on the same page it is written “... Inclusion of named subspecies in the CLECOM database results inevitably in both well-founded and even spectacular forms being mentioned along with some ‘weak’ subspecies.” Moreover, it is said that “in the exciting faunistic literature of northern, western and central Europe subspe- cies are widely, if not generally, neglected.” If so, a check-list that is based on this literature should ignore subspecies as well. One reason for including subspecies in CLECOM is related to conservation purposes: “Our only tool to make this diversity apparent to conservation authorities, researchers in applied sciences and others who require recognising its existence is the application of trinominal nomenclature.” Many conservation- ists are well aware of the need to protect in- traspecific diversity but adopt different strategies. In recent years, several approaches have been deployed using “conservation units” instead of dubious subspecies (e.g., Crandall et al., 2000; Fraser & Bernatchez, 2001). But surely it is inappropriate to name sub- species as a convenience and in the absence of well-founded data, no matter how honor- able the intentions are. (3) Data Sources In the introduction to the (sub)generic list of CLECOM 1+1 by Bank et al. (in: Falkner et al., 2001) it is said that “The list presented in this paper is based on the study of hundreds of publications ...; only a selection of them could be cited in the reference list.” | can only imag- ine how many publications the CLECOM com- 230 DAVIS mittee must have had studied. But it certainly would be useful to list all those publications as they are the primary data source and as a check-list can only be as good as the publica- tions it is built upon. Another data source that is often used in check-lists is unpublished information. Philippe Bouchet specifically acknowledged this valu- able source in his introduction to the French list of continental molluscs. However, unpub- lished data are not subject to the scrutiny of the scientific community and its quality and reliability may vary. Therefore, if unpublished data are used in check-lists, they should be clearly marked as such. Another data source of great value for check- lists is molecular data. The advance of robust population genetics, phylogeographic and phylogenetic studies has lead to numerous re- assignments of species, genera and families. Yet, these data are (still) largely neglected in check-lists. To give an example, the molecu- lar work my group has done in the past ten years on several European taxa of the super- family Rissooidea largely has been ignored by CLECOM. Instead, the rissooidean systemat- ics of that list is still mainly based on tradi- tional (mostly shell-based) data. It is not that the molecular genetic data have been ignored because they often contradict the findings of members of the CLECOM team. Rather, it is important that CLECOM incorporate new find- ings based on genetic data much more quickly, even if these findings are inconvenient. One of the declared goals of the CLECOM list is to promote a stable nomenclature of European non-marine molluscs. But this can only be achieved by an objective assessment of all data available. With regard to promotion of a stable nomen- clature (one of the two primary goals of the ICZN), it is important that check-lists are not used to introduce changes in the names of various taxa, species in particular. Apparent novelties in this field must be the subject of careful evaluation by the scientific community before they are proposed as unquestionable to a vast public of non-specialists. Stability of nomenclature is not necessarily achieved by an uncritical application of the law of priority, but through the conservation of the names in use, no matter if they are not the oldest names available, as clearly seen in many findings of the ICZN over the past few decades. The adoption of new systematic hierarchical order- ing in check lists (as done by CLECOM) that were neither adopted before nor previously checked by the scientific community is another critical point we should raise. As such new systems are unknown in the scientific litera- ture or in private or public collections, they in- evitably are a source of confusion, particularly for non-specialists. Moreover, as experience shows, they are inevitably subject to rapid change when they are subjected to rigorous scientific evaluation. Check lists should be a catalogue of all the species of a certain group for a given geo- graphical area, not a vehicle for promoting the reconstruction of phylogenetic history or pro- moting a particular phylogenetic hypothesis. (4) Need for Further Research The introduction to CLECOM begins with the headline “The need for a uniform catalogue to promote biodiversity studies”. And indeed, comparing the various activities ongoing at national levels and formulating coherent syn- theses that can be used by both scientists and policy-makers is a declared goal of the CLECOM committee. In order to indicate taxa that need further research, the CLECOM list uses question marks for those taxa. However, it appears as if those question marks are heavily under- utilized in CLECOM. To use the Bythiospeum example above, none of the 36 dubious spe- cies and subspecies listed for Germany car- ries a question mark. In fact, none of the numerous German taxa of the superfamily Rissooidea (a highly controversial group with many cryptic radiations) is, according to the CLECOM list, in need of further studies. But numerous recent publication using molecular markers have shown that many of these groups are in urgent need of revision. We do not know the reason the above omis- sions, but it seems that CLECOM is trying to suggest that almost everything is known about the systematics and taxonomy of the European non-marine mollusks. This certainly will not promote further research and it will not help to protect biodiversity either. In fact, it only dis- courages biologists from asking meaningful questions and therefore contradicts the de- clared goals of the same CLECOM. In conclusion, the above points highlight some of the problems | see with recently pub- lished check-lists. Success and acceptance of those lists depend not only on the quality and SPECIES CHECK-LISTS 231 quantity of the databases, but also on the abil- ity to consolidate conflicting information and ensure a critical assessment of their own work. In order to promote nomenclatural stability, it is also necessary to clearly state the subspe- cies and species concepts upon which the list is based and the operational criteria used to implement those concepts. LITERATURE CITED BLACKWELDER, R.E., 1967, Taxonomy. A text and reference book. John Wiley & Sons, New York, London, Sydney. BOUCHET, P., 2002, Mollusques terrestres et aquatiques de France: un nouveau référentiel taxonomique, un nouveau départ, de nouvelles perspectives. Pp. 5-20, т: ©. FALKNER, Т. Е. J. RIPKEN 8 М. FALKNER, Mollusques continentaux de France. Liste de référence annotée et Bibliographie. Publications scientifiques du Museum National d'Histoire Naturelle, Paris. CRANDALL, К. A., O. R. P. BININDA-EMONDS, С. М. MACE & БК. К. WAYNE, 2000, Consider- ing evolutionary processes in conservation bi- ology. Trends in Ecology and Evolution, 15: 290-295. DANCE, 5. P., 1970, “Le fanatisme du nobis”: a study of J.-R. Bourguignat and the “Nouvelle Ecole”. Journal of Conchology, 27: 65-86. DANCE, 5. P., 1986, A history of shell collecting. E. J. Brill 8 W. Backhuis, Leiden. DAVIS, G. M., 1979, The origin and evolution of the gastropod family Pomatiopsidae, with em- phasis on the Mekong River Triculinae. Mono- graph of the Academy of Natural Sciences of Philadelphia, 20: i-viii, 1-120. DAVIS, G. M., 1994, Molecular genetics and taxonomic discrimination. Nautilus, 8, Suppl. 2: 3-23. DOBZHANSKY, T., Е. J. AYALA, С. L. STEBBINS 8 J. W. VALENTINE, 1977, Evolution. W.H. Freeman and Company, San Francisco. FALKNER, G., A. BANK 8 T. v. PROSCHWITZ, 2001, Clecom-Project. Check-list of the non- marine molluscan species-group taxa of the States of Northern, Atlantic and Central Europe (CLECOM |). Heldia, 4: 1-76. FALKNER, G., Т.Е. J. RIPKEN 8 М. FALKNER, 2002, Mollusques continentaux de France. Liste de référence annotée et bibliographie. Publications scientifiques du Muséum National d'Histoire Naturelle, Paris. FRASER, D. J. & Е. BERNATCHEZ, 2001, Adap- tive evolutionary conservation: towards a uni- fied concept for defining conservation units. Molecular Ecology, 10: 2741-2752. GIUSTI, Е. & G. MANGANELLI, 1992, The prob- lem of the species in malacology after clear evidence of the limits of morphological system- atics. Pp. 153-172, in: E. GITTENBERGER & J. GOULD, eds., Proceedings of the Ninth Inter- national Malacological Congress, Edinburgh, 1992, Unitas Malacologica, Leiden. HAWKSWORTH, О. L. 8 Е.А. BISBY, 1988, Sys- tematics the keystone of biology. Pp. 3-30, in: D.L. HAWKSWORTH, ed., Prospects in System- atics. The Systematic Association, special vol- ume no. 36, Clarendon Press, Oxford. HULL, D. L., 1997, The ideal species concept — and why we can't get it. Pp. 357-380, in: М. Е. CLARIDGE, H. A. DAWAH 8 M. R. WILSON, eds., Species: the units of biodiversity, Chapman & Hall, London. KOBELT, W., 1881, Exkursionen in Süditalien. Die Sicilianischen lberus. Jahrbücher der Deut- schen Malakologischen Gesellschaft, 8: 50-67. MAYR, E., 1940, Speciation phenomena in birds. American Naturalist, 74: 249-278. MAYR, E., 1963, Animal species and evolution. The Belknap Press of Harvard University Press, Cambridge (Massachusetts, USA), Lon- don (Englana). MAYR, E., 1982, The growth of biological thought: diversity, evolution and inheritance. The Belknap Press of Harvard University Press, Cambridge (Massachusetts, USA), Lon- don (Englana). O'BRIEN, $5. J. & Е. MAYR, 1991, Bureaucratic mischief: recognizing endangered species and subspecies. Science, 251: 1187-1188. Revised ms. accepted 3 June 2004 MALACOLOGIA International Journal of Malacology in > o | po Vol. 46(1) 4 E Publication dates Vol. 36, No. 1-2 8 Jan. 1995 Vol. 37, No. 1 13 Nov. 1995 Vol. 37, No. 2 8 Mar. 1996 Vol. 38, No. 1-2 17 Dec. 1996 Vol. 39, Мо. 1-2 13 May 1998 Vol. 40, No. 1-2 17 Dec. 1998 Vol. 41, No. 1 22 Sep. 1999 Vol. 41, No. 2 31 Dec. 1999 Vol. 42, No. 1-2 18 Oct. 2000 Vol. 43, No. 1-2 20 Aug. 2001 Vol. 44, No. 1 8 Feb. 2002 Vol. 44, No. 2 30 Aug. 2002 Vol. 45, No. 1 29 Aug. 2003 Vol. 45, No. 2 22 Mar. 2004 VOL. 46, МО. 1 MALACOLOGIA 2004 CONTENTS NORBERT BUYSSENS HocomotionriniielixcaSpelrsar Er cic ae En cie 211 MAXIMILIANO CLEDON, LUIZ RICARDO L. SIMONE, & PABLO E. PENCHASZADEH Crepidula cachimilla (Mollusca: Gastropoda), A New Species from BatagoniawArgentiinag mest cas nc cd Male aun. 185 GEORGE M. DAVIS р Species Check-Lists: Death or Revivlal of the Nouvelle Ecole? .......... 227 GEORGE А. EVSEEV, NATALYA К. KOLOTUKHINA, & OLGA YA. SEMENIKHINA Anatomy of a Small Clam, Alveinus ojianus (Bivalvia: Kelliellidae), with a Discussion on the Taxonomic Status of the Family ................... 1 GENNADY M. KAMENEV New Species of the Genus Kellia (Bivalvia: Kellidae) from the Commander Islands, with Notes on Kellia comandorica Scarlato, 1981 .............. 57 GENNADY M. KAMENEV New Species of the Genus Abrina (Bivalvia: Semelidae) from the CommandermancdikuniltislandSass An. o rarer terns oe 157 STEFFEN KIEL Shell Structures of Selected Gastropods from Hydrothermal Vents and SI N A A NA A A ОВ 169 JORIS М. KOENE & IGOR V. MURATOV Revision of the Reproductive Morphology of Three Leptaxis Species (Gastropoda, Pulmonata, Hygromiidae) and lts Implication on Dart Evolution 73 GIUSEPPE MANGANELLI, SIMONE CIANFANELLI, NICOLA SALOMONE, & FOLCO GIUSTI Morphological and Molecular Analysis of the Status and Relationships of Oxychilus paulucciae (De Stefani, 1883) (Gastropoda: Pulmonata: LONA CE ROUE о sie PP sities een tee Neve Lc tml Noster ian re 19 CHRISTOPHER P. MEYER Toward Comprehensiveness: Increased Molecular Sampling within Cypraeidae and Its Phylogenetic Implications ....................... 127, BRIAN MORTON The Biology and Functional Morphology of Foegia novaezelandiae (Bivalvia: Anomalodesmata: Clavagelloidea) from Western Australia ............. 37 IRA RICHLING Coloration in Helicinidae (Mollusca: Gastropoda: Neritopsina) .......... 247 PAWEL WOZNICKI Chromosomes of the Chinese Mussel Anodonta woodiana (Lea 1834) (Bivalvia, Unionidae) from the Heated Konin Lakes System in Poland .... 205 MIN WU Preliminary Phylogenetic Study of Bradybaenidae (Gastropoda: Sy iommatoprora ее ева. 79 ¿ аа | M Ar | E | Li ¡WWW : wi AED peo! y are . az Im mine M bat o RD ИЕ EN no р Lane BT oe ï ul tempor AAA ous © e el A MALACOLOGIA SUBSCRIPTION AND PAST ISSUE ORDER FORM Name: Address: Personal rates: Per volume Subscription $ 56.00 Single volumes $ 56.00 Institutional rates: Subscription $ 75.00 Single volumes $ 75.00 Postage & handling per volume for single/past volumes $ 5.00 e Subcriptions begin with the current volume. 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Regular subscribers are those who have paid-up subscriptions for the current issue and the following issue. Students (including individuals submitting dissertations) must identify themselves at the time of manu- script submission and also provide the e-mail address of their advisor. VOL. 46, МО. 1 MALACOLOGIA 2004 CONTENTS GEORGE А. EVSEEV, NATALYA К. KOLOTUKHINA, & OLGA YA. SEMENIKHINA Anatomy of a Small Clam, Alveinus ojianus (Bivalvia: Kelliellidae), with a Discussion on the Taxonomic Status of the Family ................... 1 GIUSEPPE MANGANELLI, SIMONE CIANFANELLI, NICOLA SALOMONE, & FOLCO GIUSTI Morphological and Molecular Analysis of the Status and Relationships of Oxychilus paulucciae (De Stefani, 1883) (Gastropoda: Pulmonata: Zenitidae) na A NN RENE EE A 19 BRIAN MORTON The Biology and Functional Morphology of Foegia novaezelandiae (Bivalvia: Anomalodesmata: Clavagelloidea) from Western Australia ............. 37 GENNADY M. KAMENEV New Species of the Genus Kellia (Bivalvia: Kellidae) from the Commander Islands, with Notes on Kellia comandorica Scarlato, 1981 .............. 57 JORIS M. KOENE & IGOR V. MURATOV Revision of the Reproductive Morphology of Three Leptaxis Species (Gastropoda, Pulmonata, Hygromiidae) and Its Implication on Dart Evolution 73 MIN WU Preliminary Phylogenetic Study of Bradybaenidae (Gastropoda: Stylommatophora: Helicoidea). ее 79 CHRISTOPHER P. MEYER Toward Comprehensiveness: Increased Molecular Sampling within Cypraeidae and Its Phylogenetic Implications ....................... 127 GENNADY M. KAMENEV New Species of the Genus Abrina (Bivalvia: Semelidae) from the Commander and'Kuñnil Islands : 2. - ааа ол = oe eee eee 157 STEFFEN KIEL Shell Structures of Selected Gastropods from Hydrothermal Vents and SECDS «cw sess scala ee sob co cu : - 169 MAXIMILIANO CLEDON, LUIZ RICARDO L. SIMONE, & PABLO E. PENCHASZADEH Crepidula cachimilla (Mollusca: Gastropoda), A New Species from Patagonia, Argentina ...... u... Jess 185 RESEARCH NOTES PAWEL WOZNICKI Chromosomes of the Chinese Mussel Anodonta woodiana (Lea 1834) (Bivalvia, Unionidae) from the Heated Konin Lakes System in Poland .... 205 NORBERT BUYSSENS Locomotion in Helix aspersa .„.............:- ола а 211 IRA RICHLING Coloration in Helicinidae (Mollusca: Gastropoda: Neritopsina) .......... 217 LETTER FROM THE EDITOR GEORGE M. DAVIS р Species Check-Lists: Death or Вемма! of the Nouvelle Ecole? .......... 227 VAL НЯ y MALACOLOGI UE. Proceedings of the | | Long Key, Florida, . : Edited by Rudiger Bieler and Paula M. Mikkelsen Vol. 46(2) 2004 ча. ? MALACOLOGIA http:\\malacologia.fmnh.org EDITOR-IN-CHIEF: GEORGE M. DAVIS Editorial Office Malacologia РО. Box 1222 West Falmouth, MA 02574-1222 Copy Editor: EUGENE COAN California Academy of Sciences San Francisco, CA Managing Editor: CARYL HESTERMAN Haddonfield, NJ Graphics Editor: THOMAS WILKE Justus Liebig University Giessen, Germany Business & Subscription Office Malacologia P.O. Box 385 Haddonfield, NJ 08033-0309 Associate Editor: JOHN B. BURCH University of Michigan Ann Arbor Assistant Business Managers: KEVIN ROE & STAFF Malacology Department Delaware Museum of Natural History Wilmington, DE MALACOLOGIA is published by the INSTITUTE OF MALACOLOGY, the Sponsor Members of which (also serving as editors) are: RUDIGER BIELER Field Museum, Chicago JOHN BURCH MELBOURNE R. CARRIKER University of Delaware, Lewes GEORGE M. DAVIS Secretary and Treasurer CAROLE S. HICKMAN President University of California, Berkeley ALAN KOHN Vice President University of Washington, Seattle JAMES NYBAKKEN President Elect Moss Landing Marine Laboratory, California CLYDE F. Е. ROPER Smithsonian Institution, Washington, D.C. SHI-KUEI WU University of Colorado Museum, Boulder Participating Members PETER MORDAN Secretary, UNITAS MALACOLOGICA The Natural History Museum London, United Kingdom J. FRANCES ALLEN, Emerita Environmental Protection Agency Washington, D.C. KENNETH J. BOSS Museum of Comparative Zoology Cambridge, Massachusetts JACKIE L. VAN GOETHEM Treasurer, UNITAS MALACOLOGICA Koninklijk Belgisch Instituut voor Natuurwetenschappen Brussel, Belgium Emeritus Members ROBERT ROBERTSON The Academy of Natural Sciences Philadelphia, Pennsylvania W. D. RUSSELL-HUNTER Easton, Maryland Copyright © 2004 by the Institute of Malacology ISSN:0076-2997 J. А. ALLEN Marine Biological Station Millport, United Kingdom Jallen@udcf.gla.ac.uk Е. Е. BINDER Museum d'Histoire Naturelle Geneve, Switzerland P. BOUCHET Muséum National d'Histoire Naturelle Paris, France bouchet@cimrs1.mnhn.fr P. CALOW University of Sheffield United Kingdom R. CAMERON Sheffield United Kingdom R.Cameron@sheffield. ac.uk J. G. CARTER University of North Carolina Chapel Hill, U.S.A. M. CHARRIER Universite de Rennes France maryvonne.charrier@univ-rennes1.fr В. H. COWIE University of Hawaii Honolulu, HI., U.S.A. A. H. CLARKE, Jr. Portland, Texas, U.S.A. B. C. CLARKE University of Nottingham United Kingdom R. DILLON College of Charleston SC, U.S.A. C. J. DUNCAN University of Liverpool United Kingdom D. J. EERNISSE California State University Fullerton, U.S.A. E. GITTENBERGER Rijksmuseum van Natuurlijke Historie Leiden, Netherlands sbu2eg@rulsfb.leidenuniv.de F. GIUSTI Universita di Siena, Italy giustif@unisi.it MCZ 2004 EDITORIAL BOARD LIBRA RY ВА n MANE HAN 10 2005 A. N. GOLIKOV HARVARD Zoological Institute U М IVE RSI ДУ St. Petersburg, Russia A. V. GROSSU Universitatea Bucuresti Romania T. HABE Тока! University Shimizu, Japan R. HANLON Marine Biological Laboratory Woods Hole, Mass., U.S.A. G. HASZPRUNAR Zoologische Staatssammlung München München, Germany haszi@zi.biologie. uni-muenchen. de J. М. HEALY University of Queensland Australia Jhealy@zoology.uq.edu.au D. М. HILLIS University of Texas Austin, U.S.A. К. Е. HOAGLAND West Falmouth, U.S.A. B. HUBENDICK Naturhistoriska Museet Göteborg, Sweden S. HUNT Lancashire United Kingdom R. JANSSEN Forschungsinstitut Senckenberg, Frankfurt am Main, Germany M. S. JOHNSON University of Western Australia Nedlands, WA, Australia msj@cyllene.uwa.edu.au В. М. KILBURN Natal Museum Pietermaritzburg, South Africa M. A. KLAPPENBACH Museo Nacional de Historia Natural Montevideo, Uruguay J. KNUDSEN Zoologisk Institut Museum Kobenhavn, Denmark С. LYDEARD University of Alabama Tuscaloosa, U.S.A. clydeard@biology.as.ua.edu C. MEIER-BROOK Tropenmedizinisches Institut Tübingen, Germany H. K. MIENIS Hebrew University of Jerusalem Israel J. E MORTON The University Auckland, New Zealand J. J. MURRAY, Jr. University of Virginia Charlottesville, U. S.A. R. NATARAJAN Marine Biological Station Porto Novo, India D. Ó FOIGHIL University of Michigan Ann Arbor, U.S.A. J. OKLAND University of Oslo Norway T. OKUTANI University of Fisheries Tokyo, Japan W. L. PARAENSE Instituto Oswaldo Cruz, Rio de Janeiro Brazil J. J. PARODIZ Carnegie Museum Pittsburgh, U.S.A. R. PIPE Plymouth Marine Laboratory Devon, United Kingdom RKPI@wpo.nerc.ac.uk J.P POINTIER Ecole Pratique des Hautes Etudes Perpignan Cedex, France pointier@gala. univ-perp. fr W. F. PONDER Australian Museum Sydney QUIZA Academia Sinica Qingdao, People's Republic of China D. G. REID The Natural History Museum London, United Kingdom S. G. SEGERSTRÁLE Institute of Marine Research Helsinki, Finland A. STANCZYKOWSKA Siedice, Poland F. STARMÚHLNER Zoologisches Institut der Universität Wien, Austria У. |. STAROBOGATOV Zoological Institute St.Petersburg, Russia J. STUARDO Universidad de Chile Valparaiso C. THIRIOT University Pet M.Curie Villefranche-sur-Mer, France thiriot@obs-vifr. fr S. TILLIER Museum National d'Histoire Naturelle Paris, France J. À. M. VAN DEN BIGGELAAR University of Utrecht The Netherlands N. Н. VERDONK Rijksuniversiteit Utrecht, Netherlands Н. WÁGELE Ruhr-Universität Bochum Germany Heike.Waegele@ruhr-uni-bochum.de A. WAREN Swedish Museum of Natural History Stockholm, Sweden В. В. WILSON Dept. Conservation and Land Management Kallaroo, Western Australia H. ZEISSLER Leipzig, Germany A. ZILCH Forschungsinstitut Senckenberg Frankfurt am Main, Germany BIVALVE STUDIES IN THE FLORIDAKEYS Proceedings of the International Marine Bivalve Workshop Long Key, Florida, July 2002 GUEST EDITORS Rüdiger Bieler Department of Zoology (Invertebrates) Field Museum of Natural History, Chicago Paula M. Mikkelsen Division of Invertebrate Zoology American Museum of Natural History, New York \ MALACOLOGIA, 2004, 46(2): 241-248 INTERNATIONAL MARINE BIVALVE WORKSHOP 2002: INTRODUCTION AND SUMMARY Paula M. Mikkelsen’ & Rüdiger Bieler? In July 2002, a two-week workshop on ma- rine bivalves, with an emphasis on systemat- ics, anatomy, and natural history, was organized to further knowledge of living ma- rine bivalves and to train graduate-level stu- dents in this understudied field of modern malacology. With support from and in the spirit of the National Science Foundation's Райпег- ships in Enhancing Expertise in Taxonomy (PEET) program, students worked one-on-one in teams with expert scientists on selected bi- valve species or groups of species. This vol- ume, for the most part comprising papers co-authored by the scientist-student research teams, represents the scientific results of projects initiated at the workshop. The Florida Keys at the southernmost tip of peninsular Florida, a region emphasized by the organizers' joint research program since 1994, formed a biologically diverse and logistically convenient site for a workshop of this type. As defined by this research venture, the Florida Keys includes the entire island chain and sur- rounding waters, from Broad Creek at the northern end of Key Largo (including Card and Barnes sounds, but not Biscayne Bay) through and including the Dry Tortugas, plus the ap- proximate southeastern half of Florida Bay (ex- cluding the more brackish areas in the outfall of the Florida Everglades), and offshore ar- eas to the reef line and beyond (with collec- tion and literature records to a maximum depth FIG. 1. Field stations (black diamonds) sampled during the International Marine Bivalve Workshop, July 2002. Re-sampled or neighboring stations are here combined; see text for individual station data. Station FK-619 (Lake Surprise), about 48 km northeast, is not shown. West-to-east extent of the displayed area, ranging from Big Pine Key to Upper Matecumbe, is approximately 85 km. ‘Division of Invertebrate Zoology, American Museum of Natural History, Central Park West at 79th Street, New York, New York 10024-5192, U.S.A.; mikkel@amnh.org ¿Department of Zoology, Division of Invertebrates, Field Museum of Natural History, 1400 $. Lake Shore Drive, Chicago, Illinois 60605-2496, U.S.A.; bieler@fieldmuseum.org 242 MIKKELSEN & BIELER of 300 т). Included within these limits is the Florida Keys National Marine Sanctuary, the second largest marine sanctuary in the United States, as well as a variety of other jurisdic- tions protecting various terrestrial and marine sites. The workshop was held at the Keys Ma- rine Laboratory on Long Key, a venue centrally located in the Keys allowing ready access to a large portion of the archipelago including off- shore and bayside habitats (Fig. 1). Easily ac- cessible habitats included intertidal rocks, sand and seagrass flats, rock ledges, seawalls, mangroves, mud channels, patch reefs, back reefs, artificial reefs (shipwrecks), and the only living near-shore coral reefs in the continental United States. Tourism is the leading industry of Florida and the Florida Keys has long been a favorite des- tination for ocean-oriented vacationing, diving, sport fishing, and shell collecting. In spite of a century of avid shell collecting and molluscan research in the Keys, Levy et al. (1996) noted, “except for a few ecological inventories that include mollusks, there is a lack of compre- hensive, ecosystem-wide species inventories in the Florida Keys.” Coral reef conservation efforts stress corals, sponges, algae, spiny lobster and fish, and, except for a few mem- bers of the “charismatic macrofauna” [e.g., the queen conch, Strombus gigas Linnaeus, 1758, and Flamingo Tongue, Cyphoma gibbosum (Linnaeus, 1758)], regularly ignore mollusks. This deficiency has been acutely sensed since the establishment of the Florida Keys National Marine Sanctuary (FKNMS) in 1990, created to protect and restore fragile marine habitats from the environmental impact of human use. The FKNMS Draft Management Plan listed only 630 marine species (Lyons & Quinn, 1995), only slightly fewer than a considerably earlier yet little-known, privately published list of 710 species by Lermond (1936). The mol- luscan species list compiled by the organiz- ers’ research program has more than doubled to nearly 1,700 species through original field- work, museum collection surveys, and exten- sive literature research; approximately 400 of these species are bivalves (Mikkelsen & Bieler, 2000; Bieler & Mikkelsen, 2004). Twelve international specialists in marine bivalve systematics participated by mentoring a student during the workshop. They included scientists with a wide range of specialties, spanning functional morphology, phylo- genetics, molecular biology, and faunal diver- sity research. А list of readily obtained Florida Keys bivalves was provided early in the plan- ning process, allowing each scientist to select and prepare materials to study a taxon in the field. The 12 student participants were selected from 49 applicants in response to notices dis- tributed at meetings and on internet listservers. Research teams were formed by pairing an alphabetical list of scientists with a reverse- alphabetical list of students. Together with the six members of the organizing team, this was a highly international group of 30 individuals representing 17 nations and five continents of origin or residence (Fig. 2). The participants were: Mr. Kyle Bennett, Rutgers University, New Brunswick, New Jersey, U.S.A. Dr. Rudiger Bieler, Field Museum of Natural History, Chicago, Illinois, U.S.A. [organizer] Mr. Gregorio Bigatti, Universidad de Buenos Aires, Argentina. Mr. Matthew Campbell, Indiana University, Bloomington, Indiana, U.S.A. Mr. Anton Chichvarkhin, Rossiiskoi Akademii Nauk, Vladivostok, Russia. Ms. Louise Crowley, City University of New York and American Museum of Natural His- tory, New York, New York, U.S.A. [organiz- ing team]. Ms. Grete Dinesen, University of Aarhus, Den- mark. Dr. Osmar Domaneschi, Universidade de Sao Paulo, Brazil. Ms. Joanne Dougherty, Villanova University, Villanova, Pennsylvania, U.S.A. Dr. Emily Glover, The Natural History Museum, London, United Kingdom. Ms. Johanna Jarnegren, Norges Teknisk- Naturvitenskapelige Universitet, Trondheim, Norway. Ms. Isabella Kappner, University of Illinois at Chicago and Field Museum of Natural History, Chicago, Illinois, U.S.A. [organizing team]. Ms. Lisa Kirkendale, Florida Museum of Natu- ral History, Gainesville, Florida, U.S.A. Ms. Martina Knapp, Universitat Wien, Austria. Dr. José H. Leal, The Bailey-Matthews Shell Museum, Sanibel, Florida, U.S.A. Ms. Amy Maxmen, Harvard University, Cam- bridge, Massachusetts, U.S.A. Dr. Paula M. Mikkelsen, American Museum of Natural History, New York, New York, U.S.A. [organizer]. Dr. Russell Minton, Field Museum of Natural History, Chicago, Illinois, U.S.A. [organizing team]. IMBW 2002 — INTRODUCTION AND SUMMARY 243 Ms. Juri А. Miyamae, Swarthmore College, Pennsylvania, and American Museum of Natural History, NSF-REU Summer Intern Program, New York, New York, U.S.A. [or- ganizing team]. Prof. Brian Morton, Swire Institute of Marine Science, University of Hong Kong, China. Dr. Diarmaid О Foighil, Museum of Zoology, University of Michigan, Ann Arbor, Michigan, U.S.A. Dr. P. Graham Oliver, National Museum of Wales, Cardiff, United Kingdom. Ms. Melita Peharda, Institut za Oceanografiju ¡ Ribarstvo, Split, Croatia. Dr. Jay Schneider, George Washington Uni- versity, Washington, DC, U.S.A. Ms. Elizabeth K. Shea, Bryn Mawr College, Bryn Mawr, Pennsylvania, U.S.A. Dr. Luiz Ricardo L. Simone, Museu de Zoologia, Universidade de Sáo Paulo, Brazil. Dr. Gerhard Steiner, Universitát Wien, Austria. Prof. John Taylor, The Natural History Mu- seum, London, United Kingdom. Mr. Paul Valentich Scott, Santa Barbara Mu- seum of Natural History, Santa Barbara, California, U.S.A. Dr. Richard Willan, Northern Territory Museum of Arts & Sciences, Darwin, Australia. The workshop occupied nearly the entire Keys Marine Laboratory facility, including dor- mitories, wet laboratory for sorting, dry labo- ratory for microscope work and photography, classroom for presentations and discussion, and small boats for snorkeling trips. Various vehicles facilitated land travel for collecting and other group events; some scuba trips utilized commercial dive boats. Group and team ac- tivities included collecting by snorkeling, scuba, shovel-and-sieving (Fig. 3), and crack- ing dead coral rocks, followed by appropriate laboratory study (Fig. 4) and sharing their find- ings through discussions and presentations. Each research team included at least one cer- tified scuba diver and this allowed exploration of additional habitats, including an offshore FIG. 2. Participants of the IMBW gather for a group photograph at Pigeon Key (photograph Бу |. Simone). 244 MIKKELSEN & BIELER FIG. 3. Shovel-and-sieving typified collecting efforts for shallow-water bivalves in the Florida Keys. FIG. 4. Following a day's collecting efforts, the laboratory filled with a variety of study activities. IMBW 2002 - INTRODUCTION AND SUMMARY 245 wreck that had become a habitat for several deeper-water bivalve species. In deference to the sanctuary location, no dredging was con- ducted, and no protected species were col- lected; workshop activities were intentionally designed around relatively common, shallow- water species. Lectures were presented on most evenings, either by guest speakers or the participants. Each scientist spoke of his/ her research or laboratory, and toward the end of the workshop, each student summarized the results of their team investigations. The IMBW documented 121 species of bivalves from 48 field stations (Fig. 1), includ- ing several previously unrecognized taxa and others of poorly known distribution and habi- tat. Voucher specimens are deposited in the mollusk collections at AMNH, FMNH, and the home institutions of some participants (includ- ing Florida Museum of Natural History, Gainesville; Bailey-Matthews Shell Museum, Sanibel Island, Florida; Museum of Zoology, University of Michigan, Ann Arbor; Santa Bar- bara Museum of Natural History, California; The Natural History Museum, London, United King- dom; Museu de Zoologia, Universidade de Sáo Paulo, Brazil; and Northern Territory Museum of Arts & Sciences, Darwin, Australia). WORKSHOP STATIONS IMBW-FK-619, 14-VII-02, Lake Surprise, Key Largo, MM 107.5, NE end of U.S. Rte. 1 causeway across lake, 25°10.9’М, 80°23.0’М/, off mangroves at side of road, by hand on shallow subtidal rocks. IMBW-FK-620, 16- & 18-VII-02, Old Dan Bank, bayside of Long Key, 24°50.45’N, 80°49.63'W, Thalassia seagrass bed with Halimeda calcareous algae, Porites finger- coral, sponges, hydroids, patches of sand/ Halimeda shell hash, by hand, 0.3-0.6 т, R/V FLORIDAYS. IMBW-FK-621, 17-VII-02, “Long Key Artificial Reefs”, oceanside of Long Key, 24*44.78'N, 80°50.00’W, sand plain with Thalassia/ Syringodeum seagrass patches, scuba, by hand, 7 m, R/V FLORIDAYS. IMBW-FK-622, 20-VII-02, directly off Keys Marine Laboratory, bayside of Long Key, 24°49.5’М, 80°48.9’W, seagrass bed with coral rubble, snorkeling, sieving, by hand, 0-1.5 m. IMBW-FK-622A, 22-VII-02, off Keys Marine Laboratory, bayside of Long Key, 24°49.5’N, 80748.9'W, about 30 m from shore, thin sand over rock, with Halodule, Thalassia, Syringo- dium, Halimeda, shovel/sieving, 0.5-1 m. IMBW-FK-623, 20-VII-02, Long Key State Park, oceanside, 24°48.67’N, 80°49.68’W, seagrass bed (predominantly Thalassia) on muddy sand, snorkeling, by hand, 0-0.75 m. IMBW-FK-624, 20-VII-02, Horseshoe Reef, off Fat Deer Key, 24°39.91’N, 80°59.56’W, patch reef with sandy bottom, scuba, 7.3 m, M/V SHUTTERBUG II. IMBW-FK-625, 20-VII-02, Coffins Patch Sanc- tuary Preservation Area, off Crawl Key, 24°40.92’ N, 80°58.26’ W, patch reef with sand patches, gorgonian, pillar coral, scuba, 6.4 m, M/V SHUTTERBUG II. IMBW-FK-626, 21-VII-02, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), ММ 35, 24°39.3’N, 81°18.2’W, “hole” at center of quarry, rock wall and soft sediment, snorkeling and scuba, to ca. 6.1 m. IMBW-FK-627, 21-VII-02, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), MM 35, 24°39.3’N, 81°18.2’W, mangrove area, soft sediment and detritus, hand dredge, < 1 т. IMBW-FK-628, 21-VII-02, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), ММ 35, 24°39.3’N, 81°18.2’W, Thalassia seagrass, shovel/ sieve, ca. 1 m. IMBW-FK-629, 21- 8 26-VII-02, “The Horse- shoe” site, bayside of West Summerland Key (Spanish Harbor Keys), ММ 35, 24°39.3’N, 81°18.2’W, among rocks along arms of quarry, by hand, snorkeling, to ca. 1 т. IMBW-FK-630, 22-VII-02, roadside quarry М of Keys Marine Laboratory, Long Key, 24°49.78’М, 80%48.51'W, rock wall and scuzzy algae, hot (36°С) water layer, snor- keling, by hand, > 1 т to horizontal ledge on wall [total depth of quarry not assessed], S = 31 ppt. IMBW-FK-631, 22-VII-02, Burnt Point, bayside, N point of Long Point Park, 24°45.56’ М, 80°59.14’ W, rocky bottom, soft coral/sponges, patches of seagrass, snor- keling, 0.6-1.2 m [2.4 m in channel along shore]. IMBW-FK-632, 22-VII-02, Bahia Honda State Park, oceanside, just E of old bridge, 24739.25'N, 81°16.83’W, seagrass beds with sand blowholes, snorkeling at low tide, 0.6 m. IMBW-FK-633, 22-VII-02, Missouri Key, 24°40.5’М, 81°14.3’W, coral rubble and 246 MIKKELSEN & BIELER seagrass beds, snorkeling and by hand, in- tertidal zone to 1 m. IMBW-FK-634, 22-VII-02, Bahia Honda State Park, oceanside, 24°39.69’М, 81°16.11’W, at small roadway bridge over channel W before Sandspur Campground, sandy bottom at low tide, sparse seagrass, by hand, 0-0.1 т. IMBW-FK-635, 22-VII-02, Veteran's Beach, oceanside, Little Duck Key, 24°40.87'N, 81°13.82’W, Thalassia/Halodule seagrass on silty sand, shovel/sieve, low intertidal zone to shallow subtidal. IMBW-FK-636, 22-VII-02, Long Key State Park, oceanside, 24°48.67’М, 80°49.68’W, sandy beach, seagrass, by hand, snorkel- ing, intertidal zone. IMBW-FK-637, 22-VII-02, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), MM 35, 24°39.3’М, 81°18.2’W, among rocks/rubble along arms of quarry, by hand, intertidal zone. IMBW-FK-638, 23-VII-02, Anne’s Beach, oceanside, Craig Key, MM 72, 24°50.95’М, 80°44.40'W, Thalassia/Halodule seagrass, shovel/sieve, by hand, 0.5-1 т. IMBW-FK-638A, 26-VII-02, Anne's Beach, oceanside, Craig Key, MM 72, 24°50.95'N, 80°44.40’W, Thalassia/Halodule seagrass, shovel/sieve, by hand, 0.5-1 т. IMBW-FK-639, 23-VII-02, Coral Gardens in- shore patch reef, oceanside off Lower Matecumbe Key, 24°50.23’М, 80°43.77’W, snorkeling, 3.6-4.6 m, Keys Marine Labora- tory boat. IMBW-FK-640, 23-VII-02, oceanside off Craig Key, 24°49.81’М, 80°45.73’W, nearshore patch reef, hardbottom, snorkeling, 0.1-1.2 m, Keys Marine Laboratory boat. IMBW-FK-641, 23-VII-02, Tennessee Reef, off Long Key, 24°44.75’М, 80°46.95’W, hard bottom with coral, scuba, 7 m, R/V FLORIDAYS. IMBW-FK-642, 23-VII-02, “The Billboard” site, oceanside, Lower Matecumbe Key, MM 74.5, 24°51.4’N, 80°43.7’W, thin sand cover on rock platform, small coral, Thalassia/ Halodule seagrass, Sargassum, wading, snorkeling, shovel/sieving, 0.5-1 m. IMBW-FK-643, 23-VII-02, Fiesta Key cause- way, oceanside, 24°50.41’М, 80°46.95’W, at turnoff W of Channel #5 bridge, rocky shore, sand, Thalassia seagrass, snorkeling, by hand, 0-3 т. IMBW-FK-644, 23-VII-02, Fiesta Key cause- way, bayside, 24°50.41’М, 80°46.95’W, at turnoff W of Channel #5 bridge, rocky shore, concrete pilings, snorkeling, by hand, 0-3 m. IMBW-FK-645, 24-VII-02, Grassy Key, oceanside, 24°46.60’N, 80°55.44’W, at turn- off before Tom’s Harbor Channel, attached to underside of large, algal-encrusted boul- ders/rocks along shore, by hand, snorkeling, 1-3 m. IMBW-FK-646, 25-VII-02, “The Billboard” site, oceanside, Lower Matecumbe Key, MM 74.5, 24°51.4’М, 80°43.7’W, rubble and sand with seagrass, wading, shovel/sieving, 0.5- 0.75 m. IMBW-FK-646A, 24-VII-02, “The Billboard” site, oceanside, Lower Matecumbe Key, MM 74.5, 24°51.4’N, 80°43.7’W, thin sand on rock plat- form, with Thalassia, Halodule, Halimeda, Penicillus, shovel/sieving, 0.5-1 т. IMBW-FK-646B, 27-VII-02, “The Billboard” site, oceanside, Lower Matecumbe Key, MM 74.5, 24°51.4’N, 80°43.7’W, thin sand on rock plat- form, with Thalassia, Halodule, Halimeda, Penicillus, shovel/sieving, 0.5-1 т. IMBW-FK-647, 25-VII-02, W side of Pigeon Key, 24°42.2’N, 81°09.3’W, Thalassia/ Halodule/Syringodeum seagrass on sand/ rubble, concrete bridge piers, by hand, snor- keling, shovel/sieving, 0.5-1 m. IMBW-FK-648, 26-VII-02, Tennessee Reef Light, off Long Key, 24°44.75’М, 80°46.95’W, patch reef, sand, rubble, scuba, 4-7 т, R/V FLORIDAYS. IMBW-FK-649, 27-VII-02, Sprigger Bank, bayside, just W of Everglades National Park border, 24°54.75’М, 80°56.24’W, Thalassia/ Syringodeum seagrass, snorkeling, shovel/ sieving, 0.1-0.9 т, Keys Marine Laboratory boat. IMBW-FK-650, 27-VII-02, wreck of “Thunder- bolt”, approx. 6 nmi $ of Marathon, 24°39.68’N, 80%57.82'W, steel wreck with fouling bivalves, alcyonarians and hydroids, orange/ red sponge overcoating most specimens, scuba, 34.1 m, M/V SHUTTERBUG II. IMBW-FK-651, 27-VII-02, “Samantha’s patch reef’, approx. 5 nmi S of Marathon, 24°39.49’N, 81°00.32’W, coral rock inter- spersed with sandy channels, scuba, 7.6 m, M/V SHUTTERBUG ll. IMBW-FK-652, 27-VII-02, Long Key State Park, oceanside, 24°48.67’N, 80°49.68’W, seagrass bed (predominantly Thalassia) on muddy sand, wading, shovel/sieving, less than 1 т. IMBW-FK-653, 27-VII-02, Old Dan Bank, bayside of Long Key, 24°50.45’N, 80°49.63'W, Thalassia seagrass bed with Halimeda calcareous algae, Porites finger- coral, sponges, hydroids, patches of sand/ IMBW 2002 — INTRODUCTION AND SUMMARY 247 Halimeda shell hash, snorkeling, 0.3-1.5 т, R/V LAST MANGO. IMBW-FK-654, 28-VII-02, East Turtle Shoal, oceanside off Grassy Key, 24°43.49’N, 80°56.00’W, at Marker “45” in Hawk Chan- nel, silty patch reef, scuba, 7.5 m, R/V FLORIDAYS. IMBW-FK-655, 28-VII-02, Veteran’s Beach, oceanside, Little Duck Key, 24°40.87’N, 81°13.82’W, Thalassia/Halodule seagrass on silty sand, shovel/sieve, low intertidal zone to shallow subtidal. IMBW-FK-656, 28-VII-02, mangrove channel near Goshen House, South Layton Drive, Layton, Long Key, 24°49.40’N, 80°48.77’W, red mangrove roots, snorkeling, 0.6-1.2 m. IMBW-FK-657, 28-VII-02, Pigeon Key, 24°42.2’N, 81°09.3'W, seagrass, sand, rubble, by hand, wading, 0-0.5 т. IMBW-FK-658, 26-VII-02, E end of Big Pine Key, Spanish Harbor Channel, 24°38.89’N, 81°19.80’W, pier/pilings, algae-covered rocks, snorkeling, hammer/chisel, 0-2 т. IMBW-FK-659, 28-VII-02, Pigeon Key, 24°42.2'N, 81°09.3’W, seagrass, scuba, 0.6- 1.2 m. IMBW-FK-660, 28-VII-02, Old Dan Bank, bayside of Long Key, 24°50.08’М, 80°49.63’W, Thalassia seagrass bed with Halimeda calcareous algae, Porites finger- coral, sponges, hydroids, patches of sand/ Halimeda shell hash, snorkeling, 0.3-1.5 m, R/V LAST MANGO. The contributions to this proceedings vol- ume reflect the specialized interests of the par- ticipants. The projects were initiated at the 12-day workshop but required substantial fol- low-up between scientist and student, often communicating and even visiting across con- tinents or oceans. The majority of the studies focused on detailed investigations of the com- parative and functional anatomy/morphology of exemplar species in the families Arcidae, Donacidae, Psammobiidae, Pteriidae, and Veneridae. Others took a somewhat broader taxonomic approach and developed regional systematic studies based on morphology and/ or molecules. These resulted in reviews of Florida Keys oysters (Gryphaeidae and Ostreidae), boring bivalves (Gastrochaenidae, Mytilidae, and Petricolidae), and western At- lantic Chamidae. Again other teams, using different experimental setups and analytical approaches, studied predator-prey interac- tions between naticid gastropods and venerid bivalves, or made fine- and ultrastructural in- vestigations into aspects of periostracal mor- phology, and oocyte and sperm development in the family Lucinidae. Two additional papers on the entire bivalve fauna of the region, com- piled from the organizers’ long-term research program, provide a broader look at the diver- sity of the Florida Keys bivalve fauna. ACKNOWLEDGMENTS Major funding for this workshop was pro- vided by the National Science Foundation PEET program (DEB-9978119), as part of a grant on marine bivalves to RB and PMM. Additional support was provided by the Ber- tha LeBus Charitable Trust, Comer Science 8 Education Foundation, Field Museum's Women’s Board, as well as other institutional funds from AMNH and FMNH. José Leal gen- erously provided supplemental transportation through use of the Bailey-Matthews Shell Museum vehicle. Collections were supported by permits from the Florida Keys National Marine Sanctuary (educational event permit FKNMS-2002-079), the State of Florida (in- dividual saltwater fishing licenses to all par- ticipants), Florida Department of Environmental Protection (FDEP 5-02-43; for Long Key and Bahia Honda State Parks), and Pigeon Key Foundation. The organizers thank members of the organizing team as listed above, as well as Julia Sigwart (AMNH), for their assistance with numerous aspects of planning and running the work- shop. Special thanks are extended to three internationally recognized bivalve specialists, Drs. John Allen (University Marine Biologi- cal Station Millport, Scotland), Kenneth J. Boss (Museum of Comparative Zoology, Harvard University), and Eugene V. Coan (Palo Alto, California), who assisted in re- viewing the student applicants' qualifications and choosing the awardees. Thanks are also due to the numerous colleagues who cri- tiqued and improved the manuscripts through rigorous peer review, and to George M. Davis and Eugene V. Coan who coordinated re- views of the editors’ own papers. Workshop activities were facilitated and enhanced by Billy Causey (Superintendent, Florida Keys National Marine Sanctuary), Dr. Erich Mueller (Mote Marine Laboratory, Center for Tropi- cal Research) and the staff of Keys Marine Laboratory. 248 MIKKELSEN & BIELER LITERATURE CITED BIELER, К. 8 P. M. MIKKELSEN, 2004, Marine bivalves of the Florida Keys: a qualitative fau- nal analysis based on original collections, mu- seum holdings and literature data. In: к. BIELER & P. M. MIKKELSEN, eds., Bivalve Studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 503-544. LERMOND, М. W., 1936, Check list of Florida marine shells. Privately published, Gulfport, Florida. 56 pp. LEVY, J. М., M. CHIAPPONE & К. M. SULLIVAN, 1996, Invertebrate infauna and epifauna of the Florida Keys and Florida Bay. Site character- ization for the Florida Keys National Marine Sanctuary and environs, vol. 5: 1-166, The Na- ture Conservancy, Florida and Caribbean Ma- rine Conservation Science Center, University of Miami & The Preserver, Zenda, Wisconsin. LYONS, W. С. & J. Е. QUINN, Jr., 1995, Appen- dix J. Marine and terrestrial species and algae: Phylum Mollusca. J-10 — J-26, in: Florida Keys National Marine Sanctuary Draft Management Plan/Environmental Impact Statement, Vol. III. United States Government Printing Office, Washington, D.C. MIKKELSEN, P. M. 8 R. BIELER, 2000, Marine bivalves of the Florida Keys: discovered biodiversity. Pp. 367—387, in: The evolution- ary biology of the Bivalvia [Proceedings of Bi- ology & Evolution of the Bivalvia, an international symposium organized by the Malacological Society of London, 14-17 Sep- tember 1999, Cambridge, UK], E. M. HARPER, J. D. TAYLOR & J. A. CRAME, eds. Geological So- ciety, London, Special Publication 177. MALACOLOGIA, 2004, 46(2): 249-275 SHELL MORPHOMETRY OF WESTERN ATLANTIC AND INDO-WEST PACIFIC ASAPHIS; FUNCTIONAL MORPHOLOGY AND ECOLOGICAL ASPECTS OF А. DEFLORATA FROM FLORIDA KEYS, U.S.A. (BIVALVIA: PSAMMOBIIDAE) Osmar Domaneschi' & Elizabeth К. Shea? ABSTRACT The genus Asaphis has long been considered monotypic, with A. deflorata having a worldwide, tropical distribution. Recent research has provided evidence for a tropical, western Atlantic species, A. deflorata, and a tropical Indo-West Pacific species, A. violascens. Ecological and other aspects of the biology of these species have been studied exten- sively but morpho-functional features have been known for the Indo-West Pacific species only, so that separation of them has been based on shell sculpture alone. This paper examines the shell morphometry of western Atlantic and Indo-West Pacific specimens of Asaphis, and the functional morphology and ecological aspects of a population of the genus present in the Florida Keys, USA. It is our aim to improve knowledge about the biology of the western Atlantic Asaphis and identify new characters that may support either their monotypy or the two valid species hypothesis. In addition to confirming that shell sculpture may be a good character in distinguishing both forms, our ecological and mor- pho-functional data also concur in validating A. deflorata as a distinct species. Growth rates, maturity levels and predominantly upper shore intertidal position of the Keys popu- lation are consistent with a previous study of the Bahamas population of A. deflorata. т both studied areas, A. deflorata constitutes the sole bivalve present in the upper shore; conversely, most specimens of the Indo-West Pacific A. violascens occupy an intermedi- ate to subtidal position, and share the intertidal region horizontally and vertically with other species of bivalves. The functional anatomy of A. deflorata is very similar to that of A. violascens; however, the hind gut provides a useful parameter for separation of both spe- cies, as it progressively widens, coils and spirals in the Atlantic form, whereas it has an extraordinary dilation in its proximal end only, in the Indo-West Pacific A. violascens. Key words: functional morphology, shell morphometry, ecology, Asaphis, Psammobiidae, Bivalvia. INTRODUCTION Bivalves of the genus Asaphis Modeer, 1793, are commonly found intertidally in gravelly sand, cobble-covered sediments (Depledge, 1985; Britton, 1985; Berg & Alatalo, 1985) or around mangrove roots (Coomans, 1969; Stanley, 1970; Berg € Alatalo, 1985). Populations at- tain sufficiently high densities to support sus- tained collection for human consumption (Fisher, 1978; Berg € Alatalo, 1985; Willan, 1993), and their viability as an aquaculture re- source has been investigated (Berg & Alatalo, 1981, 1985). The natural variability in shell color gives the Atlantic Asaphis its common name, gaudy asaphis, and a place in shellcraft indus- try (Abbott, 1974; Berg & Alatalo, 1985). Prashad (1932) considered the Indo-Pacific specimens of Asaphis as belonging to A. dichotoma (Anton, 1838), and distinct from those living in the Western Atlantic assigned to A. deflorata (Linné, 1758). Abbott (1950) considered both Indo-Pacific and western At- lantic forms to be conspecific, and proposed that the genus is monotypic, with A. deflorata having a worldwide, tropical distribution. This hypothesis was codified in Abbott (1974), but more recent research (Willan, 1993) provides evidence for a tropical western Atlantic spe- cies, A. deflorata, and a tropical Indo-West Pacific species, A. violascens (Forsskál, 1775). ‘Departamento de Zoologia, Instituto de Biociéncias, Universidade de Sáo Paulo, PO Box 11461, CEP 05422-970, Sao Paulo (SP), Brazil; domanesc@ib.usp.br ¿Department of Biology, Bryn Mawr College, Bryn Mawr, Pennsylvania, U.S.A.; eshea@brynmawr.edu 250 DOMANESCHI & SHEA The Indo-Pacific Asaphis was generally known as A. dichotoma until the early 1970s (Willan, 1993), when Cernohorsky (1972) al- tered its name, without explanation, to A. violascens. Willan (1993) has vindicated such an alteration. In this paper, Willan observed that the separation between A. deflorata and A. violascens was based on shell sculpture alone. Actually, there have been no compre- hensive studies to date on the functional mor- phology of the Atlantic Asaphis that allow comparison with that performed by Purchon (1960) on the stomach, and by Narchi (1980) on the functional anatomy of the Indo-Pacific species. Specimens of Asaphis from the Indo- Pacific Ocean (Singapore?), were sent by Purchon to R. Tucker Abbott, Pennsylvania, USA, who identified them as A. deflorata (Purchon, 1960). Ecological and biological data on Asaphis spp. have been provided by Stanley (1970), Narchi (1980), Britton (1985), Depledge (1985), Berg & Alatalo (1985), Soemodihardjo & Matsukuma (1989), Willan (1993), and Kurihara et al. (2000, 2001). This paper examines the shell morphometry of the two Asaphis spp. sensu Willan (1993), and the functional morphology and ecological aspects of Asaphis deflorata from the Florida Keys, USA. It is also the aim of this study to identify characters that may support Abbott's (1950) or Willan's (1993) proposal. MATERIALS AND METHODS Survey Site Asaphis deflorata was collected during the International Marine Bivalve Workshop (IMBW) held in the Florida Keys, USA, 19-30 July 2002 from Station IMBW-FK-629, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), MM 35, Monroe County, Florida Keys 24°39.3’М, 81°18.2’W. Mikkelsen & Bieler (2004) provide a listing of all stations and a map of the studied area. Collections were made in accordance with permit requirements of the State of Florida, under a Research/Collecting Permit issued by the U. S. Department of Commerce, National Oceanic and Atmospheric Administration, Na- tional Ocean Service to Drs. Paula Mikkelsen and Rudiger Bieler (Permit FKNMS 2002-079). Individual Florida Saltwater Fishing Licenses (FSFL) numbers were “M-N112Y018675” (OD) and “M-N1A79018604” (EKS). Field Survey Sampling was performed at low tide when the clams' intertidal habitat was fully exposed; waters receded - 4 т from the high tide mark during the sampling period. Specimens were collected along a 4 m transect, from four 0.25 m? quadrats. The first quadrat was placed at the low water mark, and three additional samples were taken at 0.5 m intervals moving toward the high tide mark. The quadrat area was excavated to 10-12 cm, the maximum depth allowed by the local rocky gravel sub- stratum along the transect. All sediments were sieved through two sieves, with mesh open- ings of 10.0 mm and 0.4 mm, respectively. Behavior Behavior of A. deflorata was observed in the Keys Marine Laboratory on Long Key, and tak- ing photographs in the field and snorkeling dur- ing high tide. Results were compared with those obtained by Stanley (1970) and Berg & Alatalo (1985) observing specimens both in laboratory and in field. Laboratory observations included analyses of the burrowing period (Stanley, 1970) of seven individuals each of 1.0, 1.1, 1.3, 1.8, 2.5, 4.5, and 5.7 cm shell length, lying either on the right or left shell valve on a coarse sand substratum free from natural obstacles as pebbles, shells and rubbles. The ability of the species to surmount sediment deposits through maximum extension of the siphons was evalu- ated by firmly trapping ten specimens (shell length range: 2.5-5.7 cm) posterior end up among pebbles on the bottom of small aquaria. This procedure simulates the condition in which several specimens were found in nature: wedged and trapped both in crevices and among pebbles within the sediment. A new 1-2 cm-thick layer of coarse sand was added every time the siphons tip reached sediment surface, simulating catastrophic burial by sand as it 1$ known to happen in nature (Stanley, 1970; Berg 8 Alatalo, 1985). This procedure was repeated till the specimens failed to reach the water col- umn. Extrusion of both siphons from the sub- stratum and the ability of the inhalant to take either suspended or deposited material in were observed in - 30 specimens kept buried for eight days in clean, coarse sand in aquaria. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 251 Morphometrics To evaluate if growth data are taxonomically important in distinguishing the Atlantic А. deflorata from Indo-West Pacific A. violascens, shell length, height and width of complete shells from ten different localities in the Indo-West Pacific, and deposited at the Delaware Museum of Natural History - DMNH, USA (п = 23; shell length range (sir): 28.5-74 mm), and of our material (n = 32; sir: 20.6-59.2 mm) were measured to 0.1 mm with dial calipers, and recorded in an Excel spreadsheet. Such growth data were mod- eled using a Model II regression analysis in Systat v. 5.2.1 for Macintosh. To accomplish this, the loss function was changed from the ordinary least squares regression equation to: LOSS = (Y - (BO + В1*Х))^2/АВ$ (B1), where Y and X are the dependent and inde- pendent variables, and ВО and B1 are the two parameters to be estimated. To evaluate if radial ribs are taxonomically important, ribs were counted in the best-pre- served shell valves toward the umbo and at the valve margin of the Atlantic Asaphis (n = 10; shell length range: 35-59.2 mm) (our ma- terial) and the Indo-West Pacific Asaphis (n = 10; sir: 41-56.5 mm) (the Delaware Museum of Natural History - DMNH, USA collection). The degree of branching was recorded as a ratio of margin/umbo ribs, where a larger num- ber indicates more branching. Principal components and cluster analyses (McCune & Mefford, 1999) were used to as- sess the degree of similarity or difference be- tween the Atlantic and the Indo-Pacific Asaphis spp. Six shell characters were included in the analysis: shell length, height, width, number of ribs at margin, number of ribs at umbo, and the ratio of margin/umbo ribs. T-tests were used to assess significance in character dif- ferences. Museum Collections Asaphis violascens. Delaware Museum of Natural History, USA: Solomons Island, lots 185776 (1 v., 2 spec.) and 129978 (1 v., 1 spec.); Fiji Islands, lots 185057 (1 spec.), 211943 (2 v.), 205760 (2 spec.) and 166675 (2 spec.); Japan, lots 185777 (2 spec.), 109642 (1 spec.); Western Australia, lot 175710 (3 spec.); Guam Island, lot 174675 (1 spec.); Philippines, lots 152564 (1 v.), 122836 (1 v.), 122499 (1 v.) and 194390 (1 spec.); Ambon Island, lot 168461 (4 spec.); Palau, lot 211968 (1 v., 1 spec.); Northern Australia, lot 181892 (1 spec.); Malaysia, lot 201332 (1 spec.); Singapore, lot 21545 (2 v.); Moorea Island, lot 152461 (1 v.). Instituto de Biociéncias, Universidade de Sáo Paulo (IBUSP), Brazil: four whole specimens and fourteen single shell valves from Hong Kong, China, not numbered, on which Narchi (1980) based part of his work on A. violascens; material qualitatively ana- lyzed for shell features. The following museum molluscan database were examined in mid-January 2003 to assess the historical and recent distribution of A. deflorata: the Academy of Natural Sciences of Philadelphia (ANSP), National Museum of Natural History (USNM), Florida Museum of Natural History (FLMNH), the University of Miami Rosentiel School of Marine and Atmo- spheric Sciences (RSMAS), and the Florida Marine Research Institute (FMRI). Specimen identifications in the museum col- lections were assumed to be correct and were not verified by the authors; however, speci- men identifications of the RSMAS collection were verified by P. M. Mikkelsen, to whom we are grateful, those of the DMNH collection by E. K. Shea, and of the Instituto de Biociéncias, Universidade de Sáo Paulo by O. Domaneschi. Anatomy Live specimens (shell length range: 6.9-59.2 mm) were dissected and examined for the presence of developed gonads and eggs or sperm using a compound microscope. Speci- mens were recorded as immature when no gonad tissue could be located; mature males and females were identified when sperm or eggs were present. Studies of the anatomical features and draw- ings were made based on living and relaxed and preserved specimens. Magnesium sulfate and refrigeration were used as relaxing agents. Ciliary currents of feeding and cleansing were observed in live specimens using both colloi- dal graphite and carmine powder suspensions, carborundum grade F3 and graded sand par- ticles. Complete serial histological sections (4 to 8 um thick) were taken from a specimen 1.5 cm in shell length fixed in Bouin acetic and stained with Ehrlich's haematoxylin and eosin. 252 DOMANESCHI & SHEA RESULTS Ecology At the “The Horseshoe” site, A. deflorata is restricted to the intertidal zone, in patches of gravelly coarse sand covered with pebbles and rubble on a coral ground. The beach slope of the sampled area is slightly larger than 10° and waters were calm and receded approximately 4 m from the high tide mark during the studied period. Live, sparse unburied specimens were found lying by the high tide mark; buried specimens occurred till a maximum depth of 10-12 cm, the latter determined by the rocky, impenetrable substratum underneath. Crevices and restricted spaces among pebbles within the substratum were usually occupied by individuals. Depths of 5 to 15 cm (Berg & Alatalo, 1985) and deeper (Stanley, 1970) have been registered for the species. Our experiments showed the species can extend their siphons as long as 1.5 times the shell length. This allows us to predict that the largest specimens (- 8 cm in shell length - Stanley, 1970; Berg & Alatalo, 1985) burrow as deep as - 20 cm. In pockets of sand crowded 100 80 60 Number 40 with Asaphis, larger and smaller specimens were intermingled indifferently, occupying the substratum without horizontal or vertical segregation according to size. Their normal life position was with the posterior end up and the longitudinal axis at an angle of between 10° to 30° from the vertical. This angle increased to 90” in some specimens buried shallowly both in crowded or shallow pockets of sand. This supports Stanley's (1970) statements that life position and burial depth of A. deflorata in nature are in part controlled by boundary effects. Along the transect, each station (quadrat) sampled had a different sediment composition, and a correspondingly different population density and structure (Fig.1). Station 1 was composed of rubbles and pebbles and no living specimen was present (n = 0). Station 2 was composed of pebble-covered, gravelly silt; a few (n = 27) living specimens were present, intermingled among a larger number (not recorded) of buried, recently dead shells retained in their life position. Station 3 was composed of gravelly coarse sand with silt; in spite of being dominated by one extremely large piece of coral rubble, it yielded the largest number of specimens (n = 238). Station 4 was Size (mm) оЗаНоп 1 a Station 2 а Station 3 m Station 4 EIG an deflorata. Size distribution of live specimens collected at Stations (St.) 1-4 in July 2002, from the “The Horseshoe” site population, Florida Keys, USA. Total number of specimens by station: St. 1 (n = 0); St. 2 (n = 27); St. 3 (п = 238); St. 4 (п = 137). MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 253 composed of gravelly coarse sand and had a smaller yield (n = 134). Dead shells were scarce in stations 3 and 4. Over 93% of the 399 specimens collected came from stations 3 and 4, the nearest to the high tide mark and regularly exposed during ebb tides. Morphometrics Shell length ranged between 6.9-59.2 mm (n = 399). Almost 70% of the specimens collected were 30-50 mm in shell length; approximately 25% of the population was between 5-30 mm, and only 2.5% was > 50 mm. Based on the measurements of living and > о y = 0.640x + 1.039 В: = 0.997 Measurement (тт) № © о о > о © dead collected specimens, the height and width of А. deflorata grow gradually without obvious interruption or change in the growth trajectory over the size range of 6.9-59.2 mm (Fig. 2A). Overall, shell length of А. deflorata is 1.54 times the height, and 2.26 times the width. Parameters of the growth equations that describe A. violascens (DMNH collection) fall within the 95% confidence intervals for A. deflorata, reflecting the overall similarity in ontogenetic trajectories (Fig. 2B); overall shell ratios are also similar, with the shell length of A. violascens 1.44 times the height and 2.22 times the width. y =0.551x - 1.997 В? = 0.984 Length (mm) > © o a o y = 0.681 + 0.649 В? = 0.998 > o Measurement (mm) № & o © - © © 20 30 40 o height (mm) = width (mm) y = 0.468 - 0.648 В? = 0.994 50 60 70 80 Length (тт) В o height (mm) = width (mm) FIG. 2. Model II regression analysis of height and width vs. length for: A, Asaphis deflorata from “The Horseshoe” site population, Florida Keys, and В, А. violascens at Delaware Museum of Natural History, U.S.A. Although the regression equations are different for each variable, the slope and intercept are within the 95% confidence intervals of each other, and thus the growth trajectories are essentially the same. 254 DOMANESCHI & SHEA TABLE 1. Comparison of observed rib counts (our data - D & S) made on well-pre- served shells of Asaphis deflorata (п = 10; sir: 35-59.2 mm) from the “The Horseshoe” site population, Florida Keys, USA and of A. violascens (п = 10; sir: 41-56.5 mm) in the collection at the Delaware Museum of Natural History, USA, and rib counts pre- dicted for both species by Willan (1993 - И/); п, total number; SD, standard deviation; sir, shell length range; X, average. Asaphis deflorata Asaphis violascens Shell character range (X + SD) range (X + SD) Rib number (W) Rib number — at umbo (D & $) Rib number - at margin (D €. $) 80-102 (92.8 + 7.9) 52-100 (68.4 + 12.9) Rib branching (W) less frequent more frequent Rib branching index (D & S) 1.35 2.58.(1.70=:0.38) 7 125-291 (22515015) 60-90 48-80 (56.3 + 10.9) 40-60 22 60225 +100) Rib counts both at the umbo and at margin of A. deflorata from the “The Horseshoe” site and that is solely composed of А. violascens. lt is also evident that À. violascens generally has of A. violascens (DMNH collection) have a wider range than predicted for both species by Willan (1993), and the ranges overlap (Table 1). In spite of this, principal components analysis (PCA) (Fig. 3) shows that two groups are consistently found: one that contains A. deflorata specimens with a few A. violascens specimens, and one fewer, more branching, ribs than A. deflorata, and these differences are significant at P = 0.05 (t-test: ribs near umbo P = 0.002; ribs at margin P = 0.000; branching ratio P = 0.018). Cluster analysis (Fig. 4) shows that all A. deflorata specimens are clustered in a group that shares < 25% of the information with the majority of A. O A. deflorata A ee % А. violascens 9 Fiji Palau 4 Sol.Is.4 y N. Aus. Malaysia 4 ФЕЙ ФЕ! EN % Japan PCA axis 2(rib number) $01.15. 4 -4 -2 0 2 4 6 PCA axis 1 (size) FIG. 3. First two axes of the principal components analysis account for > 85% of the overall variance. Size (length, width and height) increases from left to right along axis 1. Umbo and margin rib counts decrease from bottom to top along axis 2, whereas the branching ratio increases from bottom to top. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 255 Information Remaining (%) 100 75 50 25 0 AA —— AQ —__—— _—————- —_ _ _———_—o—o—ouoe0 ee ————_——_—————————————— A] = 5 000 00000000000000800000000O Malaysia O A. deflorata O A. violascens FIG. 4. Cluster diagram (Jaccard distance measure, group average linkage) showing the overall similarities between each specimen in length, height, width, number of ribs at umbo, number of ribs at margin, and branching ratio. Branch points at 0 indicate no relationship between variables; branch points at 100% mean the specimens were virtually identical overall. FL, Florida, USA; Sol. Is., Solomons Island; Philip., Philippines; W. Aus., Western Australia; N. Aus., Northern Australia. violascens; however, two anomalous specimens of A. violascens cluster within the A. deflorata branch. Thus, there is no relationship between the two major groups, even though PCA analysis shows that several A. violascens group within that of A. deflorata. Regardless of these outliers, t-tests show that the number of ribs at the umbo and at margin, and the degree of branching distinguish the species and that size measurements do not. Other useful shell characters in distinguishing A. deflorata from A. violascens are: the presence of a discernible, rounded posterior radial ridge and posterior slope in the Atlantic Asaphis (our material), not discernible (Willan, 1993; IBUSP collection) in the Indo-West Pacific specimens; a smooth inner surface in the Atlantic Asaphis, contrasting with the well-marked, ridged inner surface in specimens from Hong Kong (IBUSP collection). Sexual Maturity In the sampled population, all specimens < 24 mm in shell length were immature, and all specimens > 32 mm had recognizable eggs or sperm in the gonad tissue. The gonads of most specimens > 24 mm in shell length were identifiable as male or female; just two specimens of 26 mm and 34 mm in shell length were not. Distribution Asaphis deflorata has historically been collected along the Atlantic and Gulf coasts of Florida, as far north as Saint Augustine Beach (FLMNH 16923), and as far south as the Dry Tortugas (FLMNH 16919) as well as in the Bahamas. Rios (1994) registered the species for Atol das Rocas, off northeast Brazil. Most of these records have collection dates between 1878 and 1966. None of the collections assessed had specimens of A. deflorata collected after 1975, although extant populations exist at other Caribbean sites including the Bahamas (FLMNH 247323), Cuba (ANSP 192408), and Trinidad & Tobago (FLMNH 226465). Functional Anatomy Shell: The shell of A. deflorata from the “The Horseshoe” site population (Fig. 5), matches the general shell characterization described 256 DOMANESCHI & SHEA by Abbott (1974) and Rios (1994) for the species in Atlantic waters of the Caribbean region, Bermuda and off northeast Brazil. Shell oval-elongate, equivalve, moderately inflated; umbos subcentral anterior. Maximum shell height at the umbonal-ventral axis; maximum width at the level of umbos. Posterior margin truncate; anterior margin broadly rounded; ventral margin straight to slightly convex; shell ends gaping slightly. Shell with rounded posterior ridges and discernible posterior slope marked by stronger radial ribs; posterior ridges more conspicuous at the dorsal half of the shell, running diagonally to and fading as it meets the acute- rounded confluence of both posterior and ventral shell margins. Outer surface sculptured with numerous fine, radial ribs ranging in number from 80-102 (n = 10; shell length range: 35-59.2 mm) at shell margin (Table 1). Few ribs with a weak tendency to fork; radial ribs stronger, wider apart and scaly to slightly nodulose posteriorly. Fine commarginal ridges smoothly crossing the radial elements and FIG. 5. Asaphis deflorata. External view of the left shell valves of specimens from the “The Horseshoe” abs population, Florida Keys, USA, showing radial ornamentation and little variation in shell outline. cale bar = 5 ст. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS expanding into scale-like or nodulose processes on the posterior radial ribs. Exterior dull; yellowish to creamish white predominate in the population. Periostracum thin, dehiscent. Interior often glossy, brightly colored; radial ornamentation on the outer surface not interfering on the inner surface, which 1$ smooth throughout. A deep violet blotch often present, spreading over from the nymph through the posterior margin; a similar, smaller, often fading blotch can be present at the level of the anterior adductor muscle scar. Anterior adductor muscle scar elliptical, elongate dorso-ventrally (Fig. 6); posterior one elliptical to rounded; anterior and posterior retractors pedal muscle scars fused dorsally to the corresponding adductor scars. Pallial line recessed deeply from within the smooth shell margin. Cruciform muscle scars faint to invisible. Pallial sinus broad, extending almost to the level with the posterior cardinal tooth; sri 297 upper limb straight to slightly curved; anterior margin rounded; lower limb detached from pallial line, descending obliquely to coalesce with the latter far from the rear end of the pallial sinus. Extension of the fusion lower limb-pallial line corresponding to one third of the pallial sinus depth; rear end of this extension reaching level with the anterior half of the posterior adductor scar. Striking impression of the fan- shaped siphonal retractor muscle present within the pallial sinus. Hinge plate with two cardinal teeth in each valve (Fig. 7); nymph broad; ligament elongate, thick, tough. Right anterior cardinal tooth conspicuous, emerging from the hinge plate as a knob-like or thick, plate-like projection; right posterior cardinal stronger, elongate, deeply bifid and separated from the anterior tooth by a deep triangular socket. Left anterior cardinal tooth strong, elongated and deeply bifid; left posterior cardinal emerging as a knob-like or thick, plate-like projection. pet | VA cms FIG. 6. Asaphis deflorata. Internal view of the right shell valve. Abbreviations: aas, anterior adductor muscle scar; act, anterior cardinal tooth; ars, anterior pedal retractor muscle scar; cms, cruciform muscle scars; |, ligament; п, nymph; pas, posterior adductor muscle scar; pct, posterior cardinal tooth; pl, pallial line; prs, posterior pedal retractor muscle scar; ps, pallial sinus; sri, siphonal retractor muscle impression; u, umbo. Scale bar = 1 cm. 258 DOMANESCHI & SHEA Mantle: The mantle lobes are thin and trans- lucent with the usual three folds along their free ventral edges. Asaphis deflorata lacks the ad- ditional folds that isolate the rejection channel for pseudofaeces (waste canal of Kellogg, 1915), which is present in some Tellinidae and Semelidae (Yonge, 1949), and in species of Mesodesmatidae and Veneridae (Narchi, 1981, 2002). Graham (1934a) and Domaneschi (1992) have given a detailed picture and de- scription of the mantle edges in species of Gari. The mantle edges in A. deflorata are histologi- cally similar to those described and illustrated in detail by Domaneschi (1992) for Gari solida (Gray, 1828). The outer fold is the least developed; the inner fold is moderately higher and the only to be involved in siphon formation. The middle is a huge, sensory fold, which bears a single row of cylindrical, cup-shaped tentacles and the best supplied with pallial muscles. When the foot and siphons are protracted, the middle folds are extended well beyond the limits of the shell valves, exhibiting an outer surface lined by a thin, translucent periostracum. The periostracal groove lies outside, adjacent and parallel to row of the tentacles. Similar organi- zation has been noted in the other Tellinoidea in the semelid Ervilia castanea (Montagu, 1803) by Morton (1990) and in other Psammobiidae — in Gari solida by Domaneschi (1992) and in Heterodonax bimaculatus (Linné, 1758) by Narchi & Domaneschi (1993). Each bundle of fibers that comprise the pal- lial retractor musculature splits into two sets immediately after they originate at the pallial line. Both sets supply mainly the middle fold and respective tentacles, and meet again ad- jacent to the periostracal groove, as in Gari tellinella (Lamarck, 1818) (Graham, 1934b) and С solida (Domaneschi, 1992). A few fi- bers that supply the outer mantle fold arise from the set running adjacent to the outer mantle epithelium. Muscle fibers that retract the inner mantle fold arise from the inner set underlying the inner mantle epithelium. In addition to these pallial retractors, there are bundles of fibers running transversely across the mantle edge and folds, and bundles of longitudinal fibers restricted to the inner and middle folds, with the bulk of them along the inner face of middle fold. The pallial nerve cord lies embedded in the connective tissue contained by the two sets of pallial retractors. Mucous-gland cells form a well-defined, glandular region along the pedal opening, just dorsal to the base of the inner mantle fold. Mucus-secreting cells are more abundant on both ends of the pedal opening. FIG. 7. Asaphis deflorata. Hinge plate morphology of the right (bottom) and left (top) shell valves. Abbreviations: act, anterior cardinal tooth; |, ligament; n, nymph; pct, posterior cardinal tooth. Scale bar = 2 mm. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 259 FIG. 8. Asaphis deflorata. The animal viewed from the left side after removal of the left shell valve and mantle lobe. The siphons and foot are shown somewhat contracted. Arrows show the direction of the ciliary currents. Abbreviations: aam, anterior adductor muscle; cm, cruciform muscle; dd, digestive diverticula; dh, dorsal hood; ex, exhalant siphon; f, foot; id, inner demibranch; ilp, inner labial palp; in, inhalant siphon; lg, lateral oral groove; mi, mantle isthmus; od, outer demibranch; olp, outer labial palp; pam, posterior adductor muscle; pm, pallial retractor muscles; pr, pericardial region; prm, posterior pedal retractor muscle; so, sense organ. Scale bar = 1 cm. The cruciform muscle with its specialized sensory organs occurs postero-ventrally, be- tween the base of the inhalant siphon and the pedal gape (Fig. 8). Both structures are diag- nostic for the Tellinoidea (Ihering, 1900; Frenkiel, 1979; Morton, 1984, 1990; Morton & Scott, 1990). The sensory organs, which lie at the posterior arms of the cross open directly to the siphonal space at the summit of minute papillae as described by Graham (1934a) and Domaneschi (1992) for Gari, and Domaneschi (1995) for Semele. The cruciform muscle and its sensory organs have been described by Mouéza 8 Frenkiel (1974, 1976, 1977), Frenkiel 8 Mouéza (1977, 1984), Frenkiel (1979), Morton (1990). Cilia are present all over the inner epithe- lium of the mantle lobes. They are more con- centrated along a well-defined, wide ciliated gutter that lies parallel to the origin of the pal- lial retractor muscles. Cleansing ciliary cur- rents (Fig. 9) sweep particles forward and downward from the dorsal area adjacent to the posterior adductor muscle to a vigorous C- shaped rejection tract. This rejection tract ex- actly follows the line of origin of the siphonal retractor muscle. Weak cleansing currents within the limits of the C-shaped tract, and on the dorsal area adjacent to the anterior ad- ductor muscle convey particles downward and backward to the base of the inhalant siphon. Minute, isolated particles coming into contact with inner mantle fold are carried slowly up- ward to join those coming downward on the mantle surface. Despite the presence of the ciliated gutter, A. deflorata lacks a well-defined, vigorous, rejection current running backward parallel to the mantle edge. Such a vigorous current 1$ present in A. violascens, Неегодопах bimaculatus, and Сап solida (Psammobiidae) (Narchi, 1980; Narchi & Domaneschi, 1993; Domaneschi, 1992, respectively). Siphons: The siphons, type Aof Yonge (1957, 1982), are wide, separate throughout their extent (Fig. 8) and up to 1.5 times the shell length. The inhalant is slightly longer, as in other Psammobiidae (Yonge, 1949; Domaneschi, 1992; Narchi & Domaneschi, 1993). In a week's time in the laboratory, only the inhalant siphon was extruded clear of the sediment surface, but not as far as the 3-5 cm 260 DOMANESCHI & SHEA FIG. 9. Asaphis deflorata. Ciliary cleansing currents on the inner surface of the right mantle lobe. Scale bar = 1 cm. as reported by Berg & Alatalo (1985). When extruded and undisturbed, the inhalant was held passively for many hours, with the ring of tentacles either straight or curled inward; re- tracted, its aperture either flushed with or was kept below the sediment surface. This latter behavior 1$ the rule for the exhalant tip. The inhalant was never seen bending down onto or along the bottom sediment sucking in de- posited material as in typical deposit-feeding tellinoideans described by Yonge (1949). Many specimens buried within the sediment in aquaria had the tip of the inhalant siphon com- pletely covered by a 1 cm-thick layer of coarse sand, but maintained an active current flow- ing through the interstices of the sand grains, as confirmed by pouring carmine powder suspension into the water. Such an ability al- lows the species to survive buried within either rock-, or cobble-covered gravelly sediment without extruding the siphons into the water column. Berg & Alatalo (1985) stated that they have never observed the tip of the inhalant exposed into the water column in the field. The inhalant aperture is fringed with twelve simple, finger-like tentacles, six longer alter- nating regularly with six shorter. From the dis- tal end of each longer tentacles arises a double row of minute, cylindrical, cup-shaped papil- lae which extend down the length of the si- phon. The exhalant aperture is fringed with eight longer tentacles interspersed with eight shorter tentacles. Single rows of minute papil- lae extend throughout the length of the exhal- ant siphon, each row associated with longer tentacles. Brilliant, golden-yellow pigmentation is present in the inner wall of both siphons. Pig- ments may simply create a speckled pattern, or be grouped into rounded-elongate, regularly arranged spots, or a mixture of both. The larger concentration of pigments in the tips of the si- phons gives them a vivid, golden-yellow color, which fades away toward the bases. Narchi (1980) and Domaneschi (1992) also noted a yellow color for the siphons of A. violascens and Сап solida, respectively. The siphons of A. deflorata are sensitive to touch. Although light sensitive, they exhibit a similar response either to high or low luminosity coming from an electronic flash, or a microscope Шитта- tor, respectively. Labial Palps: The labial palps (Fig. 8) are approximately one half the shell length. When completely expanded, their free distal tips do reach and even surround the posterior border of the visceral mass. The ventral half of the inner surfaces of the palps are obliquely folded and separated from the dorsal, smooth half by a longitudinal fleshy cord that overhangs slightly the dorsal extremi- ties of the folds. The inner demibranchs of the ctenidia project deeply between the palps, but the ventral tips of their anteriormost filaments are not inserted into a distal oral groove. Thus, the labial palps-ctenidial junction is of Category Ш (Stasek, 1963). Exceedingly large palps, provided with dif- ferent ciliary tracts indicate that A. deflorata processes large amounts of particles in the mantle cavity. The palps play an important selective function, even though the wide ctenidia exert a previous selection of the bulk of material entering the mantle cavity as in other suspension feeding tellinoideans. The sorting mechanisms of the palps are shown in Figures 8 and 10. Particles coming into contact with the smooth outer face of the palps (Fig. 10A) are carried dorsalward (current “a”) and then passed to the internal, smooth, dorsal half of the organs. Here, transverse, ventrally directed currents transfer them onto the folded area (Fig. 10B) to be selected. Transversely directed currents (b), operat- ing obliquely oralward and markedly ventralward across the crests of the folds, to either accept or reject particles, depending upon the size or total volume of particles. On the aboral face of the folds a current (c) carries isolated particles and small agglom- erations of particles dorsally. As they move dorsally, they are influenced by transverse currents “b” and removed anteriorly. Only minute, isolated particles are caught by cilia on the adoral faces of the folds and are MORPHOLOGY AND SHELL MORPHOMETRY OF ASAPHIS 261 carried dorsalward. Traveling on this current “d”, particles reach the dorsal extremity of the folds, where a conspicuous oralward current (e) transports them onto the lateral oral groove, between the palps. Particles escaping from the action of the pre- vious currents “b”, “с”, “4”, and reaching the floor of the grooves between adjacent folds, are driven ventrally (current “f”) onto a rejec- tion current (g) present along the narrow, smooth ventral edges of the palps. Excess material on current “b” also converges onto this current “д”. Currents “с” and “а” function as resorting devices; in combination with current “b”, they keep the food material away from rejection tracts, and allow the agglomerations to be disintegrated, resorted, and useful material in- gested prior to being discarded as pseudo- faeces. Muscular contractions of the palps increase their sorting efficiency in processing different amounts of particles drawn into the mantle cavity through the inhalant current. By bend- ing laterally, the inner palps touch the visceral mass epithelium where particles are being carried ventrally. Once trapped by currents “a”, such particles are passed to the folded sur- a face of the organ and sorted. The outer labial palps do the same, touching the mantle epi- thelium. Muscular activity is also responsible either for bringing folds closer or forcing them apart, and in keeping them erect or bending them over. Such devices permit total exposure of the ciliary tracts, favoring resorting and in- gestion of scarce profitable food, as well as their concealment, which favors rejection when the inhalant flow of water contains excess material. Foot and Visceral Mass: The foot is a huge, axe-shaped muscular organ that expands far beyond the anteroventral margin of the shell. At the posterior end of its narrow, ventral edge is a distinguishable slit-like opening, related to a shallow depression (duct), both remnant of the byssal complex, as identified by Pelseneer (1911), Graham (1934b), and Domaneschi (1992) for Gari spp. No trace of byssus gland was detected within the foot of A. deflorata. Pelseneer (1911) and Narchi & Domaneschi (1993) did not find even a byssal groove aper- ture in Asaphis violascens and Heterodonax bimaculatus, respectively. Narchi (1980) made no reference to byssal complex in A. violascens. Every time it was provided a soft substratum, the foot in all laboratory specimens aboral FIG. 10. Asaphis deflorata. A, sketch of the left inner labial palp showing the ciliary currents (a), (e) and (g). B, diagrammatic representation of three folds and respective ciliary currents (b), (c), (d) and (f). For lettering (a) through (g), see text on labial palps. 262 DOMANESCHI & SHEA of А. deflorata (shell length range: 6.9 to 57 mm) was used exclusively for burrowing, this being slower in larger individuals. Burrowing period (Stanley, 1970) lasted from 2 to 5 hours among individuals < 2.5 cm in shell length (n = 5), and 23 hours and 35 hours in individuals 4.5 and 5.7 cm in shell length, respectively. The upper, visceral portion of the foot is cov- ered with a low, smooth epithelium. Ciliary currents were detected in all dissected speci- mens, even though only sparse patches of cilia could be seen in histological sections. Intense cleansing ciliary currents on this visceral por- tion of the foot sweep and concentrate par- ticles downward over a narrow longitudinal area, juxtaposed with the outer dorsal mar- gins of the inner labial palps and the free edges of the inner demibranchs. Only a weak dorsalward current was observed on a nar- row area juxtaposed with the very proximal portion of the inner labial palps. Particles trav- eling on this current are caught by cilia on the outer, smooth face of the palps and passed to the opposite face of the organs (Fig. 10A). The most ventral, predominantly muscular portion of the foot is obliquely ridged and lined with a densely ciliated epithelium (cilia 5.7 um long). Despite being intensively ciliated, no ciliary currents could be detected on this por- tion of the foot; Domaneschi (1982) stated they are absent in Gari solida. Only Pohlo (1972: fig.1) has depicted dorsalward ciliary currents on the most ventral portion of the foot of a psammobiid. The portion of the foot juxtaposed mainly with the labial palps has a 28 um-depth, longitudi- nally striated epithelium. Lack of cilia on this portion was confirmed through histological sections and by observing live specimens. Ctenidia: The ctenidia are large and occupy most of the mantle cavity when completely expanded (Fig. 8). They are eulamellibranch, plicate and heterorhabidic. The morphology of both demibranchs is much the same, the outer being more shallowly plicate. The ascending lamellae of both inner demibranchs are attached to the epithelium of the visceral region of the foot by ciliary junc- tions; behind the foot they connect to each side of a thin, wide triangular membrane the base of which surrounds and attaches to the foot by ciliary junctions. The ascending lamellae of both outer demibranchs connect to the vis- ceral mass epithelium by tissue fusion. Histological sections prepared from a speci- men 1.5 cm in shell length revealed that the deepest plicae (400 um depth) of the inner demibranch are formed by 35 ordinary fila- ments. Each filament bears 4.3 um-long, fron- tal cilia, these bordered latero-frontally by 17.2 um-long, latero-frontal cilia. The lateral cilia responsible for the inhalant current of water are roughly 11.5 um long. Terminal cilia (23 um long) form a fringe along the lateral walls of the marginal food groove. This is 34.2 um depth on average and evenly carpeted with 5.7 um-long cilia. The frontal surface of a principal filament varies in form throughout its extent; it is ridged near the free ventral margin of both demibranchs, and changes to a broad, shal- low gutter toward the ctenidial axis. The more stretched a particular region of the principal filament is, its frontal gutter is more flattened and broad, the sides of the filament frequently sloping away nearly at the same plane as that of the groove. In strongly con- tracted plicae, the frontal gutter of the princi- pal filament changes to a narrow, deep central groove. This is flanked by the sides of the fila- ment, which lie at the same plane of that of the frontal surface, as described by Ridewood (1903) for Gari vespertina (Gmelin, 1791). The morphology and respective ciliary mechanisms of the ctenidium (Fig. 11) are of type C (1a) (Atkins, 1937), characteristic of a variety of eulamellibranchs, including the Tellinoidea, as in the Psammobiidae analyzed by Atkins (1937) and Narchi & Domaneschi (1993). Acceptance oralward currents are thus restricted both to the marginal food groove of the inner demibranch and to the ctenidial axis. Frontal ciliary currents on both ordinary and principal filaments of the outer demibranch are exclusively ventralward on the ascending lamella, and round the bend at the free edge, and dorsalward on the descending lamella. Frontal currents are exclusively ventralward on both lamellae of the inner demibranch. There is a tendency for large particles or masses of particles that reach the free edge of the outer demibranch to be passed straight off onto the inner demibranch. However, an incipient oralward current was also registered along the free ventral margin of the outer demibranch. Particles on this current are car- ried for short distances anteriorly, then devi- ate ventralward under the influence of the frontal currents on the inner demibranch. Another incipient oralward current does ex- ist along and outside the marginal food groove of the inner demibranch. Material traveling on this current usually falls off on the rejections currents of the mantle epithelium. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 263 FIG. 11. Asaphis deflorata. Diagrammatic trans- verse section showing the form of the ctenidium and directions of the frontal ciliary currents (ar- rows). Solid circles, oralward currents; hollow circles, incipient oralward currents. Adductor Muscles: The anterior adductor muscle (aam) is elliptical and dorsoventrally elongate; the posterior adductor (рат) is subelliptical to rounded and thicker than the anterior (Fig. 12). Pedal Musculature: The extrinsic pedal mus- culature (Fig. 12) consists of bilateral pairs of almost equally developed anterior and poste- rior pedal retractors, one pair of anterior pedal protractor, and one pair of vestigial pedal el- evator muscles. On each side of the roof of the visceral mass, a thin layer of muscle fibers (el) converge dor- sally and insert deep on the umbonal cavity, where they leave a single, well-impressed scar, or more than a single scar. Such a mus- cular layer is functionally not significant com- pared to the other extrinsic pedal muscles; it corresponds by its insertion on the shell valve to the functional elevator pedal muscle in many other bivalves. In A. deflorata, the elevator pedal muscles are atrophied as in other Tellinoidea, for example, Gari spp. (Pelseneer, 1911; Bloomer, 1911; Domaneschi, 1992), Tellina foliacea (Linné, 1758) (Pelseneer, 1911), Semele spp. (Domaneschi, 1982, 1995). Narchi (1980) and Narchi & Domaneschi (1993) did not find pedal elevators muscles in A. violascens and Heterodonax bimaculatus, respectively. The anterior and posterior pairs of pedal re- tractors are equally developed, each attached under the hinge plate internal and contiguous to the adductor, where their edges and respec- tive scars coalesce. The right and the left posterior pedal retractors (prm) pass antero- ventrally, converging and meeting in the sag- ittal plane, almost completely enveloped in the kidneys. Where these muscles meet, their most internal bundle of fibers intersect; the bundles coming from the right pass deeply into the left side of the foot and vice-versa. The most external bundles of each posterior re- tractor pass directly into the foot. As a whole, the posterior retractors form the innermost muscular layers within the foot. The anterior pedal retractors (arm) also con- verge on the sagittal plane, where a few inter- nal bundles of fibers intersect each other and pass deep into the opposite side of the foot. The majority of bundles from each muscle pass directly into the foot, on its corresponding side. Within the foot the anterior retractors form an outer layer in relation to the posterior pedal retractors. The outermost muscular layer of the foot is composed of fibers coming from a pair of an- terior pedal protractor muscles (ppm). Differ- ently from other psammobiids, on each side of A. deflorata the anterior protractor is com- posed of two separate sets (branches) of bundles: one ventral, slender set attaches to the shell juxtaposed to and slightly inserted in the postero-dorsal surface of the anterior ad- ductor; the other set, dorsally placed, gathers the bulk of the fibers. Fibers of the dorsal set insert on the shell valve embracing the ven- tral half of the origin of the anterior retractor. From its origin on the shell valve, the slen- der, ventral set of the protractor muscle passes horizontally into the foot, where the bulk of its fibers go backward, while the remainder twist abruptly, most downward and a few upward, spreading like a fan. Since the bulk of its fi- bers lies almost to the level with the base of the labial palps, this ventral branch of the pro- 264 DOMANESCHI & SHEA tractor may easily be misjudged as related to the palps. However, careful dissections re- vealed the protractor fibers penetrating exclu- sively into the visceral region of the foot. The dorsal set of the protractor has the bulk of its fibers extending almost horizontally and running backward parallel to the longitudinal musculature of the ctenidial axis. Such a juxta- position makes the identification of both muscles almost impossible. The dorsal set has a large number of its fibers spreading fanwise downward, where they mask those coming from the ventral set. The remaining fibers of the dor- sal set spread fanwise dorsally and intermingle with those coming from the vestigial, “elevator” pedal muscle. Narchi (1980) stated that pro- tractor muscles are lacking in A. violascens. Apart from the extrinsic pedal muscles, the visceral mass and the most ventral parts of the foot contain a large number of isolated, transverse bundle of fibers (intrinsic muscles of Bloomer (1911)). The bundles crossing the aam 7 | visceral mass are particularly numerous, thick and long in A. deflorata. Such transverse bundles play an important role in moving blood, and moving materials within the organs. Ctenidial Retractor and Longitudinal, Ctenidial Axis Muscles: The ctenidial retrac- tors (cr) are a pair of thin, but conspicuous muscles, with an origin slightly posterior to the insertion of each vestigial “elevator” pedal muscle on the umbonal cavity. The ctenidial retractor fibers pass downward, meet and in- termingle with those muscular fibers running longitudinally throughout the ctenidial axis (cam). The combined action of both, ctenidial retractor and longitudinal, ctenidial axis muscles shortens the ctenidia as a whole and lifts especially the inner demibranchs. Bloomer (1907), Villarroel & Stuardo (1977), and Domaneschi (1982, 1995) described and de- pict a similar muscle in Tagelus divisus (Spengler, 1794), T. dombeii (Lamarck, 1818) and Semele spp., respectively. ar cam FIG. 12. Asaphis deflorata. Musculature, as seen from the left side. Abbreviations: aam, anterior adductor muscle; arm, anterior pedal retractor muscle; cam, longitudinal, ctenidial axis muscle; cr, ctenidial retractor muscle; el, “elevator muscle”; pam, posterior adductor muscle; ppm, ventral and dorsal sets of the pedal protactor muscles; prm, ventral and dorsal sets of the posterior pedal retractor muscle. Scale bar = 1 cm. MORPHOLOGY AND SHELL MORPHOMETRY ОЕ ASAPHIS 265 Alimentary Canal: The stomach and style sac in А. deflorata follow the general psammobiid pattern described by Purchon (1960), Narchi (1980), Domaneschi (1992), and Narchi & Domaneschi (1993), whereas the intestine parallels only that of А. violascens, in which the hind gut dilates to store faeces (Purchon, 1960; Narchi, 1980). The general configuration of the alimentary canal of A. deflorata is shown in Figures 13 and 14. The main difference distinguishing it from A. violascens lies in the hind gut, as shown in Table 2. The measurements provided below were taken from a transversely sec- tioned, 1.5 cm-long specimen; dimensions and total numbers of faecal pellets were taken from a 5.2 cm-long specimen. Essentially the intestine comprises two coiled sections. The first section (mg) lies posterior and adjacent to the distal half of the style sac and is separated from this by gonad follicles, digestive diverticula, and a number of cross strands of muscle fibers. lts external diameter is almost uniform and very reduced (500 um in average), with a 36 um-thick wall, the 25 um-tall columnar epithelium richly provided with 16 um-long cilia. A feeble layer of circu- lar, muscular fibers is present around the mid gut walls. The cilia cause rotation and reloca- tion of material coming from the stomach and wrap it in a viscous mass. Muscular fibers may be responsible for peristalsis that contributes to compacting, molding and relocating dis- carded material. This way faecal pellets are completely formed within the mid gut. Faecal pellets are similar in appearance to the con- tents within the appendix, described below in the stomach section; however, they gather a larger number of inorganic particles that include whitish, hard corpuscles and small quantity of non-identified organic debris and microorgan- isms. Most sponge spicules and diatom frag- ments were not affected when the contents of the appendix of the stomach, and faecal pel- lets were submitted to weak acid solution (HCl); however, the whitish hard fragments dissolved FIG. 13. Asaphis deflorata. Configuration of the alimentary canal and part of the excretory and circulatory systems, as seen from the left side. Abbreviations: ab, aortic bulb; ax, appendix; dh, dorsal hood; f, foot; hg, hind gut; к, kidney; Ic, left caecum; Ip, left pouch; mg, mid gut; mo, mouth; o, esophagus; pc, pericardial cavity; r, rectum; ss, style-sac; st, stomach; v, ventricle. Scale bar = 1 cm. 266 DOMANESCHI & SHEA completely, revealing their calcareous compo- sition. Midway to the stomach (st), the mid gut por- tion ends and the intestine dilates to form its second coiled section, the hind gut (hg), dor- sally placed. This second section is more ex- tensive, more expanded and more intricately coiled throughout its extension (Fig. 13); its walls comprise а 5.5 um-thick epithelium sur- rounded by an either equal or thinner layer of circular, muscular fibers, which allow widen- ing/narrowing of the hind gut diameter. Cilia could not be detected in histological sections of the hind gut. The section walls are so thin and so closely applied together that the ex- amination and tracking of the hind gut course was extremely difficult. Polarized light allowed confirmation of the presence of muscular fi- bers surrounding the alimentary canal. The hind gut accumulates small, very regu- lar, rod-shaped faecal pellets. From its proxi- mal end (Fig. 13) the hind gut extends dorsal- and backward to the floor of the pericardial cavity (pc). From here it passes downward to the right side of the animal (Fig. 14), spirals both in a tight and clockwise way, increasing to its maximum width of 6.6 mm, that is, about 14 times wider than the mid gut (Fig. 13). This enormous swelling extends ventrally, where it reduces in diameter and turns abruptly both forward and dorsalward (Fig. 14) passing to the posterior right side of the appendix (ax) of the stomach. Here it returns to a narrow ex- ternal diameter (~ 250 um, empty condition; lining epithelium deeply folded), penetrates the pericardium (pc) and terminates in the anal papillae on the posterodorsal face of the pos- terior adductor muscle. The epithelial cells lin- ing this very rear portion of the intestine are 8-10 um in height and densely ciliated (cilia 8.3 um long). Similar to the esophagus, this portion has a deep folded epithelium, sur- rounded by a thick fibrous-like layer, which includes circular muscular fibers; the muscu- lar layer is thicker around the esophagus. Approximately 650 faecal pellets, with a most frequent length of 1.5 mm (range: 0.5-2.5 mm) were recovered from a 5.2 cm-long specimen. FIG. 14. Asaphis deflorata. Configuration of the alimentary canal and part of the excretory and circulatory systems, as seen from the right side. Abbreviations: rc, right caecum. For other lettering, see Fig. 13. Scale bar = 1 cm. MORPHOLOGY AND SHELL MORPHOMETRY OF ASAPHIS 267 FIG. 15. Asaphis deflorata. А. Interior of the stomach after being opened by a longitudinal incision in the dorsal wall. В, С. Internal anatomies of the right and left caeca, respectively. С. anatomical differ- ences found in two different specimens. Abbreviations: ax, appendix; bp, blind pocket; cs, crystalline style; dh, dorsal hood; e, semi-circular elevation on floor of the stomach; el, long forwardly projecting elevation; fp, fleshy pads within the appendix; gs, gastric shield; ig, intestinal groove; Ic, left caecum; Ip, left pouch; mg, mid gut opening; mt, minor typhlosole; о, esophagus; г, broad fold passing from the anterior floor of the stomach to the interior of the dorsal hood; r1, ridge passing from the posteroventral blind pocket to the interior of the dorsal hood; rc, right caecum; rm, rim to the esophageal orifice; rt, rejection tract; s, swelling on the left anterior wall of the stomach; sa, sa3, заб, sa7, sa8, sa10, sa11, sorting areas; ty, major typhlosole. 268 DOMANESCHI & SHEA Arranged end to end, all these pellets would perform a beaded, 98 cm-long thread (650 x 1.5 mm). Stomach: The morphology and functioning of the stomach of А. deflorata (Fig. 15) are so similar to those described in details by Purchon (1960) and Narchi (1980) for А. violascens that a complete description of the stomach of the former species is here assumed to be unes- sential. Table 2 shows the main differences between the organs in both species. Apart from those differences, the following aspects of the morphology and functioning of the stomach of A. deflorata deserve mention: - The semi-circular elevation (e) [= shortest branch of the major typhlosole (ty)] borders a shallow, supporting gutter for the rotating crystalline style (cs); the gastric shield (gs) protects this gutter against abrasive materi- als adhering to the style. - Entering the right caecum (rc), the longest branch of the major typhlosole sends flares into the openings of five ducts coming from the digestive diverticula; entering the left caecum (Ic), it sends flares into the mouth of the largest of such ducts only. The left caecum does not receive a constant number of ducts coming from the digestive diverticula in different specimens (Fig. 15C). Freshly incised stomachs show that the fold ‘r” prolongs backward the gutter formed by the floor of the esophagus. The free edges of this fold touch both the crystalline style and the semi-circular branch of the major typhlosole, posteriorly, and a low, broad swelling ($) on the anterior left wall of the stomach. The fold *r” isolates ventrally the entrances of both right and left caeca, the entrance of the left pouch and the transverse section of the intestinal groove, as observed by Purchon (1960) in A. violascens. So stra- tegically positioned, the fold *r” favors mate- rial entering the esophagus to be caught by the rotating style and mixed with the disinte- grating, gelatinous tip of the latter. Thence, material is passed into the dorsal hood (dh), TABLE 2. Analysis of morphological variation in the alimentary canal of Asaphis deflorata [our data] and A. violascens [Purchon’s (1960 P) and Narchi's (1980 N) data]; г, fold; sa, sorting area on the inner wall of the stomach. Hind gut Stomach “Гл” Sa “sa8” “sa11” Appendix Left pouch Blind pouch A. deflorata widens progressively as it coils and spirals in a tight way throughout its extension; turns to a narrow diameter only within the heart present; passes deep into the dorsal hood present; lying posterior to and throughout the extension of the long sorting area “sa3” present; extending from below the esophagus opening deep into the left pouch present; at the stomach roof and anterior wall of the dorsal hood present; a single sorting area outward ciliary currents present on its fleshy pads sorting areas present (sa6, sa7) present; a wide cone-shaped depression on the stomach floor A. violascens (P) (№) extraordinarily ballooned in its proximal end, where mid gut enters dorsally and a narrow, non-coiled hind gut leaves ventrally (P) (№) present; does not enter the dorsal hood (P) described and depicted a similar fold shorter than “sa3”; didn't name it (N) just depicted it throughout the extension of “sa3” (P) (N) present; just below the esophagus opening (P) (№) neither described nor depicted it (P) (N) present; two sorting areas (P) (N) no outward ciliary currents are present Sorting areas present [заб (P) (N)]; two others unnamed (P) (P) (N) did not refer to it MORPHOLOGY AND SHELL MORPHOMETRY OF ASAPHIS 269 following storage of excess material in the appendix (ax). Another important role of the fold “r” is to prevent material entering from the esophagus to be caught earlier by cilia on the intestinal groove. - The full packed appendix equals the volume of the main cavity of the stomach. A thick, gelatinous mass constitutes the bulk of ma- terial often present within this pocket. Lots of amorphous greenish-brown debris and minute, white, mineral corpuscles, many sponge microspicules and broken megascleres, some algae debris, diatoms, a few foraminiferans and unidentified micro- organisms and eggs were the most frequent material entangled in a such viscous mass. Outward ciliary currents are present on the two fleshy pads (fp) that protects the appen- dix entrance and neck respectively. Such currents are not vigorous enough to deal with (remove) the bulk of material and the vis- cous mass stored in the appendix. Contrac- tions of the appendix walls, as well as of the closest intrinsic, tansverse musculature of the visceral portion of the foot probably play an important role in such emptying process. Purchon (1960) stated that the contents of the appendix of A. violascens are possibly discharged into the stomach by muscular contraction of its walls. - The sorting areas sa10 and sa11 are !-ае- fined and the least conspicuous within the stomach. The same is true for sa7, in its portion below the esophagus only. Only care- ful analyses of several live and preserved specimens allowed confirmation of their presence in the stomach of A. deflorata. Organs of the Pericardium: The heart lies at the level of the shell ligament (Figs. 13, 14). This organ comprises a ventricle (v) pen- etrated by the rear end of the hind gut, and a pair of auricles. The posterior aorta dilates just after its emergence from the ventricle to form the aortic bulb (ab), the latter as long as the ventricle (0.1 of the shell length). From the pericardium arise a pair of reno-pericardial apertures which drain primary urine into a pair of kidneys (k) located between the posterior retractor pedal muscles and the floor of the pericardium. The kidneys open into the supra- branchial chamber at the summit of minute papillae, between the ctenidial axis and the line of attachment of the ascending lamella of the inner demibranch to the visceral mass. Close and ventral to the renal apertures are the slit-like gonopores. DISCUSSION In considering the Atlantic and the Pacific specimens of Asaphis to be conspecific, Abbott (1950) pointed out that, when or if differences can be demonstrated between them, it would be wise perhaps to retain the name A. deflorata Linné, 1758, for the western Atlantic speci- mens and apply A. violascens (Forsskál, 1775) to the Pacific ones. Willan (1993) considered that the Atlantic and Pacific Asaphis share a common ancestor from which divergence oc- curred relatively recently. Lacking anatomical studies on the Atlantic specimens has re- stricted separation between them exclusively on the basis of shell sculpture. The analysis of Asaphis from the “The Horse- shoe” site population, West Summerland Key, Florida, USA, has shown that, in addition to shell sculpture, as predicted by Prashad (1932) and Willan (1993), the western Atlan- tic Asaphis also has ecological and morpho- functional characters that distinguish it from its Indo-West Pacific, close relative. Such a population at the “The Horseshoe” site is restrict to the intertidal region, where it lives buried at moderate depths (0-12 cm) and densely aggregated in gravelly sand, cobble covered environments in the high intertidal zone. The species constitutes the sole bivalve present in the upper shore; no specimen was found subtidally. These characteristics are consistent with previous reports of this Atlan- tic species (Stanley, 1970; Berg & Аза, 1985). In contrast, populations of A. violascens in Hong Kong and Indonesia share the inter- tidal region both horizontally (Narchi, 1980; Britton, 1985; Depledge, 1985) and vertically (Soemodihardjo & Matsukuma, 1989) with other bivalve species, and may be found in sandy environments. Most specimens of A. violascens occupy an intermediate intertidal, or even subtidal position (Narchi, 1980; Soemodihardjo & Matsukuma, 1989; Willan, 1993) and are found more deeply buried than specimens of other bivalve species, at an av- erage depth of 20 cm (Narchi, 1980; Soemodihardjo & Matsukuma, 1989). Zonation patterns in intertidal bivalves have been attributed to differences in physiological tolerances to desiccation, salinity and heat stress (Britton, 1985; Depledge, 1985, and ci- tations there). Because A. deflorata and A. violascens have different horizontal distribu- tions, they likely have different physiological tolerances. The responses of A. violascens to 270 DOMANESCHI & SHEA temperature, salinity and desiccation have been assessed (Briton, 1985; Depledge, 1985), but similar experiments have not been conducted on À. deflorata. When found in gravelly sand or cobble cov- ered substrata, both A. deflorata and A. vio- lascens are densely aggregated (Stanley, 1970; Britton, 1985; Berg & Alatalo, 1985; Kurihara et al., 2000, 2001; our data). Asaphis violascens collected from sandy beaches, or beaches ei- ther with reduced cobble coverage or increas- ing deposition of sand were drastically lower in density (Soemodihardjo & Matsukuma, 1989; Kurihara et al., 2001). Willan (1993) observed that A. violascens is strictly intertidal and in- habits the lower shore where it prefers muddy sand substrata with incorporated gravel or coral rubble; uniform muddy or sandy substrata ap- pear inimical to habitation. Stanley (1970) re- ported a casual observation of Asaphis deflorata in Bermudan sand flats; Berg & Alatalo (1985) could not confirm such behavior for the spe- cies population living in the Bahamas beaches. These studies suggest Asaphis deflorata 1$ more strongly adapted to a coarse gravelly, cobble-covered sediment than is A. violascens. The presence of live, unburied specimens of A. deflorata lying by the high tide mark sug- gests that the “The Horseshoe” site population faced recent, natural disturbance of the sedi- ment and/or that specimens are able to move out spontaneously. Berg & Аза (1985) have never found live individuals lying on the sub- stratum surface in the field and considered this indicative of little natural disturbance in the sedi- ment. In the laboratory, a few specimens moved in and up within the coarse-sand substratum, this being free from natural obstacles such as pebbles, shells and rocks. Berg & Alatalo (1985) observed that the species can move in and out in disturbed substratum, such as in and around the tag-recapture plots in the field, but have difficulty penetrating the natural, un- disturbed, coarse gravel nearby. The large number of dead shells of A. deflorata retained in their life position in sta- tion 2, and their scarce presence in stations 3 and 4 (present work) may indicate that mor- tality in station 2 occurred either from senes- cence, or possibly from recent, catastrophic modification of the substratum along the low tide water, leading to suffocation or starvation of the trapped animals. Future field investiga- tion on the life cycle of A. deflorata inhabiting Florida Keys are necessary to distinguish be- tween these alternatives. Catastrophic, long-term burial by sand is thought to kill entire local populations of A. deflorata (Stanley, 1970). Berg & Alatalo (1985) identified size-independent mortality caused by movements of sand over the habi- tat of A. deflorata in the Bahamas beaches. Long-term burial by a thick layer of sand makes it difficult for clams wedged among pebbles to move to the surface (Berg & Alatalo, 1985). Although restricted to two days of field work, during a 12-day workshop period, our quanti- tative data on the “The Horseshoe” site popu- lation of Asaphis provided reliable results, as they fit well with previous ecological and bio- logical data of Berg € Alatalo (1985) on A. deflorata population in the Bahamas. Com- pared with those data obtained from January 1981 through January 1983 by Berg & Alatalo (1985) on the Bahamas population, our data reveal that both have very similar life history characteristics, including growth rates and spawning times. The population structure in July 2002 of the “The Horseshoe” site is bi- modal, as it is in the July population of the Bahamas (Berg & Alatalo, 1985), although the most frequently encountered size is smaller in the former. Size at maturity is similar for both populations, with maturation of the gonads occurring at approximately 24-25 mm in shell length, which corresponds to an age of 2-3 years (Berg 8 Alatalo, 1985). Gonad condi- tions at the “The Horseshoe” site in July were consistent with the July data from Berg and Alatalo (1985) and indicate that spawning was imminent. Statistical analyses show that shell rib num- ber and tendency to fork are good characters in distinguishing А. deflorata and А. violascens, supporting previous qualitative assessment (Prashad, 1932; Willan, 1993) of these shell characters. However, we found both species have considerably larger rib ranges than pre- dicted, and that these ranges can overlap. Furthermore, the five outlier А. violascens specimens that nest within the А. deflorata specimens suggest that rib counts and indi- ces may be better regarded as emergent prop- erties of a population, not definitive characters of the individual. Further examination of the effects of environment and geography on rib number are warranted. Distinguishing characters other than shell sculpture are: the presence of a discernible, rounded posterior radial ridge and posterior slope in the Atlantic Asaphis (our material), not discernible (Willan, 1993; IBUSP collection) in MORPHOLOGY AND SHELL MORPHOMETRY OF ASAPHIS 271 the Indo-West Pacific specimens, and a smooth inner surface in the Atlantic Asaphis, contrasting with the well-marked, ridged inner surface in specimens from Hong Kong (IBUSP collection). The effects of the environment and geography on these characters also deserve further examination. Asaphis deflorata is a native species that has been periodically collected in southern Florida and the Florida Keys for over 100 years. Asaphis deflorata should be соттоту en- countered in the Florida Keys because suit- able environmental condition occur, including gravelly sand, cobble rich intertidal habitat. The two-week planktonic larval period (Berg & Alatalo, 1985) should provide ample time for dispersal. Dispersal routes are dependent on local current regimes. Overall, current patterns in the Florida Keys move from Florida Bay, through the Keys and then join a southwest- erly countercurrent flow (http:// oceanexplorer.noaa.gov, accessed 14 Janu- ary 2003). These patterns suggest that north- ern populations could serve as a source of larvae to populate more southerly areas. Prior to the mid 1970s, A. deflorata was com- monly collected from Key West to Key Biscayne (Miami), and possibly included the Dry Tortugas (FLMNH 16919, collected in 1937). Many pre-1975 collection records sug- gest Key Biscayne had a robust population of adult A. deflorata specimens. Our search did not find any post-1975 records attributed to Key Biscayne. Although the lack of museum records does not prove Asaphis is not present in the field, the combination of no records and a single population found after extensive sam- pling of the molluscan fauna in the Florida Keys (P. Mikkelsen, pers. commun.) suggests the distribution of A. deflorata has diminished greatly. Although our data are preliminary, they point to a drastic decline in the distribution of Asaphis populations in the Florida Keys. Why the persistent “The Horseshoe” population does not act as a source population requires further investigation. The two-week larval stage of Asaphis should provide ample time for distribution to more southerly locations. However, if the spawning season does not coincide with sufficiently strong and appropri- ately oriented currents, the larvae may never be moved outside the confines of the sheltered “The Horseshoe” site. Because appropriate substrata and quiet waters are found through- out the Keys, this physical barrier is the most likely explanation for the lack of additional southerly populations. Future studies on the distribution of Asaphis deflorata should incor- porate a systematic examination of the sum- mertime current regime around the “The Horseshoe” site. The functional morphology of the Atlantic Asaphis is very similar to that of the Indo-West Pacific species as described by Narchi (1980). The correct identification of both species based upon the tissues requires the analysis of the alimentary canal. Great similarities sup- port Willan's (1993) opinion that both species share a common ancestor from which morpho- logical and ecological divergence has occurred relatively recently. The most striking anatomi- cal difference lies in the hind gut configura- tion: progressively widening, and intricately coiled and spiraled throughout its extension in the Atlantic Asaphis (our data); extraordi- nary ballooning of its proximal end only, where the mid gut enters dorsally and a narrow, non- coiled hind gut leaves ventrally, in the Indo- West Pacific specimens (Purchon, 1960; Narchi, 1980). The long, wide, coiled hind gut of A. deflorata and the dilation of that of A. violascens are both devices for retention of large amount of faecal pellets. It is difficult to understand how the compacted, randomly positioned faecal pellets are relocated from a wide compartment toward a narrow one, which is even more re- stricted in its passage through the ventricle. Peristalsis of the hind gut walls and the trans- verse musculature within the visceral portion of the foot may be the mechanism to achieve relocation of such compacted, rod-shaped fae- cal pellets. Expansion/contraction of the ven- tricle, peristalsis, and ciliary action of the very rear sector of the intestine force faecal pellets oriented end-to-end or in small groups to be expelled. Most specimens of A. deflorata examined for alimentary canal morphology had the hind gut packed with faeces, these always fully formed in their origin within the midgut. It is quite prob- able that faeces storage in Asaphis is not cor- related with the necessity to consolidate faecal pellets as proposed by Yonge (1949) for de- posit-feeding tellinoideans. Comparing the gross morphology of Abra profundorum (Smith, 1885) (Tellinoidea) and that of north Atlantic species of the genus, Allen 8 Sanders (1966) found a clear correlation between gut length and volume, size of de- posit-feeding bivalves and respective palps and ctenidia, and depth they inhabit. A reduc- tion in thickness of the gut walls, an increase of the lumen diameter allowing faecal pellets 272 DOMANESCHI & SHEA to be randomly positioned, and the gut taking up an increasingly proportion ofthe body were correlated with depth by these authors. Allen & Sanders (1966) stated that among the spe- cies of Abra they compared, “the limit of evolu- tion ofthe hind gut is reached in A. profundorum where many ofthe adjoining walls ofthe coiled gut are lost and much of the posterior half of the body above the muscles of the foot be- comes а sac containing faeces”. Asaphis deflorata inhabits shallow waters and preferentially the upper shore; neverthe- less, this species shares most of the morpho- logical features of the gut referred to by Allen & Sanders (1966) for tellinoideans inhabiting deep waters, except for a loss of walls where the coils and spiral are tightly applied to each other. Asaphis deflorata has a long intestine, com- pared with the suspension-feeding psammobiid Heterodonax bimaculatus from intertidal, coarse sand (Narchi & Domaneschi, 1993), and only moderately long compared to Gari solida from subtidal coarse sand sub- strata (Domaneschi, 1992). The exceedingly large palps, the presence of simple, digitiform tentacles around the inhalant aperture, but especially the enormous amount of faecal pellets retained within the hind gut suggest that the species deals with and ingests large amounts of material entering via the inhalant siphon. Asaphis deflorata shares most of the morpho-functional features considered by Pohlo (1982) to be characteristic of typical suspension feeding tellinoideans: large gills, no waste canal, outer demibranch not up- turned, marginal food groove present, animal lying in a vertical position within the sediment and the inhalant siphon does not take depos- ited material actively. Once extruded into the water column this siphon in A. deflorata is kept passively, much the same as in the suspen- sion-feeders tellinoideans studied by Yonge (1949), Pohlo (1972), Domaneschi (1992, 1995). However, its non-straining, curled in- ward tentacles may allow entrance of large amount of suspended material brought by the rising and falling tides, as well as of dense material lifted from the bottom. Living in the constraint of a cobble-covered sediment, the inhalant aperture more often is flush with, or slightly below the sediment surface. It contrib- utes to the intake of dense, mineral particles, benthonic microorganisms and organic and inorganic debris deposited either outside around the aperture of, or lining the passage of the inhalant siphon through the substratum. Such behavior supports Berg & Alatalo’s (1985) statement that A. deflorata is a suspen- sion and facultative deposit feeder. Phy- toplankton and C, plants detritus constitute most of its diet (Berg & Alatalo, 1985). Like A. violascens (Narchi, 1980), Gari solida (Domaneschi, 1992), and other Tellinoidea (Pohlo, 1982; Domaneschi, 1995), A. deflorata shares a mosaic of morphological features with suspension and specialized deposit-feeding tellinoideans. Such a condition was considered to represent (Pohlo, 1982) an intermediate step in the evolution of the Tellinoidea, the primitive forms represented by early suspen- sion-feeding species, and the most derived, represented by highly specialized, deposit- feeding species. When present in the beach, Asaphis deflorata has been detected to be the only bi- valve occupying the upper shore (Stanley, 1970; Berg & Alatalo, 1985; our data). The possession of a huge appendix in the stom- ach, and of a capacious hind gut allows stor- age of excess, ingested food and retention of an exceedingly large amount of faecal pellets, respectively. Both features are, probably, ad- aptations to upper shore life and non-selec- tive feeding habit, as during high tides the species can make the most of the period of submersion, both to get rid of lots of faecal pellets and take into the mantle cavity a large amount of suspended and re-suspended ma- terial, processes and ingests it. Asaphis feeds only during the period of submergence by high tides and fasts the remainder of the time (Berg & Alatalo, 1985). The huge appendix provides room for the bulk of material entering the stom- ach and mixed with enzymes liberated by the dissolving head of the style. It also prevents blockage of the main cavity of the stomach, allowing its normal functioning as already pro- posed by Yonge (1949) for the appendix of other tellinoideans. The capacious hind gut allows retention of a corresponding great vol- ume of material coming from the stomach and being molded into faecal pellets within the mid gut. Faeces have to be eliminated during sub- mergence periods only, when the exhalant current of water takes them far away from the animal; conversely, during low tides the ani- mal is at risk of faeces sedimentation within its own mantle cavity. Purchon (1960) consid- ered the capacious stomach and intestine of A. violascens a device for retention of food and faecal material during low tides, as well as for survival when the habitat is covered with sand. Capacious stomach provided with an MORPHOLOGY AND SHELL MORPHOMETRY OF ASAPHIS 273 appendix structurally and functionally similar to the appendix of the tellinoidean bivalves is also shared by the Pholadidae, Xylophaginidae and Teredinidae (Pholadoidea) (Purchon, 1941, 1955), the appendix being exceedingly long and broad to store mainly wood particles in many teredinids (Lopes et al., 2000, and citations there). Purchon (1941, 1955) con- cluded from that similarities that these struc- ture are homologous and provide reliable evidence of a relationship between these groups. Lopes et al. (2000) discussed the prob- able implications of the specialization of the appendix, as well as of the digestive system as a whole in the evolution of the xylophagous habit of two species of Teredinidae. The presence of a thin-walled, huge hind gut suggests that the compacted mass of faecal pellets of A. deflorata cannot be promptly and completely eliminated. As retention is not re- lated to the necessity to consolidate faecal pellets, it may be that another purpose is served by such an intriguing hind gut than to store, then eliminate faeces during convenient periods: for example, to allow enough time for the breakdown and consumption of material with a food value present in the faecal pellets; the enzymes required for this process could be the same as those present in the stomach and passed onto the intestine along with dis- carded material. Similar storage of faeces oc- curs in some teredinid bivalves (Lopes et al., 2000, and citations there). Long residence of faeces within the anal canal of Neoteredo reynei (Bartsch, 1920) was considered by Lopes et al. (2000) as a probable device al- lowing both enzymatic degradation of mate- rial of a food value and absorption to be continued; the presence of epithelial cells richly supplied with microvili, and the highly vascularized anal canal walls of the species giving support to this latter hypothesis. Allen & Sanders (1966) detected the pres- ence of large numbers of amoebocytes in the gut and surrounding the faecal pellets in Abra profundorum, “which would possibly indicate that the pellets provide a surface, upon which bacteria would thrive, attack and convert to a digestible form the carbon compounds indi- gestible to the mollusc.” According to Allen & Sanders (1966), “Newell (1965) has shown that faecal pellets of both Hydrobia (gastro- pod) and Macoma (tellinoidean bivalve) are excellent substrates for bacteria”, and “that the nitrogenous compounds built up by the growth of the bacteria can be digested by the mol- lusc”. Neither amoebocytes nor bacteria could be identified within the hind gut of A. deflorata through the methodology adopted in histologi- cal preparations. This cannot be taken as a definitive result, nor the hypotheses of the presence of amoebocytes and of a possible relationship of the species with symbiotic bac- teria can be discarded. Morphological and functional similarities between the gut of this psammobiid and of A. profundorum address those hypotheses. Appropriate methodologies were not employed to test the occurrence of such living elements in the gut of A. deflorata, as this was not the aim of the present work. The significance of pellet storage and the hy- pothesis that digestion and absorption of nu- trients continue within the hind gut of A. deflorata via endogenous enzymes and/or symbiosis remain to be elucidated; it cannot be fully explained until further experimental examination with live specimens has been carried out. ACKNOWLEDGMENTS We are greatly indebted to Dr. Paula Mikkelsen and Dr. Rüdiger Bieler for inviting us to participate in the International Marine Bivalve Workshop; to the members of the IMBW and Keys Marine Laboratory staff for the support and hospitality they provided us. The authors gratefully acknowledge the U. S. National Sci- ence Foundation - PEET Program (DEB- 9978119) that provided major funding for this workshop, as part of a grant to the co-organiz- ers R. B. 8 P.M., as well as the Bertha LeBus Charitable Trust, the Comer Science 8 Educa- tion Foundation, the Field Museum of Natural History, and the American Museum of Natural History for additional support. We thank Enio Matos, IBUSP, for technical assistance in the histological preparations, and Flávio D. Passos, André F. Sartori, José E. R. Marian, Sónia M. Montini, advisees of the author (OD), for their support with photographs, in mounting plates and gathering literature. 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A 7 - у y = \ 18 u О у 7 0 dl a id E Ú : “ ” LA EC 6 7 ”u o = a MALACOLOGIA, 2004, 46(2): 277-294 EXTRAORDINARY FLEXIBLE SHELL SCULPTURE: THE STRUCTURE AND FORMATION OF CALCIFIED PERIOSTRACAL LAMELLAE IN LUCINA PENSYLVANICA (BIVALVIA: LUCINIDAE) John D. Taylor'*, Emily Glover’, Melita Peharda?, Gregorio Bigatti? & Alex Ball! ABSTRACT The lucinid bivalve Lucina pensylvanica possesses an unusual flexible commarginal shell sculpture formed from calcified periostracal lamellae. The lamellae comprise thick, recurved, periostracal extensions with distal calcified scales. The periostracum is also densely embedded with calcareous granules around 2.0-2.5 um in diameter and a thin (10 um) layer of prismatic aragonite covers the ventral face of each lamella. Other species of Lucina in the western Atlantic possess calcified scales but with different morphologies and the continuous commarginal ridges of the eastern Atlantic Lucina adansoni and other Afri- can species are similarly constructed and homologous. The periostracal lamellae are a probable apomorphy of the genus Lucina and morphology of the calcified structures pro- vides a set of systematic characters of importance in the discrimination of species. Key words: Lucina pensylvanica, periostracum, calcification, shell growth, systematics. INTRODUCTION Lucina pensylvanica (Linnaeus, 1758) 1$ one of ten species of chemosymbiotic lucinid bivalves inhabiting intertidal and shallow subtidal habitats in the middle Florida Keys. Remarkably, the shell sculpture consists of closely spaced commarginal lamellae, faced with triangular, calcareous scales that are slightly flexible in live animals. The scales and lamellae become brittle after death and in beach-collected shells the surface 1$ white, relatively smooth with low, thin, commarginal ridges, sometimes with traces of periostracum. Our initial observations suggested that both lamellae and scales were a form of periostracal orextra-periostracal calcification, distinct from the normal shell. Because of the rarity of periostracal calcification in bivalves in general and the probable apomorphy of this character for Lucina spp., we decided to investigate the structure and formation ofthe lamellae in more detail and, if possible, determine the periodic- ity of their secretion. Additionally, we wanted to compare the form of the periostracal lamel- lae between Lucina species, both to establish the homology of these as well as investigate their possible use as systematic characters. Detailed understanding of lamellar formation may also suggest hypotheses about their pos- sible function. Periostracal and extraperiostracal calcifica- tion is an unusual feature of bivalves but has been described in different forms from a vari- ety of families. Usually in Lucinidae the periostracum is relatively thin (Harper, 1997), although exceptionally the genus Rasta has a dense, shaggy periostracum extended into numerous long pipes (Taylor & Glover, 1997). Prominent, sculpture-forming calcified periostracum appears restricted to the genus Lucina, of which L. pensylvanica is the type species (ICZN, 1977). The morphology of the calcified scales has been used by Gibson- Smith 8 Gibson-Smith (1982) as a character to divide “Lucina pensylvanica” of the west- ern Atlantic into four separate species. Amongst other bivalve families, Veneridae, such as Lioconcha and Callocardia possess encrustations formed of fine aragonitic needles projecting through the periostracum (Ohno, 1996; Morton 2000); others such as Granicorium and Samarangia secrete extra- ‘Department of Zoology, The Natural History Museum, London SW7 5BD, United Kingdom Institute of Oceanography and Fisheries, РО Box 500, 21000 Split, Croatia “Departado Biologia, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Pab Il, C1428EHA Buenos Aires, Argentina “Corresponding author: j.taylor@nhm.ac.uk 278 TAYLOR ET AL. periostracal calcareous cements to form a crust of sediment on the shell (Taylor et al., 1999; Braithwaite et al., 2000). Many Anomalodesmata, such as Laternula and Lyonsia, possess spines formed within the periostracum as do some Gastrochaenidae, such as Spengleria rostrata (Spengler) (Carter & Aller, 1975). Amongst the Mytilidae, intraperiostracal aragonitic granules and pro- jecting spikes have been described in Trichomya and Brachidontes (Carter & Aller, 1975; Bottjer & Carter, 1980; Carter et al. 1990), while intra- and extraperiostracal cal- cified structures are a feature of various spe- cies of Lithophaginae (Carter et al., 1990). Little is known of biology of Lucina pensylvanica. Stanley (1970) demonstrated using x-rays that animals burrowed with the anterior part of the shell lying uppermost in the sediment, an unusual life orientation for Lucinidae. The general anatomy was de- scribed by Allen (1958) and Gros et al. (1996) made a detailed description of the gill ultra- structure and chemosymbiotic bacteria. Addi- tionally, Taylor & Glover (2000) illustrated the large bipectinate mantle gills that lie alongside the pallial blood vessel. Lucina pensylvanica and its close allies are often referred to in the literature under the generic name Linga. However, the name should correctly be Lucina as Lucina pensylvanica was designated the type species of the genus in 1977 (ICZN, 1977). MATERIALS AND METHODS Lucina pensylvanica was live collected from a number of oceanside intertidal and shallow water sites in the Florida Keys during the In- ternational Marine Bivalve Workshop (IMBW) in 2002 (Mikkelsen & Bieler, 2004, fig. 1 — map). Live animals were abundant only at Sta- tion IMBW-FK-642, mile marker 74.5 (24°51.4’М, 80%43.7'W) on Lower Matecumbe Key. Here they occurred in low intertidal to shallow, subtidal pockets of medium to coarse sand, located on a wide, coral-rock platform. The area was vegetated with patches of Thalassia and Halodule, as well as growths of Penicillus and Halimeda. Despite similar col- lecting effort, Lucina pensylvanica was much less common at other sites, such as Anne's Beach, Upper Matecumbe Key (Station IMBW- FK-638) from Thalassia-covered sand and Pigeon Key (Station IMBW-FK-657) in a tidal stream with Thalassia and Syringodium. No live animals were found at any bayside sta- tions. Animals were collected by extensive dig- ging and hand sieving. Voucher specimens held in BMNH, London. Live animals were fixed in 75% ethanol, 5% seawater formalin or Bouin's fluid. Tissue samples were also fixed in 2.5% solution of glutaraldehyde in phosphate buffer. Sections of mantle were stained with Mallory's triple. For optical microscopy of the shell, geological thin sections were made from fresh specimens embedded in resin. Pieces of the same em- bedded shell were also examined by scanning electron microscopy (SEM) after cutting, pol- ishing and etching in EDTA. Shell sections were also examined by con- focal microscopy using a Leica SP NT in re- flected light mode. Simultaneous images were collected at several different wavelengths, and a reference image was obtained with the trans- mitted light detector. We also carried out an initial test for autofluorescence using a wave- length (lambda) scan. The section was scanned at a single focal plane with each la- ser in turn. The detector was programmed to step through 25 pre-determined 10 nm-wide detection windows at wavelengths from 495— 750 nm that produced an intensity profile for each emission wavelength. This optimised la- ser detector position and line. The best results were obtained with the 488 nm Argon laser and this was used for all subsequent imaging. No autofluorescence was detected from within the shell matrix, so the first detector window was set at 486-507 nm. This wavelength gave a direct reflection image of the sample and was false coloured in green. Strong autofluorescence from the periostracum was detected at around 550 nm, so the second detector window was set at 537-568 nm and the images coloured red. A stack of 30 images was collected at ~0.4 um intervals. Each frame was scanned three times and run through a frame-averaging filter to reduce background noise. For single images, the z-axis (depth) data from the entire stack was combined and the brightest pixel from each point computed and displayed (maximum projection image). Growth Periodicity Twenty valves from live collected animals were used to study growth periodicity. We embedded these in MET20 resin (Struers Ltd), sectioned them transversely from the umbo to the ventral edge. They were then ground, polished and etched for 20 min in 0.01M HCI SHELL SCULPTURE IN LUCINA PENSYLVANICA 2119 and acetate peel replicas prepared following Richardson (2001). Distances between suc- cessive periostracal lamellae were measured to the nearest 0.05 mm on 11 shells. Distinc- tive major lines in the ощег and middle shell layers and in the umbonal region (Fig. 22) were correlated with the formation of closely spaced or uncalcified periostracal lamellae. Three separate observers used these major growth marks in both umbo and valve to estimate the age in years of the animal (Richardson, 1993). The major growth increments were treated as annual lines by comparison with a similar study of Codakia orbicularis from the Bahamas (Berg 8 Alatalo, 1984). RESULTS Shell Microstructure The shell consists of three aragonitic layers. The outermost layer is composed of a pris- matic layer of irregular acicular crystals, their long axes inclined towards the shell margin (irregular spherulitic structure of Carter & Clark, 1985). This is followed by a middle layer of finely lamellate, crossed-lamellar structure and, within the pallial myostracum comprising irregular prisms, there is an inner layer formed of complex crossed-lamellar structure, inter- calated with thin prismatic sheets. This se- FIG. 1. Lucina pensylvanica exterior of right valve showing commarginal periostracal lamellae with projecting calcareous scales. Shell height = 22.8 mm. Station IMBW-FK-642, Mile Marker 74.5, 24°51.4’М, 80°43.7’W, on Lower Matecumbe Key. quence of shell layers resembles most other Lucinidae (Taylor et al., 1973). Calcified Periostracal Lamellae Periostracal lamellae (hereafter referred to as lamellae) consist of an extended periostracum sheet faced with prominent cal- cified scales (Fig. 1). The lamellae recurve dorsally and are regularly spaced at intervals of 400-1500 um, extending about 1,000 um from the shell surface. Interspaces between the lamellae are relatively smooth (Figs. 2, 3) and in live collected specimens are packed with sediment grains (Figs. 8, 9). The discrete, closely-spaced calcareous scales (Fig. 8) are around 600-1,000 um in height and seemingly embedded into the periostracum. In shape, the scales are triangular to lanceolate, broad at the base (varying between 500-950 um) and taper distally. When newly formed, they are usually pointed at the tips (Fig. 4) but become truncated with wear. Scale shape varies around the shell; those on the posterior dor- sal area are usually broader, more closely spaced and less recurved. Over most of the shell surface, lamellae recurve dorsally but when first formed they extend straight out from the shell margins, with the scales embedded in the sheet of periostracum (Fig. 4). Subse- quently, lamellae become progressively re- curved away from the commissure (Fig. 4), and the periostracum erodes away from the scales (Figs. 5, 6). On juvenile shells, the scales are differently shaped (Fig. 7) being lower and quadrate with narrower spaces between, so that they form an almost continuous ridge. The quadrate scales change to a triangular shape at a shell height of around 4.5-5.0 mm. Sections Optical, scanning and confocal microscopy shows that each lamella is composed of a periostracal extension in which the calcare- ous scales occupy the distal ventral face (Figs. 8-11). Each lamella projects from a thin ridge in the true shell (Figs. 8-10). Within a lamella the periostracum is about 55 um thick and continuous with that of the outer shell surface. Between successive lamellae the periostracum gradually increases in thickness from around 1-2 um at the termination of one extension to about 50 um at the base of the succeeding extension (Fig. 12). Higher mag- nification of the calcareous scales reveals a 280 TAYLOR ET AL. FIGS. 2-4. Lucina pensylvanica. FIG. 2: Surface view of successive commarginal lamellae with scales. Scale bar = 500 um; FIG. 3: Periostracal lamellae on posterior of shell with pointed scales with smooth periostracal surface between lamellae. Scale bar = 500 um; FIG. 4: Site of formation of periostracal lamellae at valve margins showing lamellae lying parallel with shell margin but becoming recurved dorsally away from the edge. Scale bar = 500 um. SHELL SCULPTURE IN LUCINA PENSYLVANICA 281 FIGS. 5, 6. Lucina pensylvanica. FIG. 5: Ventral view of forming lamella at shell margin showing row of scales embedded in periostracum stretched between them, but in the preceding row this has disappeared. Scale bar = 250 um; FIG. 6: View of posterior shell margin with pointed scales joined by a membrane of periostracum. Scale bar = 250 pm. FIG. 7. Lucina pensylvanica, juvenile shell (shell height 3.5 mm) with lamellae formed of closely spaced, quadrate scales. Scale bar = 200 um. FIGS. 8, 9. Lucina pensylvanica. FIG. 8: Transverse section of shell showing two lamellae. Note ridges in shell and sediment trapped behind lamellae. Scale bar = 250 um; FIG. 9: Transverse section of a single lamella. Scale bar = 250 um. thin (1.5-2.0 um) initial periostracal sheet fol- lowed by a layer of aragonitic spherulitic mi- crostructure (Fig. 14). Each scale is about 220 um thick tapering distally. Within the spheru- litic layer of the scale, interpenetrant bundles of long, thin crystals radiate from nucleation sites on the inner periostracal surface. Fine growth lines indicate that the scales are se- creted incrementally. Another calcified layer (10-15 um thick), of short, prismatic arago- nite crystals embedded in periostracum, forms the ventral face of each completed lamella (asii 13.18). Sections of the basal periostracal part of the lamella show that it is densely embedded with tiny calcareous granules about 2-2.5 um in diameter consisting of aggregations of crys- talline aragonite (Figs. 13, 16, 19). Granules are absent in the outermost of part of the periostracum but at about 10 um from the edge ofthe lamella increase in abundance (Fig. 12). 282 TAYLOR ЕТ AL. FIGS. 10, 11. Lucina pensylvanica. FIG. 10: Confocal image of transverse section through a periostracal lamella. Periostracum red; calcified structures green. Scale bar = 100 ит. Abbreviations: pf, calcified prismatic front of lamella; pl, periostracum of lamella; ps, periostracum above shell; r, ridge in outer shell layer; s, shell; sc, scale; FIG. 11: Confocal image of the proximal region of a periostracal lamella, showing detail of the periostracum and the calcified front of the lamella. Scale bar = 50 um. Abbreviations: as for Fig. 10; gz, granule zone; oz, ощег granule-free periostracal zone. FIG. 12. Lucina pensylvanica, SEM image of transverse section through base of a lamella showing shell ridge and thinned periostracum that thickens towards the succeeding lamella. Scale bar = 100 um. Abbreviations: osl, outer shell layer; р, periostracum; pf, prismatic front of lamella; pl, periostracum of lamella; tp, thin periostracum. SHELL SCULPTURE IN LUCINA PENSYLVANICA 283 These granules are also present in the nor- mal periostracum secreted above the outer shell layer and gradually increase in frequency between successive lamellae. Sections of the junction between the calcar- eous scales and the periostracal lamella show that lines representing growth increments in- terdigitate from periostracum into the calcified scales and also that the granules increase in density and fuse at the transitional boundary (Figs. 13, 15). The calcified scales are thus secreted contemporaneously with the periostracal layers of the lamella and not laid down subsequent to it. Images clearly show a covering of periostracum eroding from the scale surfaces. We conclude from these ob- servations that both the granules and scales are forms of periostracal calcification. FIGS. 13-15. Lucina pensylvanica. FIG. 13: SEM image of a transverse section through junction between calcareous scale and proximal part of the lamella showing interdigitation of calcareous layer with periostracum and granules. Scale bar = 50 ит; FIG. 14: Section through a calcareous scale showing spherulitic crystal growth arising from thin periostracum layer below. Scale bar = 70 um; FIG. 15: Section through junction of calcareous scale and periostracum showing continuity of growth increments from the calcified portion into the periostracum. Scale bar = 50 pm. Abbreviations: gz, granule zone of periostracum; р, periostracum; sp, spherulitic crystal growth. 284 TAYLOR ET AL. FIGS. 16-19. Lucina pensylvanica. FIG. 16: Surface of a forming periostracal lamella at shell margin showing aragonitic granules embedded in surface. Scale bar = 50 um; FIG.17: Higher magnification image of granules showing crystalline form. Scale bar = 15 um; FIG.18: Section of periostracal lamella showing discrete aragonitic granules in periostracum and the fringe of prismatic aragonite crystals along the front of the lamella. Scale bar = 20 um; FIG. 19: Detail of discrete granules embedded in periostracum. Scale bar = 2 um. Mantle Edge gations indicating the potential for consider- able extension. Epithelial cells at the margin The mantle edge of L. pensylvanica is thick are tall, with nuclei located towards the mid- and divided into several folds (Fig. 20). The point, but decrease in height dorsally to the large outer fold (of) is thrown into deep corru- short, cuboidal cells of the general outer SHELL SCULPTURE IN LUCINA PENSYLVANICA 285 FIG. 20. Lucina pensylvanica. Transverse section of anterior mantle edge. Mallory's triple stain. Scale bar = 250 um. Abbreviations: с, cuticle; fme, corrugated mantle epithelium of outer fold; gz, glandular zone; if, inner mantle fold; Im, longitudinal pallial muscles; mf(1) & mf(2), lobes of middle mantle fold; of, outer mantle fold; ome, outer mantle epithelium; p, periostracum; pg, periostracal groove; rm, radial pallial muscles. mantle surface. The outer fold is separated from the middle fold by a deep periostracal groove, with the forming periostracum lying against the outer surface of the middle fold. The middle fold is divided into two distinct lobes with the outermost of these (mf 1) form- ing a short, slender lobe whilst the other (mf 2) is broad and longer. The inner fold (if) is a small, low ridge. Cells of the middle lobes are shorter than those of the outer fold and pos- sess basal nuclei. The epithelium of the middle folds is overlain by a thin cuticle (ct) that ex- tends almost to the inner fold. The mantle sur- face within the inner fold is ciliated. Two well-defined bundles of radial muscles extend into the outer and middle folds respec- tively and a thick bundle of longitudinal pallial muscles (Im) is located near the inner fold (seen in transverse section in Fig. 20). The inner part of the mantle within the inner fold is highly glandular with subepithelial gland cells opening to the inner mantle surface. Two types of gland cell are present; one type, staining blue, is located superficially while the other dark green type lie more deeply. Periodicity of Lamellae The lamellae appear regularly spaced but measurements taken from acetate peels of shell sections show that the increments are variable in width and furthermore change with age. Figure 21 demonstrates that for eight live- collected shells widths between successive lamellae increase steadily from around 200- 450 um to a maximum (up to 1,800 um) at around 25-30 mm shell height. Thereafter, interlamellar spacing becomes much narrower but more variable. Observations of the outer surfaces of larger, dead-collected shells show 2- 2 ] 1.8 4 1.8 Specimen 3 Specimen 5 8161 or ae © vw € 1.47 Е 14 © iy eur] 2 2 о oO Oe aid 2 1 Е 5 E 0.8 4 2 0.81 8 0.6 8 0.6 | 8 0.4 504 D TD = 0.24 0.2 - 0 + - т т = 0+ т т ДА 0 10 20 30 40 0 G 10 15 20 25 30 335 40 cumulative distance cumulative distance 27 DIE is | 1.8 | Specimen 6 Specimen 8 © 16 916 Г Е ] Е 1.44 Е 1.4 = 152 | Е 152%] U ® Ф 1 2 1 3 © 208] 30.8 ® ® = 0.6 9 Olen 2 2 on 0.4 m 0.4 TD ho] | 0:23] 0.2 0 + т т т 1 DE т т т т —— т т — 0 10 20 30 40 0 5 10 15 20 25 30 35 40 cumulative distance cumulative distance 27 р 18 1.8 1 (0) v : S 1.6 Specmen 10 a 1.6 4 Specimen 12 vo 4 = 1.4 7 Е 1.4 = 12 = 172%] 91 | 271 Е Е 2 0.8 - 2 0.8 U ® = 0.6 | = 0.6 » 0.4 w 0.4 TD DT 0:2 0.2 0 je T T T 7 0 т oe oe SS en ee 0 10 20 30 40 0 5 10 15 20 25 30 35 40 cumulative distance cumulative distance 25 DE 1.8 | 1.8 - Specimen 20 o 4 [0] ES ne Specimen 18 & Es uo uo Е 1.47 Е 1.4 © 5 12 = 1.22 u ры 3 3 3 0.8 y E 0.84 Ф i ® = 0.6 © 0.6 © д 0.4. Я 0.4 Le) ho] 0.2 0.24 о + - 1 : à 0 +— oH _— и — 0 10 20 30 40 0 5 10 15 20 25 30 35 40 cumulative distance cumulative distance FIG. 21. Lucina pensylvanica, interval between successive lamellae plotted against cumulative length around shell circumference for eight individual Lucina pensylvanica. Measurements made from acetate peels of transverse sections. SHELL SCULPTURE IN LUCINA PENSYLVANICA 287 + + “il 5 к 3 à es : Pt RR, RTE \ x RAS). ЧН FIG. 22. Lucina pensylvanica, acetate рее! of transverse section of shell showing major growth line extending through outer and middle shell layers. Scale bar = 500 pm. that this change in the interlamellar interval is Frequently, major growth halts are marked by visible on all individuals at shell heights of the secretion of a sequence of several un- around 22-27 mm. In older individuals the in- calcified periostracal extensions (Figs. 22, 23). terval between major growth halts is narrower Our interpretation of this growth pattern is that with fewer lamellae (Fig. 21: specimen 12). shell accretes rapidly and uninterrupted to a FIG. 23. Lucina pensylvanica, semidiagrammatic summary drawing (based on camera lucida image) of transverse section through shell showing successive lamellae and two growth halts where only uncalcified periostracal sheets were secreted. Scale bar = 1.0 mm. Abbreviations: cf, calcified front of lamella; msl, middle shell layer; osl, outer shell layer; ps, periostracum above shell; pl, periostracum of lamella; r, ridge in outer shell layer; sc, scales; upl, uncalcified periostracal lamellae. 288 TAYLOR ET AL. FIG. 24. Lucina pensylvanica, semidiagrammatic transverse section through a single periostracal lamella. Scale bar = 500 um. Abbreviations: gz, granule zone; osl, outer shell layer; pf, thin prismatic ventral fringe to lamella; tp, thin periostracum. size of around 25 mm. Thereafter, growth rates decline and become more variable. Study of gonads from our small sample indicates that sexual maturity occurs in these bivalves at shell heights of around 20-25 mm (Bigatti et al., 2004). The major change in shell growth pattern may thus coincide with time of first spawning. A study of growth in Codakia orbicularis (Linnaeus, 1758) from the Bahamas showed that prominent growth rings in the shell were annual (Berg & Alatalo, 1984). Following this, the major growth halt lines seen in shell sec- tions (Figs. 22, 23) in our sample could be ten- tatively interpreted as annual marks and used to estimate the ages of the animals. Table 1 indicates that 20 sectioned shells show be- tween 0-4 major lines and the interpretation is that the animals vary between one and four years old. Proper age estimation should be done using marked and calibrated shells but this was impossible in the time available for the study. Sequence of Secretory Events The structure of the commarginal lamellae 1$ summarized diagrammatically in Figures 23- 24. Each commarginal lamella represents an extension of the mantle beyond the normal shell profile. Although the lamellae in L. pensylvanica are recurved dorsally, observations at the site of secretion show that the lamellae initially project more or less straight from the valve margin and curve dorsally later (Fig. 4). Thus, the mantle is not extended and reflected dor- sally as it would be if secreting commarginal lamellae formed from normal shell layers as seen in other bivalves such as the venerid Placamen calophyllum (Philippi, 1836) (Checa, 2002). Initially, the mantle secretes a thin, periostracal sheet, followed by calcification of the distal portion with spherulitic aragonite crys- tals. Calcification of the distal edge of the lamella is localised, presumably to groups of cells, so that individual scales are formed. At the same time the proximal part of the lamella is laid down as periostracum, embedded with crystalline granules. Finally, the mantle withdraws from the extended position, leaving a thin layer of pris- matic crystals along the ventral face of the lamella. The withdrawal of the mantle is marked by a low, commarginal ridge in the shell profile (Figs. 12, 24). Following termination of a lamella, the periostracum is very thin but gradu- ally thickens and becomes densely embedded with granules prior to the next lamellar exten- sion (Fig. 24). Periodically, there are major growth breaks where only extended uncalcified periostracal sheets are formed (Fig. 23). Comparison with Lucina adansoni and Other Species An interesting comparison may be made with another species, Lucina adansoni (Orbigny, 1839) from West Africa. This has a thick, subspherical shell, sculptured, with closely spaced, broad commarginal lamellae about 300 um in width (Figs. 25, 26). These are often eroded, detached or absent in dead-collected shells or museum specimens. Each lamella is divided into sections (up to 500 рт long) by narrow sutures aligned between successive lamellae. Interspaces between lamellae are often packed with sediment. 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PL A9} aquinoajeyy Jamo] Zÿ9 97 С € peumeds 3 GIL 902 906 9 Kay эашпоззеи\ Jamo7 Z9 LZ + 0 Burumeds 4 о Gl 90.9 AS} эашпозэуеи JAMO] Zÿ9 € € eine 3 о а OC Ey, Kay эашпоззеи Jamo7 су9 05 est | Bulumeds/ Jen 3 дор EGS LC $ Ady эашпээе\ лэмо] 59 Gal 0 Buinjen 3 18 ЗА ПИ A8} equinoajey Jamo] Zÿ9 € С psumeds 3 IA OUEN цоцеэо7 uoneysS (suauioads pajoajas) — э}ец]5э saul ymoJb a}e}s x2S M H 7 al -44 эе|эше!| paniasqo N aby Jofew N aAnonpoide} "вошелИзиэа eulsn7 и! эе|эше| рэллэ$4о jo лэашпи pue saul UJMOAB jo UOIISOd ‘uorIpuo9 элцопрол9э/ 'xas ‘abe ‘ezis |э4$ ‘р FIGVL 290 TAYLOR ЕТ AL. that is triangular in cross section (Fig. 27). The tally in the calcified unit. This is more heavily lamellae are tilted towards the ventral shell calcified than the scales of L. pensylvanica but margin rather than recurved dorsally as in L. similarly constructed of spherulitic crystal pensylvanica. Each lamella is composed of a growth. The periostracal extensions are shorter thick periostracal extension that terminates dis- than L. pensylvanica but similarly embedded FIGS. 25-27. Lucina adansoni. FIG. 25: Right valve (Leiden RMNH 12179). Cape Verde Islands, SE of Boa Vista 15°59’N, 22°44’W, depth 36 m. Shell height = 32.5 mm; FIG. 26: Detail of commarginal lamellae. Arrows mark suture lines between sections along lamellae. Note sediment grains packed into interspaces between lamellae. Scale bar = 1.0 mm; FIG. 27: Transverse section of a commarginal lamella. Scale bar = 500 um. Abbreviations: pl, periostracal lamella; ps, periostracum above shell; $, shell; sc, calcareous scale. SHELL SCULPTURE IN LUCINA PENSYLVANICA 291 with calcareous granules about 2 pm in diam- eter (Fig. 28). Also, the periostracum gradu- ally increases in thickness between successive lamellae and then thins dramatically at their termination (Fig. 28). Beneath each lamella the outer shell layer forms a steep-faced lip (Fig. 28) about 200 pm high. In worn shells this is the only shell sculpture remaining after the lamellae have become detached. Calcified periostracal commarginal lamellae similar to those of L. adansoni have been ob- served (BMNH collections) in the southern African species Lucina carnosa Dunker, 1858, and L. roscoeorum (Kilburn, 1974). The lamel- lae in the latter species are described (Kilburn, 1974: 340-341, figs, 4, 5) as being “...apically imbricate, rendering their crests somewhat tabulate (i.e. in cross section each would re- semble an inverted *L”)...” and “... the crests of the lamellae are regularly but superficially incised transversely...”. DISCUSSION AND CONCLUSIONS We have demonstrated that the structurally complex commarginal shell sculpture of Lucina pensylvanica is a form of periostracal calcifi- cation, a rather unusual feature amongst bivalves. The calcareous granules within the periostracum were briefly mentioned by Bottjer & Carter (1980), but no details were given. We are not aware of any similar structures in any other lucinid. Most Lucinidae lack prominent commarginal shell sculpture but two species of Lamellolucina, namely L. dentifera (Jonas, 1846) from the Red Sea and L. gemma (Reeve, 1850) from the Philippines possess thin, elevated lamellae with spinose edges (Taylor 8 Glover, 2002: fig. 6) reminiscent of the lamellae in L. pensylvanica. However, the lamellae and spines of Lamellolucina are en- tirely calcareous and comprise extensions of the outer shell layer rather than periostracal FIG. 28. Lucina adansoni, confocal image of transverse section of a commarginal lamella. Periostracum red, calcareous components green. Scale bar = 100 um. Abbreviations: gz, granule zone of periostracum; pl, periostracal lamella; ps, periostracum above shell; r, ridge in outer shell at base of lamella; s, shell; sc, calcareous scale; tp, thin periostracum. 292 TAYLOR ET AL. structures. Similarly, Lucinisca species from the western Atlantic and eastern Pacific pos- sess spinose commarginal lamellae, but again these are formed from the outer shell layer rather than periostracum. A diversity of instances of periostracal calci- fication has been described from a wide range of different bivalve families (Carter & Aller, 1975; Bottjer & Carter, 1980; Carter et al., 1990; Ohno, 1996; Morton, 2000), but none is сот- parable with L. pensylvanica. Analogous cal- careous granules embedded in periostracum have been illustrated for the mytilids Brachidontes granulatus (Bottjer 8 Carter, 1980: fig. 3) and Trichomya hirsuta (Carter & Aller, 1975: fig. 1c). Little attention has been paid to this calcification either functionally or as a set of systematic characters and in many cases it is routinely cleaned off specimens. Function of the Lamellae Although we have no experimental evidence, we suggest by analogy with sculpture on other bivalves that there might be at least three pos- sible functions of the commarginal lamellae. These include acting as a sculptural aid to burrowing, maintaining stability in the sediment and as a possible deterrent to predators. Un- usually amongst bivalves, the commarginal lamellae of L. pensylvanica are flexible in life and this property may have added but un- known functional significance. As demonstrated by Stanley (1970), some lucinids, including Lucina pensylvanica, bur- row into the sediment vertically with the hinge axis parallel to the sediment surface and rock from side to side to gain purchase into the sand. Unusually for lucinids, L. pensylvanica rotates posteriorly after penetrating the sedi- ment to lie with the anterior part of the shell uppermost. The recurved, flexible lamellae and scales might aid this process but we have no experimental evidence similar to that available for the divaricate-ribbed Divaricella quadrisulcata (Orbigny, 1846) (Stanley, 1970). However, the external lamellae of L. pensylvanica are easily removed to enable a comparison of burrowing performance to be made with and without the structures. In shallow burrowing bivalves, the ridges and spines on the shell surface have been shown to reduce the effects of scour and may pre- vent dislodgement from the sediment (Bottjer 8 Carter, 1980; Stanley, 1981). We have no experimental observations but in Lucina pensylvanica and L. adansoni the lamellae are extremely effective in trapping sediment close to the shell surface (Figs. 8, 26) and in most live-collected specimens the interlamellar spaces are full of sediment. Compared to other lucinids of similar size from the Florida Keys, Lucina pensylvanica is the most shallowly burrowed, living in medium to coarse, mobile sands rather than the thicker Thalassia-bound sediments favoured by Codakia orbicularis and Anodontia alba. A further possible function of the lamellae might be to deter predation. Strong commar- ginal lamellae on the venerid Placamen calophylum have been shown to deter shell drilling predatory gastropods (Ansell 8 Morton, 1985). Any test of this suggestion would need experimental analysis. The function of the discrete aragonitic gran- ules embedded in the periostracum and periostracal extensions of L. pensylvanica and L. adansoni is unclear, but they may provide additional stiffness to the largely proteinaceous part of the lamellae that supports the more heavily calcified distal scales or ridge. Further- more, the thin calcified layer along the ventral face of the lamellae may also provide stiffness but, additionally, the differential mechanical properties on either face of the lamella may cause the lamellae to curve dorsally. Systematic Implications of Commarginal Lamellae in Lucina Although Lucina pensylvanica is thought to be widely distributed around the Western At- lantic and Caribbean area, from North Caro- lina to Brazil (Britton, 1970; Abbott, 1974; Bretsky, 1976), it is much more likely that a complex of several species exists. J. Gibson Smith 8 W. Gibson Smith (1982) used the morphology of the calcareous scales to divide the “L. pensylvanica” of the western Atlantic, naming three new species on the basis of dif- ferences in the form of the scales. These they distinguished from L. pensylvanica, assuming its type locality to be Florida. All the species are similar in general shell morphology but differ in the form of the calcified periostracal lamellae. We have examined the types of the Gibson-Smith species and also the syntypes of Lucina pensylvanica (Linnaeus, 1758), but unfortunately the latter material is heavily worn without any trace of lamellae. Firstly, Lucina belizana J Gibson-Smith 8 W Gibson-Smith, 1982 (Holotype: BMNH 1980103) from Belize is characterised by fine, close lamellae with delicately pointed, lightly SHELL SCULPTURE IN LUCINA PENSYLVANICA 298 calcified spines. Secondly, Lucina roquesana J Gibson-Smith & W Gibson-Smith, 1982 (Ho- lotype and paratype: BMNH 1980105/1-2) from Venezuela has calcified periostracal lamellae, but these bear broad closely spaced, blunt-ended scales that are arranged in a ra- dial rows in successive lamellae. Lucina podagrina caymanana J Gibson-Smith & W Gibson-Smith, 1982 (Holotype: BMNH 1980104/1) from the Cayman islands is simi- lar to L. roquesana, but the periostracum 1$ pale brown and the shell less globose (Lucina podagrina podagrina Dall, 1903, is a Pliocene fossil species.). J. Gibson Smith 8 W. Gibson Smith (1982) have undoubtedly highlighted the existence of a species complex within the former “Lucina pensylvanica”, but in our opin- ion the taxonomy is even more complicated. For example, another species from the west- ern Atlantic, Lucina aurantia Deshayes, 1830, which is usually synonymised with L.: pensylvanica (Abbott, 1974; Britton, 1971; Bretsky, 1976), has many distinctive shell char- acters including size and shape, dentition and colour. Some unworn shells have remnants of fine, pointed scales. We are confident that this is yet another unregarded species. Another likely distinct species from the Bahamas has been confused with L. pensylvanica but it can readily distinguished by extremely fine pointed scales (specimens from Blue Hole Cay, off Andros Is., collected by P. Mikkelsen and С. Hendler). Athorough systematic revision of the “Lucina pensylvanica” complex in the western Atlantic using live-collected animals with mor- phological and molecular analysis is needed. On the other side of the Atlantic, Lucina adansoni, L. carnosa, and L. rosceorum seem to form another possibly related clade, linked by the possession of calcified periostracal lamellae that form continuous ridges. As we have demonstrated, these ridges differ in mor- phology but are similarly constructed and thus homologous with the lamellae of the western Atlantic “L. pensylvanica” group. The relation- ships of the two clades need clarification. It should be emphasized that in museum specimens the periostracal calcified structures so diagnostic of these Lucina species are usu- ally damaged or in the case of beach collected shells, completely worn way. In dried shells, the periostracal lamellae become brittle and are easily damaged without special curatorial care. We recommend wet preservation as the most satisfactory method of preserving these structures. ACKNOWLEDGEMENTS We are indebted to Paula Mikkelsen and Rudiger Bieler for organising and inviting us to participate in the International Marine Bio- logical Workshop. We thank Dave Cooper and Tony Wighton for making the thin sections of mantle and shells; A. Pallaoro and M. Kraljevic helped with analysis of shell sections, and A. Zuljevic produced Figure 22. We are grateful to E. Gittenberger and J. Goud (Nationaal Natuurhistorisch Museum, Leiden) for the loan of Lucina adansoni specimens and for permis- sion to section a shell. P. Mikkelsen kindly sent additional specimens from the Bahamas. Fi- nally, thanks are due to our colleagues at the workshop for good humoured company and advice on sampling locations. Specimens were collected under Permit FKNMS-2002.079. The International Marine Bivalve Workshop, held in the Florida Keys, 19-30 July 2002, was funded by U.S. National Science Foundation award DEB-9978119 (to co-organizers R. Bieler and P. M. Mikkelsen), as part of the Partnerships in Enhancing Expertise in Tax- onomy (PEET) Program. Additional support was provided by the Bertha LeBus Charitable Trust, the Comer Science & Education Foun- dation, the Field Museum of Natural History, and the American Museum of Natural History. LITERATURE CITED ABBOTT, К. T., 1974, American seashells, 2" ed. Van Nostrand Reinhold Co., New York, 663 pp. ALLEN, J. A., 1958, On the basic form and ad- aptations to habitat in the Lucinacea (Eulamellibranchia). Philosophical Transac- tions of the Royal Society of London, Ser. B, 241: 421-484. ANSELL, A. D. & B. MORTON, 1985, Aspects of naticid predation in Hong Kong with special reference to the defensive adaptations of Bassina (Callanaitis) calophylla (Bivalvia). Pp. 635-660, т: В. MORTON & D. DUDGEON, eds., Proceedings of the Second International Work- shop on the Malacofauna of Hong Kong and Southern China, Hong Kong 1983. Hong Kong University Press, Hong Kong. BERG, C. J. & P. ALATALO, 1984, Potential of chemosynthesis in molluscan mariculture. Aquaculture, 39: 165-179. BIGATTI, G., M. PEHARDA & J. D. TAYLOR, 2004, Size at first maturity and external mor- phology of sperm in three species of Lucinidae (Mollusca: Bivalvia) from Florida Keys, USA. Malacologia, 46(2): 417-426. BOTTJER, D. J. & J. С. CARTER, 1980, Func- tional and phylogenetic significance of project- 294 TAYLOR ЕТ AL. ing periostracal structures in the Bivalvia (Mol- lusca). Journal of Paleontology, 54: 200-216. BRAITHWAITE, С. J. R., J. D. TAYLOR 4 Е. A. GLOVER, 2000, Marine carbonate cements, biofilms, biomineralization and skeletogenesis: some bivalves do it all. Journal of Sedimen- tary Research, 70: 1129-1138. BRETSKY, S. S., 1976, Evolution and classifica- tion of the Lucinidae (Mollusca; Bivalvia). Palaeontographica Americana, 8(50): 219-337. BRITTON, J. C., 1970, The Lucinidae (Mollusca: Bivalvia) of the western Atlantic Ocean. PhD Dissertation, George Washington University. University Microfilms 71-12,288. 566 pp., 23 pls. CARTER, J. С. & К. С. ALLER, 1975, Calcifica- tion in the bivalve periostracum. Lethaia, 8: 315-320. CARTER, y. С. & С. R. CLARK, 1985, Classifi- cation and phylogenetic significance of mollus- can shell microstructure. University of Tennessee Department of Geological Sciences Studies in Geology, 13: 50-71. CARTER, J. G., R. A. LUTZ 8 M. J. S. TEVESZ, 1990, Shell microstructural data for the Bivalvia. Part VI. Orders Modiomorphoida and Mytiloida. Pp. 391-411, in: J. G. CARTER, ed., Skeletal biomineralization: patterns processes and evolutionary trends, Vol. 1. Van Nostrand Reinhold, New York. CHECA, A. G., 2002, Fabricational morphology of oblique ribs in bivalves. Journal of Morphol- оду, 254: 195-209. GIBSON-SMITH, 4. & W. GIBSON-SMITH, 1982, Lucina s.s. (Mollusca: Bivalvia) in the western Atlantic: a reappraisal. Veliger, 25: 139-148. GROS, O., Е. FRENKIEL & M. MOUEZA, 1996, Gill ultrastucture and symbiotic bacteria in the tropical lucinid, Linga pensylvanica (Linné). Symbiosis, 20: 259-280. HARPER, E. M., 1997, The molluscan peri- ostracum: an important constraint in bivalve evolution. Palaeontology, 40: 71-97. ICZN, 1977, Opinion 1095 — Designation under the plenary powers of Venus pensylvanica Linnaeus, 1758, as type species of Lucina Bruguière, 1797 (Mollusca, Bivalvia). Bulletin of Zoological Nomenclature, 34: 150-154. KILBURN, R. N., 1974, Taxonomic notes on South African marine Mollusca (4): Bivalvia, with descriptions of new species of Lucinidae. Annals of the Natal Museum, 22: 335-348. MIKKELSEN, Р. М. & К. BIELER, 2004, Interna- tional Marine Bivalve Workshop 2002: Intro- duction and Summary. In: R. BIELER 4 P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 241-248. MORTON, B., 2000, The anatomy of Callocardia hungerfordi (Bivalvia: Veneridae) and the ori- gin of its shell camouflage. Journal of Mollus- can Studies, 66: 21-31. OHNO, T., 1996, Intra-periostracal calcified needles of the bivalve family Veneridae. Bulle- tin de l’Institut Océanographique, Monaco, No. Spécial 14, 4: 305-314. RICHARDSON, C.A., 1993, Bivalve shells: chro- nometers of environmental change. Pp. 419- 435, in: B. MORTON, ed., The marine biology of the South China Sea. Hong Kong University Press, Hong Kong. RICHARDSON, С. A., 2001, Molluscs аз аг- chives of environmental change. Oceanogra- phy and Marine Biology: an Annual Review, 39: 103-164. STANLEY, S. M., 1970, Relation of shell form to life habits of the Bivalvia (Mollusca). Geologi- cal Society of America Memoir, 125: 1-296. STANLEY, S. M., 1981, Infaunal survival: alter- native functions of shell ornamentation in the Bivalvia (Mollusca). Paleobiology, 7: 384-393. TAYLOR, J. D. 8 Е.А. GLOVER, 1997, А спето- symbiotic lucinid bivalve (Bivalvia: Lucinoidea) with periostracal pipes: functional morphology and description of a new genus and species. Pp. 335-361, in: F. E. WELLS, ed., The marine flora and fauna ofthe Houtman Abrolhos, West- ern Australia, Western Australian Museum, Perth. TAYLOR, y. D. 8 E. A. GLOVER, 2000, Func- tional anatomy, chemosymbiosis and evolution of the Lucinida, in: E. M. HARPER, J. D. TAYLOR 8 J. A. CRAME, eds., The evolutionary biology of the Bivalvia. Geological Society of London Special Publication, 177: 207-225. TAYLOR, J. D. 8 Е. А. GLOVER, 2002, Lamello- lucina: a new genus of lucinid bivalve with four new species from the Indo-West Pacific. Jour- nal of Conchology, 37: 317-336. TAYLOR, J. В., Е. A. GLOVERIS С. В. BRAITHWAITE, 1999, Bivalves with “concrete overcoats', Granicorium and Samarangia. Acta Zoologica, 80: 285-300. TAYLOR, J. D., W. J. KENNEDY & А. HALL, 1973, The shell structure and mineralogy of the Bivalvia. Il. Lucinacea- Clavagellacea, Conclu- sions. Bulletin of the British Museum (Natural History) Zoology Series, 22: 225-294. Revised ms. accepted 31 October 2003 MALACOLOGIA, 2004, 46(2): 295-307 PREDATOR-PREY INTERACTIONS BETWEEN CHIONE ELEVATA (BIVALVIA: CHIONINAE) AND NATICARIUS CANRENA (GASTROPODA: NATICIDAE) IN THE FLORIDA KEYS, U.S.A. Brian Morton* & Martina Knapp? ABSTRACT Field samples of Chione elevata (Veneridae) were collected from two sites (10 x 25 x 25 cm quadrats) on the Atlantic coast of the Florida Keys at Long Key State Park, Long Key, and Anne's Beach, Lower Matecumbe Key. Shells were divided into living, empty (non- drilled) and drilled categories and measured along their greatest lengths. The shells of other bivalve species were also so separated. Generally, the C. elevata samples from the four sites were similar to each other, except with regard to the numbers of drill holes, that is, with a significantly higher level of predation at Long Key. The shells of nine species of bivalves, but especially Chione elevata, were drilled in an approximately stereotypical manner, that is, equally on both valves, usually postero-dor- sally. Most drillings were successful, and the predator 1$ believed to be Naticarius canrena - this being the only naticid shown experimentally to be capable of drilling C. elevata in the postero-dorsal location. Chione elevata possesses an array of potential anti-predator shell defences, notably elevated shell lamellae, similar to those of Placamen calophyllum in the Indo-West Pacific. Placamen calophyllum is immune from naticid attack except by species of Polinices, which drill it at the valve margin. Naticarius canrena attacks C. elevata by drilling between the shell lamella, at the interspaces, postero-dorsally. In this the thickest region of the bivalve shell, the few incomplete drill holes suggest that N. canrena is highly successful. There were very few marginal drill holes. Mid-ventrally, C. elevata possesses large, paired pallial glands, and it is possible that this is an anti-predator device inhibiting marginal drilling. Naticarius canrena, therefore, has to drill this commonest of all intertidal bivalves postero- dorsally but does so by attacking its prey selectively, that is, smaller individuals attacking smaller bivalves so that drill hole size is always narrower than interlamellar distance. Key words: Predator-prey interactions, Naticarius, Chione, Florida Keys, drill hole posi- tion, shell defenses. INTRODUCTION That many naticid gastropods drill holes in bivalve and some gastropod shells to access the tissues inside is well documented (Taylor, 1998) and has led to the concept of an “arms race” between predator and prey as the former strives to overcome the defences of the latter and the bivalve seeks to improve its ability to defend itself against the gastropod (Vermeij, 1978, 1980). Species of Austroginella (Marginellidae) drill holes in their prey (Pon- der & Taylor, 1992), and it has been shown recently that juvenile Nassarius festivus (Nassariidae) can also drill the shells of con- specifics (Morton & Chan, 1997). There are, however, two major groups of intertidal drill- ing predatory gastropods: representatives of the Muricidae, generally, on rocky shores and species of Naticidae on soft ones. On Malay- sian shores, Anadara granosa is drilled by both Natica maculosa (Naticidae) and Thais carinifera (Muricidae) (Broom, 1981). Muricids drill vertically straight holes in shells of their prey, naticids countersunk ones, and both at- tack in a stereotypical way to gain access. Thus, Arua (1989) was able to show for an Eocene molluscan community from Nigeria that the most common prey of naticids was the epifaunal, strongly ornamented bivalve The Swire Institute of Marine Science, The University of Hong Kong, Hong Kong, China; present address: Department of Aquatic Zoology, The Western Australian Museum, Perth, Western Australia; prof_bsmorton@hotmail.com Institute of Zoology, University of Vienna, Althanstrasse 14, A-1090 Vienna, Austria 296 MORTON & КМАРР Tivelina newtoni, whereas the principal prey of muricids was the epifaunal, but also strongly ornamented gastropod Bonellitia amekiensis. Today, on a southwestern Australian rocky shore, the muricid Lepsiella flindersi attacks the mussel Xenostrobus pulex in a stereotypi- cal manner on the posterodorsal side of left and right valves equally (Morton, 1999). Simi- larly, in a southwestern Australian marsh, Lepsiella vinosa attacks another mussel, X. inconstans, also in a stereotypical manner posteriorly (Morton, 2004). In the Azores, Thais haemastoma drills the intertidal mussel Trichomusculus semigranatus only atthe pos- terior margin (Morton, 1995). On Hong Kong shores, Morula musiva attacks an array of shallow-water bivalves by drilling at specific locations on each of their shells (Harper & Morton, 1997). Similar observations have been made upon naticids. In Malaysia, Berry (1982) showed that Natica maculosa drills the trochid gastropod Umbonium vestiarium. In Europe, Ansell (1982a, b) demonstrated that the bivalves Venus striatula and Tellina tenuis are drilled by Polinices alderi, whereas on the coast of Massachusetts, Edwards & Huebner, (1977) showed that Mya arenaria is drilled by P. duplicatus. Virtually all species of bivalves are vulner- able to either muricid or naticid attack and there has been much research conducted on the predator/prey relationship between them. This is because both are readily identifiable in the fossil record (Kitchell et al., 1981), drilled shells are obvious in modern field-collected bivalve assemblages, and the stereotypical feeding behaviour of the predator makes both it and its prey eminently suitable experimental animals. For example, shell height/length pa- rameters can be correlated readily with tissue weights. Thus, Taylor (1970), examined the feeding habits of predatory boring gastropods in a Tertiary mollusc assemblage, and Negus (1975) was able to analyse the drill holes made by Natica catena in a collection of Donax vittatus shells. Morton (1990a) showed that in the Azores, the commonest shallow subtidal bivalve, Ervilia castanea, was drilled by Polinices alderi in a stereotypical, posterior, position. Ansell & Morton (1987) demonstrated that various species of Indo-West Pacific naticids attacked different species of bivalve prey in a different sequence according to in- tra-specific size, inter- and intra-specific shell thicknesses and at different locations on the shells. In an elegant series of laboratory stud- ies, Ansell (1982a, b, c) examined the ener- getic relationship between P. alderi (and P. catena) and its bivalve prey. With respect to the Florida Keys, Roopnarine & Beussink (1999) suggested that during a Pliocene-Pleistocene extinction, Chione erosa Dall, 1903, was replaced by C. cancellata Linnaeus, 1767. When this occurred, there was an increase in the size of Chione selected by naticid predators, and C. cancellata re- sponded to this by significantly increasing rela- tive shell thickness. Mikkelsen & Bieler (2000) identified three species of Chione from the Florida Keys, with C. cancellata being the most widespread. The paper on western Atlantic Chione by Roopnarine & Vermeij (2000), how- ever, suggests that C. cancellata is a Caribbean species, whereas the Floridian species is C. elevata (Say, 1822). There is, thus, a large lit- erature on what was considered, in Florida and north to North Carolina, to be C. cancellata, but which actually refers to C. elevata. Species of Chione (Chioninae) are charac- terized by elevated concentric lamellae on the shell valves. These are produced periodically and may represent a defense against drilling predators. For example, Atlantic species of Chione (Roopnarine & Vermeij, 2000) are very similar to Indo-West Pacific species of Bassina and Placamen (Matsukuma & Yoosukh, 1988), and it has been shown for P. calophyllum (Philippi, 1836) that its shell lamellae protect it from side-drilling naticid predators (Ansell & Morton, 1985; Morton, 1985). Polinices tumidus could, however, attack it by edge drill- ing. Moreover, when the lamellae of P. calophyllum were removed, side-drilling preda- tors could attack it with impunity. This may not be the case in species of Atlantic Chione, how- ever, аз Roopnarine & Beussink (1999, fig. 1) illustrate a shell of Pliocene C. erosa with a drill hole located between adjacent lamellae. During a ten-day period of study in the Florida Keys, one of the most commonly en- countered species of bivalves was Chione elevata. In field samples, moreover, many С. elevata shells were found to have been drilled by a naticid predator. It was decided, there- fore, to undertake a study of this bivalve, its drill holes and try to identify the predator that made them. The overall question that we at- tempted to answer, however, was: like Placamen calophyllum, do the shell lamellae of C. elevata protect it from naticid predation (Morton, 1985)? If not, why not? CHIONE ELEVATA-NATICARIUS CANRENA INTERACTIONS 297 MATERIALS AND METHODS For a short period from 20-29 July 2002, a research visit was paid to the Florida Keys. It was determined to research the anatomy of Chione elevata, undertake an analysis of drill holes in the shells of field-collected samples of bivalves and attempt to determine which predator made them. Anatomy Living individuals of Chione elevata were dissected. A single specimen was also fixed in 5% formalin, decalcified and, following rou- tine histological procedures, sectioned trans- versely at 6 um, and alternate slides stained in Ehrlich’s haematoxylin and eosin and Masson's trichrome. Field Studies On the southeastern coast of the keys, that is, the Atlantic side, two shores were chosen for study after reconnaissance of sites on both sides and when very few Chione elevata were collected from the Gulf of Mexico side. The two chosen shores were Long Key State Park on Long Key and Anne’s Beach on Lower Matecumbe Key, that is, station numbers IMBW-FK-623 and IMBW-FK-638, respec- tively (Mikkelsen & Bieler, 2004: fig.1). At each of two sampling locations at these two sites, ten 25 x 25 cm quadrats were laid haphaz- ardly on the sandy shore covering the full range of tidal heights. A total of, thus, 40 quad- rats was examined by excavating their con- tents to a depth of ~ 10 cm and sieving them through a one millimeter mesh sieve. All living and empty bivalve shells (plus any naticid shells) were sorted from the samples. These were analyzed in the following manner. Shells and living individuals of Chione elevata were measured along their greatest lengths; in the case of empty valves, only right ones were measured. Empty valves were ex- amined for drill holes. Where these were en- countered, the following records were made: (i) which valve; (ii) the location of each drill hole was plotted on a master illustration of the valves; (iii) the outer diameter of the drill hole; and (iv) the distance between adjacent lamel- lae at the drilling location. All other bivalve shells with drill holes were identified to species and the locations of them on the left and right valves similarly plotted on master illustrations. Although several other species of naticids are known to inhabit the shallow waters of the Florida Keys (Lyons & Quinn, 1995; Bieler & Mikkelsen, pers. comm.), the only shells ever collected were those of Naticarius canrena (Linnaeus, 1758). Similarly, but one living individual of this spe- cies was collected from any habitat during the ten-day period of study. Voucher specimens of Chione elevata (108 preserved individuals) and N. canrena (16 shells) have been depos- ited in the collections of the Natural History Museum, London (Reg. No’s: 20030605 and 20030609, respectively). Statistical Analyses The dataset comprising the numbers of liv- ing, empty and drilled shell valves of Chione elevata among the four locations was tested for normality and homogeneity of variances using the Shapiro-Wilk test and Levene sta- tistic, respectively, both at the 0.05 level of sig- nificance before ANOVA. One-way ANOVA's were performed on the dataset to test the null hypothesis that there were no significant dif- ferences in these variables among locations. Where differences were detected, Student's Newman-Keuls (SNK) tests were carried out to identify where the differences lay. For the drilled shells, the relationships between shell length and (i) lamella distance and (ii) drill hole diameter were evaluated by regressions. RESULTS Anatomy Shell: The shell of Chione elevata is illus- trated from various perspectives in Figure 1. In general terms, the shell is approximately equivalve and equilateral, and thus isomyarian, but slightly pointed posteriorly. There is a fine radial sculpture, and each valve is strongly commarginally lamellate (Fig. 1A). In anterior view (Fig. 1B), there is a small an- terior lunule (as defined by Carter, 1967) each valve here interlocking by means of marginal denticles. In dorsal view (Fig. 1C), the poste- rior escutcheon (also as defined by Carter, 1967) is much longer than the lunule and in the case of the former but not the latter, the dorsal edge of the left valve overlaps that of the right. This is also illustrated in posterior view (Fig. 1D). Finally, the shell valve margins 298 МОКТОМ & КМАРР В ar at RIT PAS ASS 4 ET La e ам À IS > ? Y y a 2 BE Я ae E у N De nr AN SA a) © À > M Pas en SARA ST Cs ee ED DIENT. 10 mm FIG. 1. Chione elevata. The shell as seen from: A, the right side; B, anteriorly; C, dorsally; D, poste- riorly and E, ventrally. are interlocked ventrally by the expanding ra- ure 2 from the right side. The shell is, as de- dial ribs (Fig. 1E). The shell valves of C. scribed, radially striate and commarginally elevata are thus very difficult to separate. lamellate. Rays of light purple pattern the gen- | erally white-cream outer surface. Anteriorly, ‚Siphons and Mantle Cavity: A living indi- there is a large digging foot. Posteriorly, there vidual of Chione elevata is illustrated in Fig- is a pair of separated siphons. The exhalant СНОМЕ ELEVATA-NATICARIUS CANRENA INTERACTIONS 299 FIG. 2. Chione elevata. An individual seen from the right side with siphons and foot extended. siphon is conical and a ring of short, thin ten- tacles sub-apically surrounds its aperture. The inhalant siphon is much larger in diameter, but shorter and is fringed apically by a circlet of long siphonal tentacles and papillae. Mid-ven- trally, the mantle possesses a line of papillae and pallial fusions, where they occur, are of the inner folds only, that is, type A (Yonge, 1982). Jones (1979) provided additional details on the gross anatomy of Chione elevata (as C. cancellata) and other chionine species. MORTON & КМАРР 300 FIG. 3. Chione elevata. А transverse section through the left mantle lobe at the margin. H, haemocoel; IMF[1], inner component of the inner mantle fold; IMF[2], outer component of the inner mantle fold; MMF, line; PRM, pallial retractor muscle; PN, pallial nerve; RT, rejection tract; middle mantle fold; OMF, outer mantle fold; P, periostracum; PL, pallial SC, secretory cell; TF, transverse fibres; VC, vacuolated cell. СНОМЕ ELEVATA-NATICARIUS CANRENA INTERACTIONS 301 Mantle Margin: In transverse section, the general surface of the mantle comprises epi- thelia, cross-connected by transverse fibers (Fig. 3, TF) and enclosing a capacious hae- mocoel (H). Jones (1979, fig. 21) provided a simple illustration of a transverse section through the mantle margin of Chione elevata (as C. cancellata). Here, that of C. elevata is shown to be very large and complex. It com- prises the usual three folds (Yonge, 1982). The outer fold (OMF) 1$ large, and its inner sur- face secretes a very thin periostracum (P) against the template of the outer surface of the middle mantle fold (MMF). The template is a long thin sheet of tissue arising from the major element of the middle fold. The inner fold is divided into two components, a smaller inner (IMF[1]) and a larger outer (IMF[2]) that also gives rise to the mantle papillae which fringe the mantle margin especially ventrally. The greatest component of the mantle mar- gin of Chione elevata is a large gland com- prising elongate vacuolated cells (VC) that lie beneath the inner epithelium and stain a light red in Masson's trichrome: apically, such cells are actively secretory (SC). The gland does not therefore appear to be secreting mucus to bind up pseudofaeces: it is too large for such a purpose and the habitat of coarse coral sand would not necessitate such rejectory capabili- ties, but otherwise its function is unknown. The illustrations of the sectioned ventral mantle margins of C. elevata (as C. cancellata) and C. undatella (Sowerby, 1835) by Jones (1979, figs. 21, 24) do not identify whether such a gland is present although they appear swol- len as described herein. Field Studies Overall, 2.3 (+ 2.4) living, 13.7 (+ 9.6) empty shells and 4.5 (+ 4.40) drilled right valves of Chione elevata were encountered per quad- rat (Table 1). The numbers of living, empty and drilled valves differed among locations, how- ever (Table 2A, p < 0.05). Results of a Student-Newman-Keuls test (Table 3) showed that the two Anne's Beach samples were similar to Long Key State Park A, but also that the two Long Key State Park locations were similar to Anne's Beach A in terms of living Chione elevata. Similar results were obtained for the empty shells, that is, the two Anne's Beach sites were similar, as were the two Long Key State Park locations but these were also similar to Anne's Beach B. Only with regard to the drilled valves was a clear intersite difference obtained, that 1$, the two Long Key State Park locations were simi- lar (6.0 and 8.8 drilled valves-quadrat*), as were the Anne's Beach ones (1.4 and 1.7 drilled valves:quadrat*) (Table 1), but also that the two pairs of locations were different from each other (Table 3). That is, there appears to TABLE 1. Results of the statistical analysis of the Chione elevata shell dataset (living, empty and drilled) obtained from the four stations in the Florida Keys (10 x 25 cm x 25 cm quadrats). Shell Type Location Living Anne's Beach Site A Anne's Beach Site B Average Empty Anne's Beach Site A Anne's Beach Site B Average Drilled Anne's Beach Site A Anne's Beach Site B Average Long Key State Park Site A Long Key State Park Site B Long Key State Park Site A Long Key State Park Site B Long Key State Park Site A Long Key State Park Site B Mean Standard Deviation 2.7 1.70 3.4 2.76 2.5 2.99 0.6 0.70 2.3 2.39 8.9 8.21 9.9 8.14 21.4 8.62 14.7 9.04 13.7 9.59 1.4 1.07 1127 VIT 6.0 2.98 8.8 SS 4.5 4.40 302 МОКТОМ & КМАРР TABLE 2. Results of a one-way ANOVA оп the Chione elevata shell dataset (living, empty and drilled) obtained from the four stations in the Florida Keys. Shell Type Comparison df Living Between groups > Within groups 36 Total 39 Empty Between groups 3 Within groups 36 Total 39 Drilled Between groups 3 Within groups 36 Total 39 be a higher level of drilling predation, possibly by Naticarius canrena, at the Long Key State Park than at the Anne's Beach locations. Location of Drill Holes: In addition to Chione elevata, the field-collected bivalve shells with naticid drill holes comprised nine species. Outlines of the shells of these species are il- lustrated in Figure 4. Also illustrated are the positions of the drill holes on both the left (o) and right (+) valves. The two lucinids, Ctena orbiculata (Montagu, 1808) and Lucinisca nassula (Conrad, 1846), were both drilled close to the ventral shell margin. All seven other bivalves, that 1$, the glycymerid Tucetona pectinata (Gmelin, 1791), the carditid Pleuromeris tridentata (Say, 1832), the cardiid Laevicardium mortoni (Conrad, 1830), the venerid Pitar simpsoni (Dall, 1895), and three tellinids — Tellina mera Say, 1834, T. iris Say, 1822, and T. similis J. Sowerby, 1806, were all side drilled in most cases close to the umbones and, again mostly, posterior to them. Figure 5 illustrates diagrammatically the empty right (A) and left (B) valves of Chione elevata with the positions of the total numbers of drill holes on them identified. It is obvious that there is, first, an approximately equal dis- tribution of drill holes (and attempts) between the two valves, that is, right 99, left 98. Sec- ond, most of the drill holes were, again, ap- proximately equally, in terms of the two valves, distributed around the postero-dorsal region of the shell with a few scattered over the rest of the surface. A third important point is that there are very few failed drill holes, that is, two in the right and four in the left valves. Fourth, only a very few of the drill holes were over the shell lamellae, that is, two on the right Mean Square F Significance (р = 0.05) 14.33 2.88 0.049 4.98 325.89 4.50 0.009 72.45 127.29 12.52 0.000 10.34 and four on the left valves and, in the latter case, these were also at the marginal lamellae. Shell Measurements: Figure 6A shows the relationship between shell length and the dis- tance between adjacent lamellae demarcat- ing a drill hole site on the shell of Chione elevata. The relationship is linear, suggesting that the predator is, for a prey of a particular size, choosing а site appropriate for drilling. That is, smaller predators (making smaller drill holes) chose positions on the С. elevata shells where there are smaller interlamellar distances in these younger, smaller bivalve prey. Con- versely, larger predators (making larger drill holes) chose positions on shells where there are larger interlamellar distances, that is, older, larger bivalve prey. This conclusion is substan- tiated in Figure 6B, where it is further shown that drill hole diameter is correlated positively with shell length in С. elevata. That is, smaller predators, making smaller drill holes, attack smaller individuals of C. elevata. In summary, therefore, it would appear from this analysis of naticid-drilled shells that TABLE 3. Student-Newman-Keuls test grouping of the four Florida Keys locations (1 = Long Key State Park Site A, 2 = Long Key State Park Site B, 3 = Anne’s Beach Site A, 4 = Anne’s Beach Site B) into subsets in terms of the mean num- ber of living, empty and drilled shells of Chione elevata in an ascending order. Shell Type Subsets Living 4=3=1<3=1=2 Empty 1=2=4<4=3 Drilled 1=2<3=4 СНОМЕ ELEVATA-NATICARIUS CANRENA INTERACTIONS 303 À Tucetona Tellina pectinata тега Le œ Pitar Pleuromeris simpsoni tridentata Ce Laevicardium mortoni TEN Chione Naticarius , elevata Tellina canrena iris Ех Ctena orbiculata Tellina % similis о Left valve Lucinisca ® Right valve nassula FIG. 4. The positions of drill holes on the left and right valves of empty shells of nine other species of bivalves collected with the Chione elevata field samples. Also shown is a ventral view of Naticarius canrena and the drill hole one individual of this predator made in the shell of an aquarium-held Chione elevata. Chione elevata (and probably other resident ing; that is, the predator avoids the structur- bivalves) is attacked in a stereotypical man- ally very similar margins and lamellar edges, ner at the posterodorsal margin, and at inter- unlike the situation of Placamen calophyllum lamellar spaces. There were virtually no in the Indo-West Pacific (Ansell & Morton, examples of either marginal or lamellar drill- 1985). 304 МОКТОМ & КМАРР FIG. 5. Chione elevata. The composite pattern of drill holes in the field-collected left and right valves of empty shells (0 non-lamella borehole, + attempted borehole, + lamella borehole). a y = 0.0657x + 0.6413 R? = 0.5595 Inter-lamellar spaces (mm) o 00016 8 & © 0 5 10 15 20 25 30 35 2 . y = 0.0406x + 0.2849 1.8 . e В? = 0.5418 Borehole diameter (mm) 0 5 10 15 20 25 30 35 Shell length (mm) FIG. 6. Chione elevata. The relationships between shell length and A, the distance between two lamellae at the position of an interlamellar drill hole and B, the outer diameter of the drill hole. CHIONE ELEVATA-NATICARIUS CANRENA INTERACTIONS 305 Laboratory Studies Although 16 empty shells of Naticarius canrena were collected from the 40 quadrats, only one living individual was obtained. This was placed in an aquarium with small individu- als of Chione elevata and one drill hole was made, on the right valve postero-dorsally (Fig. 4, С. elevata shell), that 1$, in the position typi- cal of the locations of the field collected bivalves of this species (Fig. 5). No other naticid species (not even shells) was ever col- lected alive. It thus seems possible, at least, that N. canrena made the drill holes on all the field-collected bivalves, but especially C. elevata, collected during the course of this study. DISCUSSION Chione elevata has a shell that, superficially, would appear to offer much protection. Pro- tective characteristics include a tightly fitting margin, with ventrally interlocking ribs, similar interlocking marginal denticles at the antero- dorsal lunule, a marginally overlapping es- cutcheon and internally large, strong cardinal teeth (Jones, 1979). Closed C. elevata are extremely difficult to open, even by the authors! Each adductor muscle also has a large slow component for sustained adduction, and the pallial line is deeply inset within the shell mar- gin. With the commarginal lamellae on the outside of a solid, thick shell, C. elevata would, superficially, appear to be impregnable. Such characters are also possessed by the Indo- West Pacific Placamen calophyllum and are very effective in protecting it from naticid predators (Ansell & Morton, 1985, 1987). Lamellar protection is not afforded to Chione elevata, however, and the only predator iden- tified as being capable of attacking and drill- ing this species at the study sites on the Florida Keys is Naticarius canrena and it does this at the interlamellar spaces. This is the opposite of the situation described for the Indo-West Pacific Placamen calophyllum, which 1$ virtu- ally immune from attack by side-drilling naticids because of the shell lamellae but 1$ vulnerable to the edge-drilling Polinices tumidus, as described by Ansell 8 Morton (1985). It is strange therefore that the lamel- lae of the Atlantic C. elevata do not confer any protection from N. canrena and further strange that there were very few identified attempts to drill the bivalve at the valve margins, espe- cially ventrally. The ventral mantle margin has a huge gland discharging onto the inner but widely open surfaces of the mantle: does the secretion from this constitute a further defen- sive adaptation? In the absence of any knowl- edge about the composition of the secretion from this gland, it is impossible to hypothesize further, but it does not stain for simple mucus. As noted above, it is thought possible that the major predator of Chione elevata on the Atlantic side of the Florida Keys is Naticarius сапгепа. In this study, one such aquarium-held predator did attack the bivalve successfully in the position identified for the great majority of the field-collected empty and drilled valves, that is, postero-dorsally. lt is not known for certain, however, if this species made the drill holes atthe marginal lamellae. However, since very few drillings represented failed attempts, it is clear that if N. canrena is the predator, it has apparently successfully overcome the seemingly impenetrable defenses of C. elevata by attacking it at the posterodorsal interlamel- lar spaces. Younger predators also attack younger prey as with juvenile Polinices duplicatus feeding on Gemma gemma, that is, drill hole diameter is related directly to preda- tor size (Wiltse, 1980). This has also been demonstrated for Polinices lewisii feeding on the littleneck clam, Protothaca staminea by Peitso et al. (1994). A thick shell characteristi- cally protects bivalves from drilling predators, for example, Corbula crassa in Hong Kong (Morton, 1990b), although Borzone (1988) showed that a species of Polinices, as dem- onstrated here for N. canrena, selectively drilled its prey, Venus antiqua, in the thickest region of the shell, that is, umbonally. Although no measurements were taken of C. elevata shell thickness at the drill hole sites, the posterodorsal region is the thickest, and N. canrena has therefore clearly overcome the bivalve’s defenses by selective drilling accord- ing to relative prey size and shell location char- acteristics. The most interesting question derived from this study, however, is: how is it that Indo-Pa- cific species of Bassina and Placamen have evolved strong shell lamellae that protect them from side-drilling species of Natica but have been overcome by edge-boring species of Polinices (Morton, 1985; Ansell & Morton, 1985, 1987), whereas the same shell defences of Chione offer no protection from side-drill- ing Naticarius in the western Atlantic? 306 MORTON & КМАРР ACKNOWLEDGEMENTS The International Marine Bivalve Workshop, held in the Florida Keys from 19-30 July 2002, was funded by the U.S. National Science Foun- dation award DEB-9978119 (to organizers К. Bieler and P. M. Mikkelsen), as part of the Part- nerships in Enhancing Expertise in Taxonomy [РЕЕТ] program. The Bertha LeBus Charitable Trust, the Comer Science and Education Foun- dation, the Field Museum of Natural History, and the American Museum of Natural History provided additional support. We are grateful to Paula Mikkelsen (American Museum of Natu- ral History, New York) and Rüdiger Bieler (Field Museum, Chicago) and their colleagues and students for organizing the workshop and much practical help and kindness during it. MK also thanks the organizers for providing a travel grant to Florida. BM thanks Dr. Katherine Lam (The Swire Institute of Marine Science, The Упмег- sity of Hong Kong) for statistical advice and help. LITERATURE CITED ANSELL, A. D., 1982a, Experimental studies of a benthic predator-prey relationship. |. Feed- ing, growth, and egg collar production in long- term cultures of the gastropod drill Polinices alderi (Forbes) feeding on the bivalve Tellina tenuis da Costa. Journal of Experimental Ma- rine Biology and Ecology, 56: 235-255. ANSELL, A. D., 1982b, Experimental studies of a benthic predator-prey relationship. Il. 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VERMEIJ, 2000, One species becomes two: the case of Chione cancellata, the resurrected C. elevata and a phylogenetic analysis of Chione. Journal of Molluscan Studies, 66: 517-534. TAYLOR, J. D., 1970, Feeding habits of preda- tory gastropods in a Tertiary (Eocene) mollus- can assemblage from the Paris Basin. Palaeontology, 13: 254-260. TAYLOR, J. D., 1998, Understanding biodiversity: adaptive radiations of predatory marine gas- tropods. Pp.187-206, in: B. MORTON, ed., The marine biology of the South China Sea. Pro- ceedings of the Third International Conference on the Marine Biology of the South China Sea, Hong Kong 1996. Hong Kong University Press, Hong Kong. VERMEIJ, С. J., 1978, Biogeography and adap- tation: patterns of marine life. Harvard Univer- sity Press, Cambridge, Massachusetts and London, England. VERMEIJ, С. J., 1980, Drilling predation in Guam: some paleo-ecological implications. Malacologia, 19: 329-334. WILTSE, W. 1., 1980, Predation by juvenile Polinices duplicatus (Say) on Gemma gemma (Totten). Journal of Experimental Marine Biol- ogy and Ecology, 42: 187-199. YONGE, C. M., 1982, Mantle margins with a re- vision of siphonal types in the Bivalvia. Jour- nal of Molluscan Studies, 48: 102-103. Revised ms. accepted 4 February 2004 . u $ MES > u qe o р E 7 ss = =i й o О 0 x e 09% ©. rs ¿MT й | | = "ais rte ine - E o 7 u 7 +: + u | - aA rg ar 5 „` 09 ST a : ES. YA Ls PTE Pe un 6 O ae >. 1 i = = u u he > = a u . vi a The | y aA { + : = | : Lat ai q Be ce: № . u u И a or: te} A a . TV ¿Led CAT 0 у ne: | | | os dur EDR %, SAGE. В + an 2 ne e eat MALACOLOGIA, 2004, 46(2): 309-326 OYSTERS OF THE CONCH REPUBLIC (FLORIDA KEYS): À MOLECULAR PHYLOGENETIC STUDY OF PARAHYOTISSA MCGINTYI, TESKEYOSTREA WEBERI AND OSTREOLA EQUESTRIS Lisa Kirkendale*, Taehwan Lee?, Patrick Baker? & Diarmaid O Foighil?* ABSTRACT We investigated the evolutionary relationships of three species of Florida Keys oysters, Parahyotissa mcgintyi, Teskeyostrea weberi, and Ostreola equestris, using nuclear and mitochondrial (mt) phylogenetic trees. Both 28S (nuclear) and 16$ (mt) ribosomal gene trees consistently recovered a paraphyletic Parahyotissa in which P mcgintyi, the type species, was robustly sister to a tip clade containing P numisma and Hyotissa hyotis. This topology implies that there is no phylogenetic basis for Parahyotissa Harry, 1985, and we therefore recommend that all hyotissinid taxa be returned to the genus Hyotissa Stenzel, 1971. Phylogenetic placement of T. weberi within brooding oyster mt 16S gene trees con- clusively demonstrated that it is a distinct ostreinid lineage, lacking any obvious candidate sister species, and falsified the hypothesis that it is a free-living ecomorph of the sponge commensal Cryptostrea permollis. Population-level mt COI sequence analysis of Ameri- can Ostreola equestris and New Zealand Ostrea aupouria revealed that these two globally disjunct ostreinids, though remarkably close relatives, are reciprocally monophyletic sister taxa. Unlike a large fraction of the Floridian nearshore marine biota, O. equestris shows no evidence of a vicariant phylogenetic break distinguishing Gulf of Mexico and Atlantic popu- lations. Our results imply that its present day Gulf/Atlantic distribution has been achieved by range extension from source Atlantic populations followed by a demographic growth pulse in the new Florida Keys/Gulf of Mexico habitats. Ostreola equestris individuals dis- play an impressive range of shell morphs and coloration, some externally resembling 7. weberi, and we present a plate of genotyped individuals that document this diversity. Key words: Ostreidae, Gryphaeidae, systematics, biogeography, Florida, molecular phylogeny. INTRODUCTION The Florida Keys archipelago extends 362 km SW from the tip of peninsular Florida, sepa- rating Florida Bay from the Straits of Florida. This subtropical island chain represents the exposed surface layer of a much larger car- bonate platform and has a rich bivalve fauna, estimated at approximately 325 species (Mikkelsen & Bieler, 2000). The strategic goal of the International Marine Bivalve Workshop, held at the Keys Marine Laboratory (Long Key) from 19-30 July 2002, was to expand our knowledge of targeted segments of this fauna. We elected to study the local oyster taxa, or at least that fraction accessible by wading, snorkeling and SCUBA diving during our lim- ited sampling window. Although oysters are among the most stud- ied marine invertebrate taxa, their taxonomy and systematics is still fraught with uncer- tainty due to their xenomorphic post-larval growth patterns (Ranson, 1951; Quayle, 1988; Yamaguchi, 1994), relative dearth of tractable anatomical characters, and exten- sive anthropogenic global transfer (Dinamani, 1971; Edwards, 1976; Buroker et al., 1979: Chew, 1990; Carlton & Mann, 1996). Harry’s (1985) ambitious taxonomic revision, based largely on morphology, represents the most recent comprehensive reclassification of liv- ing oysters. Subsequently, a number of de- Department of Malacology, Florida Museum of Natural History, University of Florida, Gainesville, Florida 32611, U.S.A. *Museum of Zoology and Department of Ecology and Evolutionary Biology, University of Michigan, Ann Arbor, Michigan 48109, U.S.A. “Department of Fisheries and Aquatic Sciences, University of Florida, 7922 NW 71st St, Gainesville, Florida 32653, U.S.A. *Corresponding author: diarmaid@umich.edu 310 KIRKENDALE ЕТ AL. tailed paleontological studies (Malchus, 1990; Malchus 8 Aberhan, 1998; Dhondt et al. 1999), together with a steady trickle of mo- lecular phylogenetic analyses (Reeb & Avise, 1990; Littlewood 1994; Banks et al., 1993, 1994; Anderson 8 Adlard, 1994; Hare 8. Avise, 1998; Boudry et al., 1998; O Foighil et al., 1998; Jozefowicz & О Foighil, 1998; О Foighil 8 Taylor, 2000; Campbell, 2000; Steiner & Hammer, 2000; Lam 8 Morton, 2001; Giribet & Wheeler, 2002; Lapegue et al., 2002), have significantly refined our understanding of many aspects of ostreoidean evolution and system- atics. The ostreoidean fauna of the Florida Keys is atypical in that the ecologically dominant cupped oysters of the adjacent Caribbean and Atlantic seaboards are almost completely ab- sent. Although isolated records occur in the Keys (Mikkelsen 8 Bieler, 2000), we did not encounter specimens of either the temperate Crassostrea virginica (Gmelin, 1791), or the tropical C. rhizophorae (Guilding, 1828) (Ostreidae, Crassostreinae). Crassostrea virginica populations are critically dependent on estuarine conditions, absent from the Keys, where salinity variation acts to reduce biotic competition and parasitism (Galtsoff, 1964; Ford 8 Tripp, 1996; Shumway, 1996). Our sampling efforts yielded three distinct oyster groupings. By far the most common were small flat oysters (Ostreidae, Ostreinae), displaying an impressively diverse and over- lapping range of shell morphology and colora- tion. Based on shell phenotype, many of these were readily identifiable as either Ostreola equestris (Say, 1834) or Teskyostrea weberi (Olsson, 1951); however, quite a few individu- als were difficult to place with confidence. During dives, we encountered specimens of the gorgonian-associated Dendostrea frons (Linné, 1758) (Ostreidae, Lophinae) and the equally distinctive Parahyotissa mcgintyi Harry, 1985 (Gryphaeidae, Pycnodonteinae). We focused our efforts on the gryphaeid and flat oysters as they require the most system- atic attention. In particular, we addressed the following four questions. Systematic Placement of Parahyotissa mcgintyi Harry, 1985 Harry (1985) reorganized the gryphaeid (pycnodonteinid) tribe Hyotissini into the mo- notypic Indo-Pacific genus Hyotissa and a new genus Parahyotissa (containing three subgen- era and four species) which includes the tropi- cal Atlantic type species P. (Parahyotissa) mcgintyi, and the Indo-West-Pacific P. (Numismoida) numisma (Lamarck, 1819). He distinguished among the two hyotisssinid gen- era mainly by the relative degree of opening of the left promyal passage: open but reduced in Hyotissa, closed in Parahyotissa. We aimed to test the phylogenetic robustness of this ge- neric reorganization by constructing nuclear and mitochondrial ribosomal gene trees incor- porating these three taxa together with a neopycnodontinid gryphaeid, Neopycnodonte cochlear (Poli, 1795), that is sister to the Hyotissini (O Foighil & Taylor, 2000). Phylogenetic Status of Teskyostrea weberi Olsson (1951) considered Ostrea weberi to be the most distinctive regional species of oyster, and designated Key West as its type locatity. Harry (1985) supported its taxonomic distinctiveness, placing it in a monotypic new genus, Teskeyostrea. Alternatively, Abbott (1974) regarded T. weberi as a free-living ecophenotype, and junior synonym, of the sponge commensal Cryptostrea permollis (G. В. Sowerby II, 1871), and this taxonomic in- terpretation has been largely followed in the subsequent literature (Carriker & Gaffney, 1996). Cryptostrea permollis is recorded from the northeastern Gulf of Mexico and off North Carolina (Harry, 1985), and we did not encoun- ter it in the Florida Keys. There are multiple records of C. permollis in the Florida Keys (Mikkelsen & Bieler, 2000); however, these refer to free-living, 7. weberi (К. Bieler, pers. comm.). Jozefowicz & O Foighil (1998) incor- porated, for comparative purposes, Keys specimens they identified as Т. weberi in their molecular study of Southern Hemisphere flat oysters. However, they were unaware that the range of shell ecomorphs produced by another Keys ostreinid, Ostreola equestris, overlaps with that of 7. weberi. Subsequent unpublished work by one of the authors (P. Baker) showed conclusively that the “Т. weber” specimens sequenced by Jozefowicz & О Foighil (1998) were actually O. equestris. The phylogenetic placement of Т. weberi therefore still remains to be established. We revisited this issue by generating mitochondrial genotypes - large ribosomal subunit (16$) — from authentic 7. иерей and incorporating them, together with C. permollis and O. equestris genotypes, into a phylogenetic analysis of brooding oysters. FLORIDA KEYS OYSTERS 311 Biogeographic Relationships of Ostreola eques- tris and Ostrea aupouria (Dinamani & Beu, 1981) Jozefowicz & О Foighil (1998) uncovered a number of unexpectedly close phylogenetic re- lationships among geographically disjunct ostreinid taxa. Their Keys Ostreola equestris samples (misidentified as Teskeyostrea weberi, see above) differed from specimens of the New Zealand O. aupouria by as little as a single trans- version in their mt 16$ large subunit ribosomal gene fragments. We aimed to revisit this sur- prising biogeographic pairing by utilizing Cyto- chrome Oxidase | (COI), a faster-evolving mt gene fragment more useful in resolving oyster tip taxa (O Foighil et al., 1998), and by incorpo- rating samples of O. equestris spanning the well- defined Gulf/Atlantic marine biogeographic break in southeastern Florida (Avise, 1992, 2000; Cunningham & Collins, 1994). In the ab- sence of post-separation gene flow, the process of lineage sorting is expected to sequentially lead newly formed daughter populations from initial polyphyly, to paraphyly, and ultimately to reciprocal monophyly (Avise, 2000). We were interested in establishing whether these disjunct New Zealand/American populations were recip- rocally monophyletic, or if one was a recent founder of the other. Another objective was to determine how the aupourialequestris genetic disjunction scaled relative to the anticipated Gulf/Atlantic break in O. equestris. Two hypo- thetical topologies, each containing an O. equestris Gulf/Atlantic disjunction, are presented as exemplars in Figure 1. There are of course many other topological possibilities. Shell Phenotype Variation in Ostreola equestris Ostreola equestris is commonly known as the “crested” oyster and, as its informal name im- plies, it is described as having a shell with raised crenulated margins (Abbott, 1974). We encoun- tered this morph in intertidal Keys habitat; how- ever, subtidal individuals, genotyped in this study for mt markers, were usually cemented to the substratum along their entire left valves, yield- ing a very thin, contour-hugging, morph that ex- hibited a wide variety of coloration and sculptural texture, some of which closely approximated the Teskeyostrea weberi phenotype (Olsson, 1951; Harry, 1985). Employing genotyped individuals only, we aimed to give a photographic summary of the impressive range of shell phenotypes dis- played by our samples of this species. G NZ At NZ a G At NZ Fe b FIG. 1. Two exemplary unrooted mitochondrial tree topologies predicted by distinct hypotheses of historical relationships among geographically disjunct sister populations of New Zealand (Ostrea aupouria) and American (Ostreola equestris) ostreinids. Both hypotheses assume a priori that О. equestris has undergone clado- genesis into distinct Atlantic (At) and Gulf (G) lineages, a well-documented pattern among coastal Floridian marine taxa (Avise, 1992, 2000; Cunningham & Collins, 1994). There are of course many other hypothetical topologies that could be entertained. a, O. aupouria (NZ) represents a recent founder population of Gulf O. equestris (G) and genotypes of the former are predicted to nest within a Gulf tip clade; b, O. aupouria (NZ) has experienced a distinct evolutionary history that predates the origin of the Gulf/Atlantic disjunction in O. equestris and all three groupings are predicted to be recipro- cally monophyletic with the stem branch lead- ing to O. aupouria (NZ) being the most pronounced. MATERIALS AND METHODS A summary of sampling locations and of voucher specimen information 1$ outlined in Table 1, and specific sampling details for Flo- ridian taxa are given in the following para- graphs. For specimens collected in the Florida Keys, all collections were made ма snorkel- ing in depths from 1-5 m, except collections from IMBW-FK-650 where SCUBA was used to sample specimens from roughly 30 т. These specimens were preserved in 95% de- natured alcohol and then transferred to 95% non-denatured alcohol upon return to the De- partment of Malacology а the Florida Museum of Natural History. Specimens collected else- where were sampled from shore and pre- served in > 70% ethanol. 312 KIRKENDALE ET AL. TABLE 1. Species identification and sampling locality data, together with voucher specimen information. UMMZ and FLMNH numbers respectively refer to the voucher specimen catalog numbers of the Mollusk Division, University of Michigan Museum of Zoology, and the Department of Malacology, Florida Museum of Natural History. See Mikkelsen & Bieler (2004) for specific details concerning the International Marine Bivalve Workshop (IMBW-FL) sampling stations. Taxa Location Family Gryphaeidae Subfamily Pycnodonteinae Parahyotissa mcgintyi IMBW-FK-650 Parahyotissa numisma Guam Hyotissa hyotis Guam Neopycnodonte cochlear Maui, Hawaii Family Ostreidae Subfamily Ostreinae Teskeyostrea weberi IMBW-FK-645 Ostreola equestris IMBW-FK-629 Ostreola equestris IMBW-FK-644 Ostreola equestris IMBW-FK-649 Ostreola equestris Ostreola equestris Ostrea aupouria Cryptostrea permollis Subfamily Crassostreinae Crassostrea virginica Crassostrea virginica Parahyotissa mcgintyi Numerous specimens of the gryphaeid Parahyotissa mcgintyi were sampled (by L. Kirkendale and G. Steiner) from the superstruc- ture epibenthos of the sunken vessel Thunder- bolt (IMBW-FK-650; Table 1) — apparently this species' first record from the Florida Keys (Mikkelsen 8 Bieler, 2000). Parahyotissa mcgintyi is easily distinguished from other re- gional oysters Бу its frequently plicated shell margins, absence of clasper spines, typically pycnodonteinid vesicular shell structure (Fig. 2), and presence (in live adult specimens) of a bright orange pigment in ovarian tissue (Harry, 1985). In order to test Harry's (1985) taxonomic rearrangement of the Hyotissini, we sequenced a 941nt (post-alignment length) fragment of nuclear 28$ rDNA, added it to О Foighil & Taylor's (2000) homologous 28$ ostreoidean matrix, and analyzed the resulting dataset uti- lizing pterioid outgroups (Giribet & Distel, 2003). A complementary gryphaeid mt 16$ rDNA data set was constructed and then phy- logenetically analyzed using Neopycnodonte cochlear, a sister taxon to the Hyotissini (Ó roighil & Taylor, 2000), as an outgroup. Skidaway River, Georgia Cedar Key, Florida Hauraki Gulf, New Zealand 12 Panacea, Florida Skidaway River, Georgia 3 Panacea, Florida # of individuals sequenced Catalog # UMMZ 300092 UMMZ 265996 UMMZ 265995 UMMZ 265997 4 FLMNH 298644 E FLMNH 298643 1 FLMNH 298645 1 FLMNH 298640 dl UMMZ 300093 10 UMMIZ 300094 UMMZ 255404 2 UMMZ 255410 UMMZ 300095 2 UMMZ 300096 Teskeyostrea weberi Specimens of Teskeyostrea weberi were re- covered (by L. Kirkendale) from one of our sampling sites: the ocean-side shore of Grassy Key (IMBW-FK-645; Table 1), where it was locally abundant attached to the underside of large boulders at depths of 1-3 m. Positive identification of this species was made not only on the basis of its shell characters — flat, thin apricot-colored shell ornamented with fine ra- dial ribbing and thin lamellose extensions (Olsson, 1951; Harry, 1985) — but also on its lack of an anal appendage, a prominent ana- tomical feature of Ostreola equestris (Harry, 1985). To place Teskeyostrea weberi phylo- genetically, we generated mt 16$ sequences for four individuals, yielding two haplotypes, which were incorporated into Jozefowicz & O Foighil's (1998) brooding oyster 16$ matrix. This matrix was further supplemented by 16$ sequences (two haplotypes) generated from 11 Florida Keys Ostreola equestris specimens sampled from three locations in the Florida Keys (Table 1). These latter specimens collec- tively displayed a wide variety of shell morphs, including Т. weberi look-alikes, but exhibited FLORIDA KEYS OYSTERS 313 FIG. 2. Views of gross shell morphologies of adult and juvenile Parahyotissa mcgintyi specimens sampled from IMBW-FK-650. a, internal view of the left valve of an adult preserved in 95% ethanol after shucking (Note the prominent plication of the ventral valve margin); b, external view of the right valve of specimen depicted in 2a (Note the heavy fouling which obscures the valve outline); c, detail of anterio-dorsal inner edge of left valve of adult (see boxed area in 2a) showing the distinctive vesicular substructure characteristic of pycnodonteinid gryphaeids (Harry, 1985); d, external view of intact juvenile (note straight hinge line, flattened D-shaped profile and the vesicular substructure evident in abraded surface areas). distinct anal appendages (mainly digitform, Ostreola equestris some more cardiform in outline). Finally, we added to the single available 16S haplotype In order to more fully resolve the phyloge- of the sponge commensal Cryptostrea netic relationships of these geographically dis- permollis by sequencing two additional speci- junct, polytomous (at least for 16$, Fig. 4), mens (Table 1). New Zealand/American tip taxa, a mt COI 314 KIRKENDALE ЕТ AL. gene fragment (626 nt) data set was gener- ated for a total of 44 individual oysters. Twelve New Zealand Ostrea aupouria — reliably dis- tinguished by their possession of an anal ap- pendage (Dinamani 8 Beu, 1981) from the co-occurring Ostrea chilensis (Philippi, 1844) — were sequenced, yielding 6 haplotypes, as were 32 Ostreola equestris specimens which collectively contained 15 haplotypes. We were interested in establishing if Ostreola equestris exhibits a regional Gulf/Atlantic ge- netic break in southeastern Florida in common with many other co-occurring nearshore ma- rine taxa (Avise, 1992, 2000; Cunningham & Collins, 1994) and, if so, how it might scale relative to the equestris/aupouria disjunction. In addition to Florida Keys specimens (N = 11, six haplotypes), our 32 O. equestris individu- als sequenced for СО! also included speci- mens from the northeastern Gulf of Mexico (Cedar Key, N = 11, seven haplotypes) and from the Atlantic coast of Georgia (Skidaway River estuary, N = 10, six haplotypes). To pro- vide a phylogeographic yardstick, we also generated homologous COI sequences (598 nt) for a token number of replicate Gulf (Pana- cea, Florida Panhandle, N = 2, one haplotype) and Atlantic (Skidaway River, N = 3, 2 haplo- types) specimens of the cupped oyster Crassostrea virginica. This ecologically domi- nant regional oyster species displays a well- characterized Gulf/Atlantic mt disjunction centered on southeastern Florida (Reeb & Avise, 1990). Molecular Methods Specimens utilized in this study were pro- cessed for molecular characterization either at the University of Florida (by L. Kirkendale) or the University of Michigan (by T. Lee). As a result, there were some minor methodologi- cal distinctions associated with DNA template preparation and PCR amplification as re- ferred to below. All novel DNA sequences were generated at the University of Michigan’s DNA Sequencing Core and have been deposited in GenBank (Accession #s AY376596-AY376635). Genomic extractions and amplifications of flat oyster samples collected during the Florida Keys Bivalve Workshop were conducted by L. Kirkendale at the Florida Museum of Natu- ral History Molecular Phylogenetics Lab at the University of Florida (UF). Total genomic DNA was obtained from ethanol-preserved mantle tissue using modifications of standard proto- cols. Roughly 20-30 mg of tissue was finely cut, ground with a mortar and pestle and placed in 750 uL of DNAzol with 5-20 uL of 5— 20 mg/ml proteinase K (Molecular Research Center, Inc.). Tissue was gently shaken over- night on an orbital shaker and following three rounds of ethanol extraction and centrifuga- tion, the pellet was eluted in 100 mL ddH20 (for further details of DNAzol extraction pro- cedure, refer to Chomczynski et al. 1997). Universal primers were used to amplify 16S and СО! gene regions sequenced from the above- mentioned samples and were as follows: 16Sar 5’-CGCCTGTTTATCAAAAACAT-3’ and 16Sbr 5’-GCCGGTCTGAACTCAGATCACGT-3’ (Kessing et al. 1989) and LCO1490 5’- GGTCAACAAATCATAAAGATATTGG-3' and HCO2198 5’-ТАААСТТСАСССТСАССА AAAAATCA-3’ (Folmer et al., 1994). Reactions included 1uL of genomic DNA template and 31.8 uL ddH20, 5 pL of 10X TAQ PCR buffer (Perkin Elmer), 5 uL of dNTPS (10 mM stock), 2 pL of each primer (10 uM stock), 3 pL of MgCl2 solution (25 mM stock, Perkin Elmer) and 0.2 uL TAQ enzyme (Perkin Elmer). Re- actions for 16S were initially denatured at 96°C for 150 sec, followed by 37 cycles of 94°C for 40 sec, 52°C for 35 sec, and 72°C for 60 sec. Reactions for СО! were handled similarly ex- cept that the initial denaturation step was at 95°C for 120 sec and that 40 cycles of ampli- fication were employed with a 40°C annealing temperature. All amplifications were run with positive and negative (no template) controls. PCR products were visualized by electro- phoresis on 1% TBE agarose gels, stained with ethidium bromide solution and photo- documented. Successful PCR products were cleaned for cycle sequencing using Wizard PCR Preps (Promega), following described protocols. Verification of the cleaned PCR product occurred in the same manner as for initial PCR products. Ostreola equestris samples from Cedar Key were extracted at UF, as above, but amplified at the Museum of Zoology, University of Michigan (UMMZ), by T. Lee, along with Skidaway River O. equestris samples, using specifically de- signed СО! primers: 5'-GATATTGGACGGTTTT ATAT-3' and 5’-ССАААААТСААААСААТССТ- 3’ (Lee, unpublished). DNA template prepara- tion methods utilized at the UMMZ are detailed in Lee 8 Ó Foighil (2003). Other target gene fragments amplified at the UMMZ were mt 16S from Cryptostrea permollis and from the four FLORIDA KEYS OYSTERS 315 gryphaeid study species (Table 1) using Kessing et al. (1989) primers, 28$ nuclear ri- bosomal domains 1-3 from Parahyotissa mcgintyi using О Foighil & Taylor's (2000) primer set, and mt COI from Ostrea aupouria, and Crassostrea virginica Gulf (Panacea) and Atlantic (Skidaway River) samples using Folmer et al. (1994) primers. A touchdown (Palumbi, 1996) protocol was used for all UMMZ PCR reactions [after 4 min denaturation at 94°C, the initial annealing temperature of 65°C was decreased by 2°C/cycle (40 sec denaturing at 94°C, 40 sec annealing and 1.5 min extension at 72°С) until the final anneal- ing temperature (45°C for COI, 50°C for 16S and 52°С for 28$) was reached and subse- quently maintained for an additional 30 cycles]. Phylogenetic Methods Initial alignments were constructed using Clustal X (Thompson et al., 1997) using de- fault parameters and then adjusted by eye to minimize mismatches in the ribosomal gene datasets. Phylogenetic analyses were con- ducted on each of six molecular datasets — (1) gryphaeid 2895, (2) gryphaeid 16S, (3) Ostreid/Lophinid 165, (4) Ostrea aupouria/ Ostreola equestris COI, (5) O. equestris COI, and (6) Crassostrea virginica СО! — under the maximum parsimony (MP) optimality criterion using PAUP*4.0b10 (Swofford 2002). While unrooted anayses were performed on COI datasets, the pterioid taxa, Neopycnodonte cochlear, and lophinid taxa were designated as outgroup for gryphaeid 28S, gryphaeid16S and ostreid 16S datasets respectively. MP analyses were performed using heuristic search option with 100 random stepwise ad- ditions and tree bisection-reconnection (TBR) branch-swapping. Gaps were treated as a missing state, character states were treated as unordered and equal weights were as- sumed. Branch support was estimated by bootstrapping (Felsenstein, 1985) (500 repli- cates, heuristic searches, 10 random additions each) and decay indices (Bremer, 1994), gen- erated in TreeRot (Sorenson, 1996). We wished to construct unrooted gene net- works for three COI datasets (Ostreola equestris and O. aupouria; O. equestris alone, Crassostrea virginica alone) and took a Maxi- mum likelihood (ML) approach because two of the three (O. equestris and O. aupouria; O. equestris alone) produced multiple equally most parsimonious trees. A MP tree was first used to estimate the log-likelihood scores us- ing PAUP*. The best-fit ML model for each partition was then determined by hierarchical likelihood ratio tests (hLRTs) using Modeltest 3.06 (Posada & Crandall, 1998). ML analyses were conducted using a heuristic search op- tion in which the parameter values under the best-fit model were fixed and a MP tree was used as a Starting point for TBR branch swap- ping. The K81uf model [K81 model (Kimura, 1981) with unequal base frequencies] + Г [gamma-distributed heterogeneity of the sub- stitution rate across sites (Yang, 1994)] was chosen as the best-fit model for the combined Ostreola equestris and O. aupouria dataset. For the O. equestris and С. virginica СО! datasets, the respective best-fit models chosen were K81uf and HKY (Hasegawa et al., 1985). RESULTS Systematic Placement of Parahyotissa mcgintyi Figure 3 shows the most parsimonious gene tree obtained when a Р mcgintyi 28$ geno- type was added to, and analyzed with, Ó Foighil & Taylor's (2000) ostreoidean 28$ dataset. We obtained a paraphyletic Parahyotissa and a robust terminal sister re- lationship for the two Pacific Hyotissini: P. numisma and Hyotissa hyotis. A congruent topology was recovered when the 16$ se- quences for the four gryphaeid taxa at our dis- posal (Table 1) were subjected to a maximum parsimony analysis (Fig. 3). The earlier study (О Foighil & Taylor, 2000) should be consulted for a detailed discussion of the ostreid clade topology. Phylogenetic Status of Teskyostrea weberi Figure 4 shows the strict consensus topol- ogy о the 54 most parsimonious trees obtained when the brooding oyster 16S matrix was ana- lyzed using the lophine taxa as outgroups. Major elements of the topology are congruent with that obtained, and discussed at length, in an earlier study (Jozefowicz & O Foighil, 1998) and will not be reiterated here. The salient features of the topology concern the relative placement of the three Floridian flat oyster taxa (labeled in bold text). All three occur in distinct, well-sup- ported terminal clades: Teskeyostrea weberion its own, Ostreola equestris in a terminal polytomy with the New Zealand Ostrea 316 KIRKENDALE ET AL. Isognomon alatus Pinctada imbricata Gryphaeidae 285 165 Neopycnodonte cochlear 100 28 Parahyotissa macgintyi Hyotissa hyotis Parahyotissa mumisma 100 / 60 100 Crassostrea rhizophorae ) С. virginica — 10 changes Striostrea margaritacea 100 66 31 3 100 Saccostrea commercialis 18 Saccostrea cucullata 64 100 Crassostrea ariakensis 14 C. gigas Ostrea chilensis 99 99 р 8 О. angasi 12 Е Ostreidae О. edulis 78 5 100 ©. conchaphila 10 О. puelchana 1 O. denselamellosa 1 O. algoensis 5 Dendostrea frons 3 D. folium 14 Alectryonella plicatula 1 (| Lopha cristagalli A FIG. 3. The single most parsimonious tree (809 steps, Cl = 0.668, RI = 0.779) obtained by heuristic unweighted searches of 28S genotypes for 22 oyster taxa, including 4 gryphaeid species, with the two pterioids, Pinctada and /sognomon, designated as outgroups. See also the juxtaposed single most parsimonious tree (173 steps, Cl = 0.948, RI = 0.710) obtained by heuristic unweighted searches of gryphaeid mt 16S genotypes, in which Neopycnodonte cochlear was the designated outgroup. Numbers above the branches represent bootstrap values (> 50) and numbers below indicate decay index values. aupouria, Cryptostrea permollis in a terminal frons) polytomy captures the branch support- polytomy with the Argentine Ostrea puelchana. ing the T. weberi tip clade (Fig. 4), thereby ob- A prominent basal ostreinid (+ Dendostrea scuring its sister relationships. FLORIDA KEYS OYSTERS ЭЙ Ostrea aupouria 1 O. aupouria 2 > О. aupouria 3 Ostreola equestris 1 (N=10) Ostreola equestris 2 63 Cryptostrea permollis 1 (N=2) = I 1 С. permollis 2 3 С. permollis 3 1 Ostrea puelchana O. denselamellosa Ostreola conchaphila 100 Teskeyostrea weberi 1 (N=3) T. weberi 2 Ostrea angasi 1 95 79 6 5 О. angasi 2 ; О. angasi 3 au О. edulis 1 75 О. edulis 2 3 О. edulis 3 O. chilensis 100 Ostrea algoensis 1 8 О. algoensis 2 1 58 Dendostrea frons 1 100 1 D. frons 2 D. frons 3 100 D. folium 1 66 6 D. folium 2 Alectryonella plicatula Lopha cristagalli FIG. 4. Strict consensus of 54 equally most parsimonious trees (174 steps, Cl = 0.6379, RI = 0.8437) resulting from heuristic unweighted searches of 29 brooding oyster 16S genotypes. The lophine taxa D. folium, D. frons, A. plicatula and L. cristagalli were designated as outgroups. Florida Keys ostreinid taxa are in boldface. Bootstrap values (> 50) and decay indices are shown above and below the branches, respectively. 318 KIRKENDALE ET AL. Biogeographic Relationships of Ostreola equestris and Ostrea aupouria А maximum-likelihood analysis of the com- bined American Ostreola equestris and New Zealand Ostrea aupouria СО! dataset is shown as an unrooted network in Figure 5. New Zealand and American samples were recipro- cally, and robustly, monophyletic. Note however, that the minimum cumulative branch lengths separating members о the two clades was less than that of the maximum branch lengths sepa- rating within-clade O. equestris haplotypes. Figure 6 concerns only American taxa and shows the unrooted maximum-likelihood Gulf/ Atlantic СО! networks for both Ostreola equestris and Crassostrea virginica. The Crassostrea virginica Gulf/Atlantic phylogenetic split, estimated by Reeb & Avise (1990) from whole mt genome RFLP assays at approxi- mately 2.5% divergence, was also recovered from our token sample of Gulf/Atlantic CO | gene fragment sequences (1.8%; 11 substitu- tions over 598 nt). In sharp contrast, no such disjunction was evident in Ostreola equestris. Two haplotypes were found in all three regional populations (Table 2, Fig. 6), including by far the most common mt COI genotype (AFG1; М = 13). This latter mt genotype was numerically predominant in both Gulf (Cedar Key, 6/11) and Florida Keys (5/11) samples of Ostreola equestris, but not among our Atlantic (Skidaway River sample; 2/10) specimens. № we consider the former two samples in isolation, the numeri- cally predominant haplotype was centrally placed and connected to all but one (F4) of the Ostrea aupouria 97 =o) 0.001 substitutions/site Ostreola equestris FIG. 5. Maximum likelihood network (-In = 1073.4044) of Ostrea aupouria (New Zealand) and Ostreola equestris (American) СО! haplotypes. Numbers on the branches are МР bootstrap values. FLORIDA KEYS OYSTERS 319 Skidaway River estuary Panacea Cedar Key Ostreola equestris СОТ Network At@ Atlantic haplotypes (Skidaway River) F4 F © Florida Keys haplotypes G © Gulf haplotypes (Cedar Key) At4 — 0.001 substitution/site Florida Keys Gulf/Atlantic Crassostrea virginica COI disjunction Gulf haplotype (P ) Atlantic haplotype anacea (Skidaway River) FIG. 6. Regional map showing our collection sites for Gulf/Atlantic Ostreola equestris and Crassostrea virginica samples and also the superimposed maximum likelihood networks of the resulting O. equestris (-п = 985.5091) and С. virginica (-т = 878.0842) COI haplotypes. TABLE 2. Relative distribution of the 16 СО! genotypes recovered from the three regional бий Atlantic Ostreola equestris sampling locations. The prefixes At, F, G and AtFG, respectively indicate haplotypes found solely in the Atlantic (Skidaway River) site, solely in the Florida Keys sites, solely in the Gulf (Cedar Key) site, and finally, those recovered from all three sites. See Figure 6 for map showing sampling site locations and the inferred topological relationships among the COI haplotypes. © © N (42) st > — = со Ke) co Su Error Dann бо оо Skidaway River 2. vl 4 1 1 1 - - - = = - = = = = Florida Keys Ge - - - - 1 1 1 1 = > E E 2 4 Cedar Key oa - - - - - - - - 1 1 1 1 1 1 320 KIRKENDALE ЕТ AL. other 10 COI genotypes recovered from the Gulf (Cedar Key) and Florida Keys populations by single substitutions (Fig. 6). Our Atlantic (Skidaway River) sample exhibited a different topological pattern characterized by a relatively extensive network in which the constituent haplotypes showed more pronounced collec- tive phylogenetic definition (Fig. 6). Shell Phenotype Variation in Ostreola equestris An impressive diversity of О. equestris shell phenotypes was recovered from the Florida Keys, and indeed also from single sampling sites, such as the Summerland Key Horse- shoe. Intertidal Horseshoe specimens exhib- ited a shell morphology that is typically associated with this species: gray oval shells with raised crenulated margins (Abbott, 1974). Figure 7a shows a cluster of specimens show- ing this morphology, sampled in this particu- lar case from the Skidaway River study population. Subtidal Florida Keys specimens were generally flatter in appearance, in some cases markedly so, and frequently incorpo- rated a diversity of pigmentation colors and patterns, some of which are presented in Fig- ure 7 (b-f). Exemplars spanning the range of O. equestris shell phenotypes found in the Horseshoe site, and other locations in the Keys, were genotyped using mt (16S and COl) markers and no evidence for genetic differ- entiation was evident among them. A minor- ity of O. equestris individuals displayed shell phenotypes that resembled Teskeyostrea weberi in external appearance: very thin shells with golden brown pigmentation sculptured with fine radial ribbing and lamellose exten- sions (Fig. 7). DISCUSSION Systematic Placement of Parahyotissa mcgintyi Our nuclear and mt ribosomal gene trees consistently recovered a paraphyletic Parahyotissa in which P. mcgintyi, the type species, was robustly sister to a tip clade con- taining Р numisma and Hyotissa hyotis. This topology implies that the character state used by Harry (1985) to distinguish Parahyotissa (closed left promyal passage) is plesiomorphic in extant Hyotissini, rather than a synapomorphy diagnosing a Parahyotissa clade, and that the condition in the monotypic genus Hyotissa (open but reduced left promyal passage) is autapomorphic. Based on avail- able information, there seems to be no phylo- genetic basis for Harry’s Parahyotissa. Future research incorporating P. (Parahyotissa) imbricata (Lamarck, 1819) and P. (Pliohyotissa) quercinus (G. B. Sowerby II, 1871), may un- cover more than one natural (i.e., monophyl- etic) group within the Hyotissini that can be defined by morphological synapomorphies and warrant generic status. Until then, we recom- mend that all hyotissinid taxa be returned to the genus Hyotissa Stenzel, 1971. Phylogenetic Status of Teskeyostrea weberi Our 16S strict consensus tree topology (Fig. 3) conclusively demonstrates that this species is not a free-living ecomorph of the sponge commensal Cryptostrea permollis, as thought by Abbott (1974), but is instead a distinct ostreinid lineage lacking (at present) any ob- vious candidate sister species. Olsson (1951) had proposed the eastern Pacific “Ostrea iridescens”, synonymized with Striostrea prismatica (Gray, 1825) by Harry (1985), as a putative sister species to Т. weberi, based on the similarity of the former’s juvenile shell phe- notype to that of the adult Т. weberi. However, S. prismatica’s taxonomic placement in the cupped oyster subfamily Crassostreinae (Harry, 1985), which is supported by prelimi- nary molecular data (Lee & O Foighil, unpub- lished), rules this out. A more comprehensive sampling of brooding oyster global diversity, including data from genes other than 16S, is required to better resolve T. weberïs phylo- genetic position within the Ostreinae/Lophinae. Although Teskyostrea weberi and Ostreola equestris represent very distinct lineages (Fig. 3), they co-occur in the Florida Keys, and a fraction of latter species resemble Т. weberi in their external appearance (Fig. 7). Fortu- nately, these O. equestris weberi-lookalikes can be distinguished upon dissection by their distinct anal appendage (Harry, 1985), and their relatively larger adductor muscle. Based on our preliminary observations, there may also be ecological and larval settlement dif- ferences among these two ostreinid taxa in the Florida Keys. All of the T. weberi speci- mens we encountered were attached to the underside of rocks (Harry, 1985: fig. 25) in an oceanside location, whereas O. equestris were commonly sampled from the exposed hard surfaces in bayside locations. FLORIDA KEYS OYSTERS FIG. 7. Shell phenotypes. a-f, displayed by genotyped Ostreola equestris sampled from the Skidaway River, Georgia (a, cluster of individuals), and from 2 sites in the Florida Keys (b-e, IMBW-FK-629 from rock surfaces and Г, IMBW-FK-649 epifaunal on Pinna); g, a specimen of Ostrea aupouria, New Zealand sister species of Ostreola equestris (UMMZ 255404); h, a specimen of the sponge commen- sal Cryptostrea permollis from Panacea, Florida Gulf Coast (UMMZ 255410); i and j, individuals of Teskeyostrea weberi sampled from IMBW-FK-645. 322 KIRKENDALE ET AL. Biogeographic Relationships of Ostreola equestris and Ostrea aupouria The СО! gene tree topology (Fig. 5) demon- strates that our respective study populations of New Zealand Ostrea aupouria and Gulf/At- lantic Ostreola equestris are reciprocally monophyletic. This result is sufficient, at least for now, for retention of their respective spe- cific status. Coan et al. (2000) rejected the separation of Ostreola from Ostrea based on morphological characters and the phylogenetic validity of Harry's (1985) Ostreola is question- able given that two of his three constituent species (О. equestris and О. conchaphila) are not sister taxa in our gene trees (Fig. 3). How- ever, a definitive generic designation for equestris and aupouria requires data from the Mediterranean/African-Atlantic type species Ostreola stentina (Payraudeau, 1826). Two lines of evidence indicate that the Ostreola equestris/O. aupouria disjunction re- sults from evolutionarily recent dispersal rather than ancient vicariance. Maximum within- population COI genetic divergence for the Skidaway River sample exceeds the minimum New Zealand/American divergences obtained (Fig. 5). This result implies that the age of the О. equestris/O. aupouria disjunction may be less than the haplotypic lineage sorting time window for the Atlantic population of the O. equestris. Although we do not have а fossil- calibrated lineage-specific clock for any оуз- ter, the well-studied Gulf/Atlantic Crassostrea virginica divergence has been dated, using “conventional calibrations” to approximately 1.2 myr (Reeb & Avise, 1990). Parsimony analysis of our token samples of Gulf/Atlantic С. virginica COI sequences found that they differed by 11 steps (1.83% of the 598 nt frag- ment). The minimum number of substitutions separating the New Zealand and American COI clades in parsimony analyses is six steps (0.95% of the 626 nt fragment). Although the resulting age estimate of 0.625 myr for the О. equestris/O. aupouria disjunction is undoubt- edly crude, it is over two orders of magnitude less than the vicariant separation of New Zealand from Gondwanaland (Weissel & Hayes, 1977). The Ostreola equestris/O. aupouria geo- graphic disjunction is but one of three such cases involving tip taxa in the brooding oyster 16S gene tree (Fig. 3); the other two involve Ostrea edulis/O. angasi and Crypytostrea permollis/Ostrea puelchana and are discussed in Jozefowicz & O Foighil (1998). Although an- thropogenic transoceanic oyster introductions have occurred on numerous occasions (Dinamani, 1971; Edwards, 1976; Buroker et al., 1979; Chew, 1990; Carlton & Mann, 1996; Boudry et al., 1998; O Foighil et al., 1998), we can, with some confidence, rule out such his- toric transfers among the New Zealand/Ameri- can study populations (Fig. 4). This conclusion is based on their lack of shared COI haplotypes and on their reciprocal monophyly (Fig. 5), a phylogenetic relationship that is characteristic of populations that have not experienced evo- lutionary recent gene flow (Avise, 2000). It is possible, however, that such an event may have occurred involving yet-to-be-sampled, geneti- cally differentiated portions of either species’ ranges — according to Harry (1985), O. equestris occurs from North Carolina to Argentina. Genetic Structuring of Gulf/Atlantic Ostreola equestris and Crassostrea virginica Genetic characterization of near-shore ma- rine taxa found on either flank of the Floridian peninsula have revealed cryptic phylogenetic disjunctions among diverse Gulf-Atlantic Caro- linian faunal elements (Saunders et al., 1986; Bert, 1986; Avise et al., 1987; Bert & Harrison, 1988; Dillon & Manzi, 1989; Brown & Wolfingbarger, 1989; Cunningham et al., 1991; Sarver et al., 1992; Cunningham & Collins, 1994; Felder & Staton, 1994; Bert & Arnold, 1995; Duggins et al., 1995; O Foighil et al., 1996; Schizas et al., 1999; Avise, 2000; Collin, 2001, 2002), with by far the most intensively studied exemplar being the American oyster Crassostrea virginica (Reeb & Avise, 1990; Karl & Avise, 1992; McDonald et al., 1996; Hare & Avise, 1996, 1998; Hare et al., 1996). Ostreola equestris occurs in micro-sympatry with C. virginica throughout regional estuar- ies, although prior research has shown that O. equestris tends to be abundant only at high salinity portions of estuaries (Hoese, 1960). Surprisingly, our O. equestris mt СО! data (Fig. 5, Table 2) show that this oyster species dif- fers from C. virginica, and from a large frac- tion of the regional marine biota, in lacking a Gulf/Atlantic mt genetic disjunction. Absence of genetic structuring among Gulf and Atlantic populations is not unique to O. equestris (Gold & Richardson, 1998; Avise, 2000); however, our results indicate that these two co-occur- ring oyster species have experienced signifi- cantly different regional histories. FLORIDA KEYS OYSTERS 323 Another discrepancy among the two oyster mt datasets concerns the relative topological definition of Gulf and Atlantic populations. Beckenbach (1994) performed a cladistic analysis of Reeb 8 Avise's (1990) extensive (N = 232) C. virginica mt RFLP dataset and found that both Gulf and Atlantic populations were dominated by one or two common haplotypes. These occupied central positions in their respective clades and were separated by single steps from a large number of termi- nally positioned rare haplotypes. Our Gulf (Ce- dar Key) and Florida Keys samples of Ostreola equestris showed (either separately or jointly) essentially a similar topology; however, the Atlantic (Skidaway River) sample did not (Fig. 5). In the absence of significant homoplasy, the relative lengths of individual branches within a molecular phylogenetic tree topology are rough proxies for evolutionary time. In this context, it is interesting to note the markedly longer collective branch lengths interconnect- ing Ostreola equestris Atlantic haplotypes rela- tive to the truncated area of the COI topology occupied by Gulf and Florida Keys haplotypes (Fig. 5). This topological distinction is consis- tent with an older evolutionary history for this species in the Atlantic section of its present- day regional range. The compact star-like hap- lotypic topology produced by Gulf (Cedar Key) and Florida Keys COI genotypes (Fig. 5) is characteristic of a population founded more recently by one ancestral type, presumably represented by the numerically predominant, topologically central, well-connected (Castelloe & Templeton, 1994) haplotype AFG1, found in all three study populations. Such a topology is also indicative of popula- tions that have experienced a phase of rapid demographic growth, a process associated with lowered stochastic elimination of novel/ rare lineages (Avise et al., 1984; Slatkin & Hudson, 1991; Moritz, 1996). Our mt СО! data for the three study popula- tions of Ostreola equestris paint a regional his- tory that differs in important respects from that of Crassostrea virginica and also from a large fraction of the local marine biota. The domi- nant regional theme is the presence of a Gulf- Atlantic phylogeographic break characterized by considerable geographic concordance in genetic structuring across diverse faunistic elements (Avise, 2000). This implies a coher- ent spatial patterning of vicariance and sec- ondary contact events. In contrast, О. equestris shows no evidence of a vicariant imprint and our results imply that its present day Gulf/Atlantic distribution has been achieved by range extension from source At- lantic populations followed by a demographic growth pulse in the new Florida Keys/Gulf of Mexico habitats. Shell Phenotype Variation in Ostreola equestris Though forearmed with an awareness of the fabled xenomorphism of oysters, we were sur- prised at the extent to which O. equestris, the most commonly encountered ostreid in the Florida Keys, exhibited a multitude of shell phenotypes — a repertoire far from exhausted by our limited presentation in Figure 6. This facility is also a characteristic of Ostrea aupouria, its New Zealand sister taxon (Dinamani & Beu, 1981). Although genetic characterization is a reliable method for dis- tinguishing co-occurring oyster species with overlapping shell morphs, the presence of a distinct anal appendage in O. equestris (Harry, 1985; but not all are digitiform) and in O. aupouria (Dinamani & Beu, 1981) is also particularily useful in this regard. It is unclear to what degree the phenotypic variation we observed in O. equestris reflects populational allelic diversity and/or local micro-environmen- tal parameters, or what contribution this plas- ticity makes to the local ecological success of this small species — the numerically dominant Florida Keys oyster. ACKNOWLEDGEMENTS Our thanks to Paula Mikkelsen and Rudiger Bieler for organizing the International Marine Bivalve Workshop and to Russ Minton, Louise Crowley and Isabella Kappner for their unstint- ing field assistance. We are grateful to our fel- low workshop attendees who facilitated our efforts in numerous ways and to two anony- mous reviewers whose detailed comments sig- nificantly improved the manuscript. Liath Appleton provided expert help with the photo- graphic plates. Supplementary specimens were kindly provided by Gustav Paulay (Pacific gryphaeids), Andrew Jeffs (Ostrea aupouria) and Randy Walker (Skidaway River Ostreola equestris and Crassostrea virginica). Supported by NSF awards DEB-9978119 to К. Bieler & Р. M. Mikkelsen and by OCE-0099084 to D. O Foighil. Additional support was provided by the 324 KIRKENDALE ЕТ AL. Bertha LeBus Charitable Trust, the Comer Sci- ence & Education Foundation, the Field Mu- seum of Natural History, the American Museum of Natural History and the Mollusk Division of the University of Michigan Museum of Zoology. LITERATURE CITED АВВОТТ, R. Т., 1974, American seashells: the marine Mollusca of the Atlantic and Pacific coasts of North America 2nd Ed. 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ABSTRACT A morphological study of Arcopsis adamsi was made on three populations from contrast- ing biotopes, supralittoral, sublittoral (1 m depth) and offshore (8-10 m depth). Shell morphometrics gave statistically significant differences and these were supported by sub- jective observations on shell sculpture, periostracum. and haemoglobin content. The ob- servations form the core of a discussion on the reliability of morphological characters when defining species taxonomy. It must still be recognised that bivalve growth can strongly affect shell morphology and that sculptural and anatomical differences can be the result of environmental differences. The influence of geographical separation or isolation of popu- lations probably unduly influences taxonomic decision making. Such decision making re- quires consideration of ecological factors and should now be more widely supported by molecular studies at the population level. Key words: Arcopsis, morphology, species taxonomy, growth effects. INTRODUCTION This paper is a result of a taxonomic training workshop organised under the U.S. National Science Foundation PEET Initiative and re- flects on one issue of bivalve taxonomy, namely the reliability of morphological characters at the species level. Current bivalve species taxonomy remains primarily based on morphological characters and species identification relies mostly on shell char- acters. Introducing students to bivalve taxonomy is often problematic because many characters, both shell and anatomical, are subtle parts of a continuum and not discreet. Compounding this are the widespread phenomena of ecophenotypic and geographic clinal variation. Arcopsis adamsi (Dall, 1886) is a relatively common species inhabiting both intertidal and sublittoral biotopes (Abbott, 1974) and was chosen because it or similar taxa had already appeared in morphometric analyses (Marko & Jackson, 2001; Oliver & Cosel, 1992). In these instances, the comparisons were made across isolation barriers or along large geographical ranges. Analysing morphological variation at a much smaller geographical scale has not been done within the Arcoidea. Arcopsis adamsi is a common component of intertidal, sublittoral and offshore mixed rock and sand biotopes to re- corded depths of 10 m. It is generally found attached by a weak byssus to the undersides of rocks resting on sand. It is widespread in the subtropical and tropical western Atlantic and Caribbean Sea ranging from Brazil to North Carolina. In the tropical eastern Atlantic it has a sister species, Arcopsis afra (Gmelin, 1791) (Figs. 8, 9), that occurs along with the closely related Striarca lactea (Linnaeus, 1758) (Figs. 14, 15). In the western Atlantic, in contrast, the genus Striarca is not represented in the Re- cent fauna. A similar situation occurs in the Pacific Panamic region where Arcopsis solida (Sowerby, 1833) is the sole representative of the Striaciinae. Arcopsis species may therefore fill a much broader niche in the Americas and may have radiated into cryptic species. This study examines the biotope range of Arcopsis adamsi in the Florida Keys and com- pares the morphology of populations from dif- ferent biotopes. The primary aim of this paper is to draw attention to issues that students will en- counter when undertaking species-level tax- onomy studies and is intentionally discursive in nature. Many questions about Arcopsis tax- onomy are raised by this study and both authors are aware that the solutions are not provided although methodologies are proposed. ‘Department of Biodiversity & Systematic Biology, National Museum of Wales, Cardiff, United Kingdom; graham.oliver@nmgw.ac.uk “Trondjem Biological Station, Trondjem University of Science & Technology, Norway 328 OLIVER & JÁRNEGREN MATERIALS AND METHODS Materials The materials used in the morphometric analysis are housed in the National Museum of Wales, Cardiff, under the specific numbers cited below. All additional materials are also housed in the National Museum of Wales un- der NMW.Z.2003.075. Arcopsis adamsi was found in four biotopes. (1) Supralittoral crevices Collection site: IMBW-FK-629, 21 & 26-VII-02, “The Horseshoe” bayside of West Summerland Key, MM35, Monroe County, Florida Keys, 24°39.3’N, 81°18.2’W; NMW.Z.2003.075.1. At the “The Horseshoe” site, the supralittoral fringe along the south side of the quarry con- sists in places of highly eroded friable limestones with many cavities and crevices that remained damp throughout the tidal cycle. These cavities were populated by many Arcopsis a few Brachidontes and the occasional Arca imbricata and /sognomon. The presence of the “terres- trial” snails Laemodonta cubensis (Pfeiffer, 1854) and Truncatella pulchella Pfeiffer, 1839, is highly indicative of the supralittoral environment. (2) Shallow sublittoral rubble on sand and lower littoral Collection site: IMBW-FK-629, 21 8 26-VII-02, “The Horseshoe” bayside of West Summerland Key, MM35, Monroe County, Florida Keys, 24°39.3’М, 81°18.2W; NMW.Z.2003.075.2. Other sites: IMBW-FK-622, IMBW-FK-657 On the north side of the Horseshoe quarry the very low vertical face drops onto a sloping face covered in muddy sand and blocks of quarry rubble. Arcopsis was found frequently attached to the undersides of the rubble blocks in 0.5 to 1.5 m water depth. This area was never exposed at low tide and can be consid- ered sublittoral. The upper surfaces of the blocks were covered, predominantly by large Chama, Arca, and oysters. The lower surfaces were mostly bare with an assemblage of small gastropods, Rissoidae, Cerithidae, Muricidae. Arcopsis can also be found attached to the undersides of rocks in the lower littoral often with Acar domingensis. (3) Offshore coralline sands and rubble Collection sites: IMBW-FK-651, 27-VII-02, “Samantha's patch reef” 5 nmi S of Marathon, Monroe County, Florida, 24°39.49’N, 81°00.32’W, 7.6 m; NMW.Z.2003.075.3. IMBW- FK-641, 23-VII-02, Tennessee Reef Light, off Long Key, Monroe County, Florida, 24°44.75’N 80°46.95’E, 6.1 m; NMW.Z.2003.075.4. IMBW- FK-624, 20-VII-02, Horseshoe Reef, off Fat Deer Key, Monroe County, Florida, 24*39.91'N, 80°59.56’E, 7.3 т; NMW.Z.2003.075.5 These sites are all of patch reefs with sandy bottoms and rubble blocks. Arcopsis are found attached to the undersides of the blocks lying on sand. (4) Sea grasses Arcopsis were also collected from a number of sea grass biotopes but not in sufficient num- bers for analyses. They were found attached to the base of shoots and exposed rhizomes. Methods Shell morphometric analyses were based on samples of at least 30 individuals, represent- ing the total size ranges, taken from the supralittoral, sublittoral, and offshore sites. In the hand, specimens from each biotope had a different appearance generally in the appar- ent inflation and elongation of the valves. Pa- rameters used focussed on these outward subjective views and included: shell length, shell height, and shell tumidity. The ligament appeared to be smaller in the offshore popu- lation, and the parameters inter-umbonal dis- tance, ligament length, and number of ligament bands were measured. During the process, it was suspected that the inflation of the right and left valves was not equal and consequently left and right valve tumidity were included. This analysis was not applied to the small number offshore specimens available, as the separa- tion of the valves would have destroyed the soft tissues, which were needed for anatomi- cal study. Paired parameter comparisons were made rather than a multivariate ap- proach in order to reveal specific differences. Arcoid bivalves are known to exhibit allomet- ric growth of the ligament (Thomas, 1975, 1976) and it was decided that other shell pa- rameters should be examined in relation to ontogeny. All data were analysed using Statview™. Shell sculpture and structure were examined using a scanning electron microscope. Anatomical comparisons were made from both living and preserved specimens. ARCOPSIS MORPHOLOGY AND ТАХОМОМУ 329 RESULTS Shell Morphometrics Maximum size: The maximum size of the shells differed between samples with the larg- est shells occurring in the shallow sublittoral and reaching 13.5 mm in length. In the supralittoral the maximum size recorded was 10.5 mm and for offshore itwas 9.5 mm. Sample sizes here for the supralittoral and sublittoral populations exceeded 100 individuals but for the offshore population only 30 were collected. offshore 8 sublittoral length:height ratio o. supralittoral offshore Je sublittoral supralittoral length:inter-beak distance ratio Comparisons of Shell Parameter Ratios: The most apparent difference in the hand was the shorter, more inflated form of the supralittoral shells, and to test this the length: height (Fig. 1) and length: tumidity (Fig. 2) ratios were com- pared. An ANOVA test (Fisher's PSLD) of the two ratios showed that the supralittoral popu- lation did have proportionately shorter and more tumid shells as compared to the other populations (significant at p = 0.0005 to < 0.0001) but that the difference between the sublittoral and offshore populations was not significant. 22 e = a - £2 5 2 Е = fo) 50 a = a offshore 1. sublittoral supralittoral O о > > A E > Se length:ligament length ratio о sublittoral offshore 104 JE supralittoral 8 left valve:righr valve tumidity ratio supralittoral subliforal [0] FIGS. 1-5. Box plots comparing shell morphometric ratios for populations of Arcopsis adamsi from the Florida Keys. FIG. 1: Shell length to shell height; FIG. 2: Shell length to shell tumidity; FIG. 3: Shell length to inter-umbonal distance; FIG. 4: Shell length to ligament length; FIG. 5: Left valve to right valve tumidity. 330 OLIVER & JARNEGREN Ligament length (mm) 2 4 6 8 10 12 14 Shell length (mm) Shell tumidity (mm) 2 4 6 8 10 12 14 Shell length (mm) FIGS. 6, 7. Comparisons of growth curves in three populations of Arcopsis adamsi from the Florida Keys. FIG. 6: Plot of ligament length against shell length; FIG. 7: Plot of shell tumidity against shell length. Diamond, supralittoral; dot, offshore; +, sublittoral. ARCOPSIS MORPHOLOGY AND ТАХОМОМУ 331 Comparisons of the inter-umbonal distance (length: inter-umbonal distance) gave the same pattern of results (Fig. 3) with significance at p < 0.0001. Comparison of the relative size of the ligament (length: ligament length) gave signifi- cant differences between all three populations (Fig. 4) except that this was less between the supra and sublittoral populations, p only 0.0106. Comparing the tumidity of left and right valves did reveal that Arcopsis adamsi 1$ slightly inequivalve (Fig. 5) and that the supralittoral population is significantly more inequivalve than the sublittoral population, p = 0.02. Comparisons of Growth Curves: The regres- sion plot of ligament length to shell length fits an exponential curve (Fig. 6) and confirms al- lometric growth of the ligament. Similar plots of tumidity (Fig. 7), interumbonal distance and number of ligament bands all reveal a similar pattern. In all cases, the point at which the curve most rapidly climbs is at a smaller shell size in the supralittoral population. In the offshore popu- lations the curves are least steep. Shell Sculpture and Structures Sculpture: On outward appearance the shells from the three biotopes have different aspects. The offshore shells (Figs. 10, 20) appear delicate, are clean with no apparent periostracum, white in colour with a sculpture of interlacing radial and concentric tracery. The density of the sculpture appears less in the offshore population, with the posterior area lacking the distinct semi-erect nodules seen in the sub and supralittoral populations. The sublittoral shells (Figs. 12, 18) are dirty with a pilose periostracum that retains fine sediment. The sculpture is partly obscured but when re- vealed appears primarily radial but is cancel- late. The supralittoral shells (Figs. 13, 16) are grubby, dirty white with the periostracum ei- ther lacking or persistent around the margins only. The sculpture appears primarily radial but is cancellate. A detailed examination using the scanning electron microscope reveals that all three populations have the same sculptural pattern (compare Figs. 16, 18, 20) but that the appar- ent differences are related to the expression of the concentric element. The sculpture con- sists of radial rows of elongate teardrop shaped pustules, the expanded portion coin- ciding with the intersection of the concentric element (compare Figs. 17, 19, 21). The con- centric sculpture consists of elevated bands attached to the radial elements but not to the interspaces, so when entire there 1$ a lattice effect. The observed differences consequently appear to be related to the degree and rapid- ity of erosion of the non-attached interspace concentric cords. Periostracum: The periostracum consists of flimsy concentric lamellae with fine hairs. In the offshore population, the periostracum 1$ not apparent in the hand but, when magnified, appears as a thin concentrically striated cov- ering expanded into thin lamellae along the lower edges of the concentric shell sculpture (Fig. 26). The lamellae bear thin, slightly thick- ened strap-like hairs at regular intervals. This contrasts with the obvious brown pilose cov- ering on the sublittoral and supralittoral shells (Fig. 25) where both the adherent portion and the lamellae are more strongly developed. Shell Pores: As with other arcoid bivalves, Arcopsis valves possess numerous pores (caeca) (Reindl & Haszprunar, 1996). These tubules traverse the valves and can be ob- served on the inner and outer surfaces. On the external surface the pores are visible be- tween the raised sculpture and under low magnification were most obvious in the supralittoral population. Scanning electron microscopy shows that the pores in all three populations are similar in size but that in the supralittoral shells the area around each pore is more heavily eroded and thus gives the appearance of being larger. Pore density was also examined and was observed to vary over the shell with a radial pattern present and a decrease in density to- wards the ventral margins. Comparisons be- tween populations were made by examining a strip on the internal surfaces directly below the ligament, and then at the same point on each strip (compare Figs. 27, 28, 29). A visual comparison suggests that the роге density 1$ greater in the supralittoral population. Larval Shell: The larval shells in all three populations are of the same form and same size (compare Figs. 22-24). There is a Prodissoconch | that is 105-106 um in width and is smooth. There is a Prodissoconch Il that is 174-178 um in width and has concen- tric sculpture of widely spaced raised lines. 332 OLIVER & JARNEGREN FIGS. 8-15. Scanning electron micrographs of Arcopsis and Striarca shells. FIGS. 8, 9: Arcopsis afra, Angola; FIGS. 10, 11: Arcopsis adamsi, offshore population, IMBW-FK-651; FIG. 12: A. adamsi, sublittoral population, IMBW-FK-629; FIG. 13: A. adamsi, supralittoral population, IMBW-FK-629; FIGS. 14, 15: Striarca lactea, Banyuls, Mediterranean Sea. ARCOPSIS MORPHOLOGY AND ТАХОМОМУ 333 FIGS. 16-24. Arcopsis adamsi, all scanning electron micrographs. FIGS. 16, 17: Cleaned supralittoral shell and detail of sculpture; FIGS. 18, 19: Cleaned sublittoral shell and detail; FIGS. 20, 21: Cleaned offshore shell and detail; FIGS. 22-24: Larval shells, supralittoral, sublittoral and offshore. 334 OLIVER & JARNEGREN Anatomy Gross Anatomy: The gross anatomy of Arcopsis adamsi is in all respects very similar to that of the Indo-Pacific, epibyssate Striarca symmetrica (Reeve, 1844) (Oliver, 1985) and the eastern Atlantic Striarca lactea (Oliver, pers.obs). Offshore Population (Fig. 30) — The adductor muscles are approximately of equal size, both with quick and catch portions. The posterior pedal (byssus) retractor is prominent but not large, some 5 x the size of the anterior pedal retractor. The mantle is thick, and the mantle edges are free, with the main inhalant and exhalant regions at the posterior; the frilled anterior mantle edge suggests that there is an ante- rior inhalant current. EICH” e a The foot has a developed toe and a smaller heel; the byssal groove runs along the ante- rior and median regions. The gills are paired, with both demibranchs well developed, the inner being larger than the outer. The labial palps are moderately large, with 12 to 15 well-developed palp ridges. The anus is attached to the underside of the posterior adductor muscle and is accompanied by a pair of abdominal sense organs each in the form of a simple dome. Sublittoral (Fig. 32) and Supralittoral (Fig. 31) Populations — As above, with the only appar- ent difference seen in the supralittoral popu- lation, where the inner demibranch is smaller and almost the same as the outer. It is not possible to discount differential contraction through the fixation process. === 200um FIGS. 25-29. Arcopsis adamsi, SEM of periostracum. FIG. 25: Supralittoral shell; FIG. 26: Offshore shell; FIGS. 27-29: Arcopsis adamsi, SEM of shell pores at the median area below the beaks; FIG. 27: Supralittoral; FIG. 28: Sublittoral; FIG. 29: Offshore. ARCOPSIS MORPHOLOGY AND ТАХОМОМУ 335 30 АРВ Pe PPR/BR FIGS. 30-35. Anatomy of Arcopsis adamsi. FIG. 30: Gross anatomy (left mantle removed), Samantha's Patch Reef (offshore population). A, anus; AA, anterior adductor muscle; APR, anterior pedal retrac- tor muscle; ASO, abdominal sense organ; CT, ctenidium; Е, foot; ID. inner demibranch of ctedium; LP, labial palps; ME, mantle edge; OC, ocelli; OD, outer demibranch of ctenidium; PA, posterior adductor muscle; PPR/BR, posterior pedal/byssus retractor muscle; FIGS. 31, 32: Gross anatomy of supralittoral (FIG. 31) and sublittoral (FIG. 32) specimens; FIGS. 33-35: Left mantle after shell removal showing areas of haemoglobin staining (stippled areas); FIG. 33: Offshore; FIG. 34: Sublittoral; FIG. 35: Supralittoral. 336 OLIVER & JARNEGREN Haemoglobin: The Arcoidea are one of the few bivalve superfamilies in which haemoglo- bin cells are a characteristic component of the haemocoelomic fluid. Arcopsis adamsitissues, especially those of the mantle, are tinged pink to blood red indicating the presence of hae- moglobin. The variation in colour intensity in- dicates that the haemoglobin concentrations differ between populations, but given the con- straints of the workshop we were unable to measure the actual concentrations. We ob- served that the living tissues of the offshore population are scarcely tinged pink and in fixed material a rust coloured band is present only along the mantle edge (Fig. 33). This contrasts completely with the supralittoral population, in which the living tissues, primarily the mantle, are dark blood red and in the fixed state the pigmentation covers most of the mantle (Fig. 35). The sublittoral population is intermediate in appearance but very distinctly with haemo- globin and thus more like the supralittoral population (Fig. 34). DISCUSSION The question for the taxonomist is of the degree of significance of these observations, could they reflect different species or are they result of ecophenotypic variation related to the different biotopes inhabited by the Arcopsis populations? Marko & Jackson (2001) using shell morphometrics concluded that the Pacific taxon Arcopsis solida was different from the Caribbean A. adamsi, but that this difference was primarily one of size rather than shape. Nevertheless, they nowhere suggested that the two were conspecific. Given that the two taxa are now isolated by the Panamanian isth- mus, their conclusion to maintain them as separate species was probably influenced by the geographical separation of the taxa as much as the morphological differences. Marko (2002) later showed that at the molecular level his populations of A. solida and A. adamsi were distinct. The anatomy of A. solida and A. adamsi has been described by Heath (1941), but no direct comparisons were made, and his observations were inconclusive in relation to differences between these species. Oliver & Cosel (1992), studying Striarca lactea populations along the West African coast, also used morphometrics to justify the erection of subspecies, giving outline and sculpture most credence. They did discuss the problems of erecting new taxa on such evi- dence but again were influenced by the geo- graphical separation of the populations. The morphometric data from these studies are comparable with those presented here but if they were used to create new taxa such de- cisions would be met with severe scepticism. The proximity of the populations and the data available would suggest to many that ecophenotypic variation was being observed. However, it is now widely accepted that, for example in the Littorinidae, that a number of species can live in adjacent microhabitats in close proximity to each other (Reid, 1986). This is accepted because the differences cited are from disjunct characters, such as the struc- tures of the genitalia and radula, rather than being based on statistical analyses of gradi- ents such as shell shape. Unfortunately, bivalves display few characters of this kind and one of the major problems in bivalve species taxonomy is the gradation of many characters. Greater character definition of these gradients would strengthen bivalve taxonomy, but in using statistical methods at what levels of sig- nificance do we attach a species level or popu- lation level distinction? In 1992 Oliver & Cosel did discuss the af- finities of the West African taxon A. afra and A. adamsi. They indicated sculptural differ- ences, but given the variation now seen in A. adamsi such differences are not so conclu- sive. Arcopsis afra appears more umbonate with a stronger sculpture, but in most respects difficult to separate from A. adamsi. Once again, the geographic isolation of the two taxa is giving the weight to their separation. It is therefore essential to examine in detail the possible causes of the morphological dif- ferences observed in populations that only show character gradients. Thomas (1975, 1976; Thomas et al., 2000) showed that the allometric growth of the arcoid ligament was essential to maintain a function- ing hinge. This allometry is expressed either in the progressive invasion of the ligament ventrally into the hinge plate or by further and further separation of the beaks. In Arcopsis the allometry is displayed in the separation of the beaks, and this allometry occurs at smaller shell length in the supralittoral population and is least marked in the offshore population. The growth function of the ligament is correlated not only to the relative inter-umbonal distance, ligament width and number of ligament bands, but probably also to the relative differences in shell tumidity and the degree of the inequiva- ARCOPSIS MORPHOLOGY AND ТАХОМОМУ 337 lve condition. Consequently, the morphometric data presented are all a function of growth and may be related to environmental parameters. Of the three biotopes sampled, the supra- littoral provides the most extreme conditions for a suspension feeding organism as feeding time is restricted to a short period at high tide times. Long periods of exposure will also cause potential stress from desiccation and restricted respiration. It is reasonable to suggest that restrictions in feeding time will reduce the growth rate in the supralittoral population and will result in the decrease in maximum size observed. If the allometry is age-related rather than size-related, then different growth rates will produce shells of different shapes where the parameters are linked to umbonal separa- tion. The increased relative tumidity and in- equivalve condition seen in the supralittoral population is likely to be growth-related and therefore ecophenotypic. However, the off- shore population has a similar maximum size but shows little allometry and is smaller than the sublittoral population. If we were to use ecophenotypic variation as an explanation of morphological difference, then it would be sup- portive if we could link the differences to envi- ronmental parameters. The taxonomist, therefore, should also be aware of the ecol- ogy and habitats of the taxa under study. Here we can only speculate at the ecological differ- ences between the habitats occupied by Arcopsis, but changes in growth rate and maxi- mum size are likely to be controlled by food availability. The function of the shell pores has been ten- tatively linked to respiration and haemoglobin is known to be a more efficient oxygen carrier than haemocyanin. The density of shell pores and the haemoglobin concentration are great- est in the supralittoral population and may be physiological responses to the greater time spent out of the water. When considering the sublittoral and offshore populations, it is more difficult to apply the same reasoning as both populations are permanently submerged. The sublittoral population lives under rocks embed- ded in muddy sand and may suffer oxygen depletion in the very warm surface waters along the margins of the Keys. At the offshore sites, the rocks sit on clean sand and water flow is probably much greater over the ani- mals. These differences may account for the different densities of shell pores and haemo- globin seen in the two submerged populations. Consequently, the characters that define the supralittoral population can all be related to environmental effects on growth rate and physiology, but a similar argument cannot ex- plain all the differences between the offshore population and the two shallow populations. Additionally, it is difficult to explain the differ- ence in periostracum simply through abrasion as that on the offshore population is not abraded only very thin. The sculptural density is also not readily explained by differential growth rates. Without disjunct characters, the taxonomic process has been led more and more towards ecology and physiology and increasing at- tempts to discover the functionality of the characters under review. Most of the discus- sion above is subjective and would require substantial experimentation to confirm whether or not morphology and physiology were responding to environment. Taxono- mists traditionally have not or were not able to support their decisions concerning species discrimination and intraspecific variation. Morphometric analyses do give statistical support to observations but do not resolve the issue. Environment does affect shell form and growth rate, and these may alter sculp- tural density reinforcing the need for the tax- onomist to be aware of ecology. At least in the arcoids, the ligament growth is related to the expression of many other shell charac- ters so that what may appear to be an array of characters is in reality a single one. Although morphological characters are the traditional tools of the bivalve taxonomist they must be used carefully, because many are gradients and many are influenced by envi- ronment. Without additional data from ecology, including physiology and reproductive biology, the interpretation of characters is difficult. The application of molecular techniques needs to become routine especially when ecophenotyic or geographic variation is suspected. It must be recognised that most bivalve species tax- опоту remains at the morphospecies concept and that a good species remains the product of a good taxonomist! A molecular study of the populations discussed here is in progress but the wider application of this technique now requires access to correctly preserved mate- rial, which precludes most of the collections in the worlds museums. Morphological based taxonomy will remain widespread as will the need to identify species based on readily ob- servable characters. Although molecular tech- niques are necessary, the routine use of molecular characters for identification is prob- ably a long way off. 338 OLIVER & JARNEGREN CONCLUSIONS Morphological characters in bivalves need careful assessment before conferring spe- cies level significance to differences in them. Many characters form gradients and need sta- tistical analyses to substantiate observations. Statistical differences in gradient characters do not necessarily indicate species difference. Many shell characters are inter-dependent and can all be altered by simple changes in growth. Where possible, ecology should be an inte- gral part of taxonomic studies. Molecular techniques need to be applied to complex problems to give better resolution. ACKNOWLEDGEMENTS The authors wish to thank the organisers of the International Marine Bivalve Workshop, Dr. Rüdiger Bieler and Dr. Paula M. Mikkelsen for the invitations to attend and to them and their colleagues for their considerable efforts to accommodate our research. The Workshop, held in the Florida Keys, 19-30 July 2002, was funded by U.S. National Science Foundation award DEB-9978119 to co-organisers К. Bieler and P. M. Mikkelsen, as part of the Partner- ships in Enhancing Expertise in Taxonomy [PEET] Program. Additional support was pro- vided by the Bertha LeBus Charitable Trust, the Comer Science & Education Foundation, the Field Museum of Natural History, and the American Museum of Natural History. Thanks are also due to the National Museum of Wales for additional support and facilities to complete this study. LITERATURE CITED ABBOTT, R.T., 1974, American Seashells. Van Nostrand Reinhold, New York. 663 pp. HEATH, H., 1941, The anatomy of the pelecy- pod family Arcidae. Transactions of the Ameri- can Philosophical Society, (n.s.) 31(5): 287-319, 22 pls. MARKO, P. B., 2002, Fossil calibration of mo- lecular clocks and on the divergence times of geminate species pairs separated by the Isth- mus of Panama. Molecular Biology and Evo- lution, 19(11): 2005-2021. MARKO, P. B. & J. B. C. JACKSON, 2001, Pat- terns of morphological diversity among and within arcid bivalve species pairs separated by the Isthmus of Panama. Journal of Paleontol- ogy, 75(3): 590-606. OLIVER, P. G., 1985, A comparative study of two species of Striarciinae from Hong Kong with comments on specific and generic systemat- ics. Pp. 283-310, in: B. MORTON & D. DUDGEON, eds., Proceedings of the Second International Workshop on the Malacofauna of Hong Kong and Southern China, 1983. Hong Kong Uni- versity Press, Hong Kong. OLIVER, P. G. & R. VON COSEL, 1992, Tax- onomy of tropical West African bivalves. V. Noetiidae. Bulletin du Muséum National d'Histoire Naturelle, Paris, (4)14[A](3-4): 655- 691. REID, D. G., 1986, The littorinid molluscs of mangrove forests in the Indo-Pacific Region. British Museum (Natural History) Publication 978, London. 227 pp. REINDL, S. 8 G. HASZPRUNAR, 1996, Fine structure of caeca and mantle of arcoid and limopsoid bivalves (Mollusca, Pteriomorpha). The Veliger, 39: 10-116. THOMAS, БК. D. K., 1975, Functional morphol- ogy, ecology and evolutionary conservatism in the Glycymerididae. Palaeontology, 18: 217- 254. THOMAS, БК. D. K., 1976, Constraints of liga- ment growth, form and function on evolution in the Arcoida (Mollusca: Bivalvia). Paleobiology, 2(1): 64-83. THOMAS, R. D. K., A. MADZVAMUSE, Р. К. MAINI 8 A. J. WATHEN, 2000, Growth patterns of noetiid ligaments: implications of developmen- tal models for the origin of an evolutionary nov- elty among arcoid bivalves. Pp. 279-289, in: E. M. HARPER, J. D. TAYLOR 8 J. A. CRAME, eds., The evolutionary biology of the Bivalvia. Geological Society of London, Special Publication 177. Revised ms. accepted 18 March 2004 MALACOLOGIA, 2004, 46(2): 339-354 ROCK AND CORAL BORING BIVALVIA (MOLLUSCA) OF THE MIDDLE FLORIDA KEYS, U.S.A. Paul Valentich-Scott' & Grete Elisabeth Dinesen? ABSTRACT Eight species from three bivalve families were collected and/or observed in the Middle Florida Keys. Diagnoses based on shell characters are given for Botula fusca, Lithophaga antillarum, L. aristata, and L. bisulcata in the Mytilidae, and Gastrochaena hians in the Gastrochaenidae. Shell and anatomical comparisons are made for three members of the Petricolidae, Petricola lapicida, Choristodon robustum, and Choristodon sp. A, which is not attributable to a described Recent Choristodon species. These bivalves bore into limestone and dead coral, and in one case into living coral. Observations substantiated previous findings of primary chemical boring processes in Botula and Petricola. Key words: Botula, Lithophaga, Petricola, Choristodon, Gastrochaena, endolithic, boring bivalves, Florida Keys. INTRODUCTION As an expansion of the general bivalve biodiversity study initiated by Mikkelsen & Bieler (2000), we here describe the rock and coral boring bivalve fauna of the Middle Florida Keys. The goal of this publication is to provide a guide to the identification of the rock and coral boring bivalves in the Middle Keys re- gion. Where possible, we have made obser- vations and comparisons of the living animal, the anatomy, and the habitat of each species. Middle Keys boring bivalves are represented in the families Mytilidae, Petricolidae, and Gastrochaenidae. Turner 8 Boss (1962) de- scribed the lithophagan mytilids throughout the western Atlantic, including the Florida Keys. Coan's (1997) treatment of the eastern Pa- cific Ocean Petricolidae discussed species that are also found in the Caribbean/Atlantic re- gion. The taxonomy and biology of the Gastrochaenidae are well documented in Carter (1978). Carter also provided a list of coral boring bivalves from Soldier Key, Dade County, Florida, which is only 100 km north of the site of this study (West Summerland Key). Including members in the three aforemen- tioned families, Kleemann (1980, 1990a) dis- cussed the methods of chemical boring of these bivalves in the Caribbean, eastern Pa- cific Ocean and the Great Barrier Reef. Morton (1990) presented a global overview of coral- boring bivalves, including those in the west- ern Atlantic Ocean. MATERIALS AND METHODS Limestone and coral habitats were examined for boring bivalves, intertidally and subtidally to 3 m in the Middle Florida Keys in July 2002 (Mikkelsen 8 Bieler, 2004, provide a station listing and map). Individuals were observed and/or collected primarily from the Florida Bay side of West Summerland Key (24°39’N). The limestone at this site 1$ thought to be Key Largo Limestone, which in some cases 15 overlain by the Miami Oolite facies (M. Campbell, pers. comm., March 2003). Boring bivalves were collected from limestone and dead coral sub- stratum with a rock hammer and chisel. Bivalves occurring in living coral were exam- ined, but not collected. Field observations of the living animal and their burrows were made. In addition, bivalve borers were observed at Bahia Honda State Park (24°39’N), Fat Deer Key (24°40’N), Crawl Key (24°41’N), Grassy Key (24°44’N), Long Key (24°45’М), Planta- ‘Department of Invertebrate Zoology, Santa Barbara Museum of Natural History, 2559 Puesta del Sol Road, Santa Barbara, California 93105 U.S.A.; pvscott@sbnature2.org Institute of Biological Sciences, Department of Marine Ecology University of Aarhus, Finlandsgade 14, DK-8200 Aarhus N, Denmark; grete.dinesen@biology.au.dk 340 VALENTICH-SCOTT & DINESEN tion Key (24°50’N), and Lower Matecumbe Key (24°50’N). Live animals were removed from their bur- rows, and relaxed in 7% MgCl,. Observations of the shell, ligament, siphons, mantle, and foot were made while the living animal was in a relaxed state. The right shell valve was care- fully removed with a scalpel inserted between the mantle margin and the shell. For selected species, the morphology of the labial palps, ctenidia, and siphons were recorded. Relaxed specimens were placed in 4% for- malin solution, and transferred to 70% ethyl alcohol. Ctenidial and labial palp filament counts were compared between living and pre- served specimens. Voucher specimens for all species collected in this study have been deposited at the Santa Barbara Museum of Natural History (SBMNH). Each species description includes a short diagnosis, followed by an expanded descrip- tion of the shell morphology and, where ob- served, the anatomy. Measurements and localities of specimens examined are given, along with habitats where the species were observed and collected. Additional literature citations are provided for each species, and when necessary additional remarks on the taxonomy or biology of the species are given. The following abbreviations are used in the text: AMNH, American Museum of Natural His- tory, New York, New York, USA; BMSM, Bailey- Mathews Shell Museum, Sanibel, Florida, USA; FMNH, Field Museum of Natural History, Chicago, Illinois, USA; SBMNH, Santa Barbara Museum of Natural History, Santa Barbara, California, USA. Station numbers listed in the following text refer to International Bivalve Workshop - Florida Keys (IMBW-FK) stations, as maintained by AMNH and FMNH (Mikkelsen & Bieler, 2004). SYSTEMATIC ACCOUNT Mytilidae Rafinesque, 1815 Botula fusca (Gmelin, 1791) Figures 1-4 Diagnosis Shell highly inflated; exterior dark brown; periostracum silky; beaks terminal, inturned, projecting beyond anterior margin; sculpture of commarginal striae only; without calcare- ous incrustations on exterior of shell; length of shell to 40 mm. Description Exterior —Lateral View: Shell subquadrate-elon- gate, slightly bent in the middle, slightly flar- ing posteriorly; beaks terminal, prosogyrate, inturned, pronounced, inflated; region ventral of umbones straight; posterior end rounded; broadly inflated from umbones to posterior margin, with rounded shoulders radiating from umbones to anteroventral and posterior re- gions, middle region slightly depressed; ven- tral margin incurved; byssus visible; sculpture of commarginal striae; periostracum chestnut brown, lighter brown in small specimens, strongly adherent; milky white mucus rem- nants attached to shell. Dorsal View: Ligament sunken, long, dark brown portion of ligament split for much of length; shell highly inflated. Ventral View: Umbones and prodissoconch visible from ventral side; ventral margin smooth; commarginal striae more pro- nounced near posterior margin. Interior. Shell pearly white internally; periostra- cum covering hinge under beaks; long thin, sharp posterior lateral tooth; edentulous un- der umbones; ligament deeply sunken, at- tached to a rolled nymph on the anterior portion, and a shelf-like nymph posteriorly. Anatomy Dorsal View: Foot orange anteriorly, white posteriorly, depressed in an anterior poste- rior direction, with small heel; distal portion of foot triangular, black; byssus projecting from base of foot; mantle unfused for most of ventral length; posterior rim of mantle 1$ dark brown, remainder of mantle milky white. Lateral View (with left valve and mantle re- moved): Anterior adductor muscle large for size of shell; posterior adductor circular, larger than anterior; inner fold of mantle margin very muscular, middle fold thin; labial palps short. Measurements Length 29 mm, height 13 mm, width 15 mm; length 17 mm, height 8 mm, width 9 mm; both specimens from West Summerland Key, IMBW-FK-629, 24°39.3’М, 81°18.2’W, col- lected by P. Valentich-Scott and G. Elisabeth Dinesen (SBMNH 350547, 350548). Additional observations were made at Crawl Key and Bahia Honda State Park. Four additional lots of dry specimens from the Florida Keys were examined (SBMNH). ROCK AND CORAL BORING BIVALVIA 341 Habitat In а mucus nest, boring in soft limestone. Carter (1978) reported in dead coral (Diploria). Remarks The limestone burrows of several specimens were found with dorsal keels, or with anterior notches in the limestone under the umbones (Fig. 4). Mechanical boring would not allow these keels or notches to be formed in the borehole. These findings correspond with Wil- son & Tait (1984), who suggested that Botula fusca only uses chemical means for boring. There has been much nomenclatural debate as to the correct name for the species in the western Atlantic Ocean. Wilson & Tait (1984) FIGS. 1-4. Botula fusca. FIGS. 1, 2: External left valve, internal right valve. West Summerland Key, Monroe County, Florida; 24°39.3’N, 81°18.2’W; subtidal; Station 629; length 28.8 mm; SBMNH 350546; FIG. 3: Dorsal view, Grassy Key, Monroe County, Florida; 24°45’46”N, 80%57'11”W; length 39.6 mm; SBMNH 53503; FIG. 4: In limestone substratum; arrows denote invagination below umbones and See, banding notch in limestone; locality data the same as figures 1-2; length 26.1 mm; SBMNH 47. 342 VALENTICH-SCOTT & DINESEN used Botula fusca (Gmelin, 1791) as a single global species distributed in the Indian, Atlantic, and Pacific Oceans, and placed B. cinnamonea (Gmelin, 1791) in synonymy. Nielsen (1986) con- trasted this viewpoint, seeing B. cinnamonea as valid, with a broad Northern Hemisphere distri- bution. In addition, Nielsen designated a lecto- type for B. cinnamonea, and restricting the type locality of this species to the Nicobar Islands. Additional morphological, anatomical and genetic studies are needed to solve this global issue. Literature Abbott (1974: 436), Keen (1971: 74), Mikkelsen & Bieler (2000), Nielsen (1976), Redfern (2001: 201), Soot-Ryen (1955: 86), Wilson & Tait (1984). FIGS. 5-8. Lithophaga antillarum. Missouri Key, Monroe County, Florida; 24°40.6’М, 81°14.3'W; length 77.9 mm; SBMNH 350549. FIG. 5: Dorsal view; FIG. 6: External left valve; FIG. 7: Internal right valve; FIG. 8: Ventral view. ROCK AND CORAL BORING BIVALVIA 343 Lithophaga antillarum (Orbigny, 1853) Figures 5-8 Diagnosis Shell elongate, cylindrical; beaks subtermi- nal, but not extending past anterior end; periostracum lightto medium brown, dehiscent; sculpture of fine vertical lines over most of shell, and heavy commarginal undulations postero- dorsally; without calcareous incrustations on exterior of shell; length of shell to 120 mm. Description Exterior— Lateral View: Shell cylindrical, some- what compressed laterally, sharply rounded anteriorly, broadly rounded posteriorly, slightly flaring in the middle portion; beaks subterminal, small; sculpture of fine vertical lines over entire surface except narrow re- gion from beaks to posterior end, and irregu- lar commarginal striae, commarginal undulations posterodorsally; large portions of shell eroded, especially anteriorly; periostracum dehiscent, medium brown; cal- careous incrustations not present on shell, no encrusting extensions. Dorsal View: Beaks small, not inflated or pro- truding; dorsal margin not gaping; ligament not visible from dorsal surface; with long nar- row escutcheon; lunule not well demarcated; shell widest near midline, tapering posteriorly. Ventral View: Shell tightly closing, except for very narrow, short pedal gape, and very slight posterior gape; ventral margin slightly beveled inward. Interior: Interior pearly white, translucent; edentulous; ligament dark brown, deeply sunken, extending from umbones nearly to the shell midline. Anatomy Not examined. Measurements Length 85 mm, height 25 mm, width 21 тт; specimen collected by José Leal (26 July 2002) at West Summerland Key, IMBW-FK- 629, 24°39.3’N, 81°18.2’W at 3 т depth, in soft limestone; deposited as a voucher speci- men at the Zoological Museum, University of Copenhagen, Denmark. Also observed at Fat Deer Key. Eight additional lots examined from Missouri Key (24°40’N) (SBMNH 350549), Vaca Key (24°46’М), and Barbados (all SBMNH), and Lower Matecumbe Key and Townsend Island (BMSM). Habitat Boring into soft limestone. Carter (1978) re- ported in dead coral (Diploria), and Scott (1988a) observed in dead coral and rock. Literature Turner & Boss (1962), Kleemann (1983, 1984, 1990a, b, 1996), Mikkelsen & Bieler (2000), Morton (1990), Redfern (2001: 201), Warmke & Abbott (1971: 164). Lithophaga aristata (Dillwyn, 1817) Figures 9-11 Diagnosis Shell inflated, cylindrical; beaks subterminal; with heavy calcareous incrustations over most of shell; elongated incrustations posteriorly, forming overlapping scissors-like “forceps”; length of shell to 33 mm. Description Exterior — Lateral View: Shell elongate ovate to cylindrical, sharply rounded anteriorly, ta- pering posteriorly; beaks subterminal, usu- ally eroded; sculpture of fine commarginal striae; periostracum dark brown; heavy cal- careous incrustations over entire surface, eroded in some spots; incrustations extend- ing past the posterior end, forming overlap- ping, scissors-like projections. Dorsal View: Beaks usually eroded, not ex- tending past the anterior margin; ligament black, sunken anteriorly, becoming visible near shell midline. Ventral View: Shell tightly closing, without vis- ible pedal gape; ventral margin nearly straight; posterior scissors-like incrustations easily viewed from this orientation. Interior. Shell very thin, fragile, translucent, slightly pearly white, slightly flaring dorsally; edentulous; posterior end of shell tapering, with calcareous extensions. Anatomy not examined. Morton (1993) dis- cussed various aspects of the anatomy, in- cluding a discussion on the formation of the scissors-like “forceps”. 344 VALENTICH-SCOTT & DINESEN FIGS. 9-11. Lithophaga aristata. West Summerland Key, Monroe County, Florida; 24°39.3'N, 81°18.2’W; subtidal; Station 629; length 9.9 mm; SBMNH 350550. FIG. 9: Dorsal view; FIG. 10: Ventral view; FIG. 11: External left valve. Measurements Length 9.9 mm, height 4.1 mm; specimen collected by Diarmaid O'Foighil (27 July 2002) at West Summerland Key, IMBW-FK-629, 24°39.3’М, 81°18.2’W (SBMNH 350550). Two additional Florida lots were examined, along with 50 lots from the eastern Pacific Ocean (all SBMNH). Habitat Boring into limestone and coral. Coan et al. (2000) reported boring into shell in the east- ern Pacific Ocean. Literature Coan et al. (2000: 181), Keen (1971: 70), Kleemann (1983, 1990a, b, 1996), Mikkelsen & Bieler (2000), Morton (1993), Redfern (2001: 202), Turner & Boss (1962), Yonge (1955). Lithophaga bisulcata (Orbigny, 1853) Figures 12-15 Diagnosis Shell cylindrical, with flare along dorsal mar- gin, tapering posteriorly; with feathery calcar- eous incrustations along posterodorsal slope; incrustations extending evenly past posterior end of shell; length of shell to 45 mm. Description Exterior — Lateral View: Shell cylindrical, ta- pering posteriorly, anteriorly rounded; dor- sal and ventral margin parallel for the anterior half of the shell, flaring posterodorsally and then tapering posteriorly; beaks broad, slightly projecting, near anterior end, not ter- minal; sculpture of fine commarginal striae, with broad keel running from just posterior of beaks to posterior end; periostracum chestnut dark brown; surface anterior of keel with fine granular concretions except in the umbonal region and ventral margin, poste- rior of keel with heavy concretions, concre- tions becoming heavier posteriorly, feathery concretions posteriorly, posterior concretion extension short with fine granules. Dorsal View: Ligament deeply sunken in deep long escutcheon; anterior end triangular; gaping posteriorly; concretions along entire dorsal surface. Ventral View: Ventral margin slightly incurved to slightly bowed, smooth; shell narrowly gaping posteriorly; posteroventral calcare- ous incrustations with zipper-like pattern. ROCK AND CORAL BORING BIVALVIA 345 FIGS. 12-15. Lithophaga bisulcata. Missouri Key, Monroe County, Florida; 24°40.6’N, 81°14.3’ W; length 45.0 mm; SBMNH 350551. FIG. 12: Dorsal view; FIG. 13: External left valve; FIG. 14: Internal right valve; FIG. 15: Ventral view. Interior. Shell dark brown, translucent, with straight off posterior end, not forming for- slight sheen; edentulous; ligament deeply серз. sunken, extending from beaks to the end of dorsal flare (well posterior of midline); beaks Anatomy near anterior end, but not subterminal; an- terior end broadly rounded, dorsal flaring, Not examined. Scott (1988a) detailed much posterior tapering; calcareous incrustations of the anatomy of this species. 346 VALENTICH-SCOTT & DINESEN Measurements Length 21 mm (4 mm are the forceps concre- tions); height 6.5 тт; width 6 mm; specimen collected by Diarmaid O'Foighil (27 July 2002) at West Summerland Key, IMBW-FK-629, 24739.3'N, 81°18.2’W. Six additional Florida lots examined (SBMNH), including specimens from Missouri Key (SBMNH 350551). Habitat Boring into limestone. Scott (1988a) reported from living and dead coral, and rock. Literature Kleeman (1983, 1990a, b, 1996), Mikkelsen 8. Bieler (2000), Morton (1990), Redfern (2001: 202), Scott (1985, 1988a, b), Turner & Boss (1962), Warmke 8 Abbott (1971; 164). Petricolidae Orbigny, 1840 Choristodon robustum (С. В. Sowerby |, 1834) Figures 16-19, Table 1 Diagnosis Shell ovate-elongate to trigonal, moderately inflated; inequilateral, posterior end much longer; anterior broadly rounded, posterior end tapering; sculpture of strong, irregular radial ribs, most prominent on the central portion of the shell; anterior and posterior ends gaping; siphons fused for nearly half length; length of shell to 43 mm. Description Exterior — Lateral View: Shell ovate to trigonal; moderately inflated; inequilateral, posterior end much longer; anterior end rounded, pos- terior end attenuate; umbones prosogyrate; FIGS. 16-19. Choristodon robustum. West Summerland Key, Monroe County, Florida; 24*39.3'N, 81°18.2"W; subtidal; Station 629. FIGS. 16, 17: Length 20.2 mm; SBMNH 350554; FIG. 16: External left valve; FIG. 17: Internal right valve; FIG. 18: Length 19.8 тт; dorsal view; SBMMH 350553; FIG. 19: Siphons of living animal, demarcation showing region of siphonal fusion; SBMNH 350552. 347 ROCK AND CORAL BORING BIVALVIA 20] pajulod :Aj218}e] passaiduo9 .J0]09 yoead lee moyym ‘ous ‘реола 20] ‘реола ‘HOIU} ‘10/09 эуцм pajuiod ‘jjews [eau ¿pajulod pue dieus 'jews 20] :ajos био] ‘pesseid -WOOd А|е/э}е| ‘10109 SHUM 003 эеэна Op :уочеланиер Jajmo ‘эеэна ZG :youelq -1WOP лэци! seid с :уочеланиер Jamo ‘эеэна 9, :ypueıq -циэр лэци! эеэ!а OZ :уочеланиер Jamo ‘эеэна 02 :ypueıq -циэр лэци! эеэна youeqiuag эбиело Улер HUM Ашеэю 0) yuid зчбн SHUM Ашеэю JO|O9 е!р!иэ} send ze UJIM :9ZIS unIpauu ‘SHUM эеэ!а 9) :9ZIS ||euus ‚эбиело 1461 эеэна VL Um ‘971$ |[EUUS ¿MOJ|9Á 0} yuıd 1461] sdjed ¡e1qe] Jano рэ$п; Ájuo juejeyul jo у биз! леч sqou ajoejue} Hous Â18A (2) ‘apıs UO Sedo] |jeWIS Чил^ ‘эзеэлпуа JOU 'pajurod ‘ajduuis (|) 'sadÁ] ом} Jo saj9e]ua)] ЭУ|-1эмо|} ‘PUM эле eellided лэци! ‘sjods имола улер YIM ae||ided yous ‘ajdwis ши биде эе!аеа ajduis ‘uous saj9e]ua) jeuoudis juejeuul peuouelq Alıneay sıayJo ‘зиоцеэлпуа a¡duwis auwos 'sadÁ] |елэлэ$ jo эле sajoejua] SI -JIMO|Y ‘SYM эле ae] ided Jauul :SJods umoug wep чим eellided yous ‘ajduuis ши Buoje sejlıded SHUM ‘ejduuis sope}ug} jeuoydis Juejeyx3 Ájloua]sod squ jelpes “SQU эзеэмел!р eu ‘эзелрепрап$ A\s0119}S0d JaBuoJ]s Ббишоээа ‘Кпоизие squ jelpes эиц ‘eBuey YM лоиазие ‘ела}е| -inbaqns “ajeno squ ¡e1pe, лепбэлл ‘Buoys :JaBuo] yonu риэ Jouasod 'jeuoßi 0} э}ебио|э-э}ело aunydjnos / edeus ||э4$ eploide] eJ091}9 y ‘ds иоро]5иоцэ winjsngos UOpoj}sLOYD saioeds `зАэя BPUO|4 SIPPIN SU} WO эерноэщед SY) JO злеашеш Jo sosHa}eIeYD “| 378VL 348 VALENTICH-SCOTT & DINESEN beaks broad, projecting; sculpture of irregu- lar radial ribs, weak anteriorly, strong posteri- orly; posterior ribs prominent, broad, rounded, groove between ribs shallow, broad; anterior ribs very weak, barely visible at anterior end; mid portion ribs gradually increasing in size and height, sharp, thin; commarginal striae closely spaced, making surface weakly can- cellate, slightly lamellate posteriorly. Dorsal View: Shell moderately inflated, slightly compressed posteriorly; shell inequivalve, right side larger; shell gaping anterior and posterior of beaks; ligament short, external, sunken, on nymph; lunule small, deep; prodissoconch large. Ventral view: Widely gaping except for midline; terminal end of radial ribs intermesh at mid- line; right valve convex posteriorly; left valve concave posteriorly; posterior end twisted to the left; inequivalve, left valve smaller. Interior. Hinge plate short; three cardinal teeth in left valve — two anterior teeth short, stout, posterior tooth larger, plate-like, pointing pos- teriorly; right valve with two cardinal teeth — anterior tooth short, wide, stout, posterior tooth very small, thin, plate-like; ligament in two parts, outer section beginning just below beaks, light brown, inner section attached to nymph, black. Pallial sinus broad, shallow, not extending to beaks (about 1/3 distance between adduc- tors); right valve pallial sinus slightly broader than that of left valve; pallial line continuous in sinus region, patchy along ventral margin; anterior adductor muscle scar long, moder- ate in width, pointed dorsally and ventrally; posterior adductor muscle scar nearly circu- lar; left valve with two small pedal retractor scars posterodorsally; inside shell surface chalky; inner margin weakly and irregularly crenulate behind umbonal middline, inner margin non-crenulate anteriorly. Anatomy (Table 1) External View: Siphons translucent pale yellow/ orange on outer section, milky white near mantle, with small white granules in tissue, and brown streaks and blotches; exhalant si- phon much narrower than inhalant; inhalant siphon with short, simple papillae along rim; exhalant with simple papillae along rim; middle mantle fold light orange distally, rim plicate. Internal View: Outer and inner mantle fold very thick milky white, middle fold plicate, light or- ange with sporadic white granules towards the siphons, middle mantle fold near siphons dark brown; mantle fused from siphons to line below umbones; pedal gape relatively short extending from below beaks to anterior mar- gin; labial palps small, with 14 plicae; siphons fused for approximately half of their length; ctenidia creamy white; ctenidial plicae paral- lel to dorsal margin; outer demibranch 2/3 length of inner demibranch; plicae much larger and wider than Р lapicida, approxi- mately 20 plicae on outer demibranch, about 20 on inner demibranch; foot white, laterally compressed, long sole, toe small, sharp, pointed; with small pointed heel. Measurements Length 19 mm, height 14 mm, width 10 mm; specimen collected by P. Valentich-Scott and G. Elisabeth Dinesen at West Summerland Key, IMBW-FK-629, 24°39.3’N, 81°18.2’W (SBMNH 350552). Habitat Shallow, unlined burrows in limestone rocks. Carter (1978) reported (as Rupellaria typica) in dead coral (Diploria). Literature Coan (1997), Keen (1971: 199), Lamy (1923), Redfern (2001: 240), Warmke & Abbott (1971: 199, as Rupellaria typica). Choristodon sp. A Figures 20-23, Table 1 Diagnosis Shell ovate, inflated; subequilateral; anterior end with flange, extending well beyond inner shell margin; sculpture of fine radial ribs on anterior portion, stronger radial ribs posteriorly; anterior end slightly gaping, posterior end tightly closed; siphons only fused basally; length of shell to 23 mm. Description Exterior — Lateral View: Shell ovate, highly in- flated, anterior end broad, posterior end slightly tapered; subequilateral, posterior end slightly longer; posterodorsal margin straight; anteroventral margin flared laterally; beaks broad, inflated, prosogyrate; lunule deep; sculpture of pronounced radial flat-topped ribs, interspaces deep, wide, overlain by fine commarginal striae. ROCK AND CORAL BORING BIVALVIA 349 FIGS. 20-23. Choristodon sp. A. West Summerland Key, Monroe County, Florida; 24*39.3'N, 81°18.2’W; subtidal; Station 629; length 22.8 тт; SBMNH 350556. FIG. 20: External right valve; FIG. 21: Internal left valve; FIG. 22: Dorsal view; FIG. 23: Siphons of the living animal, arrow showing fusion only at base of siphon (SBMNH 350555). Dorsal View: Ligament deeply sunken, short; shell gaping anteriorly, but closed posteriorly; equivalve. Ventral View: Anterior end slightly gaping, pos- terior end tightly closed. Interior. Right valve with two cardinal teeth, with large stout anterior tooth, fairly large plate- like posterior tooth; left valve with 3 cardinal teeth, anterior tooth small, stout, middle tooth large stout, posterior tooth thin plate-like. Anatomy (Table 1) Siphons small, short; space between siphons dark brown, dorsal of exhalent siphon dark brown, remaining area around siphons white; rim of both siphons with simple, short papillae with dark brown spots; inner papillae white, flower- like; siphons only fused for a short distance be- yond mantle; posterior portion of mantle very dark brown; outer fold thick, smooth; middle mantle fold thinner than outer, slightly plicate, pigmented towards siphons; inner mantle fold thick, milky white, smooth; pedal gape short anteriorly; inner mantle fold unfused along anterior margin, but fused for remainder of ventral margin; labial palps small, short, with 16 plicae; сета pale pink to creamy white; outer demibranch with 12 plicae, inner demibranch with 16 plicae; footthick, broad, without heel, with broad, short anterior end. Measurements Length 25 mm, height 20 mm, width 16 тт; specimen collected by P. Valentich-Scott and G. Elisabeth Dinesen at West Summerland Key, IMBW-FK-629, 24*39.3'N, 81*18.2'W (SBMNH 350555). 350 VALENTICH-SCOTT & DINESEN Habitat Boring into limestone, adjacent to Choristodon robustum. We found many shells of this species to be heavily bored by sponges and polychaetes. Remarks: Coan (1997) placed Choristodon typica Jonas, 1844, in synonymy with C. robustum, based on the figure provided by Jonas (Coan, 1997: fig. 43). The species we describe above is distinct, conchologically and anatomically, from C. robustum (e.g., shell out- line and sculpture, siphonal fusion, siphonal tentacles). As yet, we have not found a de- scribed species to correspond with our mate- rial. However, our specimens are very similar to the species illustrated by Narchi (1974), which he identified as C. typica. The Florida Keys species is not the same at Redfern's (2001: 240) Petricola sp. from the Bahamas, nor Р. stellae (Narchi, 1975) from Brazil (Narchi, 1975). Table 1 compares anatomical characters of the two species of Choristodon found in the Middle Florida Keys, along with Petricola lapicida. Literature Narchi (1974). Petricola lapicida (Gmelin, 1791) Figures 24-27 Diagnosis Shell subquadrate; inequilateral, posterior end much longer; sculpture of fine divaricate ribs over entire surface, and partial radial ribs near the posterior margin; siphons not fused; length of shell to 30 mm. FIGS. 24-27. Petricola lapicida. West Summerland Key, Monroe County, Florida; 24°39.3’N, 81°18.2’W; subtidal; Station 629; length 27.3 mm; SBMNH 350343. FIG. 24: External left valve; FIG. 25: Internal right valve; FIG. 26: Detail of external of right valve showing divaricate markings; FIG. 27: Preserved animal in limestone burrow, arrows denote burrow tightly fitting around animal (anteriorly), and the constricted posterior portion of the burrow; SBMNH 350559. ROCK AND CORAL BORING BIVALVIA 351 Description Exterior — Lateral View: Shell subquadrate, strongly prosogyrate, beaks broad, inflated; inequilateral, posterior end much longer, beaks almost at anterior end; anterior end rounded; posterior end truncate; postero- dorsal margin nearly straight; sculpture very fine, divaricate ribs over most of surface (Fig. 26); posterodorsal region with few pro- nounced, sharp radial ribs, terminating be- fore margin, sometimes wavy near ventral margin (eroded in some), interspaces be- tween radial ribs wide, flat. Dorsal View: Inflated anteriorly, more com- pressed posteriorly; equivalve; ligament deeply sunken, short; lunule deeply exca- vated posteriorly beneath ligament. Ventral View: Without ventral gape. Interior. Hinge plate short, triangular; peri- ostracum in lunular region; ligament deeply sunken, seated on a elongate infolded nymph, in two sections, both dark brown; left valve with two teeth, anterior tooth large, rectangular, posterior tooth small, thin, plate- like; right valve with two teeth, anterior small peg-like, posterior larger but plate-like; pallial sinus very broad, shallow, not reaching beaks; ventral pallial line slightly patchy, continuous in sinus area; anterior adductor muscle scar long, narrow (slightly broader than C. robustum); posterior adductor circular. Anatomy (Table 1) Mantle fused from beak to anteroventral margin, small fusion just anterior of inhalant siphon; mantle open over entire ventral re- gion from inhalant siphon to anterior margin; mantle without papillae; most of mantle milky white, except near siphons where it is dark brown in color; mantle swollen antero- ventrally, possibly a pallial gland; outer mantle fold very thin; middle fold muscular, tapering on margin, wavy, inner fold thin; mantle filled with white granules; labial palps white, me- dium length, pointed ventrally, with 32 plicae; smooth dorsally and anterior portion of palp; ctenidia dark orange; plicae parallel to dor- sal surface; outer demibranch extending to middle of inner demibranch; plicae number on demibranchs — inner 52, outer 40; siphons transparent dark gray, with embedded white granules; exhalant siphon circular in outline with tentacles of several types, some with simple bifurcations, others heavily branched; inhalant siphon elongate-ovate, gray with white spots; inhalant siphonal tentacles of two types, large compared to exhalant, simple, pointed (not bifurcate), with small lobes on side; also very short tentacle nobs projecting; inhalant siphon three times as large in diameter as exhalant; siphons barely extending beyond shell margin; foot compressed laterally, peach color, very flexible, pointed at tip. Measurements Length 27 mm, height 20 mm, width 13 mm, specimen collected by P. Valentich Scott and G. Elisabeth Dinesen at West Summerland Key, IMBW-FK-629, 24°39.3’М, 81°18.2’W (SBMNH 350558). Remarks Field observations of the burrow of Petricola lapicida have shown it lives in a constricted, flat burrow (Fig. 27). This strongly suggests the species burrows through chemical means only, and agrees with the findings of Morton 8 Scott (1988) and Morton (1990). Compari- sons between the functional morphology of P. lapicida and P. pholadiformis were pre- sented by Purchon (1955). Habitat Shallow burrow in limestone. Carter (1978) reported this species in dead coral (Diploria). Literature Abbott (1974), Bromley (1978), Kleemann (1990a), Lamy (1923), Morton (1990), Morton 8 Scott (1988), Redfern (2001: 240), Robertson 1963, Warmke 8 Abbott (1971: 191). Gastrochaenidae Gray, 1840 Gastrochaena hians (Gmelin, 1791) Figures 28-30 Diagnosis Shell ovate, white; incurved and widely gap- ing ventrally; widely gaping posteriorly; beaks terminal. Description Exterior — Lateral View: Shell inflated, ovate elongate; posterior end rounded, flaring; an- terior end narrow pointed; beaks terminal, 352 VALENTICH-SCOTT & DINESEN pointed, prosogyrate; prodissococh large, smooth; widely gaping anteroventrally, invagi- nate; shell color translucent white; sculpture of commarginal striae, stronger antero- ventrally, without radial elements; shell thicker along ventral gape. Dorsal View: Highly inflated, more compressed posteriorly; right valve overlapping the left; left valve slightly concave posteriorly; ligament external, protruding, long, one third of shell length; valves slightly gaping posterior to liga- ment. Ventral View: Periostracum thin, milky white, translucent, dehiscent; outer mantle fold thick, projecting beyond valve margin, wide gape, not fused for half shell length; middle mantle edge fused except for small pedal gape near shell midline; posterior end tightly closed, right valve overlapping left; posteriorly periostracum projecting beyond shell margin. Interior. Not examined. Anatomy See Carter (1978) for discussion of Gastro- chaena anatomy and shell features, along with diagnostic characters of related species. Measurements Length 11 mm, maximum height 6 mm, width 4.5 mm, ligament length 4.5 mm, gape length 8 mm, gape width 4 mm; specimen collected by Lisa Kirkendale on 27 July 2002, at Fiesta Key, IMBW-FK-644, 24°50.4’М, 80%47.0"W (SBMNH 350345). Three additional specimens were collected by the authors from West Summerland Key, IMBW-FK-629. Habitat In calcareous lined burrows in living and dead coral, and limestone. Carter (1978) reported in dead coral (Diploria). Remarks Coan, et al. (2000: 494, left specimen) illus- trated a Florida specimen of Gastrochaena as С. ovata, but this specimen is actually С. hians. Literature Carter (1978), Morton (1983, 1990), Redfern (2001: 242). FIGS. 28-30. Gastrochaena hians. Fiesta Key Causeway, Monroe County, Florida Keys; 24°50.4’N, 80°47.0'W; subtidal; station 644; length 11.8 mm; SBMNH 350345. FIG. 28: Dorsal view; FIG. 29: Ventral view; FIG. 30: Lateral view of left side. ROCK AND CORAL BORING BIVALVIA 353 DISCUSSION The rock and coral boring bivalves of the Middle Florida Keys are diverse and numerous. With a modest sampling effort, eight species representing three families were observed and collected. While quantitative studies were not undertaken, several limestone rocks had more than 50 individuals/m?. However, there was а distinct patchiness to the distribution of these borers, even with seemingly identical substrata in adjacent areas. Often, large limestone boul- ders were completely void of bivalve borers, where adjacent rocks were riddled with petricolids, mytilids, and gastrochaenids. Careful examination of the living bivalves and boreholes has confirmed the boring mecha- nisms of two species. In agreement with Wil- son & Tate (1984) and Kleemann (1990a), our observations indicate that Botula fusca is a chemical borer (Fig. 3). Similarly we have found strong indications of chemical boring in Petricola lapicida (Fig. 27), concurring with Morton & Scott (1988). Lithophagans were rela- tively rare in our sampling areas, and we were unable to make definitive conclusions on habi- tat or boring mechanisms of these species. Far outside the scope of this paper are con- clusions about the localized or global distribu- tions of many boring bivalve species. Among the different lineages of boring bivalves, several are thought to be represented by a single genus with one or only a few species, and distributed world- wide (Morton, 1990). Morton further discussed the evolutionary events and implications, which could explain both the presence of true cosmo- politanism of some species and restricted re- gional distribution of other species. Nomenclatural inconsistency by researchers may account for confusion between cosmopoli- tan distributions and localized endemism within the boring bivalve lineages. This is easily un- derstood, as the majority of boring bivalve spe- cies names (and most marine bivalves) were Originally designated exclusively based on shell characters. The shell morphology of boring bivalves has shown intraregional variation as large as interregional variation (Coan, 1997). This could be due to worldwide conspecifity, as has been suggested for Botula fusca by Wilson & Tait (1984), with shell plasticity as a consequence of individual morphometric adap- tation to their boring habitat. The use of shell features to discriminate between the species within different lineages of boring bivalves still needs confirmation from other methods (e.g., gross anatomy and histology, molecular se- quencing and analyses). ACKNOWLEDGEMENTS We deeply appreciate all of the efforts in or- ganizing the international workshop on the Bivalvia by Rudiger Bieler (FMNH), Paula Mikkelsen (AMNH), Russ Minton (ЕММН), Isabella Kappner (FMNH), and the staff at the Keys Marine Laboratory. Important specimens for this project were collected by Liz Kirkendale (University of Florida), and José Leal (BMSM), who provided specimens and photographs. Eugene Coan gave a very helpful review of the first draft of this manuscript. Henry Chaney and Patricia Sadeghian, both of SBMNH, gave use- ful cornments on a later draft of this paper. Thanks go to Claus Nielsen, Zoological Mu- seum of Copenhagen, for valuable discussion and help with Gmelin and Chemnitz type ma- terial and designations. The International Marine Bivalve Workshop, held in the Florida Keys, 19-30 July 2002, was funded by U.S. National Science Foundation award DEB-9978119 (to co-organizers R. Bieler and P. M. Mikkelsen), as part of the Partner- ships in Enhancing Expertise in Taxonomy [PEET] Program. Additional support was pro- vided by the Bertha LeBus Charitable Trust, the Comer Science & Education Foundation, the Field Museum of Natural History, and the American Museum of Natural History. LITERATURE CITED ABBOTT, R. T., 1974, American seashells; the marine Mollusca of the Atlantic and Pacific coasts of North America, 2nd ed. Van Nostrand Reinhold, New York. 663 pp. BROMLEY, R. G., 1978, Bioerosion of Bermuda reefs. Palaeogeography, Paleoclimatology, Palaecology, 23: 169-197. CARTER, J. G., 1978, Ecology and evolution of the Gastrochaenacea (Mollusca, Bivalvia) with notes on the evolution of the endolithic habi- tat. Peabody Museum of Natural History Bul- letin, 41: 1-92. COAN, E. V., 1997, Recent species of the ge- nus Petricola in the eastern Pacific (Bivalvia: Veneroidea). The Veliger, 40(4): 298-340. COAN, Е. V., Р. VALENTICH-SCOTT & Е. К. BER- NARD, 2000, Bivalve seashells of western North America. Marine bivalve mollusks from Arctic Alaska to Baja California. Santa Barbara Museum of Natural History, Monographs 2, viii + 764 pp. KEEN, A. M., 1971, Sea shells of tropical west America. Stanford University Press, Stanford, California. 1064 pp. KLEEMANN, K. H., 1980, Boring bivalves and their host corals from the Great Barrier Reef. Journal of Molluscan Studies, 46: 13-54. KLEEMANN, K. H., 1983, Catalogue of recent and fossil Lithophaga (Bivalvia). Journal of Molluscan Studies, Supplement 12: 1-46. 354 VALENTICH-SCOTT & DINESEN KLEEMANN, К. H., 1984, Lithophaga (Bivalvia) from dead coral from the Great Barrier Reef, Australia. Journal of Mollusca Studies, 50: 192- 230. KLEEMANN, К. H., 1990a, Boring and growth in chemically boring bivalves from the Caribbean, eastern Pacific and Australia's Great Barrier Reef. Senckenbergiana Maritima, 21(1/4): 101-154. KLEEMANN, K. H., 1990b, Evolution of chemi- cally-boring Mytilidae (Bivalvia). Pp. 111-124, in: B. MORTON, ed., The Bivalvia - Proceedings of a memorial symposium in honour of Sir Charles Maurice Yonge, Edingburgh, 1986. Hong Kong University Press, Hong Kong. KLEEMANN, K. H., 1996, Biocorrosion by bivalves. Marine Ecology, 17(1-3): 145-158. LAMY, E., 1923, Révision des Petricola vivants du Muséum National d'Histore Naturelle de Paris. Journal de Conchyliologie, 67(4): 309- 359. MIKKELSEN, P. M. 8 R. BIELER, 2000, Marine bivalves of the Florida Keys: discovered biodiversity. Pp. 367-387, in: L. HARPER, J. D. TAYLOR 4 J. A. CRAME, eds., The evolutionary biology of the Bivalvia. Geological Society, London, Special Publications 17. MIKKELSEN, P. M 8 R. BIELER, 2004, Interna- tional Marine Bivalve Workshop 2002: Intro- duction and Summary. In: В. BIELER 8 Р. M. MIKKELSEN, eds., Bivalve Studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 241-248. MORTON, B., 1983, Evolution and adaptive ra- diation in the Gastrochaenacea (Bivalvia). Jour- nal of Molluscan Studies, Supplement 12A: 117-121. MORTON, B., 1990, Corals and their bivalve bor- ers — the evolution of a symbiosis. Pp. 11-46, in: B. MORTON, ed., The Bivalvia — Proceedings of a Memorial Symposium in Honour of Sir Charles Maurice Yonge, Edinburgh, 1986. Hong Kong University Press, Hong Kong. 355 pp. MORTON, B., 1993, How the ‘forceps’ of Lithophaga aristata (Bivalvia, Mytiloidea) are formed. Journal of Zoology, London, 229: 609- 621. MORTON, В. & Р. J. В. SCOTT, 1988, Evidence for chemical boring in Petricola lapicida (Gmelin, 1791) (Bivalvia: Petricolidae). Jour- nal of Molluscan Studies, 54: 231-237. NARCHI, W., 1974, Functional morphology of Petricola (Rupellaria) typica (Bivalvia: Petricolidae). Marine Biology, 27: 123-129. NARCHI, W., 1975, Functional morphology of a new Petricola (Mollusca Bivalvia) from the lit- toral of Sao Paulo, Brazil. Proceedings of the Malacological Society, London, 41: 451-465. NIELSEN, C., 1976, Notes on the boring bivalves from Phuket, Thailand. Ophelia, 15(2): 141- 148. NIELSEN, C., 1986, Fauna associated with the coral Porites from Phuket, Thailand. (Part 1): Bivalves with description of a new species of Gastrochaena. Phuket Marine Biological Cen- ter Research Bulletin, 42: 24 pp. PURCHON, R. D., 1955, The functional morphol- ogy of the rock-boring lamellibranch Petricola pholadiformis Lamarck. Journal of the Marine Biological Association of the United Kingdom, 34: 257-278. REDFERN, C., 2001, Bahamian Seashells. A Thousand Species from Abaco, Bahamas. Bahamianseashells.com, Inc., Boca Raton, Florida. 280 pp., 124 pls. ROBERTSON, P. B., 1963, A survey of the ma- rine rock-boring fauna of southeast Florida. M.Sc. Thesis, University of Miami, Coral Cables. 169 pp. SCOTT, P. J. B., 1985, Aspects of living coral associates in Jamaica. Proceedings of the 5" International Coral Reef Congress, Tahiti, 1985, 5: 345-350. SCOTT, P. J. B., 1988a, Distribution, habitat and morphology of the Caribbean coral- and rock- boring bivalve, Lithophaga bisulcata d’Orbigny (Mytilidae: Lithophaginae). Journal of Mollus- can Studies, 54: 83-95. SCOTT, P. J. B., 1988b, Initial settlement behaviour and survivorship of Lithophaga bisulcata d'Orbigny (Mytilidae: Lithophaginae). Journal of Molluscan Studies, 54: 97-108. SOOT-RYEN, T., 1955, A report on the family Mytilidae (Pelecypoda). Allan Hancock Pacific Expeditions, 20(1): 1-175. TURNER, R. D. & K. J. BOSS, 1962, The genus Lithophaga in the western Atlantic. Johnsonia, 4: 81-116. WARMKE, С. L. & К. T. ABBOTT, 1971, Carib- bean seashells. A guide to the marine mollusks of Puerto Rico and other West Indian ilslands, Bermuda and the lower Florida Keys. Livingston Publishing Company, Narberth, Pennsylvania. 348 pp. WILSON, B. R. & R. TAIT, 1984, Systematics, anatomy and boring mechanisms of the rock- boring mytilid bivalve Botula. Proceedings of the Royal Society of Victoria, 96(3): 113-125. YONGE, C. M., 1955, Adaptation to rock boring in Botula and Lithophaga (Lamellibranchia, Mytilidae) with discussion on the evolution of this habitat. Journal of Microscopical Science, 96(3): 383-410. Revised ms. accepted 31 October 2003 MALACOLOGIA, 2004, 46(2): 355-379 COMPARATIVE MORPHOLOGICAL STUDY OF FOUR SPECIES OF BARBATIA OCCURRING ON THE SOUTHERN FLORIDA COAST (ARCOIDEA, ARCIDAE) Luiz Ricardo L. Simone! & Anton Chichvarkhin? ABSTRACT A detailed study on the morphology of the arcid genus Barbatia s.l. is performed, based on the common species occurring in Florida, complemented by samples from Brazil. The species are: B. cancellaria, B. candida, B. dominguensis, and B. tenera. The primary goal of this project is to collect comparative morphological data (especially on the internal anatomy) suitable for use in phylogenetic analysis. A complete descriptive and systematic treatment of these species is presented. A small phylogenetic analysis, based on nine characters (19 states) and seven taxa, demonstrates a closer relationship of B. cancellaria with B. can- dida, and of В. dominguensis with В. tenera. Barbatia s.l. is found to be monophyletic. Keywords: specific differentiation, phylogeny, distribution, systematics. INTRODUCTION It is often thought that bivalve anatomy is very conservative and therefore of limited use for resolving systematic problems. To test this hypothesis, a study including four sympatric species that most malacologists consider to belong to a single arcid genus was undertaken. The genus is Barbatia Gray, 1842 (type spe- cies Arca barbata Linné, 1758, by subsequent designation of Gray, 1857, from the Mediter- ranean). The four species examined are com- monly collected in intertidal and subtidal areas of rocky environments on the Florida coast, as well as in the tropical western Atlantic. Ad- ditional information about arcid biology and systematics was provided by Lamy (1907), Reinhart (1935), Heath (1941), Sullivan (1961), Coelho 8 Campos (1975), and Boyd (1998). The taxonomy of the various members of Arcidae is mostly based on conchology, as the anatomical knowledge is sparse and present in few papers (e.g., Heath, 1941). The four Barbatia species studied here are commonly allocated in different subgenera, with B. cancellaria and B. candida mostly referred to the subgenus Barbatia s.s. (alternatively, B. candida has been placed in Cucullaearca Conrad, 1865; type species: Byssoarca lima Conrad, 1848, by the subsequent designation of Stoliczka, 1871; Upper Cretaceous, east- ern United States), B. dominguensis to the subgenus Acar Gray, 1865 (type species: Arca gradata Broderip & С. В. Sowerby I, 1829, by the subsequent designation of Stoliczka, 1871; eastern Pacific), and B. tenera to the subge- nus Fugleria Reinhart, 1937 (type species by original designation: F. pseudoillota Reinhart, 1937; Pliocene of Florida). Our paper provides detailed comparative descriptions of the mor- phology of some arcid representatives, taxo- nomic treatments, and a preliminary application of these data in a small phylogenetic study. The objective of this paper is to perform a detailed anatomical study, developing the analysis ofthe data under a comparative sce- nario, showing that the study of the anatomy is valuable in comparative biology and impor- tant source of data. MATERIALS AND METHODS The specimens were collected during snor- keling and scuba dives in several localities of the Florida Keys (Mikkelsen & Bieler, 2004: fig. 1), and were maintained alive for several days. Some of the specimens were narcotized using magnesium hydroxide in the field laboratory and subsequently dissected. Other specimens were fixed in 96% ETOH, and after some days transferred to 70% ETOH for dissection at MZSP or FMNH. The dissections were per- formed using standard techniques, with speci- mens immersed in sea water or fixative. Images ‘Museu de Zoologia da Universidade de Säo Paulo, Cx. Postal 42594, 04299-970 Sao Paulo, SP, Brazil; Irsimone@usp.br “Russian Academy of Sciences, Vladivostok, Russia 399 356 SIMONE & CHICHVARKHIN were obtained using a standard digital camera directly or though a microscope; drawings were made with the aid of a camera lucida. For shell measurements, length indicates the anteropos- terior distance; lateral indicates the maximum inflation of the articulated valves; height indi- cates the dorsoventral distance originating from highest region of the umbo. The phylogenetic analysis was performed using the computer program Hennig86 (Farris, 1988) by means ofthe interface Tree Gardener (Ramos, 1998). Two arcid species were also included to test monophyly of the genus Barbatia: (1) Arca zebra (Swainson, 1833). U.S.A ; Florida; Florida Keys, Monroe County, 24°39.3’N, 81°18.2’W, “The Horseshoe’ site, bayside of West Summerland Key (Spanish Harbor Keys), 4 specimens, MZSP 36100 (FK-626, Simone coll. 26/vii/02). (2) Anadara notabilis (Róding, 1798). BRAZIL; Bahia; Salvador, Ribeira beach, MZSP 28481, 15 specimens (Simone coll., 24-27/ 1/1997). А non-arcid filibranch was used as an outgroup: /sognomon bicolor (С. В. Adams, 1845), Isognomonidae (Martins, 2000). This species is operationally used as the outgroup (rooting), the other two arcids (A. zebra, A. notabilis) are operationally analyzed as part of the ingroup. This procedure is undertaken for testing the monophyly of the genus Barbatia, as represented here. The anatomi- cal study of these three non-Barbatia species was performed in detail similar to that of the Barbatia species described herein. The following abbreviations are used in the figure captions: aa, anterior adductor muscle; an, anus; ao, abdominal organ; ap, anterior pedal protractor muscle; ar, anterior retractor muscle of foot; au, auricle; bf, byssal furrow of foot; by, byssus; ce, cerebral ganglion; cp, central gastric pad; cv, ctenidial vein; dd, ducts to digestive diverticula; dh, dorsal hood; er, esophageal rim; es, esophagus; ey, eye of mantle edge; ft, foot; ga, gill edge attached by cilia; gi, gill; gm, gill longitudinal muscle; gp, gill projection; gs, gastric shield; hi, hinge; ia, intestinal and style sac apertures; id, inner demibranch; if, inner fold of mantle edge; ih, inner hemipalp; in, intestine; ki, kidney; mb, mantle border; mf, middle fold of mantle edge; ml, mantle lobe; mo, mouth; ne, nephrostome; nv, nerve; od, outer demibranch; of, outer fold of mantle edge; oh, outer hemipalp; pa, pos- terior adductor muscle; pc, pericardium; pe, periostracum; pg, pedal ganglion; pm, pallial muscle; pp, palps; pr, posterior retractor muscle of foot; pt, pedal tentacle; rt, rectum; sa, gastric sorting area; sh, shell; ss, style sac and proximal portion of intestine; st, stomach; tm, connective between cerebral-pleural gan- glia with visceral ganglion; ty, pair of typhlo- soles separating intestine and style sac; um, umbo; ve, ventricle; vg, visceral ganglia; vm, visceral mass. Abbreviations of institutions: AMNH, Ameri- can Museum of Natural History, New York; FMNH, Field Museum of Natural History, Chi- cago; ММК), Museu Nacional da Universidade Federal do Rio de Janeiro; MZSP, Museu de Zoologia da Universidade de Sao Paulo. SYSTEMATICS Barbatia cancellaria (Lamarck, 1819) (Figs. 1-7, 33-36, 43-51) For additional synonymy, see Lamy (1907: 55). ?Barbatia barbata: Heath, 1941: 294, pl. 5, figs. 2, 7; pl. 15, fig. 12 (non Linné, 1758). Barbatia (Barbatia) cancellaria: Warmke & Abbott, 1962: 158, pl. 30, fig. j; Rios, 1970: 151; Andrews, 1971: 150, fig.; Abbott, 1974: 421— 422, fig. 4966; Rios, 1975: 192, pl. 61, fig. 939; Humfrey, 1975: 210, pl. 23, fig. 11; Rios 1985: 208, pl. 75, fig. 1061; 1994: 230, pl. 80, fig. 1136; Redfern, 2001: 203, pl. 83, fig. 831. Barbatia cancellaria: Diaz & Puyana, 1994: 46, fig. 24. Description Shell (Figs. 1-7). Medium to large size, to 100 mm. Color pale reddish brown. Main sculp- ture composed of narrow radial threads, separated from each other by furrows of approximately same width as threads (Fig. 3); commarginal sculpture weak, composed mainly of undulations. Periostracum some- what thick, with many scales along radial threads; scales longer close to shell border. Most specimens with a pattern (most clear in posterior region) of one row of long scales followed by five rows of short scales (Figs. 1-5). Periostracum extending beyond shell edges (Fig. 5). Umbos flat, located between anterior and middle thirds of hinge. Inner surface glossy, brownish violet on borders, becoming paler towards umbo. Hinge vari- able, but generally with about 30 teeth lo- cated just anterior to umbo and posteriorly (Figs. 5-7); three anteriormost teeth broader, FLORIDIAN BARBATIA MORPHOLOGY 357 weakly arched, tilted towards anterior; next three teeth with similar shape to those de- scribed, but narrower; following 8-10 teeth abruptly different, shorter, narrower, situated perpendicularly to outer edge of hinge, gradually becoming thicker, slightly longer and more tilted towards posterior; last 5-6 teeth broader, almost horizontal. Soft Part Color (Figs. 33-36): Mantle border with a mixture of dark and pale brown spots, being darker towards edge (Fig. 33); some pale cream spots randomly distributed mostly along middle region of border. Pos- terior region of mantle border more pig- mented than more anterior region, including inner surface to base of gills; this posterior- inner surface pigmented uniform beige, hav- ing irregularly sized, randomly distributed white spots (Fig. 33). This pattern also cov- ering posterior and dorsal surface of poste- rior adductor muscle and rectum; anal papilla FIGS. 1-7. Barbatia cancellaria shell. FIG. 1: Left view, MZSP 36105; FIG. 2: Same, dorsal view; FIG. 3: Same, detail of sculpture in posterior region; FIG. 4: Outer view of both valves, MZSP 32336; FIG. 5: Same, inner view; FIG. 6: Right valve; MZSP 36105, detail of hinge; FIG. 7: Same, MZSP 36211. Scale bars = 5 mm. 358 abruptly preceded by а white region. Foot and ventral region of visceral mass pig- mented by a mosaic of brownish purple, coa- lescent spots, in a pale cream base (Fig. 36). Gills uniformly colored pale purple; their pos- terior, projected region colored by brownish purple, having small to large white spots ran- domly distributed, more concentrated along middle region (Figs. 33, 34). Remaining re- gions of soft parts lacking any special pig- mentation. Main Muscle System (Figs. 43, 44): Anterior adductor muscle elliptical in cross-section, located in anterodorsal region. Posterior adductor muscle rounded in cross-section, located in posterodorsal region, almost twice as large as anterior adductor muscle. Pair of pedal protractor muscles very narrow, originating in middle region of ventral edge of anterior adductor muscle, running towards posterior attached to inner surface of integu- ment splayed insertion area near anterior foot base. Pair of anterior retractor muscles of foot narrow; originating in a small area just posterior and ventral to anterior adductor muscle, running towards posterior and ven- tral close to median line, inserting in ante- rior region of foot. Pair of posterior retractor muscles of foot very large and thick, occu- pying about 2/3 of visceral mass volume; originating in umbonal cavity just anterior to posterior adductor muscle, extending about twice adductor muscle area towards ante- rior; running towards ventral and anterior closely attached to one another (except for a narrow dorsal area along median line); in- serting along middle and posterior regions of foot. Visceral wall of visceral cavity thickly muscular, serving as base for foot. Gill muscles running in a pair along ventral sur- face of ctenidial vein, between two demi- branchs (Fig. 48). Foot and Byssus (Figs. 36, 43, 44): Foot some- what narrow and long. Byssal furrow deep and broad, running all along its ventral sur- face to pedal tip (Fig. 36), in median-poste- rior region of foot becoming broader, contouring byssus. A pair of pedal tentacles located preceding this broader region, close to byssus (Figs. 36, 43: pt); this pair of ten- tacles absent in some specimens or repre- sented by a pair of flaps in others. Each tentacle slender, stubby, located in byssal furrow edge. Byssus composed of several SIMONE & CHICHVARKHIN fused fibers (similar to Figs. 37-39); their dorsal region forming a deep concavity, its edge formed by several, aligned, flat, free filaments, which fuse with each other after some distance; left series of filaments sepa- rated from right series by deep anterior and posterior furrows; these free portions remain- ing deeply introduced in byssal gland and compressed by posterior retractor muscles of foot, each filament encased in special folds of a central, large convexity (similar to Fig. 41). Middle region of byssus forming a single, solid stem. Distal portion of byssus irregularly splayed, attached to hard sub- strata. Mantle: Edge of each mantle lobe entirely free (except in dorsal region). Mantle edge thick, mostly tri-folded (Fig. 49). Color as described above. Middle fold shorter, covered by inner and outer folds; inner fold tall, undulating, typically situated towards median line, touch- ing its pair. Remaining mantle areas very thin and transparent. Flap of mantle fitting inside hinge flat and short. Pallial Cavity (Figs. 43, 46, 48): Pallial cavity surrounding most of outer region, except a narrow area anterior to posterior adductor muscle. Gill large, filibranch, having thou- sands of narrow, uniform filaments. Inner and outer demibranch of equal size. Anterior 2/3 of gill attached to visceral mass, outer basal edge of outer demibranch and inner basal edge of inner demibranch attached, respec- tively, to mantle and visceral mass, by means of cilia (Fig. 48: ga). Posterior 1/3 of each gill attached dorsally to a long, muscular, mobile stalk (Figs. 33, 43, 46). These stalks originating side by side from ventral surface of posterior adductor muscle, relatively broad, tapering distally and posteriorly; in this region, outer basal edge of outer demibranch and inner basal edge of inner demibranch with capacity of attaching, respectively, to mantle and to its pair, by means of cilia. These stalks pigmented as described above. Visceral Mass (Fig. 44): Visceral mass broad, thick, posterior half entirely filled by poste- rior pedal retractor muscles. Anterior half filled by visceral organs. Gonad pale beige, mostly located externally, surrounding diges- tive diverticula and tubes. Digestive diver- ticula brownish green, located surrounding stomach, below umbonal cavity. FLORIDIAN BARBATIA MORPHOLOGY 359 Circulatory and Excretory Systems: Heart rela- tively small, located just anterior to origin of posterior retractor muscle of foot, in umbonal cavity (Fig. 44). Auricles originating in middle portion of gill ctenidial vein, with narrow an- terior and posterior portions of ctenidial vein. Each auricle connected to accessory au- ricles as expansions of ventricle (Fig. 50) in middle region of a broad outer surface, gradually tapering and inserting in ventricle. Ventricle narrow, surrounding intestine. Kid- ney mostly solid, brown, located along pos- terior half of gill insertion on visceral mass, on their inner edge (Fig. 46), gradually be- coming broader, crossing to region dorsal to gill on visceral mass. Nephrostome a small slit located in anterior extremity of each kid- ney (Fig. 46: ne). Digestive System (Fig. 44): Palps located con- touring anterior edge of pallial cavity, just posterior to anterior adductor muscle (Figs. 34, 35), surrounding anterior edge of gill. Palps dorsoventrally very long, antero- posteriorly short; dorsal end somewhat rounded, of uniform width along their length. Palp outer surface smooth. Palp inner sur- face with a special arrangement of folds simi- lar in outer and inner hemipalps (Figs. 35, 45); folds of dorsal half of palp transverse, situated perpendicular to posterior palp edge; folds gradually curving, situated lon- gitudinally; ventral palp half with longitudi- nal (dorsoventral) folds restricted to region close to intersection of both hemipalps, run- ning parallel to one another towards mouth. Palps’ ventral connection with each other broad, forming a sac that covers antero- ventral region of visceral mass (Figs. 43, 45). An inner smooth area contouring entire pos- terior edge of palps, broader in ventral half (Fig. 45). Mouth located at anterior end of ventral connections between palps. Esopha- gus running close to posterior surface of anterior adductor muscle, almost its entire length, relatively narrow, inner surface with low, narrow longitudinal folds. Stomach dor- soventrally elongated, situated transversely in middle region of visceral mass; stomach dorsal region spherical, positioned horizon- tally, abruptly curving towards ventral, nar- rowing gradually, continuing as style sac. Gastric inner surface (Fig. 51) separated from esophageal surface by ventral, narrow, low rim, which extends toward posterior in right side; immediately posterior to rim a broad, concave sorting area, bearing sev- eral transverse (situated anteroposteriorly) uniform folds; aperture to digestive diver- ticula as two pairs, each positioned between these folds, two on each side; gastric cen- tral pad located transversely, in mid-ventral gastric surface, as posterior limit of previ- ously described sorting area; central pad possessing longitudinal (left to right) low, uniform folds, left region of central pad fit- ting in dorsal hood, anterior and right bor- ders somewhat tall, posterior border connected to typhlosole, which runs towards intestine. Pair of typhlosoles incompletely separating intestine from style sac; anterior typhlosole originating broadly, attached to central pad; posterior typhlosole narrow, Originating as low, weak fold; a low, narrow fold dorsally surrounding aperture of style sac and intestine, located in posterior-left side of ventral gastric surface. Gastric shield located in middle region of posterodorsal gastric inner surface. Dorsal hood low, lo- cated in middle-left side of gastric dorsal surface. Digestive diverticula as described above. Intestine as continuation of style sac, curving abruptly towards right, performing three strong, but short loops located along right surface of style sac. Intestine broad in region after style sac, abruptly becoming relatively narrow, with uniform width along its length. After these loops, intestine run- ning posterodorsally to pericardial region; running horizontally though ventricle and narrowly between both posterior retractor muscles of foot. Rectum exposed in excur- rent chamber (although covered by integu- ment), running in dorsal and posterior surface of posterior adductor muscle. Anal papilla (Figs. 43, 46, 47) a long projection towards posterior, narrowing gradually to a pointed tip; located in posteroventral region of adductor muscle; anal aperture long, sub- terminal, ventral, possessing a small trans- verse fold in anterior end. Central Nervous System: Pair of cerebral-pleu- ral ganglia located compressed between esophagus and anterior adductor muscle posterior surface, situated laterally at broad distance from each other. Pair of pedal gan- glia large, located close to each other in an- terior surface of visceral mass (Fig. 44), just dorsal to anterior adductor muscle; their con- nective with cerebral-pleural ganglia very thin, relatively short. Pair of visceral ganglia located in base of both gill projections (Fig. 46) in anterior region of posterior adductor 360 SIMONE & CHICHVARKHIN muscle, close to each other and to median line; a broad nerve running along gill projec- tion axis; their connective with cerebral-pleu- ral ganglia very thin, running close to integument. А conspicuous pair of nerves originating from posterior end of visceral ganglia, running along gill stalks to their dis- tal ends. Abdominal organ as a pair of bulg- ing masses covering ventral surface of posterior adductor muscle, just posterior to visceral ganglia. Measurements of Dissected Specimens (length, lateral, height in mm): MZSP 36105(1): 47.8 by 19.4 by 26.4; (2): 47.3 by 22.4 by 28.0; MZSP 36258(4): 34.4 by 13.5 by 18.7. Distribution North Carolina, USA, to Bahia, Brazil. Habitat Rocky, from intertidal zone to about 10 m depth. Material Examined U.S.A.; Florida; Florida Keys; Monroe County, roadside quarry N of Keys Marine Laboratory, 24°49.78’М, 80%48.51'W, 2-5 т depth, MZSP 36105, 6 specimens (Simone coll. 22/vii/2002); “Long Key Artificial Reefs”, oceanside of Long Key, 24°44.78’М, 80°50.00’W, 7 m depth, MZSP 36277, 4 speci- mens (FK-621, Simone coll., 17/vii/2002); W side of Pigeon Key, 24°42.2’М, 81°09.3’W, MZSP 26171, 2 specimens (sta. 647, Simone coll., 28/vii/2002); Bahia Honda State Park, oceanside, just E of old bridge, 24°39.25’М, 81°16.83’W, MZSP 36268, 1 specimen (FK- 632, Simone coll., 22/vi/2002); Old Dan Bank, bayside of Long Key, 24°50.45’N, 80°49.63’W, MZSP 36258, 4 specimens (FK-620, Simone col, 16-18/vii/2002); “The Billboard” site, oceanside, Lower Matecumbe Key, MM 74.5, 24°51.4’М, 80°43.7’W, MZSP 36261, 4 speci- mens (FK-642, Simone coll., 23/vii/2002); oceanside off Craig Key, 24°49.81'N, 80%45.73'W, MZSP 36197, 2 specimens (FK- 640, Simone coll., 23/vii/2002); “The Horse- shoe” site, bayside of West Summerland Key (Spanish Harbor Keys), ММ 35, 24°39.3’N, 81°18.2’W, MZSP 36211, 4 specimens (FK- 629, Simone 8 Leal coll., 21-25/vii/2002), MZSP 36101, 7 specimens (FK-629, Simone coll., 26/vii/2002). BRAZIL; Pernambuco, Fernando de Noronha Archipelago (Simone 8 Souza coll.); Porto Beach, MZSP 31205, 2 specimens, 4 shells, MZSP 31247, 1 shell (17/ vii/1999); Atalaia Beach, MZSP 31166, 1 speci- men (18/vii/1999); Buraco do Inferno, Rata Island, MZSP 31071, 21 specimens (19/vii/ 1999); Bahia; Abrolhos Archipelago, MZSP 39837, 1 specimen (Luiz Pinni Neto coll., v/ 1958); Coroa Vermelha, MZSP 23698, 1 speci- men (L. Рип! NT. Coll., 11/1957); Pedra da Lixa, Parcel de Paredes, MZSP 32336, 5 specimens (Souza & Goncalves coll., /2000). Barbatia candida (Helbling, 1779) (Figs. 8-14, 26, 32, 37-39, 52-59) For additional synonymy: Coelho & Campos (1975: 40). Barbatia (Barbatia) candida: Warmke 8 Abbott, 1962: 158, pl. 30, fig. 1; Rios, 1970: 150-151; Andrews, 1971: 150-151, fig.; Abbott, 1974: 421, fig. 4965; Humfrey, 1975: 210, pl. 23, fig. 6; Redfern, 2001: 203, pl. 83, fig. 832. Barbatia (Cucullaearca) candida: Heath, 1941: 294, pl. 5, figs. 4, 5; pl. 7, fig. 9; Rios, 1975: 192, pl. 62, fig. 940; 1985: 209, pl. 75, fig. 1063; 1994: 230, pl. 80, fig. 1138. Ваграйа candida: Merlano & Hegedus, 1994: 46, fig. 23. Description Shell (Figs. 8-14, 32): Medium to large size, to 100 mm. Color beige to pale brown. Main sculpture composed of narrow radial threads (Figs. 8, 12, 32) separated from each other by furrows of approximately same width as threads; commarginal sculpture of only growth lines; threads relatively uniform in size, except posterior threads somewhat broader. Periostracum relatively thick, with many scales along radial threads, scales longer close to shell border, varying in shape from simple (in southern specimens, Fig. 32) to bifid (in northern specimens, Fig. 12). Periostracum extending beyond shell edges, mainly in region between threads (Figs. 9, 11). Umbos flat to tall, located between an- terior and middle thirds of hinge (Fig. 10). Inner surface glossy, white (Figs. 9, 11). Hinge (Figs. 9, 11, 13, 14) variable, but gen- erally with about 30 teeth located both ante- rior and posterior to umbo; three anteriormost teeth broader, straight, weakly titted towards anterior; posteriorly a series of similar-shaped teeth, becoming gradually smaller until umbo level, afterwards gradu- FLORIDIAN BARBATIA MORPHOLOGY 361 ally increasing towards posterior, becoming Foot and Byssus (Figs. 37-39). As in B. weakly tilted posteriorly; last 5-6 teeth cancellaria, except pair of pedal tentacles broader, strongly tilted posteriorly. absent, and with distinct color as described above. Foot convexity for byssal accommo- Soft Part Color. Mantle border gill stalks and dation (Fig. 41) similar to that in В. cancellaria. exposed portion of foot with similar colora- tion to those of B. cancellaria, differing in Mantle (Figs. 52, 55): Mantle edge thick but being paler, with beige base color beige. thinner than that in B. cancellaria, mostly tri- folded. Color as described above. Middle fold Main Muscle System (Fig. 54): Essentially as shorter, covered by inner and outer folds; in B. cancellaria. Other details in Heath inner fold tall, undulated, typically situated (1941: pl.7, fig. 9). towards median line, touching its pair. Most FIGS. 8-14. Barbatia candida shell. FIGS. 8-10: АММН 298092; FIG. 8: Left valve, outer view; FIG. 9: Same, inner view; FIG. 10: Whole specimen, dorsal view; FIGS. 11, 12: ЕММН 183584; FIG. 11: Right valve, inner view; FIG. 12: Detail of sculpture and periostracal scales in posterior region; FIGS. 13, 14: Detail of hinge; FIG. 13: Right valve, MZSP 32304 from Bahia, Brazil; FIG. 14: Left valve, MZSP 32336. Scale bars = 5 mm. 362 SIMONE & CHICHVARKHIN specimens with a short portion with clear undulations in ventral region of incurrent canal and in mid-ventral region. Pallial Cavity: As in B. cancellaria, except as follows. Muscular and mobile stalk of gill nar- rower and differently pigmented (as described above). Gill filaments of each demibranch maintained in their position by a transverse membrane restricted to ventral half of fila- ment, this membrane connecting inner branch of each filament to outer branch in both inner and outer demibranchs (Figs. 42, 57). Visceral Mass: As in B. cancellaria. Circulatory and Excretory Systems (Fig. 58): As in В. cancellaria, except т having shorter and paler-colored kidney. Digestive System (Fig. 53): Most characters as in B. cancellaria, except as follows. Palps with same location and outer features, in- cluding sac-like ventral region preceding mouth (Figs. 52, 56); inner folds arrangement with longer portion with transverse (antero- posterior) folds, ventral portion smooth, lack- ing folds (Fig. 56). Gastric inner surface (Fig. 58) with esophageal rim surrounding entire esophageal aperture, additionally with a branch at right surrounding a distinct duct to digestive gland, being broader, forming a small dorsal chamber. Central pad narrower, originating from right branch of esophageal rim, passing anteriorly to aperture of dorsal hood; central pad disconnected from ante- rior typhlosoles. Anterior typhlosole with about same width as posterior typhlosoles, and originating from a longitudinal, low, nar- row fold running along left gastric surface. Posterior typhlosole as continuation of fold separating gastric and style sac chambers. Intestine as continuation of style sac, abruptly curving towards anterior, perform- ing thereafter a single wide loop located an- terior and to right of style sac. Intestine relatively narrow, of uniform width along its length. Following these loops, intestine run- ning posterodorsally to pericardial region close to style sac; running horizontally though ventricle and narrowly between both poste- rior pedal retractor muscles. Anal papilla (Figs. 52, 59) as a long projection towards poste- rior, narrowing gradually to a pointed tip; lo- cated in posteroventral region of adductor muscle; its aperture wide, from its insertion in adductor muscle to its apex (Fig. 59). Central Nervous System: As in B. cancellaria. Measurements of Dissected Specimens (length, lateral, height in mm): MZSP 32304: 46.0 by 20.0 by 31.8. Distribution North Carolina, USA, to Santa Catarina, Brazil. Habitat Rocky, from intertidal zone to about 10 m depth. Material Examined U.S.A.; Florida; Miami; Miami Causeway, FMNH 183584, 1 shell (Koto coll., 1940); Sol- dier Key, FMNH 183583, 2 shells (Koto coll. 1940); Florida Keys; due E of New Ground, near Tower, 24°40’51"М, 82*16'02"W, 12.2 m depth, AMNH 298092, 2 specimens (FK-080, otter trawl; Mikkelsen et al. coll.; 22/iv/1997); Monroe County, east Turtle Shoal, ocean side off Grassy Key, 24°43’15"М, 80%55'42"W., FMNH 279038, 1 shell (FK-242, Bieler & Mikkelsen coll., 04/viii/1999); Looe Key, FMNH 279041, 1 shell (FK-275, Bieler et al. coll., viii/ 1999); 24°32.87’М, 81°24.41’W, less than 3 m depth, FMNH 279040, 3 shells (FK-276, Bieler et al. coll., 21/viii/1999); 24°32.77’М, 81°24.23’W, 7 т depth, FMNH 279039, 3 shells (FK-262, Bieler & Mikkelsen coll., 11/ vii/1999); 24*32.87'N, 81°24.41’W, less than 3 т depth, FMNH 279052, 1 shell (FK-276, Bieler et al. coll., 21/viii/1999); Coffins Patch coral reef, 24°41’05"М, 80°57’28"W, 5 т depth, FMNH 279043, 2 shells (FK-236, Bieler et al. coll., 2/viii/1999); Molasses Reef, 25°00.55’N, 80°22.58’W, FMNH 279042, 1 shell (FK-200, Bieler & Mikkelsen coll., 07/iv/1999); Key Vaca, FMNH 155511, 1 shell (Nelson coll.); Missouri Key, FMNH 183596, 7 shells, FMNH 183580, 1 shell (Koto coll.). BRAZIL; Bahia; Salvador, Banco da Panela, 16-20 т depth, MZSP 28459, 1 specimen (Simone coll., 26/ii/1997); Farol da Barra, MZSP 28539, 1 specimen (Simone coll., 22—28/ii/1997); Parcel de Paredes, Pedra da Lixa, 1-5 m depth, MZSP 32304, 5 specimens, 3 shells (Souza Jr. & Gongalves col, i/2000); Abrolhos Archipelago, MZSP 28988, 1 specimen (К. Moura coll., 9— 15/1/1998); Rio de Janeiro; Arraial do Cabo, Prainha, ММК 9775, 2 specimens (Simone & Costal col, 27/ii/2003); Cabo Frio, MZSP 35243, 4 specimens (P. Goncalves coll., v/ 2002); Niterdi, Itaipu Beach, MZSP 28739, 5 FLORIDIAN BARBATIA MORPHOLOGY 363 specimens (Simone coll. 12/vii/1997); Angra dos Reis, Vila Velha Beach, MZSP 18353, 25 specimens; Sáo Paulo, Ubatuba, MZSP 23681, 1 specimen (Klappenbach coll.); Saco da Ribeira, MZSP 23679, 1 specimen (Montouchet coll., 20/vii/1967); Ilha Bela, Sao Sebastiáo canal, MZSP 28663, 3 specimens (F.L.Silveira coll., 17/v/1997); Sao Sebastiäo, Baraqueçaba Beach, MZSP 30892, 4 speci- mens (Simone coll., 27/ix/1998); Ilha Alcatrazes, 8 m depth, MZSP 28314, 2 speci- mens (Simone coll., ix/1996). Barbatia dominguensis (Lamarck, 1819) (Figs. 15-25, 60-65) For additional зупопуту: Lamy (1907: 80-82). ?Асаг reticulata: Heath, 1941: 294, pl. 6, fig. 9; pl. 7, fig. 8 (non Gmelin, 1791). Barbatia (Acar) dominguensis: Warmke & Abbott, 1962: 158, pl. 30, fig. d; Rios, 1970: 151; Andrews, 1971: 151, fig.; Abbott, 1974: 422, fig. 4967; Rios, 1975: 192, pl. 62, fig. 941; Humfrey, 1975: 210, pl. 23, fig. 5); Rios, 1985: 208-209 (pl. 75, fig. 1062); 1994: 230, pl. 80, fig. 1137; Diaz & Puyana, 1994: 46, fig. 25; Redfern, 2001: 203, pl. 83, fig. 833. Description Shell (Figs. 15-20): Small to medium size, to 25 mm. Color pale beige. Sculpture com- posed of well-developed radial and commarginal threads, both of equal strength or with commarginal threads more weakly developed; with small nodes at intersections of radial and commarginal threads. Peri- ostracum thin, transparent, lacking scales, restricted to shell edges. Umbos flat, located from between anterior and middle thirds of hinge to middle portion of it. Inner surface glossy, white (Fig. 17). Hinge with about 15- 20 teeth located just anterior to umbo and posteriorly; 3-4 anteriormost teeth larger, tilted towards anterior; immediately follow- ing posterior teeth abruptly changing in size and slightly more dorsally located, these teeth small, perpendicular to dorsal edge, gradually becoming larger and tilted towards posterior (Figs. 17, 20). Soft Part Color. Mostly pure white or pure pale cream. Some pale purple pigmentation in posterior region of gill and posteroventral region of mantle border (Figs. 23-25). Mantle border surrounded externally by a series of minute, black eyespots, located in somewhat regular distance from each other (Figs. 23- 25, 60); eyes more developed in posterior region (about 20 in each mantle lobe), scarce in ventral and anterior regions. Main Muscle System (Fig. 62): Similar to those in B. candida, with the following differences. Posterior adductor muscle proportionally larger. Pair of anterior pedal protractor muscles thicker and separated from integu- ment, running immersed in gonad. Foot and Byssus: Both structures very similar to those in B. cancellaria, with the following remarkable features. Extra-byssal portion of foot lacking pigmentation and proportionally smaller (Fig. 60). Byssus shorter, lacking posterior furrow separating left and right se- ries of dorsal byssal bands (Figs. 21, 22). Mantle: Mantle edge features broadly similar to those in B. cancellaria, but of different color. Edge relatively thinner (Fig. 65); posteroventral portion thicker, possessing a taller and thicker median pigmented fold. Outer fold bearing a series of minute, black- colored eyespots all along its length, on both mantle lobes (Figs. 23, 24, 60). Each eye located at a regular distance from next, equivalent to about five times its width. Larger eyes located at posterior edge, ven- tral eyes smaller, sometimes absent (Figs. 23, 25). Each eye composed of a spherical distal portion, black pigmented, followed by a very short stalk, positioning it subterminally in outer mantle edge fold (Fig. 24). More details of compound eyes described by Jan- Olof (1998: fig. 1). Pallial Cavity: General features similar to those in B. cancellaria, with the following notable characters. Gill coloration different. Gill stalks supporting posterior third of gill proportionally more slender, and originating more anteriorly in posterior adductor muscle (Figs. 23, 60). Visceral Mass: As in B. candida. Circulatory and Excretory Systems: No notable distinctions to those described for B. cancellaria. Digestive System (Fig. 62): General organi- zation similar to that in B. cancellaria, with the following differences. Palps' inner sur- face with a special arrangement of folds simi- lar in outer and inner hemipalps (Fig. 61); 364 SIMONE & CHICHVARKHIN FIGS. 15-25. Barbatia dominguensis shell and anatomy. FIG. 15: Left view, MZSP 36167; FIG. 16: Same, dorsal view; FIG. 17: Same, inner view of left valve: FIG. 18: Left view, MZSP 36144; FIG. 19: Same, dorsal view; FIG. 20: Detail of hinge, left valve, MZSP 36147; FIG. 21: Isolated byssus, left view; FIG. 22: Same, apical view; FIG. 23: Detail of posterior region, left view, left valve and mantle lobe removed; FIG. 24: Detail of mantle border of incurrent region; FIG. 25: Detail of anterior region, | с ей view, left valve and mantle lobe removed. Scale bars = 2 mm. FLORIDIAN BARBATIA MORPHOLOGY 365 folds of dorsal third of palp transverse, situ- ated perpendicular to posterior palp edge; folds gradually becoming oblique in middle third; ventral palp third smooth, lacking lon- gitudinal (dorsoventral) folds; region preced- ing mouth narrow, simple (lacking sac-like morphology). Esophagus running close to posterior surface of anterior adductor muscle, almost its entire length, relatively narrow, inner surface with low, narrow longi- tudinal folds. Gastric inner surface (Fig. 63) with esophageal rim restricted to lateral and dorsal region of esophageal insertion; an- other transverse fold, narrow, long, running ventrally just anterior to esophageal rim, longer at right. Concave anteroventral sort- ing area lacking left ducts to digestive diver- ticula; left duct to digestive diverticula single, located dorsal to a transverse fold which separates it from sorting area. Another small sorting area located in middle region of gas- tric dorsal surface. Central part smooth, lack- ing folds and any connection to anterior typhlosole. Anterior typhlosole narrow, sur- rounding posterior edge of central pad; pos- terior typhlosole weakly developed. Gastric shield large, occupying most of posterior gastric surface. Dorsal hood located in pos- terior region of stomach, its aperture narrow, probably containing a sphincter. Intestine curving gradually towards right and anterior, performing a single, broad loop. Intestine relatively narrow, with uniform width along its length to pericardial region; running hori- zontally though ventricle and narrowly be- tween both posterior retractor muscles of foot. Anal papilla (Fig. 64) a long projection towards posterior, narrowing gradually to a pointed tip; located in posteroventral region of adductor muscle; anal aperture long, sub- terminal, ventral, possessing thick edges. Central Nervous System: As in B. cancellaria, except in pair of connectives between cere- bral-pleural ganglia and visceral ganglia be- ing broader, running through visceral mass, gonad and digestive diverticula. Measurements of Dissected Specimens (length, lateral, height in mm): MZSP 36279: 23.0 by 11.0 by 13.6; MZSP 36167: 14.5 by 7.7 by 10.1; MZSP 26375: 18.0 by 9.3 by 11.4. Distribution North Carolina, USA, to Bahia, Brazil. Habitat Rocky, from intertidal zone to about 10 m depth. Material Examined U.S.A.; Florida; Florida Keys; “Long Key Ar- tificial Reefs”, oceanside of Long Key, Mon- roe County, 24°44.78’М, 80°50.00’W, MZSP 36279, 2 specimens (FK-621, Simone coll. 17/ vii/2002); Pigeon Key, 24°42.2’N, 81°09.3’W, MZSP 36167, 1 specimen (FK-657, Simone coll., 28/vii/2002); “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), MM 35, 24°39.3’М, 81°18.2’W, MZSP 36144, 1 specimen (FK-629, Simone coll., 26/vii/2002); Tennessee Reef Light, 24°44.75'N, 80°46.95’W, 4-7 т depth, MZSP 36147, 9 specimens (FK-648, Simone coll., 26/ vii/2002); Old Dan Bank, bayside of Long Key, 24°50.45'N, 80°49.63’W, MZSP 36260, 1 specimen (FK-620, Simone coll., 16-18/vii/ 2002). BRAZIL; Bahia; Abrolhos Archipelago, MZSP 26375, 2 specimens, MZSP 15566, 9 specimens (L. Pinni Nt. Coll., v/1958). Barbatia tenera (C. B. Adams, 1845) (Figs. 26-31, 40—42, 66-73) Barbatia (Fugleria) {епега: Warmke 8 Abbott, 1962: 158, pl. 30, fig. 9; Andrews, 1971: 151- 152, fig.; Abbott, 1974: 422; Rios, 1975: 192, pl. 62, fig. 942; Rios, 1985: 209, pl. 76, fig. 1064; 1994: 231, pl. 80, fig. 1140; Diaz « Puyana, 1994: 47, pl. 3, fig. 26; Redfern, 2001: 203, pl. 83, fig. 834. Description Shell (Figs. 26-31): Of medium size, to 50 mm. Color pale beige. Outline trapezoidal. Sculp- ture composed of well-developed radial and weakly developed commarginal threads; a small node at intersection of radial and commarginal threads; posterior threads broader than anterior threads. Periostracum relatively thick, with many scales between radial threads, scales longer close to shell border; scales brush-like in shape (Fig. 30), divided into 4-7 pointed projections. Umbos flat, located from between anterior and middle thirds of hinge to middle portion of it. Inner surface glossy, white (Fig. 27). Hinge with about 15-25 teeth located just anterior to umbo towards posterior; 3-4 anteriormost teeth larger, tilted towards anterior; next pos- terior teeth abruptly changing in size and 366 slightly more dorsally located, these teeth small, perpendicular to dorsal edge, gradu- ally becoming larger and tilted towards pos- terior (Figs. 27, 31). SIMONE 8 CHICHVARKHIN mented than more anterior region. This pat- tern also covering posterior and dorsal sur- face of posterior adductor muscle and rectum; anal papilla abruptly preceded by a white region. Foot (Fig. 41) and ventral re- gion of visceral mass pigmented uniform orange or yellow. Gills uniformly colored pale yellow (Fig. 42), including their posterior, Soft Part Color (Fig. 40-42): Mantle border uniform pale orange or yellow. Posterior re- gion of mantle border more weakly pig- FIGS. 26-32. Barbatia shell. FIGS. 26-31: Barbatia tenera shell; FIG. 26: Left view, MZSP 36278; FIG. 27: Same, right valve, inner view; FIG. 28: Same, dorsal view; FIG. 29: Left view, MZSP 36146; FIG. 30: Detail of sculpture and periostracal scales in posterior region, MZSP 36146; FIG. 31: Detail of hinge, right valve, MZSP 36278; FIG. 32: Barbatia candida, MZSP 28314, detail of sculpture and periostracal scales in posterior region (for comparison). Scale bars = 5 mm. FLORIDIAN BARBATIA MORPHOLOGY 367 FIGS. 33-42. Details of Barbatia anatomy. FIGS. 33-36: Barbatia cancellaria; FIG. 33: Detail of posterior region, left view, left valve and mantle lobe removed; FIG. 34: Region of right palp, right view, right valve removed, right mantle lobe deflected; FIG. 35: Same, outer hemipalp deflected to show inner surface; FIG. 36: Foot, ventral view; FIGS. 37-39: Barbatia candida; FIG. 37: Byssus, left view, detail of apical region; FIG. 38: Same, whole left view; FIG. 39: Same, apical view; FIGS. 40-42: Barbatia tenera; FIG. 40: Detail of posterior region, left view, left valve and mantle lobe removed; FIG. 41: Foot, left view, detail of byssal groove; FIG. 42: Left gill isolated, ventral view. Scale bars = 3 mm. 368 SIMONE & CHICHVARKHIN projecting region. Remaining regions of soft parts lacking any special pigmentation. Main Muscle System (Figs. 40, 66, 68): Simi- lar to those in В. dominguensis, with follow- ing differences. Posterior adductor muscle slightly larger than anterior adductor muscle. Pair of anterior pedal protractor muscles thin, fused with integument; their origins thin, sur- rounding ventral edge of anterior adductor muscle insertion in each valve. Foot and Byssus (Figs. 41, 66-68): Very simi- lar to those in B. cancellaria, with the follow- ing differences. Extra-byssal portion of foot lacking pigmentation (Fig. 41) and propor- tionally smaller than that in B. cancellaria (Figs. 66, 68). Byssus shorter, lacking pos- terior furrow separating left and right series of dorsal byssal bands. Mantle: Mantle edge features broadly similar to those in B. cancellaria. Color different, as described above (Fig. 40). Edge relatively thinner; posteroventral portion thicker, pos- sessing a taller, thicker median fold (Figs. 66, 70). Pallial Cavity (Figs. 66, 67): General features similar to those in B. cancellaria, with the following notable characters. Different gill coloring as described above (Fig. 42). Gill stalks supporting posterior third of gill pro- portionally more slender, originating more anteriorly on posterior adductor muscle (Figs. 66-68). Gill filaments with ascendant and descendant branches almost free from each other (Fig. 72). Visceral Mass (Fig. 68): As т В. dominguensis. Circulatory and Excretory Systems (Figs. 67, 73): No notable distinctions to those de- scribed for B. cancellaria. Digestive System (Figs. 68, 71, 73): General organization similar to those in B. cancella- ría, with following differences. Inner folds of palps (Fig. 69) transverse in dorsal and middle thirds, situated perpendicular to pos- terior palp edge; folds gradually becoming weakly oblique in ventral third; region pre- ceding mouth narrow, simple (lacking sac- like morphology), inner surface smooth. Esophagus relatively narrow, inner surface with low, narrow longitudinal folds. Gastric inner surface (Fig. 71) with esophageal rim indistinguishable, a very deep transverse (slightly oblique) furrow located at short dis- tance posterior to esophageal insertion, 2— 3 pairs of ducts to digestive diverticula originating from this furrow; central pad broad, located in central portion of gastric ventral surface, occupying about half of ven- tral surface area; a broad fold surrounding central pad anterior, right and posterior edges; anterior edge forming border of transverse furrow, posterior edge surround- ing anterior border of combined intestine- style sac origin; this fold extending transversly on left-dorsal gastric surface, surrounding anterior edge of gastric shield; a small, anteroposteriorly elongated sorting area located in ventral surface of esoph- ageal insertion, flanked by pair of narrow, low folds that unite in posterior region bulg- ing against right edge of central pad; an- other sorting area in dorsal-right gastric region adjacent to dorsal hood; gastric shield occupying posterior and dorsal regions of stomach. Dorsal hood located obliquely in dorsal region of stomach, low, amply opened to stomach. Intestine and style sac running ventrally and, in region dorsal to foot base, curving anterodorsally in a narrow loop; in- testine free from style sac in ascendant branch of this loop; in middle level of its pre- ceding loop, intestine abruptly performing a 360° curve, running adjacent to and at right of preceding loop; this ascendant branch of intestine becoming slightly separated from style sac, passing at some distance poste- rior to stomach, penetrating pericardium. Intestine relatively narrow, with uniform width along its length to pericardial region; running horizontally though ventricle and narrowly between both posterior pedal re- tractor muscles. Anal papilla (Figs. 67, 68: an) short, located on posterior region of ven- tral surface of posterior adductor muscle, with a narrow and short distal projection. Central Nervous System (Figs. 67, 68): As in B. cancellaria, except in having broad connectives between cerebropleural and vis- ceral ganglia, running through gonad and digestive diverticula. Measurements of Dissected Specimens (length, lateral, height in mm): AMNH 298090: 1) 23.2 by 10.6 by 15.0; 2) 20.0 by 9.8 by 13.7. FLORIDIAN BARBATIA MORPHOLOGY 369 Distribution Florida, USA, to Сеага, Brazil. Habitat Rocky, from intertidal zone to about 10 m depth. Material Examined U.S.A.; Florida; Florida Keys; off Key Vaca, 24°39.30’М, 81°01.30’W, AMNH 298090, 8 specimens (FK-131, R/V “Floridays”; Bieler & Cipriani coll.; 07/viii/1997); Monroe County, “Long Key Artificial Reefs”, oceanside of Long Key, 24°44.78’М, 80°50.00'W, 7 т depth, MZSP 36278, 3 specimens (FK-621, Simone coll., 17/vii/2002); Tennessee Reef Light, 24°44.75’М, 80°46.95’W, 4-7 т depth, MZSP 36146, 8 specimens (FK-648, Simone coll., 26/ vii/2002); W side of Pigeon Key, 24°42.2’N, 81°09.3’W, MZSP 36164, 1 specimen (FK- 659, Simone coll., 28/vii/2002); between Marquesas Keys & Dry Tortugas, 24°50’36"N, 82°28'25"W to 24°48'17"N, 82°28'42"W, 30 т depth, FMNH 279044, 2 shells (FK-081, Bieler et al. coll., 22/iv/1997); East Washerwoman Shoal, 24°40’N, 81°04.3’W, to 2.7 m depth, FMNH 295616, 3 specimens (FK-115, Mikkelsen & Bieler coll. R/V “Floridays”, 12/ vii.1997); off Marathon, ММ 50, 24°39.53’N, 81°00.90’W, 7 т depth, ЕММН 295615, 2 specimens (FK-121, Mikkelsen & Bieler coll., 21/vii/1997); Looe Key coral reef, 24°32.77’М, 81°24.23’W, FMNH 295617, 1 specimen (FK- 262, Bieler & Mikkelsen coll., 11/viii/1999); Missouri Key, FMNH 183739, 4 shells, FMNH 183740, 9 shells (Koto coll., 1939), FMNH 159758, 1 shell (Bales coll.); Molasses Key, FMNH 183741, 2 specimens (Koto coll., 1955). DISCUSSION OF CHARACTERS The shell obviously has been the main sub- ject of most morphological studies on the arcids. However, the shell of these sessile animals is greatly subject to ecophenotypic variation, enlarging with growth to fit into the concavities in the rocks where it lives. For ex- ample, the outline can be flattened or wide; the anteroposterior or dorsoventral distances can be short or long. As a result, the outline Can vary in a single species from almost sym- metrical (Fig. 26) to asymmetrical (Fig. 29), or from almost straight (Fig. 18) to arched (Fig. 15). For this reason, the shell can be mislead- ing, particularly in specimens of Barbatia cancellaria and B. candida. The color pattern, however, can be useful in such cases; the in- terior of the shell of B. cancellaria, for example, is purple (Figs. 1-7), whereas those of the other three species are pale beige to white (Figs. 20-26). (1) Periostracum: 0 = glabrous (without scales); 1 = with scales but not extending beyond shell edges (A. notabilis); 2 = with scales extending beyond shell edges (B. cancellaria, B. candida, B. tenera, A. zebra) (additive). The periostracum of B. cancellaria, B. can- dida and B. tenera bears well developed scales, which extend beyond the shell edge, mainly in the region between the radial ribs (Figs. 4, 5, 27, 29-32). This feature is shared with Arca zebra. Anadara species, on the other hand, normally have small, velvet-like hairs in the periostracum, as present in A. notabilis and most other species of the ge- nus, except A. brasiliana (Lamarck, 1819). The periostracum of B. dominguensis, how- ever, is glabrous, smooth, transparent, and does not extend beyond the shell edge. This state has been found in most bivalves and was initially considered plesiomorphic. How- ever, according to the analysis presented herein, the state in B. dominguensis is a re- version. The periostracum can be destroyed by the cover of epibionts that attach to the shell, as well as by wave abrasion. An interpretation of the periostracum must therefore be pre- ceded by special care with the selection of specimens. In В. candida, an interesting variation is found in the different populations observed. In northern specimens, the peri- ostracal scales are mostly bifid, resembling a snake’s tongue (Fig. 12), whereas in south- ern specimens the scales are simple and somewhat pointed (Fig. 32). For identifica- tion purposes, the types of periostracal scales are very informative in the separa- tion of B. candida and B. tenera, because in both species the conchological attributes are very similar. (2) Ligamental area: 0 = wide; 1 = narrow (Barbatia). The ligamental area of the arcids possesses a generally weak ligament, being constricted in its growth pattern to an increase in um- bonal area either by horizontal growth, which pushes the beaks apart, or by ventral growth, 370 SIMONE & CHICHVARKHIN / A NA SS т \\\\\ IN AM NN UN ухо ут Pr vg FIGS. 43-47. Barbatia cancellaria anatomy. FIG. 43: Whole left view, left valve and mantle lobe removed; FIG. 44: Main muscular system and digestive tubes as in situ, left view; FIG. 45: Left palp, left view, outer hemipalp deflected to show inner surface; FIG. 46: Posterior region of visceral mass, ventral view, both gills deflected; FIG. 47: Detail of anus, ventral view. Scale bars = 5 mm (Figs. 43, 44), 1 mm (Figs. 45-47). FLORIDIAN BARBATIA MORPHOLOGY FIGS. 48-51. Barbatia cancellaria anatomy. FIG. 48: Left gill, mid- transverse section; FIG. 49: Mantle border, transverse section between middle and posterior regions; FIG. 50: Detail of pericar- dial region, left view, dorsal pericardium and left gill removed; FIG. 51: Stomach, dorsal view, dorsal wall sectioned longitudi- nally and deflected, inner surface exposed. Scale bars = 1 mm. 371 372 which invades the hinge plate (Thomas, 1976). The structure of the ligament also varies in producing a few rather wide chev- rons (in some Anadara and Barbatia) or many narrow chevrons (in Arca). АП ligamental characters in the present sample resulted in autapomorphies, except ligamental width, which separated the ingroup from the remaining arcids. (3) Hinge teeth: 0 = mostly perpendicular; 1 = mostly tilted (Barbatia). The narrow ligamental area, when compared to other arcids, has long been recognized as a distinctive character of Barbatia. The narrow ligamental area is associated with the inclination of the hinge dentition (Reinhart, 1935: 20, pls. 1, 2). This character is found in the examined species, however, the den- tition was very variable. Younger specimens generally possess a more typical hinge (Figs. 7, 17) that is disfigured during ontogeny and development. Sometimes the more central teeth are reduced or lost (Figs. 6, 14), whereas in other cases the more anterior and posterior teeth become horizontal (Fig. 14). Barbatia dominguensis has as the distinc- tive feature of a sudden change of the hinge axis at the cardinal level (Figs. 17, 20), which is one of the features used by some authors (e.g., Reinhart, 1935) to substantiate the subgeneric separation of Acar. Asimilar situ- ation also occurs in B. tenera, which is mostly referred to the subgenus Fugleria. Another special feature of the arcids is the nature of the byssus. In the examined arcids the byssus has fused fibers, forming a single, thick bundle. Vestiges of the fibrous nature of this structure are only found in the dorsal area, where the byssus attaches to the foot. In this region, the byssal bundle has a spe- cial arrangement as a concavity ornamented by a series of parallel folds. This concavity fits into a special protuberance in the poste- rior region of the byssal groove, which has furrows for the folds. This byssal morphol- ogy is apparently exclusive to the arcids and would be a strong synapomorphy if some genera (e.g., Anadara) did not lack the bys- sus. Arca notabilis, for example, has a byssal furrow comparable to that found in other byssate bivalves and can produce a typical byssus with separated fibers. The presence or absence of a thick byssus was also used in arcid classification (Lamy, 1907; Reinhart, 1935; Coan et al., 2000). Barbatia domi- nguensis differs in byssal morphology com- SIMONE & CHICHVARKHIN pared to other Barbatia species (Fig. 39) and to Arca, as it lacks an anterior notch in the dorsal concavity (Fig. 22). This lack of an anterior notch is an autapomorphy in the present study; this character could not be fully examined in B. tenera, because no fully preserved byssi of this species were avail- able. Although normally the color patterns of the soft parts are very variable in mollusks, the species studied here maintained a constant differentiation in the color of most regions of soft parts. Barbatia cancellaria has a greater richness of pigmentation, with dark brown, beige and white spots, which also occur in B. candida, but in paler patterns; B. tenera is in general almost invariably orange, while B. dominguensis is mostly pale to dark purple. (4) Origin of pair of pedal protractor muscles: 0 = in restricted area in posteroventral re- gion of anterior adductor muscle; 1 = sur- rounding ventral edge of anterior adductor muscle (B. dominguensis, B. tenera). The two indicated species differ from the other included species in having the pair of pedal protractor muscles apparently as part of the integumental anterior area, and their origin is along the ventral edge of the origin of the anterior adductor muscle, in a dors- oventrally narrow, but anteroposteriorly broad, region. In this state, the ventral edge of the shell scar normally attributed only to the anterior adductor muscle, actually also belongs to the pedal protractors. The main musculature of the examined ingroup species has the typical arcid mor- phology. Barbatia dominguensis, however, is distinct in having enlarged pedal protrac- tor muscles, which are detached from the integument crossing through visceral glands. (5) Mantle edge inner fold: 0 = simple; 1 = tall (Barbatia). The mantle edge of the four Barbatia spe- cies studied herein is of the basic comple- ment, with three folds. This morphology has been also found in Anadara trapezia (Deshayes, 1840) (Sullivan, 1961: fig. 4). However, the species studied here present a greater development of the inner fold, form- ing an undulated membrane in some speci- mens (Figs. 49, 55, 65, 70: of). Another attribute of the examined species is the pres- ence of well-developed eyes along the mantle edge т В. dominguensis (Figs. 23-25), which FLORIDIAN BARBATIA MORPHOLOGY 373 NS, BEER N TN SC Ñ \ \ P i Ex FIGS. 52-59. Barbatia candida anatomy. FIG. 52: Whole left view, left valve and mantle lobe removed; FIG. 53: Digestive tubes, left view, seen as in situ, with some adjacent structures; FIG. 54: Main muscular system, left view; FIG. 55: Mantle border, transverse section between middle and posterior regions; FIG. 56: Left palp, left view, outer hemipalp deflected to show inner surface; FIG. 57: Left gill, mid-transverse section; FIG. 58: Stomach and pericardium, dorsal view, dorsal wall of stomach sectioned longitudinally and deflected, inner surface exposed, dorsal pericardial wall removed; FIG. 59: Detail of anus, ventral and slightly left view. Scale bars = 5 mm (Figs. 52-54), 1 mm (Figs. 55-59). 374 are present оп the ощег fold (Fig. 65), cov- ered by the periostracum. This feature is possibly related to the transparency and sim- plicity of its periostracum. Although the pres- ence of ocelli is a common feature of the arcid mantle edge (Boss, 1982), a well-developed eye with a lens (Jan-Olof, 1998) is presently an autapomorphy of B. dominguensis. The gill of the examined species is of typical arcid morphology (Atkins, 1936; Sullivan, 1961) (Figs. 42, 48), although В. candida 1$ distinct in having a membrane on the ventral half of each filament, holding it in a determi- nate position (Fig. 57). The gill projection or stalk, inserted into the ventral region of the posterior adductor muscle, and projecting the posterior region of each gill posteriorly (Figs. 43, 46, 52, 60: gp), has been clearly illus- trated by some of authors (e.g., Rost, 1955; Sullivan, 1961; Boyd, 1998; Coan et al., 2000). However, no one has either named or called attention to this structure. The kidneys are separated from each other and are closely similar in all examined arcids. A distinct nephrostome 1$ located at the an- terior end of the branchial surface of the kid- ney, apparently surrounded by muscles, forming a low papilla. The heart of the examined arcids 1$ distinct in having the accessory auricles as clear expansions of the ventricle. The remaining region of the ventricle is small and surrounds the intestine. The aortae emanate from this portion of the ventricle. The accessory au- ricles are very expansive, almost as large as the auricle itself. Heath (1941) and Sullivan (1961: fig. 1C) clearly show the ac- cessory auricles, however, they did not pay any special attention to them, simply calling them ventricles. Boss (1982) described the arcid heart as having two lateral cavities, each one with a ventricle and an auricle. (6) Palps' ventral region preceding mouth: O = simple; 1 = wide, forming a sac (B. cancellaria, B. candida). The labial palps of arcids are unlike those of other bivalves in being dorsoventrally elon- gated, with their anterior edges attached to the visceral mass. This palp morphology is apparently present in all arcids we have seen in the literature (e.g., Lamy, 1907; Reinhart, 1935; Sullivan, 1961), and could be charac- teristic of the family. The inner folds, which typically are distributed along the entire in- ner surfaces of the palps, are mostly re- stricted to the dorsal halves in the arcids SIMONE 8 CHICHVARKHIN (Sullivan, 1961: fig. 3C; this study). This ap- parently is another distinctive feature of the family. Barbatia cancellaria; however, is unique in having some longitudinal folds on the ventral halves of the palps, concentrated on the inner borders between the two hemipalps. Barbatia cancellaria and B. can- dida are distinct as a pair in having the re- gion preceding the mouth very broad and flaccid, forming a cover on the anterior re- gion of visceral sac, with a sac in the ventral region of the palps (Figs. 35, 43, 45, 52, 56). (7) Stomach with anterior typhlosole forming a fold posterior to central pad: 0 = absent; 1 = present (Barbatia). See discussion under character 8. (8) Anterior typhlosole surrounding right and posterior edge of central pad: 0 = absent; 1 = present (B. cancellaria, B. candida) (Figs. 51, 58: ty). The stomachs of the examined species are somewhat similar to one another, referred to as type Ш (Purchon, 1957; Boss, 1982). Although they differ in a number of details, only two of these are coded as characters in the present phylogenetic analysis. Several other characters are considered autapo- morphies in the present study. One such character is the position of the dorsal hood, which is in the middle region of the gastric dorsal wall т В. cancellaria and В. candida, but more posteriorly in B. dominguensis; besides, it is widely opened in В. tenera. The dorsal hood is located more anteriorly in Anadara trapezia (Sullivan, 1961). In addi- tion, B. candida has a duct to the digestive gland located in the dorsal-right region, which is exceptionally large and can be con- fused with the dorsal hood, which is some- what reduced in this species. The arcids generally possess few intestinal loops, and the intestine is narrow, running close to the style sac. In В. cancellaria, on the other hand, the intestine 1$ slightly broader, forming a close zigzag in the style sac end, forming almost a secondary intes- tinal chamber. This is an autapomorphy in present study. The anus of the examined species 15 notable in being siphoned, that is its aperture is lo- cated at the tip of a muscular stalk (Figs. 47, 64), which has the capacity for movement and protraction. However, В. candida 1$ unique in having the anal aperture posterior to this stalk base (Fig. 59). FLORIDIAN BARBATIA MORPHOLOGY 375 (9) Connective between cerebropleural ganglia The pair of connectives uniting the cerebro- and visceral ganglia: 0 = broad, running pleural ganglia with the visceral ganglia is through visceral glands; 1 = thin, running close normally easy to distinguish in dissection, as to integument (B. cancellaria, B. candida). conspicuously iridescent cords running al- À ET À / WW ENG x E xX à £ iy 3 4 2 yf N) € A р - d FIGS. 60-65. Barbatia dominguensis anatomy. FIG. 60: Whole left view, left valve and mantle lobe removed; FIG. 61: Left palp, left view, outer hemipalp deflected to show inner surface: FIG. 62: Digestive tubes and main muscular system, left view, seen as in situ; FIG. 63: Stomach, dorsal view, dorsal wall sectioned longitudinally and deflected, inner surface exposed; FIG. 64: Detail of anus, ventral view, also showing adjacent region; FIG. 65: Mantle border, transverse section between middle and posterior regions. Scale bars = 2 mm (Figs. 60, 62), 0.5 mm (Figs. 61, 64-65). 376 SIMONE & CHICHVARKHIN FIGS. 66-73. Barbatia tenera anatomy. FIG. 66: Whole left view, left valve and mantle lobe removed; FIG. 67: Posterior region of visceral mass, ventral view, both gills deflected; FIG. 68: Digestive tubes, central nervous system and main muscular system, left view, seen as in situ, palp shown as a transparent structure; right gill stalk also shown; FIG. 69: Left palp, left view, outer hemipalp deflected to show inner surface; FIG. 70: Mantle border, transverse section between middle and posterior regions; FIG. 71: Stomach, dorsal view, dorsal wall sectioned longitudinally and deflected, inner surface exposed; FIG. 72: Mid-transverse section of gill; FIG. 73: Visceral mass, dorsal view, with dorsal portion of integument and mantle, visceral glands covering stomach, and part of pericardium removed. Scale bars = 1 mm. FLORIDIAN BARBATIA MORPHOLOGY 377 TABLE 1. Character matrix for four Barbatia species studied here and three outgroups (see text for description of characters). Character/taxon 1 2 3 4 5 6 7 8 9 Barbatia cancellaria 2 1 1 0 1 1 1 1 1 Barbatia candida 2 1 1 0 1 1 1 1 1 Barbatia dominguensis 0 1 1 1 1 0 1 0 0 Barbatia tenera 1 1 1 1 1 0 1 0 0 Arca zebra 2 0 0 0 0 0 0 0 0 Anadara notabilis 1 0 0 0 0 0 0 0 0 Isognomon bicolor 0 0 0 0 0 0 0 0 0 most straight through the visceral glands (go- ANALYSIS OF THE CLADOGRAM nad and digestive diverticula) (Fig. 62: tm). However, п В. cancellaria and В. candida this A cladistic analysis based on the matrix in pair of connectives 1$ less distinct, being very Table 1 resulted in a single most parsimoni- thin and running close to the inner surface of ous tree (Fig. 74). We emphasize, however, the integument. This state is interpreted as that a search for family characters was not apomorphic. On the other hand, the enlarge- performed, which would demand a wider ment of these connectives in В. dominguensis analysis. The main concern is to compare the is autapomorphic and could be coded as a four species of Barbatia studied herein and to third, state, a presently autapomorphy. The test how the analysis of the anatomy is useful remaining features of the central nervous sys- in phylogenetic estimation. Although the ma- tem and primary sense organs of the exam- trix is admittedly very restricted, based on the ined species are similar to those described in result, B. cancellaria and B. candida are closer the literature (e.g., Sullivan, 1961). It is inter- to each other than to the other two Barbatia esting to note the location of the visceral gan- species, as are B. dominguensis and B. glia, at the base of the posterior projections tenera; the first is a branch supported by three of the gills (Fig. 46), normally they are located synapomorphies (node 4), whereas the sec- slightly anterior to them. ond is supported by a single synapomorphy Barbatia cancellaria Barbatia candida Barbatia tenera Barbatia dominguensis Arca zebra Anadara notabilis FIG. 74. Cladogram showing the relationship of the examined species and the two arcid outgroups (below). Squares represent synapomorphies (black square = not homoplastic; empty square = reversion), superior number refers to the character, inferior number indicates the state. Larger numbers in italics are node numbers referred to in the text (length = 12; consistency index = 0.81; retention index = 0.85). 378 SIMONE & CHICHVARKHIN (node 5). The genus Barbatia $.1. is also well supported by four synapomorphies (node 3), which separates them from the other arcids, being monophyletic. Additionally, it was pos- sible to see that Arca is more closely related to Barbatia than Anadara. The single ho- moplasy detected in the analysis was the re- version in the periostracum of B. dominguensis from scaly with extensions to smooth (lacking scales) (character 1). Although all autapo- morphies were excluded of the present analy- sis, they would be useful in a wider analysis, including more species and more genera. An example is the composed eyes in the mantle border of В. dominguensis; they could be char- acter of the species, of the subgenus Acar, or something else. As stated in the synonymy listing, Barbatia cancellaria and B. candida have been consid- ered as members of the subgenus Barbatia s.s. by several authors. However, В. candida is sometimes considered in the subgenus Cucullaearca Conrad, 1865 (e.g., Rios, 1994). Barbatia dominguensis, on the other hand, has been placed consistently in the subgenus Acar, whereas B. tenera has been referred to Fugleria. An analysis of the subdivision of the genus, and/or the elevation of the subgenera to generic level is considered premature at this present level of knowledge. A recent paper (Marko, 2002) presented a molecular phylogenetic analysis of a pool of arcids that coincidently includes all species studied here, in addition to a number of Pa- cific species. The results of that analysis (Marko, 2002: figs. 1-5) are difficult to com- pare with those obtained here, since they dif- fer almost totally. The combined cladogram of CO1 and H3 sequences, despite excluding some of the presently studied species, re- vealed a closer relationship of Barbatia can- dida with Anadara spp., and the same for Arca imbricata with B. dominguensis. CONCLUSIONS (1) In this analysis of four species attributed to Barbatia s.l., Barbatia cancellaria and В. candida are found to be sister taxa, as are B. dominguensis and B. tenera. (2) Barbatia s.l. is a monophyletic group. (3) Morphological characters, including those from conchology and internal anatomy, are useful in phylogenetic analysis, even of closely related bivalve species. ACKNOWLEDGMENTS This study is one of the results of the Inter- national Marine Bivalve Workshop, held in the Florida Keys, 19-30 July 2002, funded by U.S. National Science Foundation award DEB- 9978119 to co-organizers R. Bieler and P. M. Mikkelsen, as part of the Partnerships in En- hancing Expertise in Taxonomy [PEET] Pro- gram. Additional support was provided by the Bertha Lebus Charitable Trust, the Comer Science 8 Education Foundation, the Field Museum of Natural History, and the American Museum of Natural History. This study was also partly supported by Brazilian Fundacáo de Amparo a Pesquisa do Estado de Sáo Paulo, process # 00/11074-5 and 00/11357-7, to the senior author. We are grateful to anony- mous referees and the associate editor for valuable comments and suggestions on the paper. LITERATURE CITED ABBOTT, R. T., 1974, American seashells: the marine Mollusca of the Atlantic and Pacific coasts of North America, 2nd ed. Van Nostrand Reinhold Company, New York. 663 pp., 24 pls. ANDREWS, J., 1971, Sea shells of Texas coast. University of Texas Press, Austin. 298 pp. ATKINS, D., 1936, Some new observations on the ciliary feeding mechanism of Glycymeris glycymeris (L.) and Arca tetragona Poli. Quar- terly Journal of Microscopical Science, 79: 181- 308. BOSS, К. J., 1982, Mollusca. Pp. 947-1166, in: $. Р. PARKER, ed., Synopsis and classification of living animals, vol. 2. McGraw-Hill Book Company, New York. BOYD, S. E., 1998, Order Arcoida. Pp. 253-261, in: P. L. BEESLEY, С. J. В. ROSS & А. WELLS, eds., Mollusca: the southern synthesis. Fauna of Australia, Vol. 5A. CSIRO Publishing, Melbourne. СОАМ, Е. V., P. VALENTICH SCOTT & Е. К. BER- NARD, 2000, Bivalve seashells of western North America: marine bivalve mollusks from Arctic Alaska to Baja California. Santa Barbara Mu- seum of Natural History, Santa Barbara, Cali- fornia. 764 pp. COELHO, А. С. S. & D. R. В. CAMPOS, 1975, Contribuiçäo ao conhecimento dos molsucos do Rio de Janeiro, Brasil 1- Bivalvia, Pterimorphia, Arcoida, Arcoidea. Arquivos do Museu Nacional, 55: 35-57. FARRIS, J. S., 1988, Hennig86, version 1.5. Dis- tributed by the author (computer program). Port Jeffersen Station, New York. HEATH, H., 1941, Anatomy of pelecypod family Arcidae. Transactions of the American Philo- sophical Society, 31(5): 281-319, 22 pls. FLORIDIAN BARBATIA MORPHOLOGY 379 HUMFREY, M., 1975, Sea shells of the West Indies. Taplinger Publishing Company, New York. 351 pp. JAN-OLOF, S., 1998, Comparative optics of prosobranch eyes. Doctoral dissertation, Lund University, Sweden. 86 pp. LAMY, E., 1907, Révision des Arca vivants du Muséum d'Histoire Naturelle de Paris. Jour- nal de Conchyliologie, 55: 1-111. MARTINS, C. M., 2000, Estudo da anatomia descritiva e funcional de Isognomon alatus e de sua distribuicáo pelo Litoral Brasileiro. Doc- toral Thesis, Instituto de Biociéncias da Universidade de Sáo Paulo, Sáo Paulo. 185 pp. MARKO, P. B., 2002, Fossil calibration of mo- lecular clocks and the divergence times of geminate species pairs separated by the Isth- mus of Panama. Molecular Biology and Evo- lution, 19(11): 2005-2021. DIAZ MERLANO, J. M. D. & М. PUYANA HEGEDUS, 1994, Moluscos del Caribe Colombiano. Colciencias, Fundacion Natura Colombia, Bogota. 291 pp., 74 pls. MIKKELSEN, P. M. 8 R. BIELER, 2004, Interna- tional Marine Bivalve Workshop 2002: Intro- duction and summary. In: В. BIELER 8 Р. M. MIKKELSEN, eds., Bivalve Studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 241-248. PURCHON, R. D., 1957, The stomach in the Filibranchia and Pseudolamellibranchia. Pro- ceedings of the Zoological Society of London, 129(1): 27-60. RAMOS, T., 1998, Tree Gardner, version 2.2. Distributed by author (Computer program). Sáo Paulo, Brazil. REDFERN, C., 2001, Bahamian seashells: a thousand species from Abaco, Bahamas. Bahamianseashells.com Inc., Boca Raton, Florida. x + 280 pp., 124 pls. REINHART, P. W., 1935, Classification of the pelecypod family Archidae. Bulletin du Musée Royal d'Histoire Naturelle de Belgique, 11(13): RIOS, Е. C., 1970, Coastal Brazilian seashells. Fundacáo Cidade do Rio Grande, Rio Grande, Brazil. 255 pp., 4 maps, 60 pls. RIOS, E. C., 1975, Brazilian marine mollusks iconography. Fundacgáo Cidade do Rio Grande, Rio Grande, Brazil. 331 pp., 91 pls. RIOS, E. C., 1985, Seashells of Brazil. Fundacáo Cidade do Rio Grande, Rio Grande, Brazil. 328 pp., 102 pls. RIOS, E. C., 1994, Seashells of Brazil, 2nd ed. Fundacáo Universidade do Rio Grande, Rio Grande, Brazil. 368 pp., 113 pls. ROST, H., 1955, A report on the family Arcidae (Pelecypoda). Allan Hancock Pacific Expedi- tions, 20(2): 177-249. SULLIVAN, G. E., 1961, Functional morphology, micro-anatomy, and histology of the “Sydney cockle”, Anadara trapezia (Deshayes) (Lamellibranchiata: Arcidae). Australian Jour- nal of Zoology, 9(2): 219-257. THOMAS, R. O. K., 1976, Constraints of liga- ment growth, form and function on evolution in the Arcoidea (Mollusca: Bivalvia). Paleobiology, 2(1): 64-83. WARMKE, С. L. 8 К. T. ABBOTT, 1962, Carib- bean seashells. Dover Publication, Inc. New York. 348 pp. Revised ms. accepted 21 January 2004 MALACOLOGIA, 2004, 46(2): 381-415 RECENT CHAMIDAE (BIVALVIA) FROM THE WESTERN ATLANTIC OCEAN Matthew К. Campbell’, Gerhard Steiner’, Lyle D. Campbell? & Hermann Dreyer? ABSTRACT The International Marine Bivalve Workshop, Florida Keys, July 2002, initiated an exami- nation of the bivalve genera Chama and Pseudochama. Unraveling the systematics of the seven species collected during the workshop necessitated a review of literature and col- lections for the fossil and Recent chamid taxa of the western Atlantic. Subsequent discov- ery of a small population P. inezae Bayer, 1943, enabled detailed observation of variation within a previously rare species. Given the 50:50 ratio of right-attached to left-attached specimens and the characteristic Chama prodissoconch, P. inezae is transferred to the genus Chama. Pseudochama radians (Lamarck, 1819) is returned to the genus Chama based on a complex of morphological characters and molecular data. We discuss the morphology of eight Chama and one Arcinella species. Nucleotide sequence data of the ITS region (ITS1 + 5.8S rRNA + ITS2) and the 16$ rRNA gene were obtained for six species of Chama collected during the workshop. Chama congregata Conrad, 1833, appears to include multiple species based on morphological details and molecular data. One specimen of C. congregata had two alleles of ITS ele- ments, which is the first demonstrated case of multiple alleles within the Bivalvia. Molecu- lar phylogenetic reconstructions containing С. congregata will be more complicated because gene trees and species trees may not be identical due to incomplete lineage sorting. Chama sarda Reeve, 1847, appears to include multiple species based on morphological data. Populations of several other fossil and recent species need to be examined for the range of variation within nominate species. Key words: Chamidae, Chama, Pseudochama, Arcinella, Neogene, Atlantic, attachment, transposition. INTRODUCTION This revision of the Recent Chamidae from the western Atlantic was undertaken to deter- mine the systematics of seven chamid spe- cies collected during the International Marine Bivalve Workshop (IMBW), Florida Keys, July 2002. Recent western Atlantic chamid species are found in tropical and subtropical seas at shallow to moderate depths, from Brazil to Cape Hatteras, North Carolina. Neogene dis- tribution ranged from the Miocene of Argen- tina to the Miocene of New Jersey. The earliest stratigraphic records of Chamidae are from the upper Cretaceous (Keen, 1969; Kennedy et al., 1970). In the Recent, eight species of Chama and Pseudochama are recognized from cur- rent literature for the western Atlantic, all of which occur in the study area. Four are also found north to Cape Hatteras, seven range south to the Lesser Antilles, and five reach Brazil. Three species of Arcinella have been reported from the western Atlantic basin, but two occur only south of the Florida Keys. Con- sequently, a thorough literature review of the Floridian species requires a larger, western Atlantic focus. Lamy (1928) is an important but often overlooked reference containing much critical detail concerning Recent Chamidae from all oceans. ‘Department of Geological Sciences, Indiana University, 1001 East Tenth St., Bloomington, Indiana 47405, U.S.A.; Research Associate, Carnegie Museum of Natural History, 4400 Forbes Ave, Pittsburgh, Pennsylvania 15213, U.S.A.; ecphora@indiana.edu Institute of Zoology, University of Vienna, Althanstr. 14, A-1090 Vienna, Austria “Division of Natural Sciences, University of South Carolina Spartanburg, 800 University Way, Spartanburg, South Carolina 29303, U.S.A. 382 CAMPBELL ET AL. MATERIALS AND METHODS Material Examined Living specimens of Chama congregata, C. macerophylla, С. florida, С. sinuosa, С. radi- ans, С. sarda, and С. inezae were collected during the International Marine Bivalve Work- shop (Table 1). The field collections are listed in the introduction to this volume (Mikkelsen 8 Bieler, 2004). The identifications of these seven species are based on the morphologi- cal descriptions in Abbott (1974), Bayer (1943), Dall (1886), and Redfern (2001), as well as comparison with specimens from the Florida Museum of Natural History (FLMNH) and the University of South Carolina Spartanburg (USCS). Mikkelsen 8 Bieler (2000) also re- ported C. lactuca and Arcinella cornuta from the Florida Keys, and these are included in this review. Molecular Methods We determined sequences of the nuclear ITS1 + 5.88 rRNA + ITS2 region and of the mitochondrial 16$ rRNA genes of four speci- mens of Chama macerophylla, two specimens of С. congregata, and of single specimens of С. sarda, С. florida, С. inezae, and С. radians (Table 2). Two specimens of an unidentified Chama species from the Seychelles served as the outgroup. The other available bivalve sequences of the ITS genes are too divergent to be reasonably aligned. Living specimens were removed from their shells and dehydrated in 96% ethanol. The tis- sues were washed in distilled water priorto DNA extraction. Total DNA was isolated with CTAB (Winnepenninckx et al., 1993) or with CHELEX resin (Sigma) (Steiner & Hammer, 2000). The target sequence, including ITS1, 5.85 rRNA, and ITS2, was amplified using the prim- ers ITS-F2 (5'-taa caa ggt atc cat agg tga a- 3°) and ITS-R2 (5’-tgc Ка aat {са gcg ggt-3°) or in two fragments using internal primers 5.8- В (5'-cag ctg gct gcg ctc ttc tac gac-3°) and 5.8-F (5'-gtc gta gaa gag cgc agc cag ctg-3’) when necessary. The 16S rRNA sequence was amplified with the primers 16Sf (5°-ctc gcc tgt Ца wca aaa aca t-3’) and 16Sr (5 -acg ccg gtc tka act cag-3’). The PCR-reactions were run on a Personal Cycler (Biometra) in 30 ul reac- tion mixes containing 1.75-3.0 mM MgCl,, each dNTP at 250 uM, each primer at 0.1 uM, 0.6 units Taq polymerase (Biotaq Red, Bioline) and the supplied reaction buffer at 1 x concen- tration. The standard PCR cycle conditions were: initial denaturation step of 2 min at 94°C, 35 cycles of 40 sec denaturation at 94°C, 40 sec annealing at 46°C, and 75 sec primer ex- tension at 72°C, followed by a final primer ex- tension step of 10 min at 72°C. Touchdown PCR was often more successful than the stan- dard protocol. It included stepwise lowering annealing temperatures from 56°C to 42°C within 10 cycles and a subsequent return to 56°C within the remaining 25 cycles. PCR prod- ucts were purified with the Concert Rapid PCR Purification System (Life Technologies) and sequenced with the PCR primers on an ABI 3700 at VBC-Genomics Bioscience Research GmbH, Vienna. Heterogeneous PCR products were subcloned with the TOPO cloning kit (Invitrogen), and three clones sequenced. Sequences were aligned manually and with CLUSTAL X 1.8 (Thompson et al., 1997) us- ing default parameters for the ITS sequences. The only change for aligning the 16S rRNA sequences was setting the gap opening pen- alty to 20. Phylogenetic analyses were run with PAUP* 4.0b10 (Swofford, 1998) on an IBM-PC TABLE 1. Stations where Chamidae were collected live during the IMBW. For full station data, see introduction to this volume (Mikkelsen & Bieler, 2004). Species 624 625 629 Chama congregata 1 1 11 Chama macerophylla - 1 13 Chama florida & a E Chama sarda - Chama sinuosa 1 Chama inezae 2 - - Chama radians 6 Station 630 631 641 644 650 651 1 О QOı = 1 Nt N un 1 a. 1 = of 1 WESTERN ATLANTIC RECENT CHAMIDAE 383 and on the Schródinger 1 Linux-Cluster at the Central Informatics Service, University of Vienna. Unweighted maximum parsimony analyses (MP) included an exhaustive search and 1,000 bootstrap replicates, each with three random sequence additions. Gaps were treated as missing. For maximum-likelihood analyses (ML), the most parsimonious trees (MPT) were used as starting trees for the calculation of the model parameters and subsequent branch swapping. Empirical nucleotide frequencies and the parameters for the transition/transversion ratio and the gamma shape value were esti- mated under the HKY85 model with rate het- erogeneity and four categories of substitution rates following a gamma distribution (HKY85+G model) as recommended by the likelihood ra- tio test implemented in MODELTEST (Posada & Crandall, 1998). The resulting values were then set for subtree-pruning-regrafting (SPR) branch swapping. The two phylogenetic mark- ers were analysed both separately and com- bined in a single matrix. The latter analysis 1$ justified, although the taxon samples for the individual analyses are not identical, because the resulting trees were congruent. RESULTS Molecular Genetics and Phylogeny We obtained both the near-complete ITS1 + 5.85 rRNA + ITS2 and partial 16$ rRNA se- quences from seven specimens collected in the Florida Keys belonging to Chama macero- phylla (3), C. congregata, C. sarda, C. florida, and C. radians, and two specimens of C. sp. from the Seychelles. An additional specimen of С. congregata yielded two different alleles ofthe ITS region but no 16$ sequence. Chama macerophylla and С. inezae are also герге- sented by one additional specimen each with only the 16$ sequence (Table 1). We faced considerable difficulties amplify- ing and sequencing the target sequences. Al- though we applied two different extraction protocols, PCR reactions were inhibited by some components of the DNA extract of cer- tain specimens. Other specimens of the same species subjected to identical protocols worked well. In several reactions the PCR yielded at least two different products. In ad- dition to the Chama sequence, these were identified by BLAST searches as products from parasites, Perkinsus sp. (Apicomplexa, Pro- tista) and a parasitic flatworm. In the case of С. congregata 625-1a and ЛЬ, the two prod- ucts were polymorphic copies of the ITS ele- ments. The amplified sequence lengths for the ITS region ranged from 910 bp in Chama con- gregata to 1,072 bp in C. florida, resulting in an alignment with 1,058 positions (the autapomorphic 5’-епа of the С. florida se- quence was truncated) of which 234 were par- simony informative. The 16$ rRNA sequences varied in length from 659 bp in C. sarda to 825 bp in C. macerophylla. The sequence of C. florida was incomplete and consisted of 364 TABLE 2. List of species and specimens sequenced for ITS1 + 5.8 rRNA + ITS2, and 16$ rRNA with AMNH or FMNH catalog numbers, sampling locations and Genbank Accession numbers. Genbank Accession Number Species Catalog Number Station-Specimen ITS 16$ Chama macerophylla AMNH 306419 625-3 AY230078 AY388507 Chama macerophylla FMNH 301423 629-22 AY230079 AY388511 Chama macerophylla AMNH 306415 625-4 AY230080 AY388506 Chama macerophylla FMNH 301291 Palm Beach, FL - AY388509 Chama congregata AMNH 306412 624-1a AY230081 - Chama congregata AMNH 306412 624-1b AY230082 - Chama congregata FMNH 301422 629-9 AY230083 AY388512 Chama florida AMNH 306410 641-1 AY230084 AY388508 Chama sarda AMNH 306411 629-26 AY230085 AY388505 Chama radians AMNH 306414 624-6 AY230086 AY388510 Chama тегае FMNH 301292 Palm Beach, FL - AY388513 Chama sp. - Seychelles AY230087 AY388514 Chama sp. - Seychelles AY230088 AY388515 oo, 384 CAMPBELL ET AL. Chama sp. (Seychelles) Chama sp. (Seychelles) Chama sp. (Seychelles) Chama sp. (Seychelles) Chama florida Chama macerophylla Chama macerophylla Chama macerophylla Chama macerophylla Chama congregata 624 1a Chama sarda Chama congregata 624 1b Chama inezae Chama radians Chama congregata 629 9 Chama florida Chama macerophylla Chama macerophylla Chama macerophylla Chama macerophylla Chama congregata 624 1а Chama sarda Chama congregata 624 1b Chama inezae Chama radians Chama congregata 629 9 9.1 FIG. 1. Phylogenetic analysis of the combined molecular data ITS1 + 5.88 + ITS2 and 16S rRNA sequences. A: Strict consensus tree of 50 most parsimonious trees (length = 879, Cl = 0.89, RC = 0.82). Values below branches indicate boot- strap support; В: One of four maximum likelihood trees (HKY85+G model, logL = 3416.604, ti/tv ratio = 0.771, gamma shape = 0.89824). The other trees differ only in the branch- ing order of the Chama macerophylla sequences. bp of the 3° end only. The alignment had 813 positions, of which 355 were parsimony infor- mative. Parsimony analysis of the ITS data returned six parsimonious trees (length = 304, Cl = 0.93, RC = 0.89), whereas the 16S rRNA data yielded a single most parsimonious tree (length = 575, Cl = 0.87, RC = 0.76). Except for the taxa present in only one of the data sets the ITS strict consensus tree and the 16S rRNA tree were fully congruent. The combined analysis returned 50 shortest trees (length = 879, Cl = 0.89, КС = 0.82). The strict consensus tree (Fig. 1A) separated two major groups: one contains Chama macerophylla and C. florida, the other C. sarda, C. congregata, C. inezae, and C. ra- dians. Bootstrap support for the branches in the strict consensus tree was generally high (> 90) except for the branching order of C. congregata 624-1b, C. sarda, and C. inezae. Similar congruence was found for the maxi- mum likelihood trees of the two data sets, al- WESTERN ATLANTIC RECENT CHAMIDAE 385 though they differed slightly in the position of Chama florida nesting in C. macerophylla in the ITS tree (logL = 3416.604, ti/tv ratio = 0.771, gamma shape = 0.89824) as opposed to База! to it in the 16S rRNA tree (logL = 3314.612, ti/ tv ratio = 2.576, gamma shape = 0.41149). Four maximum likelihood trees, differing only in the branching pattern ofthe С. macerophylla speci- mens, resulted from the analysis of the com- bined data (Fig. 1B; logL = 7101.247, ti/tv ratio = 1.146, gamma shape = 0.648739). Both genes supported the same basic branching pattern, independent of the phylo- genetic method employed. The Chama macerophylla specimens from three different localities form a robust monophyletic clade with C. florida as sister taxon. In contrast, the С. congregata sequences - even if from the same individual, are polyphyletic. The weak support in this area of the tree may also be due to the missing 16$ data for С. congregata 624-1 and the missing ITS data for С. inezae. Morphological Characters “The mutations within the species of Chama are quite marked. They comprise color varia- tions which are often quite striking, as lemon- yellow and pale or dark purple in C. macerophylla, profuse, sparse, or obsolete fo- liation, and such changes of form as are due to the object upon which they are fixed. In dis- criminating species these fluctuations should be taken into account by the student, but it will also be found that there are features which are tolerably constant and which, after due dis- crimination, will be found to serve as guides to specific identity” (Dall, 1903: 1397). In the Chamidae, large, adult shells from wrecks, buoys, or protected portions of quiet- water reefs are the most diagnostic specimens for deriving species concepts. Such material is rare, but spectacular of form, color, and or- nament. By contrast, the encrusted, subadult specimens typical of reef and dredge samples can be a challenge to identify. Among the ge- nus or species level discriminators given in lit- erature are right or left valve attachment, size and shape of the prodissoconch, hinge pat- terns, absolute size, presence or absence of internal margin crenulations, pattern of pallial line attachment to muscle scars, color or color pattern, patterns of sculpture, and relative in- flation of attached and free valves. Chamids live with either their left or right shell cemented to a hard substratum, which can considerably influence shell shape and surface ornament. In current literature, species which attach exclusively by the left valve; species which typically attach by the left valve, but may occasionally attach by the other (Lamprell 8 Whitehead, 1992; Matsukuma, 1996); and spe- cies which attach indiscriminately to produce a roughly 50:50 balance (Yonge, 1967) are all assigned to the genus Chama. The Indo-Pa- cific genera Eopsuma and Carditachama are separated by their primitive hinge, retaining cardinal and lateral teeth. Eopsuma includes both species that attach by the right valve only, and by right or left indiscriminately (Matsu- kuma, 1996). Spiny, equilateral chamids, at- taching by the right valve as juveniles, can be assigned to Arcinella. Classification of the exclusively right at- tached chamids with a typical Chama hinge was problematic. Depending upon the author, such species represent either a valid, mono- phyletic clade assigned to Pseudochama, a form genus using Pseudochama for conve- nience, or represent Chama with reversed а+- tachment, synonymizing Pseudochama with Chama (e.g., Yonge, 1967). The concept of Pseudochama has been controversial since its introduction (Odhner, 1919; Yonge, 1967; Matsukuma et al., 1997). Most authors who use Pseudochama apply it as a form genus. Used in this sense, Pseudochama are most diverse in the tropical eastern Pacific, with Keen (1971) documenting six Pseudochama species sympatric with eight Chama species. An intriguing feature is the transposition of hinge teeth. Dentition usually is not determined by the left-right symmetry as in other bivalves, but by which valve attaches to the substra- tum. Thus, an attached valve, whether right or left, almost always exhibits the same denti- tion. Odhner (1919) placed great emphasis on differences in the hinge formulas of Chama and Pseudochama, making the hinge a key argument for separation of the genera, but these differences appear to be nothing more than the mirror image reversals achieved by transposition (Davis, 1935; Kennedy et al., 1970). Very rarely specimens have been found with transposed shell attachment but normal dentition (Matsukuma et al., 1997). Starobogatov (1992) argued for separating Arcinella and Pseudochama into an entirely different superfamily, Arcinelloidea Scarlato 8: Starobogatov, in Nevesskaya et al. (1971: 17), in the rudist suborder Hippuritoidei, based on the supposed lack of heterodont dentition. However, the nepionic shell in Pseudochama has two cardinal teeth before metamorphosis 386 CAMPBELL ET AL. (Keen, 1971: 151). Kennedy et al. (1970: 406- 410) compared in detail chamid shell and den- tition with every family of rudists, and rejected the hypothesis of a rudist ancestry. Maximum adult size appears to be a spe- cies-specific {гай within a given fauna. п the Florida Keys we found small (Chama sarda), medium (C. congregata), and large taxa (C. macerophylla). Maximum size has been fre- quently cited as a discriminating character, for example in separating the Late Pliocene fos- sil С. willcoxii Dall from the Miocene to Re- cent C. macerophylla Gmelin. However, Allen (1977: 258) found evidence for a three-year lifespan in C. gryphoides, and Epstein 8 Lowenstam (1953: 432) documented that C. macerophylla in Bermuda produced shell growth only in the warmest season of the year. Consequently, it appears that maximum size has both genetic and environmental compo- nents. More tropical, central to southern Car- ibbean С. macerophylla may be 20 to 40 mm larger than typical Floridian material (Pilsbry 8 McGinty, 1938), perhaps reflecting a greater number of days warm enough for shell growth across a brief lifespan. In turn, the greater size of С. willcoxii may reflect the milder Florida winters during mid- to late Pliocene seas. Maximum size proves helpful when dealing with adult specimens, but subadult specimens of the larger taxa are often conformable to and easily confused with the smaller species. The presence or absence of internal margin crenulations is frequently cited as a species- level discriminator (Dall, 1903; Abbott, 1974). In our study, we have found no smooth-mar- gin species, with an occasional crenulate in- dividual. One specimen of Chama inezae (USCS collections) adjusted a wide, shell-edge lamella by pleating the inner edge, but careful examination revealed no true crenulations. We have examined over 100 specimens each of the fossil species C. congregata Conrad, 1833, and Pseudochama corticosa (Conrad, 1833), and find them to be consistently crenulate. However, five of 40 specimens of the Recent C. radians lacked the crenulations typical of that species. We conclude that presence or absence of crenulation is a constant factor in some western Atlantic species, but is variable in others. Consequently, individual specimens, including holotypes, may or may not express the normative pattern. This caution also ap- plies to the identification of single specimens from a fossil site or dredge haul. Chamid ontogeny expresses a break in shell growth corresponding to larval settlement and a second break corresponding to cementation to a hard substratum (Kennedy et al., 1970: 382-384). Some authors (Odhner, 1919, 1955; Kennedy et al., 1970; Bernard, 1976; Matsukuma, 1996) used the term “prodisso- conch’ to refer exclusively to larval growth, and the terms “dissoconch” or “dissoconch |” for shell growth between larval settlement and cementation. Other authors (Cox et al., 1969; Waller, personal communication, March 2003) apply the term “prodissoconch” to all shell growth before cementation. Here we follow the latter usage. The size and shape of the prodissoconch have been used to separate Chama from Pseudochama, with Chama species having a rounded prodissoconch of less than 1 mm but Pseudochama species having a rectangular prodissoconch of greater than 2 mm (Odhner, 1919). Subsequent investigation indicates that these characters do not appear to be defini- tive at the genus level in western Atlantic chamids. Most larger chamid specimens are encrusted, abraded, or bored, and the prodissoconch is destroyed. Smaller, post- metamorphic shells (5-7 mm in diameter) in many cases preserve the prodissoconch, but at that size they have not established diag- nostic morphology, and the species can be difficult to identify. Prodissoconch form remains undescribed for most chamid species. Among the few Chama and Pseudochama species for which the prodissoconch has been described, there are overlapping ranges of variation. Chama species that preserve this early growth stage often have a round to oval prodisso- conch, 0.5 to 2 mm in diameter, which 1$ smooth or has radial sculpture. In contrast, documented Pseudochama and Arcinella gen- erally have an oval to rectangular prodisso- conch that is 1.5 to 2.5 mm in diameter, and has concentric lamellar sculpture before ce- mentation (Odhner, 1919; Matsukuma, 1996). Odhner (1919: pl. 1, fig. 7) illustrated a “de- finitive” prodissoconch for P. ferruginea, a syn- onym of C. radians, and Redfern (2001) illustrated a large, rectangular prodissoconch for С. radians. However, Ferreira & Xavier (1981: fig. 4) illustrated a similar large, oval prodissoconch for C. macerophylla, and Dall (1903: 1395) reported a 2-mm prodissoconch for C. lactuca. Odhner (1919: 14) first invoked misidentification as the explanation for Dall's (1903) report of Chama prodissoconchs larger than 1 mm, and later described a “transition” zone in the metamorphosis of Chama (Odhner, 1955) to maintain the supposed size differ- WESTERN ATLANTIC RECENT CHAMIDAE 387 TABLE 3. Stratigraphic and geographic distribution of Chama and Pseudochama in the western Atlantic Ocean: New Jersey to Florida and Gulf of Mexico, Lower Miocene to Upper Pliocene. Distribution New Jersey to Maryland Virginia to north of Cape Hatteras, NC North Carolina to northern Florida Southern Florida and Florida Keys Western Florida and Gulf of Mexico Lower & Middle Miocene C. congregata Chama sp. C. chipolana P. draconis P. corticosa Lower & Middle Upper Miocene Pliocene Upper Pliocene - C. congregata C. congregata С. emmonsi C. emmonsi P. corticosa - C. congregata C. congregata C. emmonsi C. emmonsi Р. corticosa - C. congregata C. congregata С. emmonsi C. emmonsi Chama. sp. С. heilprini P. caloosana С. willcoxii P. caloosana - C. congregata C. emmonsi ences. Kennedy et al. (1970: pl. 71) illustrated a circular, cancellate prodissoconch for C. pellucida, an oval, lamellar prodissoconch for A. arcinella, and a small, circular lamellar prodissoconch for Chama sp. We conclude TABLE 4. Stratigraphic and geographic distribution of that a small, round prodissoconch is more typi- cal of Chama, and that a larger, rectangular stage may represent “Pseudochama” of au- thors, or Arcinella. The overlap 1$ considerable and is non-definitive. We have been unable to Chama and Pseudochama in the western Atlantic Ocean: Cuba and Greater Antilles to Argentina, Lower Miocene to Upper Pliocene. Lower € Middle Lower & Middle Distribution Miocene Upper Miocene Pliocene Upper Pliocene Cuba, Greater Antilles, - C. caimitica C. macerophylla - Bahamas C. involuta C. involuta P. riocanica Lesser Antilles - - - C. macerophylla Pseudochama sp. aff. P. caloosana Central America - - C. macerophylla - Panama C. strepta C. berjadinensis - - Colombia, Venezuela C. berjadinensis P. buchivacoana C. berjadinensis - - P. corticosaformis P. quirosana P. scheibei Brazil - Argentina - P. lazai С. paschauli - - 388 CAMPBELL ЕТ AL. TABLE 5. Stratigraphic and geographic distribution of Chama and Pseudochama in the western Atlantic Ocean: North Carolina to Central America, Lower Pleistocene to Recent. Distribution North Carolina to northern Florida Southern Florida and Florida Keys Western Florida and Gulf of Mexico Cuba, Greater Antilles, Bahamas Lesser Antilles Central America find any published description of the prodisso- conch of C. cristella Lamarck, 1819, type spe- cies of Pseudochama. The adductor muscle scars in the Chamidae are large, elongate, and are often described as reniform. In most species, the muscle scars are subequal, and are connected by a simple pallial line with no sinus. Attachment of the pallial line typically is at the center of the ven- tral margin of the adductor scars, but consid- erable variation exists within conspecific populations. Pallial line attachment can meet Lower Pleistocene C. congregata - C. macerophylla С. emmonsi Upper Pleistocene Recent C. macerophylla . congregata . macerophylla . radians . lactuca congregata . macerophylla florida sinuosa radians lactuca sarda inezae . congregata . macerophylla radians lactuca C. congregata C. macerophylla C. florida C. sinuosa C. radians congregata . macerophylla florida sinuosa radians sarda congregata macerophylla florida sinuosa radians lactuca sarda congregata . macerophylla florida sinuosa radians . Sarda 0000000000000 000000 0000 00000000 0000 the ventral margin at the outer edge, inner edge, or center; or the pallial line can extend dorsally and attach to the outer or inner lateral margin of the adductor muscle scar. The latter condition has been erroneously described as a pallial sinus. Bayer (1943) cited attachment at the ventral margin of the anterior adductor muscle scar for Chama florida and attachment at the outer lateral margin of the scar as char- acteristic of C. sarda. Although we find this at- tachment pattern more common in C. sarda, it is not a consistent trait at the species level. WESTERN ATLANTIC RECENT CHAMIDAE 389 TABLE 6. Stratigraphic and geographic distribution of Chama and Pseudochama in the western Atlantic Ocean: Panama to Argentina, Lower Pleistocene to Recent. Distribution Panama - C. sinuosa C. radians Colombia, Venezuela Brazil 2 Argentina - Systematics Zoogeographic and stratigraphic distribu- tions are given from literature (Tables 3-6). Authority for particular distribution records can be found in the literature cited in the synony- mies, and is generally not repeated in the Dis- tributions. Institutional abbreviations for specimens examined or illustrated are as fol- lows: AMNH, American Museum of Natural History, New York, New York, USA; ANSP, Academy of Natural Sciences of Philadelphia, Pennsylvania, USA; FLMNH, Florida Museum of Natural History, Gainesville, Florida, USA; FMNH, Field Museum of Natural History, Chi- cago, Illinois, USA; MNHG, Museum of Natu- ral History of Geneva, Switzerland; USCS, University of South Carolina Spartanburg, South Carolina, USA. Family Chamidae Lamarck, 1809 Genus Chama Linnaeus, 1758 Type species: Chama lazarus Linnaeus 1758 (ICZN Opinion 484, 1957). Chama lazarus is from the Indo-Pacific, not the Mediterranean, as cited by Abbott (1974: 466). Matsukuma (1996: 29) gave the following characters for Chama: “Shells usually attached by the left valve, entirely aragonitic, with outer Lower Pleistocene Upper Pleistocene Recent congregata macerophylla florida sinuosa radians sarda congregata macerophylla florida sinuosa radians congregata . macerophylla florida sinuosa radians С. iudicai - C. macerophylla C. florida C. macerophylla 00000 00000 000000 crossed-lamellar layer, middle myostracal layer, and inner complex crossed-lamellar layer. Nepionic shell small, less than 1.2 mm long, prodissoconch without ornament, early dissoconch having minute sculpture of closely spaced radiating striae, punctations and some- what more distant commarginal riblets. Hinge of the nepionic shell and adult ‘pachydont’- type; hinge formula of the nepionic shell: 3a, 3b, LPI in the right valve and 2, 4b, LPII in the left; each tooth somewhat parallel to the hinge plate. Adult free right valve: single long, nar- row cardinal (3a) + 3b, a wide ventral socket below the cardinal, parallel to hinge plate, den- ticles (5b) below nymph, and a posterior lat- eral (LPI); adult attached left valve: a broad anterior cardinal (2) parallel to the hinge plate, a dorsal socket, a weak, long, narrow, poste- rior cardinal (4b), and a posterior lateral (LPII).” Genus Pseudochama Odhner, 1917 Type species (subsequent designation: Gardner, 1926): Chama cristella Lamarck, 1819 (Figs. 11, 12). Lamarck's original handwritten label stated a locality of “Océan des grandes indes”. A more recent label with the type speci- men indicated “Ocean indien” (= Indian Ocean). The holotype of Chama cristella Lamarck 1$ from the Delessert collection (Fig. 11; MNHG 390 CAMPBELL ЕТ AL. 1087/6). In Lamarck's description, he made ап incidental reference to Chemnitz (1786: fig. 993). Odhner (1917, 1919) based his type con- cept of Pseudochama on specimens of “Chama cristella Lamarck”, but did not desig- nate a type species, listing Р. cristella (Lamarck) and P similis Odhner, 1917, in the genus. He hypothesized that all species with a typical chamid hinge and right valve attachment should be placed in Pseudochama. Gardner (1926) designated C. cristella Lamarck, 1819, as the type species, a designation indepen- dently given by Prashad (1932: 295) (Nicol, 1952, b). Sandberg & Waren (1993: 120) cited the type species of Pseudochama Odhner, 1917, as Р similis by original designation. How- ever, Odhner implied but did not state a type species, an opinion confirmed by Warén (per- sonal communication, August 2003). Despite the extensive debate over the valid- ity of Pseudochama, we have been unable to locate any illustration of undoubted Р cristella published subsequent to the mid-nineteenth century. Lamy (1928: 347) noted the nomen- clatural confusion involving Chama cristella Lamarck, 1819; C. sinistrorsa Bruguiére, 1792; and Chemnitz (1786: figs. 992 and 993). Lamy (1928) cited Chemnitz (1786) for a locality of West Indies (St. Croix) for figure 992, and of West Indies for figure 993. Bruguiere (1792: 392) cited Chemnitz figure 992 and a locality of the Greater Antilles for his new species C. sinistrorsa. Lamarck (1819: 96) described C. radians based on a specimen in his collection (Fig. 7A; MNHG 1087/3), and referenced Chemnitz figure 992 as conspecific. Similarly, Lamarck described C. cristella based on a specimen in the Delessert collection (Fig. 11; MNHG 1087/6) and also referenced Chemnitz figure 993. In Lamarck’s description, he noted that C. radians was a different species from C. sinistrorsa Bruguiére, 1792. Lamy con- cluded that С. cristella Lamarck, 1819, and С. sinistrorsa Bruguière, 1792, were conspecific with Chemnitz’ figure 993, leaving Chemnitz figure 992 as conspecific with C. radians Lamarck, 1819. Lamy listed “Chama sinistrorsa ?Bruguiére, 1792”, as a synonym under the more recently described C. cristella Lamarck, 1819. Resolution of this problem is beyond the scope of this study. Bucquoy et al. (1892: 312) thought that Chemnitz’s figure 992 could depict Chama ruppelli Reeve, 1847, but Lamy (1928) stated that C. ruppelli Reeve was a distinct species from Chemnitz’s figure 992. Clessin (1889: 38, pl. 16, figs. 3, 4) thought that C. cristella as figured in Reeve (1847) differed from C. cristella Lamarck, 1819. Clessin named Reeve's figure С. reevana Clessin, 1889, and applied Lamarck’s species С. cristella to a shell from Puerto Rico. However, Lynge (1909: 265) said that the figure of Reeve (1847) was con- specific with C. cristella, and the Clessin 1889 specimen from Puerto Rico was not C. cristella. Lynge (1909: 265) quoted Reeve (1847): “the example here figured has been satisfactorily identified with Lamarck’s original specimen in the collection of M. Delessert.” The type specimen of Chama cristella Lamarck (Fig. 11) appears to be very close, if not conspecific with Pseudochama similis Odhner, 1917, as figured by Matsukuma et al. (1997: 229, figs. 7a-d). Reeve (1846: pl. 3, fig. 10b) figured a right-attached specimen, BM(NH) 1950.11.1.50, and a left-attached specimen (his fig. 10a), BM(NH) 150.11.1.49 as two syntypes of C. pulchella Reeve, 1846. An unfigured syntype, BM(NH) 150.11.1.51, is also a left-attached specimen of C. pulchella Reeve, 1846. Odhner (1917) described P. similis based on the right-attached specimen BM(NH) 1950.11.1.50 (Reeve, 1846, pl. 3, fig. 10b). Lamprell & Whitehead (1992: pl. 23, fig. 149) synonymized P. similis with C. pulchella, but Matsukuma (1996: 36) argued that Reeve’s figure 10b represented a valid right- attached species. Matsukuma et al. (1997, figs. 7a-d, 8a-d, 9) illustrated the three Reeve (1846) syntypes for С. pulchella, with P. similis as figures 7a—d. They cited a second possible specimen of P. similis from Tonga. Matsukuma et al. (1997) argued that Odhner correctly separated the right-attached syntype as P similis, because its sculpture pattern distinctly contrasted with the two left-attached syntypes. Higo et al. (2001) also illustrated this right-at- tached British Museum syntype, but gave its number as BM(NH) 150.11.1.49, identifying it as a syntype of Chama pulchella Reeve. Matsukuma (1996) followed Odhner (1919) in emphasizing anatomical differences be- tween Chama and Pseudochama. Matsukuma (1996: 46) listed the following characters for Pseudochama: “animal without lateral caecal appendage of the stomach, nephridia with the pericardial tubes not covered on their median side by distal sacs; shell usually attached by the right valve; nepionic shell sculptured just as in Arcinella Schumacher, 1817, with rather distantly spaced commarginal lamellae, and no or very fine punctuations or only traces of radiating riblets; the size of early dissoconch more considerable, length up to 1.0-2.5 mm; WESTERN ATLANTIC RECENT CHAMIDAE 391 hinge formula of the nepionic shell: 1, 3b, LPI in the right valve and (2a), 2b, (4b), РИ in the left; hinge formula of adult shell: 1 + 3b, 5b, [РИ in the right valve and (2a), 4b, (РИ in the left.” We add no anatomical details, but note that Kennedy et al. (1979) had documented left-attached species with lateral appendages on the stomach and left-attached species with- out lateral appendages on the stomach. Matsukuma (1996) discussed “normal” and “reverse” attachment, and the various chamid genera and species in which both forms oc- cur. He restricted the genus concept of Pseudochama to Р. cristella, the type species of the genus, and to those species in strict conformity of valve attachment, hinge form, shell form, prodissoconch, and anatomy to this type species. However, prodissoconch form remains unknown for P cristella. Odhner (1917, 1919) based his type concept of Pseudochama on specimens of “Chama cristella Lamarck” which Dr. Mjoberg brought from the northwestern coast of Australia. Lamprell & Whitehead (1992) and Lamprell & Healy (1998) did not recognize P. cristella from Australian waters, but noted several species which are typically or occasionally right-at- tached. Slack-Smith (1998: 308, figs. 8.1d, e) illustrated as “Pseudochama sp.” a specimen from Australian waters that resembles P. sp. cf. P. cristella (ANSP 54808) from Java (Figs. 12A-C). Most authors who cite Р cristella in current literature limit its distribution to Thai- land and Indonesia. Lamy (1928) cited a dis- tribution of Indonesia, Thailand, the Indian Ocean, and possibly South Australia for P. cristella. The holotype of Chama cristella Lamarck, 1819 (Fig. 11), contrasts with specimens from Java identified as Pseudochama cristella and housed in the Academy of Natural Sciences Philadelphia collection (Figs. 12А-С). In both the holotype and the ANSP Javan specimens, the attached valves show a similar attachment pattern, strong fluting adjacent to the line of attachment, more subdued fronds or low flut- ing adjacent to the posterior edge, and rare to common granular nodes between the two rows of fluting. General details of the interior of the valves are conformable for both lots, although Javan specimens show a stronger knob on the hinge. However, the external sculpture of the free valves differs markedly. The holotype of P. cristella shows a free valve with large, widely spaced lamellae that are scalloped into large flutes. This fluting contrasts with the three Javan specimens that show a consistent pat- tern of fine, crowded lamellae broken up into scale-like fronds. At the other extreme, the sculpture of the type of P. similis, from Matsukuma et al. (1997: 229, fig. 7a—d) is even more fluted and elaborate, but the pattern matches that of the P. cristella holotype. The question remains whether the Javan speci- mens are Р cristella with subdued and more crowded sculpture, or whether they represent a distinct species. Such range of variation is present for some chamid species, but not in others. Given the rarity of the taxa in ques- tion, we suggest that the ANSP lot from Java be referenced as Pseudochama sp. cf. Р. cristella. Resolution of these questions is es- sential in defining the range of variability within the type species of the genus Pseudochama Odhner, 1917, but a conclusion is beyond the limits of this study. Kilburn & Rippey (1982: 174, pl. 39, fig. 9) reported Pseudochama cristella (Lamarck) from “Indo-Pacific to western Transkei”. They described and figured a small (to 30 mm), right- attached form, the size, color, and sculpture of which differ considerably from Lamarck’s holotype. This South African taxon is probably not Р cristella. Odhner’s (1919) hypothesis of a pervasive, monophyletic Pseudochama, defined by a typi- cal chamid hinge and right valve attachment, can be no longer supported. In the western Atlantic, right-attached taxa first appeared in the Middle Eocene, and continued as a scarce element into the Recent tropical and subtropi- cal faunas. The right-attached condition ap- pears to have arisen independently by valve transposition several times in western Atlan- tic and eastern Pacific fossil and Recent fau- nas. For example, the Californian Р exogyra (Conrad, 1837) and sympatric Chama arcana (Bernard, 1976) (= C. pellucida of authors, non Broderip, 1835) differ in valve attachment, but share an otherwise rare outer shell layer of calcite (Kennedy et al., 1970: 388-389) from which they were judged to be closely related. We agree with Matsukuma (1996) that a valid concept of Pseudochama must conform rig- orously to the type species, but many uncer- tainties surround P. cristella and congeneric status with Chama radians cannot be presently established. In the western Atlantic, there are no extinct Pseudochama species with right attachment that are morphologically an obvi- ous ancestor of C. radians. Pilsbry & McGinty (1938) observed that certain C. radians ap- pear to be mirror images of Recent C. congregata. Chama radians is consistently 392 CAMPBELL ET AL. right-attached, and has a rectangular prodissoconch of about 2 mm. In our molecu- lar analysis of the ITS gene, C. radians falls within a cluster of Chama species. The hypoth- esis of a right-attached monophyletic clade predicts that all species of “Pseudochama” should stand apart. Even Odhner (1919: 81- 82) informally divided his genus into a P. cristella “group” and a P. radians “section”. While recognizing that more systematic and molecular work needs to be done, including analysis of multiple individuals of several dif- ferent “Pseudochama” species, we are return- ing P. radians to the genus Chama. CHAMID SPECIES OF SOUTHERN FLORIDA Chama congregata Conrad, 1833 Figures 2А-С ?Chama foliacea Gmelin, auctt., non Gmelin, 1791: 3304, based on Lister, 1685: pl. 215, по: 51. Chama congregata Conrad, 1833: 341; Dall, 1903: 1400-1401; Glenn, 1904: 342, pl. 91, figs. 1-3; Olsson, 1922: 218, pl. 28, fig. 11; Lamy, 1928: 329; Bayer, 1943: 120, pl. 12, fig. 3; Weisbord, 1964: 235-238, pl. 31, figs. 11-14, pl. 32, figs. 5, 6 (extensive syn- опуту); Hoerle, 1970: 58; Waller, 1973: 41, 48; Abbott, 1974: 466, pl. 21, fig. 5385; Cerridwen & Jones, 1991: 100; Campbell, 1993: 30, pl. 8, fig. 76; Diaz & Puyana, 1994: 74-75, pl. 16, fig. 152; Rios, 1994: 260, spe- cies 1266, pl. 89; Redfern, 2001: 214, spe- cies 876, pl. 89. Chama congregatoides Maury, 1917: 200, pl. 33, 10. 8: Туре Locality Miocene, James River, Virginia “near Smithfield” (Conrad, 1833: 341). Emended by Campbell (1993): Pliocene, Yorktown Forma- tion, James River, Virginia. Numerous out- crops were available to Conrad in the riverbanks near Smithfield, such as Rock Wharf, a known Conrad locality with a high molluscan diversity. Remarks on Early Literature Dall (1903: 1401) said of Recent Chama congregata: “This appears to be the species said to be abundant in Cuba, which 1$ cited by Arango (1878-1880: 272) as Chama foliacea Gmelin, based chiefly on Lister’s figures. It is, however, too uncertain to be adopted even if the specific name was not so glaringly inap- propriate.” Chama foliacea Gmelin, 1791, was originally described from “Mediterranean and Americas,” based on Lister (1685: pls. 215- 217, figs. 51, 53). Lister had fossil specimens from Colonial Virginia (Campbell, 1993), and modern “Colonial Williamsburg” gravels its paths with C. congregata. Lister (1685) some- times gave localities, for example, plate 215, figure 50, portrayed a Chama with radiating color bands (probably C. florida) from “Barb. jamaic” (Barbados and Jamaica). Some of his Virginia fossils have the designation of “virg”. However, no Lister (1685) locality information accompanied figures 51 and 53, and the two are not conspecific. Lister's figure 51 was сор- ied by Klein (1753: pl. 12, fig. 81), a reference also cited by Gmelin (1791). Figure 51 is an upper valve of a Chama, showing the radial grooving of typical C. sinuosa, but lacking in all variations of C. congregata (see below). Figure 53 depicts an upper valve not showing the radial grooving. lt is larger, and shows соп- siderable bioerosion of the ventral margin. Such damage is unusual for C. congregata, but common in larger chamid specimens. The third citation given by Gmelin (1791) is Chemnitz (1783: pl. 52, fig. 521). Chemnitz figured a Chama with a very low, appressed beak, and with the attached valve hardly larger than the free valve. Chama congregata pos- sesses a high, “naticiform” beak, and is clearly not conspecific with that figure. The last figure cited by Gmelin (1791) is Lister (1685) plate 215, figure 50, which was considered above. Consequently, we concur with Dall (1903) in rejecting C. foliacea (Gmelin) as a senior syn- onym for C. congregata. Bruguiére (1792) pro- posed C. rugosa on essentially the same concepts, citing Lister (1685: fig. 53), Klein (1753: fig. 81), and “Martini” [= Chemnitz], fig. 521, along with two other references not given by Gmelin. Consequently, whatever C. foliacea is, С. rugosa would appear to be a synonym. Lamy (1928: 327) noted that C. foliacea Quoy & Gaimard (1835) is a junior homonym of C. foliacea Gmelin. A second complicating element to the spe- cies concept of Chama foliacea Gmelin from the older literature concerns C. lamellosa Lamarck, 1806. Lamarck (1806: 348) listed fossil species of Chama, beginning with C. squamosa (Brander, 1766 [= Solander]), which he renamed С. lamellosa. This usage has con- WESTERN ATLANTIC RECENT CHAMIDAE FIG. 2. Chama spp. A: Chama congregata, right and left valves, IMBW-FK-624, Horseshoe Reef, off Fat Deer Key, Florida, 7.3 т (AMNH 306412); В: С. congregata, right and left valves, IMBW-FK-629, “The Horseshoe” site, West Summerland Key (Spanish Harbor Keys), Florida (FMNH 301422); C: C. con- gregata, right and left valves, Missouri Key, Monroe Co., Florida, (FLMNH 127093); D: C. macerophylla, right and left valves, IMBW-FK-625, Coffins Patch Sanctuary Preservation Area, off Crawl Key, Florida, 6.4 m (AMNH 306419); E: C. macerophylla, right valve, IMBW-FK-629 (FMNH 301423); F: C. macerophylla, right valve, Key Largo, Florida (FLMNH 2271). Scale bars = 5 mm (А-С); 10 mm (D-F). 394 CAMPBELL ЕТ AL. tinued. Brander's species was from the Eocene Barton beds at Hampshire, England (Natural History Museum, 1975). Brander's il- lustration portrayed a medium-sized Chama with a high, strongly recurved, “naticiform” at- tached valve, similar to C. congregata. This being the only work cited by Lamarck, 1806, the species concept of C. lamellosa should be uniquely tied to C. squamosa, and become a junior synonym. However, Lamarck (1819: 98) later expanded the concept of C. lamellosa to include C. rugosa Bruguiére, 1792 [non Linnaeus, 1771] and Chemnitz (1783: pl. 52, fig. 521), which Gmelin included in С. foliacea. As none of the taxa from the 1819 expansion appear conspecific with C. squamosa (Brander), they may be rejected from C. lamellosa (sensu stricto). Chama squamosa (Brander) (= С. lamellosa Lamarck) is conver- gent, but not conspecific, with С. congregata. Remarks on Chama congregata Three or more fossil and Recent morpho- species appear to be present under the name “Chama congregata”. Additionally, we report under molecular systematics that DNA se- quencing of multiple specimens of C. congregata within the Recent Florida Keys population indicates that multiple cryptic spe- cies may be present. Here we treat each form separately and, for the present, retain all forms under the traditional name. Chama congregata Conrad, 1833, original lot of nine fossil syntypes from the James River, Yorktown Formation, Pliocene of Virginia. Description Small to medium sized (to 25 mm, rarely to 30 mm); both valves with small, short, flat, smooth scales arranged in radial rows; at- tached valve with very small attachment area, mostly free growth, producing a deeply cupped, “naticiform” valve; upper valve circu- lar, somewhat inflated, lacking or with only the slightest hint of a depressed central radial fur- row; inner margin crenulated. Distribution Miocene, Kirkwood beds, New Jersey; Mi- ocene, Calvert Formation, Maryland. Glenn (1904) figured specimens from the Calvert that appear conspecific with Yorktown material. Lower Pliocene, Yorktown Formation, Zone 1 and Zone 2, Virginia, Yorktown Formation, North Carolina; Goose Creek Limestone, Raysor Marl, Duplin (at Natural Well); Jack- son Bluff Formation, western Florida; Pinecrest bed 7, Sarasota, Florida; Upper Pliocene, Waccamaw Formation, Carolinas; Caloosa- hatchee Formation, southern Florida. We are not aware of any Recent specimens conform- able with Conrad's Yorktown Formation types. Discussion Calvert specimens from Plum Point, Mary- land (USCS collections), are larger, have raised lamellar sculpture, and are wider than high. This latter form is probably an undescribed species. А similar form was illustrated by Ward (1992) from the Belgrade Formation of the Upper Oligocene of North Carolina. Ward (1992) used the name “Chama chipolana Dall, 1903”, for these specimens, but Ward's Шиз- trated topotypes do not appear conspecific with Dall's illustrated syntypes of С. chipolana, nor with Chipola specimens on loan from the Florida Museum of Natural History. Given the possibility of multiple species, all Kirkwood and Calvert Formation records of C. congregata need to be critically reexamined. Although Conrad’s types were from the south bank, along the north bank of the James River at King's Mill (another Conrad locality), and in several localities around Williamsburg, Vir- ginia, Chama congregata is the dominant ele- ment in bioherms or reefs 5-10 m thick and 200 m or more wide. The millions of complete specimens found in these reefs give context to Conrad's name for the species. In the Chama reefs, individuals were unattached or were loosely attached to their neighbors. Other shell substrata, such as large valves of Chesapecten, Mercenaria, and Ostrea, were common in these reefs and on the Yorktown sea floor, but appear unexploited as attach- ments by this form. Similar concentrations of paired C. congregata with no reef development have been found in Early Pliocene deposits near Lynchburg and Florence, South Carolina. “Chama congregata” of Upper Pliocene to Recent deposits, North Carolina and South Carolina to Florida and the West Indies. Description Shell small to medium-sized (15 to 20 mm, rarely to 30 mm), usually attached to rock, coral, or shell substrata; attachment area large, with specimens typically having the anterior edge of attached valve in contact with substratum, WESTERN ATLANTIC RECENT CHAMIDAE 395 giving right (unattached) valve a semicircular outline; attached valve shallow-cupped, not “naticiform”; sculpture suppressed and scaly in the furrow, but coarsely frondose on the flanks; shell outline somewhat rectangular; inner mar- gin crenulated. Bayer (1943: pl. 12, fig. 3), Weis- bord (1964: pl. 32, figs. 1, 2), and Redfern (2001: pl. 89, figs. 876А, B) illustrated large specimens in which the unattached valve has a very broad and depressed central radial furrow. Distribution Upper Pliocene, lower and upper Waccamaw Formation at Calabash, North Carolina. Late Pliocene, Nashua Formation, northeastern Florida. Early Pleistocene, Bermont beds, southern Florida. Upper Pleistocene, Venezu- ela and Grand Cayman. Recent, shallow to mid- shelf, Cape Hatteras, North Carolina, to Florida, Texas, Bermuda, and the West Indies. Redfern (2001: 214) reported this morphology as com- mon in two to five m depth in the Bahamas. “Chama congregata” Recent of Brazil Description “Valves rounded (35 x 28 mm), grayish to red in color. Right valves smaller than the left. Surface with axial, wavy corrugations. Ventral margins slightly crenulated. Attached to rocks, corals, and other shells, usually in colonies at low tide and on mangrove roots” (Rios, 1994: 260). Pilsbry & McGinty (1938: 75) cited a maximum size of 36 mm. Discussion The large size and subdued sculpture sug- gest that Recent “Chama congregata” of Bra- zil is a different species. Both attributes may be a consequence of living in shallow subtidal, higher energy waters. The larger size could also be related to the effects of water tempera- ture on shell growth (Epstein 8 Lowenstam, 1953; Lowenstam, 1954), or a combination of these factors. Alternatively, this may prove to be a Recent extension of С. iudicai Pastorino, 1991, from the Pleistocene of Argentina. Chama macerophylla Gmelin, 1791 Figures 2D, 3A Chama gryphoides (ex parte) Linnaeus, 1767: 1139. Macerophylla, Flos Maris Chemnitz, 1783: 101, 149, pl. 52, figs. 514, 515. Chama macerophylla Gmelin, 1791: 3304; Dall, 1903: 1403; Woodring, 1925: 104-105, pl. 12, figs. 18, 19; Lamy, 1928: 308 (exten- sive synonymy); Weisbord, 1964: 238-241, pl. 33, figs. 1, 2 (extensive synonymy); Richards et al., 1969: 4; Hoerle, 1970: 58; Scoffin, 1972: 1281; Waller, 1973: 41, 48; Abbott, 1974: 466, pl. 21, fig. 5384; Cerridwen 8 Jones, 1991: 100; Donovan & Littlewood, 1993: 37-38; Paulay, 2003: 223, 234. Chama citrea Gmelin, 1791: 3305. Chama imbricata Lamarck, 1801: 131; Krebs, 1864: 117, non Broderip, 1835: 149. Chama lazarus Linnaeus — Lamarck, 1819: 93, non Linnaeus, 1758. Chama macrophylla [sic] Gmelin — Hanley, 1843: 226. Chama bicornis Linnaeus — Krebs, 1864: 117, non Linnaeus, 1758: 692. Chama macerophylla var. purpurascens Poulsen, 1878: 15. Chama macerophylla var. sulphurea Poulsen, 1878: 15, fig. 14, non Reeve, 1846. not Chama macerophylla Gmelin — Melvill & Standen, 1907: 840, non Gmelin, 1791. Chama cf. macerophylla Gmelin Jung, 1969: 362, pl. 24, figs. 1, 2. 2Chama bermudensis Heilprin, 1890: 141, pl. 8, figs. 1, la (see Waller, 1973: 48). Type Locality Recent, “oceano americano” (Gmelin, 1791). Description Large, very foliaceous, with profuse, erect fronds that are radially striate; fronds small and abundant, or fewer, larger, longer within a single population; left valve not deeply cupped; attachment area extensive; inner margin 1$ crenulated; colors may be uniform or combi- nations of white, lemon yellow, and/or purple; interior ventral margin almost always stained with a band of brownish purple (Fig. 2D). In dredge sample or reef-survey, this spe- cies may identified by the combination of stri- ate fronds, internal color band, and crenulate margin. Distribution Miocene and Pliocene, Dominican Republic (Woodring, 1925: 104-105); Pliocene, Bowden beds, Jamaica. Pliocene (?), Trinidad (Jung, 1969). Pliocene, Limon, Costa Rica (Woodring, 1925: 104-105). Early Pleistocene, Bermont beds, southern Florida. Late Pleis- 396 CAMPBELL ЕТ AL. tocene, Grand Cayman (Cerridwen & Jones, 1991); Jamaica (Donovan & Littlewood, 1993), the Antilles and Curacao (Woodring, 1925: 104-105), Bermuda, North Carolina, Florida, Cuba, Dominican Republic, St. Kitts, Curacao, Aruba, and Venezuela. Recent, Bermuda, Cape Hatteras, North Carolina to Brazil. Loui- siana record from Garcia 4 Lee (2002). Paulay (2003: 223, 234) documented this species as a Pacific invasive in the waters around Guam. Discussion Chama macerophylla Gmelin, 1791, is com- mon on rock, reefs, wrecks, and sea walls. Woodring (1925: 104-105) recorded it from shallow water to 525 m. The radial striations are visible and pervasive. Chama sinuosa and other associated species may have an occa- sional frond showing radial striae, but that 1$ the exception. Small, encrusted specimens of C. macerophylla may appear convergent with С. congregata, but т С. macerophylla the in- terior ventral margin is almost always stained brownish purple (Fig. 2D), a color absent in C. congregata. Lamy (1928) provided an extensive syn- onymy. He concluded that Chama macero- phylla Gmelin of Melvill 8 Standen (1907: 840) from the Persian Gulf was not С. macerophylla Gmelin and instead probably was C. damaecornis Lamarck, 1819, a junior synonym of C. lazarus Linnaeus, 1758. Lamy (1928) also stated that C. imbricata Lamarck, 1801, was a synonym of C. macerophylla Gmelin, 1791. (Chama imbricata Lamarck, 1801, 1$ a senior homonym of С. imbricata Broderip, 1835, from the Red Sea and Indian Ocean.) Chama macerophylla was present in the Late Miocene of the Dominican Republic, and in the Middle Pliocene of Venezuela and south- ern Florida. These earliest populations were probably ancestral to С. willcoxii Dall, 1903, which was endemic to the Plio-Pleistocene Caloosahatchee Formation of southern Florida. Chama willcoxii was larger, and the fused fronds were more than twice the width of those of С. macerophylla. In С. willcoxii, the fronds were appressed, contrasting with the erect sculpture of C. macerophylla. Waller (1973: 48) documented heavy, deeply cupped specimens of Chama macerophylla from Walsingham Pond, Bermuda, which he compared with Chama bermudensis Heilprin, 1890. Abbott (1974: 466) followed Bayer (1943: 122) in placing C. bermudensis as a form of С. sinuosa Broderip. Pending re-ex- amination of Heilprin's type, we tentatively re- tain С. bermudensis within С. sinuosa. Chama florida Lamarck, 1819 Figures 3B, C Chama florida Lamarck, 1819: 94; Dall, 1903: 1404; Lamy, 1928: 376 (extensive syn- onymy); Pilsbry & McGinty, 1938: 74; Bayer, 1943: 117, 119, 123, pl. 12, fig. 6; Weisbord, 1964: 241-242, pl. 33, figs. 3, 4 (extensive synonymy); Abbott, 1974: 467. ?Chama sarda Reeve - Díaz & Puyana, 1994: 74, pl. 16, fig. 154; Rios, 1994: 260, pl. 89. Type Locality Recent, Dominican Republic. Description Small to medium-sized, with crowded, con- centric rows of small, fluted fronds; upper valve typically white, with a variable number of ra- dial rows of red dots, often stained internally with red or pink; inner margin crenulated; sculpture uniform or stronger with continued growth, as illustrated by Bayer (1943: pl. 12, fig. 6). A specimen from Soldier Key (FLMNH 146889) shows first-year growth with erect, recurved scales. Second-year growth remains erect, but points ventrally. Spine wear and epizoan growth suggest a three-year life span. Distribution Pleistocene, Cuba and Venezuela (Weis- bord, 1964: 242). Recent, southeastern Florida and West Indies, eastern Brazil, 9 to 80 m (Abbott, 1974: 467). Discussion Brazilian specimens were described by Rios (1994) as having white, tubular spines, and Diaz & Puyana (1994) illustrated such a speci- men from the Atlantic coast of Colombia. Rios (1994) and Diaz & Puyana (1994) placed this southern material in Chama sarda, but the Diaz & Puyana (1994) illustration most re- sembled the Bayer (1943) figure of C. florida. This Colombian material shows larger fronds with down-turned margins. Pilsbry & McGinty (1938: 74) suggested synonymizing C. sarda and C. florida. Bayer (1943: 117) argued for separate species, and noted that the pallial WESTERN ATLANTIC RECENT CHAMIDAE FIG. 3. Chama spp. А: Chama macerophylla, right and left valves, Bear Cut, Dade County, Florida (FLMNH 238239); B: C. florida, right and left valves, IMBW-FK-641, Tennessee Reef, off Long Key, Florida, 7 m (AMNH 306410); C: C. florida, right and left valves, Sand Key Light, Key West, Florida (FLMNH 146887). Scale bars = 10 mm (A); 5 mm (B-C). 398 CAMPBELL ET AL. line attaches to the bottom of the anterior ad- ductor muscle scar in C. florida, but attaches higher on the margin in C. sarda. Based on our examination of fifteen to thirty specimens of each (FLMNH), we find all С. florida speci- mens consistent with Bayer's observation, but C. sarda shows considerable variation in the position of pallial line attachment. Deshayes, in Lamarck et al. (1835: 583) noted that Chama florida Lamarck could be a variety of “C. cornuta Chemnitz” (Chemnitz, 1783: pl. 52, fig. 518). Clessin (1889) accepted this synonymy. Lynge (1909: 265) examined Chemnitz's specimen and decided that it was a young individual, worn and hard to identify, from the Nicobar Islands, India, in contrast to C. florida from the Dominican Republic. Hanley (1843) thought some о the original specimens of C. florida resembled the figure of C. spinosa Broderip, 1835. Lamy (1928) decided that the specimens were too poorly preserved for de- finitive identification, and that Chemnitz' fig- ure 518 could be C. spinosa Broderip 1835, a synonym of C. asperella Lamarck, 1819. Lamy (1928) examined five boxes of speci- mens labeled by Lamarck as Chama florida. He concluded that the specimens in the first box were too poorly preserved for definitive identification. The second and third boxes con- tained juvenile C. macerophylla Gmelin. The fourth box contained a large (45 mm), rounded discolored shell labeled as “Chama florida? old individual”. The fifth box contained five valves of С. florida that matched Reeve's il- lustrations of that species (1847). Lamy (1928) rejected the comparison by Broderip (1835) of Chama florida with C. paci- fica Broderip, 1835, and rejected the syn- onymy by Dall (1903) of C. sarda Reeve, 1847, with C. florida Lamarck, 1819. Lamy (1928) concluded that the figures in Chenu (1846) for C. florida were inadequate and instead re- sembled C. chinensis Chenu, 1846. Chama sinuosa Broderip, 1835 Figures 4-6 Chama sinuosa Broderip, 1835: 303, pl. 39, fig. 11; Dall, 1903: 1403 (in synonymy with C. macerophylla); Lamy, 1928: 311 (extensive synonymy); Pilsbry 8 McGinty, 1938: 76, pl. 7, fig. 9; Abbott, 1974: 466, text fig. 5386, pl. 21, fig. 5386; Cerridwen & Jones, 1991: 100. Chama cistula Reeve, 1847: pl. 9, fig. 51. Chama tumulosa Reeve, 1847: pl. 9, fig. 52. Chama lamarckiana Clessin, 1889: 42, pl. 5, figs. 1,2, ?Chama bermudensis Heilprin, 1890: 141, pl. 8, figs. 1, 1a. Chama sinuosa firma Pilsbry & McGinty, 1938: 76, pl. 7, fig. 1; Bayer, 1943: 121, pl. 13, fig. 11: Chama sinuosa bermudensis Heilprin — Bayer, 1943: 118, 122, 123, pl. 14, fig. 26; ?Weis- bord, 1964: 242-243, pl. 32, figs. 10, 13. Type Locality Recent, Brazil (Broderip, 1835). Description Medium to large, thin to heavy, with large, fluted, smooth (or rarely striate) fronds in typi- cal expression; unattached valve with central, radial, depressed band, the edges of which are deep, narrow grooves; grooves visible even on surf-worn valves devoid of all other sculpture; no internal margin crenulations present. А 57 mm specimen (FLMNH 15877), labeled C. sinuosa firma, shows two years growth with surface sculpture removed by a variety of shell- degrading endolithic species (Fig. 5). The on- set of third year growth provides a fringe of fresh, undamaged fronds and lamellae, the pattern that defines the variety. Distribution Early Pleistocene, Venezuela (Weisbord, 1964). Late Pleistocene, Grand Cayman (Cerridwen & Jones, 1991); Cockburn Town, San Salvador, Bahamas (Hagey, 1991). Re- cent reef species, southern half of Florida, West Indies, Bermuda, and Brazil (Abbott, 1974: 466). Discussion The variety Chama sinuosa firma Pilsbry & McGinty, 1938, has reduced sculpture except for some large fronds at the margin, is larger, and more deeply cupped (Fig. 5). That trend, taken to an extreme, produces C. s. bermudensis Heilprin, 1890, in which the shell is very large, thin to very thick; the ventral valve very deep, and the beak high and strongly coiled (Fig. 6). The latter morphology paral- lels the Pliocene C. heilprini. The Venezuelan fossils referred to C. s. bermudensis by Weisbord (1964) more closely resemble С. $. firma. Waller (1973: 41, 48) found no speci- mens of C. sinuosa in his survey of Bermu- WESTERN ATLANTIC RECENT CHAMIDAE 399 FIG. 4. Chama sinuosa. A: Right and left valves, IMBW-FK-641, Tennessee Reef, off Long Key, Florida, 7 m (AMNH 306936); B: Right and left valves, IMBW-FK-624, Horseshoe Reef, off Fat Deer Key, Florida, 7.3 m (AMNH 306938); C: Right and left valves, off Yamoto, Palm Beach, Florida, 73 m (FLMNH 15866). Scale bars = 5 mm (A-B); 10 mm (C). FIG. 5. Chama sinuosa firma, in moat, Fort Jefferson, Dry Tortugas, Florida (FLMNH 15877). Scale bar = 10 mm. A: Right valve; B: Left valve. 400 CAMPBELL ET AL. FIG. 6. “Chama sinuosa bermudensis” (Bayer 1943, label), in moat, Fort Jefferson, Dry Tortugas, Florida (FLMNH 15882). Scale bar = 10 mm. A: Right valve; B: Left valve. WESTERN ATLANTIC RECENT CHAMIDAE 401 dian molluscs, but did find heavy specimens of C. macerophylla that strongly resembled Heilprin's taxon. Waller's specimens of С. macerophylla had crenulated margins. Neither Heilprin's text nor figures expressed the inter- nal margin crenulations characteristic of C. macerophylla. The figure showed chipping of the shell margin that can remove crenulations in large Chama specimens. Heilprin (1890: 141) had a population of C. bermudensis “dredged in large quantities in Harrington Sound”. We tentatively retain C. bermudensis within С. sinuosa pending re-examination of the type material. Tryon (1872) synonymized the dextral Chama cistula Reeve with the sinistral spe- cies C. appressa Reeve. However, based on Reeves (1847) descriptions and figures, we synonymize С. cistula with С. зтиоза, and С. appressa with С. radians Lamarck. Lamy (1928: 311) noted that Chama lamarckiana Clessin, 1889 (p. 42, pl. 5, figs. 1, 2), from St. Thomas, Virgin Islands, was so badly eroded that it was difficult to identify. Benthem Jutting (1927) considered C. lamarckiana Clessin to be a variety of C. sinuosa Broderip, 1835. She also treated C. bermudensis Heilprin, 1890 as a synonym of С. зтиоза Broderip, 1835. Lamy (1921, 1928) reported that Jousseaume had identified a Chama from Djibouti, Indian Ocean, as С. sinuosa Broderip. However, Lamy thought this specimen matched C. preetexta Reeve, 1847, figured by Reeve (1847) and Clessin (1889) from the Indian Ocean. Chama radians Lamarck, 1819 Figures 7, 8 Chama radians Lamarck, 1819: 96, also ref- erencing Chemnitz, 1786: 145, pl. 116, fig. 992. Chama ferruginea Reeve, 1846, pl. 4, sp. 21; Dall, 1903: 1404. Chama variegata Reeve, 1847a: 118; 1847b, pl. 9, fig. 50; Dall, 1903: 1404. Chama appressa Reeve, 1847b: pl. 9, fig. 55. Chama ruderalis Lamarck — Guppy, 1877: 86, non Lamarck, 1819: 96. Pseudochama ferruginea Reeve — Odhner, 1919: 39. Pseudochama radians (Lamarck) — Lamy, 1928: 375 (extensive synonymy); Pilsbry & McGinty, 1938: 77; Weisbord, 1964: 243- 246, pl. 33, figs. 5, 6, pl. 34, figs. 1-6; Waller, 1973: 41, 48; Abbott, 1974: 467, pl. 21, fig. 5395; Cerridwen & Jones, 1991: 100; Donovan & Littlewood, 1993: 37; Redfern, 2001: 215, pl. 90, figs. 880A-D. Pseudochama radians variegata Reeve — Pilsbry & McGinty, 1938: 77, pl. 7, figs. 3-5; Bayer, 1943: 122, pl. 12, fig. 4. Type Locality Recent, uncertain; possibly St. Croix, West Indies (fide Lamy, 1928: 376, see below). Description Medium to large, thin to heavy, attached along the anterior edge of right valve, com- monly from umbo to ventral margin (Abbott, 1974, pl. 21, fig. 5395); left, unattached valve obliquely oval in specimens with an extended attachment surface, more circular in speci- mens with a small attachment area and greater free growth; sculpture of rough, closely packed concentric lamellae often broken into low fronds or scales; red-brown to purple stain varying in extent on the inside of attached valve; interior margin crenulated in most speci- mens, but five of the forty specimens exam- ined lacked crenulations. Distribution Early Pleistocene, Bermont beds, southern Florida, and the Abisinia Formation, Venezu- ela. Late Pleistocene, Jamaica (Donovan & Littlewood, 1993), and Grand Cayman (Cerridwen & Jones, 1991). Recent, North Carolina to Florida, Texas, West Indies, Ber- muda, and to Brazil (Abbott, 1974: 467), Loui- siana record from Garcia & Lee (2002). Indian Ocean references (Clessin, 1889; Lamy, 1928) unlikely. Discussion The type locality is unknown. Yves Finet (e- mail, 3 March 2003) examined the labels with the holotype of Chama radians Lamarck, 1819 (Fig. 7A; MNHG 1087/3). The original label had no locality data; a more recent label said “Oc. indien?”. Lamy (1928: 376) cited Chemnitz (1786) as indicating West Indies (St. Croix) as the locality for his plate 116, figure 992. Unpigmented white shells of Chama radians are known from Eleuthra, Bahamas. A lot of six live-taken pairs of C. radians (FLMNH 127267; Grassy Key, Florida) possess a bi- zarre and confusing morphology, in which lat- eral expansion of the shell was halted at a 402 CAMPBELL ET AL. length of 25 mm and height of 30 mm (Fig. 8C), with subsequent growth by shell thicken- ing and body cavity inflation. The largest pair spans 37 mm width across both valves and 1$ superficially convergent with certain rudists. Surface detail of the free valve is chemically leached, possibly from exposure to fresh wa- ter at low tide. Pilsbry & McGinty (1938: 77-78) restricted Chama radians variegata (Reeve) to Florida (Fig. 8A), and suggested that C. r. radians and C. r. ferruginea are more typically West Indian taxa. However, Honduras was Reeve's (1847) type locality for C. variegata. Most current authors consider these to represent a single, variable species. Molecular sequencing could test the validity and affinities of these forms. Lamy (1928: 312) said that Chama ferruginea Reeve was similar to C. macerophylla Gmelin. Lamy cited Tryon (1872) as synonymizing C. ferruginea with С. sinuosa Broderip, and Lamy said that Tryon cited a Broderip figure that did not exist. Clessin (1889: 48, pl. 19, figs. 1-3) described C. rotunda as a sinistral species from Veracruz, Mexico. Lamy said that the Clessin figures were insufficient and showed a worn specimen, but that C. rotunda was probably a synonym of C. ferruginea Reeve. Chama appressa Reeve, 1847, represented a large specimen of C. radians with a small attachment surface, freeing it for more circu- lar growth. Tryon (1872) combined the dextral species C. cistula Reeve, 1847, with the sin- istral species C. appressa Reeve, 1847. Here FIG. 8. Chama radians. A: Right and left valves, South of Boyton Inlet, Palm Beach, Florida (FLMNH 135589); В: Right and left valves, Marco, Collier County, Florida (FLMNH 199101); С: Right and left ee ce Key (Gulf side), Monroe County, Florida (FLMNH 127267). Scale bars = 10 mm (A- В); 5 mm : WESTERN ATLANTIC RECENT CHAMIDAE 403 we synonymize C. appressa Reeve, 1847, with C. radians Lamarck, 1819, and C. cistula Reeve, 1847, with C. sinuosa Broderip, 1835. Clessin (1889) and Lamy (1928: 376) identi- fied specimens from the Indian Ocean and Aden, Red Sea as Chama radians Lamarck. Lamy (1928: 376) noted that this was in contrast to the West Indies distribution cited in Chemnitz (1786). Chama sinistrorsa Bruguière, 1792, may be an older name for С. radians. Deshayes, in Lamarck et al. (1835: 587) synonymized C. sinistrorsa Brocchi, 1814, non Вгидшеге, 1792, with C. gryphina Lamarck, 1819 (see Discus- sion under Pseudochama cristella). Chama lactuca Dall, 1886 Figure 9A, B Chama lactuca Dall, 1886: 268, 1903: 1404; Pilsbry & McGinty, 1938: 74; Bayer, 1943: 120—121, pl.12, figs. 1, 2; Abbott, 1974: 467, text fig. 5391. Type Locality Recent, Barbados (Dall, 1886). Description Small, white, sometimes with brown bands; attached valve deeply cupped, “naticiform”, sculptured with concentric lamellae; lamellae most often entire, but in some specimens cut into broad fronds; upper valve with radiating and concentric, small, short, spine-like fronds; lamellae of attached valve and fronds of free valve with radial grooving underneath. Distribution Recent, North Carolina to Florida, Gulf of Mexico, and to Barbados; 25-200 m or more. Louisiana record from Garcia & Lee (2002). Chama sarda Reeve, 1847 Figures 9C—F, 13A Three different recent forms appear under this name in current literature. Apart from some red pigmentation and small size, they have little in common. Chama sarda Reeve, 1847 (attached valve shallow, stained deep red) Chama sarda Reeve, 1847b: pl. 7, fig. 40; Dall, 1903: 1404; Lamy, 1928: 379 (extensive syn- onymy); Pilsbry & McGinty, 1938: 74; Bayer, 1943: 119, pl. 14, figs. 15, 16; Waller, 1973: 41, 48; Abbott, 1974: 466, pl. 21, fig. 5387. Chama sarda lutea Lamy, 1928: 380. Type Locality Recent, Honduras (Reeve, 1847b). Description “Shell somewhat orbicular, both valves pe- culiarly faintly obliquely striated, with sharp remote short scales; bright coral-red within and without. Hab.: Honduras (attached to coral). Rich in colour and very characteristic in sculp- ture, being crossed in an oblique direction throughout with faint striae, and roughened here and there with short scales like the asparaties of a coarse file” (Reeve, 1847b). Reeve described a rather variable, often misshapen small Chama with the attached valve shallow and extensively secured to the substratum. The attached valve is partially or completely stained bright, deep red. The right, unattached valve is commonly shades of red, red-pink, or cream, with low concentric corru- gations, and in some cases, white, flat fronds near the edge of adult growth. Distribution Recent, Bermuda, northern Florida (H. Lee, personal communication, July 2003) to Florida Keys and from Bahamas at least to Honduras and the Dominican Republic. Discussion This is the form described by Reeve, and clearly illustrated by Bayer (1943), Abbott (1974), and Redfern (2001). Lamy (1928) re- jected the synonymy by Dall (1903) of Chama sarda Reeve, 1847, with C. florida Lamarck, 1819. Lamy (1928) noted that previous reports were from the Antilles, and then reported 11 specimens from Île du Prince, West Africa, with a yellow specimen from there indicated as “variété jaune”. Two lots from the Florida Museum of Natu- ral History — ten pairs from Key Largo (FLMNH 135580) and six pairs from off Palm Beach (FLMNH 127241) - clarify and extend the range of morphologies possible in this spe- cies. The Key Largo specimens show first- and second-year growth reaching about 20 mm (Fig. 9D). The paired shells are wedge- CAMPBELL ЕТ AL. FIG. 9. Chama spp. A: Chama lactuca, right and left valves, off Dodge Estates, Palm Beach, Florida, 45 m (FLMNH 135481); B: C. lactuca, left valve, off Radio tower to Breakers Hotel, Palm Beach County, Florida, 137 m (FLMNH 186363); C: C. sarda, four pairs, right and left valves, Key Largo, Florida, 11 m (FLMNH 135580); D: C. sarda, right and left valves, IMBW-FK-629, “The Horseshoe” site, West Summerland Key (Spanish Harbor Keys), Florida (AMNH 306411); E: C. sarda, right and left valves, Boynton Beach, Palm Beach County, Florida, on beached gorgonian after storm (FLMNH 135582); Е: С. sarda, right and left valves, Key West, Florida, SW Channel (FLMNH 246260). Scale bars = 5 mm (А, В, 0); 10 mm (С, E, Е). WESTERN ATLANTIC RECENT CHAMIDAE 405 shaped. Color ranges from uniform red or yel- low, to red-and-white, to nearly white shells. Clusters of long, typically white spines may flank the posterior radial groove on both valves, with shorter spines covering the re- maining shell. The Palm Beach lot exhibits more standard chamid morphologies. The larg- est specimen reaches a maximum elevation of 38 mm, but most specimens are less than 25 mm. The radial lines mentioned by Reeve, 1847, are weakly developed on some of the attached valves, but are lacking on the free valves. The irregular form and limited spine development (Reeve, 1847) seem most typi- cal of first year growth, but sometimes persist with continued growth. More typically, spine development becomes more frequent and regular with second-year growth. Kennedy et al. (1970: 395, text fig. 6) illus- trated the chamid microstructure, noting that myostracal pillars may extend from the pallial myostracum through the inner shell layer and create bosses or bumps on the inner surface of the shell. These bosses are common to a number of species, including Chama radians and C. macerophylla, but we find them to be particularly large and frequent in young C. sarda. “Chama sarda Reeve, 1847” (attached valve deeply cupped with ventral sulcus) Figure 13A Chama sarda Reeve, 1847 - Abbott, 1974: 466 (in part, deep cupped morphologies); Rehder, 1981: 729, fig. 654. Description Small, with the attached valve deeply cupped, and with a square, box-like outline due to a strong ventral sulcus, so that in outline it resembles the fossil species Chama emmonsi Nicol, 1953; left valve with a small attachment surface, allowing for much free growth; shell uniformly cream colored, with none of the red blotches typical of C. sarda, and with radial rows of tightly appressed, narrow, red fronds that may become erect at the margin of adult growth; upper valve cream color, with finer and more numerous radial rows of red rectangular appressed fronds, some of which elevate into low scales; upper valve with two radial furrows, a wider central one that merges with the ven- tral sulcus, and a smaller, narrower posterior furrow; myostracal pillars fused into linear ra- dial ridges bordering inner edge of pallial line. Distribution Bahia, Brazil, and Caribbean, precise locali- ties uncertain. Discussion The smaller, but uniform size, consistent quadrate outline with ventral sulcus, appressed sculpture, and lack of red blotching separate this form from typical Chama sarda. Typical C. sarda has myostracal pillars that form elevated oval bosses on the shell interior, not the fused, linear patterns found in this form. Rehder (1981) illustrated the diagnostic outer surface of the attached valve of this form. Olsson & Harbison (1953: 76) reported a Recent Chama (ANSP 84720) from Bahia, Bra- zil, resembling their new species C. gardnerae (= C. emmonsi Nicol, 1953). Paul Callomon located this lot and provided illustrations (Fig. 13A; ANSP 84720). The label reads: “C. lingua- felis Rve.” crossed out and replaced with “C. florida Lam. (ballast) Bahia, Brazil. J. G. Malone, 1903”. The two pairs are conspecific with our lot of 16 specimens (USCS collections) of uncertain Caribbean locality and with Rehder’s field guide specimen of unstated provenance. Based on the material in hand, this could be a previously unrecognized, distinct species with a distribution from Florida to Brazil. A search of museum collections will be neces- sary to determine its distribution and full range of variation. A review, especially of the 19th century literature, will be necessary to ascer- tain if it is undescribed. “?Chama sarda Reeve” (tubular spine form) Chama sarda Reeve, 1847; Diaz & Puyana, 1994: 74, pl. 16, fig. 154; Rios, 1994: 260, pl. 89. Discussion The taxon identified as Chama sarda by Rios (1994: 260) and by Diaz & Puyana (1994: 75) is probably C. florida, as discussed above under that species. Chama inezae (Bayer, 1943) Figure 10 Pseudochama inezae Bayer, 1943: 22, pl. 15, figs. 1-4; Abbott, 1974: 467, species 5396; Mikkelsen & Bieler, 2000: 372. 406 CAMPBELL ET AL. Type Locality Recent, Carysfort Reef, Key Largo, Florida Keys, Florida (Bayer, 1943). Description Large, thin, with very thin, widely spaced, concentric, erect, ruffled lamellae; sculpture on both valves of broad, concentric draper- ies, or of lamellae becoming frondose at outer margins, or of very wide, fused fronds; broad, flaring lamellae most spectacular on smaller, apparently second-year, fresh-growth indi- viduals; full-grown, third year specimens typi- cally with wide lamellae trimmed by bioerosion; radial lines and threads some- times present on the lamellae, but are of var- ied and limited extent; shell alabaster-white, lacking interior margin crenulations; early prodissoconch circular, about one millimeter in diameter, punctate, a pattern more typical of the genus Chama. A population from Palm Beach, Florida of 16 specimens has eight right-attached and eight left-attached individuals. FIG. 10. Chama inezae. A: Right and left valves (FLMNH 135544); В: Right attached and left free valves, off Palm Beach, Florida, 20 m, under coral ledges (AMNH 308078); C: Right free and left attached valves, off Palm Beach, Florida, 20 m, under coral ledges (AMNH 308077). Scale bars = 5 mm (A); 10 mm (В-С). WESTERN ATLANTIC RECENT CHAMIDAE 407 FIG. 11. Holotype, Pseudochama cristella, (MNHG 1087/6, photographs courtesy of Yves Finet). Lamarck's original handwritten label stated “Océan des grandes indes”. A more recent label indicated “Ocean indien” (= Indian Ocean). Scale bar = 10 mm. A: Right valve; B: Left valve. FIG. 12. Pseudochama sp. cf. P. cristella. A-C: Three pairs, right and left valves, Java (ANSP 54808, photographs courtesy of Paul Callomon). Scale bars = 10 mm. 408 CAMPBELL ET AL. Distribution Recent, southeastern Florida (Abbott, 1974); Palm Beach County to Florida Keys. Discussion Chama inezae was originally based on a right-attached, unique holotype, and remains a rare, endemic species of mid-depth reefs, hard grounds, and wrecks off Florida. Bayer assigned his species to Pseudochama be- cause of the right attachment. However, the specimen of С. inezae in the Florida Museum of Natural History (Fig. 10A; FLMNH 135544) proved to be left-attached. Two contrasting specimens are insufficient to define the nor- mal attachment pattern, and we began a search for additional specimens of this rare species. We learned that Tom Honker had dis- covered a small population in 20 m under dead coral ledges off Palm Beach, Florida. Our 16 specimens (AMNH 308077-308078; USCS collections) were collected by Tom Honker in May 2002, and showed a 50:50 ratio of right valve and left valve attachment (Figs. 10B, С). This 1$ the first known example of this attach- ment pattern for a Recent species from the western Atlantic. Yonge (1967) reported 50:50 ratios in the Indo-Pacific Chama ruderalis Lamarck, 1819, in Australia. Fossil species preserving similar ratios of right and left at- tached valves are known from Mississippi and France. Prevaling interpretation in literature places such specimens in the genus Chama. Left-attached specimens of С. inezae express the typical Chama hinge pattern, having the same arrangement of teeth and sockets as found in C. sarda and C. macerophylla. Right- attached specimens show that transposition of the hinge is linked to transposition of the shell in this species. Examination of the hinge in a right-attached valve demonstrates that the teeth and sockets are mirror images of those in a left-attached valve, but are less vigorously developed. The presence of both right and left valve at- tachment patterns complicates identification. Any small to large, all white, Recent Floridian right-attached chamid lacking marginal crenu- lations may be referred to Chama inezae. Among left-attached species, С. inezae is strik- ingly alabaster-white, has broad, widely spaced lamellae, and has concentric sculpture domi- nant on both valves. Most of the other “white” Chama species in the region are cream, multi- colored, or dirty white. The attached valve of C. lactuca has similar concentric lamellae, but the lamellae are proportionally shorter, and the valve is more deeply cupped and 1$ internally crenulate. Chama lactuca is also much smaller than С. inezae. Chama macerophylla may be alabaster-white, but the crowded, frondose sculpture sets it apart from С. inezae. Genus Arcinella Schumacher, 1817 Type species by tautonomy: Chama arcinella Linnaeus, 1767. Not preoccupied by Arcinella Oken, 1815, re- jected by ICZN Opinion 417 (1956). Synonym: Echinochama P. Fischer, 1887. “Shell attached to the substrate only in the early stages of growth. Lunule prominent, bor- dered by an incised line. Sculpture of radial rows of thin spines” (Abbott, 1974: 467). All Recent species, including Arcinella cornuta, A. arcinella from the central Caribbean to south- ern Brazil, and A. brasiliana (Nicol, 1953), from southeastern Brazil, attach by the right valve in early growth stages, are pitted between rows of spines, and are nearly equivalve (Nicol, 1953). In contrast, fossil species of Arcinella have a wider range of morphologies and sculp- tures and may lack a lunule, may lack spines, and/or may be strikingly inequivalve. J. Gibson- Smith 8 W. Gibson-Smith (1979) provided a review of the western Atlantic species. Arcinella cornuta Conrad, 1866 Figure 13B Arcinella cornuta Conrad, 1866: 105; Abbott, 1974: 467, pl. 21, fig. 5400; J. Gibson-Smith & W. Gibson-Smith, 1979: 18, pl. 3, figs. 13-15. Chama arcinella Linnaeus - Tuomey & Holmes, 1856: 22-23, pl. 7, figs. 4-6; Mansfield, 1916, pl. 113, figs. 11, 12. Echinochama arcinella (Linnaeus, 1767) — Dall, 1903: 1405-1406; Mansfield, 1932: 92— 93: pl. 18, figs. 1, 4. Pseudochama (Echinochama) arcinella (Linnaeus) — Gardner, 1926: 94-95, pl. 17, figs. 14-16. Echinochama arcinella cornuta (Conrad) — Pilsbry & McGinty, 1938: 78-79, pl. 7, fig. 7. Echinochama cornuta (Conrad) — Nicol, 1952c: 809-810, pl. 118, fig. 2, pl. 119, fig. 7. Type Locality Pliocene, Royal Landing, Waccamaw River, South Carolina (Tuomey 8 Holmes, 1856). WESTERN ATLANTIC RECENT CHAMIDAE Description Large, quadrate, equivalve, with a pro- nounced lunule, and typically with 7-9 radial rows of large spines. Distribution Middle Miocene, Shoal River Formation, western Florida. Lower Pliocene, Goose Creek Formation and Raysor Marl, South Carolina: Ecphora zone, Jackson Bluff Formation, west- ern Florida; Tamiami Limestone, southern Florida. Middle Pliocene, Duplin Formation, North and South Carolina; Cancellaria zone, Jackson Bluff Formation, western Florida; Pinecrest beds, southern Florida. Upper Pliocene, Waccamaw Formation, North and South Carolina; Nashua Formation, north- eastern Florida; Caloosahatchee Formation, southern Florida. Recent, Cape Hatteras to Florida and Texas to northern Yucatan, Mexico. Discussion Arcinella cornuta differs principally from the more southern А. arcinella in having fewer spine rows across the posterior half of the shell. Arcinella cornuta from shallow, high energy environments have stubby spines; specimens from deep, quieter waters may develop strikingly long, thin, recurved spines. The species is remarkably long-lived, ranging from the Middle Miocene to Recent. No specimens were recovered during the 409 FIG. 13. Chamidae. A: Chama sarda deeply cupped form, right and left valves, ballast, Bahia, Brazil (ANSP 84720, photographs courtesy of Paul Callomon); B: Arcinella cornuta, right and left valves, Bahia Honda Key, Florida, collected live on ocean side (AMNH 209917). Scale bars = 10 mm (A-B). 410 CAMPBELL ET AL. Bivalve Workshop, but it is well documented from the Florida Keys (Mikkelsen & Bieler, 2000). The J. Gibson-Smith & W. Gibson- Smith (1979: 18) reference to А. (Nicolia) cornuta in their figure caption to Plate 3 was a lapse with no bearing on the affinity of the spe- cies. Nicol (1952c) found one left-attached shell among over 1,000 normal specimens. Dubious or erroneous reports of Chama and Pseudochama species from the western Atlantic Ocean Chama lobata Broderip, 1835, is a large, trapezoidal species based on syntypes he la- beled “Nevis Island, Leeward Island, West Indies”. Tryon (1872) repeated this locality in- formation. Reeve (1847b) figured a specimen of this species and stated; “Mr. Broderip has recorded the ‘Island of Nevis, West Indies’ as the habitat of this very characteristic species; this must surely be an error for there are sev- eral well-authenticated specimens in the Brit- ish Museum, brought from China by John Reeves, Esq. and | cannot learn that it has been received from any other locality.” Lamy (1928) and Matsukuma (1996: 34, 45, figs. 16a-b) noted the original locality data and confirmed Reeve's correction. Chama lazarus Linnaeus was listed as be- ing from the Carolinas by Kurtz (1860). This record 1$ typical of a common misidentification in nineteenth century literature. Clessin (1889: 30) reported Chama rubea Reeve, 1847, from the Gulf of Mexico. Lamy (1928: 318-319) decided that Clessin's figures included not only true C. rubea Reeve from the Philippines but also a specimen of C. producta Broderip, 1835. Lamy noted that C. producta Broderip was a Pacific species, in contrast to the locality given by Clessin (1889). Clessin (1889: 38, pl. 16, figs. 3, 4) reported Chama cristella Lamarck, 1819, from Puerto Rico. Lynge (1909) said that the Clessin Puerto Rico specimen was not C. cristella. Vanatta (1914) reported Chama lingua-felis Reeve, 1847, from a Pleistocene deposit near Sierra Nueva, Dominican Republic. This record 1$ not С. lingua-felis Reeve, described from the Recent of Guimaras, Philippines. Peile (1926: 96) said that C. lingua-felis as recorded by Rice (1884) from Bermuda was probably a juvenile C. macerophylla. Chama coralliophaga in the index for Maury (1917) referred to Coralliophaga coralliophaga (Gmelin, 1791). Chama crassa Chenu, 1846 (pl. 5, fig. 3) is a Recent taxon with no locality data. (Chenu figured Recent taxa, including C. crassa in color, and fossil taxa in black and white — Gary Rosenberg, e-mail, 25 March 2003.) The list- ing of Chama crassa Chenu as a Florida fos- sil in Kennedy et al. (1970: 391) undoubtedly is a lapse for С. crassa Heilprin, 1886, and both are junior homonyms of C. crassa Smith, 1817. Chama crassa Heilprin, 1886, was re- named С. heilprini by Nicol (1953). DISCUSSION Although the present molecular results are limited, they allow drawing some preliminary conclusions. Both the nuclear ITS sequences and the mitochondrial ribosomal sequences support the same basic topology. All trees show four distinct sequence clusters, the Chama species from the Seychelles, the C. macerophylla group, C. florida, and the C. congregata-C. sarda-C. inezae-C. radians group. Chama congregata is polyphyletic and also shows different alleles of ITS elements in one individual. The difference between these two alleles is greater than that between C. sarda and C. congregata 624 1b indicating that the origin of the allelic difference predates the speciation. The one sequence from C. radi- ans clusters with one of the C. congregata sequences and they form the sister group to C. inezae. This suggests that the genus Pseudochama as defined by attachment pat- tern is polyphyletic, but additional sequence data are needed to confirm this tentative re- sult. The presence of different alleles of ITS ele- ments in a single specimen was reported for several species of the vetigastropod genus Haliotis by Coleman & Vacquier (2002). Our report of allelic differences in Chama congregata is the first demonstrated case for the Bivalvia, but not unique for the Mollusca. Gene trees and species trees may not be iden- tical due to incomplete lineage sorting, which will complicate phylogenetic reconstructions. The significance and impact on phylogenetic reconstructions remain to be determined by a broad assessment of this feature in chamids. Bayer (1943) provided the most recent re- view of Recent Chama and Pseudochama in the western Atlantic Ocean, and he did not include fossil taxa. Dall (1903) was the last to review fossil species. Subsequent authors WESTERN ATLANTIC RECENT CHAMIDAE 411 have doubled the number of nominal fossil species. We intend to publish a review of the fossil taxa in a separate paper. Including Mi- ocene to Recent taxa, there are eighteen spe- cies of Chama and nine species of “Pseudochama” in this region (Tables 3-6). Ten species of Chama are known only from the fossil record, with С. chipolana Dall, 1903; C. strepta Woodring, 1982; C. berjadinensis F. Hodson, 1927; C. paschuali Brunet, 1986; C. caimitica Maury, 1917; C. involuta Guppy, 1873; С. heilprini Nicol, 1953; С. emmonsi Nicol, 1953; and С. willcoxii Dall, 1900, from the Neogene and C. iudicai Pastorino, 1991, from the Pleistocene. Additionally, С. emmonsi has one doubtful record from the Recent. Five species, C. congregata, C. macerophylla, C. florida, С. sinuosa, and С. radians are known from both the fossil record and the Recent. Three species, С. lactuca, С. sarda, and С. inezae are known only from the Recent. Nine species of Pseudochama, P. draconis (Dall, 1903); Р buchivacoana (Е. Hodson, 1927); P. corticosaformis (Weisbord, 1929); P. lazai Brunet, 1986; Р. quirosana (Е. Hodson, 1927); P. scheibei (Anderson, 1929); P. riocanica (Maury, 1917); Р. corticosa (Conrad, 1833); and P. caloosana (Dall, 1903) are known only from the Neogene fossil record. In the genus Pseudochama, many species do not have obvious phylogenetic affinities with other fossil or Recent taxa from the region. Pseudochama corticosa and P. caloosana do not appear to be closely related to C. radians. Lineages are lacking. The taxa described from the Neogene of Panama, Venezuela, Argentina, Colombia, and the Dominican Republic (Chama strepta, C. berjadinensis, C. paschuali, C. caimitica, Pseudochama buchivacoana, P. corticosa- formis, P. lazai, Р quirosana, Р scheibei, and P. riocanica), are in need of further study, more detailed morphological descriptions, and more precise stratigraphic data. CONCLUSIONS The species Chama congregata, C. macerophylla, C. florida, C. sinuosa, C. radi- ans, C. sarda, and C. inezae were collected during the International Marine Bivalve Work- shop held in the Florida Keys, U.S.A., 19-30 July 2002. Mikkelsen & Bieler (2000) also re- ported С. lactuca and Arcinella cornuta from the Florida Keys. The genus-level characters cited by Matsukuma (1996), particularly soft anatomy, prodissoconch and hinge structure, need fur- ther testing among taxa in this region. Prodissoconch characters overlap to some degree in western Atlantic species historically assigned to Pseudochama and Chama. Based on the small prodissoconch and on equal num- bers of right-attached and left-attached speci- mens, Р inezae Bayer, 1943, is reassigned to the genus Chama. Chama inezae (Bayer) is the first Recent species reported from the western Atlantic Ocean with a roughly 50:50 ratio of right-attached to left-attached speci- mens. Chama radians Lamarck is returned to the genus Chama based on morphologic, hinge, and ITS and 16S rRNA sequence data. Chama congregata appears to include mul- tiple species based on morphological and molecular data. One specimen of C. congregata had two alleles of ITS elements, the first demonstrated case for the Bivalvia of ITS allelic differences within an individual. Molecular phylogenetic reconstructions con- taining C. congregata will be complicated be- cause incomplete lineage sorting could cause differences between gene trees and species trees. Chama sarda also appears to include multiple species based on morphological data. Additional morphological and molecular study is needed for populations of both species. Western Atlantic chamids are taxonomically challenging. Our study underscores the neces- sity of reviewing both the fossil and the Re- cent literature, and the necessity of synthesizing morphological and molecular data. Several taxa require further population analy- ses across a broad range of distribution in space and time. These include the Recent taxa Chama congregata, C. sarda, C. radians, and the more obscure forms described by Reeve (1846-1847). Such studies should clarify whether odd morphologies, named varieties and subspecies, and various synonyms rep- resent variation along a continuum or stand apart as separate species. ACKNOWLEDGEMENTS The International Marine Bivalve Workshop, held in the Florida Keys, U.S.A., 19-30 July 2002, was funded by U.S. National Science Foundation Award DEB-9978119 (to co-orga- nizers R. Bieler and P. M. Mikkelsen) as part 412 CAMPBELL ET AL. of the Partnerships in Enhancing Expertise in Taxonomy (PEET) Program. Additional sup- port was provided by the Bertha LeBus Chari- table Trust, the Comer Science and Education Foundation, the Field Museum of Natural His- tory, and the American Museum of Natural History. We thank all workshop participants for their camaraderie. Richard Petit and David Campbell provided copies of pertinent literature and helpful discus- sion. Paul Valentich Scott and Guido Pastorino supplied copies of pertinent literature. Gary Rosenberg (The Academy of Natural Sciences, Philadelphia) provided information on Chenu (1846). Sarah Campbell, David Campbell, and Andrew Campbell edited the manuscript. An- drew Campbell assisted with photography. Roger Portell, Gustav Paulay, and Tina Bell of the Florida Museum of Natural History provided important supplemental collections to the study. Thomas R. Waller (National Museum of Natu- ral History) allowed examination of his Bermu- dian chamids. Yves Finet (Museum of Natural History of Geneva) photographed the holotypes of Chama radians Lamarck and Pseudochama cristella (Lamarck). Paul Callomon (The Acad- emy of Natural Sciences, Philadelphia) photo- graphed Pseudochama sp. cf. P. cristella and C. sarda deeply cupped form from the ANSP collection. Tom Honker provided specimens and data for the population of С. inezae. We thank Eugene Coan and two anonymous re- viewers for their suggestions. LITERATURE CITED ABBOTT, В. T., 1974, American seashells, 2" ed. Van Nostrand Reinhold Co., New York. ALLEN, J. A., 1977, On the biology and func- tional morphology of Chama gryphoides Linné (Bivalvia: Chamidae). Vie et Milieu, (A)26(2): 243-260. 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Carnegie Institution of Washing- ton, Publication 366: v + 222 pp., 28 pls. YONGE, С. M., 1967, Form, habitat, and evolu- tion in the Chamidae (Bivalvia) with reference to conditions in the rudists (Hippuritacea). Philosophical Transactions of the Royal Soci- ety of London, (B)775 (252): 49-105. Revised ms. accepted 31 October 2003 MALACOLOGIA, 2004, 46(2): 417—426 SIZE AT FIRST MATURITY, OOCYTE ENVELOPES AND EXTERNAL MORPHOLOGY OF SPERM IN THREE SPECIES OF LUCINIDAE (MOLLUSCA: BIVALVIA) FROM FLORIDA KEYS, U.S.A. Gregorio Bigatti’, Melita Peharda? & John Taylor? ABSTRACT Gonads in the tropical lucinid bivalves Ctena orbiculata, Codakia orbicularis, and Lucina pensylvanica are located in the cephalopodial mass above the foot and behind the gills. Age estimates for Lucina pensylvanica suggest that individuals less than one year old are immature, as also is C. orbicularis, and that both males and females mature at two years. For Ctena orbiculata, we have no age data, but we consider that this species may also have early development. In July 2002 when water temperatures were high, mature indi- viduals of all three species were spawning or in resorption, and oocyte envelopes were present. Parasites were found in the digestive diverticula of C. orbiculata and L. pensylvanica. External ultrastructure of the spermatozoon shows differences between the three species. Codakia orbicularis and L. pensylvanica have a long-headed spermatozoa, whereas Ctena orbiculata has a middle-size head. Codakia orbicularis possesses a flagel- lum with a lateral undulating membrane, absent from the two other species. Key words: Lucinidae, reproduction, oocytes, sperm. INTRODUCTION The Lucinidae is the most diverse and geo- graphically widespread of the bivalve families . possessing chemoautotrophic sulphide- oxidising bacteria (Distel, 1998). Species live in a wide variety of marine habitats ranging from intertidal mangrove muds to hydrother- mal vents, and the family has a long and rich fossil history (Taylor & Glover, 2000). Many tropical lucinids live in close association with shallow water seagrass beds and are particu- larly diverse in the Florida Keys area, where 21 species have been recorded from all depths and habitats (Mikkelsen & Bieler, 2000). Of these, ten species were found living in the shallow water habitats of the Middle Keys sampled during the International Marine Bi- valve Workshop (IMBW) in 2002 (Mikkelsen & Bieler, 2004). Three species, Ctena orbiculata (Montagu, 1803), Codakia orbicularis (Linnaeus, 1758), and Lucina pensylvanica (Linnaeus, 1758) were sufficiently abundant and present in a range of size classes to at- tempt an investigation of some aspects of their reproductive biology. The objectives of this study were to determine the size of maturity, the presence of oocyte envelopes and to de- scribe the external morphology of their sperm. Additionally, the maturation sizes of L. pensylvanica were related to the age of the individual as estimated from acetate peel rep- licas of sections through the shell. Previous research on Lucinidae has prima- rily focused on aspects of functional anatomy, chemosymbiosis and evolution, and there are few studies of their reproductive biology. The exception is Codakia orbicularis, one of the most studied species. General aspects of the reproductive biology of this large, edible clam have previously been studied by Alatalo et al. (1984), Berg & Alatalo (1984), and Prieto et al. (1999), larval development by Gros et al. (1997), and sperm morphology by Mouéza & Frenkiel (1995). The potential role of chemo- synthesis in molluscan mariculture has also been investigated for this species (Berg & Alatalo, 1984). Many invertebrates possess oocyte enve- lopes with different layers. An inner protein- polysaccharide layer (vitelline envelope) can be formed by the Golgi apparatus of the оо- ‘CONICET - Facultad de Ciencias Exactas y Naturales. UBA. Ciudad Universitaria, Pab Il. C1428EHA. Buenos Aires, Argentina; gbigatti@bg.fcen.uba.ar “Institute of Oceanography and Fisheries, P. O. Box 500, 21000 Split, Croatia “Department of Zoology, The Natural History Museum, London SW7 5BD, United Kingdom; j.taylor@nhm.ac.uk 418 BIGATTI ET AL. cyte itself and deposited at the outer surface by exocytosis (Jong-Brink et al., 1983; Huebner & Anderson, 1976). When present, a thick jelly coat covers the outer surface of the vitelline envelope, and appears as a loose association of striated fibrous material (Hodgson & Eckelbarger, 2000). Follicle cells are also believed to produce secondary com- pounds or cellular egg envelopes around оо- cytes (Eckelbarger, 1994). In Codakia orbicularis, the jelly coat that covers the оо- cyte is formed by an inner layer (the vitelline envelope) and an ощег layer (Gros et al., 1997). In another lucinid, Phacoides pectinata (Gmelin, 1791), this jelly coat is made up of glycoproteins and proteoglycans (Frenkiel et al., 1997), synthesized by the oocyte during vitellogenesis (Frenkiel, unpubl.). No informa- tion exists concerning the composition and formation of oocyte envelopes in Ctena orbiculata and L. pensylvanica, and the repro- ductive biology of these species has received no attention probably due to their smaller size and lack of commercial importance. MATERIAL AND METHODS The lucinid bivalves Codakia orbicularis, Ctena orbiculata and Lucina pensylvanica were collected between July 2002, at differ- ent localities in Florida Keys, USA, during the International Marine Bivalve Workshop 2002. Sampled habitats included sandy bottoms at 6 m depth (sampled by scuba diving) and shal- low subtidal sandy substrata colonized by the seagrasses Thalassia, Halodule, Syringodium and Halophila. Most of the seagrass-covered sediments were anoxic with hydrogen sulphide concentrations detectable by smell. Bivalves were collected by digging and sieving sedi- ment through 2 mm mesh screens. Sampling localities included a number of oceanside in- tertidal and shallow water sites in the Middle Keys (IMBW-FK-622, 628, 635, 638, 642, 647, 649; Mikkelsen & Bieler, 2004: fig. 1 for map). Ctena orbiculata was most abundant at in- shore bayside sites in sparsely vegetated sand patches; Codakia orbicularis at several oceanside sites with thick Thalassia growths, and Lucina pensylvanica was found commonly only at oceanside Station IMBW-FK-642 on Lower Matecumbe Key in shallow sand on rock. Twenty individuals of each species were sexed macroscopically observing the texture of gonads (females had granulose and males homogeneous texture) and their shell param- eters measured. A preliminary scale of gonad maturation was compiled from light micro- scope observations of fresh tissue, and this was used for comparison with thin sections of gonads prepared later. External gonad mor- phology was described from fresh animals. Samples of gonad tissue were fixed in Bouin's solution for 48 h and stored in 70% alcohol. In order to determine the first maturation stage and the presence of oocyte envelopes, sec- tions of the gonads were cut at 6 um with a Leitz microtome and stained with hematoxilin and eosin. We use the term oocyte or egg envelope for the inner layer and jelly coat for the outer layer of the oocyte. For scanning electron microscopy (SEM) of spermatozoon ultrastructure, pieces of male gonad were cold fixed in 2.5% glutaraldehyde solution in Зогепзоп’$ phosphate buffer. Slices of gonad were then cut with a razor blade, dehydrated through an ascending acetone series, critical point dried, sputter coated with gold, and ex- amined by scanning electron microscopy (SEM) with a Philips XL30 field emission SEM operated at 5kV. For age determination, 20 dry shell valves of Lucina pensylvanica were embedded in MET20 resin (Struers Ltd), sectioned from the umbo to the ventral edge, ground, polished, and etched for 20 min in 0.01M НС! and ac- etate peel replicas prepared (Richardson, 2001). The age of each shell was estimated by three observers using the major growth lines present in acetate peels of the umbonal region and the outer prismatic shell layer (Richardson, 2001). These major growth lines were treated as annual lines by comparison with the results of a study of Codakia orbicu- laris from the Bahamas by Berg & Alatalo (1984). Data were fitted to the von Bertalanffy growth function L, = L_(1-e*(*) using the Fish- eries Programme “Fisat”. Voucher specimens of the species studied are deposited in the Mollusca collections of the Department of Zoology, The Natural His- tory Museum, London. RESULTS Ctena orbiculata Specimens collected measured between 5.5 mm and 13.8 mm in length. No gonad devel- opment was observed in individuals smaller than 5.6 тт. А! males larger than 5.6 mm ООСУТЕ ENVELOPES AND SPERM IN LUCINIDAE 419 FIGS. 1-4. Ctena orbiculata. FIG. 1: Living mature oocytes covered by the vitelline envelope. Scale bar = 50 um; FIG. 2: Female gamete release. Scale bar = 3 mm; FIG. 3: Sperm in follicles. Scale bar = 100 um; FIG. 4: SEM image of sperm. Scale bar =10 um. Gr, gamete release; Ve, vitelline envelope. were mature. The spermatozoa were con- tained in follicles and orientated in relation to the lumen of the follicle (Fig. 3). Females less than 7.1 mm in length were immature; oocytes of individuals of 7.1 mm had a mean diameter of 20 mm (SD = 1.8) and no envelope was present. Developing oocytes were present in specimens between 7.2 mm and 7.8 mm length with maturity reached at 7.9 mm. Some individuals were spawning at this size, but no resorption was observed. The maximum oo- cyte diameter measured was 137.5 mm, with an oocyte envelope width of 10 mm, with no jelly coat covering it. When the gonad is ripe, an egg mass full of gametes forms a thin layer, that covers the ctenidia and sometimes all of the pallial cavity (Fig. 2). The egg mass has no defined organization. At this stage, the oo- cytes are sticky and covered by a jelly coat and have a total mean diameter of 360 um (Fig. 1). This corresponds to the time of ga- mete release and possibly the oocytes are re- tained by mucus in the pallial cavity until fertili- zation. After release, the oocytes are covered by the jelly coating and this probably provides protection to the egg until fertilization occurs. No evidence of either protandry or simultaneous hermaphroditism was found in this species. Male gametes are released as sperm strings comprising hundreds of sperm attached to each other at the head. SEM images (Fig. 4) show that sperm cells are relatively short- headed, with a long, cylindrical flagellum. The heads are cylindrical and tapering, slightly curved with a visible, short acrosome. Heads have a mean length of 7.5 um and a width of 1-1.2 um, with the acrosome about 0.6 um long and the mid piece 0.9 um. The tail mea- sures around 28 um, giving a total length for the sperm cell of around 36 pm. Parasites were observed in the digestive di- verticula lying adjacent to the gonad follicles 420 BIGATTI ЕТ AL. FIGS. 5-8. Codakia orbicularis. FIG. 5: Mature oocyte with jelly coat. Scale bar =100 um; FIG. 6: Female gamete release. Scale bar =500 um (arrow shows the finger-shaped mass); FIG. 7: Sperm ready to spawn. Scale bar = 50 um; FIG. 8: SEM image of sperm. Scale bar = 10 um. Gr, gamete release; Jc, jelly coat; Tb, tubules. (Fig. 13). They are probably cercaria of the digenean family Monorchiidae (D. Gibson, personal communication). We also found yel- low granules in the digestive diverticula, flux- ing to the gonads. Codakia orbicularis The bivalves sampled measured between 11.3 mm and 64.8 mm shell length. Individuals less than 12.5 mm were immature, while one individual of 12.6 mm had started to form fol- licles. Males larger than 13.2 mm had begun to mature with some sperm visible. All males larger than 25.9 mm were mature. In individuals larger than 46 mm, sperm were observed in tubules ready to spawn (Fig. 7). All females sampled had already spawned with visible oocyte resorp- tion. Females larger than 46.8 mm had oocytes of 100 um average diameter (SD = 5.8), cov- ered by an oocyte envelope of about 7.5 um thick. The largest male sampled was 47.1 mm in length. Unfortunately, no larger-sized Codakia were available to determine if protandry occurs in this species. No hermaph- rodites were found. Release of female gametes occurs in a mucous mass held within the mantle cavity similar to that of Ctena orbiculata, but in С. orbicularis it is finger shaped (Fig. 6). At this stage, the oocytes are sticky and slightly nega- tively buoyant. Upon release from the gonads, oocytes are spherical and covered by а 50 um thick jelly coat that, together with the oocyte envelope, forms the external envelope (Fig. 5). Further expansion of the envelopes results in an egg with a total diameter of around 200 um. ООСУТЕ ENVELOPES AND SPERM IN LUCINIDAE 421 FIGS. 9-12. Lucina pensylvanica. FIG. 9: Mature oocyte with irregular jelly coat. Scale bar = 100 pm; FIG. 10: Ova release. Scale bar = 5 тт; FIG. 11: Mature sperm in gonads. Scale bar = 50 um; FIG. 12: SEM image of sperm cells. Scale bar = 10 pm. ft, foot; g, gonad; Jc, jelly coat; Oo, oocyte; Ve, vitelline envelope. The sperm have long, slender, tapering, curved heads (Fig. 8) with a long flagellum that possesses a narrow, lateral, undulating mem- brane to either side (Fig. 14). The head mea- sures around 14-15 um in length with a width of 0.8 um at the posterior end. The lengths of the flagella were difficult to measure in our preparations, but were at least 25 um long, with the undulating membrane having a width of 0.42 um. Lucina pensylvanica Individuals sampled measured between 12.8-42.6 mm shell length. No gonad devel- opment was observed in individuals of less than 13 mm. Sexual differentiation with in- cipient follicular formation begins at a size of 13.8 mm (specimens of one to two years old). Individuals larger than 15.9 mm had well-de- veloped gonads; males were all mature and females exhibited different maturity stages. Male gametes are released as sperm strings comprising hundreds of sperm attached at the head as in Ctena orbiculata (Fig. 11). Females begin maturation at a shell length of 17.7 mm and reach maturity at around 26.6 mm. Indi- viduals having oocytes with a mean diameter of 130 um (SD = 6.7) with an oocyte enve- lope of approximately 10 um were classified as mature females. Release from the gonads occurs as a finger-shaped mucous mass con- taining the ova, similar to that of Codakia or- bicularis (Fig. 10). After release from the gonad, oocytes are spherical with an irregu- lar jelly coat (Fig. 9). This jelly coat is approxi- mately 47 um thick, covering the ova, which has an external maximum diameter of 245 um. Sperm have long, curved, tapering heads (Fig. 12), with a long flagellum without a lat- eral membrane. The mean length of the head 422 BIGATTI ET AL. FIG. 13. Parasite found in the digestive diver- ticula of C. orbiculata and L. pensylvanica. Scale bar = 100 um. is 15.5 ут and the width at the posterior end is 1.1 um. Flagella measured around 33 um in length. Cercaria similar to those of Ctena orbiculata were found as parasites in the di- gestive diverticula adjacent to the gonad fol- licles. Shell length (mm) N о FIG. 14. ЗЕМ image showing the lateral undu- lating membrane (Lm) of the flagellum of C. or- bicularis. Scale bar = 1 pm. From the analysis of major growth halts ob- served in shell sections, the oldest individual studied had attained an age of 5 years. Analysed shells showed individual variations in growth (Taylor et al., 2004). The asymptotic shell length was 38.37 + 9.36 mm (Fig. 15). 3 4 5 6 Relative age FIG. 15. Growth curve for Lucina pensylvanica fitted using the Von Bertalanffy growth equation: L=38.37 (1-e-0.60t+0.37)| ООСУТЕ ENVELOPES AND SPERM IN LUCINIDAE 423 DISCUSSION It should be emphasised that this was a very short-term study conducted over a period of only two weeks. Nevertheless, the results show that interesting similarities and differ- ences exist in the reproductive biology among the three coexisting lucinid species. For mature individuals of the three lucinid species studied, the gonads were ripe, spawn- ing or in resorption in July 2002. This spawn- ing period corresponds to the hottest time of the year, and surface water temperatures measured by the National Data Buoy Center (LONF1, Long Key, Florida) ranged between 29-30°C for the period 20-30 July. At the sam- pling sites, temperatures reached around 35°C in water depths of between 0.5-1 meter. According to Berg & Alatalo (1984), the an- nual size classes of Codakia orbicularis at Grand Bahama Island are 12, 29, 42, 52 and 63 mm for years 1 to 5, respectively. By com- parison with these results, our data suggest that C. orbicularis from the Florida Keys are immature at less than one year old, with males maturing between 1 and 2 years and females spawning at 3 years. In the Bahamian Codakia, shell growth was not continuous throughout the year, as evidenced from sharp growth rings on the external surface of the shell, and the number of these external growth rings coin- cided with annual year classes predicted from the von Bertalanffy growth equation (Berg & Alatalo, 1984). Spawning and recruitment of C. orbicularis occur over a long period, with variation in individual growth rates, and at Grand Bahama Island growth rates decreased during winter months. Follicular development in C. orbicularis was first observed at a shell length of 19.8 mm, corresponding to an age of 1.5 years, and the smallest animal with fully developed gametes measured 25.4 mm (Berg & Alatalo, 1984). In our study, we were unable to record the size of first maturity for females, but one individual of 12.6 mm length was ob- served with incipient follicular formation. Ac- cording to Berg & Alatalo (1984), this corresponds to an individual of about one year old. They also recorded three hermaphrodites from a total sample of 224 bivalves analysed. We recorded no hermaphrodites in our Florida Keys sample, but all individuals examined less than 46.6 mm long were males, with the larger sizes all females. This result suggests protandry for this population, but more data are needed for confirmation. The observation from the Bahamas that most animals greater than 30 mm were ripe during the summer months indicates the same maturity pattern for this species as at Florida Keys, for we found maturity at sizes above 25.9 mm length. Ani- mals within the population at Gold Rock Creek, Bahamas, did not spawn completely, nor si- multaneously, but appeared to continue spawning over a period of at least a month (Berg & Alatalo, 1984). The spawning season for С. orbicularis in Bahamas (Alatalo et al., 1984) and in Venezuela, Sucre State (Prieto et al., 1999), agrees with our results for July 2002 in Florida Keys. In our case, we found spawned females towards the end of July, with oocyte resorption visible, but only in individu- als greater than 46.8 mm. Observations on animals of both smaller and larger sizes are necessary to complete this study. Parasites observed in the digestive diver- ticula of Ctena orbiculata and Codakia orbicu- laris resembling the general body form of Cercaria caribbea LXIV of Cable (1963) were previously recorded from the lucinid bivalve host Ctena pectinella. Cable suggests that the adult is a species of Proctotrema that occurs in the porkfish Anisotremus virginicus. The cercaria were observed only in vivo, and fur- ther studies are needed to confirm this. The yellow granules found in the digestive diver- ticula of Ctena orbiculata could be sulphur granules, related to the chemoautotrophic habits of this species. Our age estimates for Lucina pensylvanica suggest that individuals less than one year old are immature, as in C. orbicularis, and that both males and females mature at two years. For the smallest species, Ctena orbiculata, we have no age data, but from the size range we consider that this species may also have early development. Selection for rapid gonad devel- opment could assure the recruitment of new spat when environmental conditions are favourable. Unfortunately, no further data is available concerning reproductive biology of Ctena orbiculata and L. pensylvanica. Another lucinid from the western Atlantic, the large Phacoides pectinata, a protandric spe- cies inhabiting mangrove swamps and reduc- ing mud, has permanently mature gonads (Frenkiel et al., 1997). This reproductive strat- egy, which, in addition to the sulphur oxidising bacterial endosymbionts and high bacteriocyte haemoglobin concentration, is considered an adaptation to a high-stress environment. Re- sorption of oocytes and recovery of metabo- 424 BIGATTI ET AL. lites through the follicular cell lysosomal func- tion appears to be the most efficient means to minimize the metabolic cost of maintaining the state of maturity. In the cases of Ctena orbiculata and L. pensylvanica, resorption of oocytes was found, suggesting the same re- productive strategy in these species also liv- ing in similar hypoxic habitats. In the case of C. orbiculata, we observed only females spawning, but larger sizes were not found. Probably the same strategy occurs in this spe- cies, as a possible adaptation to reducing sub- strata. We consider the oocyte envelope as a vi- telline envelope. The composition and loca- tion of its synthesis is still unknown for Ctena orbiculata and Lucina pensylvanica. For Codakia orbicularis, Gros et al. (1997) con- sidered that the jelly coat is made up of gly- coproteins and proteoglycans synthesised by the oocyte itself during vitellogenesis. After spawning, it is swollen by hydration of the proteoglycan components. In Phacoides pectinata, the same jelly coat is observed and the glycoproteins are likely to support recog- nition receptors for sperm (Frenkiel et al., 1997). In bivalves from Antarctica, such as Laternula elliptica (King & Broderip, 1832), a similar oocyte envelope is observed, allow- ing storage of mature oocytes for the whole year until environmental conditions are favourable (Bigatti et al., 2001). This vitelline envelope and the jelly coating protect the embryos of Codakia orbicularis in the veliger stage (Alatalo et al., 1984) and are digested by enzymes from the larva (Gros et al., 1997). This is another possible adaptive response to a high-stress environment. For Ctena orbiculata and Lucina pensylvanica, studies concerning oogenesis and the origin and com- position of oocytes envelopes are needed, but as they are members of the same family and live in similar environments, a similar pattern could be expected. For the Florida lucinids, our observations of the release of gametes adhering within the pallial cavity suggests that these species re- tain oocytes until fertilization. However, this observation does not suggest the existence of a brood chamber, such as occurs in Ostrea spp. (Morriconi & Calvo, 1979), because no embryos or divided cells were identified. Prob- ably the oocytes were recently spawned when we observed them. This could be another ad- aptation that enhances reproductive success after the eggs are released to the exterior en- vironment. Electron microscopy of molluscan sperm has provided an important set of characters for phylogenetic studies (Healy, 1995, 1996). Most detail is provided by TEM studies, but scan- ning microscopy of external features reveals a set of morphological characters useful for generic and specific differentiation. Our obser- vations show that Lucina pensylvanica and Codakia orbicularis have similarly proportioned large sperm with long, curved, tapering heads, but the latter differs in having an undulating membrane to the flagellum. By contrast, Ctena orbiculata sperm have much shorter and less tapering heads. The length of the spermatozoa of Lucina pensylvanica and Codakia orbicularis is com- parable with that of Scrobicularia plana (da Costa, 1778) (Souza et al., 1989), which is the longest described spermatozoon in bivalves. TEM studies show that Codakia orbicularis sperm have a short acrosome (Mouéza & Frenkiel, 1995), as does Codakia punctata (Linnaeus, 1758) (Healy, 1995) and Loripes lucinalis (Lamarck, 1818) (Johnson et al., 1996). It has been suggested by Mouéza & Frenkiel (1995) that occurrence of this feature, along with long tapering heads, in species having large oocytes with a gelatinous coat might be adaptations to facilitate penetration of the spermatozoa through this coating and the vitelline envelope. The undulating lateral membrane of the fla- gellum in C. orbicularis (Fig. 14), previously described by Mouéza & Frenkiel (1995), is an unusual feature of bivalve sperm, although well-developed undulating membranes appear characteristic of corbiculid sperm (Komaru & Konishi, 1996; Konishi et al., 1998). Mouéza & Frenkiel (1995) suggest that the structure is somehow related with sperm locomotion prior to fertilization. The sperm of too few lucinid species has been studied to evaluate the pos- sible systematic significance of this feature. Finally, longer-term studies of the reproduc- tive biology of Lucina pensylvancia and Ctena orbiculata are necessary to confirm and ex- pand our preliminary findings based on the two-week survey. Both of these species are abundant, but poorly studied, chemosymbiotic bivalves inhabiting the shallow waters of the Florida Keys and the populations are highly vulnerable to environmental disturbance of these fragile habitats. OOCYTE ENVELOPES AND SPERM IN LUCINIDAE 425 ACKNOWLEDGEMENTS We are indebted Rúdiger Bieler and Paula Mikkelsen for organising the Workshop and for providing us with the opportunity to work in the Florida Keys. Thanks are due to Lic. Carlos Sanchez Antelo for help with histological sec- tioning, to Pablo Penchaszadeh for his help- ful suggestions and to Guido Pastorino, Cristián Ituarte and Olivier Gros who helped with pertinent literature. Marcel Mouéza and Liliane Frenkiel provided literature and com- ments before the IMBW, 2002 and Armin Palaoro and Miro Kraljevic helped with the ageing of L. pensylvanica. Emily Glover helped with collecting the samples and provided con- stant encouragement. John Healy and an anonymous reviewer provided critical evalua- tions of the final manuscript. Bivalves were collected under Permit FKNMS-2002.079. The International Marine Bivalve Workshop, held in the Florida Keys, 19-30 July 2002, was funded by U.S. National Science Foundation award DEB-9978119 (to co-organizers R. Bieler and P. M. Mikkelsen), as part of the Partnerships in Enhancing Expertise in Tax- onomy (PEET) Program. Additional support was provided by the Bertha LeBus Charitable Trust, the Comer Science & Education Foun- dation, the Field Museum of Natural History, and the American Museum of Natural History. LITERATURE CITED ALATALO, P., С. J. BERG & С. N. D'ASARO, 1984, Reproduction and development in the lucinid clam Codakia orbicularis (Linné, 1758). Bulletin of Marine Science, 34: 424-434. BERG, C.J. 8 P. ALATALO, 1984, Potential of chemosynthesis in molluscan mariculture. Aquaculture, 39: 165-179. BIGATTI, G., P. E. PENCHASZADEH 4 G. MERCURI, 2001, Aspects of the gonadal cycle in the Antarctic bivalve Laternula elliptica. Jour- nal of Shellfish Research, 20: 283-287. CABLE, R. M., 1963, Marine Cercariae from Curacao and Jamaica. Zeitschrift fúr Parasitenkunde, 23: 429—469. DISTEL, D. L., 1998, Evolution of chem- autotrophic endosymbioses in bivalves. Bio- science, 48: 277-286. ECKELBARGER, K. J., 1984, Comparative as- pects of oogenesis in polychaetes. Pp. 123- 148, in: A. FISCHER & H. D. PFANNENSTIEL, eds., Polychaete reproduction: progress in compara- tive reproductive biology. Fortschritte der Zoologie, Band 29. Gustav Fischer Verlag, Stuttgart, New York. FRENKIEL, L., O. GROS 8 M. 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ECKELBARGER, 2000, Ultrastructure of the ovary and oogenesis in six species of patellid limpets (Gastropoda: Patellogastropoda) from South Africa. Inverte- brate Biology, 119: 265-277. HUEBNER, E. 8 E. ANDERSON, 1976, Com- parative spiralian oogenesis. Structural as- pects: an overview. American Zoologist, 16: 315-343. JOHNSON, М. J., М. CASSE & М. LePENNEC, 1996, Spermatogenesis in the endosymbiont- bearing bivalve Loripes lucinalis (Veneroidea: Lucinidae). /nvertebrate Reproduction and Development, 45: 476-484. JONG-BRINK de, M., Н.Н. BOER 4 J. JOOSSE, 1983, Mollusca. Pp. 297-355, in: K. G. ADIYODI & В. С. ADIYODI, eds., Reproductive biology of invertebrates. Volume I: Oogenesis, oviposi- tion, and oosorption. John Wiley & Sons Ltd., London. KOMARU, A. & K. KONISHI, 1996, Ultrastruc- ture of biflagellate spermatozoa in the fresh- water clam, Corbicula leana (Prime). Invertebrate Reproduction and Development, 29: 193-197. KONISHI, K., К. KAWAMURA, Н. FURUITA & А. KOMARU, 1998, Spermatogenesis of the freshwater clam Corbicula aff. fluminea Müller (Bivalvia: Corbiculidae). Journal of Shellfish Research, 17: 185-189. MIKKELSEN, P. M. & R. BIELER, 2000, Marine bivalves of the Florida Keys: discovered biodiversity. In: E. M. HARPER, J. D. TAYLOR & J. A. CRAME, eds., The evolutionary biology of the Bivalvia. Geological Society of London, Spe- cial Publications, 177: 367-387. MIKKELSEN, Р. М & R. BIELER, 2004, Interna- tional Marine Bivalve Workshop 2002: Intro- duction and Summary. In: R. BIELER & P. M. MIKKELSEN, eds., Bivalve Studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 241-248. 426 BIGATTI ET AL. MORRICONI, E. & J. CALVO, 1979, Reproduc- tive cycle and sex alternation in Ostrea puelchana. Physis, 38: 1-17. MOUEZA, М. & L. FRENKIEL, 1995, Ultrastruc- tural study of the spermatozoon in a tropical lucinid bivalve: Codakia orbicularis L. Inverte- brate Reproduction and Development, 27: 205-211. PRIETO, А. S., J. VILLALBA & L. J. RUIZ, 1999, Produccion especifica de la almeja Codakia orbicularis (Veneroida: Lucinidae) en una poblacion del Golfo Cariaco, Estado de Sucre, Venezuela. Boletin del Instituto Oceanografico de Venezuela,Cumana, 38: 63-72. RICHARDSON, C. A., 2001, Molluscs as ar- chives of environmental change. Oceanogra- phy and Marine Biology: an Annual Review, 39: 103-164. SOUZA, M., L. CORRAL & C. AZEVEDO, 1989, Ultrastructural and cytochemical study of sper- matogenesis in Scrobicularia plana (Mollusca: Bivalvia). Gamete Research, 24: 393-401. TAYLOR, J. D. & E. A. GLOVER, 2000, Func- tional anatomy, chemosymbiosis and evolution of the Lucinidae. In: E. M. HARPER, J. D. TAYLOR & J.A. CRAME, eds., The evolutionary biology of the Bivalvia. Geological Society of London, Special Publications, 177: 207-255. TAYLOR, J. D., E. A. GLOVER, M. PEHARDA, С. BIGATTI & A. BALL, 2004, Extraordinary flexible shell sculpture; the structure and for- mation of calcified periostracal lamellae in Lucina pensylvanica (Bivalvia: Lucinidae). In: В. BIELER & P. М. MIKKELSEN, eds., Bivalve Stud- ies in the Florida Keys, Proceedings of the In- ternational Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 277-294. Revised ms. accepted 31 October 2003 MALACOLOGIA, 2004, 46(2): 427-458 PERIGLYPTA LISTERI (J. E. GRAY, 1838) (BIVALVIA: VENERIDAE) IN THE WESTERN ATLANTIC: TAXONOMY, АМАТОМУ, LIFE HABITS, AND DISTRIBUTION Rüdiger Bieler'*, Isabella Kappner' & Paula M. Mikkelsen? ABSTRACT Periglypta listeri (J. Е. Gray, 1838), one of the largest and most distinctive western Atlantic venerids, and the only Atlantic member of the genus, is redescribed based on original mate- rial from the Florida Keys, museum specimens, and literature records. Conchologically, this species agrees with previously described venerids in having a well-developed escutcheon and lunule, and a hinge with three cardinal teeth in each valve. Within the genus, it is unique in having internal purplish brown coloration, and in the frequent presence of a purplish brown “hinge dot” on the anterior lateral tooth. This is the first anatomical study for any species in the genus Periglypta, and the most complete so far for any member of Venerinae. Periglypta listeri agrees with previously described venerids in most anatomical characteristics, and no- tably features an undulating mantle edge that can close in “zipper” fashion, tentacles at the anterior mantle edge, and branching tentacles at the tips of the unfused siphons, type B mantle fusion, type C(2) ctenidia, and a type V stomach. Although empty shells are commonly collected, Р /isteri unusually (for venerids) lives cryptically in rubble or sand among rocks, and/or in reef settings. Thus far, the presence of an anterior lateral hinge tooth is the sole morphological feature separating the subfamily Venerinae from the closely allied Chioninae. Key Words: Florida Keys, Mollusca, Caribbean, infaunal, clam, sanctuary. INTRODUCTION Veneridae is the largest marine family of bivalves, with many of the more than 500 living species forming key components in the world’s clam fisheries. The nominate subfamily Venerinae currently comprises 14 genus-group taxa (e. g., Venus, Periglypta, Globivenus, Ventricoloidea; Keen, 1969) with more than 140 nominal extant and fossil species. Members of this subfamily live in a wide range of benthic habitats, in coarse sand, mud, or gravel be- tween tide lines to depths over 150 m, and from temperate to tropical seas. Delimiting shell char- acteristics are the presence of both radial and concentric sculpture and an anterior lateral tooth in the left valve (Keen, 1969; see hinge discussion below). The nominal subfamily Chioninae, another large group comprising such genera as Chione, Mercenaria, and Proto- thaca, has recently been synonymized with Venerinae by some authors (Coan & Scott, 1997; Coan et al., 2000). Relationships between these and among other venerid subfamilies remain unresolved, due to a surprising paucity of comparative morphological work. Despite their relative abundance and commercial im- portance, only about 50 venerid species have some published anatomical data. Most publi- cations focus on a few species traditionally grouped in the Chioninae, such as representa- tives of Mercenaria (Kellogg, 1892, 1903, 1915; Morse, 1919; Jones, 1979), Chione (Kellogg, 1915; Jones, 1979; Narchi & Gabrieli, 1980), Timoclea (Ansell, 1961; Narchi, 1980), Lirophora (Jones, 1979), Tawera (Burne, 1920), Chamelea (Odhner, 1912; Ansell, 1961), Anomalocardia (Narchi, 1972; Purchon, 1985), Bassina (Morton, 1985; Purchon, 1985), Protothaca (Guerón & Narchi, 2000), and Clausinella (Ansell, 1961). Anatomical details for members of other nominal venerid subfami- lies are much sparser, with data available for individual species of Gouldiinae [= “Circinae”] (Pelseneer, 1911; Ansell, 1961; Fishelson, 2000), Cyclininae (Purchon, 1985), Dosiniinae (Thiele, 1886; Ansell, 1961; Guéron & Coelho, 1989; Fishelson, 2000); Gemminae (Morse, ‘Department of Zoology, Division of Invertebrates, Field Museum of Natural History, 1400 $. Lake Shore Drive, Chicago, Illinois 60605-2496, U.S.A. *Division of Invertebrate Zoology, American Museum of Natural History, Central Park West at 79th Street, New York, New York 10024-5192, U.S.A. *Corresponding author: bieler@fieldmuseum.org 428 BIELER ET AL. 1919; Sellmer, 1967; Narchi, 1971), Meretricinae (Kellogg, 1915; Narchi, 1972; S. Gray, 1982; Narchi & Dario, 2002), Pitarinae (С. В. Sowerby |, 1854; Thiele, 1886; Pelseneer, 1911; Kellogg, 1915; Morse, 1919; Narchi, 1971; Е. R. Ber- nard, 1982; S. Gray, 1982; Fishelson, 2000; Morton, 2000), and Tapetinae (G. B. Sowerby ||, 1854; Carrière, 1879; Pelseneer, 1894, 1897, 1911, 1923, 1931; Berkeley, 1959; Ansell, 1961; Nielsen, 1963; Joshi € Bal, 1965а, b; Morton, 1985; Fishelson, 2000). The morphological diversity of the nominate subfamily, Venerinae, remains largely unex- plored, with published anatomical data re- stricted to Circomphalus casina (Linnaeus, 1758) (Ansell, 1961), Venus verrucosa Linna- eus, 1758 (type species of Venus Linnaeus, 1758, and of Clausina Brown, 1827; Pelseneer, 1894, 1897), and Globivenus toreuma (Gould, 1850) (Pelseneer, 1911). The current paper fo- cuses on a venerine species currently classi- fied in the genus Periglypta Jukes-Brown, 1914, a group not previously studied anatomically. Periglypta listeri (J. E. Gray, 1838), also known as “Lister's venus” or “princess venus”, is the second largest Caribbean venerid species, only exceeded in shell size by Mercenaria campechiensis (Gmelin, 1791), which ranges from the mid-Atlantic coast to the Gulf of Mexico and extends into the Caribbean. A shallow-wa- ter species with a very conspicuous shell, P. listeri had at one point even been declared the type species of the genus Venus (Stoliczka, 1871: xvii; Venus verrucosa Linnaeus, 1758, was subsequently fixed as the type by ICZN Opinion 195, 1954). Empty shells of Р listeri are commonly collected, but living specimens are less frequently encountered, due in part to their relatively cryptic infaunal habitat in seagrass areas and algae-covered rubble near reefs. This paper reviews the taxonomy and geo- graphic distribution of this species, its anatomy, and life habits, based on original information from living specimens from the Florida Keys, together with a re-evaluation of existing litera- ture and selected museum data. Comparisons are drawn with sympatric large-bodied venerids in the western Atlantic, with selected worldwide species of Periglypta, and with known anatomi- cal data for the family. MATERIALS AND METHODS This study is part of an ongoing investigation of marine molluscan biodiversity in peninsular Florida and the Florida Keys, formally initiated by RB and PMM in 1994. Consecutively num- bered stations comprising these collections are preceded by ап “ЕК” acronym in the following text. Living animals and empty shells were col- lected by hand mainly during scuba diving on coral reefs and shallow-water (2-10 m) patch reefs, rubble areas, and ledges. The majority of live observations were made on specimens from the “Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), com- parable to stations IMBW-FK-629 and -637 re- ported by Mikkelsen & Bieler (2004). Interpre- tations of distribution records based Henderson's Eolis expeditions are taken from the recent compilation by Bieler £ Mikkelsen (2003). Specimen photography used a variety of equipment and techniques. /n situ photographs of living animals (Fig. 24) were taken using a Nikonos V underwater camera with close-up lens. Other living animals (Figs. 25, 26) were photographed in aquaria using standard 35 mm single-lens reflex or electronic cameras. Whole- valve and detail light micrography (Figs. 3-12, 31, 32) used a Microptics® micro/macro imag- ing system based on a high-resolution Nikon) single-lens reflex digital camera. Excised pre- served tissues were prepared for scanning elec- tron microscopy (SEM) by critical point drying and gold sputter coating, then viewed at beam acceleration voltages of 10kV on a Amray 1810 scanning electron microscope at FMNH. For anatomical observation, specimens were relaxed by chilling in a household refrigerator assisted by the addition of magnesium sulfate crystals (Epsom salts) to their seawater sup- ply, or in an isotonic aqueous magnesium chlo- ride solution. Ciliary currents were studied using carmine particles. Anatomy was observed un- der a dissecting microscope; preserved tissues were dyed for better contrast with neutral red or methylene blue. Voucher FK specimens were fixed in 5% formalin, later transferred to 70% ethanol, and are deposited in the Field Museum of Natural History (FMNH), Chicago, and the American Museum of Natural History (AMNH), New York. All measurements and meristics were taken from the left valve whenever possible. Shell measurements, taken with calipers or with ocu- lar micrometer on a stereomicroscope, include: maximum height from umbo to farthest distal point on free edge, and maximum length (= width) perpendicular to axis of height. Size is expressed as shell length unless otherwise noted. Radial ribs were counted at the growth PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 429 edge on the main body of the shell. For mor- phometric analyses (length to height ratio), 32 shells were measured and the linear regres- sion calculated using Microsoft® Excel 2000. Other cited repositories include: ANSP Academy of Natural Sciences of Philadelphia, Pennsylvania, U.S.A. The Natural History Museum [= Brit- ish Museum (Natural History)], Lon- don, United Kingdom Bailey-Matthews Shell Museum, Sanibel Island, Florida, U.S.A. Carnegie Museum of Natural History, Pittsburgh, Pennsylvania, U.S.A. Delaware Museum of Natural History, Wilmington, U.S.A. Museum of Comparative Zoology, Harvard University, Cambridge, Mas- sachusetts, U.S.A. Museum National d'Histoire Natu- relle, Paris, France Staatliche Naturhistorische Sammlun- gen, Museum fúr Tierkunde, Dresden, Germany North Carolina State Museum of Natural Sciences, Raleigh, North Carolina, U.S.A. Northern Territory Museum of Arts and Sciences, Darwin, Australia SBMNH Santa Barbara Museum of Natural History, Santa Barbara, California, U.S.A. Rosenstiel School of Marine and At- mospheric Science [= University of Miami Marine Laboratory], University of Miami, Florida, U.S.A. BMNH BMSM CMNH DMNH MCZ MNHN MTD NCSM NTM UMML USNM National Museum of Natural History. [= United States National Мизеит], Smithsonian Institution, Washington, DC., U.S.A. ZMB Museum für Naturkunde [= Zoolo- gisches Museum, Berlin], Humboldt- Universität, Berlin, Germany Other abbreviations: alc fluid-preserved (alcohol) specimen frag shell fragment juv juvenile or subadult LV left valve pair an empty (dead) complete shell (2 valves) RV right valve spm a live-collected specimen valve an empty (dead) single valve RESULTS Veneroidea: Veneridae: Venerinae Rafinesque, 1815: 146 (as Veneridia) Periglypta Jukes-Brown, 1914: 72 Type species, by original designation: Venus puerpera Linnaeus, 1758. Periglypta was т- troduced as a subgenus of Antigona Schu- macher, 1817, by Jukes-Brown (1914: 72). Cytherea Róding (1798), non Fabricius, 1794 (Diptera: Bombylidae). Type species (subse- quent designation of Dall, 1902): Venus puer- pera Linnaeus. Periglypta listeri (J. E. Gray, 1838) Selected synonymy: ?”pectunculus admodum crassus ...” Lister, 1687: Liber Ш, fig. 178 [pl. 341, fig. 178 in later editions]. [unlabelled figure] — Lamarck, 1797: pl. 378, fig. 2a-b. Venus puerpera (2) Var. — Lamarck, 1818: 584-585 [referring to Lister (1687) and Lamarck (1797) figures; non Venus puerpera Linnaeus, 1771]. Venus puerpera Var. 2 - Deshayes, 1832: 1112 [referring to Lister (1687) and Lamarck (1797) figs.]; - Deshayes, 1835: 335. Venus puerpera — Schramm, 1869: 20. Dosina Listeri J. E. Gray, 1838: 308. Venus listeri— Hanley, 1843, in 1842-1856: 110; — Deshayes, 1853a: 106 [distribution (Philip- pines, Australia) erroneous]; — G. B. Sowerby Il, 1853 [in part]: 705, pl. 152, fig. 8 [excluding figs. 7, 9; distribution (Philippines, Australia) erroneous]; — Reeve, 1863: no. 14, pl. 5, fig. 14 [“Hab. Philippine Islands; Cuming” errone- ous]; — Krebs, 1864: 97; — Pfeiffer, 1869: 141, pl. 8, figs. 8, 9 [distribution (Nicobares, Philip- pines) erroneous]; — Simpson, 1889: 64; - F. С. Baker, 1891: 47; — Cockerell, 1894: 118; — Benthem Jutting, 1927: 34; - McLean, 1936b: 119; — Dance, 1974: 263-264, fig.; — Fischer- Piette, 1975: 36-37; — Dance, 1977: 264, fig. Venus (Periglypta) listeri - Lamy, 1929: 205. Omphaloclathrum listeri — Mórch, 1853: 24. Venus (Chione [Omphaloclathrum]) listeri — Römer, 1867: 32 [“Insulae Philippinae” in error]. Venus crispata — Dall, 1889: 54 [non Venus crispata Deshayes, 18535]. Cytherea (Cytherea) listeri - Dall, 1902: 372. Cytherea listeri — Dall, 1903: 1275-1276, 1279; — Maury, 1920: 103. 430 BIELER ET AL. Antigona (Periglypta) listeri — Jukes-Brown, 1914: 72; - Warmke & Abbott, 1961: 185, pl. 38 fig. L; - Humfrey, 1975: 248, pl. 30, fig. 3. Antigona (Dosina) listeri — Palmer, 1927: 337 [129]; - Palmer, 1929: pl. 28, figs. 2, 11; — Abbott, 1954: 404, pl. 32, fig. m; — Abbott, 1958: 129. Antigona listeri - Weisbord, 1926: 83; — Johnson, 1934: 48; — Lermond, 1936: 6; — Clench 8 McLean, 1936: 166; — McLean, 1936a: 41; — Clench 8 McLean, 1937: 39- 40; — М. Smith, 1937: 53, pl. 21, fig. 11; — М. Smith, 1940: 110, fig. 1445; Jaume & Perez Farfante, 1942: 39 [Pleistocene]; — Jaume, 1946: 101; - Aguayo & Jaume, 1949: 1; — Pulley, 1952: 150, pl. 13, fig. 7; — Nowell- Usticke, 1959: 14; — Abbott, 1961: 162; — Weber, 1961: 58; — Rice & Kornicker, 1962: 382, pl. 7, fig. 6a-b; — Moulding, 1967: 83; — Brooks, 1968: 8; — J. A. Baker, 1969: 3-4; — Ross, 1969: 8; — Voss et al., 1969: 71; - Abbott, 1970: 162; — Stanley, 1970: 160, pl. 21, figs. 12, 13; — McGinty, 1970: 58 [Lower Pleistocene]; — Magnotte, [1970-1979]: 63, fig. 12; - Woods, 1970: 2-3; — McGinty 8 Nelson, 1972: 13; — Godcharles & Jaap, 1973: 37; — Zischke, 1973: 35; — Zischke, 1977a: 29; — Zischke, 1977b: 338, fig. A.14— 67; - Romashko, 1974: 49, fig. 17; - Ekdale, 1974: 657; — Eisenberg, 1981: 169, pl. 151, fig. 16; — Abbott, 1986: 230, fig. 5; — Sutty, 1990: 92, fig.; — Prieto et al., 2001: 593. Antigona (Antigona) listeri -McLean, 1951: 82, рЁ 15, 19. 5. Periglypta aff. listeri — Weisbord, 1964: 300- 302, pl. 43, figs. 7, 8 [Pliocene]. Periglypta listeri — Morris, 1973: 58, pl. 24, fig. 13; — Abbott, 1974: 521, color pl. 24, fig. 5852; — Emerson 8 Jacobson, 1976: 429, pl. 42, fig. 18 [AMNH 106142; seen by authors]; — Parodiz, 1976: 20 [Mayan ruins]; — Lozet 8 Petron, 1977: 129, fig. 247; - Edwards, 1980: 3; — Theroux 8 Wigley, 1983: 47, fig. 85 (map); — Voss et al., 1983: 316, 429; - Romashko, 1984: 96, fig. р. 97; — H. E. Vokes 8 E. H. Vokes, 1984: 43, pl. 45, fig. 9; - Abbott, 1984: 54, fig. 9; — Lipe & Abbott, 1991: 76, fig. 9; - Lawson, 1993: 53; — Espinosa et al., 1994: 123; — Diaz М. & Puyana H., 1994: 78, pl. 18, fig. 170; — Abbott 8 Morris, 1995: 60, pl. 30; — Lyons & Quinn, 1995: J-13; — Alvarez, 1998: 103 ff.; Pointier 8 Lamy, 1998: 214, fig.; — Tremor, 1998: 7; — Turgeon et al., 1998: 48; — Mikkelsen & Bieler, 2000: 379; — Redfern, 2001: 236, pl. 101, fig. 965. Periglyphus (sic) listeri - Voss et al., 1983: 79, 183. Distribution (Figs. 1, 2) North Carolina, southeastern and western Florida, including the Florida Keys, Texas, the West Indies, Caribbean Central America and northern South America; apparently not reach- ing Brazil. lt appears to have a largely Carib- bean island (as opposed to Gulf of Mexico or mainland North or South American) distribu- tion. Itis rare off Texas and other Gulf of Mexico locations. It extends along the entire island chain of the Florida Keys, from Key Largo to the Dry Tortugas. Localities (*= unverified): *North Carolina (Pul- ley, 1952). Florida: east coast (AMNH, ANSP, FMNH, USNM; Dall, 1902; M. Smith, 1937; Pul- ley, 1952; Stanley, 1970; McGinty & Nelson, 1972), Florida Keys (AMNH, ANSP, BMSM, CMNH, DMNH, ЕММН, USNM, this study; Simpson, 1889; Dall, 1902; Palmer, 1927; Lermond, 1936; M. Smith, 1937; Pulley, 1952; Abbott, 1961; Brooks, 1968; Ross, 1969; Abbott, 1970; Magnotte, 1970-1979; Woods, 1970; Godcharles & Jaap, 1973; Zischke, 1973, 1977a, b; Edwards, 1980; Voss et al., 1983; Lyons & Quinn, 1995; Tremor, 1998; W. G. Lyons, pers. comm.), Dry Tortugas (USNM, this study), *west coast (M. Smith, 1937; Pulley, 1952); Texas (ANSP); Caribbean: Bahamas (AMNH, ANSP, FMNH, USNM; Clench 8 McLean, 1936; McLean, 1936b; Clench & McLean, 1937; McLean, 1938; Moulding, 1967; J. A. Baker, 1969; Lawson, 1993; Redfern, 2001), Cuba (ANSP, FMNH, MTD, USNM); McLean, 1936a; Aguayo 8 Jaume, 1949; Pulley, 1952), Cayman Islands (ANSP; Abbott, 1958), Jamaica (USNM), Hispaniola — Haiti and Dominican Republic (AMNH, ANSP, USNM; Palmer, 1927), Puerto Rico (ANSP, USNM; Clench & McLean, 1937; McLean, 1951; Warmke 8 Abbott, 1961), Virgin Islands (AMNH, ANSP, FMNH, USNM; Krebs, 1864; Dall, 1902; McLean, 1951; Nowell-Usticke, 1959; Weber, 1961), Antigua (USNM), *Martinique (Lamy, 1929; Fischer-Piette, 1975); “Guadeloupe (Schramm, 1869; Pointier 8 Lamy, 1998); Grenada (ANSP); Netherlands Antilles (DMNH, FMNH, USNM; Benthem Jutting, 1927); Central America: Mexico (AMNH; F. C. Baker, 1891; Weisbord, 1926; Jaume, 1946; Rice 8 Kornicker, 1962; Ekdale, 1974; Parodiz, 1976 [Mayan ruins]; H. E. Vokes & E. H. Vokes, 1984); Belize (ANSP, USNM), Honduras (USNM; Alvarez, 1998); Costa Rica (USNM); Panama (USNM); South America: Colombia (AMNH, FMNH, USNM; Díaz M. 8 Puyana H., 1994), “Venezuela (Prieto et al., 2001); “French Guyana (Femorale, 2003). PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 431 Atlantic Ocean FIG. 1. Distribution of Periglypta listeri; * denotes record for region, without details. South Florida Gulf of Mexico A SOE у À x ses JN e C4 %° 2 a POS SE e © > .. PE ar % ) In je ® Florida Straits FIG. 2. Distribution of Periglypta listeri т the Florida Keys; (+) living records, (A) dead records. 432 BIELER ETAL. Fossil Record Lower Pleistocene (McGinty, 1970: 58) and Upper Pleistocene (M. Campbell, in lit., Feb. 2003) of southern Florida; Pleistocene of Cuba (Jaume & Perez Farfante, 1942) and Santo Domingo [Hispaniola] (Dall, 1903). It was col- lected from Mayan archaeological sites (but not from concurrent Recent collections) in Yucatan, Mexico, by Parodiz (1976). Weisbord (1964: 302) noted that the species was “reported from the Pleistocene of ... Barbados, and the Island of Tortuga, Venezuela”, but did not include source citations for these records; Weisbord examined additional fragments from the Lower Mare for- mation (Pliocene, possibly Lower Pliocene) at Quebrada Mare Abajo, northern Venezuela. Cassab (1984) recorded the eastern Pacific P. multicostata (G. В. Sowerby |, 1835) from several Tertiary western Atlantic locations — the Chipola Formation (Lower Miocene) of Florida, the Pirabas Formation (Lower Miocene) of Para State, Brazil, and the Middle Miocene of Costa Rica; verification of the species-level identity of these materials, which date prior to the clo- sure of the Panamanian landbridge, 1$ war- ranted. All post-closure western Atlantic Periglypta records should most definitely be reconsidered against P listeri; Dall (1903: 1279) noted that Р multicostata “has been enumer- ated as one of the reef Pleistocene fossils of St. Domingo, but doubtless through a mis- identification, perhaps of [P] listeri.” A large- shelled form described as P. tamiamensis Olsson & Petit, 1964, from Florida's Late Mi- ocene and Pliocene Tamiami formation seems closely related. The relationship of Р listeri to extinct western Atlantic species from older geo- logical formations [e.g., Antigona dominica Palmer, 1928 (= A. caribbeana Anderson, 1927), Miocene of Santo Domingo, fide Hertlein & Strong, 1948; P. tarquinia (Бай, 1900), Oli- gocene of western Florida and Santo Domingo, called a small “precursor” of Р. listeri by Dall, 1903; P mauryae (H. E. Vokes, 1938), Upper Miocene of Trinidad], awaits a more compre- hensive review of the genus. Type Material No original material located (see Taxonomic Remarks, below). Material Examined See Appendix 1. Diagnosis Large western Atlantic venerid, with thick- walled, inflated, trapezoid shell, with external sculpture of erect commarginal ridges crenu- lated by underlying radial sculpture, and with posterior end vertically truncated. Exterior cream-colored, with scattered brown speck- les, blotches, or flames. Interior yellowish white with more-or-less strongly developed purplish brown stain around posterior adductor muscle scar, posterior margin, and above the pallial line. Anterior lateral hinge tooth often with a purplish brown “hinge dot”. Description Shell relatively heavy, equivalve, longer than high, trapezoid, with nearly straight dorsal margin and bluntly truncated posterior margin; inequilateral with low, rounded umbones ap- proximately 1/3 of the shell length from the anterior end (Figs. 3, 4). Length to height ratio very regular throughout lifespan (R? = 0.9929, n = 32). Inflated, with posterodorsal slope somewhat concave. External sculpture con- sisting of prominent erect commarginal ridges, regularly spaced, reflected umbonally, those along posterior margin higher and not re- flected; ridges more-or-less alternating in strength at anterior and posterior margins as well as on later growth of shells (beginning at 2.5-3 cm or after first 15-25 commarginal ridges); ridges crenulated by underlying radial sculpture consisting of uniform flattened ribs separated by narrow grooves approximately 1/2-1/3 width of ribs (Fig. 5). Lunule broadly spindle- to teardrop-shaped (Figs. 6, 13), with deeply incised margins, asymmetric, right half slightly larger than left, with many very fine commarginal lamellae (continuing the much coarser ridge pattern of the anterior shell), without radial elements. Escutcheon distinct (Fig. 6), delimited by a marginal groove, right half overlapping left in the posteriormost third, both halves finely obliquely grooved. Externally cream-colored with brown speckles, blotches, or flames, sometimes darker posterodorsally, occasionally radially merging into (often three) broadening radial stripes; escutcheon and lunule with color of surrounding shell. Internal margin finely crenulated, continuing onto lunular margin. Anterior adductor muscle scar oval; posterior adductor muscle scar bean- shaped, somewhat flattened dorsally; poste- rior scar larger and more curved than anterior; PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 433 FIGS. 3-9. Periglypta listeri, Florida Keys specimens. FIGS. 3, 4: External shell; ЕММН 176372, Little Duck Key, 75 тт; FIG. 5: Sculptural detail of shell in Fig. 3. Scale bar = 5 тт; FIG. 6: Umbonal aspect, FMNH 296695, Rachel Bank, 66 mm; FIGS. 7, 8: Internal shell: FMNH 296695, Rachel Bank, 62 mm; FIG. 9: Loose pearls from 78 mm shell; АММН 295199, Spanish Harbor Keys, largest pearl with maximum dimension of 5.8 mm. (AMMS, accessory mantle muscle scars; E, escutcheon; GrL, Pigmented growth lines on posterior adductor muscle scar; HD, hinge dot; L, lunule). 434 BIELER ЕТ AL. FIGS. 10-12. Periglypta listeri, details of specimen in Figs. 7, 8. FIGS. 10, 11: Close-up of hinge teeth; FIG. 12: Articulated shells. (1, right middle cardinal tooth; 2a, left anterior cardinal tooth; 2b, left middle cardinal tooth; 3a, right anterior cardinal tooth; 3b, right posterior cardinal tooth; 4b, left posterior cardinal tooth; HD, hinge dot; All, left anterior lateral tooth). Scale bars = 5 mm. dorsalmost portion of posterior scar formed by pedal retractor muscle scar, separated by faint demarcation. Anterior pedal retractor muscle scar on ventral side of hinge plate, dorsomedial to anterior adductor muscle scar, just below anterior cardinal tooth (Fig. 12: 3a). Pallial line entire (Figs. 7, 8); pallial sinus wide, roundly pointed anteriorly. Accessory pallial muscle scars just inside pallial line, more-or-less regu- larly spaced. Internal color yellowish white with purplish brown stain around posterior adduc- FIGS. 13, 14. Periglypta listeri, details of umbo and prodissoconch; AMNH 296531, Looe Key reef, Florida Keys, 9.5 mm. FIG. 13: Anterior aspect with lunule. Arrows point to transition from wide commarginal lamellae on surface to fine striae on lunule; FIG. 14: Transition between prodissoconch | and II (arrow). Scale bars = 1 mm (Fig. 13), 100 um (Fig. 14). tor muscle scar and posterior margin, extend- ing dorsally above the pallial line, sometimes also ventrally below, occasionally also sur- rounding anterior adductor and pedal retrac- tor muscle scars; additional faint purplish brown “lines” often extending toward umbo from an- terior limits of pallial sinus and anterior pedal retractor muscle scar (Figs. 7, 8). Live-collected specimens that are pure white internally also noted (e.g., USNM 464243, 890543, 890776). Color pattern within posterior adductor muscle scar sometimes showing distinct growth lines (Fig. 7). Hinge teeth of left valve (Figs. 11, 12) comprising minute anterior lateral [A//] (see Discussion below), prominent triangular ante- rior cardinal (2a), slightly smaller bifid middle cardinal (2b; with posterior part 1/4-1/3 less prominent), and lamellate posterior cardinal (4b). Hinge teeth of right valve (Figs. 10, 12) comprising well-defined, relatively small ante- rior cardinal (3a), larger bifid middle cardinal (1; with posterior part narrower and smaller), PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 435 FIGS. 15-18. Diagrammatic anatomy of Periglypta listeri. FIG. 15: Organs of the mantle cavity, with RV and most of right mantle removed. Arrows on gills and labial palps indicate direction of particle flow. Outer labial palp (OLP) is reflected to show structure and particle flow over palps; FIG. 16: General dissection of alimentary system, with RV and right mantle, siphons, gills and labial palps removed; FIG. 17: Gut-loop variation in two additional dissected specimens (FK-273, FK-357); FIG. 18: Vertical section through gill, illustrating direction of food currents along inner and outer demibranchs. (A, anus; AAM, anterior adductor muscle; APRM, anterior pedal retractor muscle; AU, auricle of heart; BA, bulbus arteriosus; CG, cerebral ganglia; Cut M, cut mantle edge; DG, digestive gland; E, esophagus; EX, excurrent siphon; F, foot; GL, gut loop; |, intestine; IDB, inner demibranch; ILP, inner labial palp; IN, incurrent siphon; K, kidney; M, mantle edge; MO, mouth; ODB, outer demibranch; OLP, outer labial palp; PAM, posterior adductor muscle; PG, pedal ganglia; PPRM, posterior pedal retractor muscle; S, stomach; SS, style sack; V, ventricle of heart; VG, visceral ganglia). 436 BIELER ЕТ AL. and prominent, wide, equally bifid posterior cardinal (3b). Anterior lateral tooth at base of anterior cardinal of LV, and its corresponding socket in RV, often each with purplish brown “hinge dot” (Figs. 8, 10, 11). Prodissoconch | visible at tip of umbo (Figs. 13, 14), 155 um in length (n = 1; AMNH 296531, FK-269). Uncertain border between prodisso- conch II and juvenile shells (at about 1.3 mm). Juvenile sculpture (Figs. 13, 23) initially smooth, followed by fine commarginal ridges with wide bands of radial ribs between. Sur- face pattern of brown to orange speckles par- ticularly noticeable on juvenile shell (Fig. 23). Foot large (Fig. 15), with brown pigment (in living specimens) between muscular part and visceral mass, with pedal gland opening slightly anterior of midpoint; pedal groove extending from ventral posterior end toward anterior third. Mantle muscles attaching mantle to shell, some extending dorsalward to produce so-called accessory muscle scars dorsal to pallial line (approx. 1.5 mm dorsal, in larger specimens). Attachment of anteriorly pointed siphonal re- tractor muscles producing (and reflecting shape of) pallial sinus on shell. Anterior and posterior adductor muscles (AAM, PAM, re- spectively) oval; PAM slightly larger than AAM. Posterior pedal retractor muscle (PPRM) round, arising anterodorsal to PAM. Fibers of PAM differently colored in living material, with posterior third darker than remainder. Anterior pedal retractor muscle (APRM) attaching to underside of anterior hinge plate and inserted into anterodorsal part of foot a short distance posterior to AAM. Visceral retractor muscle attached to inner surface of umbo behind hinge plate, inserting into roof of visceral mass. Mantle with four mantle folds as previously described for other venerids (Ansell, 1961; Yonge, 1957). Ventral mantle margins distinctly wavy (Figs. 15, 20, 26) in living and preserved specimens; capable of closing in “zipper” fash- ion and held closely appressed in living indi- FIGS. 19-22. Periglypta listeri, scanning electron micrographs of critical-point dried tissues; AMNH 295199, West Summerland Key, 73.2 mm. FIG. 19: Mid-ventral portion of the outer demibranch, with food groove (arrows); FIG. 20: Ventral mantle margin with wavy inner median mantle fold; FIG. 21: Anteriormost portion of mantle margin with all four mantle folds; FIG. 22: Detail of anterior mantle margin with triangular tentacles. (IF, inner mantle fold; M1, inner middle mantle fold; M2, outer middle mantle fold; OF, outer mantle fold. Scale bars = 100 um (Figs. 20, 22), 1 mm (Figs. 19, 21). PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 437 viduals at rest. Pedal gape extending ventrally from base of incurrent siphon to AAM. Anterior 1/8 of inner median mantle fold with small tri- angular tentacles (Figs. 21, 22). Siphons sepa- rated (Figs. 15, 26), each with distinct black pigment (persisting in preserved specimens) between tentacles internally and externally; pig- ment stronger at terminal margin and fading toward basal region; internally with densely placed yellow-white surface papillae; each si- phon with terminal digitate tentacles and basal siphonal membrane consisting of a thin tissue flap narrowing the lumen, with membrane of incurrent siphon forming a double ridge. Siphonal mantle fusion of type B (Yonge, 1957, 1982); union of inner and middle folds expos- ing outer surface of middle folds, with common outer ring of sensory tentacles. Incurrent (ven- tral) siphon slightly larger in diameter and length than excurrent (dorsal) siphon (Figs. 24-26); excurrent siphon tapering to narrower terminal diameter than wider, slightly flaring incurrent FIGS. 23-26. Periglypta listeri, juvenile shell and living specimens. FIG. 23: Juvenile shell with color pattern; FMNH 296719, West Summerland Key, Florida Keys, 7.2 mm; FIG. 24: Siphons of living specimen /7 situ, muddy sand and algal cover, 2 т depth; West Summerland Key, not collected; FIG. 25: Siphons of living specimen in sand in laboratory tank; FMNH 301448, West Summerland Key, 56 mm; FIG. 26: Living specimen in laboratory tank; ЕММН 295706, off Stirrup Key, Florida Keys, 59 mm. 438 BIELER EMA siphon. Siphonal tentacles digitate, those оп incurrent siphon with 5-11, on excurrent with 3-5, lateral papillae, interspersed with addi- tional small, simple tentacles; both kinds often- tacles on excurrent siphon with distal opening of unknown function (Figs. 29, 30); complex tentacles on incurrent siphon (Figs. 24-28) 30— 75% larger, slightly more pointed and more numerous (approx. 40 tentacles of various sizes in 73 mm specimen, FK-273; AMNH 295199) than those on smaller excurrent siphon (approx. 30 tentacles, same specimen; Figs. 24-26, 29- 30). Distal end of excurrent siphon with pointed dome-shaped valve surrounded by ring of ten- tacles, presumably allowing control of current (Figs. 24-26, 30). Demibranchs smooth (not plicated; Fig. 15), with axes nearly vertical dorsoventrally; inner demibranch slightly larger than outer, each with numerous interlamellar junctions. Surface cur- rents moving particles ventrally on the outer surface of each demibranch; currents on inner surfaces uncertain. Food grooves at distal edges of inner and outer demibranchs (Figs. 18, 19), with oralward currents, indicating gills and ciliation of type C(2) (Atkins, 1937); longi- tudinal oralward current also found between bases of adjacent demibranchs. Triangular palps with narrow lamellae on inner surface (33-37 lamellae, n = 2; FK-273, AMNH 295199, 73 mm; FK-352, FMNH 283534, 67 mm); margins smooth at ventral and dorsal side; outer surface smooth. Acceptance cur- rents on palps along lamellae directed ven- trally, and on ventral boundary oralward. Rejection currents counter-oralward on smooth ventral and dorsal margins of palps. Ctenidial/labial palp association of type II (Stasek, 1963); anteroventral tips of inner demibranch inserted and fused to distal oral groove between labial palps. Esophagus relatively short, with 7-8 longi- tudinal rugae, leading into anterior part of stomach (Fig. 16); stomach embedded in di- FIGS. 27-30. Periglypta listeri, scanning electron micrographs of critical point dried siphonal papillae; AMNH 295199, West Summerland Key, 73.2 mm. FIG. 27: Digitate tentacles on incurrent siphon, exterior aspect of siphon; FIG. 28: Same, viewed from interior of siphon; FIG. 29: Detail of simple tentacle of excurrent siphon with orifice of unknown function; FIG. 30: Digitate tentacles on excurrent a front of flap-like valve (arrow). Scale bars = 1 mm (Fig. 27), 100 um (Figs. 28, 30), 10 um 102229); PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 439 gestive diverticula, located posterior to labial palps. Stomach of type V (Purchon, 1985), with two caeca. Right caecum with seven ducts; left caecum with 11 ducts opening into diges- tive diverticula; four ducts of digestive diver- ticula opening into left pouch. Major typhlosole (MT) and intestinal groove extending from in- testine into stomach, penetrating right caecum; MT crossing stomach floor beneath esoph- ageal opening into left саесит; МТ continu- ing from stomach into intestine, ending just after gut loop. Minor typhlosole projecting into stomach but ending close to midgut opening. Gastric shield located opposite crystalline style sac at roof and left side of stomach, with flanges that pass into dorsal hood and left pouch. Dorsal hood with left and right sorting areas, located over esophageal opening. Style sac combined with midgut, together exiting posteroventrally from posterior end of stom- ach, then turning anteriorly and coiling on ven- tral side of stomach, turning again posteriorly and crossing style sac, then ascending (as hindgut, penetrating pericardium, ventricle and bulbus arteriosus) to continue dorsally and posteriorly across surface of PAM (Fig. 16). Gut loops varying somewhat (Fig. 17) in con- figuration between individuals, but always in- cluding a single ventral loop. Ventricle of heart surrounding intestine, with lateral auricles connecting to outer limbs of kidney (Fig. 16). Kidney positioned along ven- tral edges of pericardium and anterior to PAM (Fig. 16). Three pairs of ganglia (Fig. 16), with cerebral ganglia between APRM and AAM, joined by supraesophagal commissure; vis- ceral ganglia ventral and slightly anterior to PAM; pedal ganglia extensively fused, anteroventral to the ventral gut loop. Gonad surrounding stomach and intestine within vis- ceral mass; reproductive system not otherwise investigated. Dimensions and Maximum Recorded Size Median length approximately 65 mm, rang- ing 13-100 mm (mean 63.3 + 14.7 mm SD, п = 103). Maximum 100.2 mm, St. Thomas, Virgin Islands; AMNH 31868 [registered specimen, K. Hutsell (San Diego, California), Registry of World Record Size Shells]. Largest Florida Keys specimen, 96 mm, FK-601, FMNH 296718. Habitat and Ecology Based on observations in southeastern Florida's Biscayne Bay, Stanley (1970: 160) described this species as “widespread in in- tertidal and shallow subtidal settings, in large numbers; it is generally restricted to coarse substrate and grassy areas”. Voss et al. (1969: 71) reported the species, for the same region, from the highly unlikely habitat of “rocks; pil- ings; seawalls” as well as from seagrass beds, open sand, and lagoonal patch reefs. Other habitat records include sand (FMNH, this study; Abbott, 1974; Humfrey, 1975; Dance, 1977; Tremor, 1998), mud (Humfrey, 1975), loose rock/rubble (this study; Godcharles & Jaap, 1973; Zischke, 1973; Voss et al., 1983), and seagrass (this study; Clench & McLean, 1937; Zischke, 1973; H. E. Vokes & E. H. Vokes, 1984). Clench & McLean (1937: 39) noted that P. listeri was “exceedingly abun- dant” in Savannah Sound (Eleuthera, Baha- mas) but generally uncommon elsewhere. At Savannah Sound, they lived buried 5-10 cm below the surface of seagrass-covered sand bars; native Bahamians were observed using “a short stick as a probe [to locate the clams] ... [they are not] eaten by the natives though they are gathered in this way for fish bait” (Clench & McLean, 1937: 37). In the present study, one living specimen was found nestled in red algae within the cavity of a large barrel sponge at 8 m depth (FK-395), although the favored habitat for the species seems to be sand with loose rubble. It undoubtedly is a shallow-water species, in contrast to the analy- sis by Theroux & Wigley (1983: 47) who placed this species based on two dredge records “in the 50-99 т depth range grouping’. Collect- ing records of all (living and dead) material range from shallow water to 84 m (Theroux & Wigley, 1983), with the deepest confirmed live- collected specimen in this study from 8 m (FK- 395). Stanley (1970: 160) described the slow burrowing process of this species and noted that (in Biscayne Bay) the depth of burial is to some extent dependent upon interference by the subsurface rhizomes of surrounding turtlegrass (Thalassia testudinum König). In the laboratory, when permitted to burrow in its native sediment with the seagrass removed, а 5.8 cm-long animal assumed a position with the posterior shell margin 4.5 cm beneath the sediment surface. Specimens observed in the present study were often found hindered from deep burrowing by rubble, and had their shells barely covered by substratum. In addition to an herbivorous diet, evidenced through the presence of a crystalline style, P. listeri ap- pears to opportunistically ingest Zooplankton: two copepods were found in the stomach of 440 BIELER ET AL: one individual (FK-352). In Puerto Rico, this species is a “favourite food of the Slipper Lob- ster” (Sutty, 1990: 92). In the Florida Keys, empty shells have been frequently found as- sociated with octopus middens at scuba depths, or with beveled drill holes, the latter indicative of predation by naticid gastropods. Pearls A single living animal was found with a se- ries of loose pearls lining the center of the in- side shell, apparently in response to the remains of an intruding worm-shaped organ- ism (Fig. 9). The 78 mm specimen (FK-273, АММН 295199) contained 11 irregular pearls, the largest with a maximum dimension of 5.8 mm. This 1$ the first record of pearls from Periglypta; both free and attached pearls have previously been reported from several other venerids, especially Mercenaria spp. (Haas, 1931; Shirai, 1994; Hill, 1996; Landman et al., 2001; Mienis, 2001). Taxonomic Remarks Although there is general consensus in the recent literature of applying the species name “listerT to this western Atlantic taxon, the taxo- nomic history of that name 1$ not without com- plications. Dosina listeri was introduced by J. E. Gray (1838: 308). He placed it in an un- named section of Dosina J. E. Gray, 1835, characterized by “anterior lateral tooth small, sometimes obliterated”, together with three pre- viously described species: Venus verrucosa Linnaeus, 1758 (as “Dosina veerrucosa [sic], Venus veerruicosa [sic], Linn.”); Venus reticulata Linnaeus, 1758; and Venus puerpera Linnaeus, 1771. While other species in the same article were expressly identified as new species descriptions (labeled as “n.s.”, accom- panied by a textual description and including an indication of the originating collection; e.g., as done for Grateloupea cuneata J. E. Gray, 1838: 304), the name D. listeri appears as a new name referring to prior literature data (1838: 308). The complete “description”, cryp- tic by today’s standards, introducing the name listeri for a previously unnamed variety of Ve- nus puerpera Linnaeus, 1771, reads as follows: “Dosina Listeri, V. puerpura [sic] var., Linn. Sow. Gen. f. Ency. Meth. t. 278, f. 2”. Venus puer- pera was introduced by Linnaeus (1771: 545), in the Mantissa. Linnaeus himself did not men- tion varieties in the original description and re- ferred to two prior illustrations — Gualtieri (1742: pl. 83, fig. F) and Argenville (1742: pl. 26, fig. F). In Hanley’s words (1855: 453): “Neither of the very dissimilar figures referred to bears the least resemblance to the shell which has been universally accepted for the species.” In any case, Venus puerpera clearly is an Indo-Pa- cific, not Atlantic, species (Fig. 31). Lamarck (1818: 584-585) and Deshayes (1832: 1112) distinguished two varieties of V. puerpera; their “Venus puerpera var. 2” referred to the cited figure of Lister (1687), as well as to an illustra- tion by Lamarck (1797: pl. 278, fig. 2a, b) in the Encyclopédie Méthodique. J. E. Gray (1838) had named Dosina listeri for Martin Lister, author of the Historia Conchyliorum (1685-1692). The first refer- ence to a figure by Lister in conjunction with Venus puerpera, and the first reference to a distinct variety of this species (as y), appeared in the 13" edition of Linnaeus’ Systema naturae. The latter was authored by Gmelin (1791: sp. 3276), and it is the entry in this work that Gray seemed to have meant with his “var., Linn.”. The indicated Lister figure (1687: pl. 341, fig. 178) is in agreement with today’s con- cept of Periglypta listeri, but lacks detail to dis- tinguish it from similar bivalve shells. J. E. Gray’s (1838) introduction of the new name Dosina listeri included references to two works. As outlined above, he did not seem to have actual specimens before him, and it ap- pears that only the specimens referred to in these two works (and, possibly, Lister’s origi- nal material) qualify as the type series (ICZN, 1999: Art. 72.4). One reference, and the only one indicated by Gray with actual plate and figure numbers, was to Lamarck’s figures (1797: pl. 278, fig. 2a, b) as cited earlier by Lamarck (1818) and Deshayes (1832) for “va- riety 2” of Venus puerpera. The latter are ex- cellent illustrations of an external right valve and of the external hinge area of an articu- lated specimen. Lamarck’s illustration matches today’s concept of western Atlantic Periglypta listeri. Somewhat confusingly, Deshayes (1835: 334) stated that Lamarck’s specimen of “var. 2” and the figure 2a, b of the Encyclopédie fit well with the typical V. puer- pera of Linnaeus. Lamy & Fischer-Piette (1938: 292) agreed and considered all such Lamarck material in the MNHN collection to represent V. puerpera [as well as (from a speci- men agreeing with figs. 1a, b of Encyclopédie pl. 278) Venus magnifica Hanley, 1845]. The interior features of the shell, particularly its coloration, are unknown; the specimen on which Lamarck’s excellent illustrations were PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 441 based cannot currently be located (B. Metivier, 10/2002 in lit.). The other work, cited Бу J. E. Gray as “Sow. Gen. f.”, is С. В. Sowerby l's Genera of Re- cent and fossil shells (1834). In it, figure 1 of the Venus plate shows detailed color illustra- tions of the inside valves of a species identi- fied as V. puerpera. However, the illustrated valves lack the brown or purple stains usually present in Periglypta puerpera and P. listeri and instead are shown with extensive orange coloration below the umbo. The illustrated specimen or specimens (considerable differ- ences in the shape of mantle line and muscle scars indicate that these drawings of valves might originate from different individuals) have not been located in the BMNH collection. The orange interior coloration is matched by a specimen of P. puerpera from J. E. Gray's col- lection (BMNH 1991135, unknown locality, 49 mm; seen by authors). However, this shell dif- fers considerably from Sowerby's illustrations in having dark brown markings at the shell margin, oval (not angular) anterior muscle scars, and a much more angular posterior shoulder. Another specimen at BMNH, fur- nished as a potential syntype of Dosinia listeri (К. Way, 02/2003 in lit; BMNH 1840.7.1.41, 76 mm, no locality, old British Museum collec- tion; see Appendix), is a member of P. listeri in today’s sense, but likewise not a shell illus- trated by С. В. Sowerby | (1834). It is here, for reasons outlined above, not considered part of the syntypic series. Because the identity of P. listeri is not currently in dispute, and the fig- ured specimens of Lamarck and Sowerby can- not be located at present, we refrain from designating a lectotype (or neotype) in the context of this study and defer to a future ge- nus-wide revision. Subsequent British authors, particularly Hanley (1843, in: 1842-1856: 110) and G. B. Sowerby II (1853: 705), referred to Venus listeri as a species distinct from V. puerpera. How- ever, their concept of this nominal species was broader and included differently colored forms with dark rayed patterns on the shells, as evi- denced by the range of included illustrations. Venus listeri was thought to hail from the “In- dian Seas”, from the Philippines and Austra- lia. Nevertheless, С. В. Sowerby И (1853: 705) expressly endorsed Lamarck's oft-cited Шиз- tration for V. listeri by stating “The figure in the Encyclopaedia is very exact for the type.” Reeve (1863: pl. 5, no. 14) narrowed the т- terpretation of V. listeri to shells that are “fulvous-white, obscurely freckled with flesh- brown”, but still assumed the waters near the Philippines Islands as its home. It was in the context of an Indo-Pacific interpretation of М listeri that Reeve (1863: pl. 3) stated that he doubted that eastern Pacific Periglypta multicostata “is anything more than a variety of V. listeri, in which the ribs are more timidly thickened and recurved.” Fischer-Piette (1975) assigned most of the Indo-Pacific records of Venus/Dosinia/Chione/Cytherea/Antigona listeri to P. puerpera and restricted Р listeri to Atlantic records. Comparative Remarks Periglypta listeri is one of the largest west- ern Atlantic venerids, second only to Merce- naria campechiensis in maximum adult size. Its sculpture, of flattened radial ribs between sharp commarginal ridges (which are further crenulated by the radials), renders it distinct from other sympatric venerids: M. campe- chiensis lacks discernible sculpture between the commarginals, whereas Globivenus (for- merly Ventricolaria) rugatina (Heilprin, 1886), С. rigida (Dillwyn, 1817), and Circomphalus strigillinus (Dall, 1902) have fine concentric striae between the commarginal ridges and are also rounder in general shell shape. Periglypta listeri is part of a worldwide com- plex of morphologically similar species, the relationships of which far exceed the scope of this paper. Rómer (1867: 32, here translated) referred to similarities so great “that the differ- ences often can barely be put into words” when comparing “Venus” listerito other nominal spe- cies of Venus — Venus lacerata Hanley, 1845; V. clathrata Deshayes, 1853; V. crispata Deshayes, 1853; V. multicostata G. B. Sowerby |, 1853; М. laqueata С. В. Sowerby ll, 1853; V. resticulata С. В. Sowerby Il, 1853; V. chemnitzii Hanley, 1844; V. sowerbyi Deshayes, 1853; V. reticulata Linnaeus, 1758; and V. monilifera С. В. Sowerby II, 1851. Cit- ing a high degree of variability, E. A. Smith (1885: 120-121) considered “Venus” resti- culata, V. aegrota Reeve, 1863, V. lacerata Hanley, V. sowerbyi, V. clathrata, V. crispata, V. puerpera, V. listeri, and probably also V. multicostata and V. magnifica Hanley, 1845, as “races” of a single biological species, al- though he fell short of formalizing this action in not placing these names in synonymy un- der V. puerpera. Modern revisionary treatment of the Indo-Pacific taxa is urgently needed. Periglypta listeri is the only living member of the genus in the Atlantic Ocean. It differs from 442 BIEEER-ERAE: FIGS. 31, 32. Contrasting shell morphology in Periglypta spp. FIG. 31: Р puerpera; FMNH 82478, Mindanao, Philippines, 59 mm; FIG. 32: Р multicostata; FMNH 165936, Pacific Panama, 111 mm. the Indo-Pacific P puerpera (Fig. 31), type species of the genus, in general shell shape and coloration. Periglypta puerpera is less anteroposteriorly elongated, and generally more rounded in outline. Its external sculpture appears smoother, a result of its less promi- nent commarginal ridges. Externally it is cream- colored, with or without scattered radial brown flecks, often with 1-3 darker brown rays, one almost always extensively covering the poste- rior third. Internally it is white with a bright purple (not purplish brown) stain at the posterior mar- gin below the PAM, sometimes also with a yel- low or peach-colored flush at the center (although internally pure-white specimens also occur). As earlier noted by E. A. Smith (1885: 120-121), P puerpera has a “V-shaped purple mark upon the apex of the umbones”, refer- ring to two radial color bands on the prodissoconch and earliest juvenile stage; this is lacking in examined specimens of P listeri. Weisbord (1964: 302) called Periglypta listeri the western Atlantic analog of Panamic P. multicostata, citing a difference in the outline of the posterior end (obliquely vertical and trun- cated in P. listeri, subtruncated and more rounded in P. multicostata; see Fig. 32); al- though based on many museum specimens (AMNH, n = 31), this difference is only obvi- ous among the largest specimens. Periglypta multicostata attains a larger size (maximum observed 120 mm, АММН 248600, Baja Cali- fornia Norte, Mexico) than P listeri. Externally, the commarginal ridges of P multicostata are decidedly coarser and more prominently dor- sally reflected than those of P listeri, render- ing the crenulations and radial ribs less obvious. Internally, P. multicostata is white (or very occasionally flushed with pink, not pur- plish brown, either centrally or in an oblique posterior streak from umbo to margin, but not prominently surrounding the posterior muscle scars) and marginal crenulations are less no- ticeable or absent in the largest specimens (although the latter are quite prominent in specimens < 65 mm). DISCUSSION Periglypta listeri agrees with previously de- scribed venerids in conchological features, such as a well-developed escutcheon and lunule, and a hinge with three cardinal teeth in each valve (Keen, 1969). Although Lamprell (1998) cited differences in the shape of the pallial sinus, we found that distinguishing char- acters at the species level reside in sculpture and color/color pattern. So far as is known, the pattern of the internal purplish brown col- oration and the purplish brown “hinge dot” are unique to P. listeri within the genus. PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 443 This paper presents the first anatomical study for any species in the genus Periglypta, and is arguably the most extensive anatomical de- scription yet available for any member of the subfamily Venerinae. Anatomical information is published for only three other venerines. Pelseneer (1894, 1897) provided minor details about the anatomy of Venus verrucosa (tongue-shaped foot without byssus, very small labial palps, siphons more or less fused). He later (Pelseneer, 1911) presented similarly cur- sory details (siphons short and united, with retractors; short “byssal” groove on posterior foot; very narrow external demibranch) for Venus (now Globivenus) toreuma. Ansell (1961: fig. 8) illustrated (but did not extensively discuss) the gross anatomy and stomach struc- ture of Circomphalus casina (Linnaeus, 1758), which are closely similar to those in P. listeri with respect to the gills, labial palps, foot, stom- ach, and adductor muscles, as well as the flow of particles over the gills and labial palps. Like P. listeri, C. casina features a structurally com- plex ventral mantle edge, but its function was not mentioned. Its siphons appear short, un- like the longer, separated siphons of P. listeri. Ansell (1961; also Pelseneer, 1911) considered the degree of siphonal fusion to vary greatly within Veneridae, and from these minimal data this could also be true within Venerinae. The anatomy of Periglypta listeri agrees with these and other previously described venerids in most features. As elaborated by Yonge (1957), Ansell (1961), and Narchi (1971), the mantle edge has four folds (outer, OF; inner middle, M1; outer middle, M2; and inner, IF). The periostracum is secreted between OF and M2. In Р listeri, M1 is undulating, with its an- terior part elaborated into distinctly triangular tentacles. The wavy structure and tentacles can close in “zipper” fashion (Fig. 26), presum- ably to protect the organs of the mantle cavity from intrusion of particles and small organisms. At the anterior end of the pedal gape, OF fuses with M2, and M1 fuses with IF. At the posterior end, M1 and IF are fused to form the siphons; this configuration typifies venerids of type B fusion (Yonge, 1948, 1957, 1982; Ansell, 1961). The conical valve on the tip of the ex- current siphon is the elaborated IF, while the ring of tentacles on the siphon represents M1 (Yonge, 1957; Ansell, 1961; Jones, 1979). The tentacles on the incurrent siphon are not dis- tinguishable into inner and outer rings of ten- tacles, in contrast to Jones’ (1979) findings in members of Chione, Mercenaria, and Austrovenus. The siphonal tentacles of P. listeri are long and digitate, serving as an ef- fective screen to block intrusion of particles. Ansell (1961) contrasted the digitate tentacles of Circomphalus, Timoclea, Clausinella, Venerupis, and Gafrarium spp., which live in gravelly and stony bottom habitats, against the simple tentacles of Chamelea and Dosinia spp., which live in cleaner sand or gravelly bottoms. Periglypta listeri is not congruent with this pattern; it carries digitate tentacles but lives in a gravel and sand habitat without a high amount of detritus in suspension. At the proximal end of each siphon, Periglypta listeri has а siphonal membrane or valve comprised of thin tissue flaps, as de- scribed for other venerids (Ansell, 1961; Jones, 1979). That of the incurrent siphon consists of a basal double ridge; Ansell (1961) described this double ridge in Circomphalus casina and, like others (Kellogg, 1915; Jones, 1979; Narchi & Dario, 2002) postulated that such a valve can be opened to admit water inflow, or closed to direct water ventrally and flush out accu- mulated pseudofeces. The anterior end of the gill in Periglypta listeri is inserted into the distal oral groove, in the midline of the labial palps to which the gill is fused (Stasek, 1963). The ctenidia, with food grooves on the edges of both demibranchs, correspond to Atkins’ (1937) type C(2) along with members of Paphia, Mercenaria, Chione, Tivela, and Saxidomus. Other venerines, such as Venus verrucosa and Circomphalus casina, as well as other venerids (in C/ausinella, Dosinia, Gafrarium, Chamelea, and Paphia), lack a groove on the outer demibranch and have therefore been assigned to type C(1b). However, it must be noted that Atkins (1937) found this character variable within genera (e.g., Paphia) and species (e.g., only one of four specimens of C. casina had a groove on the outer demibranch), indicating that this char- acter requires larger sample sizes and further research. The circulatory and nervous systems are broadly similar to the species studied by Jones (1979), as is the general plan of the digestive system and configuration of the gut loop. The type V stomach of Р listeri agrees overall with the descriptions by numerous authors (Ansell, 1961; Dinamani, 1967; Narchi, 1971, 1972; Jones, 1979; Purchon, 1987; Narchi & Dario, 2002) for other members of the family. A com- bined style sac and midgut is typical of venerids, except for Placamen tiara (Dillwyn, 1817), in which the openings, although close together, lead into separate tubes (Dinamani, 444 BIELER ET AL. 1967). The numbers of ducts entering the left and right caeca vary among species and range, respectively, from a minimum of one and two in Nutricola tantilla (Gould, 1853) to a maximum of 13 and seven in Venerupis pullastra (Montagu, 1803) (Purchon, 1987); P. listeri is at the high end of this range with 11 and seven ducts. Four additional ducts open into the left pouch in P. listeri, in contrast to two or three in Meretrix, Katelysia, Placamen, Sunetta, and /rus spp. (Dinamani, 1967), five ducts in Tivela and Gafrarium spp., or eight in Dosinia spp. (Purchon, 1987). No additional ducts enter the stomach near the right cae- cum in P. listeri, unlike in Clausinella and Callista spp. (Purchon, 1987). Periglypta listeri lives in rubble or sand among rocks, and/or in reef settings (as op- posed to soft bottoms like most other venerids). This is also true for the similar east- ern Pacific P. multicostata, aptly named “giant reef clam’, described аз a common species in 3-6 m depth in Baja California Sur (Garcia- Dominguez et al., 1998), and noted from sand among rocks (Keen, 1971). Other Periglypta species recorded for rocky or reef habitats are P. puerpera and P reticulata (fide Whitehead, 1983; Graham, 1995). Periglypta was originally described as a sub- genus of Antigona, and Harte (1998: 358) maintained that that is its proper placement. However, although Antigona Schumacher (type species by original designation, A. lamellaris Schumacher, 1817, Indo-Pacific) shares radial threads between the commar- ginal shell ridges with Periglypta, its members have radial ribs extended onto the lunule, no groove around the escutcheon, a more trian- gular pallial sinus, and hinge teeth with a much wider 3b and smaller 2b. Periglypta also dif- fers from Dosina J. E. Gray (type species, D. zelandica J. E. Gray, 1835, South Pacific, by subsequent designation of Frizzell, 1936), in which Р. listeri was originally described, by the predominantly concentric sculpture (Keen, 1969) and a weakly or undeveloped escutch- eon in that genus (pers. obs.). Most authors (e.g., Keen, 1969) refer to an anterior lateral tooth in the left valve as char- acteristic of venerines, but an anterior lateral tooth is also present in members of other venerid subfamilies (e.g., Gouldiinae [= “Circinae”], Sunettinae, Meretricinae, Pitarinae, and Dosiniinae; Keen, 1954). There are indi- cations that such lateral teeth result from dif- ferent ontogenetic pathways and might not be homologous. Felix Bernard (1895: 127) pre- sented an ontogenetic series of hinge devel- opment for a Miocene species of Gouldia, which (together with its numbering system) became the model for tooth development in the “corbiculoid” hinge type sensu Cox (1969: N54). According to this model, the venerine anterior lateral tooth of the left valve (desig- nated as All) derives from lamella //. Whereas F. Bernard (1895: 127) noticed differing hinge morphologies of other venerids, such as Macrocallista, he still recognized the anterior lateral tooth in the latter as a lamella // deriva- tive, and thus an All. Marwick (1927: 598-599), in another ontogenetic comparison, showed that the Oligocene venerine Kuia vellicata (Hutton, 1873) displayed the same All devel- opment as shown in F. Bernard’s Gouldia study. However, Marwick (1927: 598) main- tained that the left anterior lateral tooth of Macrocallista, a continuation of a low ridge . proceeding from below the umbo, is “in no way connected with the anterior cardinal”. Some subsequent authors (Frizzell, 1936) followed Marwick’s argument of non-homologous an- terior lateral teeth in the Veneridae; Grant & Gale (1931: 316) referred to the venerine All of Antigona as a “pseudolateral”, in contrast to the situation in groups such as Macro- callista. lt appears that venerid “anterior lat- eral teeth” might variously be derived from either lamella // or IV and homology assump- tions of these structures need closer scrutiny. Chionines do not have anterior lateral or pseudolateral teeth even though their overall shell morphology is very similar to that of venerines. Anatomical studies carried out by Jones (1979) showed that chionine siphons are usually fused along their length in con- trast to the unfused siphons of Periglypta listeri seen in this study; the high degree of variability in siphonal fusion, as well as the presumably fused siphons in the venerine Circomphalus casina (see above) render this a weak distinguishing character. At present, the presence of an А// lateral hinge tooth 1$ postulated as the only reliable morphological feature separating these two nominal subfami- lies. Although recently synonymized by Coan 8 Scott (1997), preliminary molecular studies (16S gene; I. Kappner, unpubl.) indicate a dis- tinct grouping that might warrant retention of these subfamilial units. Studies of additional taxa, and more anatomical and molecular characters are needed for resolution of subfamilial synapomorphies. PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 445 ACKNOWLEDGMENTS This work is part of an ongoing investigation of the molluscan diversity of the Florida Keys; regional surveys and collections were sup- ported by permits from the Florida Keys Na- tional Marine Sanctuary (080-98, 2000-036, 2002-078, and 2002-079); in the vicinity of Pigeon Key (on National Register of Historic Places) under the auspices of the Pigeon Key Foundation; in the Dry Tortugas National Park, under collecting permits DRTO-19970030 and 2002-SCI-0005; in Long Key State Park (Long Key, Florida Keys) under Florida Department of Environmental Protection permit 5-02-43; in Key West National Wildlife Refuge (near Sand Key, Florida Keys) under United States Fish and Wildlife Service permit 41580-01-07. Additional collecting was sanctioned under Florida Fish and Wildlife Conservation Com- mission permit 99S-024 to affiliates of The Bailey-Matthews Shell Museum (Sanibel, Florida) and permit 01S-056 (as well as an- nual permits for prior years of this study) to affiliates of the Smithsonian Marine Station (Ft. Pierce, Florida; logistic support by Mary E. Rice and staff is much appreciated). We thank Timothy Collins, Timothy Rawlings, Roberto Cipriani, Deirdre Gonsalves-Jackson, Cecelia Miles, Louise Crowley, Daniel Miller, - Jim Culter, the Smithsonian Marine Station at Fort Pierce, and the captains and crews of R/ V EUGENIE CLARK (Mote Marine Laboratory, Sarasota, Florida) and R/V CORAL REEF II (Shedd Aquarium, Chicago) for collecting as- sistance. Data gathering from other museum collections was facilitated by Gary Rosenberg (ANSP), José Leal and Tina Petrikas (BMSM), Charles Sturm (CMNH), Timothy Pearce, Leslie Skibinski, and Albert Chadwick (DMNH), Katrin Schniebs (MTD), Nancy Voss (UMML), Jerry Harasewych (USNM), Ronald Jansen (Senckenberg Museum, Frankfurt), Matthias Glaubrecht and Lothar Maitas (ZMB). Bernard Metivier (Museum National d’Histoire Naturelle, Paris), and Kathie Way (BMNH) pro- vided type specimen information and loans. Richard E. Petit, as so often before, helped us disentangle obscure literature references. IK greatly appreciates Osmar Domaneschi's advice on anatomical questions and thanks Gustav Paulay and Rudo von Cosel for very fruitful discussions. The research, in part, was Supported by NSF-PEET DEB-9978119 and Comer Science and Education Foundation grants to RB and PMM. 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Revised ms. accepted 31 October 2003 APPENDIX 1: Material Examined Material examined (this study, Florida Keys) RB unnumb., М of Bahia Honda State Park, shallow water, sand, 25 Mar 1989 (1 spm alc, FMNH 288810); FK-018, “The Stakes”, off Middle Florida Keys, scuba, 20 ft (6.1 m), patch reef/ledges, sand patches, M/V SITE FINDER, 08 July 1995 (1 pair, AMNH 308088); FK-035, Indian Key Fill, 24°53’25"М, 80°40’28"W, bayside, Thalassia seagrass bed, 1 m, shovel/ sieve, 10 March 1996 (1 juv valve, AMNH 296528), FK-039, Crawl Key, 24°44’36"М, 80°58'47"W, oceanside, beach inside channel, shallow sand, seagrass, seawall, by hand and hand dredge, 12 March 1996 (1 зрт alc, FMNH 301424); FK-047, Channel marker 50A off Ramrod Key, 24°35.80’N, 81°27.24’W, rubble and patch reef, 15 ft (4.6 m), scuba, M/V THE SNAIL, 21 September 1996 (1 pair, 1 valve, AMNH 295179); FK-068, bayside of West Summerland Key (Spanish Harbor Keys), 24°39’19"М, 81°18'13"W, bayside, shovel/ sieve in Thalassia + beach combing, 0.5-1 m, 18 April 1997 (3 valves, 1 frag, FMNH 279531); FK-069, Channel marker 50A off Ramrod Key, 24°35.80’М, 81°27.24’W, rubble and patch reef, 18 ft (5.5 m), scuba, R/V FLORIDAYS, 19 April 1997 (1 pair, FMNH 279521); FK-090, Garden Key, Dry Tortugas, 24°37’50"N, 8252’20"W, sand beach with stone pilings, 2.5 m, by hand and snorkeling, 23 April 1997 (1 pair, AMNH 296520), FK-091, Fort Jefferson, Garden Key, Dry Tortugas, 24°37’50"М, 82°52’24"W, beach at mote and mote wall, by hand and snorkel- ing, 23 April 1997 (2 pair, AMNH 290122); FK- 115, East Washerwoman Shoal (Channel Marker 49), off Marathon, 24°40’N, 8104.3’W, 9 ft (2.7 m), scuba, R/V FLORIDAYS, 12 July 1997 (1 valve, FMNH 279523); FK-117, Key Vaca, channel west of Stirrup Key, 24°44.19’N, 81°02.92’W, bayside, in channel, 15 ft (4.6 т), 452 BIELER ET AL. plus in surrounding shallow Thalassia/sand, scuba, 14 July 1997 (1 spm alc [photographed alive], FMNH 295706); FK-119, “The Slabs” patch reef between “outer patches” and “coral humps’ off Marathon, 24°39.53’N, 81°00.90’W, scuba, 23 ft (7.0 т), R/V FLORIDAYS, 20 July 1997 (1 valve, FMNH 279528); FK-121, “The Slabs” patch reef between “outer patches” and “coral humps” off Marathon, 24°39.53’N, 8100.90'W, 23 ft (7.0 m), scuba, R/V FLORIDAYS, 21 July 1997 (1 pair, 3 valves, AMNH 296525); FK-131, off Key Vaca, oceanside, “outer patches” south of Hawk Channel, 24°39.30’М, 81°01.30’W, rubble, gorgonians, sponges, 21 ft (6.4 m), scuba, R/ V FLORIDAYS, 07 August 1997 (3 valves, AMNH 296530); FK-135, east of Bethel Bank, Florida Bay, 2443.86'N, 81°07.41’W to 24°43.60’М, 81°07.47’W, sand/sparse sea- grass, 8 ft (2.4 m), dredge, R/V FLORIDAYS, 07 August 1997 (1 valve, FMNH 279525); FK- 149, exposed flats outside channel off Crawl Key, 24°44’29"М, 80°58’24"W, oceanside, clean sand + Thalassia/Syringodeum sea- grass, Penicillus, 0.5 ft (0.15 m), by hand, R/V FLORIDAYS, 22 August 1997 (2 valves, FMNH 279530); FK-169, Tavernier Creek, near bayside entrance, west side (Plantation Key), 25°00.76’М, 80°32.68’W, sand/Thalassia, 6- 8 ft (1.8-2.4 m), ponar grab and dredge, R/V FLORIDAYS, 17 September 1998 (1 valve, AMNH 296522); FK-171, just off mouth of Tavernier Creek, oceanside, near marker #7, 24°59.67’М, 80°31.72’W, sand, Thalassia/ Syringodeum seagrass, 0-3 ft (0-0.9 m), snor- keling/sieving, R/V FLORIDAYS, 17 Septem- рег 1998 (observed pair); FK-178, east end of Rodriguez Key, oceanside of Key Largo, 25°03.13’М, 80°26.49’W, Thalassia seagrass, sand, small rubble, 2-3 ft (0.6-0.9 m), snor- keling, R/V FLORIDAYS, 20 September 1998 (2 pair, 1 valve, AMNH 295178); FK-179, Lower Matecumbe Key, 24°51’24"М, 80°43’40"W, oceanside, from beach front 3 days after Hurricane Georges, in Thalassia droves (1-3 ft deep) washed ashore by storm, 28 September 1998 (1 valve, FMNH 279529); FK=205, Carysfort Веер . 25-.18.25N: 80°12.78’W, coral rubble, sand patches, coral heads, 6-15 ft (1.8-4.6 m), scuba, R/V FLORIDAYS, 10 April 1999 (1 spm alc, AMNH 298893); FK-207, east end of Rodriguez Key, 25°03.13’М, 80°26.49'W, sand, seagrass, rubble, 1-2 m, snorkeling, R/V FLORIDAYS, 11 April 1999 (3 pair, AMNH 296519); FK-228, Old Dan Bank, bayside off Long Key, 24°49.66’М, 80°50.18’W, Thalassia/Porites/ Halimeda, 2-4 ft (0.6-1.2 m), snorkeling, R/V FLORIDAYS, 31 July 1999 (2 pair, AMNH 296521); FK-233, Old Dan Bank, bayside of Long Key, north of marker 2X, 24°49’57"М, 80°49'45"W, Thalassia seagrass, Halimeda, Porites, 2-4 ft (0.6-1.2 т), snorkeling/sieving, R/V FLORIDAYS, 01 August 1999 (2 pair, AMNH 296527); FK-236, Coffins Patch, oceanside of Grassy Key, 24°41’05"М, 80°57'28"W, algae-covered coral reef, 16 ft (4.9 m), scuba, R/V FLORIDAYS, 02 August 1999 (3 valves, FMNH 279522); FK-244, bayside of West Summerland Key (Spanish Harbor Keys), 24°39’19"М, 81°18’13"W, bayside, rock wall and sand slope, 23 ft (7.0 m), scuba, 05 August 1999 (1 valve, AMNH 296524); FK-246, bayside of West Summerland Key (Spanish Harbor Keys), along inner shore of western arm of horse- shoe, 24°39'19"N, 81°18'13"W, beach, 05 August 1999 (1 frag, AMNH 296523); FK-255, Friend Key Bank, bayside of Bahia Honda Key, north side at crest of bank, 24°42.58’М, 81°16.77'W, Thalassia/Syringodeum sea- grass, sand patches, 0.5-2 ft (0.15-0.6 т), snorkeling, R/V FLORIDAYS, 09 August 1999 (3 valves, FMNH 279520); FK-260, Looe Key coral reef, oceanside of Ramrod Key, 24°32.80’М, 81°24.80'W, spur and groove reef, 24-25ft(7.3-7.6 m), scuba, R/V FLORIDAYS, 10 August 1999 (1 valve, FMNH 279526); FK- 268, Looe Key back reef, 24°32.79’М, 81°24.33’W, seagrass, sand patches, rubble, 2-8 Н (0.6-2.4 т), snorkeling, R/V FLORI- DAYS, 19 August 1999 (1 valve, FMNH 279527); FK-269, Looe Key back reef, 24°32.79’М, 81°24.33’W, sediment sample, 8 ft (2.4 m), snorkeling, R/V FLORIDAYS (1 juv pair, AMNH 296531); FK-273, bayside of West Summerland Key (Spanish Harbor Keys), at outermost point of western arm of horseshoe, 24°39.35’М, 81°18.22’W, 1-4 ft (0.3-1.2 m), snorkeling, hand collecting, 19 August 1999 (3 spm alc [1 dissected, 1 with pearls, 1 SEM, AMNH 295199; 1 partial animal alc [body of spm with pearls, see prior; DNA], FMNH 296696; 1 juv pair, AMNH 296526); FK-275, Looe Key back reef, 24°32.87’М, 81°24.41’W, 5-10 ft (1.5-3.0 т), snorkeling and hand-col- lecting, R/V FLORIDAYS, 20 August 1999 (1 valve, FMNH 279524); FK-287, bayside of West Summerland Key (Spanish Harbor Keys), inside outermost arm of horseshoe, 24°39.35’N, 81°18.22’W, shallow subtidal, by hand, snorkeling, 10 April 2000 (1 spm alc, FMNH 289982; 1 live-collected pair, FMNH 301426; 9 pair, FMNH 289939); FK-298, Looe PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 453 Key National Marine Sanctuary, southwest, ca. 24°32.5’М, 81°24.7’W, 30 ft (9.1 m), scuba, R/ V EUGENIE CLARK, 02 July 2000 (1 juv pair; АММН 308080); FK-350, Looe Key National Marine Sanctuary, southwest corner of core area, 24°32.61’М, 81°24.66 W, 32 ft (9.7 m), scuba, R/V EUGENIE CLARK, 07 July 2000 (1 valve, 1 juv valve, AMNH 299545); FK-351, Looe Key back reef, 24°32.87’N, 81°24.41’W, rubble, 3-7 ft (0.9-2.1 т), snorkeling, R/V FLORIDAYS, 08 July 2000 (1 pair, AMNH 299489); FK-352, bayside of West Summerland Key (Spanish Harbor Keys), south arm, 24°39’19"М, 81°18'13"W, bayside, rubble, to 1.5 m, snorkeling, 08 July 2000 (3 spm alc [dissected], FMNH 283534; 6 juv pair, 1 valve, 2 juv valves, AMNH 299467); FK-357, American Shoals, northwest of lighthouse, 2431.54’М, 81°31.26 W, Thalassia seagrass with large coral rubble, 9-11 ft (2.7-3.3 т), scuba, R/V FLORIDAYS, 09 July 2000 (3 spm alc [1 dissected], FMNH 301428); 1 frag, АММН 299581); FK-359, American Shoals, 24°31.56'М, 81°31.10’W, Thalassia/ Syringodeum seagrass with rubble, rocks, 11— 12 ft (3.3-3.6 т), scuba, R/V FLORIDAYS, 10 July 2000 (observed spm; 2 pair, 2 valves, AMNH 299421); FK-360, coral lumps off Newfound Harbor Keys, off Big Munson Key, 24°36.96’N, 81°23.64’W, sand, seagrass, patch reef, gorgonians, 9 ft (2.7 m), scuba, R/ V FLORIDAYS, 11 July 2000 (1 pair, AMNH 299520); FK-363, east end of Rodriguez Key, oceanside of Key Largo, 25°03.27’М, 80°26.66'W, Thalassia seagrass, sand, small rubble, 6 ft (1.8 m), snorkeling, R/V FLORIDAYS, 07 October 2000 (2 pair, AMNH 307612); FK-364, Dove Key (just southwest of Rodriguez Key), oceanside of Key Largo, 25°02.94’М, 80°28.27’W, sand, algae, sponges, Sargassum, 2-6 ft (0.6-1.8 m), snor- keling, R/V FLORIDAYS, 07 October 2000 (1 pair, AMNH 307613); FK-367, northeast cor- ner of Conch Reef, oceanside of Key Largo, 24°57.895’N, 80°27.248’W, rubble, Thalassia seagrass, 9 ft (2.7 m), snorkeling, R/V FLORIDAYS, 08 October 2000 (1 frag, AMNH 307736); FK-392, Lower Matecumbe Key, 24°51'24"N, 80°43’40"W, oceanside, by hand in wrack line, 20 October 2000 (9 valves, FMNH 301429); FK-395, Snappers Ledge reef, 24°58.88’N, 80°25.36’W, in cavity of large sponge with red algae, 26 ft (7.9 m), scuba, M/S REPUBLIC IV, 27 March 2001 (1 pair, FMNH 301430); FK-459, Ft. Zacchary Taylor State Park, Key West, beach, shells washed ashore among rubble, 02 May 2001 (1 frag, AMNH 307737); FK-463, American Shoals, 24°31.541'М, 61°33: 218° WW. 9 ft (lS mi); Thalassia/Syringodium seagrass and rubble, scuba, R/V FLORIDAYS, 20 July 2001 (3 pair, 4 valves, 2 frag, FMNH 301431); FK-499, Sand Key, 24°27.18’М, 81°52.79’W, beach to 16 ft (4.9 m) under ship, sand, rocks, patch reef, snorkeling, R/V EUGENIE CLARK, 24 July 2001 (2 frag, AMNH 307738); FK-539, south of Bahia Honda Key, west of Looe Key reef, 24°34.24’М, 81°16.64’W, 30.2-34.1 m (99-112 ft), sand and rubble, pipe dredge and triangle dredge, R/V EUGENIE CLARK, 28 July 2001 (1 уу valve, AMNH 308089); FK-547, Looe Key reef, 24°32.809'N, 81°24.158’W, 25 ft (7.6 m), spur and groove reef, scuba, R/V FLORIDAYS, 30 July 2001 (2 valves, AMNH 307614); FK-559, south beach of Loggerhead Key, Dry Tortugas, 24°37.790’N, 82°55.400’W, wrack line to 7 ft (2.1 m), by hand and snor- keling, R/V CORAL REEF Il, 15 April 2002 (3 valves, FMNH 301432); FK-581, north shore of Loggerhead Key, Dry Tortugas, 24°37.871N, 82°55.447’W, wrack line and shallow subtidal, by hand and snorkeling, R/V CORAL REEF II, 16 April 2002 (5 valves, FMNH 301433); FK-601, Hospital Key, Dry Tortugas, 24°38.970’М, 82°51.284’W, patch reef, sand, 16 ft (4.9 т), scuba and snorkel- ing, R/V CORAL REEF ll, 18 April 2002 (1 valve, 3 frag, AMNH 307615; 1 pair, FMNH 296718); FK-606, southwest of Dry Tortugas, 24°30.009’М, 82%59.914'W to 24°29.970'N, 82°59.687’W, 28 m, triangle dredge, R/V CORAL REEF Il, 18 April 2002 (1 frag, AMNH 307616); FK-615, Cosgrove Shoal, 24°27.486'N, 82°11.039'W, 9.7 т (32 ft), patchy rubbly reef with sponges, gorgonians, overhangs, sand flat, scuba, R/V CORAL REEF ||, 20 April 2002 (2 pair, 3 valves, AMNH 307617); FK-620, Old Dan Bank, bayside of Long Key, 24°50.45’М, 80°49.63’W, Thalassia seagrass with Halimeda, Porites, sponges, hydroids, patches of sand/Halimeda hash, 1- 2 ft (0.3-0.6 т), by hand, R/V FLORIDAYS, 16 and 18 July 2002 (2 valves, FMNH 301445); FK-622, directly off Keys Marine Laboratory, bayside of Long Key, 24°49.5’N, 80°48.9'W, seagrass bed with coral rubble, snorkeling, sieving, by hand, 0-1.5 m, 20 July 2002 (valves observed); FK-624, Horseshoe Reef, off Fat Deer Key, 24°39.91’N, 80°59.56’W, patch reef with sandy bottom, 24 ft (7.3 m), scuba, M/V SHUTTERBUG II, 20 July 2002 (3 valves, FMNH 301446); FK-625, Coffins Patch Sanctuary Preservation Area, off Crawl Key, 24°40.92’М, 80°58.26’W, patch reef with 454 BIELER ET AL. sand patches, gorgonian, pillar coral, 21 ft (6.4 т), scuba, M/V SHUTTERBUG ll, 20 July 2002 (5 valves, FMNH 301447); FK-629, bayside of West Summerland Key (Spanish Harbor Keys), 24°39.3’N, 8118.2'W, among rocks along arms of quarry, to ca. 1 m, by hand, snorkeling, 21 and 26 July 2002 (1 spm alc [siphons photographed in sand, dissected], FMNH 301448; 5 pair, 4 valves, 2 juv valves [incl. photo voucher], FMNH 296719); FK-639, Coral Gardens inshore patch reef, oceanside of Lower Matecumbe Key, 24°50.23’М, 80°43.77’W, snorkeling, 1215 ft (3.6-4.6 т), Keys Marine Laboratory boat, 23 July 2002 (valves observed); FK-647, west side of Pi- geon Key, 24°42.2’М, 81°09.3'W, Thalassia/ Halodule/Syringodeum seagrass on sand/ rubble, concrete bridge piers, 0.5-1 m, by hand, snorkeling, shovel/sieving (1 pair, NTM); FK-649, Sprigger Bank, bayside, just W of Everglades National Park border, 24°54.75’N, 80°56.24’W, Thalassia/ Syringodeum seagrass, 1-3 ft (0.1-0.9 т), snorkeling, shovel/sieving, Keys Marine Labo- ratory boat, 27 July 2002 (2 pair, 1 valve, FMNH 301449); FK-659, Pigeon Key, 24°42.2’М, 81°09.3'W, seagrass, scuba, 2-4 ft (0.6-1.2 т), 28 July 2002 (valves observed); FK-660, Old Dan Bank, bayside of Long Key, 24°50.08’М, 80°49.63’W, Thalassia seagrass with Halimeda, Porites, sponges, hydroids, patches of sand/Halimeda hash, 1-5 ft (0.3- 1.5 т), snorkeling, R/V LAST MANGO, 28 July 2002 (2 pair, 5 valves, 3 frag, FMNH 301450); FK-661, Molasses Keys, south of center of Seven-Mile Bridge, north of westernmost is- land, 24°41.070’М, 81°11.483’W, sandy bot- tom, coral rubble, Thalassia seagrass, hot water (> 30°C), snorkeling, 1-6 ft (0.3-1.8 т), R/V FLORIDAYS, 04 August 2002 (5 valves [2 with breakage by predator], FMNH 301434); FK-662, Sombrero Reef, vicinity of buoy SO- 3, 5 nmi south of Knights Key, 24°37.619’М, 81°06.528’W, sandy bottom adjacent to coral reef; scuba, 17—23. (52-7. mie Ri FLORIDAYS, 05 August 2002 (1 valve, 1 frag, FMNH 301435); FK-664, Molasses Keys, south of center of Seven-Mile Bridge, north of westernmost island, 24°41.070'N, 81°11.483’W, sandy bottom, coral rubble, Thalassia seagrass, strong current, snorkel- ing, 2-5 ft (0.6-1.5 т), R/V FLORIDAYS, 06 August 2002 (2 valves, FMNH 301436); FK- 665, Coffins Patch pillars, south of Crawl Key, 24°40.899’М, 80°58.246’W, coral reef, sand plains, some Thalassia seagrass, scuba, 18- 23 ft (5.5-7.0 т), R/V FLORIDAYS, 07 August 2002 (2 valves, FMNH 301437); FK-668, Money Key, oceanside of west end of Seven- Mile Bridge, off north and west ends of island, 24°41.009’М, 81°12.955’W, “ironshore” beach rock, sand, Thalassia seagrass, snorkeling, 0— 5 ft (0-1.5 т), R/V FLORIDAYS, 09 August 2002 (1 pair, FMNH 301444); FK-672, John Sawyer Bank, 3 nmi north of western part of Key Vaca, bayside, 24°45.498’N, 81°06.621'W, coral rubble, Thalassia seagrass, strong cur- rent over shoal, snorkeling, 2-6 ft (0.6-1.8 т); R/V FLORIDAYS, 11 August 2002 (1 pair, 2 valves, 1 fresh frag, FMNH 301438); FK-673, Bethel Bank, 2 nmi N of Knights Key Channel, 24°43.796'N, 81°07.588'W, bayside, coral rubble, Thalassia seagrass, strong current over shoal, 3-5 ft (0.9-1.5 т), snorkeling, R/V FLORIDAYS, 11 August 2002 (1 valve, FMNH 301439); FK-674, Sombrero Reef, southwest area of lighthouse reef, 24°37.555’М, 81°06.729’W, spur and groove reef, rubble, . scuba, 22-26 ft (6.7-7.9 т), R/V FLORIDAYS, 12 August 2002 (4 valves, 1 fresh frag, FMNH 301440); FK-675, Red Bay Bank, 3 nmi north of Pigeon Key, bayside, 24°45.1000’М, 81°08.647’W, very diverse habitat: coral, seagrass, many algae, snorkeling, 1-6 ft (0.3- 1.8 т), R/V FLORIDAYS, 13 August 2002 (2 spm, 2 valves, FMNH 301441); FK-676, Rachel Bank, 2 nmi north of Key Vaca, bayside, 24°44.714’N, 81°04.599’W, muddy sand, seagrass, coral rubble, snorkeling, 35 ft (0.9-1.5 т), R/V FLORIDAYS, 13 August 2002 (5 spm, 7 valves, 1 fresh hinged frag, FMNH 296695); FK-677, Doughnut Reef, oval reef mass east of Coffins Patch (off Crawl Key), 24°41.507'N, 80°56.835’W, sand flat at reef edge, scuba, 23 ft (7.0 т), M/V SHUTTERBUG |, 13 August 2002 (1 valve, FMNH 301442); FK-678, bayside of West Summerland Key (Spanish Harbor Keys), 24°39.3’М, 81°18.2’W, inside western arm, scuba, 3-10 ft (0.9-3.0 m), 14 August 2002 (1 spm alc [dissected], 4 valves, FMNH 301443); FK-688, Conch Reef, oceanside of Key Largo, 24°57.380'N, 80°29.428'W, algae-covered reef, vertical walls and adjacent sand plains, max. 28 ft (8.5 m), scuba, R/V FLORIDAYS, 05 June 2003 (1 V, FMNH 302086; 2 LV, AMNH 308081); FK- 689, northeast of Dove Key, oceanside of Key Largo, 25°03.055’N, 80°28.220’W, hard bot- tom with silty sand, sponges, gorgonians, 0.5— 1.0 m, snorkeling, R/V FLORIDAYS, 06 June 2003 (1 juv pair, AMNH 308084); FK-692, Hen and Chickens patch reef, oceanside of Plan- tation Key, 24°56.099’N, 80°32.920’W, тах. 21 ft (6.4 m), scuba and snorkeling, R/V PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 455 FLORIDAYS, 08 June 2003 (1 RV, 1 LV, AMNH 308087); FK-693, off Dove Key, oceanside of Key Largo, 25°03.039’М, 80°28.151’W, silty Thalassia seagrass and sand, 3-5 ft (0.9-1.5 m), snorkeling, R/V FLORIDAYS, 08 June 2003 (3 pair, 1 RV, 1 LV, AMNH 308083); FK- 696, Sand Island (near Molasses Reef), 25°01.116’М, 80°22.046’W, patch reef with rubble, max. 22 ft (6.7 m), scuba, R/V FLORIDAYS, 10 June 2003 (1 pair, AMNH 308085); FK-698, Wolfe mooring buoy (near Three Sisters Reef), 25°01.311'N, 80%23.774'W, patch reef with adjacent Thalassia seagrass, max. 16 ft (4.9 т), scuba, R/V FLORIDAYS, 11 June 2003 (2 RV, 1 frag- ment, AMNH 308082); FK-701, off Dove Key, oceanside of Key Largo, 25°03.011'N, 80°28.163 W, silty Thalassia seagrass and sand, 1-2 ft (0.3-0.6 m), snorkeling, R/V FLORIDAYS, 12 June 2003 (5 pair, 1 RV, 1 LV, AMNH 308086). FK-702, seagrass flat off Yellowtail Inn, oceanside of Grassy Key, mile marker 58.3, 24°45.494’N, 80°57.179'W, 1-5 ft (0.3-1.5 m), snorkeling, 5-23 August 2003 (1 pair, 1 LV, FMNH 302071); FK-703, patch reef with sand pockets in vicinity of mooring buoy near “the Stake”, Coffins Patch coral reef, oceanside of Grassy Key, 24°41.159’М, 80°57.836'W, 8-18 ft (2.4-5.5 m), snorkeling, R/V FLORIDAYS, 7 August 2003 (1 pair, 1 RV, - FMNH 302079); FK-704, Sombrero Reef, vi- cinity of buoy SO-4, 5 nmi S of Knights Key, 2437.591'N, 81°06.563’W, sandy bottom im- mediately adjacent to spur & groove coral reef; scuba, 17-25 ft (5.2-7.6 m); R/V FLORIDAYS, 08 August 2003 (1 LV, FMNH 302072); FK- 705, Doughnut Reef, oval reef mass E of Cof- fins Patch (off Crawl Key); 24°41.439’N, 80°56.862’W, patch reef surrounded by sand flats, scuba, 24 ft (7.3 m); M/S SEAFARI, 09 August 2003 (1 LV, FMNH 302073); FK-706, Elbow Reef, E of Coffins Patch (off Crawl Key); 24%41.551'N, 80°56.789’W, patch reef sur- rounded by sand flats, scuba, 19 ft (5.8 m); M/S SEAFARI, 09 August 2003 (1 RV, FMNH 302074); FK-710, “Porkfish” and “Hammer Ledge” reefs, E of Coffins Patch (off Conch Key); 24°42.038’М, 80°53.578’W, scuba, 19- 25 ft (5.8-7.6 т); M/S SEAFARI, 11 August 2003 (2 RV, 2 frag, FMNH 302075); FK-711, patch reef with sand pockets in vicinity of moor- ing buoy 9, Coffins Patch coral reef, oceanside of Grassy Key, 24°41.131’N, 80°57.818'W, 18— 20 ft (5.5-6.1 m), scuba; R/V FLORIDAYS, 12 August 2003 (1 LV, FMNH 302076); FK-714, vicinity of mooring buoy 1 at “Marker 48” patch sand and seagrass; 24°41.505’N, 81°01.528’W, 16-24 ft (4.9-7.3 т), scuba; R/V FLORIDAYS, 17 August 2003 (1 pair, 5 LV, FMNH 302077); FK-719, beach near moat wall, Fort Jefferson, Garden Key, Dry Tortugas, 24°37’50"N, 82°52’24"W, by hand, 21 August 2003 (1 RV, FMNH 302078); Other Material Examined Florida: Boynton, Lake Worth, T. L. McGinty! ANSP 195919 (1 pair); Lake Worth Inlet, C. T. Simpson! UMML 28.1729 (1 pair); South Lake Worth near Boynton, Kline! July 1944, DMNH 72175 (1 pair); Lake Worth, Lermond! August 1941, DMNH 153846 (3 pair); Lake Worth, south end, dredged in 6 ft (1.8 m), Lermond! 14 September 1937, DMNH 153848 (2 pair); Lake Worth, south end, Doremus! 1942, DMNH 63779 (2 pair); Lake Worth, south end, Е. Lyman family! Summer 1944, FMNH 144097 (1 pair); Lake Worth, South Inlet, oyster reef, F. M. Bayer! USNM 890543 (1 pair with tissue); Boynton in Lake Worth, Е Fal, sand pocket in rock reef, Т. L. McGinty! June 1940, USNM 599285 (1 pair); Lake Worth, White collection, USNM 153360 (1 pair); Lake Worth, Singer Bridge, F. M. Bayer! USNM 890617 (2 juv pair); opposite Lemon City, C. T. Simpson! UMML 28.1731 (1 LV); Bal Harbor, Broad Causeway, dredgings, Poh! June 1975, DMNH 118270 (1 pair, 1 valve); Coral Gables, FMNH 54675 (1 pair); off Cape Florida, Florida, Finger Channel flats, Crovo! 05 April 1970, DMNH 30152 (1 pair); Cape Florida, Biscayne Bay, Т. Е. McGinty! 1937, ANSP 264016 (1 pair); Miami Bay area, flats off Cape Florida, Ingalls Family! 12 March 1967, AMNH 140432 (1 pair); Miami area, Biscayne Bay, flats off Cape Florida, W. E. Old! 12 March 1967, AMNH 136053 (2 pair); Cape Florida, T. L. McGinty! FMNH 26427 (2 valves); Cape Florida, near Miami, T. L. McGinty! June 1936, USNM 599308 (1 pair); Bear Cut, Key Biscayne, Miami, W. S. Bitler! 1963, AMNH 142323 (1 pair); Bear Cut, Key Biscayne, S. Sokoloff! 18 November 1961, AMNH 261423 (1 pair); Bear Cut, April 1939, UMML 28.45 (1 pair); Bear Cut, Hepler! DMNH 45331 (2 pair); Caesar Creek, Whitney! November 1956, DMNH 97311 (1 pair); Florida Keys, Lermond! DMNH 153830 (1 pair); Angelfish Creek [north of Key Largo, connecting Card Sound and Atlantic Ocean], Wisoff Collec- tion, AMNH 120395 (1 pair); Key Largo, Nelson collection, FMNH 155544 (1 pair); Key Largo, T. L. Moise! August 1950, ANSP 456 193815 (1 juv pair); Key Largo, F. M. Bayer! June 1940, USNM 890827 (1 pair); Windley Key, Whale Harbor Channel, near bridge in 5-10 ft (1.5-3.0 т), Т. В. Waller! Sta. 3, 31 August 1971, USNM 707750 (1 valve); Up- per Matecumbe Key, Islamorada, on beach, 13 January 1978, BMSM 26108 (5 pair); In- dian Key, D. V. Stingley! May 1959, BMSM 26109 (1 pair); Lower Matecumbe Key, Hausman! AMNH 133675 (1 pair); Conch Key, south end, J. J. Parodiz 8 Winters! 08 July 1976, CMNH 43728 (4 pair); Grassy Key, М. & $. Snyder! July 1966, ANSP 309750 (2 pair); Grassy Key, Minzak! February 1972, DMNH 93337 (3 pair); Crawl Key, 24°44’36"N, 80°58'47"W, beach, P. M. Mikkelsen 8 R. Bieler! 22 September 1996, AMNH 295180 (1 pair); Crawl Key, Richardson! DMNH 85844 (1 pair); Crawl Key, shallow water, J. M. Bijur! May 1964, АММН 248309 (1 pair); Crawl Key, bayside, on flats at 1 ft (0.3 m), Raeihle! November 1961, AMNH 106142 (1 pair, figured Emerson 8 Jacobson, 1976: pl. 42, fig. 18); Crawl Key, bayside, on beach, D. Raeihle! November 1973, AMNH 179278 (1 pair); Crawl Key, bayside, D. Raeihle! AMNH 116658 (1 pair); Crawl Key, bayside, November 1959, Raeihle! AMNH 307848 (3 pair, 1 valve, live- collected); Crawl Key, С. Dingerkus & (. D. Uhler! 06 January 1977, AMNH 267424 (1 spm alc); Crawl Key, bayside, edge of bor- row pit Y mi north of mile marker 56, in weeds, M. J. de Maintenon! 24 June 1985, AMNH 307571 (1 pair); Bonefish Key, A. Koto! AMNH 133664 (1 pair) and FMNH 176293 (1 pair, 1 valve); Marathon Key, Florida Straits, K. C. Vaught Collection, AMNH 250783 (1 pair); Marathon, 1959, BMSM 26110 (2 pair); Marathon Key, south end, Jensen! 1962, DMNH 42547 (1 pair); Sombrero Reef, 25-30 ft (7.6-9.1 m), P. S. Mikkelsen! 21 May 1980, DMNH 180066 (2 valves); Washerwoman's patch reef, 24°39'54"N, 81°04'14"W, M. Snyder! August 1966, ANSP 398069 (1 pair); Pigeon Key, 24°42'N, 81°09’W, M. Snyder! July 1966, ANSP 398068 (2 pair); Little Duck Key, A. Koto! 1955, FMNH 176330 (1 pair) and FMNH 176372 (1 pair [photo voucher]); Little Duck Key, shallow water, sand, F. Schilling! July 1968, FMNH 288716 (1 pair); Missouri Key, Richardson! DMNH 85835 (1 pair), Mis- souri Key, snorkeling, A. D. Barlow! 05 March 1967, AMNH 243914 (2 pair); Missouri Key, sand, F. Schilling! 16 July 1970, FMNH 288718 (1 pair); Bahia Honda State Park, BIELER ET AL. Germer Collection, 11 July 1973, AMNH 269541 (1 pair); north of Bahia Honda State Park, shallow water, sand, R. Bieler 8 P. M. Mikkelsen! 25 March 1989, FMNH 288810 (1 spm alc); Bahia Honda Key, BMSM 26111 (1 pair); Spanish Harbor Key, beach, Piech! July 1980, DMNH143838 (1 pair); West Summerland Key, at entrance to dredge hole, L. Scheu Collection, 1984, AMNH 230097 (2 pair); Newfound Harbor Keys, living in shal- low water, sand, November 1968, Raeihle! AMNH 307849 (1 pair); Torch Key channel, C. T. Simpson! UMML 28.1716 (4 pair); Sugar- loaf Key, J. В. Clark! 23-24 Мау 1921, ANSP 9634 (1 juv pair); Boca Chica Key, H. A. Pilsbry! ANSP 100273 (1 valve); Stock Island, Е. В. Kirtland! 1936, ANSP 167777 (1 pair); Key West, A. Koto! FMNH 176402 (1 pair); Key West, Nelson! FMNH 166745 (1 pair); Key West, coral banks at low tide, J. W. Milner! USNM 127385 (1 pair); Key West, Hawk . Channel, 3-20 ft (0.9-6.1 m), Eolis sta. 65, J. B. Henderson Jr.! 15 May 1913, USNM 448344 (2 juv valves); Key West, reefs, rare, H. Hemphill! USNM 95672 (2 pair); Key West, Smith Shoals, Eolis sta. 335, J. B. Henderson Jr.! 1916, USNM 448343 (1 juv pair); Key West, М side, beach collecting, Eolis sta. 35, J. В. Henderson Jr.! 30 May (or 6 June) 1911, USNM 48340 (1 juv pair); Key West, USNM 406825 (2 pair); Key West, H. Hemphill! ANSP 52199 (2 pair); Key West, H. A. Pilsbry! March- April 1940, ANSP 175943 (1 pair); Key West, Sand Point, B. R. Bales! 1946, DMNH 21225 (2 pair) and ANSP 285406 (6 pair); Key West, at low tide, 3—4 ft (0.9-1.2 т), Е. Schilling! 21 June 1967, FMNH 288719 (1 pair); Key West, С. T. Simpson! UMML 28.1748 (1 pair); Sand Key, Key West, Ostheimer! DMNH110654 (3 pair); near Boca Grande Key, UMML 28.1740, С. Т. Simpson! (1 pair); Tortugas, Stm.! USNM 36403 (4 pair, 1 valve); Dry Tortugas, off Log- gerhead Key, north end, beach collecting, Eolis sta. 367, J. B. Henderson Jr.! 13 June 1911, USNM 448347 (1 valve); Dry Tortugas, Garden Key, 3 mi out from red sea buoy, 5 dredge hauls, 14—15 fms (25.6-27.4 т), Eolis sta. 34, J. B. Henderson Jr.! 09 June 1911, USNM 448341 (4 pair, 1 valve); Tortugas, Bush Key, F. M. Bayer! USNM 890776 (1 pair); off Carabelle [Franklin County, panhandle of Florida], 29°15’М, 84°40'W, 90-100 ft (27.4— 30.5 m), dredged, J Moore! ex M. & B. Naide, August 1966, ANSP 402130 (1 pair). Texas: off Freeport, 28°13’N, 94°51’W, 27 ft (8.2 m), dredge, A. Kight! ANSP 338392 (1 pair). PERIGLYPTA LISTERI IN THE WESTERN ATLANTIC 457 Bahamas: Bahamas, J. В. Henderson Jr.! USNM 448342 (2 pair); PMM-1047 (wp277- R rubble), off Andros, 24°54’44.42"М, 77°53'51.80"W, rubble in back reef pave- ment zone with gorgonians, 7 ft (2.1 m), scuba, P. M. Mikkelsen, et al.! 29 August 2000, AMNH 305616 (1 valve); PMM-1090 (wp415-R), off Andros, 24°53’32.2"М, 77°53'51.4"W, thick Thalassia seagrass, 12 ft (3.6 m), scuba/snorkeling, P. M. Mikkelsen, et al.! 04 September 2000, AMNH 305617 (1 pair), ЕММН 301425 (1 spm alc [95%]); PMM-1079 (wp427-R), off Andros, 24°55'24.8"N, 77°55'19.8"W, sand/algal plain, 5 ft (1.5 m), scuba/snorkeling, P. M. Mikkelsen, et al.! 02 September 2000, FMNH 296720 (1 pair); PMM-1063 (BH-R seagrass), oceanic blue hole off Blue Hole Фау Off Andros, 24°53’55.2"N, 77°55'12.1"W, Thalassia seagrass, 4 ft (1.2 m), scuba/snorkeling, P. М. Mikkelsen, et al.! 31 August 2000, AMNH 307572 (1 pair); East Andros Island, Calabash Bay, Abbott! Feb- ruary 1971, DMNH 29242 (1 pair); East Andros Island, Small Hope Bay, Abbott! DMNH 41253, March 1971 (1 valve); Chub Cay, Berry Islands, Moise! ANSP 193106 (1 pair); Chub Cay, Periwinkle Beach, K. C. Vaught Collection, April 1977, AMNH 250782 (1 pair); Grand Bahama Island, 26°31’N, 78°46 30"W, J. М. Worsfold! ANSP 375213 (2 pair); Grand Bahama Island, 26°31’00"N, 78°46'30"W, J. М. Worsfold! ANSP 375212 (1 juv valve); Grand Bahama Island, Run- ning Mon Canal, 26°29’45"N, 78°41’45"W, J. М. Worsfold! ANSP 369788 (1 pair); Grand Bahama Island, C. C. Allen! 1922-1923, ANSP 133697 (1 pair); Grand Bahama is- land, Bottle Bay Canal, 26°39’30"М, 78°57'00"W, sediment, 5 ft (1.5 т), J. Worsfold! ANSP 371908 (1 pair); east end of Grand Bahama Island, Deep Water Cay, ca. 2.5 mi northwest of Sweetings Cay Light, intertidal sand and rocks, V. O. Maes! Janu- ary 1965, ANSP 307688 (1 juv pair); Grand Bahama Island, F. H. Low Collection, AMNH 113790 (1 pair); Nassau, New Providence Island, Wards! (before 1893), FMNH 2741 (1 pair, 1 valve); Nassau, New Providence Island, Pope! FMNH 187147 (2 valves); Nassau, C. C. Allen! USNM 36617 (1 pair); New Providence Island, Dicks Point, McLean & Russell! July 1936, ANSP 169917 (2 valves); New Providence [Island], Lyford Cay, AMNH 80763 (2 pair); north coast of Hog Island, north of New Providence Island, R. Robertson! 11 September 1955, ANSP 299657 (1 juv pair); Great Abaco Island and Green Turtle Key, Abaco Islands, Cherokee flats, Great Abaco and Mendelson’s flats, Heilman! DMNH 37728 (3 pair); Great Abaco Island and Green Turtle Cay, Cherokee flats, DMNH 37977 (5 pair); Great Abaco, Mendelson’s flats, Heilman! February 1958, DMNH 86223 (5 pair, 2 valves); Great Abaco, Parrot Cays, west of Elbow [Little Guana] Cay, near octopus hole, R. Robertson! 14 August 1953, ANSP 299066 (3 pair); Great Abaco, west coast of north end of Elbow [Little Guana] Cay, mud/sand, Halimeda re- mains, Thalassia seagrass, 0.5-3 ft (0.15- 0.9 т) and near octopus hole, К. Robertson! 04 September 1953, ANSP 298847 (1 pair, 1 juv pair); Abaco Island, Green Turtle Cay, Gwillim Bay, on sand on exposed sand bar at low tide, A. & A. Taxson! 10 June 1964, AMNH 111921 (4 pair) and AMNH 269540 (2 pair, ex D. Germer Collection); Abaco, Crab Cay, 2-4 ft (0.6-1.2 т), Е. |. Wright! 1974, USNM 846377 (1 pair); Abaco, Marsh Harbor, O. Bryant! USNM 180541 (1 pair); east-central Eleuthera, north end of Half Sound, 25°07'45"М, 76°09: 00) Wy В. Robertson! 18 April 1984, ANSP 359292 (2 pair); Eleuthera, Savannah Island, Santy Point, W. J. Clench! May 1936, ANSP 173811 (3 pair); Eleuthera, Savannah Sound, Sandy Point, Cora Staples Collec- tion, AMNH 306250 (1 pair); Eleuthera, Cur- rent, Current Club, A. Ross! 29 July 1963, AMNH 100174 (1 valve); Eleuthera Island, Doremus! DMNH 63778 (2 pair); Eleuthera Island, Sandy Point, Savannah Sound, Doremus! May 1936, DMNH 63780 (2 pair); Harbour Island, N end Eleuthera Island, Loc.114, Kline! 17 June 1949, DMNH72173 (1 pair); Eleuthera, N end Half Sound, E cen- tral Eleuthera Island, Abbott! June 1976 DMNH 115333 (1 valve); Exuma Island, Rolle Town, Loc.16, Kline! 11 July 1951, DMNH 72174 (1 pair); southern Exuma Cays, north of Leaf Cay, R. Robertson! 07 July 1957, ANSP 285738 (1 pair); Bimini, near Bailey Town, Bimini Lagoon, R. Robertson! 1957-1958, ANSP 326271 (1 valve); South Bimini, east of Nixon’s Harbour, R. Robertson! 1957-1958, ANSP 325603 (1 pair); Bimini, 1.75 mi southeast of Orange Cay, 23 ft (7.0 т), К. Robertson! 1957-1958, ANSP 325698 (1 juv valve); Bimini, South Cat Cay, grassy, T. L. Moise! ANSP 193577 (1 pair); Bimini, around Risty Causeway, Pi- 458 BIELER ET AE. geon Cay, Steger! April 1956, DMNH 107635 (1 pair); San Salvador Island, W Pigeon Creek, beach, Piech! February 1977, DMNH 143251 (1 pair). Turks and Caicos: Providenciales, Water Cay, beach, Piech! February 1978, DMNH 144123 (1 pair). Cuba: east of Tarallones de Arena, near Santiago, sand beach, R. E. Dickerson! ANSP 182932 (3 valves); Paradise Island, Oriente, Christofferson! 17 April 1949, FMNH 144071 (4 valves); west of Guardalavaca, eastern shore near Playa Esmeralda, Prov- ince Holguin, K. & Ch. Schniebs! December 2001, 1 pair, MTD 43828; Guantanamo, E. O. Mitchell et al.! 1930, USNM 405334 (1 valve); Cayo Hutia Reef, Barrera Expedition sta. 218, USNM 448345 (2 pair); Esperanza, 2-3 fms (3.6-5.5 т), Barrera Expedition sta. 210, USNM 448346 (1 pair); Varadero Beach, Barrera Expedition sta. 213, USNM 448348 (1 juv valve). Cayman Islands: Grand Cayman Island, Gun Bay, near Blakes’, mud and turtlegrass flats, A. J. Ostheimer Ш! ANSP 199513 (1 pair); Grand Cayman, North Sound, Jensen! Au- gust 1970, DMNH 39561 (1 pair). Jamaica: Harboreale, near Annotta Bay, St. Mary, Orcutt! USNM 440717 (1 valve); Black River, St. Elizabeth, Orcutt! USNM 441413 (1 valve). Hispaniola: Haiti, off Port-au-Prince, southeast side of Grand Bans, east side of reef, G. Goodfriend! 25 June 1972, AMNH 177703 (1 pair); Haiti, Cape Haitien, American Haitien Dev. Company, Krieger! USNM 487861 (1 valve); Santo Domingo, Monte Christi, W. J. Clench et al.! July 1937, ANSP 173105 (2 valves); Santo Domingo, Monte Cristi, Doremus! July 1937, DMNH 63777 (2 pair). Puerto Rico: Puerto Rico, Stearns! USNM 54091 (1 pair); El Deseches Island, Mayaguez B., F. A. Gallardo! USNM 464243 (1 valve); Bahia Bramadero [south of Mayaguez], С. L. Warmke! November 1956, ANSP 222755 (1 valve); Puerto Rico, Richardson, DMNH-85845 (2 pair). U.S. Virgin Islands: St. Thomas, W. A. Haines Collection, pre-1895, AMNH 31868 (9 pair, including largest recorded specimen); St. Thomas, M. Petit! USNM 250151 (1 pair); St. Thomas, Petit collection, USNM 530502 (1 pair, 2 valves, 1 juv valve); St. Thomas, Lindberg Beach, D. M. Barringer! 1936, ANSP 166919 (1 valve); St. Thomas, Swift Collection, ANSP 53580 (5 pair, 2 juv pair); St. Thomas, ZMB-104283 (1 pair); St. Tho- mas, ZMB-104287 (1 pair). British Virgin Islands: Peter Island, Little Harbour, R. H. Pine! July 1976, FMNH 197453 (1 valve); Seal Dog Islands, R. H. Pine! August 1976, FMNH 197476 (1 pair); Tortola, east of Roadtown, H. G. Richards! ANSP 244999 (1 juv valve); Tortola, Kjaer! USNM 3208 (1 pair); Tortola, K. Lamprell! 25 September 1980, АММН 303476 (1 pair); Gorda Island, Colquhon Reef, Bredin- Smithsonian Expedition sta. 37-58, Schmitt et al.! 07 April 1958, USNM 735909 (1 pair). Antigua: Falmouth Harbor, beach, SUI Expe- dition, J. B. Henderson Jr.! 1918, USNM 500994 (1 pair). Grenada: South Grenada, Little Bacaye Har- bor, silt, Thalassia seagrass, sand patches, R. Ostheimer and Buerk! 23 January 1964, ANSP 297064 (1 pair). Mexico: Isla Mujeres, K. С. Vaught Collection, . AMNH 250781 (1 pair). Belize: east-southeast of Punta Negra, 16°16’15"М, 88°32’10"W, К. Robertson! 23 August 1961, ANSP 281836 (1 juv valve); north of Tarpon Cay, shallow Acropora cervicornis reef, 16°37’05"М, 88°09’05"W, К. Robertson! 17 August 1961, ANSP 282574 (1 valve); Glovers Reef, NE Cay, R. S. Houbrick! USNM 771205 (1 pair). Honduras: NW shore Bonacca Island, L. Kornicker! July 1963, USNM 667961 (4 valves). Costa Rica: Port Limon, Wailes! USNM 187276 (1 pair). Panama: Payardi Island, NW end, W. P. Woodring! 13 December 1959, USNM 67821 (1 valve); Payardi Island, NW end, 6 mi NE of Colon, suction dredge at refinery site, W. P. Woodring! 13 December 1959, USNM 637931 (1 valve); Payardi Island, Minas Bay, E of Colon, R. H. Stewart! USNM 734522 (3 valves). Colombia: Old Providence Island, Sid Ander- son! January—March 1966, AMNH 138019 (1 pair); San Andres Island, S. Anderson! Janu- ary 1966, AMNH 137541 (1 valve, 1 frag); vicinity of Cartagena, T. A. Link! USNM 364301 (3 valves); Cartagena, R. Pfaff! 1959, FMNH 78686 (1 pair). Netherlands Antilles: Aruba, M. R. Barnes! USNM 619369 (1 valve); Bonaire, Abbott! November 1972, DMNH 72707 (1 pair); Bonaire, Abbott! February 1973, DMNH 72794 (1 pair); Curagao, N. Dearborn! 1908, FMNH 12764 (1 valve, subfossil?). MALACOLOGIA, 2004, 46(2): 459-472 ANATOMY AND SYSTEMATICS OF NORTHWESTERN ATLANTIC DONAX (BIVALVIA, VENEROIDEA, DONACIDAE) Luiz Ricardo L. Simone? & Joanne К. Dougherty? ABSTRACT A morphological examination of two nominal species of northwestern Atlantic donacids, Donax fossor and D. variabilis, was performed to resolve current taxonomic discrepan- cies. Specimens from New Jersey, South Carolina, and Florida were studied confirming the typical anatomical bauplan for the family as previously reported. Detailed investigation of all organ systems revealed a series of differences, mainly in the shell, mantle border papillae, siphonal tentacles and papillae, and digestive system, supporting separation of the two species. Other shared morphological features, such as the gill muscle, pallial muscles of the siphonal chamber, the glandular dorsal gastric caecum, and length of the style sac, have potential value for further functional and systematic studies. Key words: Donax fossor, Donax variabilis, differentiation, distribution, western Altantic. INTRODUCTION The systematics of the genus Donax along the Atlantic coast of the United States has been problematic for the last three decades. The nomenclature of Donax variabilis Say, 1822, which occupies the intertidal zone of sandy beaches from Virginia to Mississippi, has been particularly confusing. Donax variabilis became a primary junior homonym when Latona variabilis Schumacher, 1817, was proposed as a new name for Donax cuneatus Linnaeus, 1758 (Morrison, 1971). Because Latona is considered a subgenus of Donax, the name D. variabilis Say is thus preoccupied. Morrison’s (1971) revision of the group identified the next available name for this species, D. protracta Conrad, 1849. Morrison considered D. protracta, from the southeastern coasts of the United States and eastern Gulf of Mexico, to be a subspecies of D. roemeri Philippi, 1849, from the northern and western Gulf of Mexico. Based on morphological differences between the two forms, Morrison designated the eastern forms as D. roemeri protacta Conrad, 1849, and the western forms as D. roemeri roemeri Philippi, 1849, because the publication date of D. roemeri was five months prior to that of D. protracta. Later, Boss (1970) proposed conservation of the name D. variabilis Say, 1822, to the International Commission on Zoological Nomenclature and it was subsequently conserved (Melville, 1976). In addition to his recognition of D. гоетеп roemeri and D. roemeri protacta, Morrison (1971) recognized four other species of Donax inhabiting the eastern shores of the United States: Donax fossor Say, 1822, from Cape Hatteras, North Carolina, to New Jersey and occasionally the southern shores of Long Island; D. parvula Philippi, 1849, from North Carolina to southern Florida; D. dorotheae Morrison, 1971, along the shores of the northeastern Gulf of Mexico; and D. texasianus Philippi, 1847, along the shores of Louisiana, Texas and Mexico. Subsequent analysis of RAPD DNA markers failed to support Morrison's distinction between the subspecies D. roemeri roemeri and D. roemeri protracta (Adamkewicz & Harasewych, 1996). The analysis also demonstrated that D. parvula was indistinguishable from D. fossor, and D. dorotheae was indistinguishable from D. texasianus, with the latter of each pair having taxonomic priority. That analysis simplified the biogeography of Donax; D. variabilis shares the Atlantic coast with D. fossor and the Gulf coast with D. texasianus. Even in older literature, taxonomic problems are notable. Say (1822) described two similar species of Donax, D. fossor (“the digger”), a northern form inhabiting the coasts of New Jersey and Maryland, and D. variabilis (“highly ¡Museu de Zoologia da Universidade de Säo Paulo, Cx. Postal 42594, 04299-970 Sao Paulo, SP, Brazil; Irsimone@usp.br “Department of Biology, Villanova University, Villanova, Pennsylvanica 19085, USA; Joanne.dougherty@villanova.edu 460 SIMONE & DOUGHERTY variable”), a southern form from the coasts of Georgia and eastern shores of Florida. Based on the original species descriptions, differen- tiating the two species is quite difficult. Size, color, sculpture, and thickness of the shell valves are most often used to differentiate the two species (Say, 1822; Chanley, 1969). Donax variabilis reaches a length of 19 mm, exhibits an wide variety of colors, and displays radial shell sculpture that is more pronounced on the posterior slope (Say, 1822; Chanley, 1969; Morrison, 1971). Donax fossor reaches 13 mm length, exhibits only “yellowish” or “whitish” colors, has smooth radial sculptur- ing over the entire shell and exhibits thickened valves at the anterior end to produce “lips” (Say, 1822; Chanley, 1969; Morrison, 1971). The escutcheon area of D. variabilis juveniles of about 5 mm in length is more rounded to- ward the vertical, whereas in D. fossor, the same is regularly sloping parallel to the rounded posterior ridge (Morrison, 1971). Donax fossor is believed to inhabit the surf and subtidal zones in the winter, while D. variabilis inhabits the intertidal zone through- out the year (Morrison, 1971). These two species are so similar that the populations of New Jersey have often been labeled D. variabilis (e.g., Johnson, 1927; A. E. Wood 8 H. E. Wood, 1927; McDermott, 1983; Alexander et al., 1993), which exacer- bates taxonomic confusion in the northern limit of Donax on the Atlantic coast of North America. Some authors accept the validity of these two species (e.g., Johnson, 1934; Mor- ris, 1947; Miner, 1950; Morrison, 1971; Abbott 8 Morris, 1995), while others suggest that the two are conspecific (Abbott, 1954, 1974; Chanley, 1969). For example, Chanley (1969) suggests that D. fossor is merely a summer range extension of D. variabilis, based on spo- radic populations of Donax on Long Island, New York, that do not overwinter. Chanley (1969) hypothesized that these northern populations were actually D. variabilis re- cruited from larvae swept north of the sustain- able species range due to fortuitous warm-water currents, and that conchological differences between the two species are merely ecophenotypic. However, Morrison's (1971) revision of Chanley's specimens con- cluded that D. fossor is not a summer range extension of D. variabilis, further supporting the distinction between the two species. According to Morrison (1971), Donax variabilis is not found north of Virginia Beach, Virginia, and D. fossor is not found south of Nag's Head, North Carolina. Thus, their ranges do not overlap until Virginia Beach, Virginia. The objective of this study was to compare the anatomy of specimens from Florida and South Carolina (supposedly D. variabilis) to that of specimens from Avalon, New Jersey (supposedly D. fossor), to provide evidence for confirming or refuting the biological valid- ity of the two species along the east coast of the United States. Anatomical investigations have been per- formed for about ten species of donacids (e.g., Ridewood, 1903; Pelseneer, 1911; Graham, 1934; Yonge, 1949; Duval, 1963; Nakazima, 1965; Wade, 1969; Narchi, 1972, 1978; Mouéza 8 Frenkiel, 1974, 1976, 1978; Odiete, 1981; Hodgson, 1982; Ansell, 1983; Salas- Casanova 8 Hergueta, 1990; Passos, 1998), however, none has been published on the species analyzed here. Those papers provide a secure scenario for discussion of the ana- . tomical characters at the species and family level. MATERIAL AND METHODS Specimens were collected and fixed directly in 70% ethanol. Gross dissections were per- formed with the specimen immersed in fixa- tive under a stereomicroscope. Histological 5-um serial sections of partial regions were stained with Mallory's tristain. All drawings were made with the aid of a camera lucida. Abbreviations used in figures: am, anterior adductor muscle; an, anus; au, auricle; cm, cruciform muscle; cv, ctenidial (efferent) ves- sel; dd, ducts to digestive diverticula; dg, di- gestive diverticula; dh, dorsal hood; di, inner demibranch; do, outer demibranch; dv, dorsal portion of outer demibranch covering visceral mass; es, esophagus; fe, foot elevator muscle; ff, fecal furrow; fm, posterior foot retractor muscle; fp, foot protractor muscle; fr, anterior foot retractor muscle; ft, foot; gf, ventral gas- tric fold; gm, gill retractor muscle; go, gonad; gs, gastric shield; id, insertion of outer demibranch in mantle; in, intestine; р, inner hemipalp; is, septum in siphonal base sepa- rating infra- and suprabranchial chambers; ki, kidney; mb, mantle border; mm, mantle muscles of siphonal chamber; mp, mantle pa- pillae; op, outer hemipalp; pa, posterior ad- ductor muscle; pc, pericardium; pd, dorsal caecum; pp, palp; sc, siphonal chamber; se, excurrent siphon; ri, ridge in esophageal in- sertion in stomach: rt, rectum; sh, shell; si, in- NORTHWESTERN ATLANTIC DONAX 461 current siphon; sm, siphonal retractor muscle; ss, style sac; st, stomach; ty, typhlosole; um, fusion between left and right mantle lobes; ve, ventricle; vg, visceral ganglia; vm, visceral mass. Abbreviations of institutions: FMNH, Field Museum of Natural History, Chicago; MZSP, Museu de Zoologia da Universidade de Sáo Paulo, Brazil; USNM, National Museum of Natural History [United States National Mu- seum], Washington, DC. SYSTEMATICS Donax fossor Say, 1822 (Figs. 1-5, 11, 12, 14, 16-26) Synonymy (for additional references, see Morrison, 1971: 456): Donax fossor: Abbott, 1974: 509 (as form of D. variabilis); Emerson & Jacobson, 1976: 415-416, pl. 43, fig. 15; Abbott & Morris, 1995: 91; Adamkewicz 8 Harasewych, 1996: 97-103. Donax variabilis: McDermott, 1983: 529-538; Alexander et al., 1993: 289-303. Diagnosis Shell triangular; anterior (pedal) edge very thick; posterior region flattened, with a flattened posterior margin. Mantle edge and siphons with large number of papillae. Gastric style sac almost straight, with distal region positioned in ventral region of visceral sac. Intestine bearing few undulations. Description Shell (Figs. 1-5, 14): Up to 13 mm length. Color varying from pure white to yellowish. Outline somewhat triangular; general shape cuneiform; anterior region with very thick edge, producing a strong slope. Umbo lo- cated in posterior third of hinge, weakly pro- truded, rounded. Outer surface smooth, bearing only concentric undulations. Hinge with three teeth in left valve, including small tooth just anterior to umbo, another two car- dinal teeth at posterior level of ligament (Fig. 14, arrows) of similar size, transverse, sepa- rated by short depression. Right valve with single small, transverse tooth articulating between two cardinal teeth of left valve; also bearing sockets for teeth of left valve. Mantle: Mantle border of somewhat uniform width along its length; mostly not fused, ex- cept in siphonal area and short portion ven- tral to them (Figs. 18, 20, um). Mantle border with two folds, each with series of small pa- pillae of uniform size; each papilla long, slen- der (Figs. 11, 18, 20), with longitudinal, narrow furrow along outer side; tip concave, edges slightly projecting. Siphons separated from each other, similarly sized (Figs. 18, 20), each protected by cavity formed by mantle, depth about one-quarter of animal's length. Siphonal walls thickly muscular, basal region thinner, with muscle fibers arranged radially like a fan (Figs. 11, 20), originating from pallial sinus of shell. Incurrent siphon with 6-7 larger folds projecting inwardly, each bearing several papillae on outer sur- face (Figs. 12, 16, 17, 20), smaller and sim- pler tentacles among large papillae (Figs. 16, 17). Excurrent siphon with simpler tentacles than incurrent siphon (Fig. 20); tentacles narrow, with 2-3 papillae on distal end; fe- cal groove narrow, shallow, running longitu- dinally along internal ventral surface of excurrent siphon (Fig. 20, ff), terminating in a furrow on siphonal edge between two ten- tacles. Cruciform muscle located on ventral edge between middle and posterior thirds of mantle edge, at base of incurrent siphon (Figs. 18, 20), inside mantle fusion (Fig. 21); anterior branches longer and narrower, т- serted on shell tangentially; posterior branches broader, shorter, inserted on shell almost perpendicularly (Fig. 21). Several radial muscle fibers connecting posterior edge of posterior adductor muscle with mantle border, becoming successively larger ventrally, abruptly terminating in middle re- gion of siphonal area; ventral fibers thicker, branched distally (Fig. 23). Mantle Organs: Pallial cavity very ample (with only narrow dorsal portion not covered by cavity) (Fig. 18). Gills small, occupying about one-third of pallial cavity (Fig. 18). Outer demibranch shorter than inner demibranch anteriorly, gradually becoming about same width posteriorly. Both demibranch ventral edges simple, lacking food grooves (Figs. 18, 19). Gill insertion on visceral mass be- tween demibranchs, that of inner demi- branch (on visceral mass) more ventral; outer demibranch with portion dorsal to gill insertion, covering visceral mass (Fig. 19), with shallow longitudinal furrow separating 462 SIMONE & DOUGHERTY FIGS. 1-13. Donax shells and mantle. FIGS. 1-5: Donax fossor shells, two specimens MZSP 36508 (New Jersey); FIGS. 1, 2: Right lateral view; FIGS. 3, 4: Ventral view; FIG. 5: Detail of anterior region of Fig. 4; FIGS. 6-10: Donax variabilis shells, two specimens MZSP 36509 (South Carolina); FIGS. 6, 7: Right lateral view; FIGS. 8, 9: Ventral view; FIG. 10: Detail of anterior (foot) region of Fig. 9; FIG. 11: D. fossor, left mantle lobe, inner view, detail of siphonal basal region; FIG. 12: D. fossor, incurrent siphon, detail of apical region opened longitudinally, showing tentacles and papillae on inner edge; FIG. 13: Same for D. variabilis, showing more weakly developed tentacles and papillae. Scales = 2 mm. this portion from remaining demibranch, con- necting to visceral mass far dorsal of gill in- sertion. Gill dorsal and ventral connections to visceral mass cilial only. Gills connected to one another posterior to visceral mass in median line; no other anatomical gill connec- tion, either with mantle (only by cilia) or with posterior adductor muscle. A communication between infra- and suprabranchial chambers remaining in contracted gill condition (Fig. 20). Mantle transverse septum in ventral base of excurrent siphon, separating infra- NORTHWESTERN ATLANTIC DONAX 463 FIGS. 14-17. Donax hinges and siphons. FIG. 14: Donax fossor hinge, left valve at left, arrows indicating teeth. Scale = 1 mm; FIG. 15: Same for D. variabilis; FIG. 16: D. fossor, extended incurrent siphon, dorsal-slightly apical view. Scale = 0.5 mm; FIG. 17: Same, apical view, showing fully extended tentacles and papillae. Scale = 0.5 тт. 464 SIMONE & DOUGHERTY FIGS. 18-23. Donax fossor anatomy. FIG. 18: Whole specimen, left view, left mantle lobe partially removed (except for portion in siphonal base); FIG. 19: Left gill, transverse section at mid-region, with some adjacent structures; FIG. 20: Siphonal region, left view, both siphons opened longitudinally along their left side; FIG. 21: Detail of posteroventral union of mantle lobes, at base of incurrent siphon, inner view, with inner layer of tissue removed; FIG. 22: Left labial palp, outer hemipalp deflected, with adjacent region of inner demibranch; FIG. 23: Foot and visceral mass, left view, emphasizing main muscle system, pericardial structures, topology of visceral glands, gill muscle and mantle muscles shown in situ. Scale bars = 1 mm. NORTHWESTERN ATLANTIC DONAX 465 and suprabranchial chambers, somewhat short (Fig. 20, is). Palps long, сигуеа, slightly triangular, located in anteroventral corner of inner demibranch, relatively small (Fig. 18); outer surface smooth; inner surface (Fig. 22) with several uniform transverse folds, some- what parallel to palp posterodorsal edge; dorsal portion of folds very narrow, ventral portion broader, a short transverse whitish furrow located in distal end of each fold of outer demibranch (Fig. 22, op); folds ending short distance from palp inner edge, produc- ing a narrow smooth margin. Palps and palp folds gradually becoming shorter toward an- terior, a smooth inner area in palp portion surrounding mouth. Foot and Main Muscle System: Foot large, about half of body size; triangular, tip broadly pointed, laterally flattened, bent in retracted condition (Figs. 18, 23). Adductor muscles similarly sized (Figs. 18, 23); each with ven- tral region somewhat circular, dorsal some- what pointed. Anterior adductor muscle close to anterodorsal shell edge. Posterior adduc- tor muscle at middle level of posterior shell edge. Paired anterior protractor muscles broad, thin, flat, originating in posteroventral edge of anterior adductor muscle, passing posteroventrally to insert fan-like on lateral wall of visceral mass (Figs. 18, 23, fp). Paired anterior pedal retractor muscles (Figs. 18, 23, fr) long, flat, narrow, slender, originating on shell just posterior to anterior adductor muscle, internally crossing anterior protrac- tor muscles, passing superficially posteroventrally, inserting fan-like in middle region of transitional area of foot-visceral mass. Paired pedal levator muscles (Figs. 18, 23, 25, fe) narrow, long, cylindrical, origi- nating in umbonal cavity at some distance posterior to anterior adductor muscle, run- ning ventrally and anteriorly close to median line and close to one another, covered later- ally by anterior retractor muscles, inserting in transitional area of foot-visceral mass. Paired posterior pedal retractor muscles (Figs. 18, 23, fm) originating on shell just dorsal to posterior adductor muscle, pass- ing anteroventrally, narrow in posterior half, gradually becoming broad in anterior half, inserting on outer surface of posterior region of visceral mass. Paired gill retractor muscles very narrow, thin (Figs. 18, 19, 23, gm), origi- nating on very small area of umbonal cavity at some distance posterior to levator muscles, penetrating pallial cavity between demibranchs, passing along gill to posterior end, becoming thinner and more diffuse (Fig. 23). Visceral Mass: Internal organs visible Бу trans- parency only in narrow dorsal umbonal re- gion (Fig. 18). Digestive diverticula pale green in preserved specimens, somewhat small, surrounding gastric area in dorsal re- gion of visceral cavity. Gonad very large (Fig. 23), cream-colored, occupying most of vis- ceral cavity, surrounding all visceral struc- tures except renopericardial organs and some portions of digestive diverticula. Circulatory and Excretory Systems: Pericar- dial cavity relatively small, located just ante- rior to posterior adductor muscle (Figs. 18, 23). Paired auricles triangular, with thin, transparent walls, central region connecting directly to gill, anterior and posterior verti- ces connecting to relatively short efferent gill vessels (Fig. 23). Auricles connecting to ven- tricle laterally. Ventricle surrounding intes- tine. Kidney whitish, mostly solid, located ventral to pericardium, compressed by pos- terior pedal retractor muscles, gonad and posterior adductor muscle. Digestive System: Mouth somewhat small, in central region between palps. Esophagus relatively long, dorsoventrally flattened, away from anterior adductor muscle (Fig. 24). Stomach (Figs. 24-26) ovoid, located in umbonal region of visceral mass totally sur- rounded by digestive gland. Gastric dorsal hood narrow, about half of stomach length, originating close to median line, situated on left side covering dorsal surface of stomach (Figs. 24, 25, dh); inner surface smooth, except for broad, low longitudinal fold on ventral surface (Figs. 25-26). Dorsal gas- tric caecum (Figs. 25, 26, pd) small, gener- ally bifid; walls whitish, glandular, with inner space narrow; connecting to stomach by very narrow duct to anterior third of stomach left- dorsal side (Fig. 25). Inner surface of poste- rior esophagus smooth; esophageal junction with stomach marked by tall, transverse typhlosole almost entirely surrounding this insertion, except at two narrow portions in lateroventral region where pair of furrows begin, running toward ducts of digestive di- verticula (Fig. 26). Digestive diverticular ap- ertures located lateroventrally in anterior gastric region. Dorsal hood aperture just dorsal of left apertures of digestive diver- 466 SIMONE & DOUGHERTY dg dh dd st es FIGS. 24-26. Donax fossor anatomy. FIG. 24: Visceral mass showing digestive system in situ, left view, with some adjacent structures; left wall of visceral sac, gonad and part of digestive diverticula removed; FIG. 25: Stomach, dorsal view, with some adjacent structures; with transverse sections of dorsal hood and dorsal caecum; FIG. 26: Same, left wall sectioned longitudinally and deflected, exposing inner surface. Scale bars = 1 mm. ticula; transverse, low fold (closer to dorsal hood aperture) separating the two digestive diverticula apertures, bearing short projec- tion of gastric shield (Fig. 26). Aperture of gastric dorsal caecum immediately dorsal to right aperture of digestive diverticula; deep, narrow furrow running posteriorly from right aperture of digestive diverticula, along ven- tral-right inner gastric surface, to intestinal origin. Gastric shield occupying about one- third of inner gastric surface, located in ven- tral and left inner regions (Fig. 26). Intestine and style sac origins adjacent (that of intes- tine right-anterior); narrow low fold almost NORTHWESTERN ATLANTIC DONAX 467 entirely surrounding style sac origin, except for short portion adjacent to intestinal origin (Fig. 26). Style sac entirely separated from intestine, very long (longer than dorsoven- tral height of visceral cavity), passing ven- trally, gradually narrowing, with tip somewhat pointed, curved forward (Fig. 24). Intestine mostly narrow, from origin in posteroventral region of stomach, to right of style sac, pass- ing anteriorly, contouring ventral gastric re- gion to left-anterior side, abruptly twisting towards right, passing sinuously posteroven- trally, crossing right side of style sac ventral third; in posteroventral region of visceral cavity curving dorsally, to region just poste- rior to stomach; curving abruptly toward ante- rior, crossing pericardium and posterodorsal surface of posterior adductor muscle (Fig. 24). Anus in middle region of posterior ad- ductor muscle, bearing short longitudinal, narrow notch on median line (Fig. 24, an). Genital System: Gonad apparently dioecious. Central Nervous System: Not seen in detail, except for pair of large visceral ganglia (Fig. 23) close to one another on posteroventral surface of posterior adductor muscle. Measurements (length x dorsoventral height х lateral width, in mm): MZSP 36508: no. 7, ISSO x 5.8; no. 8, 10.4 x 6.0 x 3.6. Material Examined: U.S.A.: New Jersey; 66" St., Avalon, MZSP 36508, 13 specimens (Joanne Dougherty, coll., 01/ix/2001). Donax variabilis Say, 1822 (Figs. 6-10, 13, 15, 27) Synonymy (for additional references, see Morrison, 1971: 550-551): Donax variabilis: Boss, 1970: 205-206; Tif- fany, 1971: 82-85; Melville, 1976: 19-21; Abbott, 1974: 509, fig. 5753; Emerson & Jacobson, 1976: 414, pl. 43, fig. 14; Mikkelsen, 1981: 230-239; Leber 1982: 297-301; Mikkelsen, 1985: 308-311; Vega & Tunnell, 1987: 97-135; Ruppert & Fox, 1988: 158, pl. B28; Estes & Adamkewicz, 1991: 321-332; Bonsdorff & Nelson, 1992: 358-365; Nelson et al., 1993: 317-322; Adamkewicz & Harasewych, 1994: 97-103; Meinkoth, 1995: 556, pl. 321; Ellers, 1995a: 120-127; 1995b: 128-137, 1995c: 138-147; Abbott & Morris, 1995: 91, pl. 4, fig. 37; Adamkewicz & Harasewych, 1996: 97-103; Wilson, 1999: 61-83; Manning & Lindquist, 2003: 415—422. Diagnosis Shell elongated-elliptic; anterior (pedal) edge thin; posterior region compressed, anterior slope weak. Mantle edge and siphons with few weakly branched papillae. Gastric style sac curved, with distal region positioned in antero- dorsal region of visceral sac. Intestine bear- ing several loops. Description Shell (Figs. 6-10, 15): Up to 20 mm. Color very variable, from pure white to yellowish, grayish, brownish and reddish (see Morrison, 1971, for further comments). Outline some- what elongated and elliptical; general shape weakly cuneiform, somewhat compressed; anterior (pedal) region with edge about half as thin as that of preceding species. Umbos located between middle and posterior third of hinge, weakly protruded, rounded. Re- maining characters similar to those of Donax fossor, differing in: (1) larger size; (2) sur- face with more developed radial sculpture; (3) anterior pedal region thinner, narrowing gradually, with a more rounded anterior mar- gin (Figs. 8-10); (4) posterior siphonal slope longer (Figs. 6, 7), less abrupt than in D. fossor. This last difference is reflected in the position of umbo, which is in posterior quar- ter of dorsal edge in D. fossor, between middle and posterior third of dorsal edge in D. variabilis; (5) cardinal teeth shorter (Fig. 15), total hinge somewhat narrower. Mantle: Features similar to those of D. fossor, with the following differences: Mantle bor- der entirely bearing single series of papillae on inner fold; outer fold double, smooth or with very small, low papillae between papil- lae of inner fold, with two series of papillae in region of siphonal chamber similar to those in D. fossor. Mantle border papillae similar to those of D. fossor, but smaller. Distal end of incurrent siphon with 5-6 folds proportion- ally smaller than those of D. fossor, bearing fewer papillae on outer surface (Fig. 13); that of excurrent siphon with 4—5 folds lacking papillae or bearing few, very small papillae; fecal furrow also present (Fig. 27, se). Mantle Organs: Similar to those of D. fossor, including characters of gill and palps (Fig. 27). 468 SIMONE & DOUGHERTY FIG. 27. Donax variabilis anatomy, whole specimen, left view, left mantle lobe partially removed (except at siphonal base), left gill, left wall of visceral sac, gonad and digestive diverticula removed, palps deflected. Scale = 2 mm. Foot and Main Muscle System, Visceral Mass, and Circulatory and Excretory Systems: Similar to those of D. fossor (Fig. 27). Digestive System: General organization simi- lar to that of D. fossor, with following differ- ences. Dorsal gastric caecum proportionally larger, with glandular portion more greatly developed. Gastric style sac proportionally longer (Fig. 27, ss), passing surrounding ventral border of visceral sac, decreasing gradually, in mid-ventral region of visceral mass ascending anteriorly, slightly sinuously, with distal end in mid-dorsal region of vis- ceral sac at short distance from mouth. In- testine more convolute (Fig. 27, in), bearing a strong loop in region ventral to stomach. Measurements (length x dorsoventral height X lateral width, in mm): MZSP 36509: no. 1, A ADO A LES OS SL Material Examined: U.S.A.: South Carolina; Pawleys Islands, south end, MZSP 36509, 15 specimens (David Bushek, coll., 11/viii/2002). Florida; Franklin County, south corner of St. George Island, MZSP 36507, 12 specimens (Harry G. Lee & R. L. Whipple, coll., 29/vii/ 2002); Florida Keys; Monroe County, Mate- cumbe Key, FMNH 202068, 17 shells, 202070, 12 shells, 202071, 3 shells (A. Koto, coll.). DISCUSSION Donax fossor and D. variabilis are very simi- lar in their morphology. The most significant difference in the shell is the pedal region, which is thicker and with a narrow slope in D. variabilis (Fig. 5), whereas in D. fossor is of a shape more typical of donacids, and with a thinner shell border (Fig. 10). The outline is also different; D. fossor has a more developed posterior slope and this region is shorter and more blunt (Figs. 1-2, 6-7). Donax fossor has many more papillae both at the mantle bor- der and the siphons, with two series of some- what long papillae (Fig. 11), while most specimens of О. variabilis have a single row, although some bear very small and short pa- pillae on the outer mantle border fold. The incurrent siphon of D. fossor (Figs. 14, 15) has larger apical folds with proportionally more papillae on the outer surface than in D. variabilis (Figs. 12, 13). The tip of the excur- rent siphon generally lacks papillae in D. variabilis (Fig. 27), or has very small papillae, while that of D. fossor has well-developed papillae (Fig. 20). The gastric style sac is longer in D. variabilis, being curved and slightly sinuous, with its distal end close to the mouth (Fig. 27); in D. fossor, the style sac is almost straight, with the distal end in the ventral region of the visceral mass (Fig. 24). NORTHWESTERN ATLANTIC DONAX 469 It is important to emphasize that the style sac proportions were consistent among speci- mens of different sizes of both species. The intestine of D. variabilis is more highly looped than that of D. fossor, mainly in its proximal region ventral to the stomach (Fig. 27). Both Donax species show anatomy consis- tent with previous reports for the family, such as the siphons being separated from each other beginning at their base, the presence of a cruciform muscle, and the separation of the gastric style sac from the adjacent intes- tine. The circulatory system is very similar to that of Donax trunculus (Linnaeus, 1758) (Mouëza & Frenkiel, 1978). The cruciform muscle (Fig. 21) is cited as a character of the superfamily Tellinoidea (Yonge, 1949); it matches that previously reported in both spe- cies studied (Mouëza & Frenkiel, 1974: figs. 2-5). The siphonal constitution is also similar to those described for other species of the fam- ily, with a clear siphonal septum separating supra- and infrabranchial chambers (Mouëza & Frenkiel, 1978; Hodgson, 1982). This sep- tum (Fig. 20, id) aids in directing water flow to the gills, because the gills are not anatomi- cally connected to the siphons. This study is the first report of a fecal furrow (Figs. 20, 27, ff) in any donacid species. The shape, num- Бег and constitution of the siphonal tentacles and papillae (Figs. 11, 12) are obviously as- sociated with the high energetic environment that donacids normally inhabit; their differ- ences have been very useful in comparative analysis among sympatric species (e.g., Ansell, 1981; 1983: fig. 6). Species of the gen- era Iphigenia (Narchi, 1972) and Egeria (Purchon, 1963) have weakly developed siphonal papillae and inhabit low energy en- vironments. The well-developed radial pallial muscles connecting the posterior edge of the posterior adductor muscle with the mantle edge (Figs. 22, 27, mm) is a unique feature of the two species studied herein, and has not been described for any other species. The more dorsal of these muscles is short and thin, gradually becoming thicker, longer and more distinct ventrally, where it abruptly terminates. A portion of the outer demibranch covering the visceral sac (Fig. 19, dv) is as previously reported in other donacids; this portion has been called supra-axial extension of the ctenidium (Ansell, 1983). The two species studied here differ from other known donacids in lacking a food groove on the ventral edge of the inner demibranch; this groove has been found in other species (Purchon, 1963; Yoloye, 1977; Ansell, 1981; Passos, 1998). Another interesting feature is the well-devel- oped gill muscle (Fig. 22, gm), which occurs in both studied species, originating inside the umbonal cavity (Figs. 18, 27) and penetrat- ing the mantle along the gill just between the demibranchs. The posterior region of the gill muscle is more diffuse and thin. Only one simi- lar structure has been reported in the litera- ture on donacids, the so-called “demibranch muscle” in Donax gouldii Dall, 1921, studied by Pohlo (1967). However, some confusion with the levator muscle of the foot exists in that description (Pohlo, 1967: 330). The gill muscle could be an exclusive feature of the three species (D. fossor, D. variabilis, D. gouldii). The stomach of Donax variabilis and D. fossor (Figs. 25, 26) is also typical of donacids, having a transverse typhlosole in the esoph- ageal insertion and a dorsal hood at left. How- ever, the dorsal caecum found on the right side of stomach can be absent in some spe- cies (Nakazima, 1965) and has been called a stomach appendix (Pohlo, 1967; Wade, 1969; Narchi, 1972, 1978; Passos, 1998) or poste- rodorsal caecum (Yonge, 1949; Purchon, 1963; Mouéza & Frenkiel, 1976; Salas- Casanova & Hergueta, 1990). These last au- thors demonstrated that in D. venustus (Poli, 1795) sand grains and similar coarse particles pass by the dorsal caecum; in the present species the caecum appears to be a gland, because it is almost entirely filled by glandu- lar tissue and has a narrow duct separating it from the stomach (Figs. 25, 26, pd). Dorsal caeca have been reported in members of other tellinoidean families (Yonge, 1949: figs. 28—29), but they differ from those of donacids in being larger and amply opened to the stom- ach. The species studied here also lack tall gastric typhlosoles as those reported in the above-mentioned papers. The highly curved style sac is a unique feature of D. variabilis (Fig. 27); however, Mouéza & Frenkiel (1976) showed a long style sac for D. trunculus, fig- uring it as a semi-circle. The configuration of the intestinal coils is also useful for species distinction (e.g., Ansell, 1983: fig. 7) reinforcing the distinction be- tween the two species studied here. The donacid intestine is normally weakly coiled, however, Egeria radiata (Lamarck, 1804) is an exception (Purchon, 1963: fig. 10). 470 УМОМЕ & DOUGHERTY CONCLUSIONS (1) Specimens from New Jersey (attributable to Donax fossor) and South Carolina and Florida (attributable to D. variabilis) are con- firmed as separate species, distinguishable by morphological features of shell and soft parts, in agreement with the molecular find- ings of Adamkewicz & Harasewych (1996). (2) Morphological study of Donax fossor and D. variabilis confirmed the typical bauplan of the family as revealed by previous au- thors. Detailed investigation of all organ systems showed useful distinguishable dif- ferences that are consistent in each sample and sufficient for specific separation. (3) At the present time, the radial pallial muscles of the siphonal chamber are unique to Donax fossor and D. variabilis, not hav- ing been described for any other donacid species. The gill muscle is an exclusive fea- ture of these two species and D. gouldii. ACKNOWLEDGMENTS This study is one result of the International Marine Bivalve Workshop, held in the Florida Keys, 19-30 July 2002, funded by U.S. National Science Foundation award DEB-9978119 (to co-organizers R. Bieler and P. M. Mikkelsen), as part of the Partnerships in Enhancing Ех- pertise in Taxonomy [PEET] Program. Addi- tional support was provided by the Bertha LeBus Charitable Trust, the Comer Science 8 Education Foundation, the Field Museum of Natural History, and the American Museum of Natural History. This study was also part of the junior author master's thesis, supported by the Lerner-Gray Fund of the American Museum of Natural History, Conchologists of America, and Sigma Xi, and 1$ in part supported by Brazilian Fundacáo de Amparo a Pesquisa do Estado de Sáo Paulo, process nos. 00/11074-5 and 00/11357-7, to the senior author. We are grate- ful to Flavio Dias Passos (Instituto de Biociéncias, Universidade de Sáo Paulo) for bibliographical support and suggestions related to anatomy. We are also grateful to David Bushek and Harry Lee for providing specimens. LITERATURE CITED ABBOTT, R. T., 1954, American seashells. Van Se Princeton, New Jersey. 541 pp., 32 pls. ABBOTT, К. T., 1974, American seashells, 2" ed. Van Nostrand Reinhold Company, New York. 663 pp., 24 pls. АВВОТТ, К. Т. & P. A. MORRIS, 1995, Shells of the Atlantic & Gulf coasts & the West Indies. Peterson Field Guide, Houghton Mifflin Com- pany, Boston, Massachusetts. 350 pp., 74 pls. ADAMKEWICZ, L. & M. G. 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Philosophical Transactions of the Royal Society, 234B: 29-76. Revised ms. accepted 22 January 2004 MALACOLOGIA, 2004, 46(2): 473-501 PINCTADA LONGISQUAMOSA (DUNKER, 1852) (BIVALVIA: PTERIIDAE), AN UNRECOGNIZED PEARL OYSTER IN THE WESTERN ATLANTIC Paula M. Mikkelsen’, Пуа Témkin?, Rudiger Bieler? & William С. Lyons* ABSTRACT Pinctada longisquamosa (Dunker, 1852) is redescribed based on original collections from the Florida Keys, type material, and other museum specimens. Conchological and anatomical features support its transfer from the genus Репа (originally Avicula) to the genus Pinctada. А unique periostracal structure, with elements corresponding to the indi- vidual prismatic structures of the outer shell layer is described and illustrated. Comparison is made between this species and Pinctada imbricata and Pteria colymbus, with which it co-occurs and has often been confounded. Its preferred habitat in Florida Bay is seagrass, often intermixed with macroalgae, to which individuals byssally attach substantially off the silt-laden bottom. Quantitative and qualitative data from Florida Bay populations show fluctuating population densities, from absence to over 300 individuals/m?, sometimes within a few months. These pronounced changes could be seasonal and/or influenced by the extremes of high and low salinity that sometimes occur in the Florida Bay estuarine sys- tem. Key words: Florida Keys, Florida Bay, sanctuary, Pteria, Mollusca, systematics, anatomy. INTRODUCTION Members of the bivalve family Pteriidae, in- cluding pearl oysters and wing oysters, are characterized by obliquely ovate shells with a triangular wing-like projection both anterior and posterior to the straight hinge line. They are monomyarian and epibyssate, their shells in- equivalve, inequilateral, and interiorly nacre- ous. Hinge teeth are small to obscure, and the exterior surface is often adorned with layers of overlapping lamellae arranged in radial rows. The fossil record extends from the Tri- assic. Three extant genera (Pteria Scopoli, 1777; Pinctada Róding, 1798; Electroma Stoliczka, 1871; Hertlein & Cox, 1969) and about 50 living species are currently recog- nized. Mainly tropical and subtropical in distri- bution, pteriids are relatively common and well-recognized bivalves by virtue of their his- torical and current roles as sources of nacre (mother-of-pearl) and natural or cultured pearls (Landman et al., 2001). Despite this familiar- ity, anatomical characters have received little attention and detailed work has mostly been restricted to a few species of commercial in- terest mainly in the genus Pinctada (Shiino, 1952; Hynd, 1955; Shirai, 1994). No phyloge- netic framework yet exists for the family. In the tropical western Atlantic, Pteriidae is represented mainly by the Atlantic pearl oys- ter, Pinctada imbricata Róding, 1798 (which some authors, e.g., Shirai, 1994, consider circumtropical), and the Atlantic wing oyster, Pteria colymbus (Réding, 1798). One other species of Pinctada, P. margaritifera (Linnaeus, 1758), widespread throughout the Indo-Pacific, has been introduced to Florida (Chesler, 1994; Carlton, 1996; Camp et al., 1998), but without evidence of established reproductive populations. One unidentified Electroma species was reported as well es- tablished in coastal Colombia (Borrero & Diaz, 1998), also presumably introduced from the Indo-Pacific. Three other nominal species of Репа, P. vitrea (Reeve, 1857), P. hirundo (Linnaeus, 1758), and P. longisquamosa (Dunker, 1852), have also been reported as ‘Division of Invertebrate Zoology, American Museum of Natural History, Central Park West at 79th Street, New York, New York 10024-5192, U.S.A.; mikkel@amnh.org “AMNH, and BRIDGES Program, Department of Biology, New York University, 1009 Main Building, 100 Washington Square East, New York, New York 10003, U.S.A. “Department of Zoology, Division of Invertebrates, Field Museum of Natural History, 1400 S. Lake Shore Drive, Chicago, Illinois 60605-2496, U.S.A. “Florida Marine Research Institute (retired); present address: 4227 Porpoise Drive SE, St. Petersburg, Florida 33705, U.S.A. 474 MIKKELSEN ЕТ AL. indigenous western Atlantic species (Hayes, 1972; Rios, 1994; Turgeon et al., 1998), but are far less represented т the literature and collections. This paper reviews the taxonomy and geographic distribution of one of these poorly known species, P. longisquamosa, es- tablishing its generic placement, and describ- ing its anatomy and life habits, based on original collections of living specimens from the Florida Keys together with a re-evaluation of existing literature and selected museum data. Comparisons are drawn with sympatric pteriids within its geographical range and with known anatomical data for the family. MATERIALS AND METHODS This study is part of an ongoing investiga- tion of marine molluscan biodiversity in pen- insular Florida and the Florida Keys, formally initiated by PMM and RB in 1994. Consecu- tively numbered stations comprising these collections are preceded by ап “ЕК” acronym in the following text and Appendix. Living ani- mals and empty shells of Pinctada longi- squamosa were collected mainly by hand during snorkeling on shallow-water (2-10 m) seagrass flats; the majority of observations on living specimens were made from beds of Thalassia testudinum Kónig and Syringodium filiforme Kützing in Florida Bay off the Upper Florida Keys. Pinctada imbricata and Репа colymbus were collected for comparative pur- poses from locations throughout the Florida Keys. Voucher FK specimens were fixed in 5% formalin, later transferred to 70% ethanol (or fixed directly in 95-100% ethanol for potential molecular investigation), and are deposited in the American Museum of Natural History (AMNH), New York, and Field Museum of Natural History (FMNH), Chicago. Our attention was first drawn to this species by its prominent occurrence in a study con- ducted by the Florida Marine Research Insti- tute (FMRI) to assess the responses of molluscan populations to changes in salinity in Florida Bay during periods of drought and floods. Results, provided here by WGL, were obtained during quantitative sampling con- ducted at 101 sites spaced evenly throughout Florida Bay (Fig. 30) during the summers of . 1994 (a drought year; salinity range at sites 14.8-51.8 ppt) and 1996 (year following a flood year; site salinity range 0.8-40.9 ppt). Fifteen samples were taken using a large-bore coring device at each site, with 0.27 m? of bottom area collectively sampled during each site visit. Sam- pling and analytical techniques were described by Lyons (1998), and a preliminary summary of results was presented by Lyons (1999). Live- collected specimens from this study are stored FIG. 1. Diagrammatic shells of Pinctada longisquamosa, showing general exterior (left) and interior (right) features and methods of shell measurement. (aa, anterior auricle; ams, adductor muscle scar; bn, byssal notch; br, byssal ridge; cs, commarginal stripe; Ig, ligament; nb, border of nacreous layer; pa, posterior auricle; pl, primary lamella; plms, pedal levator muscle scar (anterior pedal levator muscle scar not shown, situated within ambonal arch); pm, prismatic margin; pms, pallial muscle scars; pr, posterior ridge; rms, pedal retractor muscle scar; s, socket of anterior dentition (corre- sponding to tooth in RV, not shown); sl, secondary lamella; u, umbo). PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 475 in 70% ethanol in the Specimen Reference Collection of the Florida Marine Research In- stitute, Florida Fish and Wildlife Conservation Commission, St. Petersburg (FSBC |). Living specimens used for anatomical ob- servations in the field were relaxed by chilling in a household refrigerator assisted by the addition of magnesium sulfate crystals (Epsom salts) to their seawater supply, or in an iso- tonic aqueous magnesium chloride solution. The anatomy of preserved specimens was studied using a combination of histology and gross dissection in various planes. For histol- ogy, shells were removed manually from for- malin-fixed, ethanol-preserved specimens; tissues were dehydrated through a graded ethanol series, followed by clearing in xylene substitute, and embedding in paraffin. Com- plete 7-um serial sections were produced for intact individuals in transverse and sagittal planes, and stained with PAS (Alcian Blue/ Periodic Acid/Schiff’s) trichrome stain. Dried shells and excised preserved tissues were prepared for scanning electron microscopy (SEM) by critical point drying (for tissues only) and gold-palladium sputter coating, and were then viewed on a Zeiss DSM-950 scanning electron microscope at AMNH. Specimen photography used a variety of equipment and techniques. Laboratory photo- graphs of living animals (Fig. 4) were taken in aquaria or finger bowls with a 35 mm single- lens reflex camera and electronic flashes; in situ underwater photos (Fig. 29) were accomplished with the same equipment in an underwater housing. Whole-valve and detail light microgra- phy (Figs. 5-9) used a Microptics® micro/macro imaging system based on a high-resolution Nikon® single-lens reflex digital camera. Shell measurements (taken with digital cali- pers or with ocular micrometer on a stereomi- croscope) and meristics were taken from the right valve. Maximum shell height was mea- sured perpendicular to the hinge line to the most distal point of the ventral shell margin; maximum shell length was taken parallel to the hinge line (Fig. 1). Primary radial sculp- tural elements were counted on the main body of the shell. Size is expressed as shell length unless otherwise noted. Although the hinge teeth of pteriids have been called “cardinal” and “lateral” teeth by some authors (e.g., Hayes, 1972), we use the phrases “anterior dentition” and “posterior dentition” to avoid unsupported assumptions of homology with the teeth of heterodont bivalves. Other cited repositories include: ANSP — Academy of Natural Sciences of Philadelphia, Pennsylvania; BMNH - The Natural History Museum, London [= British Museum (Natural History)]; BMSM - Bailey-Matthews Shell Museum, Sanibel Island, Florida; DMNH — Delaware Museum of Natural History, Wilmington; HMNS - Houston Museum of Natural Sciences, Texas; UMML - Rosenstiel School of Marine and Atmospheric Science, University of Miami, Florida [= University of Miami Marine Laboratory]; USNM - National Museum of Natural History, Smithsonian In- stitution, Washington, DC [= United States National Museum]; and ZMB - Institute of Systematic Zoology, Museum für Naturkunde, Berlin, Germany [= Zoologisches Museum Berlin]. Other abbreviations and conventions used in the text (other than figure labels, explained in the figure legends) are: alc — fluid-preserved (alcohol) specimen; D — “diameter” or shell inflation; frag — shell fragment; H — shell height; juv — juvenile or subadult; L — shell length (of- ten called “width”); LV - left valve; pair — an empty (dead) complete shell (2 valves); RV — right valve; som - a living specimen; and valve — an empty (dead) single valve. SYSTEMATIC RESULTS Pterioidea J. E. Gray, 1847 (1820) Pteriidae J. E. Gray, 1847: 199 [as Pteriadae] (1820) Aviculidae Goldfuss, 1820, is an available older name, but was replaced before 1961 when the name Avicula Вгидшеге, 1792, was deemed a junior synonym of Pteria Scopoli, 1777. Because Pteriidae has won general acceptance, it is maintained under ICZN (1999) Art. 40.2. Pinctada Réding, 1798: 166 (pearl oysters) Synonyms: Margaritiphora Megerle von Mühlfeld, 1811; Margarita Leach, 1814; Perlamater Schumacher, 1817; Meleagrina Lamarck, 1819 (for further information on these and other synonyms; see Hertlein 8 Cox, 1969: N304). Type species by subsequent designation (Iredale, 1915: 305): Mytilus margaritiferus Linnaeus, 1758. 476 MIKKELSEN ЕТ AL. Pinctada longisquamosa (Dunker, 1852) (scaly wing oyster) Synonymy Avicula (Meleagrina) longisquamosa Dunker, 1852; 76-77: 1872: 12 р. 2,85. 6: Avicula longisquamosa Dunker. — Petit de la Saussaye, 1856: 151 [name only]; — Beau, 1858: 21 [name only]; — Dall, 1885: 34 [name only, citing Dunker (1852), Petit de la Saussaye (1856), Beau (1858), and Krebs (1864)]. Avicula longisqvamosa Dunker [error pro longisquamosa]. — Krebs, 1864: 131-132 [name only, citing Beau, 1858]. Meleagrina longisquamosa “d’Orbigny.” — Arango y Molina, 1878-1880: 268 [name only]. Репа longisquamosa (Dunker, 1852). — Hayes, 1972 [unpubl.]: 52-58, pl. 2, fig. 2, pls. 6-8, 11f; — Abbott, 1974: 440, по. 5121; — Abbott 8 Dance, 1982: 301, fig.; - Espinosa et al., 1994: 114 [frare”, name only, citing Arango y Molina, 1878-1880]; - Camp etal., 1998: 9 [name only]; — Brewster-Wingard et al., 2001: 210-212, 214-216, 218, 220, 223- 225, 227, 228, 230; Trappe & Brewster- Wingard, 2001: fig. 3, table 1. Pinctada longisquamosa (Dunker, 1852). — Mikkelsen & Bieler, 2000: 376 (table 1). Pteria viridizona Dall, 1916a: 15 [nomen nu- dum]; 1916b: 403; — Abbott, 1974: 440, no. 5119; - Keen, 1937: 25 [“extralimital” to east- ern Pacific fauna]. Pteria viridozona [error pro viridizona] Dall, 1916. - Dall, 1921: 17; — Oldroyd, 1925: 48; — Burch, 1944: 8 [“Specimens in the Golisch collection taken from the backs of deep sea crabs off San Pedro ... a very questionable species with no member of the club sure of what it is”]; — Burch, 1945: 5 [name only; “questionable member of our [eastern Pa- cific] fauna”). Pteria xanthia Schwengel, 1942: pl. 3, fig. 1, 1a [July, nomen nudum], 64 [October]; — Aguayo 8 Jaume, 1948a: 1; — Fischer-Piette, 1982: 174. Pinctada xanthia (Schwengel, 1942). - McGinty & Nelson, 1972: 11 [‘rare”; name only]. Pinctada sp. — Brewster-Wingard & Ishman, 1999: 374 [“important Florida Bay faunal constituent”]. Pinctada radiata [non Pinctada radiata (Leach, 1814)]. - Smith, 1937: pl. 5, fig. 6; — Pulley, 1952b: pl. 4, fig. 14; — ?G. L. Voss & М. A. Voss, 1955: 226; — ?Abbott, 1958: 115; - Hudson et al., 1970: 7; — Turney & Perkins, 1972: 7, 9, 10, 12-16, 30, 31, figs. 6, 8, table 3; — Wingard et al., 1995: 7; — Ishman et al., 1996: table 2; — Brewster-Wingard et al., 1996: 18, 19, 21, 22, tables 3, 4; 1997: 9, 11, table 2; 1998a: 164, 166; 1998b: 6, 9, 12, 14, table 2, figs. 5, 6; — Brewster-Wingard € Ishman, 1999: 374-376, fig. 4. Репа colymbus [non Репа colymbus (Róding, 1798)]. — Tabb & Manning, 1961: 584. Material Examined Type Material: Avicula longisquamosa Dunker, 1852, holotype (1 pair, olive green [as originally described/figured by Dunker, 1872] with byssus attached to RV; RV 27.6 mm H, 35.9 mm L; LV 34.0 mm H, 40.2 mm L [figured by Dunker, 1872: pl. 2, fig. 6, given in text as 33 mm H, 46 mm L, probably referring to LV with lamellae]; broken and repaired according to a note by К. Kilias, dated 1972), ZMB Moll. . 101 674, from “Venezuelan beach at Puerto Cabello” (Dunker, 1852: 77, here translated). Pteria viridizona Dall, 1916b, 5 syntypes USNM 172600 (vidi, “holotype” L 25 mm, H 13 mm, D 5 mm, figured by Hayes, 1972: pl. 7), Long Beach, California [erroneous, according to Hayes, 1972], H. М. Lowe!. Pteria xanthia Schwengel, 1942, holotype ANSP 178717 (L 35 mm, Н 18 mm, exclusive of projecting lamel- lae; figured by Schwengel, 1942 [July]: pl. 3, figs. 1-1a, and Hayes, 1972: pl. 10), dredged off Captive [error pro Captiva] Island, Florida, Alice D. Miner!, December 1941. Other Material Examined: See Appendix. Distribution Bermuda, Florida (from St. Augustine to the Florida Keys to the panhandle), Texas, Baha- mas, Greater and Lesser Antilles, Caribbean coast of Mexico, Colombia, and Venezuela. Localities (*= unverified): Bermuda (USNM; Hayes, 1972; Abbott, 1974). Florida: east Florida [including St. Johns County (AMNH), Volusia County (HMNS), *Palm Beach County (Hayes, 1972), *Broward County (McGinty 8 Nelson, 1972), *Dade County (Pulley, 1952b; Hayes, 1972)], Florida Keys (AMNH, ANSP, BMSM, DMNH, FMNH, FSBC 1, UMML, USNM, this study; Hudson et al., 1970; Hayes, 1972; Abbott, 1974), Dry Tortugas (USNM, this study; Hayes, 1972), west Florida (Smith, 1937) [in- cluding Lee County (AMNH, Schwengel, 1942; Hayes, 1972), Sarasota County (AMNH); Hillsborough County (AMNH, USNM; Hayes, 1972), Wakulla County (USNM; Hayes, 1972), PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC FIGS. 2-9. Pinctada longisquamosa, showing variation in ornamentation and shell coloration. FIG. 2: Holotype (ZMB Moll. 101 674, 35.9 mm); FIG. 3: Dunker's (1872: pl. 2, fig. 6) original illustration of the holotype; FIG. 4: Living animal (FMNH 295709, 23.5 mm total L), showing tentacles at shell margin and extreme elongation of posteroventral lamella; FIGS. 5-9: Representative Florida specimens, Showing variation in shell shape, ornamentation, and color. FIG. 5, AMNH 264528, 23.7 mm; FIG. 6: AMNH 308109, 27.7 mm; FIG. 7: AMNH 308109, 23.3 mm; FIG. 8: AMNH 308109, 21.9 mm; FIG. 9: AMNH 308109, 29.2 mm. 478 MIKKELSEN ET AL. Franklin County (USNM; Hayes, 1972), *Okaloosa County (Hayes, 1972)]. Texas (HMNS, USNM). Caribbean: Bahamas (AMNH, USNM; Hayes, 1972); Cuba (AMNH, USNM; Arango y Molina, 1878-1880; Aguayo & Jaume, 1948a; Hayes, 1972; Espinosa et al., 1994), *Grand Cayman Island (?Abbott, 1958), Ja- maica (HMNS; Hayes, 1972), Puerto Rico (AMNH), Virgin Islands (AMNH), “Guadeloupe (S. Petit, 1856; Beau, 1858; Krebs, 1864), *Dominica (Hayes, 1972), Netherlands Antilles (USNM). Caribbean Central America: Mexico (USNM; Hayes, 1972). Caribbean South America: *Colombia (Hayes, 1972), Venezuela (AMNH, HMNS, ZMB; Dunker, 1852). Dimensions and Maximum Recorded Size Mean dimensions (from a single population, AMNH 308109, n = 195: length 16.34-29.75 mm, mean 23.31 + 2.64 mm SD, median 23.38 mm, mode 23.92 mm; height 14.55-25.10 mm, mean 19.86 + 2.00 mm SD, median 19.87 mm, mode 16.79 mm). Largest recorded specimen 39.34 mm L, 28.66 mm H (AMNH 308234). Diagnosis Small western Atlantic pteriid, with radial rows of narrow shell lamellae, generally bright coloration (commonly green to yellow), nacre thin (allowing external color and ornamenta- tion to show through shell), a relatively strong ridge interiorly delimiting anterior auricle of LV, and anterior dentition with tooth in RV and corresponding socket in LV; intestine with twisted loop within visceral mass and passing dorsal to heart; pallial tentacles simple. Description Shell (Figs. 2-10) obliquely ovate to round, thin-shelled, fragile (especially when dried), compressed, inequivalve with LV more strongly convex. Hinge line straight, extended into anterior and posterior auricles. Anterior auricle of RV small, triangular, with slightly concave anterior edge, strongly demarcated from anterior shell margin by groove; that of LV extending ventrally past right auricle, al- though flattened, not clearly demarcated from anterior margin and continuous with it. Pro- longed posterior auricles relatively small, slightly sinuated, supporting elongated lamel- lae. Byssal notch very narrow, present in RV only. Anterior margin (just posterior to ante- rior auricle and byssal notch) straight or slightly concave; ventral and posterior margins rounded convex and continuous with poste- rior auricle. Ventral margin (distal to nacreous . layer) of RV flexible, capable of bending dur- ing closure to create tight seal against LV, with no gap present when shell is closed. Valves concordant in color, usually green with vari- ants to yellow or brown (Figs. 6-9; from single population, AMNH 308109, n = 195: 88.7% green, 9.2% yellow, 1.5% white, 0.5% purple- brown), with commarginal (sometimes zigzag) green, brown, and/or yellow stripes of varied thickness generally corresponding to valve color but darker; sometimes indistinct. Juve- niles often with opaque white randomly dis- tributed, irregular blotches, and without conspicuous lamellae (Fig. 10). Commarginal growth lamellae inconspicuous. Umbones just posterior to anterior auricle, prosogyrous, slightly projecting beyond hinge margin, with FIG. 10. Pinctada longisquamosa, juvenile shell, showing color pattern and absence of radial lamellae (AMNH 296429, 3.7 mm). PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 479 that of LV projecting slightly more dorsally than that of RV. Umbones often without color pat- tern, turning opaque white as shell abrades with age. Prodissoconch | (Fig. 11, inset) ap- proximately 50 pm L. Subtriangular prodisso- conch || with regularly spaced commarginal growth lines, slanting slightly posteriorly, 178.1-184.4 um H, 212.5-225.0 um L, (mean 218.8 + 8.8 um, n = 2) (Fig. 11); presumably aragonitic (as shown for Р fucata martensii (Dunker, 1872) by Kobayashi, 1980). Outer shell layer simple, prismatic, comprised of single layer of regular, vertical, polygonal (mostly hexagonal) prisms, each 30 um wide, assumed calcitic as shown for other pteriids (Carter, 1990) (Fig. 12). Periostracum with concentric rings of uniform thickness, with one roughly outlining each prism (Figs. 12, 13-17). Periostracal rings close to shell margin vary- ing in shape and size, not closely adjacent to one another (Fig. 17); more proximally, reach- ing subequal size and shape (closely reflect- ing diameter of prisms), and becoming more regularly and densely arranged (Fig. 16); periostracum progressively wearing off toward older part of shell (Figs. 14-15), ultimately ab- sent in umbonal region exposing prismatic layer (Fig. 13). Each valve externally ornamented with roughly equal number of radial rows of flat, thin, elongated, flexible lamellae, from approxi- mately mid-valve to distal edge, typically ap- proximately 9 major rows per RV (from single population, AMNH 308109, п = 186: range 6- 14, mean 9.1 + 1.50 SD, median 9, mode 9; given as 10-12 by Dunker, 1872), relatively regularly spaced; progressively increasing in length toward shell margin. Lamellae usually of same color as valves but darker and often speckled with small brown dots. Rows adja- cent to anterior margin and on auricles much more narrowly spaced and comprised of very small, semitransparent lamellae more densely packed in RV than LV. Shorter lamellae in mid- valve typically oriented toward distal shell mar- gin parallel to valve surface or at slight angle (few almost perpendicular to valve surface), especially those in posterior region. Lamellae along ventral limit extending beyond shell mar- gin and curving medially, interdigitating with those of other valve; those along posterior au- ricle and posterior margin also extending be- yond shell margin but flaring laterally (with only few interdigitating) and directed slightly dor- sally. Secondary or intermediate rows often present between major rows, composed of shorter and narrower lamellae, beginning and extending more distally than major rows, ter- minating at margin thereby producing “fringed” edge. Single row of lamellae terminating at posteroventral “corner” frequently bearing dis- tally flaring lamellae (versus distally tapering lamellae in all other rows), often noticeably wider and longer than lamellae of other major rows (Fig. 4), sometimes conspicuously col- ored differently from valve (usually dark green or dark brown) (Fig. 8), often distinct only in one valve. In general, posterior lamellae longer than those on remaining part of valve, espe- cially on dorsal margin of posterior auricle. Interior shell layers nacreous, iridescent, but thinly so with external color visible through FIGS. 11, 12. Pinctada longisquamosa, details of prodissoconch and shell microstructure (SEM; AMNH 308118, 10.7 mm). FIG. 11: Left (top) and right (bottom) prodissoconch; arrow indicates prodissoconch II border. Inset, closeup of left prodissoconch, showing prodissoconch | border (arrow); FIG. 12: Shell microstructure, showing prismatic layer (P) and nacre (N). Scale bars = 100 um (Fig. 11), 50 um (Fig. 12). 480 MIKKELSEN ЕТ AL. nacreous layer. Non-nacreous prismatic mar- gin widest ventrally, approximately 1/5 of shell height, often considerably wider in RV (possi- bly to allow flexure during tight shell closure). Hinge plate narrow, widest in anterior part slightly posterior to umbo and gradually taper- ing toward posterior auricle. Ligament alivincular (restricted to hinge area), internal, lying in depression in hinge plate extending from area slightly posterior to umbones to pos- terior auricle. Anterior dentition (Fig. 18) rep- resented by single rounded crenule in RV and complementary socket in LV. Posterior denti- tion in LV represented by single elongated ridge oriented nearly parallel to hinge line origi- nating under posterior end of ligamental pit; complementary socket т RV comprised of two ridges oriented in same fashion. Anterior au- ricle interiorly often delimited on LV by very strong ridge extending from umbo to ventral FIGS. 13-17. Pinctada longisquamosa, exterior surface of LV, sequence from umbo to margin showing gradual wear of periostracum to reveal surface of prismatic layer, with shell diagram indicating relative position (SEM; AMNH 308118, 9.5 mm). Scale bars = 10 um. PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 481 FIG. 18. Pinctada longisquamosa, hinge (RV top, LV bottom) (AMNH 308118, 10.3 mm); (br, byssal ridge; c, crenule of anterior dentition; lg, ligament; pr, ridge of posterior dentition; ps, socket of posterior dentition; s, socket of anterior dentition; u, umbo). Scale bar = 1.0 mm. edge of byssal notch; corresponding ridge in RV much less conspicuous; two ridges to- gether nearly closing byssal gape when valves are closed. Adductor muscle scar (and corresponding adductor muscle) bean- to kidney- to cres- cent-shaped, with tapering dorsal end and widely rounded ventral end. Circular pedal retractor muscle and scar inset within adduc- tor muscle and scar, producing fused three- lobed scar located subcentrally along dorsoventral axis, slightly posterior to antero- posterior axis (Fig. 1). Variable number of small discontinuous pallial muscle scars of irregular shape extending (a) from dorsal ex- tremity of large adductor-retractor scar dor- sally to area ventral to posterior dentition, and (b) from anteroventral extremity of large ad- ductor-retractor scar along curve leading to pedal levator muscle scars. Two small, dis- tinct, round pedal levator muscle scars, one within umbonal arch, other slightly ventral and posterior to umbo. Adductor muscle heterogeneous, comprised of subequal anterior and posterior lobes cor- responding to “quick” and “catch” muscles re- spectively, the former comprised of denser and finer transverse fibers in cross-section (Figs. 19, 20). Two anterior pedal levator muscles narrowing distally, extending from visceral mass at posterior side of base of foot to at- tachment sites in umbonal arches of each valve. Left levator much stronger than right levator and passing anterior to the latter. Two posterior pedal levators branching off anterior pedal levators just dorsolateral to mouth, pass- ing posteriorly, attaching to valves postero- ventral to umbones. Two symmetrical pedal retractor muscles extending from root of byssal gland to valves in concavity of adductor muscle scar. Pallial muscles radiating fan-like within the mantle to its edge from attachments at pallial muscle scars on inner valve surfaces. Mantle points of attachment including pallial muscles, adductor and retractor muscles, pedal levator muscles, dorsal posterior part of labial palps, dorsal edge of outer demibranchs, and lateral surface of visceral mass. Mantle lobes fused dorsally, anterior and posterior to ligament along length of hinge line; remaining mantle margin free; inner and middle folds each equipped with single row of generally 482 MIKKELSEN ET AL. af | hi g st ddd e PINS о A à E = п ktyh gs ff tty wre plm ат ATEO, € LS af I IN? Ip amc ddd ата 20 cs br pg FIGS. 19, 20. Pinctada longisquamosa, internal anatomy. FIG. 19: Histological longitudinal section, close to midline, anterior at right (7 um, PAS stain; AMNH 298904, 18.1 mm). Scale bar = 1 mm; FIG. 20: Diagrammatic anatomy from right side, with shell, mantle, and right ctenidia removed; stomach opened from right side; other structures (foot, intestine) depicted as though transpar- ent; (af, anal funnel; alm, anterior levator muscle; am, adductor muscle; amc, adductor muscle (“catch” portion); amq, adductor muscle (“quick” portion); b, byssus; br, byssal root; cg, cerebral ganglia; cs, crystalline style; ct, ctenidia; ddd, duct of digestive diverticula; dg, digestive gland; e, esophagus; +, foot; ff, fleshy fold; g, gonad; gs, gastric shield; h, heart; in, intestine (dashed line indi- cates extent of typhlosole); ip, intestinal pouch; К, kidney; Ip, labial palps; т, mantle; mo, mouth; pg, pedal ganglia; plm, posterior levator muscle; rm, pedal retractor muscle; ss, style sack; st, stomach; ty, typhlosole; tty, tongue of typhlo- sole; vg, visceral ganglia; wr, wavy ridge). PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 483 FIGS. 21, 22. Pinctada longisquamosa, mantle margin and ctenidia (SEM; AMNH 298903, 9.7 mm). FIG. 21: Mantle margin showing simple pallial tentacles; FIG. 22: Mid-ventral portion of the ctenidia, showing food grooves on both inner and outer demibranchs (arrows). Inset, closeup of gap in outer demibranch (circle) showing interlamellar tissue connections (arrows). Scale bars = 250 um (Fig. 21), 1 um (Fig. 22). alternating large and small simple tentacles (Fig. 21), creating fringed pallial veil. Pallial veil translucent in living animals with alternat- ing bars of black and white blotches; irregular blotches of brown-orange pigment often present on and between inner and middle mantle folds. Remaining mantle lacking pig- mentation and effectively translucent. Pallial fold directed toward tips of ctenidia on poste- rior side of mantle lobes. Tentacles of inner fold ionger than those of middle fold. Larger tentacles with short pointed lateral processes in larger animals (Fig. 21). Larger inner fold tentacles in living specimens frequently with brown-orange blotches at base, this color pat- tern retained in preserved specimens. Periostracum secreted from deep groove be- tween outer and middle folds. Labial palps projecting dorsoventrally on ei- ther side of anterodorsal visceral mass sur- rounding mouth area; each consisting of pair of elongated folds, wider at base, smooth on exterior surface and plicated by about 15 trans- verse lamellae on inner surface (Figs. 19, 20). Association of labial palps and ctenidia of Category Ш of Stasek (1963), characterized by anterior filament of inner demibranch not inserted into distal oral groove. Ctenidia large, plicate, broadly sickle-shaped, encircling ven- tral half of pallial cavity. Inner and outer demibranchs subequal, each with marginal food groove (Fig. 22); eulamellibranch with regularly spaced interfilamental connections; heterorhabdic with regularly spaced (occurring at intervals of 4-7 plica), large, U-shaped (in cross-section) principal filaments connected by Interlamellar septa extending full height of demibranch and often but not always marked by brown pigmentation on dorsal edge. Neigh- boring filaments predominantly connected by continuous stretches of tissue; junctions also mediated by ciliated disks. Filaments of cor- responding ascending and descending lamel- lae joined by regularly spaced interlamellar junctions, varying in number depending on fila- ment length, reaching maximum of nine in outer and six in inner demibranch. Dorsal edges of inner demibranchs connected medi- ally via ciliated junctions from point immedi- ately ventral to foot to posteriormost extremity; dorsal edges of outer demibranchs attached laterally to mantle by ciliated junctions. Color of ctenidial ventral edge in life sometimes matching that of pallial veil (i.e., with white and dark bars), in other specimens, edge of outer demibranchs white along entire length while inner demibranchs translucent anteriorly and darker posteroventrally; such variability in color pattern can occur between ctenidia in single individual. Overall gill morphology correspond- ing to Type B (1b), characterized by frontal currents dorsalward in plical grooves and ventralward on crests (Atkins, 1937). Muscular foot emerging anteriorly from vis- ceral mass, with tip pointing dorsally and byssal groove on ventral side, extending from base to tip (Fig. 20). Foot curving noticeably to the left (observed in preserved specimens), possibly to accommodate foot extension and byssus deposition through byssal notch (RV only). Foot speckled dorsally and laterally with brown spots persisting after preservation that become darker and denser on dorsal side. Byssal gland in posteroventral portion of foot 484 MIKKELSEN ЕТ AL. (Figs. 19, 20). Golden to green shiny byssus of partially fused threads, oval in cross-section. emerging from groove in foot base, comprised Byssal threads oval in cross-section, each ter- of discrete threads extending outward; internal minating in subtriangular flat fan with rounded portion (within foot) bundled into twisted mass corners oriented perpendicular to thread axis. FIGS. 23-27. Pinctada longisquamosa, anal funnel. FIGS. 23-26: Surface, showing various degrees of surface ciliation at tip (Fig. 24), mid-region (Fig. 25) and base (Fig. 26); FIG. 27: Cross-section (7 um, PAS stain; AMNH 298904, same specimen as Fig. 19); a, anus; am, adductor muscle; f, anal funnel: in, intestine; т, mantle. Scale bars = 100 um (Fig. 23), 5 um (Figs. 24-26), 0.5 mm (Fig. 27). PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 485 Small living specimens very active in labo- ratory conditions, extending very agile foot to drag shell across surface of finger bowl. One museum label (AMNH 133648) noted, “when they were first put into a collecting jar full of salt water they moved their shells in and out, much like a bird flapping its wings”; this be- havior not confirmed in this study. Mouth concealed by dorsal and ventral lips produced, respectively, by outer and inner folds of labial palps. Esophagus connecting mouth to anterior stomach located asymmetrically in left portion of visceral mass. Overall stomach morphology corresponding to Type III of Purchon (1957) (Figs. 19, 20). “Wavy ridge” structure [as described for Pinctada vulgaris (Schumacher, 1817); Purchon, 1957], on ven- tral side of esophageal orifice but not extend- ing downward toward left anterior group of ducts to digestive diverticula. Stomach lumen divided into anterior and posterior parts by fleshy fold extending from mid-dorsal to mid- ventral section along left wall. Marginal groove on right side marking division between ante- rior and posterior regions, extending toward right side from posteroventral region above embayments leading to ducts of digestive di- verticula anterior to partition wall. Posterior part of partition wall extending anteriorly and toward right wall, emerging from ventral side of orifice _ of co-joined style sac and midgut on postero- ventral wall of stomach, connected to poste- rior part of fleshy fold by a ridge. Minor and major typhlosoles emerging from right-ventral side of style sac/midgut; minor typhlosole fus- ing to ventral side of partition wall; major typhlo- sole passing along edge of partition wall to terminate at its ventral-most part. From this point, tongue of major typhlosole accompanied by intestinal grooves on each side, passing on ventral side from area to left of embayments, making a loop around ciliated finger-like ex- tensions of fleshy fold (interpreted as food-sort- ing area by Purchon, 1957), turning left just anterior to fleshy fold, and passing along left wall to apex of ciliated food-sorting caecum. Gastric shield on left wall of posterior stomach consisting of dentate and membranous parts divided by cleft opening into stirring hollow; bordered anteriorly by dorsal hood, separated from fleshy fold by dorsal groove that branches off marginal groove in mid-lateral part of right wall, lining stomach roof, descending into stir- ring hollow just posterior to fleshy fold on left ventral side. Five groups of ducts to digestive diverticula exiting stomach: (1) one on ventral side of left pouch of posterior region (below gastric shield) extending posteriorly and left- ward; (2-3) two from embayments at anterior part of partition wall, branching posteriorly, ven- trally, and rightward; (4) one from ventral re- gion just posterior and to left of esophageal orifice, extending anteriorly and laterally; and (5) one located sub-centrally on ventral wall of anterior stomach, leading ventrally and right- ward. Intestine descending ventrally from posteroventral stomach wall into small intesti- nal pouch, turning left, twisting over itself, as- cending to posterodorsal extremity, exiting visceral mass, passing dorsal to pericardial cavity, descending to anus along posterodorsal midline of adductor muscle. Intestine terminat- ing in posteriorly oriented membranous process (anal funnel; Figs. 23-27) surrounding anus. Anal funnel flat, with tapering tip, facing ven- trally, perpendicular to posterior surface of ad- ductor muscle; anal opening at base. Distal inner surface of funnel (facing anal opening) ciliated; base smooth (Figs. 24-26). Digestive gland occupying most of visceral mass. Gonadal alveolar tissue underlying epi- thelium and enveloping visceral mass from anteriormost region just above esophagus to posteriormost visceral mass to anterior area ventral to foot. Heart (Fig. 28) within pericardium located posterior to visceral mass, dorsal to intestinal pouch (produced slightly posteriorly), anterior to upper part of adductor muscle; consisting of thick-walled ventricle attached to intestine at dorsal extremity, plus two symmetrical au- ricles situated ventral to ventricle, each with membranous extensions connecting with ven- tral pericardium and efferent blood vessels entering from posteroventral visceral mass. Major blood vessels including (1) anterior and posterior aortae, the former directed anteriorly, passing to dorsal midline over left side of in- testine, branching into visceral mass and most anteriorly dividing into two pallial arteries along mantle edge; the latter passing backward along right side of intestine, branching into interior of adductor muscle above anus, (2) paired branchial afferent and efferent vessels passing longitudinally through ctenidial axis at dorsal junction of inner and outer demibranchs. Paired nephridia laterally compressed, on ei- ther side of posterior visceral mass below heart, connected to pericardium by wide ducts at dorsal extremity. Ventral edge fused with axis of dorsal junction of inner and outer demibranchs. Nervous system conforming to general bi- valve bauplan, with three pair of ganglia (Figs. 19, 20): (1) cerebropleural ganglia surround- ing esophagus, (2) fused pedal ganglia at base 486 MIKKELSEN ЕТ AL. Ming EPA nn ei u Da Br % Pm в ne m ie FIG. 28. Pinctada longisquamosa, histological sec- tion of heart, showing relative position of intestine (7 um, PAS stain; AMNH 298904, same specimen as Fig. 19); (au, auricle; in, intestine; p, pericardial cavity; v, ventricle). Scale bar = 0.5 mm. of foot, and (3) visceral ganglia at anteroventral side of adductor muscle. Cerebropleural gan- glia connected to visceral ganglia by cerebrovisceral connectives and to pedal gan- glia via cerebropedal connectives. Branchial nerves passing dorsally to blood vessels of ctenidial axis at dorsal junction of inner and outer demibranchs; pallial nerves transversing mantle edge. Habitat and Ecology In the Florida Keys, Pinctada longisquamosa is more typical of shallow Florida Bay than of areas off the Atlantic Ocean side of the islands. It occurs on the oceanside, but only in near- shore shallows (agreeing with Abbott & Dance, 1982), not on the patch or other coral reefs. It has been found most often associated with shallow (1-2 т) Thalassia testudinum seagrass beds, but has also been recorded in Halimeda clumps, in mixed algae on mangrove roots and rocks, associated with sponge and gorgonian stalks, an artificial reef, or attached to floating Sargassum that had washed ashore. Living specimens from “eelgrass” at Rock Harbor (AMNH 133956) and Teatable Key (AMNH 308111) in the Upper Florida Keys (also reported by Hayes, 1972) are reinter- preted here as from turtlegrass, Thalassia testudinum; true eelgrass, Zostera marina Linnaeus, extends only to northern Florida (Kaplan, 1988). Individuals byssally attach, usually substantially off the silt- or sand-laden sea bottom. Hayes (1972: 146) noted that one collector described their attachment to Thalassia in Rock Harbor as “attached [by] its byssus to 3 blades ... at the point of crossing, holding the grass in the position of an elon- gated X.” Their predominantly green color has been noted as “the same as the marine plants to which they were attached” (label data, USNM 129177). Brewster-Wingard et al. (2001: 223), in reference to its occurrence on benthic macroalgae (especially Chondria and Laurencia), construed that the species “has the ability to camouflage itself to match the color of the vegetation to which it attaches.” The deepest record for living specimens in FK samples is 4 т off the bayside of Tavernier : (FK-170; in agreement with Hayes, 1972, rang- ing 1-4 m). Empty shells are a frequent com- ponent of beach drift, and were also found in one shallow offshore sediment sample from 12 m (FK-296), perhaps due to the transport qualities of its lightweight shell. P. longi- squamosa was reported in a dredge sample from 10 fms (18.3 m) off Broward County, Florida (McGinty & Nelson, 1972, as Pinctada xanthia), although the condition of the speci- mens (live-collected or empty shells) was not indicated; empty shells are also known from a dredge sample (5.5-6.1 т) in Apalachee Bay, Florida (HMNS 38564). Pinctada longisquamosa populations show some indication of seasonality at certain loca- tions. Hayes (1972: 149) noted that the spe- cies was scarce in February 1971 in an area of Biscayne Bay, Dade County, Florida, where according to collectors the species had been abundant during the previous November; she attributed its absence to “drastic changes in environmental conditions” caused by dredg- ing or pollution. During the present study, a similar situation was noted near Pigeon Key, off the Florida Bay side of Tavernier, Key Largo [not to be confused with the better-known Pi- geon Key near the center of the Seven Mile Bridge], where an especially abundant popu- lation was sampled in September and Octo- ber 1998 (FK-165, 183) and again in October 2000 (FK-368). Following a particularly cold spring, no living specimens were located at this site in April 2003 (FK-680), although nu- merous small (> 17 mm) specimens were again present two months later (FK-684, 691, 700; salinity 35 ppt) in macroalgal clumps PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 487 FIG. 29. Pinctada longisquamosa, living juvenile in situ on macroalgae (off Pigeon Key [bayside of Tavernier], Florida Keys, FK-700, approxi- mately 10 mm). within the seagrass bed (mixed Thalassia testudinum and Syringodium filiforme) (Fig. _ 29); population density in this newly recruited population was measured at 324 and 342 spm/ m? (n = 2). One of the largest recorded speci- mens was found concurrently on a dead gor- gonian stalk at 1.5 т oceanside of Key Largo in June 2003 (FK-693) suggesting that popu- lations in more environmentally stable offshore locations might serve as recruitment sources for the bay. From this anecdotal evidence, P. longisquamosa densities at this location might be fluctuating in response to seasonal tem- perature or salinity changes; the population fluctuations noted by Hayes (1972) might be similar phenomena. Such a hypothesis, how- ever, has not been rigorously investigated. More compelling evidence exists for the ef- fect of changing salinity on the presence and local abundance of Pinctada longisquamosa. Brewster-Wingard et al. (2001: 223, 224) in- cluded P longisquamosa among eight mollusks considered to be important biological indicators of environmental conditions in Florida Bay. Their “Репа assemblage” (centered on Pinctada longisquamosa) was “typically found on the sides of mudbanks (40-150 cm of water) in dense Thalassia beds, relatively clear water, and salinities between 20 and 40 ppt.” They noted that the species “seems unable to sur- vive in water of diminished quality”, and believed distribution of the assemblage to be “controlled by a combination of salinity, substrate, water depth, and water clarity.” Those authors re- ported a mean salinity of 29.3 ppt, based on 62 field observations of P. longisquamosa sites. In the 1994-1996 FMRI study, living P. longisquamosa was found in salinities from 18— 42 ppt (mean 32 ppt; 65 observations). Its dis- tribution seemed strongly influenced by the availability of suitable salinity. During the drought year of 1994, when hyperhaline condi- tions (salinity > 40-52 ppt) occurred through- out central Florida Bay (Lyons, 1998), P. longisquamosa was found only at 18 sites in the eastern bay, where more moderate salini- ties (18—30 ppt) prevailed (Fig. 30, top). In 1996, P. longisquamosa was found at 34 sites (Fig. 30, bottom), the increase attributed almost en- tirely to expansion into the central bay, where salinities returned to more moderate conditions. During both years of sampling, the species was most abundant immediately north of central Key Largo, where the highest density for the spe- cies (396 spm/m?) was recorded in 1996 in a sample site immediately northeast of that con- taining Pigeon Key sampled by Mikkelsen/ Bieler (Fig. 30, bottom). A secondary center of abundance was recorded in the central bay, where density was 156 spm/m? at one site, also in 1996. The five greatest densities recorded for Р longisquamosa in 1994 ranged from 41— 74 spm/m? at sites of salinity 36.8-41.0 ppt (mean 38.4 ppt); the five greatest densities in 1996 (41-396 spm/m?), including the two maxima mentioned above, occurred at sites of substantially lower salinity (27.0-33.4 ppt, mean 30.4 ppt). Similar patterns of retracting and ex- panding populations in response to increasing and decreasing salinity were discerned for sev- eral other Florida Bay mollusks, principally bivalves (Lyons, 1998). Brewster-Wingard et al. (2001) questioned whether the occurrence of Pinctada longisqua- mosa in relatively clear water indicates that the species favors clear water, or that the wa- ter is cleared by the filtering activity of Р. longisquamosa. Both explanations could be valid, especially in very shallow water. How- ever, P longisquamosa in the FMRI study was relatively abundant at several deeper (2.5-3.1 m) sites in northeastern Florida Bay. Water clarity at those sites was generally excellent, but similar levels of clarity also occurred nearby at sites where P. longisquamosa was uncommon or absent. In that area of the bay, high water clarity could be as much a factor of nutrient scarcity in the water column as it is a factor of bivalve abundance. 488 MIKKELSEN ET AL. Pigeon Key 1996 FIG. 30. Distribution of living Pinctada longisquamosa in Florida Bay in 1994 (top, drought year) and 1996 (bottom, year following flood year). Florida mainland is at top of each map, with Cape Sable at upper left. Cross-hatched hexagons represent 101 quantitative sites sampled by FMRI (clear hexagons not sampled); solid black hexagons are sites at which living P. longisquamosa was collected. White star and white “x” indicate sites of highest (396 spm/m?) and second highest (156 spm/m?) recorded densities, respectively. PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 489 DISCUSSION Pinctada longisquamosa has most frequently been placed in the genus Репа likely due at least in part to its oblique shell shape. In col- lections, it is frequently misidentified as one of the two other common western Atlantic pteriids, Pinctada imbricata and Pteria colymbus (Figs. 31-33). Its confusion with the former no doubt stems from its sculpture of radial lamellae. Aguayo 8 Jaume (1948b) in- cluded P. longisquamosa as a synonym of Pinctada radiata (Leach, 1814), a species gen- erally recognized today as synonymous with P. imbricata. Conchological, anatomical and ecological data for Pinctada longisquamosa, P. imbricata, and Репа colymbus are presented in Table 1. These data reveal character sets that ap- pear consistent at the generic level (I. Témkin, unpubl. data). Most informative are anterior and posterior dentition (tooth/socket in RV or LV), ornamentation (shell or periostracal), in- testinal loop (with or without twist), intestinal path (dorsal to or through heart), shape of the adductor muscle (bean-shaped or oval), and byssal structure (filamentous or stalk-like). Pinctada longisquamosa shares these char- acters in common with P imbricata and other examined species of Pinctada, and on this basis is here transferred to the genus Pinctada. Molecular data based on the 18S rDNA gene also support this conclusion, and will be re- ported in context of a larger analysis else- where. For routine identification purposes, Pinctada longisquamosa can be distinguished from Pteria colymbus by the genus-level features (dentition, ornamentation) discussed above. It is separable from Pinctada imbricata by thin- ner and denser radial lamellae, and by thin- ner nacre, allowing external color and ornamentation to show through the valve. This latter character is useful in comparison with both Р imbricata and P. colymbus, even т small subadults. The prodissoconch of Р longi- squamosa is noticeably smaller in length than that of Р imbricata (as reported by Waller 8 Macintyre, 1982; Table 1), although these data are based on very few specimens in both cases. The prodissoconch of Malleus can- deanus (Orbigny, 1842) (Pterioidea, Malleidae) is remarkably similar in size and overall mor- phology (Waller 8 Macintyre, 1982: fig. 213). FIGS. 31-33. Contrasting shell morphology in the three common western Atlantic Pteriidae. FIG. 31: Pinctada imbricata (FMNH 227467, Ohio-Missouri Key Channel, Florida Keys, coral rocks, 57.2 mm); FIG. 32: Pinctada longisquamosa (FMNH 302080, 40.1 mm including lamellae); FIG. 33: Pteria colymbus (FMNH 183297, Missouri Key, 56.0 mm). MIKKELSEN ЕТ AL. 490 —_—_=____——— un. WONOQ-yo Ajjenueysqns ‘зэлюпд$ Jayjo pue зиеиоблоб ЭУ!|-У[е}$ payoueiq yesy yuBnosu} SIM} JNOUJIM Жо! ¡eno A1Uljaxoos ‘AY Ul эбры Ny Ul 38205 ‘АЛ UI 4100} umouyun зэбри jeoesjsoued jeipes pue 91JU39UO9 чулл Jays YJOOWS А|элце|э/ pajsnjoua usyo '(uwnoessouad) umoiq Ajueulwopaid (ww 08 ed) abe} WONOQ-Yo Ajjenueysqns ‘эебе/з$елбеэз зпозиэше|у sıdwıs резч о} ¡esjop SIM} UM lleys чбполц} MoUs о} цоцезуиэшеило pue 10109 бимое ‘и!ц} рэдецз-иеэа Ay UI 1900$ ‘AT и! эбры AT UI 1200$ “AY ul 4400} бус 38/1918] |эу$ молеи ‘био| Jo SMOJ |ере/ usal6 Аплешияа (ww се чеэц/) штрэш-нец$ шп]едзап$ Jesu ‘9j “Jays ‘HOO1 зезаен sno}uswell aınyonys jesskg рэцэочела Sajoe}ue} jellled eau о} |езлор yyed ¡eulsajul }SIM} YM doo] jeursequ] 1914) SIOEN pedeys-ueaeq Ay Ч! 38008 ‘AT Ul эбры AT Ul 1920$ ‘AY U! yyoo] (2861 'SIÁUIDEIn Y JelleM) 90€ эе|эше| |эч$ apim А|элце|эл Jo SMOI |е1рел spueg syymyyoeig |еблеш yym ‘мо|эл 0} umolq 1461] (ww 09 ed) able] Jeos ajosnu лозэпрр\у цоцвиер 1011э}50а LONNUSP лоиаи\у (un ‘7 ueaw) цэцоэо$рола иоцеиэшешо 109109 |ISUS [еи.эх3 azIS SnquuiÁO9 BLIA}Y esowenbsibuo] epejould в]еэридии epej9uld ‘eepile}d4 эциеп\ чле}зэм чошиоэ ээлц} BY} JO зэцзиэелецо [8291601098 pue jeoiBojoydiow anpejeduo) | 3191 PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 491 The type locality of Pinctada longisquamosa is Venezuela, the southernmost point of the species’ geographical range. Several standard references on the malacofauna of Venezuela (Work, 1969; Lodeiros S. et al., 1999) do not list Pinctada longisquamosa, although they in- clude P imbricata and Репа colymbus. Three additional specimens from Venezuela have been examined (AMNH 203057, HMNS 30309, the latter topotypes); they differ from the type specimen by a somewhat smaller size and having a zigzag pattern of commarginal ornament. This degree of variation is within that noted for Florida specimens. Pteria viridizona, clearly conspecific with Pinctada longisquamosa but based on speci- mens supposedly from Long Beach, Califor- nia, was previously determined to be based on specimens with erroneous locality data (Hayes, 1972; Coan et al., 2000). Prior to that determination, Fischer-Piette (1982) listed P viridizona as a synonym of Pteria sterna (Gould, 1851), in turn treated by that author as a subspecies in a circumglobally distributed Pteria colymbus. Pteria xanthia is a distinctive cadmium yel- low color variant of Pinctada longisquamosa. Hayes (1972: 57) noted that some collectors have labeled such specimens as Avicula crocata (Swainson, 1831), a yellow-shelled - species described from Ceylon that has been synonymized with P. imbricata (see Ranson, 1961; Shirai, 1994). Pinctada longisquamosa is similar to Pteria vitrea (Reeve, 1857), “which it most nearly resembles, though it is not as oblique, the color is yellow instead of opaque white, and the lami- nations are much longer, though not as pro- fuse” (Schwengel, 1942: 64). Hayes (1972: 58) also noted that Р. vitrea has an extended pos- terior auricle, whereas P. longisquamosa has a short posterior auricle adorned with elon- gated lamellae. The differences in hinge teeth and ornamentation, discussed below in a ge- neric context, also serve to distinguish these two nominal species as separate. Avicula guadalupensis Orbigny (1842: pl. 28, figs. 23, 24), from Guadeloupe (Beau, 1858), was investigated as a possible earlier name for Pinctada longisquamosa. Hayes (1972: 87) deemed the description of A. guadaloupensis inadequate and the figure too stylized for posi- tive identification. Examination of the holotype (BMNH 1854.10.4.611, 1 pair), which is off-white to gray in color, indicated that d’Orbigny’s draw- ing, schematic as it is, is accurate in showing overlapping commarginal fringes of short, well- preserved prismatic scales continuous across the shell surface that are alternately arranged, as opposed to the long, radially arranged lamel- lae of P. longisquamosa. We therefore do not consider it conspecific. Full systematic revision of this and other nominal western Atlantic pteriids awaits a more comprehensive review of the genus, currently underway. The unusual periostracal structure revealed by SEM during this study does not appear to have been previously reported. This configu- ration could have resulted from the growing shell prisms pushing aside the still-fluid periostracum as they increase in diameter (J. Carter, in lit., July 2003). The general anatomical features of Pinctada longisquamosa was described and illustrated by Hayes (1972: figs. 6-8). Our results agree in all respects except in the presence of the catch component of the adductor muscle that Hayes (1972) stated as being absent. Anatomical characteristics of Pinctada longisquamosa correspond well to those de- scribed for other pteriid species, with some notable variation. In the gills of Р longisqua- mosa, interfilamental junctions consist of tis- sue plus ciliated connections, as was previously reported for Pteria argentea (Reeve, 1857) (Ridewood, 1903), Pinctada marga- ritifera (see Atkins, 1938), and Pinctada vul- garis (see Herdman, 1904, 1905). However, interfilamental junctions in Pteria hirundo are exclusively cilial except for the area uniting principal and ordinary filaments in the dorsal part of a demibranch (Atkins, 1936, 1938). The presence of laterofrontal tracts of cilia on the gill filaments and the varied nature of interfila- mental connections in these species led Atkins (1938) to consider pteriid gills as an interme- diate grade between filibranch and eulamelli- branch types. However, the extensive organic fusion in Pinctada longisquamosa is charac- teristically eulamellibranch. The ciliated lateral attachment of the dorsal inner demibranch margins with the mantle is as described for other pteriids (Herdman, 1904; Atkins, 1936). The degree of fusion var- ies among individuals from complete attach- ment to entire absence, possibly indicating an inherent weakness of the cilial junctions lead- ing to easy dissociation of the structures with minimal force. Herdman (1905: 226) described a similar condition in Pinctada vulgaris, noting that “slight pressure with dissecting-needles is generally sufficient to force the parts asun- 492 MIKKELSEN ЕТ AL. der, and they are seen to separate with clean- cut broad edges or seams and leave no ap- pearance of tearing.” Overall stomach morphology closely re- sembles that of other pteriids (Herdman, 1904; Purchon, 1957, 1985; Kuwatani, 1965). As in Pinctada martensii (but not in P. vulgaris; Purchon, 1957; Kuwatani, 1965), the major typhlosole does not enter a blind pocket on the right stomach wall before turning ventrally. Our histological study of the anal funnel of Pinctada longisquamosa confirmed previous observations in P. imbricata and Pteria colymbus (Hayes, 1972) that it consists of mem- branous loose connective tissue covered by thin epithelium; no muscular bundles or glands are present. In P longisquamosa, the anal funnel is comparatively large and extends at a right angle from the intestine and posterior surface of the adductor muscle, passing beyond the gills and mantle margin. This suggests that it plays a role in directing the passage of fecal pellets, preventing their deposition in the pallial cavity, as proposed by Hayes (1972). The distributional range of this species 1$ in part difficult to establish. Because we have found it routinely confounded with Репа colymbus and Pinctada imbricata in museum collections, we conclude that records of its occurrence in the literature cannot be trusted without verification through voucher speci- mens or published illustrations. A good ex- атр!е 1$ the presence of Pinctada longisquamosa in Texas, currently the westernmost point of its range and only U. $. record outside of Florida. Pulley's (1952b) unpublished dissertation on Gulf of Mexico bivalves figured P. longisquamosa under the name P. radiata (now P. imbricata), but the fig- ure caption gave that specimen as from Co- conut Grove, Florida, and the accompanying description could refer to either nominal spe- cies. Pulley’s (1952a: 130, pl. 7, figs. 3, 4, as P. radiata) Texas checklist definitely illustrated P. imbricata “occasionally found on Mustang and Padre Islands”, providing no support that Pulley saw P. longisquamosa specimens west of Florida. Other standard works on the mol- lusks of Texas (Odé, 1979; Andrews, 1981, and earlier editions) did not list or illustrate this species. One museum lot (USNM 465343) of P. longisquamosa was verified, but with an indeterminate locality (“Texas coast”). Finally, the HMNS collection, well known for Texas and Gulf of Mexico material, provided the needed confirmation in the form of two lots from South Padre Island (HMNS 14589, 41598). Several other published records suggest misidentified Pinctada longisquamosa but defy confirmation. Abbott (1958: 115, as P. radiata) described P. imbricata from Grand Cayman Island as “in shallow, inshore waters ... under one inch in length, very fragile and quite ob- lique in shape”. This is most similar to P. longisquamosa, although it cannot be con- firmed in the absence of a published figure. Voucher specimens for Abbott (1958) are at both ANSP and USNM (and МСА, not seen), yet none identified as P longisquamosa were confirmed by Hayes (1972), who used the USNM collection extensively, or by the authors during this study. Ekdale’s (1974: 653, as P radiata) description of P. imbricata’s habitat as “common in lagoons attached to Thalassia grass blades” in Yucatan, Mexico, is more simi- lar in our experience to that of Р. longi- squamosa than P. imbricata, although the referenced figure (Warmke & Abbott, 1961: pl. 32, fig. b) is definitely that of P imbricata. The same argument can be made about G. L. Voss & М.А. Voss (1955: 226) who listed P radiata “living on Thalassia” off the southeast quad- rant of Soldier Key off Biscayne Bay, Florida. A substantial body of literature (see syn- onymy & literature cited) has reported the oc- currence and importance of a species in Florida Bay reported principally as Pinctada radiata. However, Brewster-Wingard et al. (2001) established that the extensive records of Turney & Perkins (1972) and by Brewster- Wingard herself in earlier papers (Brewster- Wingard & Ishman, 1999; Brewster-Wingard et al., 1996, 1997, 1998b; Ishman et al., 1996; Wingard et al., 1995) were actually of P. longisquamosa, and a photographed shell by Turney & Perkins (1972: 12, fig. 6) supports that contention. Lyons (1998; unpublished) and Brewster-Wingard and her collaborators made several hundred collections of mollusks at sites throughout Florida Bay during several years in the 1990s. Pinctada longisquamosa was encountered during every year and ev- ery season (as both living specimens and dead shells in sediment cores dated to the mid- 1800s), but no other pteriid species was en- countered in the bay. Given this dominance by Р longisquamosa, it is reasonable to con- clude that other Florida Bay records (e.g., Tabb & Manning, 1961; Hudson et al., 1970) also represent P. longisquamosa. Pinctada longisquamosa appears highly (al- though not exclusively) specific to seagrass habitats. In shallow Florida Bay, this complex three-dimensional habitat provides a refuge PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 493 for the thin-shelled species against larger predators, and also provides attachment places substantially off the substratum, away from gill-fouling sediment. The pallial tentacles of P longisquamosa have few branches сот- pared to the dendritic tentacles of larger-bod- ied species (e.g., Репа colymbus, Pinctada imbricata), although it is uncertain if this is a factor of habitat, where highly branched ten- tacles can serve as a screen against siltation. Nevertheless, such a shallow-water habitat undergoes serious physical stresses associ- ated with changing water temperatures, aerial exposure during low tide, stagnation causing low dissolved oxygen levels, and nutrient- loading eutrophication (natural and anthropo- genic) during hot Florida summers (Sousa, 2001; Williams 8 Heck, 2001), and salinity ex- tremes (Lyons, 1998; Brewster-Wingard et al., 2001). Anecdotal observations on populations during this study also suggest that unusually cold Florida winters are similarly impactful. Re- gardless of the effect of these factors on the pteriids themselves, seasonal sloughing off of older Thalassia blades probably influences the size and structure of Р longisquamosa popu- lations on a nearly annual cycle. The aperi- odic die-offs of Thalassia beds that have occurred in Florida Bay in recent years (Robblee et al., 1991) must have been accom- - panied by local extirpation of pteriids and other seagrass dwellers from those affected areas. ACKNOWLEDGMENTS This work is part of an ongoing investigation of the molluscan diversity of the Florida Keys; regional surveys and collections were sup- ported by permits from the Florida Keys Na- tional Marine Sanctuary (080-98, 2000-036, 2002-078, and 2002-079); within John Pennekamp Coral Reef State Park under Florida Department of Environmental Protec- tion (FDEP) permit 5-01-22; Everglades Na- tional Park under National Park Service permit 2000073 to Tom Frankovich; in Dry Tortugas National Park under permit DRTO-19970030 to Tim Collins; in Long Key State Park (Long Key, Florida Keys) under FDEP permit 5-02- 43; and additional collecting under Florida Fish and Wildlife Conservation Commission permit 99S-024 to affiliates of The Bailey-Matthews Shell Museum (Sanibel, Florida) and permit 01S-056 (as well as annual permits for prior years of this study) to affiliates of the Smithsonian Marine Station (Ft. Pierce, Florida; logistic support by Mary E. Rice and staff is much appreciated). For collecting assistance, we thank Timothy Collins, Timothy Rawlings, Tom Frankovich, Roberto Cipriani, Deirdre Gonsalves-Jackson, Cecelia Miles, Brett Kubricht, Rebecca Price, Russell Minton, Louise Crowley, Juri Miyamae, Ed Yastrow (and his boat PATCH HAPPY), Raymond Baiz (and his commercial fishing vessel STRANGE BRU), staff of the Keys Marine Laboratory (Layton, Long Key), the captains and crews of R/V EUGENIE CLARK (Mote Marine Laboratory, Sarasota, Florida) and R/V BELLOWS (Florida Institute of Oceanography, St. Petersburg, Florida). David Eaken, Nancy Diersing, Kim Bjorgo, Robert Tempe, David Camp, and Ed Matheson provided major sampling support during the FMRI Florida Bay field program, and several other persons are thanked for filling in during brief periods; staff of the Keys Marine Laboratory and the FMRI South Florida Re- gional Laboratory at Marathon also provided logistic support and other assistance that fa- cilitated the sampling effort. Data gathering from other museum collections was facilitated by Gary Rosenberg (ANSP), Matthias Glaubrecht (ZMB); José Leal (BMSM), John Wise (HMNS), Nancy Voss (UMML), and Timothy Pearce, Leslie Skibinski, and Albert Chadwick (DMNH). Photography was facilitated by David Grimaldi and Tam Nguyen (AMNH; light photography) and Kevin Frischmann (AMNH; SEM). Joseph Carter (University of North Carolina, Chapel Hill) offered comments on the periostracal structure described herein. This research was supported in part by NSF-PEET DEB-9978119 and Comer Science and Education Founda- tion grants to RB and PMM. Additional field- work support from the Bertha LeBus Charitable Trust, Field Museum’s Women’s Board, Field Museum's Zoology Department’s Marshall Field Fund and AMNH’s Proctor-Old-Sage Ma- lacology Fund is also gratefully acknowledged. LITERATURE CITED ABBOTT, R. T., 1958, The marine mollusks of Grand Cayman Island, British West Indies. Monographs of the Academy of Natural Sci- ences of Philadelphia, 11: 1-138, pls. 1-5. ABBOTT, R. T., 1974, American seashells: the marine Mollusca of the Atlantic and Pacific coasts of North America, 2nd ed. Van Nostrand Reinhold, New York. 663 pp., 24 pls. ABBOTT, Б. Т. & S. P. DANCE, 1982, Compen- dium of seashells: a color guide to more than 4,200 of the world’s marine'shells. Е.Р. Dutton, New York. 411 pp. 494 MIKKELSEN ET AL. AGUAYO, С. С. 8 М. L. JAUME, 1948a, Pele- cypoda - Pteriidae. 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Florida Keys Diversity Study: FK-021, Lake Surprise, Key Largo, mile marker 107.5, NE end of U.S. Rte. 1 causeway across lake, 25°10.9’М, 80°23’W, off mangroves at side of road, hand dredge in sediment/algae, ca. 1.5 m, and by hand on shallow subtidal rocks, S = 22 ppt, 09 July 1995 (6 juv valves, AMNH 296437); FK-032, “Billboard” site, Lower Matecumbe Key, mile marker 74.5, oceanside, 24°51.4’N, 80°43.7’W, hand picked from flats and wrack line, 08 March 1996 (1 juv pair, AMNH 308210); FK-035, Indian Key Fill, mile marker 79, bayside, 24°53.4’N, 80°40.5’W, Thalassia, 1 m, shovel and sieve, 10 March 1996 (1 pair, AMNH 296438); FK-036, same as FK-032, hand scooped from wrack line on beach, 10 March 1996 (1 pair, 8 juv valves, АММН 308209); FK-043, same as FK-021, off mangroves at side of road, shovel/sieve in sediment/algae (ca. 1.5 m) + shallow subtidal rocks by hand, 19 September 1996 (1 valve, FMNH 279467); FK-064, center of Coupon Bight, bayside off Big Pine Key/Newfound Harbor, 24°38.6’М, 81°22.2’W, 1.5 m, mud/ Thalassia, bottom grab, R/V FLORIDAYS, 15 April 1997 (1 juv spm alc, FMNH 295709; 4 valves, FMNH 279466); FK-126, on old lob- ster traps being sold near “Billboard” site, Lower Matecumbe Key, mile marker 74.5, traps formerly used in “Florida Keys waters”, 24 July 1997 (1 pair, AMNH 308099); FK-128, Rachel Shoal, bayside off Key Vaca, 24°45’N, 81°05'W, sand with algae/sponges, 1.5 m, dredge (2 tows), R/V FLORIDAYS, 25 July 1997 (1 pair, FMNH 279462); FK-136, E of Bethel Bank, bayside, 24°44.8’N, 81°04.7’W to 24°44.7’М, 81°04.7’W, sand/sparse seagrass, gorgonians, algae, 1.8-2.1m, 10 min dredge, R/V FLORIDAYS, 07 July 1997 (1 valve, AMNH 296433); FK-163, Tavernier, Key Largo, bayside, 25°03.6’N, 80°30.0’W to 25°03.5’М, 80°30.2’W, sandy mud/ Thalassia/ chicken liver sponge/Dasycladus, 1.7 m, dredge, R/V FLORIDAYS, 12 September 1998 (13 juv spm alc, AMNH 298903; 22 pair, 34 valves, AMNH 296432); FK-165, off W shore of Pigeon Key (bayside of Tavernier), 25°03.3’N, 80°30.8'W, Thalassia, 1.4 m, scuba, by hand, R/V FLORIDAYS, 13 Septem- ber 1998 (32 spm alc, 2 valves alc, AMNH 298904); FK-166, Tavernier, Key Largo, bayside, mile marker 95.5, 25°03.2’N, 80°29.1’W, on Acetabularia on rocks, 0-1 m, by hand, 14 September 1998 (2 spm alc, FMNH 295714); FK-168, Tavernier Creek, near bayside entrance, west side (Plantation Key), 25°00.7’М, 80°32.7’W, mangrove root scrapings, snorkeling, R/V FLORIDAYS, 17 September 1998 (1 spm alc, FMNH 295715); FK-170, Hawk Channel, E of mouth of Tavernier Creek, 24°58.8’N, 80°30.9’W to 24°58.7’М, 80°30.8’W, Thalassia droves, 3.3— 4.0 m, dredge, two tows of 3 min each, R/V FLORIDAYS, 17 September 1998 (2 juv spm alc, FMNH 295718); FK-172, Cowpens An- chorage, bayside of Plantation Key, 24°59.1'N, 80°34.4'W, grey soupy mud/Thalassia, 1.8- 2.1 m, ponar grab and dredge, R/V FLORI- DAYS, 18 September 1998 (4 spm alc, FMNH 295716; 2 pair, 4 valves, AMNH 296428; 9 valves, FMNH 288688); FK-174, same as FK- 165, Thalassia, 1.4 m, scuba, by hand, R/V FLORIDAYS, 18 September 1998 (12 spm alc, AMNH 298902; 100 spm [empty] ас, ЕММН 290099); FK-181, Rock Harbor, Key Largo, mile marker 98, bayside, 25°04.7’М, 80°27.7’W, shallow water off beach, seagrass/ sand/shell gravel, snorkeling, 08 October 1998 (11 spm alc, ЕММН 288839); FK-183, N of Key Largo, S end of Pigeon Key, bayside, 498 MIKKELSEN ET AL. 25°03.3’М, 80°30.6’W, dense to sparse seagrass, 0.3-1.5 m, snorkeling, R/V FLORIDAYS, 10 October 1998 (20 spm alc, ЕММН 288845); FK-184, off Key Largo, W side of North Nest Key, bayside, 25°09.1’М, 80°30.8’W, seagrass flats near dock, R/V FLORIDAYS, 12 October 1998 (spm obs, 6 pair, ЕММН 288613); FK-186, NW of Planta- tion Key, Cowpens Cut, bayside, 24°59.8’М, 80°33.6’W, seagrass beds, 1.2-1.8 т, hand- sieving, R/V FLORIDAYS, 14 October 1998 (1 spm alc, FMNH 288867); FK-188, Black- water Sound, east of Bush Point, 25°08.8’N, 80°25.3’W, seagrass, 1.8 m, bottom sample, R/V FLORIDAYS, 14 October 1998 (13 spm alc, FMNH 288844); FK-191, Key Largo, bayside, small mangrove key off Hammer Point, near border of Everglades National Park, 25°02.0’М, 80°31.3’W, wrackline to 0.3 m, at extreme low tide, mats of brown and green algae, R/V FLORIDAYS, 31 March 1999 (7 juv ас, AMNH 298905; 4 pair, AMNH 296435); FK-192, same as FK-184, clay and sand beach, mangroves, by hand, R/V FLORIDAYS, 01 April 1999 (5 pair, FMNH 279460); FK-193, off Key Largo, bayside, E of Buttonwood Sound, Swash Keys, E shore of unnamed middle key between Whaleback Key and Shell Key, 25°07.3’М, 80°28.8’W, clay and sand beach, mangrove roots with oysters, by hand, R/V FLORIDAYS, 01 April 1999 (3 pair, AMNH 296431); FK-201, Key Largo, bayside, mile marker 95.5, 25°03.2’N, 80729.1'W, sand/wrack line, intertidal, by hand, 06-13 April 1999 (8 juv pair, АММН 296429); FK-209, same as FK-191, intertidal algae/ seagrass/mangrove roots, by hand, R/V FLORIDAYS, 11 April 1999 (9 juv spm alc, 1 juv pair, FMNH 295717); FK-210, same as FK- 032, beach drift, 13 April 1999 (1 pair, AMNH 296430); FK-211, unnamed bay between Shark Key and Big Coppitt Key, bayside, 24°36.4’М, 81°39.2’W, Thalassia, bottom sample, R/V FLORIDAYS, 17 April 1999 (1 juv valve, FMNH 279470); FK-224, W end of Mis- souri Key, oceanside, 24°40.5’М, 81°14.3’W, rocks and beach line, 0-1 m, by hand, 20 April 1999 (1 pair, FMNH 279465); FK-226, Saddlebunch Harbor, off W tip of Saddlebunch Keys, 24°35.2’N, 81°37.9’W, sand with vari- ous algae and sponges, Thalassia, 0.3 m, hand dredge, R/V FLORIDAYS, 20 April 1999 (1 frag, FMNH 279463); FK-249, E end of Ohio Key, oceanside, mile marker 39, 24°40.3’N, 81°14.5’W, beachcombing in wrack line, 05 August 1999 (3 valves, AMNH 308098); FK- 255, Friend Key Bank, bayside of Bahia Honda Key, N side at crest of bank, 24°42.6'N, 81°16.8'W, Thalassia/Syringodium with sand patches, 0.15-0.6 т, snorkeling, R/V FLORIDAYS, 09 August 1999 (1 alc, FMNH 289986); FK-258, Friend Key, bayside of Ba- hia Honda Key, off М shore, 24°43.4’N, 81°17.3'W, sparse Thalassia with sponges and sand patches, 0.3-0.6 т, snorkeling, R/V FLORIDAYS, 09 August 1999 (1 valve, AMNH 296434); FK-270, Ohio Key camp- ground, mile marker 39, bayside, 24*40.5'N, 81°14.7’W, mangrove shore, by hand, 19 Au- gust 1999 (1 pair, 3 valves, AMNH 296436); FK-278, Ohio Key, ocean side, beach facing Seven Mile bridge, 24°40.3’М, 81°14.5’W, beach drift, algae, silt-covered platform ex- posed at low tide, by hand, 06 April 2000 (1 pair, AMNH 308101); FK-286, same as FK- 201, by hand, sand/wrack line, intertidal, 09 April 2000 (2 juv valves, AMNH 308103); FK- 289, Cowpens Anchorage, bayside of Plan- tation Key, 24°58.7’М, 80°34.5’W, sand/sparse seagrass/Penicillus, 1.7 m, petit ponar grabs, M/V PATCH HAPPY, 11 June 2000 (8 valves, AMNH 308100); FK-353, “Horseshoe” site (outside S arm), Spanish Harbor Keys, 24°39.3’М, 81°18.2’W, gulf side, to 1.5 m, hand dredge, Thalassia/Halodule seagrass, 08 July 2000 (1 valve, AMNH 299574); ЕК- 357, American Shoals, NW of lighthouse, 24°31.5'N, 81°31.2’W, Thalassia with large coral rubble, 2.7-3.3 m, scuba, R/V FLORIDAYS, 09 July 2000 (1 pair, AMNH 299590); FK-368, off W shore of Pigeon Key (bayside of Tavernier), 25°03.3’М, 80°30.7'W, live-collected in Thalassia and Syringodium seagrass, 0.3-0.9 m, snorkeling, R/V FLORIDAYS, 08 October 2000 (ca. 400 spm, AMNH 308109 and FMNH 302080); FK-392, same as FK-032, wrack line, by hand, 20 October 2000 (1 valve, AMNH 308102); FK- 428, Infaunal Mollusk Survey, Windley Key transect sta. W-14-FK-428, 24%53.3'N, 80°31.5’W, 46.9 m, shelly mud, small pipe scoop, M/V STRANGE BRU, 25 April 2001 (1 valve, 1 fragment, AMNH 296439); IMBW-FK- 629, “The Horseshoe” site, bayside of West Summerland Key (Spanish Harbor Keys), mile marker 35, 24°39.3’N, 81°18.2’W, among rocks along arms of quarry, by hand, snorkel- ing, to ca. 1 m, 21 and 26 July 2002 (2 pair, AMNH 308106); IMBW-FK-649, Sprigger Bank, bayside, just W of Everglades National Park border, 24°54.7’N, 80°56.2’W, Thalassia/ Syringodium, snorkeling, shovel/sieving, 0.1— PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 499 0.9 m, Keys Marine Laboratory boat, 27 July 2002 (1 juv pair with dried tissue, AMNH 308105); IMBW-FK-652, Long Key State Park, oceanside, 24°48.7’М, 80°49.7’W, seagrass bed (predominantly Thalassia) on muddy sand, > 1 m, wading, shovel/sieving, 27 July 2002 (1 pair, AMNH 308107); IMBW-FK-656, mangrove channel off South Layton Drive, oceanside of Long Key, 24°49.4’N, 80°48.8’W, red mangrove roots, snorkeling, 0.6-1.2 m, 28 July 2002 (4 spm alc, AMNH 305172); FK-680, W of Pigeon Key (bayside of Tavernier), 25°03.304’N, 80°30.715’W, 2 ft (0.6 m), sweep- net through mixed algae on Thalassia seagrass, R/V FLORIDAYS, 12 April 2003 (1 valve, AMNH 308097); FK-681, bayside of Tavernier, off small mangrove island near bor- der of Everglades National Park, 25°01.95’N, 80°31.17 W, 2 ft (0.6 т), pavement with sand, sparse to thick Thalassia seagrass, snorkel- ing, by hand, R/V FLORIDAYS, 12 April 2003 (26 pair, 40 valves, AMNH 308496; 7 pair, FMNH 296727); FK-683, W of Tavernier Creek, bayside, fast-flowing mangrove-lined tidal channel in midst of Cross Bank leading into Cowpen’s Cut, 25°00.345’N, 80°33.468’W, 0.5-1.0 т, snorkeling, Thalassia seagrass, R/V FLORIDAYS, 03 June 2003 (1 juv pair, 60 juv valves, AMNH 308120); FK-684, W of Pigeon Key (bayside of Tavernier), _ 25°03.304’N, 80°30.715’W, 0.6 m, snorkeling, dipnet, mixed algae on Thalassia seagrass, R/V FLORIDAYS, 03 June 2003 (12 alc, 11 juv pairs, 1 fragment, AMNH 308119/308235/ 308236); FK-690, E of Pigeon Key (bayside of Tavernier), 25°03.400’N, 80°30.617’W, 0.9 m, snorkeling/dipnet, sparse Thalassia/ Halodule seagrass with algae, some Gracilaria, R/V FLORIDAYS, 07 June 2003 (2L alc, AMNH 308231); FK-691, W of Pigeon Key (bayside of Tavernier), 25°03.304’N, 80°30.715’W, 0.6 m, snorkeling, dipnet, mixed algae on Thalassia seagrass, R/V FLORIDAYS, 07 June 2003 (89 alc, 25+ pair, 100+ valves, AMNH 308233); FK-693, off Dove Key, oceanside of Key Largo, 25°03.039’М, 80°28.151’W, 0.9-1.5 т, snor- keling, silty Thalassia seagrass and sand, at top of dead gorgonian stalk, R/V FLORIDAYS, 08 June 2003 (1 alc, largest recorded speci- men, AMNH 308234); FK-700, W of Pigeon Key (bayside of Tavernier), 25°03.287’N, 80°30.693’W, 0.3-0.6 m, snorkeling, seagrass with algae, R/V FLORIDAYS, 12 June 2003 (1 pair, 2 valves, AMNH 308104; 70+ juvs alc, FMNH 302081). Other material examined: Bermuda: Bermuda, entrance of Harrington Sound, H. G. Richards! 29 June 1932 (2 pair, USNM 422492). Florida: Florida, Velie!, “the color when living is green — the same as the marine plants to which they were attached” (4 pair with tissue, 3 pair, USNM 129177). East- ern Florida: 20 mi S of St. Augustine, live- collected on beach in seaweed, May 1968, Miriam K. Hicks! (2 juv pair, AMNH 249011); S of Matanzas Inlet, on seaweed at high tide mark, Fall 1970, Miriam K. Hicks! (2 pair, AMNH 248983); Ormond Beach, 1955 (3 pair, HMNS 27356); off Ormond Beach, Betty Allen! 1977 (3 pair, HMNS 19433); Coconut Grove, at Pan Am Airport, 4 ft (1.2 m), Fostor! 1936 (1 spm, 1 valve, HMNS 201); Chicken Key, 6 mi S of Coconut Grove, R. Foster! (1 pair with piece of sponge (presumed substra- tum), AMNH 200267). Florida Keys: Broad Creek, Bean! 24 December 1906 (1 pair with tissue, USNM 198115); Cormorant/Jew Points, mangrove roots among Caulerpa verticillata Agardh, 07 October 1978, P. M. and P. S. Mikkelsen! (1 juv som, DMNH 179666; 6 juv alc, 8 juv pair, AMNH 308498); lower part of Barnes Sound, Harry Balknap! (2 pair, AMNH 67013); Barnes Sound, H. A. Rehder! 20 April 1947 (2 pair, USNM 778259); Key Largo, Feb- ruary 1944 (4 pair, AMNH 200266); Key Largo, Atlantic side, shallow, “found feeding on grass of bay”, J. M. Bijur collection (6 pair, AMNH 232620); Key Largo, May 1940, J. Donovan! Grace G. Eddison collection (2 pair, AMNH 293718); NW Key Largo, mangrove channel, on mangrove roots, P. S. Mikkelsen! 07 Octo- ber 1978 (3 alc); PSM-811, Key Largo, N of Rock Reef Motel, Thalassia seagrass and al- gae, by hand, 0.7-1.5 m, Р. $. Mikkelsen! 8 and 10 February 1982 (12 juv alc, 3 juv pair, AMNH 308500); Key Largo Sound, J. M. Bijur collection (3 pair, AMNH 232622); Key Largo, Largo Sound, 08 December 1956, Craig and Fanny Phillips! (1 pair with tissue, FMNH 62888); Key Largo, Molasses Reef (1 pair, AMNH 100868); Key Largo, 2 mi W of Rock Harbor, Florida Bay, alive on eelgrass (sic), 2 ft (0.6 m), firm sand bottom, November 1961, D. Raeihle! A. D’Attilio Colln (2 pair, AMNH 133956); Key Largo, 2 mi S of Rock Harbor, Florida Bay, alive on turtlegrass, 2-3 ft (0.6- 0.9 m), firm sand bottom, November 1961, D. Raeihle! (9 pair, AMNH 106140); Florida Bay, Bottlepoint [now Bottle] Key, R. P. Allen! (1 pair, USNM 533677); Key Largo, Harris Beach, W. 500 MIKKELSEN ET AL. S. Bitler! (3 pair, AMNH 308211); Key Largo, bar on gulf side, opposite Tavernier, Vilas! (7 pair, BMSM 26118); Key Largo, bar on gulf side, opposite Tavernier, Vilas collection (1 pair, BMSM 26119); Florida Bay, May 1964— July 1965, Tabb and Manning! (many small pairs, UMML 30.8885, 8890, 8893, 8916, 8922, 8930, 8936); Upper Matecumbe, oceanside, on beach, HC Porreca! (5 pair, AMNH 139428); Islamorada, Florida Straits, H. S. Feinberg! (5 pair, AMNH 308207); Islamarada (sic), 1967, Alice Denison Barlow! (5 pair, AMNH 244070); Islamarada (sic), March 1963, Mrs. Ward!, H. C. Porrecca col- lection (1 pair, BMSM 26117); east of Teatable bridge, living on eelgrass (sic), 1959, Raeihle collection (2 pair, AMNH 308111); Lower Matacumbe (sic) Key, 1956, Elinor Townsend!, Dale V. Stingley collection (2 pair, BMSM 26116); Islamorada, off mangrove roots, Eleanor Townsend! (10 pair, AMNH 139427); Lower Matacumbe (sic) Key, February 1952, Mary Brevillier! Dale V. Stingley collection #593 (6 pair, BMSM 26114); Lower Matecumbe Key, January 1965 and 1966, H. C. Porreca! (9 pair, BMSM 26123); Matecumbe Key, Wisoff col- lection (1 pair, AMNH 120106); Matacombe (sic) Key, February 1965, Bert Porrecca! (7 pair, BMSM 26113; 8 pair, BMSM 26121; 16 pair, BMSM 26122); Key Vaca, Marathon, B. R. Bales collection, donated by J. Schwengel 1958 (1 pair, ANSP 222085); Marathon, Mrs. H. McGill! via T. L. Moise (1 pair, ANSP 193909); off Marathon, on old горе (2 pair, FMNH 159957); Marathon, ocean|[side], ropes of lobster traps, E. M. Malone collection (1 juv pair, BMSM 26120); Marathon (1 pair, BMSM 26115); N of Knight’s Key [bayside], channel, 11 fms (20.1 m), U.S. Fish Commission Fishhawk sta. 7412 (1 pair with tissue, USNM 198142); Lower Florida Keys, 1947, A. Koto! ex Wickham (1 pair, FMNH 183214); PSM- 786, Big Pine Key, seawall off Bogie Channel, subtidal attached algae, by hand and sieving, 0.2 m, P. S. and P. M. Mikkelsen! 16 August 1981 (1 juv alc, AMNH 308501); Boca Chica, Н. A. Pilsbry! (2 pair, ex ANSP 100274); Boca Chica, fresh-dead in drift of turtlegrass, No- vember 1963, Raeihle collection (6 pair, AMNH 308110); Key West, pulled onto boat on sea- weed while fishing, A. D. Barlow collection (5 pair, AMNH 244074): Key West, 1937, A. Koto! (5 pair, FMNH 288737); Key West, South Beach, 18 February 1947, Mrs. Ward Brown!, ex J. D. Parker (2 pair, ex ANSP 182650); Key West, especially E side, Jan.-Feb 1958, M. Bogart! (14 pair, AMNH 184816); Key West, December 1938, Dr. and Mrs. Julius Wisof! (1 pair, AMNH 308208); Key West, Mrs. Eshnaur! (3 pair, 1 valve, USNM 404236); Key West, A. E. Mehring! sta. 6-B-5, 29 December 1949 (8 pair, USNM 700383); Key West, А. D. Clark!, June 1958 (1 pair, HMNS 199); SW of Tortugas, 35 fms (64.0 m), Schmitt! 14 August 1933 (1 pair, USNM 421671). Western Florida: Sanibel Island, О. Germer! (11 pair, AMNH 264528); Manasota Key, Butler!, Johnstone collection (1 juv pair, AMNH 210312); Tarpon Springs, brought in by sponge fishers, “sent in for identification by William Rhodes, School for the Blind, Indianapolis, Indiana” (1 pair, AMNH 70803); Tarpon Springs, P. Bartsch! 1936 (5 pair, USNM 428700); Apalachee Bay, dredged, 18-20 ft (5.5-6.1 т), near artificial reef, Gleeson, Keeler & Loftin! 30 April 1987 (1 pair, HMNS 38564); off Live Oak Island, Wakulla County, on scallop shells, J. Rudloe! 11 November 1970 (1 valve, USNM 70690); Gulf coast, Apalachicola, sta. 304/305, A. S. Pearse! June 1935 (1 pair with tissue, USNM 467928). Texas: Texas coast, J. D. Mitchell! (8 pair, 3 valves, USNM 465343); Port Isabel (gulf beach on South Padre Island), ex Betty Allen via C. E. Boone! (2 pair, HMNS 14589); South Padre Island, beach drift, W. W. Sutow! August 1966 (1 juv valve, HMNS 41598). Ba- hamas: Bahamas, 1904, Prof. W. М. Wheeler! (26 pair, 3 valves, with tissue, AMNH 27897); Bahamas, Prof. W. Wheeler! (6 pair with tis- sue, 3 valves, AMNH 27808); Andros, sta. wp126-Q5, transect from 24°54.8’М, 77°53.3'W to 24°54.8’М, 77°53.2’W, quadrat #5, Thalassia, 10 ft (3.0 m), scuba, P. M. Mikkelsen and G. Hendler! 03 September 2000 (1 valve, AMNH 308117); Andros, sta. wp126- R, 24°55.4’N, 77°54.4’W, Thalassia, 10 ft (3.0 т), scuba/snorkeling, P. M. Mikkelsen et al.! 03 September 2000 (3 spm alc, AMNH 308499); Andros, sta. мр415-К sediment, 24°53.5'N, 77°53.8'W, thick Thalassia, 12 ft (3.6 m), scuba/snorkeling, P. M. Mikkelsen et al.! 04 September 2000 (2 juv valves, AMNH 308497); Andros, sta. PMM-1039, on beach in front of Forfar Field Station, on Sargassum washed ashore, by hand, P. M. Mikkelsen! 28 August 2000 (12 pair, AMNH 308118); Andros, inside Golding Key, P. Bartsch! 03 May 1912 (1 pair with tissue, USNM 269347); Spanish Wells, across from Galliot Cay, NW end of Cape Santa Maria, Long Island, Mrs. J. Stout! 12-18 March 1967 (1 pair, AMNH 136184); Eleuthera, M. Bogart! (1 pair, AMNH 184885); PINCTADA LONGISQUAMOSA IN THE WESTERN ATLANTIC 501 shores E of Fox Hill, Nassau, February 1928, William $. Treator! (1 pair, АММН 270630); W shore of North Bimini, near entrance to Cavelle P[on]d, on tips of mangrove roots out of water at very low tide, W. Schwarting, 20 April 1950 (2 pair with tissue, AMNH 87385). Cuba: on flats between Cuba and Isle of Pines, live-col- lected on sea weed, 1 ft (0.3 т), Е. H. Low! (7 juv pair, AMNH 113648); Santa Lucia, NW Cuba, 2-4 fms (3.6-7.3 т), Barrera Expedi- tion sta. 200 (7 pair, USNM 456990); Punta Tolete, NW Cuba, 2-3 fms (3.6-5.5 т), Barrera Expedition sta. 205 (5 pair, USNM 45683); sand bar off Arroyos, NW Cuba, Barrera Ex- pedition sta. 206 (1 pair with tissue, 1 pair, 7 valves, USNM 457007); Santa Rosa, NW Cuba, 3-6 fms (5.5-11.0 m), Barrera Expedi- tion sta. 209 (14 pair, 43 valves, USNM 456989); Cape Cajon, NW Cuba, Barrera Ex- pedition sta. 211 (7 pair with tissue, 4 pair, 14 valves, USNM 456987); Varadero Beach, Cuba, Barrera Expedition sta. 213 (1 pair, USNM 456985); Esperanza, NW Cuba, 4-6 ft (1.2-1.8 m), Barrera Expedition sta. 217 (1 pair, USNM 456986); Bay of Santa Rosa, NW Cuba, 1-3 fms (1.8-5.5 m), Barrera Expedi- tion sta. 219 (2 pair, USNM 456982); Los Ar- royos, NW Cuba, 3 fms (5.5 т), Barrera Expedition sta. 229 (1 pair, USNM 456988). Jamaica: Kingston, south shore, Betty Walden! _ September 1956 (1 valve, HMNS 15401). Puerto Rico: Mangrove Island, W of Magueyes (sic) Island, on Rhizophora mangle, 13 Sep- tember 1962, H. E. Coomans! (1 pair with tis- sue, AMNH 109620); E of Carib Cayo, 5.5-8.75 fms (10-16 т), 25 June 1915, R. С. Osburn! (1 juv pair, AMNH 1036). Virgin Is- lands: St. Thomas, Alice Denison Barlow! (2 pair, AMNH 244072). Mexico: in bay E of larger island S of village, Mujeres Island, Quintana Roo, dredged, eelgrass bottom, Bredin-S.!. Expedition sta. 26-60, 31 March 1960 (4 pair with tissue, 1 valve, USNM 662550); Allen Point, Ascension Bay, Quintana Roo, Bredin- S. |. Expedition sta. 68—80, Schmitt! 13 April 1960 (1 pair with tissue, USNM 736070); shore of small bay behind Halfway Point, N end As- cension Bay, Quintana Roo, Bredin-S.l. Ex- pedition sta. 76-60, Bousfield and Rehder! (2 valves, USNM 736096); shore near Halfway Point, N end Ascension Bay, Quintana Roo, Вгедт-$.1. Expedition sta. 77+93, Schmitt et al.! 15, 18 April 1960 (1 pair with tissue, USNM 736157). Netherlands Antilles: Curacao, Spaanse water near Brakkeput Ariba, on Isognomon alatus in mangroves, К. В. Meyer! 07 February 1971 (3 pair, USNM 702277). Venezuela: Chichiribiche, live-collected on al- gae washed ashore, August 1975, Edo. Falcon! F. Fernandez H. collection (2 pair, AMNH 203057); Puerto Cabello, R. W. Barker! October 1949 (1 pair, HMNS 30309). MALACOLOGIA, 2004, 46(2): 503-544 MARINE BIVALVES OF THE FLORIDA KEYS: A QUALITATIVE FAUNAL ANALYSIS BASED ON ORIGINAL COLLECTIONS, MUSEUM HOLDINGS AND LITERATURE DATA Rudiger Bieler' & Paula М. Mikkelsen? ABSTRACT Marine bivalve biodiversity in the waters surrounding the Florida Keys, an island archi- pelago off southern Florida, including the Florida Keys National Marine Sanctuary, was studied from ten years of original collections as well as from a critical review of museum specimens and literature data. A database of more than 12,000 records representing 389 species (half of which were ranked as abundant or common) was assembled and ana- lyzed, resulting in a 139% increase of the known bivalve fauna of this region compared to the most recent prior (1995) checklist. Of the 389 species, 42% have not been positively recorded as live-collected, and 12.5% are represented only as singletons or doubletons. Using multivariate non-metric statistics and a priori geographic groupings along the island chain (Upper, Middle, Lower Keys; Dry Tortugas) and across the island chain (Florida Bay, shallow Atlantic waters [< 35m], deeper Atlantic waters [35-300 m]), the data showed distinct differences in benthic community structure across several spatial gradients. A pro- nounced northeast-to-southwest gradient was found on the Florida Bay-side of the island chain, although none was evident along the oceanside in either shallow or deep depth zones. Although they shared dominant species, the shallow-water communities of bayside and oceanside differed significantly in the percentage distributions of co-occurring spe- cies. In contrast, the deeper oceanside community differed substantially from both shal- low-water groups in supporting a different set of species. A comparison of the bivalve fauna of the Keys with other well-documented faunas of the western Atlantic indicated that the Florida Keys fauna groups more closely to the Gulf of Mexico and Cuba than to eastern peninsular Florida, Yucatan, or the Bahamas. The impact of the heterogeneous nature of the dataset (live-specimen, dead-shell; and original collections, museum, and literature) is discussed and compared to analyses based on live-only data: the latter resulting in less spatial resolution but the same general patterns. In a comparison of data sources (original collections, museum records, gray literature and traditional literature), original collections were least effective (51%) in capturing the total species list despite representing approxi- mately half of the total records. Literature was most successful (90%) in capturing the list but only when gray literature was included. Rapid assessment methods contrasted against the long-term results showed effectiveness when based on a range of sample types and habitats. Key words: spatial patterns, community analysis, western Atlantic, Florida Keys National Marine Sanctuary, inventory, rapid assessment. INTRODUCTION Type of Analysis Regional diversity can be assessed in dif- ferent ways. Community ecologists use a “sample and estimate” approach, extrapolat- ing species diversity from standardized sam- pling data. Systematists, on the other hand, often prefer a “find-them-all” approach and use a variety of information sources. In the case of shelled mollusks, the two academic ap- proaches usually also differ in that benthic ecologists restrict their analyses to living or- Department of Zoology, Division of Invertebrates, Field Museum of Natural History, 1400 $. Lake Shore Drive, Chicago, Illinois 60605-2496, U.S.A.; bieler@fieldmuseum.org “Division of Invertebrate Zoology, American Museum of Natural History, Central Park West at 79th Street, New York, New York 10024-5192, U.S.A. 504 BIELER & MIKKELSEN ganisms, whereas systematists integrate data from both live-collected animals and empty “recently dead” shells. The current study, on extant marine bivalves of the Florida Keys, 1$ systematically focused — we have compiled all accessible data to produce a “baseline” inven- tory that can be analyzed for its internal con- tent as well as externally with other faunal results. Any attempt to document and analyze a fauna, regardless of approach, depends on a sound taxonomic foundation. This naturally pertains to identification of field and museum material, in this case particularly for juvenile or small-adult “little white clams”, and also plays an important role in critical interpreta- tion of “legacy” literature data. While mono- graphic and illustrated descriptions are easily verified, unfigured references, even if deemed of trustworthy authority, must be interpreted in taxonomic and historic contexts. This not only involves actual nomenclatural synony- mies, but also “traceable” misidentifications, for example, misapplication of the name of a morphologically similar eastern Pacific or east- ern Atlantic species to specimens from Florida. Deliberation is often also necessary in cases of incomplete or enigmatic geographic infor- mation, especially with names like “Sombrero”, “Sand Key”, or “Long Key”, each of which ap- plies to multiple locales in the subtropical west- ern Atlantic. To make such decisions transparent and to allow subsequent correc- tions where necessary, we have documented our interpretations of literature data used in this study in a separate publication (Mikkelsen 8 Bieler, 2004b). In contrast to most other animal groups, it is useful and often necessary to include skel- etons, (i.e., empty shells) in a molluscan biodiversity study. Many cryptic, burrowing, stenoecious (e.g., parasitic), or seasonal spe- cies are rarely encountered alive, and in fact many species are known exclusively from empty shells (or have never been explicitly indicated, in literature or collections, as hav- ing been collected alive). Shell assemblages, with their usually high preservation potential, provide a time-averaged record that could only be mirrored by extremely intensive and long- term studies of living individuals. A labor-in- tensive molluscan diversity study in New Caledonia (Bouchet et al., 2002), involving 400 person-days of collecting effort of live and dead specimens, showed that about 30% of the to- tal recorded molluscan fauna was never en- countered alive. The inclusion of empty shells raises, however, three key issues. First, re- cent studies have explored the question of whether relative abundances in live-assem- blages are reflected in their respective dead- assemblages. Kidwell (2001) found that dead-collected species occur in statistically similar rank-order abundance to that found in the local living community as measured by a single census. Live-dead agreement (at least for specimens larger than 1.5 mm) was found to remain stable or improve as information on the composition of the living fauna increased through prolonged or otherwise intensified sampling. Bouchet et al. (2002) agreed with the ecological fidelity of dead-assemblages and stressed the improbability of large-scale dead-shell transport into their study area. As in Bouchet's study, we placed considerable effort in collecting living specimens in original collections (because of our research interest in morphology), and, where possible, we have . contrasted live-only data with the findings of the overall analysis including live plus empty shells. Second, geological age also becomes an issue when older strata are deliberately or accidentally sampled (e.g., by corers or bot- tom dredges). Data exist for Florida Bay that largely alleviate this concern: coring data for sites within (Bob Allen Keys) or immediately north of our study region (Whipray Basin) dem- onstrate a rapid sedimentation rate about 0.5 cm per year, so that shells from shallow dig- ging or accidentally exposed deeper layers still date from very recent depositions (Wingard et al., 1995; Trappe 8 Brewster-Wingard, 2001). During this study, we made efforts to collect only uppermost layers and otherwise exclude subfossils from the study. Third, the collection of empty shells has been demonstrated to substantially increase the species-level cap- ture of a biotic inventory. Bouchet et al. (2002) found that 28.5% of the total molluscan fauna in the New Caledonia study was only repre- sented by dead shells. This is remarkably simi- lar (25%) to the results of Kidwell's (2002) meta-analysis of multiple molluscan diversity studies. This percentage is even higher in our study, as will be demonstrated below. From the onset, we were aware of different levels of data reliability in our study: (1) Original Project Collections: Our original collections provide data for individual collect- ing events, clearly delimited in time and space. Specimens are fully vouchered in permanent and publicly accessible museum collections (AMNH, American Museum of Natural History, New York; FMNH, Field Museum of Natural MARINE BIVALVES OF THE FLORIDA KEYS 505 History, Chicago), and precise locations have been recorded by Global Positioning System (GPS) in the field. (2) Previous Collections: Specimens in ex- isting (museum or private) collections have been taxonomically verified as part of this study, but vary in their degree of data reliabil- ity, as well as in their accuracy of distributional and temporal information. Some specimen series have the same data quality as our project collections, while others obviously con- tain pooled specimens collected at different times. Many museum series have incomplete collection data, ranging from simply lacking a collection date (a very common omission in malacological collections that hampers recon- structing past faunas) to solely indicating a broad place name (e.g., “Florida Keys”). (3) Literature: Monographic works, especially when images are provided, often deliver the same data quality as originally collected speci- mens. However, literature data in general show the greatest range in data reliability. Distribu- tional information is often pooled (copied or summarized from earlier authors), and tem- рога! information can sometimes only be in- terpreted as “some time before the publication date”. Faunal checklists (e.g., Lermond, 1936; Lyons 8 Quinn, 1995) have been of critical importance to this project, yet works such as _ these usually provide the poorest supporting data, to the extent that the basis for a species’ inclusion on a given checklist can be enig- matic. Similar to the time-averaging effect of accu- mulated shell material, the pooling of litera- ture and museum-label data that have accumulated over decades might (1) hide changes over time (such as local extinctions or invasions), and (2) make a few chance oc- currences add up to appear as established members of the local fauna. Here we have the opportunity to compare our original fieldwork data with documented prior collecting efforts and to investigate any discrepancies. The pooled data provide the background (i.e., “all Recent bivalve species ever recorded in this region”) against which the original fieldwork results can be discussed. Any discrepancies must be the result of (1) actual changes in fau- nal composition, (2) overlooked species (then or now), (3) rare visitors or human-trans- planted material, or (4) taxonomic misidentifi- cation or other technical problems. Although we are able to make coarse abun- dance inferences and have collected ecologi- cal data during our own field research, we emphasize that ours is necessarily a qualita- tive study, in order to take advantage of the range of data sources. Because of the widely varying habitats (from intertidal mangrove root communities to mud bottoms and coral reefs), great range of individual size and life mode (ranging from a cemented 18-cm gryphaeid oyster to an infaunal nuculid clam measuring 1 mm), and different techniques employed (e.g., collecting by hand or with a bottom grab), the collecting/observation events are not di- rectly comparable. Many of the statistical methods that have been previously used, for instance, for standardized soft-bottom grab samples or for person-hours invested in sam- pling along a transect, cannot be applied to this type of analysis. Project Region — the Florida Keys The Florida Keys (Fig. 1) form a curved chain of Pleistocene limestone islands, mud islands, and reefs at the southernmost tip of the conti- nental U.S.A., extending about 360 km from Key Largo to Key West and westward to the Dry Tortugas Archipelago (about 24°20’- 25°21’М and 80°-83°W). The limestone is- lands that stretch from Key Largo to Key West are characterized by two principal surficial stratigraphic units: Key Largo Limestone in the Upper and Middle Keys, and the oolitic facies of the Miami Limestone in the Lower Keys (Hoffmeister 8 Multer, 1968). The islands in Florida Bay and the islands west of Key West (Marquesas Keys, Dry Tortugas) are not formed of limestone but are mostly accumula- tions of modern sediment, carbonate mud, sand, and mangrove peat (Randazzo & Halley, 1997). The Middle Florida Keys are charac- terized by large tidal channels that allow for significant water exchange between Florida Bay and the Atlantic Ocean. A semi-continu- ous series of offshore bank reefs forms the Florida Reef Tract that demarcates the south- ern edge of the Floridian Plateau, about 5-11 km from shore. Between the reef tract and the island chain is Hawk Channel, a wide V- shaped basin of 5-12 m depth that parallels the island chain and contains various shoals and patch reefs. Ecologically, the “Keys”, situ- ated at the intersection of Florida Bay, the Gulf of Mexico, and the Atlantic Straits of Florida, comprise diverse marine habitats including hypersaline ponds, mangrove thickets, seagrass meadows, mud banks and tidal channels, sandbars, coral reefs, patch reefs, deep sand plains, and hard bottoms. 506 BIELER & MIKKELSEN А unique natural resource that includes North America's only living barrier reef, the third long- est in the world following those of Australia and Belize (albeit discontinuous and perhaps more correctly a “bank reef system”; Jaap, 1984), the Keys are host to about three mil- lion visitors annually. More than a dozen pres- ervation areas have been established for conservation and management of local histori- cal and ecological resources. By far the larg- est of these, the Florida Keys National Marine Sanctuary (FKNMS), was established in 1990 with about 10,000 km? encompassing the en- tire island chain out to the 91-m (300-ft or 50- fm) isobath (NOAA, 1996, also http:// floridakeys.noaa.gov/). This management area surrounds the Dry Tortugas National Park in the west and borders on the Everglades Na- tional Park and the Biscayne National Park in Legend —— Project area FKNMS ~~ Depth, fathoms UMLT Subregi ubregions Gulf of Mexico the east. Our project area encloses nearly the entire FKNMS (Fig. 1). For further description and regional subdivision of the project region, see Materials and Methods, below. Molluscan Diversity Research in the Keys The relative ease of access to tropical spe- cies has made the Florida Keys a popular col- lecting site, and many formal and informal publications, including the many popular shell books by R. Tucker Abbott, have included Keys taxa. But despite its rich history of popular and professional mollusk collecting, and a vast accumulation of Florida Keys material in many museum and private collections, formal sci- entific inventorying in this region began late and with great difficulty. This has resulted in the paradoxical situation of the Florida Keys Atlantic Ocean FIG. 1. The Florida Keys. The project area of approximately 28,000 km? corresponds in size to the combined areas of the U.S. states of Massachusetts, Delaware, and Rhode Island, and is only slightly smaller than the country of Belgium. U, M, L, and T indicate the Upper Keys, Middle Keys, Lower Keys, and Dry Tortugas subregions, respectively, as employed in the study. In an alternative approach, the Lower Keys area west of Key West was united with the Dry Tortugas subregion into a Western Keys unit. MARINE BIVALVES OF THE FLORIDA KEYS 507 as one of the most sampled, but also most poorly inventoried, regions of the North Ameri- can coast. А first comprehensive attempt was made in the late 1860s by William Stimpson of Chicago's Academy of Sciences, who ac- cumulated all available records and loan ma- terial from various collections. This included the extensive holdings of the Smithsonian In- stitution and the original collections from the Straits of Florida and the Pourtales Terrace off the Florida Keys obtained by Count Louis Francois de Pourtales (of the Museum of Com- parative Zoology, Harvard University) during the U.S. Coast Survey expeditions of the 1860s (Dall, 1896; Rehder, 1999). Tragically this material, together with Stimpson's near- completed manuscript, was lost in the Great Fire that devastated Chicago in 1871 (Dall, 1883), and Stimpson never attempted to re- create the research. William Healey Dall (1883) made a second attempt, beginning by discussing the results of collecting efforts of amateur conchologist Henry Hemphill and by analyzing the works of James C. Melvill (1881; who reported on material obtained mainly in Key West during 1871-1872) and of W. W. Calkins (1878; who collected in the Keys dur- ing 1875 and 1877). Charles Torrey Simpson (1887-1889) produced the first effective Florida Keys checklist by including a separate - column in his tabulation of Florida mollusks; this included 98 bivalve species names, now recognized as 86 valid taxa. Concurrently, Dall described the renewed dredging efforts off southern Florida by the U.S. Coast Survey (Dall, 1886, 1889b), culminating in a prelimi- nary species catalog (Dall, 1889a, revised in 1903) that tabulated 225 species from the Florida Keys (plus 15 species now regarded as synonyms and 34 species out of the range of this survey). From 1910 to 1915, amateur collector John B. Henderson, Jr., sampled the molluscan fauna of the Florida Keys with his yacht Eolis, resulting in a massive collection of more than 31,000 specimen series now housed in the National Museum of Natural History, Washington, DC (Bieler & Mikkelsen, 2003). No comprehensive taxonomic treat- ment of the Eolis efforts was ever published, although numerous Eolis specimens have been cited in scattered systematic papers. A privately issued checklist by Lermond (1936) reported 247 nominal bivalve species for the shallow waters of the Keys, 214 of which are here considered as valid. No other compre- hensive attempt at summarizing the Keys fauna was made until the inception of the Florida Keys National Marine Sanctuary in the 1990s, when a taxon list including 163 bivalve species appeared in the FKNMS Draft Man- agement Plan (Lyons & Quinn, 1995). This last total was surprisingly low given the previous checklists (perhaps confounded by the plethora of archaic and synonymous names used by Simpson, Dall, and Lermond) and was, like them, undocumented as to source and specific Keys location. In 1999, we completed a preliminary assess- ment of marine bivalve diversity in the Florida Keys based on a dataset of 6,000 records criti- cally researched from literature, taxonomically updated museum holdings and five years of Original collections from 1994 to 1999 (Mikkelsen & Bieler, 2000). That paper empha- sized the importance of this fauna to studies of zoogeography, malacology, and conserva- tion efforts of the Florida Keys National Ma- rine Sanctuary. 325 species were documented, representing a 100% increase over the previ- ous tally (Lyons & Quinn, 1995), largely attrib- utable to a critical review of museum-held specimens. Literature sources (especially when non-traditional newsletters and agency reports were excluded) or original collections alone were found to be less effective in cap- turing the total fauna; these results questioned the effectiveness for inventory work of eco- logical and other studies that are so restricted in sampling effort. Since 1999, our effort to document species occurrences has continued, adding 186 origi- nal stations over another five years of collect- ing throughout the Florida Keys (including the Dry Tortugas, the region least represented in the previous analysis) and, importantly, cap- turing data from 280 additional literature sources that mention bivalves in the Keys. This has culminated, in our tenth year of study, ina database more than twice as large as that used in our prior analysis. The newly surveyed lit- erature includes important early checklists (Calkins, 1878; Simpson, 1887-1889; Lermond, 1936), results of substantial ecologi- cal surveys (e.g., Turney & Perkins, 1972; Brewster-Wingard et al., 2001), and many detailed Florida Keys collecting reports by amateurs in publications such as Frank Lyman’s Shell Notes and the Palm Beach, Florida, Shell Club’s newsletter Seafari. The full annotated bibliography appears elsewhere in this proceedings issue (Mikkelsen & Bieler, 2004b). 508 BIELER & MIKKELSEN Goals This paper addresses four major issues: (1) Characterization of the bivalve fauna of the Florida Keys, for which a complete (“ever recorded”) annotated checklist is provided. (2) Analysis and discussion of species-level similarity patterns within the Florida Keys, both in terms of geographic subregions (Upper, Middle, Lower Keys, and Dry Tortugas) and in relative position to the island chain (bayside or oceanside, the latter further divided into shallow and deep records). (3) Comparison of the total Keys bivalve fauna to other regional faunas in the western Atlantic. (4) Discussion of the respective utility of the different kinds of major data sources (original collections, museum holdings, and literature sources), and of the results obtained by “а! shells” versus live-collected records alone. MATERIALS AND METHODS Study Area The “Florida Keys” project area is here de- fined as the waters surrounding the entire is- land chain from Broad Creek (about 25°21’N, 80°15’W) at the northern end of Key Largo (in- cluding Card and Barnes Sounds but not Biscayne Bay, southwest of but not including Old Rhodes Key) to west of the Dry Tortugas (at 83°30’W). The southern half of Florida Bay is included (with a northern border at the lev- els of, from east to west, the northern end of the Nest Keys, Russell Key, and the northern limit of Rabbit Key Basin), eliminating that part of the bay that is more properly considered the southern extent of the Florida Everglades (Fig. 1). To facilitate analysis of the Keys fauna by a priori selected subregions, traditional bound- aries were employed but explicitly defined as Upper Keys [Key Largo to Craig Key], Middle Keys [Fiesta Key to the western end of Seven- Mile Bridge], and Lower Keys [Little Duck Key to Rebecca Shoal (west of the Marquesas)]; the Dry Tortugas Archipelago was treated sepa- rately. These subregions are roughly equiva- lent to those recently used in other diversity/ monitoring studies, such as that by Wheaton et al. (2003) on stony corals. An alternative scheme, that draws the western border of the Lower Keys at Key West and combines all those remaining westward with the Dry Tortugas into a Western Keys subregion, has been used in water circulation/exchange studies (e.g., Lee et al., 2003) and in several of our analyses we recoded our data to include this grouping. Depth categories used were based upon sampling techniques: shallow = wading to diving depth (0-35 m), and deep = beyond normal scuba depth (1.е., > 35 т or 100 ft); these are as pre- viously employed (Mikkelsen 8 Bieler, 2000), but an oceanward depth limit has now been set at the 300 m (= 164 fms or 984 ft) isobath (a restriction which eliminated seven previously included species). Nearby records were com- bined into 59 land-based locations (usually a representing single island or small cluster of neighboring keys) so that each location was represented by at least 25 records (fewer for the live-only analysis, see below). Each loca- tion was then coded for its general exposure (bayside of the island chain, 1.е. in Florida Bay/ Gulf of Mexico; oceanside, shallow or deep), and its position in the Upper, Middle, Lower Keys, or the Dry Tortugas. For analyses of the (considerably fewer) live-collected records, all stations represented by single records were combined with neighboring stations where fea- sible, otherwise they were eliminated (result- ing in a total of 63 stations). Data Sources and Quality Literature sources are as described in the catalog and annotated bibliography elsewhere in this volume (Mikkelsen & Bieler, 2004b). In addition to the nine museum collections pre- viously listed (Mikkelsen & Bieler, 2000), the full bivalve holdings of the Houston Museum of Natural Sciences (HMNS) and the Rosenstiel School of Marine and Atmospheric Sciences (UMML; University of Miami, Florida) were surveyed and identifications confirmed for this analysis. Original collecting methods are as described earlier (Mikkelsen 8 Bieler, 2000). All collection lots were interpreted and recorded as dead-collected unless soft tissue was still attached to the specimen, or unless the labeling explicitly indicated live-collection. Nomenclature, initially based on Turgeon et al. (1998), has been substantially modified according to subsequent systematic research, the latter partly a result of the 2002 Interna- tional Marine Bivalve Workshop (this issue; Mikkelsen & Bieler, 2004a). Synonyms and misidentifications used in the literature in the past 160 years have been carefully researched and documented (Mikkelsen & Bieler, 20045). MARINE BIVALVES OF THE FLORIDA KEYS 509 Analyses The new Florida Keys bivalve dataset is more than twice that previously analyzed (12,382 versus 6,145 records), and provides a better- balanced analysis of all three data sources, comprising 3,385 records from the literature, 3,231 records from museum lots, and 5,768 records from original collections (Fig. 2). The dataset was compiled as a Microsoft Excel* spreadsheet in the form of a square matrix (species by location). Coding within the data- base was as previously described (Mikkelsen & Bieler, 2000), including data partitioning into the aforementioned geographically defined subregions. While Dry Tortugas remained less sampled, as in the previous analysis, the Up- per, Middle, and Lower Keys subregions pro- vided relatively equal proportions of the database records (Fig. 3). It is important to note that efforts were made not to duplicate records in the database. For example, literature records describing lots that were subsequently verified in a museum col- lection were scored as museum records, that is, as the “original” (and more reliable) source. Literature records that were judged to be based on the same material were entered into the database only once, as the earliest publi- cation. As has become customary in commu- - nity analyses, we are employing the terms “singleton” and “doubleton” for species that were encountered only once or twice (here referring to species represented only once or twice in the database). Literature and museum records that provided only “Florida Keys” as Literature 26% e 27% Original 2 47% locality data were not entered into the data- base (giving rise, e.g., to “0” record entries and singletons from multiple locations in Table 1). Of the 15,296 records generated during this study from all sources, 2,914 (19%) were not entered into the database because of incom- plete identification, species of uncertain sta- tus, poor locality data, or duplication. Although this was a qualitative study, the number of collecting events or the frequency of encounters (= records-per-species in the database) is a fair approximation of species commonness or rarity. In this context, species were scored as abundant (> 50 records), com- mon (10-49 records), and rare (< 10 records), regardless of source. These data are also re- flected by a histogram of the frequency distri- bution of occurrences, plotting number of taxa versus number of occurrences (log,) binned to full integers. Within the database, records were entered as “live” if at least one specimen in the lot was live-collected (regardless of additional dead- collected specimens). The heterogeneous nature of these records is also evident in this regard. While conservation-minded collectors might preferentially collect empty shells even if live specimens were present, morphologists, anatomists, and systematists would preferen- tially collect live specimens, perhaps also leav- ing behind empty shells at the same site. Literature records and museum labels often do not indicate whether specimens were col- lected living or dead, and museum specimens are often so efficiently cleaned as to remove all traces of tissue. In all uncertain and un- Upper 30% Middle 27% | f / Tortugas / 10% Lower 3 33% FIGS. 2, 3. Percent of records in the database. FIG. 2: From each ofthe three data sources (literature, museum holdings, original collections); FIG. 3: From the Upper, Middle, Lower and Dry Tortugas regions of the Florida Keys. 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JIWN WO (2281 ‘UOSUIEMS) е/492 BQ x x x x x lp 90 as et МП WO 68/1 'esambnig веомдии Boy x = x x x $ go See INN NO (zz81 ‘Aes) езлэлзиед влереиу x x x x x се gO as 08 LINN МО (8621 'Buipoy) sijiqejou esepeuy x = x x 5 y go as SL 11-Nn INO (6981 'релиоЭ) емериоу esepeuy x € SO ICONS re: 1961 ‘маман /иешубпед esepeuy x x x x x ce gO as 06 LINN мо (6181 ‘youewe) 5/5иебишор лезу эерюл\у en x x = | =O “OST EL 11-N NO (рег ‘duepoig) sipn snusapopog x Е X x = ms qo as ir INN NO zr8L “AUBIQIO Pp xa/dwis eıwouw эеришои\у ее EEE EA E er = Pb § с 2 я 2 = 5 seloods Awe 4 ш D 3 = D Las O pel ne 8 Е a о = D > i >, : A on > oO 2 3 S 2 © © co - © 5 oO = о с f = = 8 un + sy a со o D O © m я * 8 8 7 es) “SISIMDSYO EPUO|4 лэшеэ BAY Ul UOISNJOUI pue ‘ApNjs $1} и! asuaJmooo pue uonnqusip бицоецел sauoBajes yym ‘зэмема say ePHO|4 | 3718WL 511 MARINE BIVALVES OF THE FLORIDA KEYS (зэпициоЭ) x x = = = L X С X 9 x x x = = ez x x x x = 9 x x = = = 0 SE E E x x = x = OL x x x x x 19 = = x x = 0 x x x x = vl x x x x = 9 x x x x = L x x x x x L 0 X = = x = 0 x = x x = 9 x x = = = 2 x | X X = 5 = ¿9 x x x x x re 0 x X х x x OL x x = = = с x x Xx х = 0 550005 “ERIN 49661 O81 9561 ‘руоциел $061 '26881 “eg 6881-1881 'uosdus A +udeq ип NO --WN WO wna NO wna NO 11-N NO LIN- MO лип NO wna NO wna NO 7 O wna NO AO O LIN- NO LIN 1M- wn = Wo own Ww 111 MO wna TWO LINN NO === IN- LINN TWO wn TWO LINN NO y quisIg sAay “ely ,S891n0S веа 9881 ‘leq e9nj0e, ешецэ (СУб| ‘Jekeg IN 4) эехеш ешецо 6181 H91EWE7 ериоу ешецэ $581 ‘peiuo) ejebarbuos eueyo 9981 ‘релиоЭ enulos eyjauruy (Ly ‘релио9Э) euejdiad зиэшолэ (9581 ‘AeS) eJejuapın suewoinel4 (zv81 'Аибюло.р) sisuanbulwop зие|=) 8581 ‘релиоЭ гиериоу елэшеуирле (zv81 'AÁUBIquO Pp) шпиещие еирлезоиоби | (86/1 'snaeuur7) wnjeounw штрлеэ/цоел | (9581 ‘Чыомэщиу$) шпиециошба штириеэ/цэел | (6841 ‘элэтбп.а) suuoyuejos eepuAÂdez (Gz81 ‘Aeid) ejeonsiwes espuÁdedy (8411 'usog) eje, eapuAded (1881 EA) wnjouy шпириезошем (1881 “IeQ) э/дешелаа winipseoowayy (9881 “1eG) wnoquegAs unipleoin9e7 (1981 1эцэлеч) шиза штирлеэмеэет (0581 ‘релиоЭ)ииомош штриеэмэет (96/1 'snaeuur7) шплеблее| штирлеэлэе7 (98/1 ‘JoojuBr) wnsngoi wnipiesouiq (89/1 'snaeuul7) вирэш врлериэши (0161 ‘eleiy1) Addn6 eıpiesuswuy (8611 ‘sneeuulq) шпибеш ешбиз]$0/19у saldads эершецо эерирлеэ 9EPIIPIEO Aıwe (panuyuoo) BIELER & MIKKELSEN 512 (зэпициоЭ) > = = X = | =0 а- 8 SS IN = (9/81 'зАэщег) es е^шо!рлеЭ X = = x 2 € =0 а- 9 IT 1-0 (1881 lea) eJensouad е^шо!раеЭ x 0 а as с = © (zr8L ‘AuBIqO.p) виизздешо eAWOIPseD x 0 7 ра р iss NE (9881 ‘ysng) 219/16 елшо!риеЭ X = X X = | ао =S S -IN- 1-0 (0€£81 ‘зэлецзэа) е]е//э]50> e4uwolpieo 0 =0 а- | 1 1 (zv81 ‘AuBbiquo,p) еешеуе е^шо!рлеЭ sepiuepidsno x = x x = | -O as $ 11-N мо (2981 'swepy y) eso/osds eyjajessesang xx = = = о 90 а tt dj N= (zrgL ‘AuBIqiC,P) з/5иеоииеш eyjsulsselg xX xX x x = а, do Mas. ce nn WO (pee! ‘резлиоЭ) EJE/NUN] ejjeulsseug x 0 =O «ds $ LIN WO (5061 lea) euerudnp eJjeuisselo эер!э}е$$е19 x € -© ds 7 De “NO (9881 ‘US “y 3) иаача етоиоэнел = = = x = 0 = Saas 20 1 (2981 'swepy ‘4 '9) eueisgamy епдиозиел x 7 X = = 6 go cas #e 11-Nn INO (zv8! ‘AUBIqO,p) sijuedsip епдиоэиел = = = х = 0 7 ==), 0 7 9€81 ddılıyd зи/елтЬэе ejnquose!np x = x x = 0 go cas 0 11-N TINO (2981 ‘swepy ‘4 9) euelzjaip enquosoÁ1eg = = x x z 0 = ти (10 7 (Lee! lea) е/эш/э ejnquoso/ueD X = x = z e =O) Gis G Le PQ (zzg\ ‘Aes) ejoenuos епдиоэоеЭ AS x x в | =. == | = AMES (2681 ‘swepy ‘я 9) eue4puyo einquoaoAled х = x x = G go as SL LINN NO (zpg1 'AuBIqIO p) eaeques enquosoieo aep!nquo9 xx x x x с: 90 “S$ 85 INN TINO (zreı 'AÑuBIqIO Pp) ешциеш eposawAJOy SepI|noquog xx = = = С go as zz INN NO (9681 lea) ичишз sisdopieg sepupreoo¡Ápuoo xx = = - 9 go AS E INN NO GESl ‘duepoig ESONUIS ешецо xX xX x x = @ go as SL nn INO [781 ‘элээч epses eueyo хх = =. = ee 90 as Sl Lawn NO 6181 ‘\олешел SUEIPEI eweyo хх x x x Ov gO as 19 LINN WO 1621 ‘uewo еИцаолееш eueyo о 2 = a > 5 saleds Aluue 4 U O 3 — D RS D, = Я 0 D = 2 œ $ =: + : a n S 6 2 @ 2 $ e 2 a 3 ° © я 9 cai es NN NC © D co 9 (panunuoo) 513 MARINE BIVALVES OF THE FLORIDA KEYS (sanuyuoo) € =O dS Ol IIN= TINO (9861 ‘Амен) ¡quibow essyoAy sepraeydAso x x x x = 49 90 as Ус ПМП NO (1621 ‘Uljewg) ejeunosd euoyeon | X X X X = 0 —O == Z === IN - (94/1 'snaeuu!7) ejepun sugwÄaA]g С о 95 7 п О (2961 ‘IOOIN) syeyoeds зиэш/э/|=) x x x = = G =O) 9 eg LINN “NO (86/1 ‘sneeuuly) ejessnoap suawAo//5) X = X = = 0 == dz | 15 IN — (6281 's9ueJJaq) eueouawe SUaWADA/N SepipuawioAo x x x x x y qo cas pl LINN NO ($841 ‘Ja|bueds) eyeysos гиэ/биэа$ x x x x x р ao as el LIWN NO vest ‘| Áquamos g ‘9 ejeno виэеуэодзе=) x x x x = el 90 as LE LIWN INO (L6ZL ‘UISWO) suey eueeyoonses эершэецоодзео x y go SS LIWN WO (6681 ‘!еа) euepuoy eJaygolo x | da as al EAN 10 (9881 “esneny) ejenuejd eyashy x = = x x y go SN IIN= NO (162) ‘uewO) Iuosuepe eaese7 x | zo 20 | 5550. JUS (£08L ‘nBeJUON) suejnaiquogns ejay eeployewwosjes тя х x E 0 Е ne “0 7 (9881 Ра) /99qs6rs ejjounoay Sep11]ojuy X ad X = 7 0 g- =S | FM. 150 (6781 'Znj99y) /ayes sısdoyiAyy X | а- ES с === 50 (Leg1 ‘peiuo9) eyeaeydoong sisdo/nAn aepiuassialg x 7 x x я 0 se ae. CC seal Jc (8181 ‘\олешел) eueljisesg eluebiyd] x = x x x 0 == ne AS м IW 2281 ‘Aes $//демел xeuog эерюечоа x = = x x y а- ESTO WA IW (9681 “11eG) euepuoy epiouaAD эер!рючцэ/^Э X E x X a € =0 а- С In IN = (1881 ‘иеа) зте/пиелб иоро]э/“ 0 = =) | ===> М 9981 lea вещзютеа елэиоли| = = = x = 0 se as 0 ==. - 155 (1881 eq) елэдиэше! eısuoAyy x 0 == => | 1 IN (reg ‘шел ‘3 Vv) eajuebib erauoAyy x = = x = С OMAN Mee IS (56/1 “a¡Buads) eyeysou виер!а$пЭ re = я X = 0 == а- L les NS (9781 ‘ugA07) езэдо euepidsno = go ® 5 = = 2 =) a x O sendads Анме- u 2.23 = 3 o, ® о O af | > © = g в = 3, : a ” N 5 со о us ® © es = со 5 = 2 о de) = ep A о о oO) co AR = E 8 à = co (репицио9) BIELER & MIKKELSEN 514 (senunuo9) x = x = A 0 8 959 ZN ЛИ (9/81 'э$$019) IWWBIY9S ециороиу х = x x = 2 qo cas OZ INN NO 1081 un е4/е едиороиу эериюп7 x E =O, “US 19 LEN NO 8681 'ysng Y IIHISA “3 ‘Y eyegjns 515407 X г > X = 0 =0 Ge £ O) IN = 958, ıddılıyd eynuiw sısdowiT x с = x = 0 а № мп Wo 9/81 'sAayyar erejsus sisdow!7 x = 3 x = 0 > sal | Ne (ÿL8L 1429048) вшпе sısdowiT eepisdou 0 == а- | уе IN = 9881 ‘eq геиешио/д eawı7 = = = x = 0 == а- | === IW- (208; 'nBejuoyy) eJeynaunegns ejnjeu7 Den = x = 0 == = = 10 1 9881 lea eIsyNas е/тзешит TR х = u 0 = 0 1 (S88L ‘UWS y '3) esnjuos enjeu 7 x x x x = ss 90 as est мп мо (9781 ‘swepy ‘4 ‘9) eplonjjad euewiT ) ON 1 7 (5961 ‘AUIOON) JUNO) емеиит х x x X X Le go as 85 LINN TINO Zrgl ‘Aubiqio,p esequeo eur x = = x = 0 == al | 7 (9881 lea) ешоэ!4/е ешииел!а x x x x x ez dO qs 96 wn WO (8221 ‘Uog) 9925$ sapiouajD X | a- as G LIN IN = 2861 ‘орлепз$ ¡¡nedyoues зэрюиз]9 x 0 re а- | 1 IN (9881 lea) snjejnuejd $эрюиз} x x x x x 12 gO as zı LINN NO (2081 ‘Уэлешел) sw зэрюиз} С g- Ge se ies: < $005 ‘181819 3 USSISHAIIN SISUSIWEIW SOPIOUSJI sepiur] X x X = > 9c go as 29 LINN WO (6581 'uojuy) Snjerpei иошоибоз| x x X = = LS go as 50 LINN ‘WO (Gpg| ‘swepy ‘4 9)/0/091q иошоибоз|] х X x x x Gt 80 =S pol LIWN WO (1621 “uneuwo) snjeje иошоибо$| eepluowoubos] т 5 = X = 0 = si 0 7 (1881 lea) eueze eyajelyH x = = x = L =0 as or wn WO (19/1 ‘Sheeuul]) 89/7918 в/эзен эерша}е!н x р =0 as v п МО (56.1 ‘104) /еэ/уроэ эшороиолаовм = & E Y D za 2 я a т O зэюэас Анше- ш 8 3 3 e o, pe 8 y D > 9 = о ca # : © n O [Ce] а. = = < O O [de] ES © (47) = oS m 2 © © = 8 un + eS se © a 2 O oO A =; > в 00 ch 00 oO (penunuos) 515 MARINE BIVALVES OF THE FLORIDA KEYS (sanuyuo)) 2 = x x = р a == | ste Sa (zz81 ‘Aes) зиелазе/ erunnyy x x X x = € go =S Lg INN TINO (1621 ‘чнэщ9) з/ибел ешоодоеи| x = X = = 0 == == | NT IN - (2081 'Ja¡buadg) вицеие euneuy SepiSelAl x L go 8.78 LINN = WO 6781 ‘релиоЭ виериоц eISUOAT x x x x = G go as Hr wn TWO (zv81 ‘AUBIqiO,p) eueag ewusspouz SepiisuoÁ7] X = = x = 0 =O Gas INN “NO (££8L 'релиоЭ) виериоу EIUEME]S x X X X = 9 so as zp LINN TINO (1061 'IIeQ) ejuenue eulonjoipey х = = x = 0 go as Le LINN WO (9881 'Иеа) sisuaralquios eulonjoinald x x = x = 0 а в п мо (9881 Ile) ешИэоэпа|/ ешополта| x x x = = С go as oO Wn IN- (1621 ‘Ullawg) ejeuyosd зарюэеца x x x x =; LL go as gg LINN TINO (1061 'еа) е/эиало euronjnleg x x x = x G OMS eo! LINN WW (zv81 'Аибюро.р) 8721509 eulonyinied = = = x = 0 = as р => Ne (0881 ‘UWS 3 эл ‘1 Y) Sug sısdosuAyy X = = x = 0 sae = 0 1 (9881 ‘I1eq) ereuubes eauAyy x = = x = 0 ON Gig Al LIN (1581 'uosduns) esoy ешошопт x x = x x IZ 90 as 9 LINN мо (9ÿ8L ‘релиоЭ) eınsseu eosiuon7 x x x x x y =0 =8 1 11-Й IN - (96/1 '4a¡Buadg) eJesunw еэзшиэп7 х x x x x те 90 as vz LINN TINO (86/1 ‘sneeuuly) вошелИзиаа eulon7 x x x = Е. 0 go Oy In WO (5181 ‘POOM) EJejuap eJjeoueng х X à x x | go as 16 INN NO (zve1 ‘AuBIqO,p) eyeajnsupenb eßunenig x x X x 7 0 == as LL ==" 2220 (2981 ‘swepy 'g 9) eeunoed eue) X x x x x bp 90 as 282 LINN мо (8081 ‘nBejuon) ejenorquo eue]; X x > x x 06 90 as 25 LINN мо (86/1 ‘sneeuuly) suejnaiquo амероЭ x X x x E G go as X LINN TINO (1061 ‘чозаци$ $ пеа) ерие/9 ебилеЭ x = = = = LL go as 8 wn Wo (1461 ‘Je 39 xo9 ul 'ueaeyo) эвиээх виопуеЭ SS ов = = 2 я À т S saloods Alle + w D 3 3 TD o D, TD. à. A o ао ti + : o En ее = E E % # © à @& e un © ER, ea CA Oo. Fe 5: © © Ber co (panuyuoo) BIELER & MIKKELSEN 516 (senunuo) ххх KK KK KK KK KK KK KK KKK KK KK XK 650002 “ERIN X X X = X X X = ет. « X X = 5 X X X X X X X X X X X X X X X = x x X X A X X X x Dn FR FEN Lo Nes Le X X X = X X X = X X X X X X X X D Len A Lu a A A ie, = ae a ee D oo gg + © 3 = 3 s 2 @ 9 oo see ek te & O © IR 8 6 INN IN = OC ЛП TINO er LINN TWO GG LINN TINO evt. ПАП TWO Se LWA TWO Le LINN TWO Ср LINN TWO eg LINN TINO vl LINN TWO e an W У LINN то € мо W- 8 Ale: TWO 9€ ZINN TWO CL INN TWO pee ШП МО Lv LWA TWO G San IN = С Tes 120) ÿ NN IN = | N N 81 LINN TINO | ый TA с IAS а > y e 5 x LA an < O 5 E > * (zz81 ‘AeS) esalapuod ejaoyy (9881 Ile) swepe sisdooy (zz81 ‘Aes) /елазе| snjnosnyy 296| 'Ányuadnesg snsowenbs snjoipoyy (GL&L чэрром Y YORS7 и! ‘цоеэл) зпиеэиаше SNJOIDO/A (zreı ‘AuBiqO,p) Bu ебецаоцут (zreı ‘AUBIqO,p) взеэп$!9 ебецаоцут (2181 ‘иАмиа) ejeysue ебецаоцуит (zve1 ‘AuBbiquo,p) шпиецие ебецаоцут (zz81 ‘ÂeS) snaue]seo Sn19g017 (ОСЗ ‘enbseuyey) uwnrngal wnıpeyas] (1641 ‘uyewo) ебецаоело) еэиебал=) (yL61 ‘Ш Áqiamos “g 9) ewissisoue/6 eisuaynay 1661 'Sejoo $ sejes иозлэриац LuNjnjuebaja штриюеа (3081 'nBejuoyy) eyessnoap ejjoualo (/9/| “snaeuur7) snjorpow sajuopiyoelg (8G/L 'snaeuur7) snisnxe sajuopiyoelg (1621 'uauwo) eosny ejnjog peBlL Japyay wnjeyibes штербАшу (0881 “UPLUS 9 шел “3 ‘Y) шпуюа WinjepbAwy (9yg| ‘релиоЭ) wnuAded wnjepbAwy (pGgl ‘SWepy y $ swepy 'H) sıllbeuy ешецас (2481 “AÑUBIQIO P) зпиеэариеэ SnaJjEn (1281 ‘релиоЭ) /эцалел eınsıds (8181 o9Jewe7) г//азе/а ejaey saloads эерэом эернзли! 2epAn эер!эеи\ Alle + (репицио9) 517 MARINE BIVALVES OF THE FLORIDA KEYS (зэпициоЭ) x L 0) as Др ип “IN- (6181 'Уолешел) suejoauı изоэатЬэу = = X > = 0 == >= 0 1 (себ! lea) зпоеуэц usjoadmbay x 0 50 @= el és We (2881 “MUSA “3 VW) $719//б uaysodinbet эершцоэа x € go as pl nn “NO 6961 ‘LAW 9 SSOY веуи! eIopueg = = = X Eu 0 = == 0 7 6181 ‘чэеэл sıjewejb e1opued X E —O as € AN IN = 9881 “eq eueiysng elopued С =0 == С = == VESL PeIUO9 esouale влорие эериоричеа x й go as Уд ИП = IWO (LG6L ‘uoss|CQ) Vegem eaysofeyse | x x x x = 81 go ES Le INN INO (pee! (es) sıysanba ejoaso x x x x = Ze go as 6LL LIWN WO (8911 ‘sneeuuly) suoy еэдзориэа X = X x = € go as OL wn WO (16/1 ‘uljews) еэшиблл еэд$0$$е19 x = = = x 0 == =S с = == (вез! ‘Buipjing) aesoydoziys еэд4$0$$е/9 эер!э4$0 x 0 go as GG лип TINO 2281 ‘Aes eunxold ejnonyy X 0 =© as € п ANS 998| ‘зшеру y ejejnuaso ejnony X 8 go ES L INN 1-0 1161 ‘2100ON 2/09/9/29 ejnonyy X 0 go во G TWA =40 (8081 'n6ejuoyy) sinua] ejnonuuz x 3 x x = OL dO == oh IAN = (6.81 'sAauyar) sisussbae ejnonuug эерипопм = = x x =: р — ©) as 8 =1N WO (1881 ‘Wed) vajuadieo epajadolg Zu 05 > x = 0 Fe == 40 = 7 (zr8L “AuBIqIO p) еали euepnony x = x = = 0 oo = 0 1 (9881 lea) euenuen euejnony г. 3 = = = 0 =0 (al С == _ МО (9881 ‘1I2Q) ejoeyipyos euenany E = = L = 0 =0 as =1-- 1-- (zv81 'Либало.р) sısugsıewel euejnony X € g- as OL пи 1-0 (yzg1 ‘Aes) еэщиаэио) eue/nonN x = = x 6 go as ez wn WO (ZEB ‘pesuod) ense euejnony = = = x = 0 == == 0 Sn = 8681 'ysng 3 IIHISA ‘1 VW Sinajgns ерэрэт эершетпопм = E & Y D > = = a am я saioeds Ajiwie 4 we) Se o, O D о of wm = 2 = $ °F 1 = : & n os aa 3 © - FE US |e 5 3 un Ba SS Bn 00 CA a © © in = * E A co (panuyuoo) BIELER & MIKKELSEN 518 (senunuo9) x x x x x ji go =S Or LINN WO (L6ZL 'UauWo) eprorde, eJ09194 0 => == e и IN- Er6l ‘1эрчэч eoyueye е/элэ4ооЭ x x = x x ih go -S Ly wn “NO (pee! ‘| Áqiamos ‘g 9) winjsngos иород5иоц эерноэ3э4 x = x x = 0 als =S | —-N- 10 С881 ‘1OU9SI4 шпиэиа] eworduad x — x X X 0 == == с nl IN - (LOS) A9Jewe7) шпеэешеблеш ewordusg 0 in =a 0 Л 0981 ‘uosduu}S unmepiuelAd EUS9PO/Y907) эерцешоацэа x x = = = С go as SL ми WO (2681 'ysng ишед ‘2 YW) Mpauag sAuejyooyjeds = = x x = 0 go as Ov лип NO (6781 ‘peiuod) snsobey uajoedipoyy x 0 == =S | 7 IN (£t6L чэлея) зпзо/дпишех usjoadoJA7 x x x X = G go as 69 wn “NO (8281 'poom) SNSOISNU иаэдершт X x = X = 0 == —— 0 7 (ÿ98L “Jaxung) ejewenbs nu sÁweJyomae7 x x x x я G go as СУ лип TINO (8SZL 'snaeuulT) 9ez91z eJonn3 x = x X x | go as 85 Hiwn “TINO (8681 '11eQ) //эиэле/ е/олп5 x 0 -O as С IIN= IN- (5/81 'аае9) esoelÁded yo ejonn3 x 0 ae d= v ¡AS Wi (1641 ‘uljews) диалпе! ejonn3 x x x x = | -O as bz INN W- (0061 'Biaquazineg) ayezeyo ejoang x > = x = 0 OV US" 9 LIWN IN (9881 lea) wn/6yd ueoedojdA) x X x = = ez go as р LIWN TINO (£981 '3A99Y) SNUOS зАшецэедие x x x X x 0 go =S ez INN =AWO (6181 YoJewe7) вело SAÁWeJyoequeo x x = = $ 0 =0 =S 6 INN IN- (рубр Jaeg) эералриш sÁWeJyoequeo x x x x x 0 10) as LS INN WO (16/1 ‘uljawis) везидии s{wejyoequed x x x x x 8 go as vl ли WO (5981 ‘Zn99y) Wneynue sÄwejyssjyseig X x x x = G go =S ZS лип WO (81/1 ‘Uog) snajonu uajoadobIy x x = = = vg go as с, INN ‘WO (6181 Ao9Jewe7) зиврел uajoadobIy XxX x x x y go as u LINN NO (86/1 'snaeuul7) snqqub изрэаоблу = $ © E =. = a я a a O saloods Ale + с Le) 3 — 5 DA o a 2 ‘ y = aks O N a =} 2 = t A on N = 00 O + (Y) © co а со 5 < (e) O oO Ra © [07] = 2 À = © o о o an + = EN œ CA a oO oO 1 =; A о — oO © © Ten, 00 co (panuyuoo) 519 MARINE BIVALVES OF THE FLORIDA KEYS (senunuo9) x = x x x 0 == == | = IN - (857, ‘зпэвишл) snjejnoewig хеиоролэ]эН x 0 = ©) OS ig, 6) (9/81 ‘UOION) 82410 We x x x x x v go SO INN NO (86/1 'snaeuur7) взелоуер siydesy — эериаошшеза x = — x = 0 == а- | === IN - (9981 ‘12a) wnuefes uinissnwesdold x - - x = | =0 а- G 11-N IN - (998, 'IIea) wnueisajeunod winissnwesdold = = = x 3 0 Fa > 0 1 (9881 ‘118Q) wnurssejeyg шпиззпшелиес = = = x = 0 == 2-20 ar == (9881 ‘Чиш$ “y '3) шзеезие) wnıssnweneg = = = x = 0 = ==” 0 1 (6881 lea) snjeJ6u]s usjoadoj9ho 0 go as 0 INN NO ¿681 'ysng 3 IIHISA “3 “y $пиви uasjoadoj9Ág эериззпшеэдола x 0 =O. de р IN IN Er6l ‘Japyay EJensoı EAWOIOG x = x x = | OA SA MINO (6581 'diopuajsam 3 SAN) вде/пиелб eAWOs0g = = > X = 0 La ee 1 9881 “ea ер!а/е eAwolog эер!Ашолос х х х x = y go as #8 INN NO 1081 ‘H21EWET esoqq/6 етеэ!/а эерипуеэна x x x x x 81 go as $56 INN WO 6/1 ‘uljewig eaues виша x = x x x | so as 0 INN NO (9281 ‘| Agemos ‘а '9) ejeuas Buy x X 3 x = v go A INN ГА (6181 A9Jewe7) ерпише$ eun] y x x x = = 9 go sn ce LIWN WO (9821 0044617) ериби вищу SEPIUUIA = = X X = 0 ЕН Е: 0 Л 1611 "USW sısugıyaoadwesa SeJ0Ud x = x x = 0 g- zoe MEL LINN = WO (86/1 'snaeuu!7) EJEUS EISOUEN x = x x x 0 55 + > al = (zz81 ‘AeS) зишодеипо eISSUEN Xx = X X = | ао ES 9 AN IN - (85/| '“snaeuul7) е]е]50) einajdopAg x = x x = | g- = | Ne INS (zz81 ‘Aes) eyeounsy esuleg эерреоча а —- = X z L sa Я = Poa SNS (1881 lea) sisuayyue spero seplAiqoliud x Rg = x = 0 =0 =. E “lent ae 0061 “eq -/OUIW SISUZ эермеча x =; x x я 0 3 =a | leo Мы (8181 ‘олешел) зишиодреюца еивоэщеа = D 5 в = = 2 O À a 5 salads Awe © LS) 3 T D = o, Е 2 == а WN > 2 с 6 ++ 5 0 n о oO а со 5 < O © oO 5 © [4] = Sn 5 2 © S © O co 1 = E о = oO (95) © DEA 00 co (репицио9) BIELER & MIKKELSEN 520 (senunuo9) = = = — оюоюфооокюокг чот ою т оютюям о 9681 ‘21994 Sn919]0I SnıApuods гель ‘ччециэн snueouawe зпИриоа$ zz81 ‘Aes шп/эл eAwajos 1681 'saÁeysag э/елиэрюо EAW8/OS (98/1 ‘oojur) sniagajd snjabe 1 (p6Z| ‘1эбиуэа$) snsmip snjabe 1 (1981 чэхупа) snueıßununga $пипээ/о$ (1y81 ‘эброн ul ‘релиоЭ) зарю/пэпи euljawas (L6ZL ‘чНэш9) susaseindind ajawes (66/1 ‘Aeuaynd) епэуола ajawiasg (1581 'релиоЭ) ejen]sejjeq ajawas 9881 UIUS “y 3 ejeyjeoueogns ели (9081 'nbejuo¡y) зиэуи 813 (0981 'sauoH) еэщиаэзиоэ el) 6561 ‘лэрчем ибиш/циел eıbumundg Seg, ‘| Адлэмос ‘9 ‘9 2J2J9/209 eıßunundg 868L ‘USNg 3 ишед 3 у eueouawe SIJE9IBUO] e1qy (1881 ‘112q) elo! e1qy (zz81 ‘Aes) syenbee e1qy (1981 ‘eAS94) вади виз“ (8611 'Buipoy) 19 ш//о euej4 (857 | ‘зпэвишл) еладдиебБлеш epejouig (2581 ‘чэхипа) esowenbsibug] epejould 86/1 'Buipoy eJE01qUI epejould (16/1 ‘uewo) euejounbues еивоитбие$ aepiipuods aepiuwajos Sepiungajos sepijawas aepiliald т x x = x x x x X = = X x x X = x X = X x = x X X x ri = x X x X X X X x x x X X X X = x = ee = = x x X X x Xx X X X X X = x x x X X x = = zy ie = x = X = X X = x x X = = = = Bu X = X X X X = x X x x x x г г EEE DR £ O = = 3 uw : > : 5 — N o = со O о © o © = s a 8 = un + 2 © un + о © = a o. 8 o) LINN “NO Hwa Wo LIN NO ---N W- -INN WO dwn NO nm п wn то wn WO LINN NO a LINN “NO LINN TWO LINN TWO LINN TWO 1 ий MN 11-N NO ПИ ahvi= LINN TWO ji LINN NO LINN Wo i-wn 1MN- TI о г © x fe) © о a = > * saloads Aıwe oo (panunuoy) 921 MARINE BIVALVES OF THE FLORIDA KEYS (sanuyuoo) XX XXX KK KK KK KK XK x X X X KK XK хх 650005 ‘аи x X x - x x x x х - x - = 2 > E x х = = X х x x x x x = x x x х < . E a x x Xx x x x - - x x = — 3 x x и x u = = = À x = x x pl = x er = > x X x X ES x x vi х x x X u x = 3 x x x = x = Bs “= en @ 3 = = то ‘ву © ~ 2 © 3 a A a © © Dr * 8 oOOV2TOOoOTOoONTONNOOTONOOTOON mela MAN м LINN Wo INN NO LINN “NO a m j= ate own W- тп No LINN Wo === W- (je О LINN Wo LINN Wo LINN NO IN 7 -IN WO 11-N WO LIN- 1-0 Inn 1MN- LINN TWO Al ie LINN TWO 7 =a l= 7 U D 2 x fe) 5 * (pest ‘Aes) SUOJIA91q EWO9EN (1581 ‘Aes) вемези! зцашиодет (96/1 “Ja¡Buadg) еибеш eujoise7 (857 | ‘sneeuuly) а ебилае! euljoi2e7 (8/11 'uog) eearund eujeAinz (Spel ‘swepy 'g ‘9) Suayu eureAinz (6181 ‘uUoyn]) ejesu! euljayung (1621 ‘uewo) esonbue euljjayung (zz81 ‘Aes) ejewaye euenz (0061 led) виеомеше eu!yejojd1/3 6£6L Japyay мозиэриау CULUO} SJEJUOUO еэдеш/Э (66/1 'Аэичэупа) е]5пе; eibedoay (Erg, ‘Aeyaq) ло/о3/5лал snmnbuy (0061 lea) Snuexe} sninbuy (PL8L “USA ‘3 VW) sn¡¡9ue] зпибиу (9991 ‘peiuo9) $/5иэвеаше] snınbuy (1881 lea) snopuegás snınBuy ($961 'ssog) зпиидола snınbuy (7961 'ssog) sniaweled sninbuy (veg, ‘Aes) snıaw sninbuy (2581 ‘uosduuns) s/Be sninbuy (oye! 'AÁajueH) ир/поб snjAiooy (1y81 ‘релиоЭ) ejxajold sdojjayisuZ (eye | 'Аэшен) eJeupenb ejouajseg (0581 ‘Znj994) еова!/э enora3seg saioeds seplul|jol sepieyods Ale + (panuyuoo) BIELER & MIKKELSEN 522 (sanuyuoo) x | о | IN N (2281 ‘Âe19) ejeunes emueg sepiuipala] x x x x x à go as 69 IMAN NO 86/1 'Bulpoy) ways! JUIL х x x x = 0 a ze lawn NO 6981 'зэАечзэа evayiwenbs ги! 1 x x x x x | ЕО -5 95 Lawn NO 86/1 'snaeuu!7 eJe/pEI ви! 1 x 0 LOS 02 | 7 IN 061 ‘uosduls $ ¡leg e9/s/ad eure] x x x x = 2 de -5 $5 ми IW (Zr8L ‘2пЮэч) вде}5 мо влор!/е1 x = x x x 0 = E) IN (89/1 'snaeuur7) siuoJIsid е|иби$ X x x x = 0 go -S 9 IMAN NO (Le! ddııyd) $//делш еб $ x x e = = 0 => = Е 1 7 8561 ‘AUIOON 8 и0$$10 /qqeb 2/6} $ х x x x = 0 =) = =IN= “NO (86/1 'snaeuu!7) eueuleo eylıbus x x x x x tt 90 as vec IMAN МО (9081 ‘Aqiamos ‘F) sılwıs EJNSSIOS x = x = A 6 go =S CC LINN “NO (zz81 ‘Aes) sul emssios X x = = 0 -O as 6 LINN IW (zreı ‘AuBiqiO.p) eunuqosuos enssios x x x = = 0 go =S |. €l м IN- (zve1 'ÁubiquO p) eueapued ejnssios x x x = = 0 O (GS. 9 e le O, (zreı 'AUBIQUO Pp) з/5иеошииеш eosuayy = À x - = 0 = == 0 7 (86/1 “Ya¡buads) eunjejsus ES x x x = = С go a oO LINN WW (pz81 ‘Aes) eyeujsinbae eosuoyy x = x x Ñ e g- as LL iM ИО (peg, ‘Aes) виа} ewoseyy x 0 => EP € NS 0061 ‘ed /имод/ебе} ewoseyy 0 =0 =S С 1-N= IN - 1061 'uosduwis $ jjeq e/awopnasd ewoseyy x р а- = | N N 9681 ‘IEG //эузуш ewoseyy = = x = = 0 oO as € ми “IN- 9681 ‘Hed е/пиш// ewoseyy 0 ONCE N Ne 0061 “eq ejenuajxa ewoseyy X = X = = 0 == == S =1-N IN - (26/1 ‘элэтбп.а) е]215и09 ewosey x = x x = y go =S 0 LINN NO (sy81 ‘swepy ‘4 9) euusa ewoseyy = E E о D E 2 я 7 AL я salsads Ale + ; AS = a >. : A © = (de) à о S < O O co + © = Ф = ae NE O о CRT NC A ES 2 р O © in = * о 2 =. 00 co (репициоЭ) 523 MARINE BIVALVES OF THE FLORIDA KEYS (senunuo) x XXX XXX Xx XXX x ххх ххх 550005 ‘APN 9661 ‘O81 х 9861 ‘риоциел х $061 e688! “eg 688L-/88L ‘uosduIs © = 891 < © oO y„ATJIoooowowmorowoo-oomowo hy ооо оао sooy = — + LINN Wo LINN Wo лип TW- LINN WO === D LINN NO es CE “Wn “MO --N- WO 1 1 LIN- NO 1 0, No 1 7 1 3 a N --N- W- ===) he LINN W- 1 AN NS 1 n IN 1 1 7 7 TI о D 2 x (0) 5 + Z061 ‘Ned /moÁzew эиощЭ (zz81 ‘Aes) вела эиоцЭ (0681 lea) вешМопе ejsıjed (zy81 ‘AuBiqio,p) еиеиадпе арлеэо]ешоиу (2581 ‘swepy ‘4 I) 1040$ ешиерэ^ца (ges, 'Ча!!ча) evadsejwas euuepnofjy4 (zreL 'Аибю4о.р) eueapued eyjaiuejay (zz81 ‘Aes) eyejound виороа!а 8581 ‘лэибем з1илода/эпи ejuopojdig 1061 ‘цозаци$ 3 пеа EJEJOU вшороаа (1641 ‘ujewo) ебецаоелоэ ебецао\|ело) (zv81 ‘AuBigiO,p) вепизи} елзели1 (5881 “USA 3 VW Ul US $ шел ‘2 v) $риеиб елзе/ц 1 9881 ‘leq ¡uosduys erseuy | ¿2281 ‘\>лешел euljoaseyd еэелц | , 7961 ‘Hd /UOSHLIOU e/9eJy 1 (5081 'nBeyuoyy) evojsip eroeiy | (9881 “1eg) зиебе/э elysng 9881 ‘EG /yduey sniseyjousyjs y (228 | 'uoun |) зп/оэ/еш елорэлэ | ez6| ‘чэзреа /ddejo opera ez6l 'dde¡9 1yosueq оре/э 1 (2161 'yosueg) ¡xouy 0P818J0J0N (6y81 'saBejaljeno эр) snjejeoipad snpolA7 (LEBL 'y90yY IION) е/земашиу enueg salmads SepuauanA eepiunfun aepizadeJ | aepuiseAy | SepIIoeJy Aıwe (panuyuoo) BIELER & MIKKELSEN 524 (зэпициоЭ) x x x = = 0 == == 0 ==== == (1281 зиээшл-иес Álog) елэдпа ee18qn4 x x = > = р go as 1Z LINN WO (6781| ‘peiuo9) eaindundejul ejjsuagng x x x x = ve go as ezı LIWN WO (9681 ‘112Q) /UoSduIs 18714 x x x x x 6 go as 6S LIWN TINO (8z81 ‘SHUSN) SNJEUILUIN] лес = = x x x 0 к = gE === = (8611 ‘snaeuuly) эцо/р Jeli x с а- as SL 17 7 NO (1961 1эбуэмчцэ$) зтерлоэ Jeg x 0 = -- | A (81/1 'UJOG) SNJEUIDIIO ле“ = = x x = 0 SE == 0 7 (1621 ‘инэш9) зприе лей x x X = x 124 go as LOL МП NO (8581 ei) мэ]5 eydABued x = x = = OL go as Gt LINN “NO (9781 'pesuo)) eyenbu] suejseieg x = x x = | —O =S y —-IN- W- (8s/| '“snaeuul7) виеиаэлаш eueugauayy X = X = X | —— == y == IN - (L6ZL ‘UjeWO) sısugıyoadwesa eueussuayy x = x x = | > == 2 LINN W- (98/1 Yooyybı7) esogwiu ejsıjesonsew x x X x = 8 go as LE лип =IWO (86/1 'snaeuul7) ejenoeu ejsıjesosew x = x = x с =© as al 11n IN- (1911 'snaeuu!7) ended елоцаолт x x x = = 0 go as OC 11-N WO (1y81 ‘релиоЭ) ejer e, елоцаолт7 0 == а- | NS (ZS6L ‘Aelind) /youejo елоцаолт x X X X = bl go as 82 wn МО (сут ‘swepy 'g I) EUU8, вртоэ 2 a — X = 0 OO 11-— WO (9881 ‘имапен) euyebna snuaniqo/5 x x x x = 0 =0 as y 11-N TINO (2181 ‘UAMIIIQ) ериби зпиэл!до]=) к. “= x = = 0 == =S р AW М (peg) ‘иэно1) ewweb euwes x Е x x = С go as ZL INN WO (9781 ‘peiuoD) зиеба/э ejuisoq x = x = = G = © =S i INN W- (0581 ‘2AS9M) SNISIP eluISOG x x x x — р go =S ZL лип “TINO (2981 'zn¡99y) sinua] eyaUljaAD x = x = = L =) d= 9 = ANS (2061 lea) пи//иби$ зпецашоэлЭ = © 8 = = 2 Я a au о salads Ale + us) O 3 0 xe) D o Tel - iat) yn = 9 a $ = = : © n A See 6 2 oO = de) = QQ | = 9 o о o Ze: a D [0)) co N mp E о => O Ww © En 00 co (panuyuoo) 925 MARINE BIVALVES OF THE FLORIDA KEYS ‘dep Aq uorolse1 pue ‘заоэ4$ рэцциер!-А|п;-иец]-5$9| jo uoisnioxe 'AÁwAuouAs чбполц} pajeunuja иээа элец selseds 9, ‘Аэллпз $14} Aq рэ}зн Анецбыо элэм sargads сё чбпоцим ‘000 ‘121814 Y USSIDANINss 9661 ‘UUIND 3 SUOÁT 44 ‘эзеавер и! SP10981 рэфоэ|оэ-элн JO лэашпи :3A17 y, ‘epis{eq = g ‘эр!зиеээо = O :(AJuo эзеавер) epiSs ‘deep = а ‘mojjeys = $ :(Ajuo aseqejep) deg: ‘эзеавер и! SP10981 JO лэашпи :зрлоээч,., ‘sebnyo Aiq = 1 ‘Shay 18M07 = 7 ‘shay эррии = W ‘Shey 1eddf = п :uonnqusiq sÁsy epuo|4, ‘P10981 элпуелэ}| = 7 ‘иэшюэ4$ шпэзпш = |y ‘чоцоэоэ |ешбныо = O) :S891n0S веа»„ 705 59. vlc 966 98 1627 ATA 68€ :S[8]0 | = = x x = 0 win == 0 1 1881 “eq еищио! EIPJOA OBPIIPIOA = — = x = 0 == == 0 7 (9881 ‘112Q) eaısan EÂWO9IS8A IN X = x X = 0 =0 а- ÿ 11-N WO zv81 ‘AuBiquo,p вело eunuoBr x en = x z 0 =0 а= G Al Sal (vrg8L 'Чача) ejejsooynoe eyadisouids X = = X = 0 = ÓN | ee IN = (1881 lea) eueuayssy suey 0 — (©) а- | 1 1 (1881 lea) ewiıssyueßeje eoulong эерируоээл X X X = = С go as 6 LIAN TINO с061 “eq ¡uosduys ejjeuuesuel X 0 == а- | 7 7 (1061 'uosduis $ неа) еиелда/пэ в/эииэзиел | X x X X = 0 do as 7 LIAN NO (zv81 'AuBIquO p) еиешедпо е]эииэ$иел1 X X X x X 8 go as OV LINN NO (ves! ‘иеа) ешрелиоэ в]эииэ$иел1 x 0 =e as 0 аа Е (8,8, 'Уолешел) в/эиоби] ejan! = A X X A 0 ES = | in = 8411 'UOg зэарюдоеш EJSAI] X 0 =0 =S С =a —=0) 6561 Jepyay виериоц е/эл1 | ig a € IN N 061 “IQ S/U09EqE ejanı / x x X X x y go =S Lv wn NO (8181 'Уолешел) еэешбла eejoou] X 5 x = = G go as SE LINN NO (8981 ‘зэщон) snib еэ/роии1 = Qo o = - = 2 = A au y зэюэас Ale + fee) D 3 = D a. o, tsi 8 > y DERE a re Е # : ® a © >) 00 5 < о IE ee cant A EIER 5 3 2 + oS ER S CA ©, oO) co 1 = oO => oO © 09 ze co (репипио9) 526 BIELER & MIKKELSEN known cases, the records were entered as “dead” in the database. To maintain a measure of relative abundance in our data in the face of the obviously mixed sampling effort in this heterogeneous dataset, our analyses employed standardized data, that is, data converted from raw abundances into percentage abundances. In addition, we re- peated analyses as (a) untransformed (thus allowing common species, i.e., those with greater inferred abundance [= more database records], greater influence) and (b) presence/ absence-transformed (thus giving rare and common species identical levels of influence). Results of these multiple runs are reported where relevant. All statistical calculations and resulting graphs were generated with the software pack- age PRIMER 5 (ver. 5.2.8) for Microsoft Windows” (Clarke 8 Warwick, 2001). Each species-by-location matrix was initially con- verted into a triangular location-by-location ar- ray of similarities by calculating the Bray-Curtis Similarity Index between location pairs, based on joint species abundances or presences. Further analyses employed hierarchical clus- tering into sample groups (CLUSTER; e.g., Everitt, 1980), as well as ordination by non- metric Multi-Dimensional Scaling (MDS; Kruskal & Wish, 1978), which constructs a rank-similarity-matrix-based sample configura- tion (or “map”) in a specified number of dimen- sions. The inter-point distances in this map have the same rank order as the correspond- ing dissimilarities between samples. Stress values of the two-dimensional MDS plots indi- cate the difficulty involved in compressing the sample relationships into low-dimensional space (a perfect fit has stress = 0, whereas stress > 0.3 approaches arbitrary placement in two-dimensional ordination space). Permu- tation-based hypothesis testing was performed by non-metric, one-way, pair-wise analyses of similarity (ANOSIM, an analogue of univariate ANOVA; Clarke & Green, 1988), which tests between groups of multivariate samples and was here used to determine significant differ- ences between regional samples (with К val- ues given for standardized data unless otherwise stated). When such differences were encountered, exploratory analyses of similar- ity/dissimilarity of percentages (SIMPER; Clarke, 1993) were performed to determine which species were principally responsible for within-group similarity and between-group dis- similarity. The SIMPER routine implemented in PRIMER examines the contribution each species makes to the average similarity within a group (e.g., which species “typify” the bayside samples) as well as its contribution to the av- erage dissimilarity between two groups (e.g., which species are good “discriminators” be- tween bayside and oceanside samples). Faunal Comparisons As in the previous analysis (Mikkelsen & Bieler, 2000), other western Atlantic regions for which comprehensive species lists could be compiled were used for comparison with the Florida Keys fauna. These regions were se- lected as those having similar ecological com- plexity, including estuarine/mangrove habitats, coral reefs, and shallow-to-deep water com- ponents. Their selection and delineation was admittedly arbitrary and depends greatly on available published and unpublished data. We revised the taxonomic listings of the four pre- viously used comparative areas; those for two areas were substantially modified from the pre- vious analysis, and so a full list of sources is given here: Gulf of Mexico (compiled from Steger, 1962, West Florida; Haas, 1940 and Gundersen, 1998, Sanibel Island; Lipe, 1984, Tampa Bay; Lee, 1999, Cedar Key; Lipka, 1974, Flower Garden Bank; Shelton, 1997, Alabama; Garcia & Lee, 2002, 2003, Louisi- ana; articles in Texas Conchologist, 1964- 1999, Texas; Garcia-Cubas, 1968 [which includes records from Parker, 1959], Mexico just south of Texas), Cuba (from Aguayo & Jaume 1947-1948; Espinosa et al., 1994), Yucatan (from Jaume, 1946; Rice & Kornicker, 1962, 1965; Ekdale 1974; Garcia-Cubas, 1981; Н. Е. Vokes & Е. H. Vokes 1984; Cruz-Abrego & Flores-Andolais, 1994), and eastern penin- sular Florida (from Voss et al., 1969, Biscayne National Monument; McGinty & Nelson, 1972, Pompano Beach; Reed & Mikkelsen, 1987, eastern Florida Oculina coral reefs; Lyons, 1989, Hutchinson Island; Mikkelsen et al., 1995, Indian River Lagoon). Two new listings were compiled for this study: Bahamas (from Dall, 1896; Clench & McLean, 1936; Lawson, 1993; Redfern, 2001 [plus full species list pro- vided by that author]; and unpublished data (PMM) from the island of Andros), and Ber- muda (from an unpublished species list com- piled by the late Russell H. Jensen, Delaware Museum of Natural History). Incompletely iden- tified taxa (e.g., to genus- or family-level only) were excluded from these analyses. MARINE BIVALVES ОЕ THE FLORIDA KEYS 927. RESULTS AND DISCUSSION Characterization of the Fauna A total of 389 bivalve species are now re- corded from the Florida Keys (Table 1), an in- crease of 28% over the previously recorded total (Mikkelsen & Bieler, 2000; as adjusted; Table 1), and a 139% increase over the previ- ous FKNMS checklist (Lyons & Quinn, 1995). The Florida Keys bivalve fauna presents wide taxonomic diversity, including 61 families and 212 genera. The most diverse families are Tellinidae (47 species), Veneridae (39), Lucini- dae (22), Pectinidae (22), and Mytilidae (19). The fauna includes 86% (279 species, plus six now regarded as synonyms) of the 331 shallow-water (< 37 m) bivalves of Florida (Lyons, 1997), plus 24 additional species re- corded from shallow water. It also includes 62% (347 of 557) of North American, Atlantic coast marine bivalve species from < 200 m depth, plus an additional 42 species not in- cluded on the Turgeon et al. (1998) checklist. 355 (91%) of the 389 “ever-recorded” bivalve species were represented in the database. Of these, 71 species (20%) were abundant, 116 species (33%) were common, and 168 spe- cies (47%) were rare, as defined in Materials and Methods: Analyses (Fig. 4). The most fre- quently collected species were Chione elevata (372 records), Codakia orbicularis (352), Brachidontes exustus (334), and Barbatia cancellaria (297). For comments on extremely rarely encountered species (singletons and doubletons), see below. To eliminate possible artifact in the database caused by sampling bias by other collectors, we considered these same statistics for our original collection records only (212 species; Fig. 5), and found them comparable in scale: 39 species (18%) abundant, 72 species (34%) common, and 101 species (48%) rare. The frequency distribu- tion of occurrences appears in Fig. 6. Of the 389 ever-recorded species, 163 spe- cies (43%) have never been recorded as live- collected (Table 1). This exceeds the already high percentage of the total fauna recorded by Bouchet et al. (2002; 28.5%) and Kidwell (2002; 25%). The data were insufficient to reveal strong indications of species losses or gains through- out the Florida Keys or at specific localities (largely due to the paucity of dated collection data). Our previous analysis (Mikkelsen & Bieler, 2000) noted loose correlation of records of the Lions’ Paw scallop (Nodipecten frago- y y Common 4 33% Abundant À 20% / 4 Rare 47% > Common © 34% Abundant y 18% £ / Каге 5 48% FIGS. 4-5. Percent of records in the database categorized as abundant (> 50 records), common (10-49 records), and rare (< 10 records). FIG. 4: For all collections; FIG. 5: For original collections only. Number of taxa OS TS 4 898,216 8 Number of occurrences (log-2) FIG. 6. Histogram of the frequency distribution of occurrences, plotting number of taxa versus number of occurrences (log,) binned to full integers. 528 BIELER & MIKKELSEN sus) with the popularization of scuba diving (especially on artificial reefs), and of two spe- cies of false mussels (Mytilopsis зрр.) with increased recreational boat traffic and/or fresh- water input into Florida Bay. Three additional notes can now be made. (A) An initially suspi- cious record of the Indo-Pacific black-lipped pearl mussel, Pinctada margaritifera, in the Dry Tortugas in 1893 (Nutting, 1895) has been reinforced by recent, irrefutable records along the eastern coast of Florida (Chesler, 1994; Carlton, 1996), and could thus be the first, considerably earlier, record of this introduc- tion. (B) The discovery of a living, large-bod- ied, Indo-Pacific gryphaeid oyster on a shipwreck off the Middle Florida Keys strongly indicates a recent introduction (Bieler et al., in press). (C) À predicted invasion by the green mussel, Perna viridis (Linnaeus, 1758), from now-ubiquitous populations in western Florida (Benson et al., 2001), has not yet been dis- covered but must be anticipated in sheltered, reduced-salinity habitats of Florida Bay simi- lar to those colonized by P viridis in western Florida. Faunistic Relationships — Within Subregions of the Keys Overall Regional Comparison: Within the database of 12,382 records, 99% could be assigned to one of the Florida Keys subre- gions. 354 species (91%) were recorded for at least one subregion (the remainder included 34 species coded only as Florida Keys, plus two species in the database solely for other parameters). 163 of the regionally assigned species (46%) were recorded from all Florida Keys subregions, that is are ubiquitous at this spatial scale (“UMLT” in Table 1). 74 species (19%) were restricted to a single subregion (Upper Keys 14; Middle Keys 12; Lower Keys 27; Dry Tortugas 21). The remaining 116 spe- cies (30%) were recorded from two or more subregions. None of the MDS and CLUSTER analyses of untransformed (raw numbers of occur- rences/records), standardized (converted to percentage abundances), or presence/ab- sence data demonstrated any serial pattern of community similarities along the island ar- chipelago. Likewise, CLUSTER and MDS analyses of all records showed no clear clus- tering of Upper, Middle, Lower, or Dry Tortugas records, and the ANOSIM analysis confirmed this impression (global R = 0.036, p = 0.2). The strongest R-values under any data trans- formation were generated in pair-wise tests between the westernmost records (Dry Tortugas) and the other groups. To explore this as a potential artifact of the comparatively low number of Dry Tortugas records, the analyses were rerun using only Upper, Middle, Lower, and Western Keys subregions (see Materials and Methods). While the MDS analysis indi- cated a relatively well-defined Western Keys group, the ANOSIM analysis again resulted in a low global R-value (0.17, р = 0.001), although significant differences (R > 0.53, p = 0.001) were evident between the Upper Keys and the Western Keys, as well as the Middle Keys and the Western Keys. To test for the effect of deepwater species (by omitting all deep stations, and again using the Western Keys subregion), MDS plots showed no distinct subregional-shallow groups along the Keys, with the exception of the West- ern Keys localities. The ANOSIM analysis showed low global values (R = 0.144, p = 0.004), and non-significant differences be- tween the neighboring Upper-Middle and Middle-Lower regions. We can conclude that the lack of significant differences found in pairwise comparisons of the subregions was not due to the inclusion of the deep stations, which led to the depth-related analysis dis- cussed below. Bayside Versus Oceanside: 288 (74%) of the 389 species could be coded as to whether they occur either on oceanside or bayside of the Keys axis. Because of the exposed position of the Dry Tortugas, records from this region were never coded for ocean- or bayside. Most Florida Keys bivalve species (169 species or 59%) occur on both sides of the island chain. 95 species (33%) were recorded as oceanside only, and 24 species (8%) were bayside only. An MDS analysis of all records (coded for position, i.e., bayside, shallow-oceanside, deep-oceanside, and shallow Dry Tortugas; Fig. 7) showed three groups of stations, with bayside and shallow-oceanside groups over- lapping. The single shallow Dry Tortugas sta- tion clustered with the shallow-oceanside group, which is not surprising considering its extreme position far removed from Florida Bay; in subsequent analyses it was coded as such. Three outlier stations (circled in Fig. 7) are like- wise readily explained. The Marquesas/ bayside station, located between Key West and the Dry Tortugas, is also far removed from the influence of Florida Bay and groups with the oceanic stations (and was recoded accord- MARINE BIVALVES ОЕ THE FLORIDA KEYS 529 FIG. 7. Two-dimensional MDS ordination of all station data, coded as bayside (В), shallow-oceanside (О), deep-oceanside (D), and shallow Dry Tortugas (DT), based on standardized (percentage- transformed) data and Bray-Curtis similarities (stress = 0.16). Circled outlier stations, top to bottom, representing Big-Pine-Key/oceanside, Marquesas/bayside, and Pigeon-Key/bayside, are discussed in the text. FIG. 8. Two-dimensional MDS ordination of shallow station data that were coded as bayside (В) or oceanside (O). Based on standardized (percentage-transformed) data and Bray-Curtis similarities (stress = 0.17). The two circled outliers are again Pigeon-Key/bayside, and Big-Pine-Key/oceanside. 530 BIELER & MIKKELSEN ingly for subsequent analyses). Pigeon-Key/ bayside, although farther east in the island chain in the center of the Seven-Mile Bridge, is in a relatively exposed position and subject to massive tidal changes that apparently bring shallow-oceanside elements into this part of the bay; it also groups with shallow-oceanside stations. In contrast, the Big-Pine-Key/ oceanside location is technically on the oceanside of the island chain, but includes a number of smaller islands and mudflats that provide bay-like habitats. The ANOSIM analy- sis (global R = 0.516, p = 0.001) showed sig- nificant differences between the shallow- oceanside and bayside communities (R = 0.356, р = 0.001), between shallow-oceanside and deep-oceanside communities (R = 0.874, p = 0.001), and between bayside and deep- TABLE 2a. Comparisons of similarity. Breakdown of average within-group similarities between bayside, shallow-oceanside (as in Fig. 7), and deep-oceanside stations into contributions from each species. Bold-font numbers indicate the percent contributions to within-group similarities within the top 55% of total similarity. Numbers in parentheses are values contributing to the remaining 45%, here provided for comparison. Compare to between-group dissimilarities given in Tables 2b, c. i L— SS EEE Species Bayside Oceanside Deep Chione elevata 9.97% 3.74% (0.23) Codakia orbicularis 7.18% 7.12% - Ctena orbiculata 6.13% 3.32% (1.27) Brachidontes exustus 5.33% (0.44) - Carditamera floridana 4.93% (0.61) - Angulus merus 4.76% (0.75) - Scissula similis 4.47% 2.55% (0.91) Laevicardium топот 4.46% (0.95) - Arcopsis adamsi 4.06% (0.86) (1.00) Pinctada longisquamosa 4.01% (0.88) - Barbatia cancellaria (2.93) 6.23% - Lucina pensylvanica (1.82) 4.20% - Arca imbricata (2.41) 4.10% 3.90% Агса zebra (0.97) 3.42% (0.84) Laevicardium laevigatum (1.91) 3.10% (0.89) Tucetona pectinata (277) 2.88% (0.84) Arcopagia fausta (0.50) 2.65% - Pinctada imbricata (0.44) 2.56% - Dendostrea frons (0.07) 2.36% (0.81) Lima caribaea (0.04) 2.49% - Anadara notabilis (0.88) 2.12% (0.13) Chama congregata (0.28) 2.12% (0.84) Nemocardium peramabile - - 8.96% Plicatula gibbosa (0.01) (0.51) 5.83% Pleurolucina leucocyma - - 5.37% Pleurolucina sombrerensis - - 3.96% Spondylus ictericus - (1.42) 3.82% Lucinisca nassula (3.54) (0.92) 3.72% Pandora inflata - - 3.27% Spondylus americanus - - 2.97% Cryptopecten phrygium - - 2.88% Argopecten gibbus - (0.21) 2.19% Lucinoma filosa - - 2.14% Callista eucymata - - 1.97% Semele bellastriata - (0.07) 1.96% Nodipecten fragosus - (0.12) 1.80% A ee ee ee eee eee eee Cumulative percent contribution (bold font numbers only) 55.31% 54.97% 54.73% MARINE BIVALVES OF THE FLORIDA KEYS 531 oceanside communities (R = 0.897, р = 0.001) — pairwise tests with Dry Tortugas data, rep- resented by only two stations, have low sig- nificance levels. It is thus evident that Florida Keys bivalve communities differ between bayside and oceanside, and that there is a very strong difference between the shallow (0-35 т) and deeper (> 35 т) oceanside com- munities. A shallow-to-deep signal could also be de- tected within the near-shore oceanside com- munities when the faunas of inner patch reefs (in Hawk Channel) and outer bank reefs (at the edge of the Floridian Plateau) were com- pared. Analyzing only those records identifi- able as having been collected in reef settings (1,659 records, 164 species, from 29 reefs along the Florida Keys), MDS plots show two distinct but overlapping groups (with a low R = 0.209, based on standardized data). The strong overlap is likely the result of dispropor- tionately extensive sampling of the shallow- water back-reef rubble zones associated with the outer reefs. TABLE 2b. Comparisons of dissimilarity. Breakdown of average between-group dissimilarities between shallow-oceanside and bayside stations into contributions from each species. Species are ordered in decreasing contribution within the top 55% of total dissimilarity. Average dissimilarity = 69.23. OCEANSIDE BAYSIDE Average Average Average Dissim./ Species Abundance Abundance Dissim. SD Contrib. % Cum. % Brachidontes exustus 119 10.24 2:31 0.89 3.34 3.34 Chione elevata 3.69 8.96 1.84 182 2.66 6.00 Pinctada longisquamosa 1.42 7.40 ЭЙ 0.88 DIRT, 8.27 Carditamera floridana 1588 5.08 №51 35 2.19 10.46 Laevicardium топот 1.46 6.08 1.50 1.14 216 12.63 Ctena orbiculata 3.88 5.16 1.42 1.19 2.05 14.67 Codakia orbicularis 6.38 4.72 1.37 1.26 1.98 16.65 Angulus merus 1.65 4.04 1.36 128 1.97 18.62 Barbatia cancellaria 5.92 3.16 1555 alla 1.95 20.57 Arcopsis adamsi 185 5.24 1533 1721 1.92 22.49 Tucetona pectinata 4.46 2.92 1.30 0.91 1.88 24.36 Caribachlamys sentis 4.31 0.28 1530 0.61 107 26.24 Lucina pensylvanica 5.00 1.40 1.08 1.36 1.56 27.80 Lucinisca nassula 1.58 2.76 1.05 1.17 1252 29.32 Scissula similis 2.92 3.68 1.02 1.52 1.48 30.80 Агса zebra 4.00 1.08 0.93 1.41 1.34 32.13 Cumingia vanhyningi 0.42 3.16 0.91 0.99 1882 33.45 Argopecten irradians 0.54 3.84 0.90 0.48 1.30 "34.75 Periglypta listeri 3.08 182 0.88 1.04 127 36.02 Laevicardium laevigatum 4.35 1.88 0.88 1533 1827 37.28 Nucula proxima 0.19 1.88 0.87 0.88 1.26 38.54 Pinctada imbricata 3.69 0.76 0.86 1.28 1.24 39.79 Lima caribaea 3.46 0.20 0.85 1.47 1522 41.01 Anomalocardia auberiana 0.54 3.00 0.84 0.85 1221 42.22 Limaria pellucida PT 2.12 0.83 0.69 12 43.42 Anadara notabilis 2.73 1.04 0.83 1.16 122%] 44.63 Modiolus americanus 2.31 1872 0.83 dell 1.20 45.83 Arca imbricata 4.42 1.68 0.82 1.41 IS 47.01 Dendostrea frons 3.23 0.24 0.80 1.46 1.16 48.17 Chione mazyckii Vals 1.92 Or 0.85 El 49.28 Pitar fulminatus 0.46 1.60 0.77 0.83 Wed 50.39 Chama macerophylla 312 0.92 0.76 1.16 1.10 51.49 Arcopagia fausta Salz 0.72 0415 1.48 1.09 52:58 Pitar simpsoni 1.54 2.80 0.73 1.08 1.06 53.64 Trachycardium muricatum 1.62 1.32 0.69 1.16 1.00 54.64 592 BIELER & MIKKELSEN TABLE 2c. Comparisons of dissimilarity. Breakdown of average between-group dissimilarities between shallow-oceanside and deep-oceanside stations into contributions from each species. Species are ordered in decreasing contribution to the top 55% of total dissimilarity. Average dissimilarity = 83.47. (Average dissimilarity between bayside and deep-oceanside groups: 89.21; table not shown here.) In an analysis omitting all deep stations and coding all shallow stations west of Key West as oceanside (as described above), MDS plots (of both untransformed and transformed data) exhibit overlapping clusters (Fig. 8). These differences were confirmed as significant by an ANOSIM analysis (R = 0.404, p = 0.001). Within-group similarity analyses (SIMPER; Table 2a) revealed that Brachidontes exustus, Carditamera floridana, and Angulus merus contributed primarily to bayside similarities (i.e, they were “typical” bayside species), while Barbatia cancellaria, Lucina pensylvanica, and Arca zebra contributed primarily to shallow- oceanside percentages. Both of these top- three lists include species associated with hard and soft substrata. Four species (Codakia or- bicularis, Chione elevata, Ctena orbiculata, and Scissula similis, all soft-substratum inhab- itants) contributed substantially to both ocean- and baysides. The different character of bayside and shallow-oceanside communities is thus largely based on a different fractional combination of the same group of species. Most of the within-group similarity of the deep- oceanside stations, on the other hand, was a result of a different group of species (e.g., Nemocardium peramabile, Plicatula gibbosa, and Pleurolucina spp.) that hardly overlap with the shallow-water communities. Arca imbricata is an exception in that it is a more or less “typi- cal” representative of all groups. Tables 2b-c list those species most responsible for the between-group dissimilarities. Of these, Brachidontes exustus leads the dissimilarity between bayside and shallow-oceanside groups, whereas Nemocardium peramabile leads that between shallow-oceanside and deep-oceanside groups). Different Patterns of Shallow-Water Commu- nities in Bay and Ocean: As shown above, analy- ses including all records (transformed or untransformed; with or without the deep- oceanside stations) revealed no clear cluster- ing of Upper, Middle, and Lower Keys bivalve data. However, a pattern emerged when only bayside records were analyzed (bayside records from west of Key West here omitted, see above). > An MDS analysis based on untransformed data showed clustering of each of the three groups, although with a fair amount of overlap. The ANOSIM analysis (global R = 0.326, p = 0.001) confirmed significant differences between Up- per and Middle (R = 0.384, p = 0.002), Middle and Lower (R = 0.173, p = 0.008), and Upper and Lower (К = 0.41, р = 0.001) groups. A SIM- PER analysis (Table 3a) showed that the ubiq- uitous venerid Chione elevata is an important (typical) member of all three subregions here analyzed. Brachidontes exustus, Pinctada longisquamosa, Laevicardium mortoni, and Limaria pellucida contributed most to the within- group similarity ofthe Upper Keys bayside com- munities, while other species, such as Tucetona pectinata, Modiolus americanus, and Laevicardium laevigatum, were strong contribu- tors to the Middle Keys bayside percentage. Major components of the Lower Keys bayside included Angulus merus, Lucinisca nassula, and Lucina pensylvanica. Upper and Middle Keys share a strong component of Arcopsis adamsi in their faunas, while Middle and Lower Keys share a high contribution of Codakia orbicularis. Tables 3b-c list the species most responsible for the between-group dissimilarities (lead by Brachidontes exustus and Pinctada longisqua- mosa for the dissimilarity between Upper and Middle Keys groups, and Тисеюпа pectinata and Ctena orbiculata for the dissimilarity be- tween Middle and Lower Keys groups). The bayside stations thus show a northeast- to-southwest pattern of Upper, Middle, and Lower Keys groups that appears to be driven by the relative abundances of certain species. This is not surprising because the northeast- ern part of Florida Bay is largely separated from open ocean waters by the island of Key Largo, undergoes substantial temperature and salin- ity fluctuation, and is strongly influenced by freshwater runoffs from the Everglades (Schomer & Drew, 1982), whereas the Middle and Lower Keys are (east to west) increasingly exposed to the open waters of the Gulf of Mexico and, through the interrupted island chain, to the open Atlantic. In contrast to the confirmed bayside pattern, an analysis of the shallow-oceanside stations showed no erlying MARINE BIVALVES OF THE FLORIDA KEYS Average Dissim./ 533 Species Abundance Abundance SHALLOW DEEP Average Average Nemocardium peramabile 0.00 3.14 Codakia orbicularis 6.38 0.14 Pleurolucina leucocyma 0.12 3.71 Barbatia cancellaria 5.92 0.14 Lucina pensylvanica 5.00 0.14 Pleurolucina sombrerensis 0.12 3.29 Caribachlamys sentis 4.31 0.14 Chione elevata 3.69 0.86 Ctena orbiculata 3.88 1.00 Lucinoma filosa 0.00 2.29 Lucinisca nassula 1296 2.57. Callista eucymata 0.04 1157 Pinctada imbricata 3.69 0.14 Tucetona pectinata 4.46 0.86 Arca zebra 4.00 0.71 Laevicardium laevigatum 4.35 0.71 Lima caribaea 3.46 0.00 Arcopagia fausta 3412 0.00 Plicatula gibbosa 2.55 1.86 Scissula similis 2.92 0.57 Divalinga quadrisulcata 1.69 2.14 Anadara notabilis 2.13 0.43 Gouldia cerina 1.27. 2.00 Abra lioica 0.08 0.86 Spondylus americanus 0.31 1.29 Arca imbricata 4.42 1.14 Pandora inflata 0.08 1.29 Chama macerophylla SAIZ 0.71 Periglypta listeri 3.08 0.14 Dendostrea frons 3.23 0.43 Cryptopecten phrygium 0.12 1.29 Modiolus americanus 223] 0.43 Acar domingensis 219 0.86 Isognomon bicolor 2.08 0.14 Papyridea soleniformis 1558 0.43 Dacrydium elegantulum hendersoni 0.00 1.14 Nodipecten fragosus 0.73 la Argopecten gibbus 12 2.00 Pteria colymbus 22 0.29 Chama congregata 3315 0.71 Puberella intapurpurea 073 0.71 Spondylus ictericus 2.88 1.43 Cardiomya striata 0.00 1.00 Arcopsis adamsi 1.85 0.57 Radiolucina amianta 0.50 1.57 Aequipecten lineolaris 9:12 1.29 Trachycardium muricatum 1162 0.14 Ctenoides mitis 2:62 0.57 Lindapecten muscosus 1512 UTA Eucrassatella speciosa 0.00 1257 Tellina squamifera 0.15 1.00 Carditamera floridana 1.38 0.00 Laevicardium mortoni 1.46 0.00 Americardia media Р.И и Cucullaearca candida 1.65 0.43 Dissim. 2.38 1.98 1.78 1872 1738 1.33 1225 1.14 teal) 0.98 0:95 0.93 0.90 0.89 0.89 0.88 0.86 0.85 0.85 0.83 0.81 0.79 0.76 0.76 0.76 0:72 0.71 0.70 0.70 0.67 0.64 0.62 0.62 0.62 0.62 0.61 0.61 0.60 0.59 0.59 0.58 0.58 0.57 0.57 0.57 0.56 0.55 0.55 0.54 0:53 0.52 0.52 0.52 0.51 0.50 SD 1.10 1.84 0.96 1.79 1.50 0.90 0.58 1:31 122% O2 1725 0.68 127 1.23 1.49 1.43 1.46 1.62 1.12 1.28 0.81 т 0.88 079 1.14 1.30 1.33 2 1529 1529 132 1.22 1.36 0.99 0.72 0.51 1.01 ITS 0:99 1.34 0.64 1.36 0.48 all 0.60 0.69 0.98 al 1.07 1.02 0.74 0.63 0.88 1.04 1.07 Contrib. % 2.86 Рэй 2.13 2.06 1.65 1.60 1.50 1:36 1.36 12417 1.14 ell 1.08 1.07 1.06 1.05 1705 1.02 1.01 0199 0.97 0.94 0.91 0.91 0.91 0.86 0.85 0.84 0.83 0.80 0.77 0:75 0.74 0.74 0.74 0.73 073 0.72 0.70 0.70 0.69 0.69 0.68 0.68 0.68 0.67 0.66 0.66 0.65 0.64 0.62 0.62 0.62 0.62 0.60 Cum. % 2.86 9.23 7.36 9.42 11.07 12.66 14.16 19.92 16.88 18.05 19.18 20.30 21.38 22.46 23,92 24.57 25.60 26.62 27.64 28.63 29159 30.54 31.44 32.35 33.26 34.12 34.97 35.81 36.64 37.45 38.21 38.96 39.70 40.44 41.18 41.91 42.64 43.36 44.07 44.77 45.46 46.16 46.84 47.52 48.20 48.86 49.52 50.18 50.83 51.47 52.10 O2 53.34 53:95 54.56 534 BIELER & MIKKELSEN TABLE За. Comparisons of similarity. Breakdown of average within-group similarities into contributions from each species. Bold-font numbers indicate the percentage contribution to within-group similarities of Upper, Middle, and Lower bayside stations within the top 55% of total similarity. Numbers in parentheses are values contributing to the remaining 45%, here provided for comparison. Compare to between-group dissimilarities given in Tables 3b, c. Species Upper Middle Lower Brachidontes exustus 15.75% (3.20) (1.32) Chione elevata 11.29% 4.86% 13.61% Pinctada longisquamosa 9.10% (2.66) (1.51) Laevicardium mortoni 8.20% ei) (1.67) Arcopsis adamsi 6.11% 5.08% (0.33) Limaria pellucida 4.24% (0.87) (0.38) Codakia orbicularis (3.30) 8.80% 7.56% Tucetona pectinata (0.42) 7.60% (1.75) Scissula similis (2.68) 5.72% 3.28% Ctena orbiculata (3.46) 4.98% 9.22% Barbatia cancellaria (0.75) 4.94% (3.16) Modiolus americanus (0.55) 4.41% (0.88) Laevicardium laevigatum (0.53) 3.89% (1.31) Arca imbricata (0.50) 3.82% (2.44) Angulus merus (3.55) (3.46) 5.44% Lucinisca nassula (1.76) (2.67) 5.27% Carditamera floridana (4.24) (2/15) 7.53% Lucina pensylvanica (0.17) (1.87) 4.53% Cumulative percent contribution (bold font numbers only) 54.68% 54.10% 56.45% TABLE 3b. Comparisons of dissimilarity. Breakdown of average between-group dissimilarities between Upper and Middle Keys bayside stations into contributions from each species. Species are ordered in decreasing contribution within the top 55% of total dissimilarity. Average dissimilarity = 62.07. UPPER MIDDLE Average Average Average Dissim./ Species Abundance Abundance Dissim. SD Contrib. % Cum. % Brachidontes exustus 26.38 3.22 4.44 1.62 7.15 7.15 Pinctada longisquamosa 20.38 1:67 2.90 1.42 4.67 11.82 Tucetona pectinata 1.25 5.11 2.56 125 4.13 15.95 Chione elevata 17.88 4.67 2.48 1.54 4.00 19.95 Argopecten irradians 10.25 156 2.21 0.82 351 23:51 Laevicardium mortoni 14.38 2.78 2.15 1.62 3.47 26.98 Codakia orbicularis 5413 5.11 1.79 1.30 2.88 29.87 Barbatia cancellaria 1.75 4.78 1.74 0.96 2.81 32.67 Limaria pellucida 7.13 1.33 1.51 0.81 2.43 35.10 Carditamera floridana 8.88 2.67 1.49 1.39 2.39 37.50 Anomalocardia auberiana 7.88 0.78 1.35 1.23 2.18 39.67 Cumingia vanhyningi 121%) 1.89 1.30 1.48 2.09 41.77 Arcopsis adamsi 10.00 4.11 1.28 1.31 2.06 43.83 Laevicardium laevigatum 1.38 3.00 122 1.42 1.97 45.79 Modiolus americanus 1.75 2.44 1.20 1.59 1.93 47.72 Angulus merus 6.25 3.33 1.12 1.41 1.80 49.51 Ctena orbiculata 6.38 4.00 1.08 1.09 VIS 51.26 Trachycardium muricatum 0.38 2.44 1.06 1.49 ETA 52.97 Scissula similis 4.00 4.22 1.02 1.46 1.64 54.61 MARINE BIVALVES OF THE FLORIDA KEYS 935 TABLE 3c. Comparisons of dissimilarity. Breakdown of average between-group dissimilarities between Upper and Middle Keys bayside stations into contributions from each species. Species are ordered in decreasing contribution within the top 55% of total dissimilarity. Average dissimilarity = 62.60. (Average dissimilarity between Upper and Lower Keys groups: 68.61; table not shown here.) MIDDLE LOWER Average Average Average Dissim./ Species Abundance Abundance Dissim. SD Contrib. % Cum. % Tucetona pectinata Sal 2.13 2.18 1.07 3.48 3.48 Ctena orbiculata 4.00 5.25 1.65 133 2.95 6.43 Arcopsis adamsi 4.11 1975 1.69 1.27 2.70 9.13 Codakia orbicularis Seti 3.88 1.67 1.36 2.67 11.80 Barbatia cancellaria 4.78 PTS) 1165 1.00 2.64 14.44 Carditamera floridana 2.67 4.00 1.51 1.35 2.40 16.85 Angulus merus 3.33 2.63 1.47 1.35 2.34 19.19 Chione elevata 4.67 4.88 1.34 1.49 2.14 21.33 Periglypta listeri РМ 1650 1.19 0.95 1.91 23.24 Lucinisca nassula 2.44 2.38 1.19 del 1.91 25.14 Nucula proxima 1.89 1.00 1419 1.03 1.90 27.04 Modiolus americanus 2.44 0.88 1.14 1.60 1563 28.87 Scissula similis 4.22 25 1.14 1.36 1.82 30.69 Laevicardium laevigatum 3.00 118 113 1.36 1.80 32.50 Chione mazyckii 1.44 ZO dl 1.00 178 34.27 Laevicardium mortoni PINS 1.50 1.08 1.49 173 36.00 Brachidontes exustus 322 2.00 0.96 ESA №53 9709 Arca zebra 122 1.50 0.95 1.24 153 39.06 Pitar fulminatus 1533 0.50 0.92 1.04 1.46 40.52 Anadara notabilis 1.44 0.88 0.91 1.09 1.46 41.98 Trachycardium muricatum 2.44 1.00 0.90 1.34 1.43 43.41 Pinctada longisquamosa 1.67 0.88 0.88 1.47 1.40 44.81 Lucina pensylvanica 167 1.88 0.83 1.44 132 46.14 Cumingia vanhyningi 1.89 0.63 0.78 1.06 1.25 47.38 Arca imbricata 2.11 1:63 0.76 1.60 1722 48.60 Anomalocardia auberiana 0.78 0.63 0.76 0.81 1.21 49.81 Isognomon alatus 0.89 2.00 0.73 0.79 1.17 50.98 Pitar simpsoni 1278 0.88 072 1.30 1.14 57.12 Polymesoda maritima 0.11 0.63 0.71 0.68 1.14 53.25 Modiolus squamosus 1211 1.00 0.70 1.10 e 54.37 Arcopagia fausta 0.67 1.13 0.70 1.14 1121172 55.49 significant clustering into Upper/Middle/Lower groups; many species occur in similar percent- age proportions along the Keys. The northeast- to-southwest pattern of bivalve community similarity inferred in the earlier study (Mikkelsen & Bieler, 2000) thus appears to be a reflection of differences on the bayside alone. Faunistic Relationships — Between the Florida Keys and Other Regions The bivalve species composition of the Florida Keys was compared with those of other well-documented western Atlantic locations (eastern Florida, Gulf of Mexico, Yucatan, Cuba, Bahamas, Bermuda) via a Bray-Curtis CLUS- TER similarity analysis. The non-Keys localities formed two groups (Fig. 9): Cuba with the Gulf of Mexico (72.9% similarity), and Yucatan with eastern Florida (74.9%). In the earlier analysis (Mikkelsen 8 Bieler, 2000), the Florida Keys grouped with the latter group. Here, the Florida Keys has switched affinities, now appearing closer to the Gulf of Mexico within the Cuba- GOM group. The change can be largely attrib- uted to the expanded datasets for Florida Keys, Gulf of Mexico, and Yucatan, which added spe- cies to each list, plus the revision of taxonomies, which reduced each list through synonymy. All localities except Bermuda showed a strong 536 BIELER & MIKKELSEN underlying similarity to each other (64.3% of shared taxa) that is essentially Caribbean in nature. Bermuda, although an extreme outlier and based on far fewer species than the other localities, is not considered artifactual in this analysis because it is based on the unquestion- ably thorough, multi-decadal compilation of Ber- muda malacofauna by R. H. Jensen, compiled from a similarly wide range of data sources. Our study shows a comparatively high bi- valve species richness in the context of the entire Florida Keys malacofauna, which differs from the finding of the most diverse mollus- can fauna studied so far. Bouchet et al. (2002), exploring a much smaller study area in New Caledonia (295 km’), reported 2,738 mollus- can species, of which 519 were bivalves (19%). Our study yielded proportionally more bivalve taxa: 1,684 molluscan species (current tally, in prep.), of which 389 are bivalves (23%). This could be a result of greater gastropod diversity in the tropical reef environments of the Indo-Pacific, or a reflection of the inclu- sion of more soft-bottom samples in our study. 50 60 Y oO Similarity (ee) o 90 100 Ber Bah Yucatan Relative Efficacy of Data Sources Museum and Literature-Based Data: In the expanded database analyzed here, our origi- nal collections account for approximately half of the total records, while literature and mu- seum records each comprise approximately one-quarter of the total (Fig. 2). Despite these proportions, original collections were the least successful of the three data sources in cap- turing the total species list (Fig. 10); this 1$ in close agreement with results of our previous analysis (Mikkelsen 8 Bieler, 2000). The new results differ from the previous, however, in the proportions of species recov- ered by museum collections versus the litera- ture. In the previous analysis, data derived from museum collections captured the great- est percentage of known species (77%), fol- lowed by published literature (73%). In the current analysis, we increased our coverage of bivalve-relevant literature by a factor of 4.4 (despite our prior assessment of having been “exhaustive”), and as a result, literature has E Fla Cuba FL Keys GOM FIG. 9. Bray-Curtis similarity CLUSTER dendrogram of bivalve species presence/absence in the Florida Keys (389 species, 67% of total list), eastern Florida (243 species, 42%), Gulf of Mexico (398 species, 68%), Yucatan (261 species, 45%), Cuba (356 species, 61%), Bahamas (244 species, 42%), and Bermuda (167 species, 29%). Total species list comprised 582 species. MARINE BIVALVES OF THE FLORIDA KEYS 537 Live original | Original Fe > Museum | All literature |. 10 0 10 20 30 40 50 60 70 80 90 100 41 o 40" 20° 0 40 50 60 70 80 90 100 Percent total species FIGS. 10, 11. Percent of total species list recovered. FIG. 10: By each of three main data sources (original collections, museum holdings, literature) and by live-only original collections in the present analysis; FIG. 11: By the various categories of literature: books, peer-reviewed literature, gray literature, and all literature categories combined. risen to the rank of most successful source (90%). Although we are confident in this re- sult, we are aware of a lingering conundrum involving literature-only records. In theory, if any literature-only records were based on er- ror (i.e., misidentifications, unsound locality data, etc.), then non-literature sources would not encounter those species. The success rate of literature data to capture the highest num- ber of “ever recorded” species could then be interpreted as the result of its own poor qual- ity. Nevertheless, most of these published records are here considered “reasonable” be- cause (a) they represent western Atlantic spe- cies that are known from neighboring waters (see below), (b) the volume of literature on south Florida mollusks is particularly extensive and relatively well studied (evidenced by the paucity of new species discovered by this study), and (c) the literature survey for this project can (again, but now more confidently) claim to be “exhaustive”, having canvassed relevant as well as spurious publications for more than a decade. While additional fieldwork and collections-based research are, theoreti- cally, open-ended, there is a finite number of published works with available data, and we believe we are fast approaching that limit. We therefore expect the relative species-list-recov- ery success of literature data to decline in the future, as fieldwork and collections studies continue to validate otherwise literature-only records. To interpret the rise of literature over mu- seum data, we analyzed the composition of literature in the previous and present analy- ses. We have continued to use the category “gray literature” to encompass unpublished theses and dissertations, shell club and other newsletters, agency reports, and other infor- mal checklists. The previously used category “traditional literature” is now divided into books (which usually provided Florida Keys records only as part of species ranges) and peer-re- 538 BIELER & MIKKELSEN viewed (or, for the older literature, mainstream- journal) articles. We examined both the num- ber of works within each category and the number of records generated by the works in each category. While gray literature contributed a substan- tial proportion of the records in the previous analysis (Mikkelsen & Bieler, 2000: fig. 1b), its contribution to the present, expanded dataset is dramatically greater. In this analy- sis, the proportion of gray literature to total works was nearly equal to that of peer-re- viewed literature (Fig. 12), yet it recovered 24% more of the total species list than did peer- reviewed literature (Fig. 11), and contributed almost three-quarters of the total literature- based records (Fig. 13). The importance of such “non-traditional” literature for regional biodiversity research thus cannot be over- stated, although these data can be difficult to vet. As production costs and editorial attitudes have driven the wealth of raw data out of main- stream publications, these “gray” means of Peer-reviewed 41% Books 16% Peer-reviewed 23% Books 3% 16 D FIGS. 12, 13. Percent of records. FIG.12: From peer-reviewed, book, and gray literature as part of the total works examined; FIG. 13: Total database records generated in this analysis. data deposition have become vital. The recent trend of providing data-rich electronic supple- ments (see, for example, Brewster-Wingard et al., 2001), № permanently archived, promises to help the peer-reviewed body of literature regain its information content. In view of the strength of literature-derived data, the question is posed whether the Florida Keys bivalve species list could have been ac- curately built through literature search alone. In general, the answer is positive but quali- Нед; such a search can only be effective if gray literature is included and can be adequately interpreted. Gray literature remains the most problematic to access and often is taxonomi- cally outdated, and could not have been ad- equately interpreted without a sound foundation in bivalve taxonomy and specimen- based knowledge of the fauna in question. 187 (48%) of the 389 species were found by all three data sources (museum, literature, and original collecting; “UML” in Table 1). Five spe- cies remain unique to our original collections and are thus new records for the Keys as a result of this survey: Mytilopsis leucophaeata, Ennucula tenuis, Cardiomya ornatissima, Gari circe, and Tivela floridana. Reflecting the source proportions, 17 species (4%) were unique to museum collections and 59 species (15%) were unique to literature data. Original Collection Data: There are at least two approaches to evaluate how close an origi- nal field study comes to capturing all available species. One method is to examine the пит- ber of singletons and doubletons (in this case, species recorded only once or twice in our database). Coddington et al. (1996, analyzing spiders) have argued that the number of such extremely rare taxa should decrease as sam- pling effort increases (although this could be argued against because original mollusk col- lecting, especially including empty shells, of- ten increases the taxon list with time; pers. obs.). 84 species (21%) appear as singletons (56) or doubletons (28) in this database (Table 1). Of these, 35 are not true singletons or doubletons, because they are represented by additional data sources (e.g., records indicat- ing “Florida Keys” only), although only one or two, respectively, was sufficiently robust to score for the database. 49 true singletons and doubletons (12.5%) thus remain, and although no “threshhold of sufficient sampling” was sug- gested by Coddington et al. (1996), this can be interpreted as a relatively low percentage of the fauna. This is considerabiy less than MARINE BIVALVES OF THE FLORIDA KEYS 569 the 20% singletons found by Bouchet et al.'s (2002) New Саедота field study. А closer look at the singletons revealed that in some cases, their collection records clearly explain the lack of re-collection. Of the 28 singleton lots, 13 (9 museum, 4 literature) are from offshore dredge samples and are thus not routinely captured by shallow-water collecting; this undersampled faunal component is being addressed by an ongoing deeper-water soft-bottom transect study (in prep.). Four are from a highly spe- cialized habitat (wood) that is rarely sampled; these are shipworms (Teredinidae), all of which are represented by fewer than three records in our database, which clearly 1$ an artifact of sampling. An alternate approach to evaluating how well an original field study captures all available species is to look at species-accumulation or collectors curves. In this method, the assump- tion is that a curve approaching zero slope indicates that most “possible” taxa have been captured. This has been demonstrated to work well with standardized collecting efforts in rea- sonably homogenous areas of habitat (e.g., Colwell & Coddington, 1994). However, such standardization is not feasible when explor- ing, as in our case, an entire marine mollus- can fauna and its great range of inhabited ecological niches. Here, species accumulation strongly reflects changes in effort and tech- nique over time (see below). In addition in this case, these curves are a useful tool because of the unusual circumstance that we “know” the potential total fauna from the composite dataset drawn from archived collections and 160 years of literature data. The difference between the observed endpoint of the curve and its expected ceiling (= all “ever recorded” species) could be indicative of undersampling, loss of species over time (which could mean a small- or large scale change of occurrence), or taxonomic problems/other errors in the ceil- ing records. For example, 17 species are known solely from Dall (1889a), listed as “ЕК, archibenthal, 50-800 fms” with unknown sup- porting source data: Abra longicallis ameri- cana, Bentharca sagrinata, Cyclopecten thalassinus, Juliacorbula cubaniana, Limatula setifera, Limatula subauriculata, Limea bronniana, Cratis antillensis, Myonera lamellifera, Nuculana vitrea, Pandora glacialis, Pectinella sigsbeei, Poromya albida, Propeamussium cancellatum, Thyasira grandis, Varicorbula krebsiana, and Yoldia liorhina. These species could actually originate from depths beyond our project depth limit (300 m or 164 fms), and can thus be consid- ered in most need of verification. Our flattened species accumulation curve points to a relative completeness of the col- lecting efforts. Accordingly, a log-normal fre- quency distribution of occurrences could be expected. However, the abundance frequency of Florida Keys bivalve species is not log-nor- mal (Fig. 6), but strongly right-skewed, with a median occurrence of 10 and rare taxa very numerous. To analyze our original collecting efforts ver- sus the total known species list, species ac- cumulation curves were generated for all four Keys subregions as well as the entire study region; only those for the entire region (all- original, live-original, dead-original) are pre- sented here (Fig. 14). After ten years of sampling, only 207 species (53% of total) have been collected in our original samples of all- original material. The dead-original curve closely emulates the all-original curve, indicat- ing, as do the raw numbers [5768 records, 4582 (79.4%) dead, 1186 (20.5%) live], that these two datasets are essentially identical; only seven species were never collected dead. Three abrupt increases (Fig. 14, arrows) can each be attributed to a new collecting method or new location: scuba operations (July 1995), the first Dry Tortugas cruise (April 1997) and the first samples on the Pourtales Terrace (July 2001); the reduced expression of these three events in the live species accumulation curve (Fig. 14) emphasizes the paucity of live-col- lected records. These past records predict that additional jumps in the curve will accompany each new data source as long as we can iden- tify new ways or places to sample. Two gaps to be filled have already been mentioned: float- ing wood (for shipworms) and deep-water soft substrata. The transect study mentioned above (in prep.) has already proven its worth in this regard: even in its early stages of data processing, 29 bivalve species have been added to the Florida Keys checklist. Examination of the species accumulation curve begs the question: could we have stopped collecting sooner and still accom- plished our goal to understand the bivalves of the Florida Keys? This depends on the per- cent “completeness” that one considers ad- equate, but is in fact a circular question, dependent on hindsight applied to the ten-year results. Fig. 14 shows that the curve leveled off substantially after the first Dry Tortugas cruise in April 1997, approximately three years into our study, when 75% of the total collected 540 BIELER & MIKKELSEN 1 16 31 46 61 76 91 106 121 136 151 166 181 196211 226 241 Collections FIG. 14. Florida Keys bivalve species accumulation curves (recorded species list versus original collecting events for Florida Keys) for all-original (—4—), live-original (—4—), and dead-original (— — ) records. fauna had been captured; note that this is less than 40% of the expected total. 90% of the collected total was not achieved until April 2001, and 95% was not reached until one year later (April 2002) after a second Dry Tortugas cruise that included sampling Pourtales Ter- race. Ecological studies most often only consider live-collected records, although we and oth- ers have argued earlier in strong support of including empty mollusk shells in biotic sur- veys. To press this point further, if we consider live original records only in this analysis (which comprise 20.5% of the total original records), the percent recovery of the total species list is reduced by nearly 20% to only 34% of the to- tal (131 species) — even despite the ten-year sampling period (Fig. 14). Nevertheless, analy- ses of these live-only records reproduce the same general pattern in bayside, shallow- oceanside, and deep-oceanside groups found in the full-data analysis (Fig. 15; compare to Fig. 7), giving further credence to use of empty shells in such analyses. A final advantage of the ten-year results is the ability to judge the effectiveness of Rapid Assessment (RAP) methods, such as have become standard and in fact critical to the speedy generation of diversity data necessary for environmental decision-making (Wells, 2002). Embedded in the dataset are four samples of live and dead specimens (FK-260, 261, 262, 264), taken during a four-day period (10—11 and 13 August 1999) from Looe Key Reef (Lower Florida Keys, са. 24°32.809’N, 81°24.158’W). These samples came from di- verse habitats, including spur-and-groove reef formation, rubble zone, and Thalassia seagrass beds, all in waters of 2-8 т depth, and include three components: visually located specimens, a sample of dead coral rock to be cracked for boring bivalves, and a sediment sample to be sorted for sand dwellers and smaller species. Together they can thus be considered a Rapid Assessment sample in terms of temporal and spatial dimensions, but are robust in covering a wide range of habi- tats and collecting methods. Looe Key has been historically well-sampled, and the data- base indicates a total expected species rich- ness of 104 species from 267 records (excluding those from the four RAP samples). Bivalves recorded from our four “rapid” Looe Key samples in August 1999 captured a total of 96 records and 62 species, or 60% of the expected total (Fig. 16); 11 of these species had not been previously recorded from Looe Key. Contrasted with the results of the ten-year study for the entire Florida Keys, this four-day study at Looe Key captured a higher percent- age of its expected fauna; we therefore con- sider these RAP samples as effective. In contrast, live-collected specimens in these four RAP samples recovered only 20 species (none of which were new records), or 19% of the expected fauna, which cannot be considered effective in representing bivalve diversity at this site. With regard to judging the success of RAP MARINE BIVALVES OF THE FLORIDA KEYS 541 FIG. 15. Two-dimensional MDS ordination of all live-collected station data, coded as bayside (В), shallow- oceanside (O), and deep-oceanside (D). Based on standardized (percentage-transformed) data and Bray-Curtis similarities (stress = 0.17). ANOSIM: global R = 0.443, p = 0.001. Compare to Fig. 7. sampling of marine bivalves, these results necessity of physical samples taken from underscore the great value of including empty multiple habitat types, despite the substan- shells in generating statements of species rich- tial commitment in processing time that they ness. In such studies, we further stress the require. Expected RAP RAP live 0 20 40 60 80 100 120 Looe Key Species FIG. 16. Species richness of marine bivalves at Looe Key Reef, including total recorded (= expected) species based on the total ten-year database, total species recovered by RAP samples (FK-260, 261, 262, 264) taken over a four-day period in 1999, and live-collected species from the same RAP samples. 542 BIELER & MIKKELSEN ACKNOWLEDGMENTS Major funding for this research was provided by the Comer Science and Education Foun- dation, supplemented over the course of this project by Harbor Branch Oceanographic In- stitution (Ft. Pierce, Florida), Delaware Mu- seum of Natural History, AMNH, the Bertha LeBus Charitable Trust and FMNH's Marshall Field Fund. As in the previous report (Mikkelsen & Bieler, 2000), we acknowledge FKNMS for collecting permits and general support, and many colleagues and staff mem- bers for access to their museum collections and/or assistance in collecting and process- ing data. The Bermuda species list of the late R. H. Jensen was provided by Timothy Pearce (Carnegie Museum of Natural History). Susan Kidwell reviewed an earlier draft of the manu- script; her constructive comments are highly appreciated. LITERATURE CITED AGUAYO, С. С. & М. L. JAUME, 1947-1948, Catalogo de los moluscos de Cuba, nos. 1- 363. Privately published, Havana, Cuba. BENSON, А. J., D. С. MARELLI, М. Е. FRI- SCHER, J. M. DANFORTH & J. D. WILLIAMS, 2001, Establishment of the green mussel, Perna viridis (Linnaeus, 1758) (Mollusca: Mytilidae) on the west coast of Florida. 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Institute of Marine and Atmospheric Sciences, University of Miami. 128 pp., 42 figs. WELLS, F. E., 2002, Molluscs of the Raja Ampat Islands, Papua Province, Indonesia. Pp. 37- 45, in: S. A. MCKENNA, G. R. ALLEN 8 S. SURVADI, eds., А marine rapid assessment of the Raja Ampat islands, Papua Province, Indonesia, RAP Bulletin of Biological Assessment, 22: Conservation International, Washington, D.C. WHEATON, J., W. C. JAAP, K. HACKETT, M. LYBOLT, М. К. CALLAHAN, J. KIDNEY, 5. KUPFNER, J. W. PORTER, V. KOSMYNIN, C. TSOKOS & С. YANEV, 2003, U.S. EPA/FKNMS Coral Reef Monitoring Project. Pp. 24-29, in: Sanctuary Science Report 2001: an ecosystem report card. National Oceanic and Atmospheric Administration, Florida Keys National Marine Sanctuary, United States Environmental Protec- tion Agency, State of Florida [Final version, 21 March 2003]. Available at www.fknms.nos. noaa.gov/research_monitoring/welcome.html WINGARD, С. L., $. ISHMAN, T. CRONIN, L. E. EDWARDS, D. А. WILLARD & R. В. HALLEY, 1995, Preliminary analysis of down-core biotic assemblages: Bob Allen Keys, Everglades National Park, Florida Bay. United States Geo- logical Survey Open-File Report 95-628, 30 pp. Revised ms. accepted 1 June 2004 MALACOLOGIA, 2004, 46(2): 545-623 CRITICAL CATALOG AND ANNOTATED BIBLIOGRAPHY OF MARINE BIVALVE RECORDS FOR THE FLORIDA KEYS Paula M. Mikkelsen' & Rüdiger Bieler? ABSTRACT Literature data contributing to a biodiversity survey of bivalve species of the Florida Keys are presented in the form of 361 annotated references and a documented species list. 389 species are recorded as identified to at least the species level: all except ten species can be traced to at least one literature citation. Thirty-one nominal species-level taxa were originally described from Florida Keys material or had their type localites designated as such. Annotations on synonyms, confirmed or suspected misidentifications, and a discus- sion of problematic geographic information are included, as tools for accessing and inter- preting the full body of literature, including 19" century works and 91 entries of “gray” literature (i.e., non-peer-reviewed reports, newsletters, unpublished dissertations, websites, etc.). This paper provides supporting data for an analysis of the bivalve fauna of the Florida Keys, based on a new database of over 12,000 original, museum, and literature records, included elsewhere in this volume. INTRODUCTION Biodiversity surveys rely on thorough re- views of pertinent literature, in the context of original field observations and studies of ex- isting collections. As in the case of historic collections, literature data can provide tempo- ral depth (e.g., showing the appearance or disappearance of taxa in a given region), taxo- nomic insight, and distributional information. However, literature data must be reviewed and interpreted (especially when not accompanied by verifiable illustrations or actual voucher specimens), because its information can be suspect due to the variable taxonomic exper- tise of authors as well as changes in nomen- clature over time. The Florida Keys is a highly diverse region, heavily impacted by human-induced change. For most of its invertebrate fauna, few if any baseline studies of species-level diversity ex- ist. However, for more than a century, the Keys have served as a source region for numerous academic studies of mollusks and have been extremely popular with the shell-collecting community. This has led to a steady but ex- tremely scattered outflow of formal and infor- mal publications containing distributional and natural history data for the bivalved mollusks of this region. The current catalog provides access to this wealth of information, which cumulatively can contribute to our understand- ing of past and present diversity. We have captured and critically reviewed the species records in each work, fully documenting any taxonomic interpretations of synonymy and/or potential misidentification; each species record and its interpretation, if any, are thus open to subsequent corroboration or falsification. Our goals for this literature review of molluscan diversity data are thus (1) to compile the lit- erature in an accessible format, (2) to inter- pret, and if necessary correct, the taxonomic information, (3) to interpret, and if necessary correct, the geographic information, and (4) to cross-reference the data, allowing taxo- nomic/geographic access by species. Most works, by design or necessity, take a cumulative approach in reporting distributional data and combine individual records into broad distibutional statements. In the case of west- ern Atlantic taxa, the given distribution often indicates the northern- and southernmost ex- tremes of occurrence, with little or no indica- tion whether this reflects merely two collecting events, spans a continuous area of distribu- ‘Division of Invertebrate Zoology, American Museum of Natural History, Central Park West at 79th Street, New York, New York 10024-5192, U.S.A.; mikkel@amnh.org 2Department of Zoology, Division of Invertebrates, Field Museum of Natural History, 1400 S. Lake Shore Drive, Chicago, Illinois 60605-2496, U.S.A.; bieler@fieldmuseum.org 546 MIKKELSEN & BIELER tion, or is based on extreme chance occur- rences of an otherwise more restricted range (the latter case an example of how summariz- ing accumulated information can dilute mean- ingful biogeographic data). The current work takes the opposite approach, where possible “deconstructing” literature data into occurrence information for concrete smaller subregions within the Florida Keys (i.e., Upper, Middle, and Lower Florida Keys, and Dry Tortugas). In an earlier analysis (Mikkelsen & Bieler, 2000), we evaluated the relative contribution of the different types of source data (original collections, museum records, and literature) toward capturing species-level Florida Keys bivalve diversity. The literature review included book publications (e.g., Johnson, 1934; Abbott, 1974), entire runs of scientific serials (e.g., American Malacological Bulletin, Bulletin of Marine Science, Journal of Molluscan Stud- ies, The Nautilus, The Veliger), shell club newsletters (e.g., American Conchologist, Texas Conchologist), the published papers of malacologists known to have worked in the Keys (e.g., Pilsbry, McGinty, Houbrick), and relevant agency reports (e.g., Lyons & Quinn, 1995; Vittor & Associates, 1998). Of all the bivalve species ever recorded from the Keys, literature data documented 73% of the total known species list, including 38 species not otherwise reported. “Gray” literature (non- peer-reviewed reports, newsletters, unpub- lished dissertations, websites, etc.) played a significant role in this contribution; traditional literature (books and peer-reviewed journals) recovered only 44% of the list, that is, effec- tively missing 56% of the known diversity. We concluded that multiple sources (including lit- erature) are most effective in producing a bi- otic inventory, although traditional literature was viewed as the least effective single re- source. Since completion of the earlier survey, we have continued our research toward a com- plete faunistic analysis of Florida Keys bivalves. Results of this expanded study, based on a new database of more than 12,000 original, museum, and literature records, is included elsewhere in this volume (Bieler & Mikkelsen, 2004). This catalog represents an annotated bibliography of the literature sources included in this and the earlier sur- vey, now expanded four-fold to comprise 361 bivalve literature sources, with special focus on gray literature data. Significant additions include one of the earliest surveys of Florida Keys bivalves (22 fully-identified species; Calkins, 1878), the earliest comprehensive survey (87 species [by today’s taxonomic cri- teria]; Simpson, 1887-1889), and Lermond’s (1936) checklist with 216 species, which al- though fraught with outdated nomenclature, surpasses the only other, much more recent, extensive compilation (163 species; Lyons & Quinn, 1995). It is noteworthy that both of these last two extensive compilations are gray literature; the present database includes 1886 records from 91 separate pieces of gray lit- erature [marked below by superscript “G”]. For the purpose of this study, we have in- cluded references to species only if specifi- cally listed from the Florida Keys (i.e., excluding those with broadly stated distribu- tions, for example, from Georgia to Venezu- ela, which theoretically could also include the Florida Keys). We define the Florida Keys as the waters surrounding the entire island chain from Broad Creek (about 25°21’М, 80°15’W) at the northern end of Key Largo (including Card Sound but not Biscayne Bay, southwest of but not including Old Rhodes Key) to slightly west of the Dry Tortugas (83°30’W). The bor- ders between Keys regions are here defined as between Craig Key and Fiesta Key (Upper Keys/Middle Keys), between the Seven-Mile Bridge and Little Duck Key (Middle Keys/ Lower Keys), and between Rebecca Shoal and the Dry Tortugas (Lower Keys/Dry Tortugas), with Dry Tortugas standing alone as a fourth region. A tangential east-west line was drawn through Florida Bay in the Upper and Middle Keys (at the levels of, from east to west, the Nest Keys, Russell Key, and the northern limit of Rabbit Key Basin), eliminat- ing what is more properly considered the southern end of the Florida Everglades. Oceanward, the depth limit was set at the 300 m (= 164 fms or 984 ft) isobath, which includes the historically and biologically important Pourtales Terrace. Our study area thus encom- passes (and exceeds) the Florida Keys Na- tional Marine Sanctuary, as well as all state and county parks, reserves, and management areas of this region, with the excpetion of the Biscayne Bay area and the northern parts of the Everglades National Park. Most of the 19" century records and the majority of deepwater collections off the Florida Keys stem from U.S. government surveys. The U.S. Coast Survey Steamers Corwin (1867, collections by L. Е. Pourtales), Bibb (1868- 1869, Pourtales; 1872, W. Stimpson), Bache (1872, Stimpson), and Blake (1877-1878, A. Agassiz), and the U.S. Fish Commission CRITICAL CATALOG AND BIBLIOGRAPHY 547 Steamer Albatross (1885) all sounded and dredged/trawled off the Keys. Their station records are generally well documented (e.g., S. Smith, 1889; Townsend, 1901). Most named localites (e.g., Key West, Looe Key Reef, Carysfort Reef, etc.), even if unaccompanied by geodetic coordinates, are likewise clearly part of the Florida Keys record. However, the identities of certain named localities remain problematic: First, Gordon Key (a single unnumbered site sampled at 68 fathoms (124 т) by the Ц. 5. Coast Steamer Bache in the 1870s; Dall, 1881, 1886; repeated from Dall by four subsequent authors, see below) is by all accounts part of the Florida Keys (Dall, 1903b; Johnson, 1934). However, no locality of that name exists to our knowledge on any chart or in any gazetteer, so its relative location within the Keys is un- clear. It is quite possible that the name is a corruption of Garden Key, the site of the U. S. Civil War era Fort Jefferson (and a popular ship anchorage at the time) in the Dry Tortugas (J. Clupper, pers. comm., August 2003). Accord- ing to our records, W. H. Dall never specifi- cally mentioned Garden Key in his molluscan reports until 1889 (original description of Lippistes acrilla Dall 1889: 391), so it is plau- sible that he earlier miswrote the island's name. S. Smith's (1889) official compilation of dredging stations by the U.S. Coast Survey steamers during this period shows the Bache operating off the Dry Tortugas in 1872, how- ever, no 68 fathom station 1$ evident; the Blake (the primary source of mollusks described in Dall's 1881 and 1886 reports) likewise is not recorded to have sampled any stations at this specific depth. S. Smith (1889) indicated a single 68 fathom station sampled by the Bibb in 1869, but this is charted near Alligator Reef in the Upper Keys. Without more definite data on the vessel and date of the expedition dur- ing which this deepwater station was collected, its location remains enigmatic. “Gordon Key” is the type locality of Corbula cymella. Another issue involves the confusion of the often-cited locality “off Sombrero”, which might refer to Sombrero Key/Reef in the Middle Florida Keys. “Off Sombrero, 54 fms” reported by W. H. Dall (1881, 1886) as a station col- lected by the Blake (or Bache?) sometime during 1877-1879 has been interpreted as pertaining to the island of Sombrero in the western Caribbean, east of the Virgin Islands (E. V. Coan, in Turgeon et al., 1998: 189). (The situation is further complicated by the fact that the station was unnumbered, and no 54-fm station, from either Sombrero locality, appears in S. Smith's (1889) compilation of stations dredged by the Blake.) The 54-fm Sombrero station was never called “Key” by Dall, al- though it was later and probably erroneously called Sombrero Key by Clench & Smith (1944) and Pulley (1952). In six bivalve ac- counts by Dall, the 54-fm station is combined with one at 72 fms (as “off Sombrero, 54-72 fms”), lending support to their co-identity if not their exact location. The uncertainties of these data and their source vessel led us to com- pare other contemporary expedition accounts, but these provided little additional understand- ing: Nine Bibb stations (sta. 5P-13P) were definitely sampled off Sombrero Reef in April- May 1868, but depths ranged from 111-517 fms (Pourtales, 1871: 169; Peirce 8 Patterson, 1880: 1; S. Smith, 1889: 958; with several of these stations on the official 1868 survey chart [No. 10] of the Straits of Florida). All 1872 ex- pedition stations of the British Steamer Chal- lenger from “off Sombrero” are referable to the Caribbean island (S. Smith, 1889: 973). In January of 1879, the Blake operated in the vicinity of the island of Sombrero (sta. 140- 141), listed by S. Smith (1889: 967) as “off Virgin Gorda,” and again at much deeper depths of 861 and 1,097 fms. In November of 1887, the Albatross sampled a single station (2750) south of Sombrero Island in 496 fms (Townsend, 1901: 403). For purposes of this study, both 54 and 72 fm stations listed by Dall as “off Sombrero” (the type locality of Lucina sombrerensis, L. leucocyma, Nemocardium peramabile, and Myrtea compressa), have been excluded from Florida Keys records. Three other localities hint at potential local- ity confusion but have been retained in this survey for lack of more conclusive data. (1) Turtle Harbor is a 8-9 т (25-30 ft) anchorage just inside Carysfort Reef in the Upper Florida Keys, yet Turtle Harbor at 40-50 fms (73-91 m) was listed by Hayes (1972) and Boss (1972) for two species (Pteria colymbus, Semele bellastriata). Although this could refer to a station offshore of the Upper Florida Keys (as in the case of Eolis sta. 58; Bieler & Mikkelsen, 2003), there are at least two other Turtle Harbors in the Caribbean: in the Baha- mas (Dall, 1886; Pulley, 1952), and off Isla de Utila, Honduras (a popular scuba diving site, with a wall to 300 m). (2) Long Key is a large island in the Middle Florida Keys, between Fiesta Key and Duck Key, but also refers to part of Bush Key in the Dry Tortugas (Clupper, 2003), and to the island on which the city of 548 MIKKELSEN & BIELER St. Petersburg Beach, Florida, lies, adjacent to Boca Ciega Bay. (3) Three Sand Keys are recorded for the Florida Keys: as the site of an historical lighthouse seven miles southwest of Key West, as a former name for Hospital Key in the Dry Tortugas, and as one of the Molasses Keys oceanside of the Seven-Mile Bridge (Clupper, 2003). At least one additional Sand Key exists in Florida, off Clearwater Beach in Pinellas County. In total, 389 bivalve species are included in this diversity survey, all but ten of which can be traced to at least one literature citation (those ten known to occur in the Florida Keys, but not previously recorded as such, are in- cluded in the catalog without following refer- ences). An additional 57 taxa are recorded from literature identified to the genus level, plus an additional 25 to family level; most of these probably represent previously listed species, although a few (e.g., Cymatioa sp., Semierycina sp.) are probably otherwise unrepresented. An additional four species remain of uncertain taxonomic status. Thirty-one nominal species-level taxa were originally described from Florida Keys mate- rial or had their type localites designated as such: Amphidesma laeta “Adams” Reeve, 1853 [now regarded as a synonym of Semele purpurascens (Gmelin, 1791)], had Key West designated as its type locality by Boss (1972). Amphidesma variegata Lamarck, 1818 [now regarded as a synonym of Semele purpurascens (Gmelin, 1791)], had Key West designated as its type locality by Boss (1972). Anadara springeri Rehder & Abbott, 1951 (now regarded as a synonym of A. baughmani Hertlein, 1951), from off the Dry Tortugas. Arca (Barbatia) balesi Pilsbry 8 McLean, 1939 [now regarded as a synonym of Fugleria tenera (C. B. Adams, 1845)], holotype from Missouri Key (paratypes from Key West). Argopecten irradians taylorae Petuch, 1987 [now regarded as a synonym of A. irradians concentricus (Say, 1822)], from Rabbit Key Basin [Florida Bay, off Long Key]. Asthenothaerus balesi Rehder, 1943a, from Missouri Key. Cardium (Fulvia) peramabilis Dall, 1881 (now Nemocardium), from various localities т- cluding Blake station 9 (111 fms) off Sand Key, and station 26 (110 fms), west of the Dry Tortugas. Subsequent lectotype selec- tion (Clench 8 Smith, 1944) restricted the type locality to off Yucatan, Blake station 36. Ctenoides sanctipauli Stuardo, 1982, with origi- nal (but not type) material including a speci- men from an R/V Eolis station at Sand Key Reef. Condylocardia floridensis Pilsbry 8 Olsson, 1946 [now regarded as a synonym of Carditopsis smithii (Бай, 1896)], from Ohio Key. Corbula cymella Dall, 1881, a Florida Keys endemic species, known only from 68 fms off “Gordon Key”. Cumingia tellinoides vanhyningi Rehder, 1939, from Lower Matecumbe Key. Cytherea (Ventricola) strigillinus Dall, 1902 (now Circomphalus), from off Key West. Dosinia floridana Conrad, 1866 [now regarded as a synonym of D. elegans (Conrad, 1846)] from “Florida Keys, Gulf of Mexico”. Jagonia orbiculata Var. filiata Dall, 1901 [a named form of Ctena orbiculata (Montagu, 1808)], from Florida Keys, deep water. Jagonia orbiculata var. recurvata Dall, 1901 [a named form of Ctena orbiculata (Montagu, 1808)], from Florida Keys, deep water. Modiola tulipa var. nigra Calkins, 1878 [now regarded as a synonym of Modiolus ameri- canus (Leach in Leach 8 Nodder, 1815)], an unrayed form from Key Vaccas (sic). Nucula calcicola Moore, 1977, from Key Largo. Ostrea weberi Olsson, 1951 (now Teskey- ostrea), from Key West (type locality) and Grassy Key. Pecten (Plagioctenium) gibbus var. amplicostatus Dall, 1898 (now regarded as a subspecies or variety of Argopecten irradians), Pliocene to Recent, ranging from west of the Mississippi River, on the Texas coast, and south to Cartagena, Colombia; no type locality nor type specimen was speci- fied. Schuchert et al. (1905) subsequently listed cotypes (USNM 154186) from Pliocene deposits of Monroe County, Florida (only part of which lies in the Florida Keys); USNM la- bel data more specifically places the material from the Caloosahatchie (sic, Caloosa- hatchee) beds of Monroe County (T. Waller, pers. comm., July 2003). Waller (1969) ex- plained the complicated history of type mate- rial for this species, selecting a lectotype from another type lot, USNM 106990, from Texas; he considered the Monroe County fossil lot (USNM 154186) as an unnecessary subse- quent designation without nomenclatural standing. Waller (1969) furthermore rejected all evidence of this species from the Pliocene CRITICAL CATALOG AND BIBLIOGRAPHY 549 of Florida [claiming all examined specimens from this locality were Р. irradians concen- tricus (Say, 1822), effectively removing this taxon from this list of those originally de- scribed from the Florida Keys. Pecten (Chlamys) imbricatus mildredae Е. M. “T.” Bayer, 1941 (now Caribachlamys), rang- ing from Biscayne Bay to Dry Tortugas and Bahamas; no type locality was originally specified. Although the author (Bayer, 1942) subsequently designated Biscayne Bay as the type locality, the holotype (USNM 598977) is from Long Key Reef, Dry Tortugas (Waller, 1993: fig. 9d, e). Pecten (Euvola) tereinus Dall, 1925 [now re- garded as а synonym of Euvola chazaliei (Dautzenberg, 1900)], from off Key West. Pitaria cordatus Schwengel, 1951 (now Pitar), from Key West. Pseudamusium strigillatum Dall, 1889b (now Palliolum), from “the Antilles and Florida Reefs”. Pseudochama inezae F. M. Bayer, 1943 (now Chama), from Carysfort Reef. Syndosmya lioica Dall, 1881 (now Abra), from various localities including Blake station 9 (111 fms) off Sand Key, station 5 (229 fms) south of the Marquesas Keys, and off Sand Key (30 fms). Boss et al. (1968: 188) subse- quently gave “20 miles W of Florida; and oth- ers” as the type locality. Tellina obliqua Wood, 1815 (non J. Sowerby, 1817) [now regarded as a synonym of Semele purpurascens (Gmelin, 1791)], had Key West designated as its type locality by Boss (1972). Tellina (Angulus) paramera Boss, 1964 (now Angulus), from various localities, including from off American Shoals (45 fms), Key West, Dry Tortugas (15 fms); type locality 1$ Miami Beach, Florida. Tellina (Angulus) probrina Boss, 1964 (now Angulus), from various localities, including from off Sombrero Key, off Sand Key, Key West, and Dry Tortugas; type locality 1$ off Fowey Light, Dade County, Florida. Teredo (Zopoteredo) clappi Bartsch, 1923, from off Key West, from timber. Transennella stimpsoni Dall, 1902, from Cape Hatteras, Egmont Key, and south to Key West. Boss et al. (1968) subsequently gave Egmont Key (at the mouth of Tampa Bay, Hillsborough County, western Florida) as the type locality, based on USNM 54100. Venus purpurascens Gmelin, 1791 (now Semele), had Key West designated as its type locality by Boss (1972). Names listed in the annotations are as used in the referenced work. Abbreviations used include: ANSP, Academy of Natural Sciences of Philadelphia; DT, Dry Tortugas; *, “gray” lit- erature; LFK, Lower Florida Keys; MCZ, Mu- seum of Comparative Zoology; MFK, Middle Florida Keys; UFK, Upper Florida Keys; USNM, National Museum of Natural History [United States National Museum]; *, literature included in earlier analysis (Mikkelsen & Bieler, 2000); |, “Florida Keys” literature or taxa ex- cluded from this analysis (for geographical reasons) or former taxonomic names with ref- erence to epithet in present use. ANNOTATED BIBLIOGRAPHY OF WORKS CONTAINING REFERENCES TO FLORIDA KEYS BIVALVES ABBOTT, К. T., 1954, American seashells. Van Nostrand, New York. xiv + 541 pp., 40 pls. With distributions including Florida Keys (or specific localities): Aequipecten (Plagioctenium) gibbus nucleus, A. lineolaris, A. phrygius, Chione (Timoclea) grus, C. (Lirophora) paphia, Cumingia coarctata, Isognomon bicolor, Noetia (Eontia) ponde- rosa, Nucula crenulata, Pitar (Pitarenus) cordata, Pseudocyrena floridana, Strigilla pisiformis, Tellina (Eurytellina) angulosa, T. (Scissula) candeana, T. (Eurytellina) punicea, Trachycardium magnum. ABBOTT, R. T., 1958, The marine mollusks of Grand Cayman Island, British West Indies. Monographs of the Academy of Natural Sci- ences of Philadelphia, 11: 138 pp., 5 pls. [sec- ond printing, October 1967, also contains 16 unnumbered pages of common names] Specifically occurring in the Florida Keys: Americardia guppyi, Barbatia tenera, Cumingia coarctata, Trachycardium magnum. ABBOTT, R. T., 1961, How to know the Ameri- can marine shells, rev. ed. A Signet Book, New American Library, New York. 222 pp., 12 color pls. With distributions including Florida Keys (or specific localities): Antigona listeri, Pseudocyrena floridana, Trachycardium magnum. An index list for the common spe- cies for “Miami to Key West” also includes Noetia ponderosa, Macrocallista nimbosa [Mytilus californianus and Macoma calcarea are also included here, obviously incorrectly]. ABBOTT, БК. T., 1968, Seashells of North America: a guide to field identification. Golden Press, New York. 280 pp. 550 MIKKELSEN & BIELER Map of North America (p. 35) showing fau- nal provinces includes Florida Keys in Car- ibbean Province; text (p. 37) refers specifically only to Lower Keys. With distri- butions including Florida Keys: Noetia pon- derosa, Strigilla pisiformis, Tellina candeana. ABBOTT,R.T., 1970, How to know the Атеп- can marine shells, rev. ed. А Signet Book, New American Library, Bergenfield, New Jersey. 222 pp., 12 color pls. Specifically occurring in the Florida Keys: Antigona listeri, Pseudocyrena floridana. This book also has an interesting zoogeo- graphic section, listing “Shells by Localities”. Also, with distributions including Key West: Macrocallista nimbosa, Noetia ponderosa (also indicated as occurring in this region, obviously in error, are Mytilus californianus and the northern Масота calcarea). *АВВОТТ, К. T., 1974, American seashells: the marine Mollusca of the Atlantic and Pacific coasts of North America, 2" ed. Van Nostrand Reinhold, New York. 663 pp., 24 pls. With distributions including Florida Keys (or specific localities): Aequipecten phrygium, Americardia guppyi, Argopecten irradians, Asthenothaerus balesi, Chama sarda, Chione (Chione) pubera, Codakia (Ctena) pectinella, Isognomon bicolor, Lima albicoma, Lucina (Lucinisca) muricata, Myrtea (Eulopia) sagrinata, Nuculana pusio, N. verrilliana, Ostrea weberi, Periploma anguliferum, P. tenerum, Pitar (Pitarenus) cordatus, Plectodon granulatus, Pteria longisquamosa, P. vitrea, Strigilla (Strigilla) gabbi, Tellina (Angulus) versicolor, Thracia corbuloides, Trachycardium (Acrosterigma) magnum, Transennella cubaniana, Ventricolaria rigida. АВВОТТ, К. Т. & Р. А. MORRIS, 1995, A field guide to shells, Atlantic and Gulf coasts and the West Indies. Peterson Field Guide 3. Houghton Mifflin Company, Boston & New York. 350 pp., 74 pls. With distributions including Florida Keys (or specific localities): Aequipecten acanthodes, Periploma anguliferum, Pitar cordatus, Trachycardium magnum. AGUAYO, С. С. & М. |. JAUME, 1947a, Pelecypoda - Thracidae (sic). Catalogo Moluscos de Cuba, no. 126, 1 p. Asthenothaerus (Asthenothaerus) balesi with Missouri Key as type locality. AGUAYO, С. С. & М. |. JAUME, 1947b, Pelecypoda — Arcidae. Catalogo Moluscos de Cuba, no. 143, 1 p. Arca (Barbatia) balesi with Missouri Key as type locality. AGUAYO, C. G. 8 M. L. JAUME, 1948a, Pelecypoda — Semelidae. Catalogo Moluscos de Cuba, no. 336, 1 p. Cumingia vanhyningi with Lower Matecumbe Key as type locality. *AGUAYO, С. С. & М. L. JAUME, 1948b, Pelecypoda — Veneridae. Catalogo Moluscos de Cuba, no. 525, 1 p. Transennella culebrana from Key West, deep water. AGUAYO, С. С. & М. L. JAUME, 1948с, Pelecypoda — Veneridae. Catalogo Moluscos de Cuba, no. 526, 1 p. Tivela mactroides from “Cayos de la Florida”. *AGUAYO, С. С. & М. L. JAUME, 1949a, Pelecypoda - Erycinidae. Catalogo Moluscos de Cuba, no. 567, 1 p. Lasaea rubra from Tortugas. AGUAYO, С. С. & М. |. JAUME, 1949b, Pelecypoda — Lucinidae. Catalogo Moluscos de Cuba, no. 564, 1 p. Ctena orbiculata forma recurvata from “Cayos de la Florida”. AGUAYO, С. С. & М. L. JAUME, 1949с, Pelecypoda — Veneridae. Catalogo Moluscos de Cuba, no. 566, 1 p. Antigona strigillina from Key West. AGUAYO, C. G. & M. L. JAUME, 1949d, Pelecypoda — Periplomatidae. Catalogo Moluscos de Cuba, no. 580, 1 р. Cochlodesma pyramidatum from “Cayos de la Florida”. AGUAYO, С. С. & М. L. JAUME, 1949e, Pelecypoda — Lucinidae. Catalogo Moluscos de Cuba, no. 587, 1 p. Phacoides muricatus from “Cayos de la Florida”. AGUAYO, C. G. & M. L. JAUME, 1950a, Pelecypoda — Corbulidae. Catalogo Moluscos de Cuba, no. 664, 1 p. Corbula cymella with “Gordon Key” as type locality. AGUAYO, С. С. & М. L. JAUME, 1950b, Pelecypoda — Chamidae. Catalogo Moluscos de Cuba, no. 602, 1 p. Pseudochama inezae with Carysfort Reef as type locality. AGUAYO, С. С. & М. L. JAUME, 1950с, Pelecypoda — Condylocardiidae. Catalogo Moluscos de Cuba, no. 604, 1 p. Condylocardia floridensis with Ohio Key as type locality. AGUAYO, С. С. & М. L. JAUME, 19509, Pelecypoda — Cuspidariidae. Catalogo Moluscos de Cuba, no. 606, 1 p. CRITICAL CATALOG AND BIBLIOGRAPHY Sal Leiomya granulata forma velvetina from “Cayos de la Florida”. AGUAYO, C. G. & M. L. JAUME, 1950e, Pelecypoda - Thraciidae. Catalogo Moluscos de Cuba, no. 622, 1 р. Thracia stimpsoni from “Cayos de la Florida”. AGUAYO, С. С. & M. L. JAUME, 1950f, Pelecypoda — Cuspidariidae. Catalogo Moluscos de Cuba, no. 626, 1 p. Cuspidaria rostrata from Sand Key. AGUAYO, С. С. & М. L. JAUME, 19509, Pelecypoda - Teredidae. Catalogo Moluscos de Cuba, no. 648, 1 p. Teredo clappi with Key West as type locality. AGUAYO, С. С. & М. L. JAUME, 1950h, Pelecypoda — Pectinidae. Catalogo Moluscos de Cuba, no. 649, 1 p. Pecten tereinus from Key West. ALDRICH, В. 8 E. SNYDER, 1936, Florida sea shells. Houghton Mifflin Company, Boston 4 New York, and The Riverside Press, Cam- bridge, Massachusetts. ix + 126 pp., 11 pls. With distributions including Florida Keys (or specific localities): Arca transversa, Lithophaga bisulcata, Lucina floridana, Tellina radiata. "ALLEN, D. M., 1979, Biological aspects of the calico scallop, Argopecten gibbus, deter- mined by spat monitoring. The Nautilus, 94(4): 107-119. Key West listed among commercial fishing grounds for the calico scallop. Florida Keys populations postulated as source of larvae recruited to major commercial beds off Cape Canaveral. ANDREWS, J., 1971, Sea shells of the Texas Coast. University of Texas Press, Austin 8 London. xvii + 298 pp. With distributions including Florida Keys (or specific localities): Aequipecten muscosus, Chione grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Polycyrena floridana, Noetia (Eontia) ponderosa, Rocellaria hians. ANDREWS, J., 1977, Shells and shores of Texas. University of Texas Press, Austin 8 London. xx + 365 pp. With distributions including Florida Keys (or specific localities): Aequipecten muscosus, Chione (Timoclea) grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Noetia (Eontia) ponderosa, Polymesoda (Pseudocyrena) maritima. ANDREWS, J., 1981a, A field guide to Texas shells. University of Texas Press, Austin & London. xxvi + 175 pp. With distributions including Florida Keys (or specific localities): Chione (Timoclea) grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Noetia (Eontia) ponderosa, Polymesoda (Pseudo- cyrena) maritima. ANDREWS, J., 1981b, Texas shells: a field guide. University of Texas Press, Austin. xxvi + 105 pp: With distributions including Florida Keys (ог specific localities): Chione (Timoclea) grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Noetia (Eontia) ponderosa, Polymesoda (Pseudo- cyrena) maritima. ANDREWS, J., 1992, A field guide to shells of the Texas coast. Gulf Publishing Company, Houston, Texas. xxiv + 176 pp. With distributions including Florida Keys (or specific localities): Chione (Timoclea) grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Noetia (Eontia) ponderosa, Polymesoda (Pseudo- cyrena) maritima. ANDREWS, J., 1994, A field guide to shells of the Florida coast. Gulf Publishing Company, Houston, Texas. xxiii + 182 pp. With distributions including Florida Keys (or specific localities): Aequipecten muscosus, Arca (Arca) zebra, Chione (Timoclea) grus, Isognomon bicolor, Lioberis castaneus, Lyropecten (Nodipecten) nodosus, Noetia (Eontia) ponderosa, Polymesoda (Pseudo- cyrena) maritima, Semele bellastriata. SANTONIUS, A., А. H. WEINER, J. С. HALAS & E. DAVIDSON, 1978, Looe Key Reef re- source inventory. Florida Reef Foundation, Homestead, Florida. [6 +] 63 pp. + unpag. figures. Results of a biological inventory of Looe Key, in summer-fall 1977, by Florida Reef Foun- dation, in support of the proposal of Looe Key as a National Marine Sanctuary. Data collected by visual transect-diving, with some samples taken for subsequent identification. The molluscan species list includes: Acropsis (sic) adamsi, Anadara notabilis, Arca imbricata, Arcopagia fausta, Atrina rigida (one of the most commonly observed spe- cies on reef flat), Barbatia cancellaria, B. candida, Brachiodontes (sic) exustus, Chama congregata, C. florida, C. sinosa (sic), Chione intapurpurea, Chlamys imbricata, C. sentis, Codakia orbicularis, Corbula swiftiana, Isognomon radiatus, Lima pellucida, L. scabra, Lioberus castaneus, Lithophaga aristata, L. bisulcata, L. nigra, Modiolus americanus, Phacoides pectinatus, 552 Pinctada radiata, Plicatula spondyloidea, Pseudochama radians, Tellina laevigata, Trachycardium isocardia. Includes a descrip- tion of the origin of the Florida Keys, their geology, and a physical description of Looe Key. CARTMAN, L. P., JR., 1974, Key West giant тар: guide to shells. Privately published, 1 map. Figured along with a map of Key West: Spondylus americanus. BALES, B. R., 1940, The rock dwellers of the Florida Keys. The Nautilus, 54(2): 39—42. With occurrence in Florida Keys, and char- acteristics of burrows: Botula fusca, Lithophaga antillarum, L. bisulcata, L. nigra, Petricola lapicida, Rocellaria ovata, Rupellaria typica, Spengleria rostrata. A brief summary (without mention of species names) of Bales’ presentation “The Rock Dwellers of the Florida Keys” can be found оп р. 5 of: H.R. ROBERTSON, 1940. Tenth An- nual Meeting of The American Malacologi- cal Union 1940. [Report of the] Tenth Annual Meeting, Academy of Natural Sciences, Philadelphia, Pennsylvania, June 17-21, 1940: 1-11. SBALES, B. R., 1944, Florida rock dwellers. Shell Notes, 1(7): 47-48. During several winters spent in shell collect- ing along the Florida Keys, that author has taken alive: Lithophaga antillarum, L. bisulcata, L. nigra, L. aristata, Botula fusca, Gastrochaena ovata, С. cueniformis (sic), Rupellaria typica, Petricola lapicida, Coralliophaga coralliophaga, Fundella candeana. B. fusca, Gastrochaena rostrata, С. coralliophaga are called “decidedly rarer” than others. F. candeana is found at only one [unspecified] locality. F. Lyman (ed.) notes immediately following the article that “Botula Fusca (sic) Gmelin, has long been consid- ered the finest shells to be obtained from the Florida Keys”. 5*ВАКЕ!ЕТО, B., 1990, Cognate bivalve spe- cies of the western Atlantic and eastern Pa- cific Oceans. American Conchologist, 18(2): 20-22. Papyridea soleniformis figured from Key West. BARRETT, R. & D. PATTERSON, eds., with technical assistance by the Shell Factory, Ft. Myers, Florida, 1967, Shells and shelling. Post Publications, Miami. 64 pp. Figured in color, with distributions including Florida Keys: Trachycardium magnum, Aequipecten lineolaris, Tivela mactroides, Chione paphia. MIKKELSEN & BIELER BARTSCH, P., 1923, Additions to our knowl- edge of shipworms. Proceedings of the Bio- logical Society of Washington, 36: 95-102. Teredo (Zopoteredo) clappi n. sp., with the type from “a piece of oak timber, probably an old ship’s keel ... at Key West”. SBARTSCH, P., 1937, An ecological cross- section of the lower part of Florida based largely upon its molluscan fauna. Pp. 11-25, in: Report of the Committee on Paleontology 1936-1937, Washington Research Council, Division of Geology and Geography. Also available at http://www.aoml.noaa.gov/ general/lib/cedardoc.html; last accessed 4 April 2003. Characterizes ecological units from the Ev- erglades to plankton. Bartsch “purposely avoided mentioning the numerous minute species ... selected those mollusks which force their attention upon the observer in each habitat ... represent the striking ele- ment of each association”. From mangrove fringe of the Keys on the bay side: Melina alata, Ostrea floridensis. From shallow-wa- ter sandy stretches beyond the Keys: Donax variabilis, Dosinia discus, Laevicardium топот, Strigilla flexuosa, Tellina alternata. From shallow water (hard pan) beyond the Keys: Chama sarda, Glycymeris americana. From shallow water (mud flats) beyond the Keys: Cardita floridana, Tagelus gibbus. From coral reef walls (Miami to Key West, past the Marquesas and Dry Tortugas): Chama congregata, Lima scabra. From sandy stretches between the coral walls: Anodontia alba, Dosinia elegans. From the continental shelf, 5-100 fms: Tellina lintea. From Pourtales Plateau, 90-300 fms: Arca glomerula, Euciroa elegantissima, Modiola polita, Protocardia peramabilis. Also includes a listing from the Florida Straits. “BAYER, [F. M.] “Т.”, 1941, Notes on Florida Mollusca, with descriptions of two new vari- eties. The Nautilus, 55(2): 43—46, pl. 3 (in part). Pecten (Chlamys) imbricatus mildredae nov. var., with distribution to the Dry Tortugas. *BAYER, F. M., 1942, The type locality and dates of Pecten imbricatus mildredae Bayer. The Nautilus, 55(3): 106. Supplement to original description lists Sand Key, Carysfort Reef, and Dry Tortugas. [Biscayne Bay is designated the type local- ity.] *BAYER, Е. M., 1943, The Florida species of the family Chamidae. The Nautilus, 56(4): 116-124, pls. 12-15. CRITICAL CATALOG AND BIBLIOGRAPHY 553 Chama florida, С. macerophylla, С. sarda, С. sinuosa bermudensis, С. $. firma, Pseudo- chama inezae n. sp. BEAUPERTHUY, I., 1967, Los mitilidos de Venezuela (Mollusca: Bivalvia). Boletin del Instituto Oceanografico de la Universidad de Oriente, Cumana, 6(1): 7-115. Original material of Modiolus squamosus п. sp. includes MCZ specimens from Lower Florida Keys. S*BENDER, J. F., 1965, Vacation shelling 1965. Texas Conchologist, 2(3): 6. Collected from Pigeon Key to Key West: Arca spp., Arcopagia fausta, Codakia orbicularis. S*BENDER, J. F., 1968, Shelling in Florida — 1966. Keppel Bay Tidings, 6(4): 1, 4, 7; 7(1): 3-5. Describes shelling at Bahia Honda, Little Duck Key, Boca Chica, Walkers Island, Shell Island, Pigeon Key, and Key West. Species included are: Arcopagia fausta, Argopecten nucleus, Caribachlamys sentis, Rangia flexuosa, Spondylus americanus. BENSON, A. J., D. C. MARELLI, M. E. FRISCHER, J. М. DANFORTH & J. D. WIL- LIAMS, 2001, Establishment of the green mussel, Perna viridis (Linnaeus, 1758) (Mol- lusca: Mytilidae) on the west coast of Florida. Journal of Shellfish Research, 20(1): 21-29. Perna viridis is predicted to invade the Florida Keys from its introduction point in Tampa Bay. BIBER Re, |. KAPPNER & P. М. MIKKELSEN, 2004, Periglypta listeri (Gray, 1838) (Bivalvia: Veneridae) in the western Atlantic: taxonomy, anatomy, life habits, and distribution. In: R. BIELER & P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the International Marine Bi- valve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 427—458. Periglypta listeri from Florida Keys localities [material from this study]. BIELSA, L. M. & R. F. LABISKY, 1987, Food habits of blueline tilefish Caulolatilus microps, and snowy grouper, Epinephelus niveatus, from the Lower Florida Keys. Northeast Gulf Science, 9(2): 77-87. Gut contents of blueline tilefish (Caulolatilus microps) taken off the Lower Keys included: Laevicardium sp., Nuculana sp. BIGATTI, G., M. PEHARDA & J. TAYLOR, 2004, Size at first maturity, oocyte envelopes and external morphology of sperm in three species of Lucinidae (Mollusca: Bivalvia) from Florida Keys, U.S.A. In: В. BIELER & P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the Interna- tional Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 417- 426. Codakia orbicularis, Ctena orbiculata, Lucina pensylvanica [material from this study]. SBIPPUS, А. C., JR., 1950, Real shells — real thrills — real fun. Shell Notes, 2(1012): 166- 173: Collecting in the Upper Keys by dip net, rock turner, etc., with Frank Lyman aboard the Junonia. From the outer reefs off Key Largo, 26 June 1950: Pecten imbricatus. From a reef off Tavernier, 29 June 1950: Pecten sentis. A partial species list also includes Arca barbata, A. umbonata, Chione cancellata, Donax sp., Laevicardium mortoni, L. serratum, Pteria radiata (variety?). S*BOONE, С., 1986, Search and seizure: Dendostrea frons found on mangrove roots. Texas Conchologist, 22(2): 41-45. From off Marathon: Dendostrea frons, Spondylus sp. “BOSS, К. J., 1964, New species of Tellina from the western Atlantic. Occasional Papers on Mollusks, Department of Mollusks, Mu- seum of Comparative Zoology, Harvard Uni- versity, 2(29): 309-324, pls. 54, 55. Tellina paramera and T. probrina n. spp. from Florida Keys localities. *BOSS, К. J., 1966, The subfamily Tellininae in the western Atlantic. The genus Tellina (part I). Johnsonia, 4(45): 217-272. Museum lots (ANSP, MCZ, USNM) from the Florida Keys cited for: Tellina (Merisca) aequistriata, T. (Elliptotellina) americana, T. (Arcopagia) fausta, T. (Acorylus) gouldii, T. (Laciolina) laevigata, T. (Tellinella) listeri, T. (Laciolina) magna, Т. (Merisca) martinicensis, Т. (T.) radiata, T. (Phyllodina) squamifera. SBOSS, K. J., 1967, Evolutionary sequence in Phyllodina (Bivalvia: Tellinidae) [abstract]. American Malacological Union, Annual Re- ports for 1966, 33: 21-23. Tellina (Phyllodina) squamifera from the Florida Keys. SBOSS, K. J., 1968a, On the evolution of Spengleria (Gastrochaenidae: Bivalvia) [ab- stract]. American Malacological Union, An- nual Reports for 1967, 34: 1517. Spengleria rostrata with its main distribution in the Florida Keys. *BOSS, К. J., 1968b, The subfamily Tellininae inthe western Atlantic. The genera Tellina (part II) and Tellidora. Johnsonia, 4(46): 273-344. Museum lots (ANSP, MCZ, USNM) and pri- vate collection records (ex Schmidt) from the 554 MIKKELSEN & BIELER Florida Keys cited for: Tellidora cristata, Tellina (Eurytellina) alternata, T. (Eurytellina) angulosa, Т. (Scissula) candeana, T. (Scissula) consobrina, T. (Scissula) iris, Т. (Angulus) mera, T. (Eurytellina) nitens, T. (An- gulus) probrina, T. (Scissula) similis, T. (An- gulus) sybaritica, T. (Angulus) tampaensis, T. (Angulus) texana, T. (Angulus) versicolor. *BOSS, К. J., 1969, The subfamily Tellininae in the western Atlantic. The genus Strigilla. Johnsonia, 4(47): 345-366. Museum lots (ANSP, MCZ, USNM) from the Florida Keys cited for: Strigilla carnaria, S. gabbi, S. mirabilis. BOSS, K. J., 1972, The genus Semele in the western Atlantic (Semelidae; Bivalvia). Johnsonia, 5(49): 1-32. Material examined includes lots from Florida Keys localities for Semele bellestriata (sic), Semele (Semelina) nuculoides, Semele proficua, and Semele purpurascens. The last species has Key West designated as its type locality (as well as for those of its synonyms obliqua Wood, variegata Lamarck, and /aeta Reeve). *BOSS, К. J. & А. S. MERRILL, 1965, The family Pandoridae in the western Atlantic. Johnsonia, 4(44): 181-216. Museum lots (MCZ, USNM) from the Florida Keys cited for: Pandora bushiana, P. inflata. *BOSS, К. J. 4 D. К. MOORE, 1967, Notes on Malleus (Parimalleus) candeanus (d'Orbigny) (Mollusca: Bivalvia). Bulletin of Marine Science, 17(1): 85-94. Malleus candeanus listed and figured from several Florida Keys localities. BOSS AA J ROSEWATER -& EA: RUHOFF, 1968, The zoological taxa of Wil- liam Healey Dall. United States National Museum Bulletin, no. 287, 427 pp. In a catalog of Dall's taxa, from type locali- ties in the Florida Keys: Corbula cymella, Thracia stimpsoni, Cytherea (Ventricola) strigillina, Pecten (Euvola) tereinus. “BOSS, К. J. & М. Е. WASS, 1970, Northward range extension of Cyclinella tenuis Recluz. The Nautilus, 83(3): 112-113. Localities cited from Middle, Lower Keys and Dry Tortugas. BREWSTER-WINGARD, С. Ц. 4 $. E. ISHMAN, 1999, Historical trends in salinity and substrate in central Florida Bay: a pa- leoecological reconstruction using modern analogue data. Estuaries, 22(2B): 369-383. Summary of data on Florida Bay mollusks from earlier reports by Wingard et al. (1995) and Brewster-Wingard et al. (1996, 1997). Living taxa (from Brewster-Wingard et al., 1996) are Anomalocardia sp., Arcopsis adamsi, Brachiodontes (sic) sp., Chione cancellata, Laevicardium spp., Nucula proxima, Parastarte triquetra, Pinctada ra- diata, Polymesoda sp., Transennella spp. Core 19B taxa (from Brewster-Wingard et al., 1997) are: Brachiodontes (sic) sp., Chione cancellata, Transennella sp. Core 6A taxa (from Wingard et al., 1995) are: Transennella spp., Brachiodontes (sic) sp. SBREWSTER-WINGARD, G. L., S. E. ISHMAN, L. E. EDWARDS & D. A. WILLARD, 1996, Preliminary report on the distribution of mod- ern fauna and flora at selected sites in north- central and north-eastern Florida Bay. United States Geological Survey Open-File Report 96-732, 34 pp. Electronic version available at http://pubs.usgs.gov/pdf/of/ofr96732.html; last accessed 12 September 2003. Of the 14 numbered stations sampled, 9 are within our definition of the Florida Keys [ex- cluding the approximate northern half of Florida Bay, i.e., stations 1, 2, 3, 8, and 9]. Living mollusks were collected in February and July of 1995 using push cores, and abundances are compared among stations and between the two collections. For the entire study, Transenella (sic) spp. and Brachiodontes (sic) sp. are considered “ubiq- uitous” taxa; Anomalocardia sp. and Parastarte triquetra are present in significant numbers. Cyrenoida floridana, Polymesoda sp., and Mytilopsis leucophaeata are among indicator species for oligohaline-mesohaline conditions. No obvious seasonal trends were observed from the overall molluscan data. Anomalocardia sp., Chione cancellata, and Lima sp. showed seasonal differences, per- haps related to “seasonal spawning”. Mol- luscan data are presented for stations 11-13 only (these correspond to Turney & Perkins’ (1972) “interior subenvironment”): Anomalocardia sp., Arcopsis adamsi, Brachiodontes (sic) sp., Chione cancellata, Codakia sp., Cumingia tellinoidea (sic), Laevicardium spp., Lima sp., Nucula proxima, Parastarte triquetra, Pinctada ra- diata, Polymesoda sp., Tellina spp., Transenella (sic) spp., rare pelecypods, uni- dentified pelecypod fragments. A summary of these data was published by Brewster- Wingard & Ishman (1999). BREWSTER-WINGARD, С. L., $. Е. ISHMAN & C. W. HOLMES, 1998, Environmental im- pacts on the southern Florida coastal wa- ters: a history of change in Florida Bay. CRITICAL CATALOG AND BIBLIOGRAPHY 955 Journal of Coastal Research, Special issue 26: 162-172. Summary of data on Florida Bay mollusks from earlier reports by Wingard et al. (1995) and Brewster-Wingard et al. (1997), derived from living communities and death assem- blages in sediment cores (from cores 6A and 19B). From Bob Allen Core 6A: Brachiodontes (sic) sp. From Russell Bank Core 19B: Brachiodontes (sic) sp. Modern studies (living mollusks) at monitoring sites indicate probable habitats through time in the sediment cores. Anomalocardia sp. is rela- tively abundant in mesohaline-polyhaline sites. Pinctada radiata, Transennella spp., Laevicardium sp., and Chione cancellata prefer polyhaline to euhaline conditions. Brachiodontes (sic) sp. is predominantly found on macroalgae associated with Thalssia grassbeds. Pinctada radiata is found on both Thalassia and macroalgal mats. IBREWSTER-WINGARD, С. L., S. E. ISHMAN, М. J. WAIBEL, D. A. WILLARD, L. E. EDWARDS & С. W. HOLMES, 1998, Pre- liminary paleontologic report on Core 37, from Pass Key, Everglades National Park, Florida Bay. United States Geological Sur- vey Open-File Report 98-122, 22 pp. Elec- tronic version available at http:// pubs.usgs.gov/pdf/of/ofr98122.html; last accessed 12 September 2003. This work 1$ here excluded as outside our defined area, which extends roughly half-way between the Florida Keys island chain and the tip of peninsular Florida. Pass Key 1$ north of this limit. SBREWSTER-WINGARD, С. L., 5. E. ISHMAN, D. A. WILLARD, L. E. EDWARDS 8 C. W. HOLMES, 1997, Preliminary pale- ontologic report on Cores 19A and 19B, from Russell Bank, Everglades National Park, Florida Bay. United States Geological Sur- vey Open-File Report 97-460, 29 pp. Elec- tronic version available at http:// 131.247.143.93/publications/ofr/97-460/; last accessed 12 September 2003. Study of core samples from Russell Bank, Florida Bay, dating to 115-118 yrs at the core bottom, included shells of Anomalocardia sp., Arcopsis adamsi, Brachiodontes (sic) sp., Chione cancellata, Cumingia tellinoidea (sic), Laevicardium spp., Nucula proxima, pectinid, Pinctada radiata, Tellina spp., Transenella (sic) spp., rare pelecypods, unidentified pelecypod fragments. A summary of these data was pub- lished by Brewster-Wingard et al. (1998) and Brewster-Wingard & Ishman (1999). BREWSTER-WINGARD, С. L., J. К. STONE 8 C. W. HOLMES, 2001, Molluscan faunal distribution in Florida Bay, past and present: an integration of down-core and modern data. Pp. 199-231, in: B. R. WARDLOW, ed., Paleo- ecological Studies of South Florida, Bulletins of American Paleontology, no. 361. See full data matrix at http://flaecohist/database/ Reference/synthesis; and core data at http:// sofia.usgs.gov/flaecohist. A summary and update of living and dead molluscan assemblages in Florida Bay (pre- viously published in part by Brewster- Wingard et al. (1996, 1997), Brewster-Wingard & Ishman (1999), and Wingard et al. (1995); taxonomy is much updated, acknowledging W. G. Lyons. Analysis of core versus living data suggests changes in Florida Bay ma- rine environments during the past 200 years. Most mollusks are generally presentthrough- out the study period, however the study documents fluctuations in dominance and diversity. Modern assemblages are defined and include: Brachidontes assemblage (most dominant), Pteria assemblage (on sides of mudbanks in dense Thalassia), and three “western” assemblages (near chan- nels, indicative of euhaline conditions). Con- sidered as important biological indicators of conditions in Florida Bay during mandated restoration efforts: Anomalocardia auberiana, Brachidontes exustus, Pteria longisquamosa. Brachidontes exustus is the dominant mollusk in Florida Bay, due to its tolerance of varying salinity and substrate, and poor water quality. Live-mollusk data previously presented by Brewster-Wingard et al. (1996) include Anomalocardia auberiana, Arcopsis adamsi, Brachidontes exustus, Chione cancellata, Laevicardium mortoni, Parastarte triquetra, Репа longisquamosa, tellinid, Transennella sp. Push core data previously published by Brewster-Wingard et al. (1997), Brewster- Wingard & Ishman (1999), and Wingard et al. (1995): include Anomalocardia auberiana, Arcopsis adamsi, Argopecten irradians, Brachidontes exustus, Chione cancellata, Codakia spp., Cumingia tellinoides, Laevicardium mortoni, Limaria cf. pellucida, Lucinisca nassula, Mysella planulata, Nucula proxima, Parastarte triquetra, Parvilucina multilineata, Pitar simpsoni, Pteria longisquamosa, Semele bellastriata, Tellina spp., Transennella sp. SBRITTON, J. C., 1970, The Lucinidae (Mol- lusca: Bivalvia) of the western Atlantic 556 Ocean. Ph.D. dissertation, George Washing- ton University, Washington, ОС. v + 567 pp. including 23 pls. Recorded in materials examined sections from Florida Keys localities: Anodontia (An- odontia) alba, A. (Anodontia) schrammi, Callucina (Callucina) radians, Cavilinga blanda, Codakia (Codakia) orbicularis, C. (Ctena) orbiculata, Divaricella (Divaricella) dentata, D. (Divalinga) quadrisulcata, Lucina (Pleurolucina) leucocyma, L. (Lucina) pensylvanica, L. (Pleurolucina) sombrerensis, Megaxinus floridanus, Parvilucina (Bellucina) amiantus, P. (Parvilucina) costata, P. (Parvilucina) multilineata, P. (Lucinisca) nassula, P. (Parvilucina) pectinella, Phacoides (Lucinoma) filosus, P. (Phacoides) pectinatus. SBROOKS, J., 1968a, Keys after “Abby”. Seafari [Palm Beach County Shell Club Newsletter], 10(7): 8. Results from collecting in the Middle Keys after Hurricane Abby passed between the Dry Tortugas and Key West in early June 1968 include Antigona listeri, Arcopagia fausta, Lima lima, Lithophaga antillarum, Tellina similis, tellins, many other dead bivalves. Continued by Brooks (1968b). SBROOKS, J., 1968b, Further report on Mara- thon and vicinity in June 1968. Seafari [Palm Beach County Shell Club Newsletter], 10(10): 8. Continuing notes from Brooks (1968a). Col- lected from Marathon and _ vicinity: Americardium media, Chama spp., Chlamys sentis, Codakia orbicularis, Glycymeris pectinata, Laevicardium laevigatum, Lima pellucida, Lima scabra form tenera, Lucina pensylvanica, Papyridea soleniformis, Tellina fausta, T. similis, several other tellins, Trachycardium egmontianum, T. muricatum. SBROOKS, J., 1969, The Keys in August. Seafari [Palm Beach County Shell Club Newsletter], 11(11): 7. Collected in early August [1969] from the Marathon area: Botula fusca, Chlamys sentis. SBURGGRAF, P., 1969, Broward member in the Keys. Seafari [Palm Beach County Shell Club Newsletter], 11(10): 6-7. From Lower Keys localities, June 28-29 [1969]: Chlamys sentis, Glycymeris pectinata, Tellina radiata. CALKINS, W. W., 1878, Catalogue of the ma- rine shells of Florida, with notes and descrip- tions of several new species. Proceedings of the Davenport Academy of Natural Sci- entes, 2: 232-252 pl #8! MIKKELSEN & BIELER “Тре material for the following monograph has been mainly derived from my own col- lections, and observations made during two winters spent in Florida in 1875 and 1877. The first time as member of an expedition in the interest of the Chicago Academy of Sci- ences, and in 1877 on my own account. In addition to my personal collections, | have received since my return valuable acces- sions from ту collectors living in Florida. In the determination of species | have been assisted in many instances by Mr. George W. Tryon, Jr., of Philadelphia, to whom, and also to Mr. Thomas Bland, | desire to express ту acknowledgments for kindly aid. [...] The largest part of the species enumerated are in my cabinet. A number are in the Museum of the Chicago Academy and in that of the Davenport Academy of Sciences. Other spe- cies will be deposited from time to time.” [pp. 232-233]. From Florida Keys localities are: Avicula atlantica, A. sp., Chama arcinella, C. macrophylla, Cytherea dione, Lima scabra, L. squamosa, Lithodomus antillarum, L. lithophagus, Modiola sulcata, M. tulipa, M. t. var. nigra п. var., Pectunculus pennaceus, Perna perna, Sanguinolaria sanguinolenta, Spondylus gaederopus, Strigilla flexuosa, S. pisiformis, Tellina braziliana, T. decora, T. iris, T. radiata, T. tenera, Xylotrya fimbiata (sic). *CAMPBELL, D. C., К. J. HOEKSTRA & J. С. CARTER, 1998, 18$ Ribosomal ОМА and evolutionary relationships within the Bivalvia. Рр. 75-85, in: Р.А. JOHNSTON & J. W. HAGGART, eds., Bivalves: ап eon of evolution - paleobiological studies honoring Norman D. Newell. University of Calgary Press, Calgary, Alberta, Canada. 461 pp. Molecular sequences from Barbatia cancellaria, Pinctada imbricata, Isognomon alatus from West Summerland Key; Ostrea equestris from Marathon. CAMPBELL, М. В., ©... STEINER Eb: CAMPBELL & H. Огеуег, 2004, Recent Chamidae (Bivalvia) from the western Atlan- tic Ocean. In: В. BIELER 8 P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceed- ings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 381-415. From Florida Keys localities: Arcinella cornuta, Chama congregata, C. florida, C. inezae, C. lactuca, C. macerophylla, C. ra- dians, C. sarda, C. sinuosa [some material from this study]. CANTILLO, А. Y., С. ©. LAUENSTEIN 8 Т. P. O'CONNOR, 1997, Mollusc and sediment CRITICAL CATALOG AND BIBLIOGRAPHY 557 contaminant levels and trends in South Florida coastal waters. Marine Pollution Bulletin, 34(7): 511-521. The NOAA National Status and Trends (NS&T) Program includes one station in its Mussel Watch Project at Bahia Honda Key (24°39.52’N, 81°16.43'W, three years of data through 1995). Chama sinuosa from this site was collected for analysis of organic and inorganic contaminants, although this paper does not present the results on that species. Two other Mussel Watch Project stations in Florida Bay (Flamingo Bay and Joe Bay) are too far north to be included in this survey. CARTER, J. G., 1978, Ecology and evolution of the Gastrochaenacea (Mollusca, Bivalvia) with notes on the evolution of the endolithic habitat. Peabody Museum of Natural His- tory, Yale University, Bulletin 41, 92 pp. The “three more common gastrochaenids of the Florida Keys” are Spengleria rostrata, Gastrochaena (G.) hians, and G (Rocellaria) ovata. G. hians dominates the Keys gastrochaenid fauna in terms of population density, whereas S. rostrata is generally rare. ECHAN, E. I., 1977a, The ecology of the seagrasses of South Florida: a community profile. United States Fish and Wildlife Ser- vices, FWS/OBS - 82/25, 158 pp. Reprinted September 1985. Cited by Zieman, 1982; reporting on the ef- fects of a 1975 tanker discharge SW of the Marqueses, attributed mass mortalities ofthe pearl oyster (Pinctada radiata) a grass bed inhabitant, to some soluble fraction of petro- leum. SCHAN, Е. |., 1977b, Oil pollution and tropical littoral communities: biological effects of the 1975 Florida Keys oil spill. Pp. 539-542, in: Proceedings 1977 Oil Spill Conference (Pre- vention, Behavior, Control, Cleanup), March 8-10, 1977, New Orleans, Louisiana. Reports on impact of crude oil discharge 26 mi SSW of the Marquesas, affecting the lower Florida Keys from Little Pine Key to Boca Chica Key. Molluscan species dis- cussed: Crassostrea virginica, Isognomen (sic) alatus, Pinctada radiata (with mass mortality attributed to oil spill). S*CLAMPIT, L., 1987, Florida Keys. Texas Conchologist, 24(1): 8-11. From Missouri Key, Grassy Key, Little Duck Key, and Ohio Key in July 1987: Arcidae, Chamidae, Codakia orbicularis, Linga pensylvanica, Pinnidae, Tellinidae. S*CLAMPIT, L., 1988, Florida Keys revisited. Texas Conchologist, 25(1): 28. From Missouri Key: Chama macerophylla. CLENCH, W. J., 1942, The genera Dosinia, Macrocallista and Amiantis in the western Atlantic. Johnsonia, 1(3): 1-8. Museum lots (ANSP) from the Florida Keys cited for: Dosinia floridana. *CLENCH, W. J. 8 L. С. SMITH, 1944, The family Cardiidae in the western Atlantic. Johnsonia, 1(13): 1-32. Museum lots (MCZ) and private collections (ex Van Hyning) from the Florida Keys cited for: Laevicardium laevigatum, L. mortoni, Microcardium peramabile, Papyridea semisulcata, P. hiatus, Trachycardium egmontianum, T. magnum, T. muricatum, Trigoniocardia (Americardia) media. SCLOSE, H. T., 1974, Shelling in the Florida Keys?? Of Sea 8 Shore, 5(4): 183, 200. Account of a 1973 trip to the lower Keys in- cludes Chama macerophylla from Sand Key. COHEN, S. & К. COHEN, 1991, Florida Keys divers guide — The Upper Keys. Seapen, Key Largo & Tel Aviv. 156 pp. Includes a photo (unnamed) of a living Ctenoides floridanus (with light-orange ten- tacles). COLIN, P. L., 1978, Caribbean reef inverte- brates and plants. T. F. H. Publications, Nep- tune City, New Jersey. 512 pp. With Florida Keys specifically cited in spe- cies distribution: Spondylus americanus. CONRAD, T. A., 1866, Descriptions of new marine bivalve Mollusca. American Journal of Conchology, 2(3): 280-281, pl. 15. Dosinia floridana n. sp., from Florida Keys, Gulf of Mexico. SCROVO, М. E., 1970, Gastrochaena hians Gmelin 1791. Seafari [Palm Beach County Shell Club Newsletter], 12(11): 6-7. Study includes shells examined from the Lower Florida Keys. *DALL, W. Н., 1881, Reports on the results of dredging, under the supervision of Alexander Agassiz, in the Gulf of Mexico, and in the Caribbean Sea, 1877-79, by the United States Coast Survey steamer “Blake”, Lieu- tenant-Commander С. D. Sigsbee, U.S. М., and Commander J. К. Bartlett, U.S.N., сот- manding. XV. Preliminary report on the Mol- lusca. Bulletin of the Museum of Comparative Zoology, 9(2): 33-144. Station information in this paper is insufficient to place all localities. Decisions were made on the basis of S. Smith (1889). Eighteen 558 Blake stations are mentioned with reference to localities in or off the Florida Keys or Dry Tortugas; these are stations 5, 6, 9, 10, 11, 12, 26, 27, 28, 29, 30, 31, 43, 44, 45, 46, 70, and 72. Of these, stations 44, 45, and 46 are here excluded because they are located too far northwest into the Gulf of Mexico (north of the latitude of Cape Sable); stations 29, 30, and 31 are here excluded because they are too far west (beyond 83°30’М), which is also where the 1,000 fm isobath is located (stations 29-31 are very deep); 28 and 43 are also excluded as too deep (863 and 339 fms, respectively). All remaining sta- tions (5, 6, 9, 10, 11, 12, 26, 27, 70, 72) are less than 300 m (164 fms). “Off Sombrero, 54 fms” is in the Virgin Islands, not the Florida Keys (see Introduction). Identified by S. R. Roberts for Dall, and from the Florida Keys, are Amussium lucidum, Avicula sp., Cardium (Fulvia) peramabilis n. sp., C. (F.) p. var. tinctum n. var., Corbula cymella, C. disparilis, Gouldia cubaniana, Leda carpenteri n. sp., L. jamaicensis, Neaera alternata, М. rostrata, Poromya granulata, Syndosmya lioica п. sp., Verticordia acuticostata n. sp. DALL, W. H., 1883, On a collection of shells sent from Florida by Mr. Henry Hemphill. Proceedings of the United States National Museum, 6(21): 318-342, pl. 10. “In the absence of a good collection of named specimens from the region, it is difficult and tedious work identifying specimens соп- nected, as the South Florida shells are, with the West Indian fauna. Consequently it is with a certain diffidence that | attempted, at Mr. Hemphill's request, to work up the ex- tremely interesting collection he has given to the National Museum. The only catalogues relating to South Florida are extremely im- perfect though praiseworthy attempts. Conrad's work was never complete and 1$ antiquated; the paper of Mr. Melvill is marred by the inaccuracies of identification for which the present Mr. Sowerby is famous; Mr. Calkins’ work is the best of all, but would have been more useful if the specimens ac- tually collected by him had been discrimi- nated in some way from those quoted from other authors, whose localities or identifica- tions may not have been accurate, or at least may not have been confirmed. It is known to most persons interested that the Smithsonian collection of East American shells, especially those belonging south of New York, was in the hands of Dr. Stimpson, and with his own matchless collection was MIKKELSEN & BIELER destroyed totally by the fire at Chicago in 1871. Under these circumstances, believing it bet- ter to make some sort of start at cataloging the shells of our southern coast (even а the risk some erroneous identifications) than to wait for opportunities which not seem likely to be soon offered, the present list has been prepared in the hope that its deficiencies may stimulate others to correct and enlarge it from specimens actually obtained on the spot.” (pp. 319-320). Discussed from Key West: Anomalocardia flexuosa, Arca (Barbatia) dominguensis, A. (Barbatia) gradata, Cyrena carolinensis (sic), Lucina tigerina, Mytilus exustus, Perna ephippium, Tellina mera. The list was con- tinued by Dall (1885). DALL, W. H., 1885, List of marine Mollusca comprising the Quaternary and Recent forms from American localities between Cape Hatteras and Cape Roque including the Ber- mudas. Bulletin of the United States Geo- logical Survey, 24: 1-336. Cited from Key West (all specifically refer- ring to Dall, 1883): Anomalocardia flexuosa, Arca (Barbatia) dominguensis, A. (B.) gradata, Cyrena carolinensis (sic), Lucina tigerina, Mytilus exustus, Perna ephippium, Tellina mera. *DALL, W. H., 1886, Report on the results of dredging, under the supervision of Alexander Agassiz, in the Gulf of Mexico (1877-78) and in the Caribbean Sea (1879-80), by the U.S. Coast Survey Steamer “Blake”, Lieut.-Com- mander C. D. Sigsbee, U.S.N., and Com- mander J. R. Bartlett, U.S.N., commanding. XXIX. Report on the Mollusca. Part 1. Brachiopoda and Pelecypoda. Bulletin ofthe Museum of Comparative Zoology at Harvard College, 12: 171-318, pls. 1-9. Abra lioica, Cardium laevigatum, C. (Fulvia) peramabilis, Corbula dietziana, C. disparilis, Cuspidaria rostrata, Cytherea hebraea, Leda acuta, Lima inflata, Pecten antillarum, P. dislocatus, Petricola divaricata, Semele obliqua, Tellina tenera, Thracia corbuloidea, and Venus cancellata are included from Keys localities collected by the Blake Expedition (1877-1880) or noted from other Keys lo- calities. “Off Sombrero, 54 fms” is in the Vir- gin Islands, not the Florida Keys (see Introduction). DALL, W. H., 1889a, A preliminary catalogue of the shell-bearing marine mollusks and brachiopods of the southeastern coast of the United States, with illustrations of many of CRITICAL CATALOG AND BIBLIOGRAPHY 959 the species. Bulletin of the United States National Museum, по. 37: 1-121, 74 pls. The Florida Keys, “very intimately connected, faunally, with the northern shores of Cuba opposite, and with the Bahamas, includes the region south of Biscayne Bay on the east, and south of the southern entrance to Char- lotte Harbor on the west side of the Penin- sula, to and including the Keys and Tortugas reefs and islands” (p. 10). A table of species distributions includes the Florida Keys as a single column, plus north- ern and southern extreme distributional points. Marked as “*” [known from shores, either picked up on beach or found living between high water and 50 fms]: Anomia simplex, Arca (Byssoarca) adamsi, A. (Argina) americana, A. (Scapharca) auriculata, A. (Barbatia) barbata, A. (Arca) imbricata, A. (Scapharca) incongrua, A. (S.) lienosa, A. (Arca) noae, A. (Noetia) orbignyi, A. (N.) ponderosa, A. (Byssoarca) reticulata, A. (Scapharca) transversa, Asaphis deflorata, Astarte nana, Asthenothaerus hemphillii, Avicula nitida, Basterotia quadrata, Cardita conradii, C. floridana, Cardium antillarum, Chama arcinella, С. macerophylla, Cardium isocardia, C. mag- num, C. medium, C. muricatum, Circe (Gouldia) cerina, Corbula barrattiana, C. nasuta, Crassatella floridana, C. (Eriphyla) lunulata, Crenella divaricata, Cumingia tellinoides, Cyrena (Leptosiphon) caro- linensis (sic), Cyrenoidea floridana [nomen nudum], Cytherea albida, С. (Transennella) conradina, C. (T.) cubaniana, C. (Dione) dione, С. (Callista) gigantea, С. (Tivela) mactroides, C. (Callista) maculata, C. simpsoni, Dacrydium vitreum, Diplodonta semiaspera, D. soror, D. subglobosa, Donax denticulatus, D. fossor, D. variabilis, Dosinia elegans, Ensis americana, Ervilia concentrica, E. nitens, Gastrochaena cuneiformis, G. ovata, G. (Spengleria) rostrata, Heterodonax bimaculata, Iphigenia braziliana, Labiosa canaliculata, Lima hians, L. inflata, L. scabra, L. здиатоза, L. tenera, Lithophagus antillarum, L. bisulcatus, L. caribaeus, L. forficatus, Loripes edentula, L. e. var. chrysostoma, Lucina (Lucina) costata, L. (L.) crenulata, L. (L.) floridana, L. (L.) multilineata, L. (L.) pecten, L. (L.) pennsylvanica (sic), L. (L.) squamosa, L. (L.) tigrina, L. (L.) trisulcata, Lucinopsis tenuis, Lutricola interstriata, Lyonsia beana, Macoma brevifrons, M. cerina, M. tenta, M. t. var. souleyetiana, Mactra brasiliana, M. lateralis, М. solidissima var. similis, Margaritiphora radiata, Martesia corticaria, M. cuneiformis, M. striata, Modiola (Botula) cinnamomea, M. (Amygdalum) lignea, M. (Botulina) opifex, M. (Brachydontes) sulcata, Modiolaria lateralis, Mytilus exustus, M. hamatus, Ostrea cristata, O. frons, O. virginica, Papyridea bullata, P. petitiana, P. (Liocardium) serratum, Pecten (Pecten) antillarum, Р. (Р) exasperatus, P (P_) imbricatus, Р (Р) irradians var. dislocatus, Р (Р) nodosus, Р (Р) nucleus, Р (P) ornatus, P. (Janira) ziczac, Pectunculus pectinatus, P. undatus, Periploma angulifera, Perna ephippium, P. obliqua, Petricola pholadiformis, Р. (Choristodon) robusta, P sp., Pholas campechiensis, P. (Barnea) costata, Р (B.) truncata, Pinna carnea, P. muricata, P. seminuda, Placunanomia rudis, Plicatula ramosa, Saxicava arctica, S. azaria, Semele cancellata, $. obliqua, $. reticulata, Solenomya occidentalis, Spondylus spathuliferus, Strigilla carnaria, S. flexuosa, S. pisiformis, Tagelus divisus, Tellidora cristata, Tellina alternata, T. decora, T. fausta, T. gouldii, T. interrupta, T. lineata, T. magna, T. mera, T. modesta, T. radiata, T. squamifera, T. striata, T. tenera, T. sp., Teredo thomsoni, Thracia corbuloidea, T. phaseolina, T. stimpsoni, Venus beaui, V. cancellata, V. crispata, V. mercenaria, V. m. var. mortoni, V. pygmaea, V. (Anomalocardia) rostrata, Verticordia (Trigonulina) ornata, Xylotrya fimbriata. Marked as “dagger” [archibenthal, 50-800 fms]: Abra longicallus, Arca (Byssoarca) glomerula, A. (Macrodon) sagrinata, A. (M.) sp., Asthenothaerus (Bushia) elegans, Astarte lens, A. smithii, Cardita domingensis, Cardium peramabilis, Corbula cubaniana, C. cymella, C. dietziana, C. krebsiana, Crassatella (Eriphyla) lunulata var. parva, Cryptodon obesus, C. pyriformis, Cuspidaria (Liomya, Plectodon) granulata, С. (L., P.) 9. var. velvetina, C. (Cuspidaria) obesa, C. (Cardiomya) perrostrata, C. (Cuspidaria) rostrata, C. (Cardiomya) striata, Cytherea (Veneriglossa) vesica, C. sp., Leda (Leda) acuta, L. (L.) messanensis, L. (L.) vitrea, L. (Y.) liorhina, Lima albicoma, L. (Limatula) setifera, L. (L.) subauriculata, Limopsis antillensis, L. aurita, L. cristata, L. minuta, Loripes lens, Lucina (Lucina) filosa, L. (L.) lenticula, L. (L.) leucocyma, L. (L.) pectinella, L. (Divaricella) quadrisulcata, L. (Lucina) sagrinata, L. (L.) scabra, L. (L.) sombrerensis, Modiola (Amygdalum) polita, M. (A.) p. var. sagittata, Myonera lamellifera, Nucula 560 aegeënsis, Periploma tenera, Pandora (Kennerlia) glacialis, Pecten (Amusium, Propeamussium) cancellatum, P. (A., P.) pourtalesianum, P. (A., P.) P. var. marmo- ratum, P. (A., P.) sayanum, P. (Pecten) effluens, P. (P.) phrygium, P. (Pecten, Pseudamusium) sigsbeei, P. (P., P.) thalassinus, Poromya (Cetomya) albida, P. granulata, Venus granulata, V. lamarckii, V. pilula, V. rugosa, V. r. var. rugatina, Verticordia acuticostata, V. (Haliris) fischeriana. Marked as “* + dagger” [both shallow (known from shores, either picked up on beach or found living between high water and 50 fms) and deep (archibenthal, 50-800 fms)]: Avicula atlantica, Arca (Byssoarca) nodulosa, Leda (Leda) carpenteri, Lucina (Lucina) lintea, Chama sarda, Cardium peramabilis var. tinctum, Papyridea (Liocardium) laevigatum, Pecten (Janira) hemicyclica, Cytherea hebraea, Petricola (Naranaio) lapicida, Abra lioica, Cuspidaria (Cardiomya) costellata, Corbula swiftiana. DALL, W. H., 1889b, Report on the results of dredging, under the supervision of Alexander Agassiz, in the Gulf of Mexico (1877-78) and in the Caribbean Sea (1879-80), by the U.S. Coast Survey Steamer “Blake”, Lieut.-Com- mander C. D. Sigsbee, U.S.N., and Com- mander J. R Bartlett, U.S.N., commanding. XXIX. Report on the Mollusca. Part Il. Gas- tropoda and Scaphopoda [with “Addenda and Corrigenda to Part |, 1886”, pp. 433- 452]. Bulletin of the Museum of Compara- tive Zoology, 18: 1-492, pls. 10-40. Additions and Corrigenda to Part | (Dall, 1886) includes Pseudamusium strigillatum n. sp., from the Antilles and Florida Reefs. DALL, W. H., 1890, Scientific results of explo- rations by the U.S. Fish Commission Steamer Albatross. No. VII. — Preliminary report on the collection of Mollusca and Brachiopoda obtained in 1887-88. Proceed- ings of the United States National Museum, 12(773): 219-362, pls. 5-14 [07 March 1890]. With distribution including Florida Keys: Crassatella floridana. DALL, W. H., 1896a, The mollusks and bra- chiopods of the Bahama Expedition of the State University of lowa. Bulletin from the Laboratories of Natural History of the State University of lowa, 4(1): 12-27, pl. 1. From Florida Keys localities, some general, others from stations of the May-July 1893 S.U. 1. Bahama Biological Expedition aboard the 95-foot schooner Emily E. Johnson, led by Charles C. Nutting: Arca noae, A. MIKKELSEN 8 BIELER umbonata, Avicula atlantica, A. crocata, A. radiata, Cardium medium, C. serratum, Chama (Echinochama) arcinella, Lithophagus antillarum, Macoma sp., Pecten ornatus, Perna oblique (sic), Pinna pernula, Tagelus divisus, Tellina sp., Venus (Chione) cancellata, Venus sp. A full narrative of the expedition and station data were published by Nutting (1895). DALL, W. H., 1896b, On the American spe- cies of Ervilia. The Nautilus, 10(3): 25-27. With distributions including Florida Keys: Ervilia concentrica, E. nitens. DALL, W. H., 1897, Synopsis of the Pinnidae of the United States and West Indies. The Nautilus, 11(3): 25-26. With distribution including Florida Keys: Pinna carnea. DALL, W. H., 1898, Contributions to the Ter- tiary fauna of Florida, with especial reference to the silex-beds of Tampa and the Pliocene beds of the Caloosahatchie River, including in many cases a complete revision of the ge- neric groups treated of and their American Tertiary species. Part. IV. 1. Prionodesmacea: Nucula to Julia. 2. Telodesmacea: Teredo to Ervilia. Transactions of the Wagner Free In- stitute of Science of Philadelphia, 3(4): 571- 947; pls. 23-35. As Recent or Pleistocene from the Florida Keys: Arca (Lunarca) occidentalis, A. (L.) umbonata, Pecten (Chlamys, section Nodipecten) antillarum, P. (C., section С.) ornatus, P. (C., section Plagioctenium) gib- bus var. nucleus, Pinna carnea, Scapharca (S., section S.) transversa, S. (S., section S.) auriculata. DALL, W. H., 1899a, Synopsis of the Ameri- can species of the family Diplodontidae. Journal of Conchology, 9(8): 244-246. Diplodonta soror, with distribution to the Florida Keys. DALL, W. H., 1899b, Synopsis of the Solenidae of North America and the Antilles. Proceedings of the United States National Museum, 22(1185): 107-112. Ensis directus, with distribution to Indian Key. DALL, W. H., 1900a, Contributions to the Ter- tiary fauna of Florida, with especial reference to the silex-beds of Tampa and the Pliocene beds of the Caloosahatchie River, including in many cases a complete revision of the generic groups treated of and their Ameri- can Tertiary species. Part V. Teleodesmacea: Solen to Diplodonta. Transactions of the Wagner Free Institute of Science of Phila- delphia, 3(5): 949-1218, pls. 36-47. CRITICAL CATALOG AND BIBLIOGRAPHY 561 With Recent Florida Keys included in spe- cies distributions: Cardium (Papyridea) semisulcatum, Cumingia coarctata, Donax fossor, Ensis directus, Metis intastriata. DALL, W. H., 1900b, Synopsis of the family Cardiidae and of the North American spe- cies. Proceedings of the United States Na- tional Museum, 23(1214): 381-392. Protocardia tincta, from Key West. DALL, W. H., 1900c, Synopsis of the family Tellinidae and of the North American spe- cies. Proceedings of the United States Na- tional Museum, 23(1210): 285-326, pls. 2-4. With distributions including Florida Keys (or specific localities): Strigilla pisiformis, $. rombergii, Tellina (Eurytellina) angulosa, Т. (Scissula) candeana, T. (S.) exilis, T. (S.) iris. DALL, W. H., 1901, Synopsis of the Lucinacea and of the North American species. Proceed- ings of the United States National Museum, 23(1237): 779-833, pls. 39-42. With distributions including Florida Keys (or specific localities): Codakia orbicularis, Jagonia orbiculata var. filiata п. var., J. о. var. recurvata n. var., Myrtaea (Eulopia) sagrinata, Phacoides (Lucinisca) muricatus. DALL, W. H., 1902a, Illustrations and descrip- tions of new, unfigured, or imperfectly known shells, chiefly American, in the U.S. National Museum. Proceedings of the United States National Museum, 24(1264): 499-566, pls. 27-40. Listed and figured from the Florida Keys: Meretrix (Transennella) conradina. *DALL, W. H., 1902b, Synopsis of the family Veneridae and of the North American Re- cent species. Proceedings of the United States National Museum, 26(1312): 335- 412, pls. 12-16. With distributions including Florida Keys (or specific localities): Chione (Chione) intapurpurea, C. (C.) subrostrata, C. (Timoclea) pygmaea, Cytherea (Cytherea) listeri, C. (Ventricola) rigida, Cytherea (Ventricola) strigillina n. sp., Dosinia (Dosinidia) concentrica, D. elegans, Macrocallista (Chionella) maculata, Parastarte triquetra, Transennella stimpsoni n. sp., Venus mercenaria. DALL, W. H., 1903a, Contributions to the Ter- tiary fauna of Florida with especial reference to the silex beds of Tampa and the Pliocene beds of the Caloosahatchie River, including in many cases a complete revision of the generic groups treated of and their American Tertiary species. Part VI. Concluding the work. Transactions of the Wagner Free Insti- tute of Science of Philadelphia, 3(6): 1219- 1654, pls. 48-60. As Recent or fossil from Florida Keys locali- ties: Cardita (Carditamera) floridana, Codakia orbicularis, Cyrena (Pseudocyrena) floridana, Cytherea (Cytherea, section Ventricola) rugatina, Dosinia (Dosinidia) concentrica, Dosinia (Dosinidia) elegans, Lucina chrysostoma, Macrocallista nimbosa, Parastarte triquetra, Periploma angulifera, Phacoides (Lucinisca) muricatus, Venus mercenaria var. notata. DALL, W. H., 1903b, A preliminary catalogue of the shell-bearing marine mollusks and brachiopods of the southeastern coast of the United States, with illustrations of many of the species. Reprint to which are added twenty-one plates not in the edition of 1889. Bulletin of the United States National Mu- seum, no. 37: 1-232, 95 pls. With the same text entries as the original version (Dall, 1889a), with the following ad- ditional Florida Keys reference in the added plates (pls. 75-95): Cytherea (Ventricola) Strigillina. DALL, W. H., 1903c, Synopsis of the family Astartidae, with a review of the American spe- cies. Proceedings of the United States Na- tional Museum, 26(1342): 933-951, pls. 62-63. Astarte nana, with “Florida reefs” within its distribution. *DALL, W. Н., 1925, Notes on the nomencla- ture of some of our east American species of Pecten with descriptions of new species. The Nautilus, 38(4): 112-120. With distributions including Florida Keys (or specific localities): Pecten (Plagioctenium) nucleus, Pecten (Euvola) tereinus n. sp. from Key West. DALL, W. H., 1927, Small shells from dredgings off the southeast coast of the United States by the United States Fisher- ies Steamer “Albatross” in 1885 and 1886. Proceedings of the United States National Museum, 70 (Art. 18): 1-134. With distributions including Florida Keys: Poromya granulata. SDALTON, |., 1991, Shelling Marquesas Keys. Of Sea & Shore, 13(4): 165-166, 190. Arcinella arcinella collected July 2-4? [1991] at Marquesas Keys, with Metal Detector’s Club. *DAVIS, J. D., 1973, Systematics and distri- bution of western Atlantic Ervilia (Pelecypoda: Mesodesmatidae) with notes 562 on living Ervilia subcancellata. The Veliger, 15(4): 307-313, 3 pls. Ervilia concentrica, E. nitens and E. subcancellata contrasted, including Florida Keys material. SDEMARIA, K., 1996, Changes in the Florida Keys ecosystem based upon interviews with experienced residents. The Nature Conser- vancy, Key West; and Center for Marine Conservation, Washington, D.C. [Ш +] iii + 105 +5 +21 3 pp, An interesting interview-based compilation giving the historical perspective on many topics, including the impact of landfills on water circulation, freshwater runoff from the Everglades, the C-111 canal opened in the mid-1960s, etc. “Scallops” (probably Argopecten irradians) are discussed as com- ponents of the Florida Bay fauna off the Up- per and Middle Keys in the 1950s and 1960s. SDENT, S. R., 1998, Recent mollusk shell en- crustation patterns on the South Florida shelf: indicators of environmental conditions? [abstract] Geological Society of America Abstracts with Program (32° Annual Meet- ing, North-Central Section and associated societies), 30(2): 14. Epibiont coverage compared on Chamidae and Tellina spp. from the Florida Keys. DIAZ MERLANO, J. M. & М. PUYANA HEGEDUS, 1994, Moluscos del Caribe Colombiano, un catálogo ilustrado. Colciencias y Fundación Natura Colombia, INVEMAR, Santefe de Bogota, Colombia. 291 pp., 74 pls. Caribbean Colombian mollusks with “cayos de La Florida” in species distributions: Chama sarda, Divarilima albicoma, Lucina (Lucinisca) muricata, Nucula calcicola, Transenella (sic) cubaniana, Ventricolaria rigida. DOMANESCHI, О. & С. MANTOVANI MAR- TINS, 2002, Isognomon bicolor (С. В. Adams (Bivalvia, Isognomonidae): primeiro registro para о Brasil, redescricáo da espécie e consideracóes sobre a ocorréncia e distribuiçäo de /sognomon na costa brasileira. Revista Brasileira de Zoologia, 19(2): 611-627. From Florida Keys localities: /sognomon alatus, |. bicolor. DOMANESCHI, O. & E. K. SHEA, 2004, Shell morphometry of western Atlantic and Indo- West Pacific Asaphis; functional morphology and ecological aspects of A. deflorata from Florida Keys, U.S.A. (Bivalvia: Psammo- biidae). In: R. BIELER & P. M. MIKKELSEN, eds., MIKKELSEN & BIELER Bivalve studies in the Florida Keys, Proceed- ings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 249-275. Asaphis deflorata cited from West Summerland Key [material from this study]. SEDWARDS, C. E., 1968a, Looe Key have everything, almost. Seafari [Palm Beach County Shell Club Newsletter], 10(12): 1-4. 79 species were collected in two trips to Looe Key in November 1968 (plus a short stop at American Shoals) including Chama florida, Chlamys sentis, Pinna carnea, Spondylus ictericus, and from cracking rocks: Botula fusca, Cummingia (sic) antillarum, Gastrochaena hians, Lithophagis (sic) nigra, Petricola lapicida. SEDWARDS, C. E., 1968b, Snorkeling at Ba- hia Honda. Miami Malacological Society Quarterly, 2(2): 2-5. From a collecting trip off oceanside of Bahia Honda State Park in summer 1969: Arca imbricata, А. zebra, Arcopsis adamsi, Barbatia domingensis, Chlamys sentis, Codakia orbicularis, Isognomon radiatus, Lima pellucida. Reprinted with minor modi- fications in Of Sea & Shore, 7(3): 167-168. SEDWARDS, C. E., 1969, Off Tavernier on the reefs. Seafari [Palm Beach County Shell Club Newsletter], 11(7): 2-4. From the south end of Molasses Reef in May 1969: Chama florida, Chlamys imbricata, C. ornata, C. sentis. SEDWARDS, С. E., 1970, Off Key West's Key Wester [Motel]. Seafari [Palm Beach County Shell Club Newsletter], 12(8-9): 6-7. Collected near Key West during the 1970 American Malacological Union annual meet- ing: Arca imbricata, 5 spp. of ark shells, and large and small Chama spp. S*EDWARDS, С. [E.], 1980, Convention shell- ing and other thoughts ... Conchologists of America Bulletin, (21): 3, 8, 13. From the Keys during the Conchologists of America Convention in Key West, shortly after Hurricane Allen, on Geiger Key: Arca cancellaria, A. zebra, Brachidontes modio- lus, Codakia orbicularis, Periglypta listeri, Tellina fausta, T. radiata. A small unidenti- fied Pectinidae was also mentioned, as specimens presented to banquet attendees. SEDWARDS, C. E., 1987, Key West/Marquesas Keys trip. The Busycon [Broward Shell Club, Ft. Lauderdale, Florida], 22(8): 5. Account of shelling trip on 16 May 1987, list- ing identifiable species by common names CRITICAL CATALOG AND BIBLIOGRAPHY 563 only: Mytilidae, and “pearly-inside Oyster Shells”. EMERSON, W. К. 8 М. К. JACOBSON, 1976, The American Museum of Natural History guide to shells; land, freshwater, and ma- rine, from Nova Scotia to Florida. Alfred A. Knopf, New York. 482 + xviii pp., 47 pls. With distributions including Florida Keys (or specific localities): Botula fusca, Chione grus, Cumingia antillarum, Isognomon bi- color, Lithophaga antillarum, L. aristata, L. nigra, Моейа ponderosa, Pseudocyrena floridana, Tellina fausta, Transennella cubaniana. Atrina seminuda is “strangely absent” from the Keys. S*EUBANKS, L., 1964, A shell collector's va- cation in the Florida Keys. Texas Concholo- gist, 1(1): 3-4. From a shelling trip, at unspecified localities in the Keys: Anadara nobilis (sic), Arcopagia fausta, Barbatia cancellaria, Chama congregata, C. macerophylla, Chlamys sentis, Codakia orbicularis, Glycymeris pectinata, Lima lima, L. pellucida, Lucina pensylvanica, Papyridea soleniformis, Pseudochama radians, Pteria colymbus. FORBES, М. L., 1964, Distribution of the com- mensal oyster, Ostrea permollis, and its host sponge. Bulletin of Marine Science of the Gulf and Caribbean, 14(3): 453-464. Ostrea permollis from off Molasses Reef in UMML collection (based оп С. L. Voss, pers. comm.). SFOSTER, R. W., 1945, The Museum of Com- parative Zoology — Burry Marine Museum Expedition of 1944. The American Malaco- logical Union, News Bulletin and Annual Report, 1944-1945: 5. An account of dredging operations off the “Lower Florida Keys” including 40 stations from Carysfort Light to Molasses Reef, and from Sombrero Light to Looe Key, depths 21-117 fms, July-Aug. 1944, M/M L. A. Burry [Pom- pano, Florida], yielded 4,500 lots and ~350 species, including: Modiolus politus sagittatus, Pecten tereinus. Conspicuous for the number of species and abundance of specimens were Pectinidae, Cardiidae, Aloididae. SFRUMAR, F., 2000, Treasures from South Florida. The Greater St. Louis Shell Club web site, http://www.stlshell.com/members/ FrankFumar/dredging_off the florida keys. htm; last modified June 14, 2000; last ac- cessed April 12, 2001, 2 pp. From deep water off the Florida Keys, illus- trated by color photographs: Amusium laurenti, Chlamys benedicti. SGAERTNER, М. J., 1978, Florida shelling adventure. Of Sea & Shore, 9(2): 86. From Bahia Honda Key, 1948: “40 different species”, including egg cockles, mussels, sunrise clams, turkey wings. Also gives a description of shells collected at Bahia Honda and shown to the author by a shop owner in Cocoa, Florida, including Lion’s Paws. *GILMOUR, T. H. J., 1990, The adaptive sig- nificance of foot reversal in the Limoida. Pp. 249-263, in: B.S. MORTON, ed., Proceedings of a memorial symposium in honor of Sir Charles Maurice Yonge (1899-1986), Edinburgh, 1986. Hong Kong University Press, Hong Kong. viii + 355 pp. Material includes Ctenoides scabra from Big Pine Key. *GINSBURG, R. N., 1953, Intertidal erosion on the Florida Keys. Bulletin of Marine Sci- ence of the Gulf and Caribbean, 3(1): 55- 58. Key Largo Limestone erosion is described from Indian Key, Key Largo Sound, and the bay side of Key Vaca. Arca barbata, Mytilus (Brachidontes) exustus, and Acanthopleura granulata are attributed to the “generally shallow burrows”. Arca is stated to “almost buried in the rock”. No regional differences among the three localities are provided. SGODCHARLES, M. F. & W. C. JAAP, 1973, Fauna and flora in hydraulic clam dredge collections from Florida west and southeast coasts. Florida Department of Natural Re- sources, Marine Research Laboratory, Spe- cial Scientific Report, 40: 89 pp. Report of taxa collected during an explor- atory clam survey, including stations in Hawk Channel from Indian Key to Fowey Rocks. Raw data are presented for stations and taxa but no analysis is offered. Area Ш, charts 1249 (3.1-5.2 т) and 1250 (4.0-4.6 т) in- clude 13 box dredges by the R/V Hernan Cortez, oceanside along the Upper Florida Keys, in 1971. Chart 1249 shows stations 619 through 625, off Key Largo, between shore and the 10 m isobar. Chart 1250 shows stations 617-619, off the Matecumbes, also within the 10 m isobar. W. G. Lyons identi- fied the mollusks; “all live specimens’. Bivalves include: Anadara notabilis, Antigona listeri, Arca zebra, Argopecten gibbus, A. nucleus, Barbatia domingensis, Chama congregata, Chione cancellata, C. intapurpurea, Codakia orbicularis, Glycymeris pectinata, Laevicardium laevigatum, Lima lima, Lucina pensylvanica, 564 MIKKELSEN & BIELER Modiolus americanus, Pecten ziczac, Pseudochama radians, Trachycardium egmontianum, T. muricatum. SGOLDBERG, R., 1978, Shelling in the Florida Keys. Of Sea 8 Shore, 9(2): 109-110. From a compilation of three shelling trips to the Florida Keys between January 1977 and January 1978: Arca zebra, Chama macero- phylla, Isognomon alatus, Laevicardium laevigatum, Spondylus ictericus, Tellina fausta, Ventricolaria rugatina. A change in molluscan fauna was noted between Janu- ary 1977 and January 1978 at Indian Key Fill, attributed to a widening project for the Overseas Highway. Ohio Key oceanside 1$ described as one of the best collecting sites in the Keys. GRAU, G., 1955, A rectification of Pecten no- menclature. The Nautilus, 68(4): 113-115. The holotype of Pecten tereinus Dall [= P. chazaliei Dautzenberg] is cited as from off Key West. SGUNDERSEN, R., 1997, Blinded by the color. American Conchologist, 25(4): 19. Distinguishing characters of Caribachlamys sentis and C. ornata, from the Florida Keys. HARRY, H. W., 1985, Synopsis of the supraspecific classification of living oysters (Bivalvia: Graphaeidae and Ostreidae). The Veliger, 28(2): 121-158. Teskeyostrea weberi. SHAVILAND, E., 1994, Tellin’ it like it is. Of Sea & Shore, 17(1): 21-22. From Keys localities: Tellina radiata, “candy stripe tellin” (later called 7. similis). HAYAMI, |., 1984, Natural history and evolu- tion of Cryptopecten (a Cenozoic-Recent pectinid genus). University of Tokyo Press, Tokyo. ix + 149 pp., 13 pls. Cryptopecten phrygium from off Lower Keys, from АММН collection lots. SHAYES, H. L., 1972, The Recent Pteriidae (Mollusca) of the western Atlantic and east- ern Pacific Oceans. Ph.D. Dissertation, Georgetown University, Washington, D.C. 202 pp., 14 pls. Material examined (with localities and re- positories) includes Pinctada imbricata, Репа colymbus, Р hirundo vitrea, P. longisquamosa. HEMMEN, J. & С. HEMMEN, 1979, Beitráge zur Kenntnis der Meeresmollusken-Fauna der Karibischen See. Grenada. Jahrbuch, Nassauischer Verein für Naturkunde, 104: 137-172: Annotated species list, based on own col- lecting and literature. Those with specific reference to Florida Keys occurrence include Codakia (Ctena) pectinella. HENDERSON, J. B., 1911, Extracts from the log of the Eolis. The Nautilus, 25(6): 71-72; 25(7): 81-82. From Sand Key, 1910: Avicula, Lima, Lithodomus, Pecten. From off Key West, 1911: pectens. From Tortugas Islands, 1911: Limas, Pectens. *HENDERSON, y. B., 1913, Marine shells from drift on Upper Matecumbe Key, Florida. The Nautilus, 27(5): 59-60. From shore-drift collecting during Eolis cruises in May 1913: Chione grus, Codakia orbiculata, Pleuromeris tridentata, Transenella (sic) stimpsoni. HENDLER, С., J. E. MILLER, О. Е. PAWSON & Р.М. KIER, 1995, Sea stars, sea urchins, and allies: Echinoderms of Florida and the Caribbean. Smithsonian Institution Press, Washington, DC. 390 pp. Molluscan associations with Florida Keys echinoderms [based on literature and origi- nal research at Looe Key] include: commen- sal bivalve on Ophiophragmus septus (figured); juvenile Leptonacea symbiotic on Amphioplus sepultus; Mysella sp. C symbi- otic on arm spines of Amphipholis gracillima; Neaeromya sp. commensal on Meoma ventricosa. SHERTWECK, V., 1977, The Sarasota Shell Show — 1977. Of Sea & Shore, 8(1): 36-38. Shell of the show, Xenophora conchyliophora with a perfect Glycymeris pectinata attached, was found by Mr. & Mrs. Ernest Bradley [Bradenton, Florida] at Key West. HOWARD, J. F., D. LE. KISSLING RIRE LINEBACK, 1970, Sedimentary facies and distribution of biota in Coupon Bight, Lower Florida Keys. Geological Society of America Bulletin, 81: 1929-1946. An excellent description of Coupon Bight. Whole or fragmented molluscan shells form 22-87% of sediments in all parts of Coupon Bight. “From approximately six liters of un- consolidated sediment from each station sieved through a screen, 4,200 specimens assigned to 94 species of small mollusks were gathered and identified. Of these, 57 percent were bivalves and the remainder gastropods.” No overall species list is pro- vided, and no mention is made of voucher specimens. Identifications as per Abbott (1954) and Perry (1940). An open bay as- semblage is dominated by bivalves, espe- cially Chione cancellata, Laevicardium CRITICAL CATALOG AND BIBLIOGRAPHY 565 mortoni, Nucula proxima, and Pitar cf. fulminata. Restricted bay includes Anomalocardia cuneimeris, Parastarte tri- quetra, and Polymesoda floridana. Baymouth Bank includes Tellina cf. mera. Tidal channel fauna includes Codakia orbiculata, Lucina nassula, and Tellina candeana. Specifically mentioned as absent (while present in comparable Inner Reef Tract waters) are Arca umbonata, A. zebra, Atrina rigida, Barbatia cancellaria, and Codakia orbicularis. SHUDSON:I: A; 9. М. ALLEN & Т. J. COSTELLO, 1970, The flora and fauna of a basin in Central Florida Bay. United States Fish and Wildlife Service Special Scientific Report - Fisheries, 604, iii + 14 pp. Sampling sites, 1965-1968, were mudbanks surrounding Porpoise Lake (triangular de- pression, surrounded by mudbanks, bordered by Foxtrot Keys, Panhandle Key and Bob Allen Key; bayside of Indian Key Fill, next row of keys bayside of Shell Key) in the southern part of central Florida Bay. The authors used a wide variety of techniques (sled-mounted suction sampler, slednet, pushnet, beach seine, castnet, hand collecting, roller-frame trawl from bait-shrimp trawler, snorkeling, hook and line); maximum depth of “lake” 2.1 т; “... we did not attempt to retain plants and animals less than 5 mm long or wide.” (p. 4). Robert C. Work is given as the spe- cialist who identified the mollusks. Included are Anomalocardia cuneimeris, Arcopsis adamsi, Argopecten irradians concentricus, Brachidontes exustus, Cardita floridana, Chione cancellata, Codakia orbiculata, Laevicardium mortoni, Lima pellucida, Lyonsia hyalina floridana, Pinctada radiata, Tellina lineata, T. similis, T. tampaensis, Transennella cubaniana, T. stimpsoni. S*HUGHES, M., 1976, Field trip to the Keys. Conchologists of America Bulletin, (6): 4-5. From the Keys, June 1975, by Palm Beach and Broward Shell Clubs: Chlamys sentis. HUMFREY, M., 1975, Seashells of the West Indies — A guide to the marine molluscs of the Caribbean. Taplinger Publishing Com- pany, New York. 351 pp., 32 color pls. With distributions including Florida Keys (or specific localities): Aequipecten lineolaris, Antigona (Ventricolaria) rigida, Chama sarda, Chione paphia, Isognomon bicolor, Phacoides muricatus, Tellina punicea, Trachycardium magnum. ТЕК. о. № RUTSELE-S 0. Е. PISOR, 1997, Registry of world record size shells. Snail's Pace Productions, San Diego, California. ii + 101 pp. Florida Keys specimens among world size records: Arca zebra, Caribachlamys sentis, Isognomon alatus, Lyropecten antillarum, Mercenaria campechiensis, Pteria colymbus, Tellina magna. *INGHAM, R.E. 8 J. A. ZISCHKE, 1977, Prey preferences of carnivorous intertidal snails in the Florida Keys. The Veliger, 20(1): 49- Silk Included as molluscan prey species: Brachidontes exustus, Isognomon bicolor, and I. radiatus. |ISHMAN, S. E., С. L. BREWSTER-WIN- GARD, D. A. WILLARD, T. M. CRONIN, L. Е. EDWARDS & С. W. HOLMES, 1996, Pre- liminary paleontologic report on Core T-24, Little Madeira Bay, Florida. United States Geological Survey Open-File Report 96- 543, 47 pp. Electronic version available at http://pubs.usgs.gov/pdf/of/ofr96543.html; last accessed 12 September 2003. This work is here excluded as outside our defined area, which extends roughly half-way between the Florida Keys island chain and the tip of peninsular Florida. Little Madeira Bay is north of this limit. SIVERSEN, Е. $. & М. A. ROESSLER, 1969, Survey of the biota of Card Sound. Report to the Florida Power and Light Company. Institute of Marine and Atmospheric Sci- ences, University of Miami, Miami, Florida. 51 pp. Also available at: http://www.aoml. noaa.gov/general/lib/cedardoc.html; last ac- cessed 4 April 2003. A biotic survey of Card Sound in March—May 1969 included otter-trawl and plankton samples to assess various ecological zones, including mangroves, shallows and deep basins of the Sound. Aequipecten muscosus, Americardia media, Arca imbricata, A. umbonata, A. zebra, Argopecten irradians [an exploited species; also as Aequipecten or Aeguipecten (sic)], Atrina rigida, Barbatia cancellaria, bivalve unid., Brachidontes exustus [also as Trachidontes (sic)], Chione cancellata, Codakia orbiculata, Lima pellu- cida, Lyropecten antillarum, Modiolus americanus, Pinctada imbricata, Pteria colymbus, Trachycardium sp. SJAAP, W. C., 1984, The ecology of the South Florida coral reefs: a community profile. United States Fish and Wildlife Services, FWS/OBS - 82/08, 138 pp. Describes the impact by divers and aquarium collectors in the Florida Keys as “quite heavy 566 MIKKELSEN & BIELER on colorful and distinctive species such as ... thorny oyster (Spondylus spp.)”. JACOBSON, М. К. & L. HERNANDEZ, 1973, Ап unusual habitat for the rough file shell, Lima scabra (Born, 1780). The Veliger, 16(1): 85-86, 1 pl. Lima scabra cited from the Dry Tortugas. JINDRICH, V., 1969, Recent carbonate sedi- mentation by tidal channels in the Lower Florida Keys. Journal of Sedimentary Petrol- ogy, 39(2): 531-553. Molluscan shells form a major constituent of the sediments in Bluefish Channel, north of Key West, carbonate bank on Pleistocene bedrock: Arca umbonata, Chione cancellata, Codakia orbicularis. *JOHNSON, С. W., 1934, List of marine Mol- lusca of the Atlantic coast from Labrador to Texas. Proceedings of the Boston Society of Natural History, 40(1): 1-204. From Florida Keys localities: Antigona (Circomphalus, Ventricola) rigida, A. (C., V.) strigillina, Arca auriculata, Chama sarda, Chione (Timoclea) granulata, C. pubera, C. (T.) pygmaea, Codakia (Jagonia) orbiculata filiata, C. (J.) orbiculata recurvata, C. (J.) pectinella, Cochliolepis parasitica, Congeria rossmássleri, Corbula (Caryocorbula) cymella, Donax denticulata, Dosinia concentrica, Ervilia concentrica, E. nitens, Gouldia parva, Kellia rubra, Leiomya (Plectodon) granulata granulata, L. (P.) 9. velvetina, Lucina (Lucinisca) muricata, Myrtea (Eulopia) sagrinata, Nuculana verrilliana, Pecten (Chlamys) imbricatus, P. (Lyropecten) antillarum, P. (Plagioctenium) nucleus, P. (Euvola) tereinus, Pedalion listeri, P. semiaurita, Periploma angulifera, P. tenera, Petricola lapicida, Poromya granulata granulata, Protocardia tincta, Pteria hirundo vitrea, Strigilla pisiformis, Tellina (Arcopagia, Eurytellina) angulosa, T. (Angulus, Scissula) candeana, Т. (А., $.) iris, Thracia corbuloides, T. stimpsoni, Tivela mactroides, Transenella (sic) conradina, T. cubaniana, T. stimpsoni. JOZEFOWICZ, С. J. & D. О FOIGHIL, 1998, Phylogenetic analysis of Southern Hemi- sphere flat oysters based on partial mito- chondrial 16$ гОМА gene sequences. Molecular Phylogenetics and Evolution, 10: 426-435. Includes Ostreola equestris (as Teskey- ostrea weberi, misidentification indicated by P. Baker, unpub., in Kirkendale et al., 2004) from Big Pine Key. KIRKENDALE, L., T. LEE, P. BAKER & D. О FOIGHIL, 2004, Oysters of the Conch Re- public (Florida Keys); a molecular phyloge- netic study of Parahyotissa mcgintyi, Teskeyostrea weberi and Ostreola equestris. In: В. BIELER & Р. М. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 309-326. From Florida Keys localities: Dendostrea frons, Ostreola equestris, Hyotissa mcgintyi, Pinna sp., Teskeyostrea weberi [material from this study]. KISSLING, D. L., 1965, Coral distribution on a shoal in Spanish Harbor, Florida Keys. Bul- letin of Marine Science, 15(3): 599-611. The most common bivalves in Spanish Har- bor Channel are Arca umbonata, Atrina rigida, and Codakia orbicularis. KISSLING, D. L., 1977a, [Partial list of organ- isms ... from examination of patch reefs south of Boca Chica, Newfound Harbor Keys and at Mosquito Banks]. Pp. 181-182, in: H. G. MULTER, Field guide to some carbonate rock environments — Florida Keys and west- ern Bahamas, new ed. Kendall/Hunt Pub- lishing Company, Dubuque, lowa. 415 pp. + 10 maps. Arca umbonata, A. zebra, Barbatia sp., Brachiodontes recurvus, Codakia orbicularis, Isognomen (sic) alatus, Lithophaga antillarum. KISSLING, D. L., 1977b, [A partial list of or- ganisms that inhabit the surfaces and mar- gins of Rodriguez Bank]. P. 176, in: H. G. MULTER, Field guide to some carbonate rock environments - Florida Keys and western Bahamas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp. + 10 maps. Arca umbonata, Atrina rigida, Barbatia cancellaria, Chlamys sentis, Codakia orbicu- laris, Pinctada radiata, Tellina lineata. KLEEMANN, K. H., 1983, Catalogue of Re- cent and fossil Lithophaga (Bivalvia). Jour- nal of Molluscan Studies, Suppl. 12: 1-46. With distribution including Florida Keys: Lithophaga nigra. KNUDSEN, J. W., 1982, Anomalodesmata (Mollusca, Bivalvia) from Saba Bank, the Caribbean region. Proceedings, Koninklijke Nederlandse Akademie van Wetenschappen, Ser. C, 85(1): 121-146. Cardiomya alternata is included, citing Dall's (1881) Florida Keys record. *KRAEUTER, J. N., 1973, Notes on mollusks Ostrea and Siphonaria from Georgia (U.S.A.). The Nautilus, 87(3): 75-77. Ostrea permollis from Molasses Reef. CRITICAL CATALOG AND BIBLIOGRAPHY 567 KRAUSE, М. К., W. $. ARNOLD & W. С. AMBROSE, JR., 1994, Morphological and genetic variation among three populations of calico scallops, Argopecten gibbus. Journal of Shellfish Research, 13(2): 529-537. Argopecten gibbus, from the vicinity of the Marquesas Keys, is compared to populations from Cape Canaveral and North Carolina. SKRISBERG, M. F., 1993, А holiday observa- tion trip to the Keys. New York Shell Club Notes, no. 327: 11. From the Lower Florida Keys: Chione cancellata, Chlamys sentis, Lima lima, Papyridea soleniformis, Tagelus divisus, Tellina fausta, T. lineata, T. listeri, T. radiata. LAWSON, B., 1993, Shelling San Sal. Baha- mian Field Station, San Salvador, Bahamas. ix + 63 pp. With distribution including Florida Keys [ac- companied by black-and-white sketch]: Trachycardium magnum. SLEE, V., 1969, After the Lower Keys show. Seafari [Palm Beach County Shell Club Newsletter], 11(5): 8-9. Collected from various localities in the Lower Florida Keys: Anomalocardia brasiliensis (sic), Arca imbricata, Brachidontes citrinus, Chama macerophylla, Codakia orbicularis, Codakia orbiculata, C. o. ?form filiata, Glycymeris pectinatus, Lima pellucida, Pinctada radiata, Pseudochama radians variegata, Tellina similis. SLERMOND, М. W., 1936, Check list of Florida marine shells. Privately published, Gulfport, Florida. 56 pp. 247 Florida Keys bivalve names (= 216 spe- cies here considered valid), compiled from personal collections (1913-14 through date of publication) and records from various col- lectors (A. G. Reynolds, C. B. Lungren, C. C. Allen, D. L. Emery), professionals (H. Van Hyning, Florida State Museum, Gainesville; W. J. Clench, МСС; С. T. Stimpson, “veteran naturalist” of Little River, Florida; H. A. Pilsbry and E. G. Vanatta, ANSP; W. H. Dall and W. B. Marshall, USNM). From “beaches, reefs, and in the bays” of the Florida Keys, exclud- ing deep water forms seldom encountered by collectors [covered by Dall, 1889a, 1903b; Johnson, 1934]: Abra aequalis, A. lioica, Anatina lineata, A. (Raeta) canaliculata, Animalocardia (sic) brasilana (sic), A. cuneimeris, Anomia simplex, Antigona listeri, A. rigida, A. strigillina, Apolymetis intasriata (sic), Arca admsi (sic), A. auriculata, A. barbata, A. campechiensis americana, A. candida, A. chemnitzi, A. imbricata, A. incongrua, A. occidentalis, A. ponderosa, A. reticulata, A. secticostata, A. transversa, A. umbonata, Asaphis deflorata, Astarte nana, Asthenothaerus hemphillii, Avicula atlantica, Basterotia quadrata, В. 4. granatina, Botula castanea, B. fusca, Cardita floridana, Cardium arcinella [but corrected to Chama in this copy, signed by Lermond], С. isocardia, C. magnum, C. muricatum, C. (Hemicardium) medium, C. (Laevicardium) laevigatum, C. (L.) serratum, C. (Papyridea) semisulcatum, C. (P.) spinosum, C. (Protocardia) peramabilis, C. (Trigoniocardia) antillarum, Chama congregata, С. macerophylla, С. sarda, Circe cerina, Congeria rossmassleri, Coralliophaga coralliophaga, Corbula barrattiana, C. contracta, C. cymella, C. dietziana, C. disparillis (sic), C. nasuta, C. swiftiana, Crassatellites gibbsii, Crenella divaricata, Cumingia coarctata, C. tellinoides, Cuspidaria (Cardiomya) costellata, Cyclinella tenuis, Cytherea albida, C. hebraea, C. simpsoni, C. (Dione) dione, C. (Tivela) mactroides, C. (Transennella) conradiana (sic), C. (T.) cubaniana, Dacrydium vitreum, Donax denticulata, D. fossor, D. f. protractus, D. roemeri, D. tumidus, D. variabilis, Dosinia concentrica, D. discus, D. elegans, Egeta protexta, Ervilia concentrica, E. nitens, Gastrochaena cuneiformis, С. ovata, С. rostrata, Gemma purpurea, Glycymeris americana, G. lineata, G. pectinata, G. pennacea, Gouldia cerina, G. mactracea, G. parva, Heterodonax bimaculata, Iphigenia brasiliana, Lima hians, L. inflata, L. lima, L. scabra, L. tenera, Limatula confusa, Lithodomus antillarum, L. aristata, L. bisulcatus, L. nigra, Lucina pennsylvanica (sic), L. (Anodontia) jamaicensis, L. (A.) trisulcata, L. (Bellucina) amiantus, L. (Divaricella) dentata, L. (Jagonia) costata, L. (J.) orbiculata, L. (J.) o. filiata, L. (J.) o. recurvata, L. (J.) pectinella, Е. (Loripinus) edentula, Ё. (Lo) е. chrysostoma, L. (L.) schrammi, L. (Lucinisca) muricata, L. (Parvilucina) crenella, Lyonsia beana, Macoma brevifrons, М. cerina, M. constricta, M. leptonoides (sic), M. cimula (sic), М. tenta, М. t. souleyetiana, Macrocallista maculata, M. (Callista) gigantea, Mactra fragilis, Margaritifera ra- diata, Martesia caribaea, M. cuneiformis, M. striata, Modiolaria lateralis, M. arborescens, M. demissus, M. d. granosissimus, M. opifex, M. sulcatus, M. tulipus, Mulinia lateralis, Mytilus exustus, M. recurvus, Nucula 568 aegeensis, Nuculana carpenteri, М. solidula, N. verrilliana, Ostrea cristata, O. frons, O. verginica (sic), Pecten acanthodes, P. antillarum, P. exasperatus, P. gibbus, P. heliacus, P. imbricatus, P. nodosus, P. n. fragosus, P. nucleus, P. ornatus, Р. raveneli, P sentis, P. tereinus, P. ziczac, Pedalion bi- color, P. listeri, P. semiaurita, P. (Perna) alata, Periploma angulifera, P. tenera, Petricola pholadiformis, Р. lapicida, Pholas campechiensis, P. (Barnea) costata, P. (B.) truncata, Pinna carnea, P. rigida, P. serrata, Plicatula gibbosa, Pitar encymata (sic), P. fulminata, Polodesmus (sic) decipiens, Poromya granulata, Semele bellastriata, S. nuculoides, S. proficua, S. purpurascens, Solemya occidentalis, Spisula solidissima similis, Spondylus americanus, S. echinatus, Strigilla carnaria, S. flexuosa, S. pisiformis, $. rombergii, Tagelus divisus, Taras notata, T. nucleiformis, T. punctata, T. soror, Tellidora cristata, Tellina aequistriata, T. alternata, Т. angulosa, T. candeana, T. crystallina, Т. decora, Т. fausta, T. georgiana, T. gouldi, Т. interrupta, T. iris, T. lintea, T. lineata, Т. martinicensis, T. mera, T. modesta, T. radiata, Т. sayi, Т. squamifera, T. striata, T. tenera, Т. versicolor, Teredo clappi, T. thomsoni, Thracia corbuloides, Tivela mactroides, Transenella (sic) conradina, T. cubaniana, T. stimpsoni, Venus campechiensis, V. mercenaria, V. (Chione) cancellatus, V. (C.) granulatus, V. (C.) grus, М. (C.) latiliratus, М. (C.) mazyckii, V. (C.) paphia, V. (C.) pubera, V. (C.) pygmaeus, V. (C.) subrostrata, Verticordia ornata, Xylotrya fimbriata. LEVY, JM М. CHIAPPONE & К. М. SULLIVAN, 1996, Invertebrate infauna and epifauna of the Florida Keys and Florida Bay. Pp. 1-166, in: Site characterization for the Florida Keys National Marine Sanctuary and environs, Vol. 5. The Nature Conservancy, Florida and Caribbean Marine Conservation Science Center, University of Miami & The Preserver, Zenda, Wisconsin. Initially lists 5 classes, 26 orders, 31 fami- lies, and 712 species of mollusks in the Florida Keys, based on “24” uncited and unacknowledged references. Claims that (p. 24) “Previous studies have shown that mol- lusks are well-represented and serve impor- tant roles in benthic communities of the Florida Keys and Florida Bay (Appendices 7-8). Except for a few ecological invento- ries that include mollusks, there is a lack of comprehensive, ecosystem-wide species inventories for species in the Florida Keys.” MIKKELSEN 8 BIELER Appendix 7 gives a systematic list of mol- lusks “from southern Florida” [not restricted to Florida Keys], listing 359 species of gas- tropods, 174 bivalves, 13 cephalopods, 17 polyplacophorans, and 8 scaphopods, or a total of 571 species (not 712 species as cited earlier). Appendix 8 gives an alphabetical list- ing of molluscan species “recorded in south- ern Florida”. LINEBACK, J. A., 1977, Macrofaunal and flo- ral distributions and controls in Coupon Bight, Lower Florida. P. 96, in: H. G. MULTER, Field guide to some carbonate rock environ- ments — Florida Keys and western Baha- mas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp. + 10 maps. From Coupon Bight: Anomalocardia cuneimeris, Chione cancellata, Codakia orbiculata, Laevicardium mortoni, Lucina nassula, Nucula proxima, Parastarte trique- tra, Pinctada radiata, Pitar cf. fulminata, Polymesoda floridana, Tellina candeana, T. cf. mera. LONG ISLAND SHELL CLUB, 1988, Seashells of Long Island, New York: a guide to their identification and local status. Long Island Shell Club, New York. 209 pp. With distributions including Florida Keys: Noetia ponderosa. SLYMAN, F., 1943, About finding shells. Shell Notes, 1(4): 20. Pecten sentis and other Pecten (Chlamys) in Florida Keys, usually under stones, cor- als, inside rotten rock and coral. Also Lima, Lithoghaga while turning large slabs of stone or plate coral. Pedalion listeri on underside of flat stones between high and low water marks in Lower Keys, especially near “the long bridge” [presumably the Seven-Mile Bridge]. SLYMAN, F., 1944a, Expect to find these shells living as stated. Shell Notes, 1(7): 49. Illustrated (for purchase) from the Florida Keys: Pecten imbricatus, P. nucleus, P. ornatus, P. sentis. SLYMAN, F., 1944b, Shell collector’s paradise. Shell Notes, 1(8): 57-58, 1 map. A map of Indian Key includes a map of shal- low habitats as a guide to sampling meth- ods and expected species. On the northern side is sandy bottom and boggy flats (Codakia, etc.). On the northeastern side is grassy bottom. On the eastern side are small loose rock (pectens, etc.) and grassy bot- tom. On the southeastern side is jagged rock (Arca, etc.). On the southern side is rock CRITICAL CATALOG AND BIBLIOGRAPHY 569 bottom (many shelled species). On the west- ern side are grassy bottom and boggy flats (Codakia, etc.). SLYMAN, F., 1944c, Pearls are sometimes found in the Pinna shells ... Shell Notes, 1(9): 67. Pinna carnea, almost unknown in America, except the Florida Keys, on soft mud banks. SLYMAN, F., 1945, A report of a trip to the Florida Keys. Shell Notes, 1(14): 120-125. Off Key Largo, 26 July 1945, at a known reef that sticks out of the water at low tide, Pecten (Chlamys) Mildredaea (sic) [measuring 40 x 35 mm, from an additional paragraph on p. 126]. SLYMAN, F., 1946, Pinna carnea. Shell Notes, GOR OL Giant 10-12 inch specimens sometimes oc- cur on certain flats in the Florida Keys. SLYMAN, F., 1947a, A trip to the Florida Keys from the log book of the boat Junonia. Shell Notes, 1(19): 170-175. Modiolus tulipus on the beach at Duck Key, January 1, 1947. SLYMAN, F., 1947b, A report of our second shell hunting trip to the Florida Keys in 1947 ... from the log book of the Junonia. Shell Notes, 1(20): 187-194. By dredging in February 1947: Arca occidentalis at Rodriguez Key, Glycimerus (sic) pectinatus in Key Largo Sound. SLYMAN, F., 1948a, Pecten imbricatus ... Shell Notes, 2(2-3): 36. Pecten imbricatus categorized as the rarest species of Pecten in shallow waters in Florida, often found on the outer reefs of the Florida Keys. SLYMAN, F., 1948b, Barrier reef list. Shell Notes, 2(5): 72-74. An incomplete list of mollusks taken during a four-day cruise aboard the Junonia, on the “barrier reef” off the Florida Keys: Pecten sentis, P. imbricatus, Lima lima, Lithophaga antillarum. SLYMAN, F., 1949a, As to plate, outer reef shells (Fla. Keys). 1. Shell Notes, 2(7-9): 110-111. Illustrated (for purchase) from the Florida Keys: Pecten sentis red form and purplish form. SLYMAN, F., 1949b, As to plate, outer reef shells (Fla. Keys). 2. Shell Notes, 2(7-9): 122-123. Illustrated (for purchase) from the Florida Keys: Arca barbata outer reef form and in- shore form. SLYMAN, F., 1949c, As to plate, outer reef shells (Fla. Keys). 4. Shell Notes, 2(7-9): 128-129. Illustrated (for purchase) from the Florida Keys: Semele bellastriata, S. proficua, S. purpurascens, S. radiata. SLYMAN, F., 1950, Reef shell collecting. Shell Notes, 2(10-12): 128-129. Further comments about Bippus collecting trip [off Upper Keys, June 1950; see Bippus, 1950]: red Pecten sentis. SLYMAN, F., 1951, Dredging for shells out from Key Largo. Shell Notes, 2(13-15): 128-129. Dredging at 50+ ft., 5 April 1951: black pecten. SLYONS, W. G., 1999, Responses of benthic fauna to salinity shifts in Florida Bay: evi- dence from a more robust sample of the molluscan community. Pp. 47-49, in: Pro- grams and Abstracts, 1999 Florida Bay and Adjacent Marine Systems Science Confer- ence, Key Largo, Florida. From sampling in Florida Bay, 1994 and 1996: Brachidontes exustus, dominating in 1994. SLYONS, W. “$” (err. pro G.), 1998, Changes in benthic molluscan assemblages in Florida Bay, 1994-1996. Pp. 177-187, in: T. v. ARMENTANO, ed., Proceedings, Conference on Ecological and Hydrological Assessment of the 1994-95 High Water Conditions in the Southern Everglades, Miami, Florida, 22—23 August 1996. From sampling in Florida Bay, 1994 and 1996: Anomalocardia auberiana, Brachi- dontes exustus, Chione cancellata, Tellina tampaensis. CANONS W. ©. & 4. Е. QUINN, JR: 1995, Appendix J. Marine and terrestrial species and algae: Phylum Mollusca. Pp. J-10-J-26, in: Florida Keys National Marine Sanctuary Draft Management Plan / Environmental Impact Statement, Vol. Ш. Appendices. Na- tional Oceanographic and Atmospheric Ad- ministration, Silver Spring, Maryland. 630 species of marine mollusks are listed (undocumented) from the Florida Keys, in- cluding Florida Bay to Cape Sable (W. G. Lyons, pers. comm.); 423 species are gas- tropods. 207 species are bivalves: Abra aequalis, Aequipecten acanthodes, Americardia guppyi, A. media, Anadara notabilis, Anomalocardia auberiana, Arca imbricata, A. zebra, Arcinella cornuta, Arcopsis adamsi, Argopecten gibbus, A. irradians, A. nucleus, Asaphis deflorata, Asthenothaerus balesi, Atrina rigida, A. seminuda, Barbatia cancellaria, В. domingensis, B. tenera, Basterotia elliptica, B. quadrata, Botula fusca, Brachidontes domingensis, B. modiolus, Bractechlamys 570 antillarum, Carditamera floridana, Carditopsis smithii, Chama congregata, С. lactuca, С. macerophylla, C. sarda, C. sinuosa, Chione cancellata, C. intapurpurea, C. latilirata, C. puber, С. pygmaea, Chlamys beneditcti, С. imbricata, C. mildredae, C. multisquamata, C. ornata, C. sentis, Codakia costata, C. or- bicularis, C. orbiculata, C. pectinella, Coralliophaga coralliophaga, Crassinella lunulata, C. martinicensis, Crenella divaricata, Cumingia coarctata, Cyclinella tenuis, Dendostrea frons, Diplodonta punctata, D. semiaspera, Divaricella dentata, D. quadrisulcata, Entodesma beana, Ervilia concentrica, E. nitens, Gastrochaena hians, G. ovata, Glans dominguensis, Glycymeris decussata, G. pectinata, G. undata, Gouldia cerina, Gregariella coralliophaga, Isognomon alatus, |. bicolor, | radiatus, Laevicardium laevigatum, L. mortoni, L. sybariticum, Leporimetis intastriata, Lima lima, L. pellu- cida, L. scabra scabra, L. s. tenera, Linga amiantus, L. leucocyma, L. pensylvanica, Lioberus castaneus, Lithophaga antillarum, L. aristata, L. bisulcata, L. nigra, Lucina muricata, L. nassula, L. pectinata, Macoma brevifrons, Macrocallista maculata, Mactra fragilis, Malleus candeanus, Modiolus americanus, M. modiolus squamosus, Mus- culus lateralis, Nodipecten nodosus, Ostreola equestris, Papyridea semisulcata, P. soleniformis, Parvilucina blanda, Р. multilineata, Pecten chazaliei, P. ziczac, Periglypta listeri, Petricola lapicida, Pinctada imbricata, Pinna carnea, Pitar fulminatus, Р. simpsoni, Pleuromeris tridentata, Plicatula gibbosa, Polymesoda maritima, Pseudo- chama radians, Pteria colymbus, Pteromeris perplana, Rupellaria typica, Semele bellastriata, S. proficua, S. purpurascens, Semelina nuculoides, Solemya occidentalis, Spengleria rostrata, Spondylus americanus, S. ictericus, Strigilla сатапа, $. gabbi, $. mirabilis, Tellidora cristata, Tellina aequistriata, T. alternata, T. americana, Т. angulosa, T. candeana, T. consorbrina (sic), T. fausta, T. gouldii, T. laevigata, T. lineata, T. listeri, T. magna, T. martinicensis, Т. тега, T. nitens, T. paramera, T. probina (sic), Т. radiata, T. similis, T. squamifera, T. sybaritica, T. tampaensis, T. texana, T. versicolor, Trachycardium egmontianum, T. magnum, Т. muricatum, Transennella conradina, Т. cubaniana, T. stimpsoni, Ventricolaria rigida. MAGNOTTE, G., 1970-1979 (various ver- sions, all undated), Shelling & beachcomb- MIKKELSEN & BIELER ing in southern & Caribbean waters. Inter- national Graphics, Hollywood, Florida. 96 pp. With distributions including Florida Keys (or specific localities): Aequipecten gibbus, À. irradians, А. lineolaris, А. muscosus, Americardia media, Anadara lienosa floridana, А. notabilis, Anatina plicatella, An- odontia alba, Anomalocardia cuneimeris, Anomia simplex, Antigona listeri, Arca imbricata, A. zebra, Arcinella cornuta, Asaphis deflorata, Atrina rigida, Barbatia cancellaria, B. candida, B. tenera, Barnea truncata, Brachidontes exustus, B. recurvus, Chama congregata, C. macerophylla, C. sardo (sic), Chione cancellata, C. grus, C. paphia, Chlamys imbricatus, C. mildredae, C. sentis, Codakia orbicularis, Cyrtopleura costata, Dinocardium robustum, Diplodonta punctata, Dosinia discus, D. elegans, Eucrassatella speciosa, Glycymeris americana, С. decussata, С. pectinata, Iphigenia brasiliana, Isognomon alatus, |. radiatus, Laevicardium laevigatum, L. mortoni, Lima lima, L. scabra, Lithophaga antillarum, Lucina pensylvanica, Lyropecten antillarum, Macrocallista maculata, Mercenaria campechiensis, Modio- lus americanus, Mulinia lateralis, Noetia pon- derosa, Ostrea equestris, О. frons, Pecten ziczac, Petricola pholadiformis, Phacoides pectinata, Pinctada radiata, Pinna carnea, Pitar fulminatus, Plicatula gibbosa, Pteria colymbus, Spondylus americanus, S. gussoni, Strigilla romgergi (sic), Tagelus divisus, T. plebeius, Tellidora cristata, Tellina alternata, T. laevigata, T. lineata, T. listeri, T. magna, T. mera, T. radiata, T. similis, T. tampaensis, Trachycardium egmontianum, T. muricatum. More specifically distributed is Lyropecten nodosus (to Dry Tortugas). Ac- cording to the text, the illustrated specimens are deposited at Burry's Shell Museum, Pom- pano Beach, Florida. *MARELLI, D. C., М. К. KRAUSE, W. $. ARNOLD & W. С. LYONS, 1997, System- atic relationships among Florida populations of Argopecten irradians (Lamarck, 1819) (Bivalvia: Pectinidae). The Nautilus, 110(2): 31—41. Neither morphometric nor genetic evidence supports the distinction of А. irradians taylorae from A. irradians concentricus in Florida Bay. SMASON, L., 1969, AMU report - excerpts. Seafari [Palm Beach County Shell Club Newsletter], 11(11): 12-13. On display during the conference at the Uni- CRITICAL CATALOG AND BIBLIOGRAPHY 571 versity of Wisconsin — Green Вау т Marinette, Wisconsin, is Spondylus americanus, collected from the sides and deck of a salvaged vessel that sank near Key West. MAURY, С. J., 1920, Recent molluscs of the Gulf of Mexico and Pleistocene and Pliocene species from the Gulf states. Part 1: Pelecypoda. Bulletins of American Paleon- tology, 8(34): 35-147, pl. 1. With distributions including Florida Keys (or specific localities): Donax fossor, Ensis directus, Pteria vitrea, Scapharca auriculata, Thracia stimpsoni. MAURY, C. J., 1925, A further contribution to the paleontology of Trinidad (Miocene hori- zons). Bulletins of American Paleontology, 10(42): 153-402, pls. 12-54. Scapharca (Scapharca) auriculata, with dis- tribution including Key West. AMEGINTY. Р. L.-& Т. Е. McGINTY, 1957, Dredging for deep water shells in southern Florida. The Nautilus, 71(2): 37-47. An account of dredging operations from the cabin-cruiser Triton, off Palm Beach, Som- brero Key, and Key West. Keys mollusks in- clude: Aequipecten lineolaris, Antigona strigillina, Aurinia schmitti, Chama lactuca, Pecten chazaliei (tereinus), Pecten phrygium. McGINTY, T. L., 1939, Collecting on a coral reef in Florida. The Nautilus, 53(2): 37-39. From Middle Sambo Shoal, 8 mi SE of Key West: Arca, Chama, Spondylus. EMCGINTY, Т. L., 1942, Diving as applied to shell collecting. Pp. 32-36, in: The Ameri- can Malacological Union, The Eleventh An- nual Meeting, Rockland and Thomaston, Maine, August 26-29, 1941 [with] Papers Presented at the Symposium on Methods of Collecting and Preserving Mollusks, Wednesday, August 27, 1941. An account of “an imaginary diving trip” [but based on the author's experience?] on “a wreck of an old schooner which lies in thirty feet of water along the edge of a Florida coral reef”. Using a diving helmet, the diver finds Arca sp., Chama sp., Pecten nodosus “hang- ing like a pendant, byssus attached to a bit of old spar”, Spondylus sp. McGINTY, T. L., 1955, New marine mollusks from Florida. Proceedings of the Academy of Natural Sciences of Philadelphia, 107: 75— 85. Describes material from the cruiser yacht Triton [see Thompson et al., 1951]: Semele bellastriata. *MELVILL, J. C., 1880, List of Mollusca ob- tained in South Carolina and Florida (princi- pally at the island of Key West in 1871-1872). Journal of Conchology, 3: 155- 173. Anomalocardia impressa, Arca noae, Asaphis dichotoma, Barbatia sp., Callista (Dione) gigantea, Cardita (Mytilicardia) floridana, Cardium muricatum, Chama macerophylla, Chione cancellata, Hemicardium medium, Laevicardium laevigatum, L. serratum, Lima scabra, Loripes chrysostoma, L. edentula, Lucina jamaicensis, L. tigerina, Modiola plicatula, Mytilus cubitus, Ostrea frons, О. rhizophorae, Pectunculus pectiniformis, Pholas costata, Ricinula nodulosa, Scalaria venosa, Scapharca inaequivalvis, $. occidentalis, Spondylus ramosus, Strigilla carnaria, S. pisiformis, Tellina fausta, T. interrupta, T. lineata, T. radiata, T. r. var. unimaculata, T. robusta, T. similis, T. sol, T. tenera. Dall (1889a: 21) stated: “This cata- log contains many erroneous identifications.” SMIKKELSEN, P. M., 1981, Mollusks. Pp. 45- 48, in: S. C. JAMESON, ed., Key Largo Coral Reef National Marine Sanctuary Deep Wa- ter Resource Survey, NOAA Technical Re- port CZ/SP-1, 144 pp. From Johnson-Sea-Link submersible dives as part of Key Largo Coral Reef National Marine Sanctuary Deep Water Resource Survey: Arca imbricata, Arcopsis adamsi, Barbatia (Acar) domingensis, Chama sp., Diplodonta (Diplodonta) ?punctata, Isognomon radiatus. Voucher specimens referenced in Harbor Branch Oceanographic Museum, Ft. Pierce, Florida. MIKKELSEN, Р. М. 8 К. BIELER, 2000, Ma- rine bivalves of the Florida Keys: discovered biodiversity. Pp. 367-387, in: Е. М. HARPER, J. D. TAYLOR 8 J. A. CRAME, eds., The evolution- ary biology of the Bivalvia [Proceedings of “Biology & Evolution of the Bivalvia”, an in- ternational symposium organized by the Malacological Society of London, 14-17 September 1999, Cambridge, UK]. Geologi- cal Society, London, Special Publication 177. 325 bivalve species reported for the Florida Keys: Abra aequalis, A. lioica, Aequipecten glyptus, Americardia guppyi, A. media, Amusium laurentii, A. papyraceum, Amygdalum papyrium, A. politum, A. sagittatum, Anadara baughmani, A. floridana, A. notabilis, A. ovalis, A. transversa, Anatina anatina, Anodontia alba, 572 A. philippiana, Anomalocardia auberiana, Anomia simplex, Arca imbricata, A. zebra, Arcinella cornuta, Arcopsis adamsi, Argopecten gibbus, A. irradians, A. lineolaris, A. nucleus, Asaphis deflorata, Astarte nana, Asthenothaerus balesi, A. hemphilli, Atrina rigida, A. seminuda, A. serrata, Bankia carinata, Barbatia cancellaria, B. candida, B. domingensis, Barnea truncata, Basterotia elliptica, B. quadrata, Bathyarca glomerula, Botula fusca, Brachidontes domingensis, B. exustus, B. modiolus, Brachtechlamys antillarum, Callista eucymata, Cardiomya costellata, С. glypta, С. ornatissima, С. perrostrata, Carditamera floridana, Carditopsis smithii, Caribachlamys imbricata, C. mildredae, C. ornata, C. sentis, Chama congregata, C. florida, C. lactuca, C. macerophylla, С. sarda, С. sinuosa, Chione cancellata, C. mazyckii, C. paphia, Choristodon robustum, Circomphalus strigillinus, Codakia costata, C. orbicularis, C. orbiculata, C. pectinella, Coralliophaga coralliophaga, Corbula barrattiana, C. caribaea, C. contracta, C. dietziana, C. swiftiana, Crassinella dupliniana, С. lunulata, C. martinicensis, Crassostrea rhizophorae, С. virginica, Crenella decussata, Cryptopecten phrygium, Cryptostrea permollis, Ctenoides floridanus, С. planulatatus (sic), C. sanctipauli, C. scaber, Cumingia coarctata, C. tellinoides vanhyningi, Cuspidaria gigantea, C. rostrata, Cyclinella tenuis, Cyclopecten sp., Cymatioa sp., Cymatoica orientalis hendersoni, Cyrenoida floridana, Cyrtopleura costata, Dacrydium elegantulum hendersoni, Dendostrea frons, Diplodonta punctata, D. semiaspera, Divalinga quadrisulcata, Divaricella dentata, Divarilima albicoma, Donax variabilis, Dosinia discus, D. elegans, Ennucula tenuis, Ensis minor, Entodesma beana, Ervilia concentrica, E. nitens, E. subcancellata, Eucrassatella speciosa, Euvola chazaliei, E. raveneli, E. Ziczac, Gastrochaena hians, G. ovata, Geukensia granosissima, Glans dominguensis, Globivenus rigida, G. rugatina, Glycymeris americana, G. decussata, G. pectinata, G. undata, Gouldia cerina, Gregariella coralliophaga, Halirus fischeriana, Heterodonax bimaculatus, Hiatella arctica, Iphigenia brasiliana, Ischadium recurvum, Isognomon alatus, I. bicolor, I. radiatus, Kellia suborbicularis, Laevicardium laevigatum, L. mortoni, L. pictum, L. sybariticum, MIKKELSEN & BIELER Laevichlamys multisquamata, Lasaea adansoni, Leporimetis intastriata, Lima caribaea, Limaria pellucida, Limopsis aurita, L. cristata, L. minuta, L. sulcata, Lindapecten exasperatus, L. muscosus, Lioberus castaneus, Lirophora latilirata, Lithophaga antillarum, L. aristata, L. bisulcata, L. nigra, Lucina amianta, L. floridana, L. leucocyma, L. pectinata, L. pensylvanica, L. radians, L. sombrerensis, L. trisulcata, Lucinisca muricata, L. nassula, Lucinoma filosum, Lyonsia floridana, Lyropecten kallinubilosus, Macoma brevifrons, M. cerina, M. constricta, M. mitchelli, M. tageliformis, M. tenta, Macrocallista maculata, M. nimbosa, Mactrotoma fragilis, Malleus candeanus, Martesia cuneiformis, M. striata, Mercenaria campechiensis, M. mercenaria forma notata, Modiolus americanus, М. modiolus squamosus, Musculus lateralis, Myrtea sagrinata, Mysella planulata, Mytilopsis leucophaeata, M. sallei, Nemocardium peramabile, N. tinctum, Neopycnodonte co- chlear, Nodipecten nodosus, Noetia ponde- rosa, Nototeredo knoxi, Nucula aegeensis, N. calcicola, N. crenulata, N. proxima, Nuculana acuta, N. concentrica, N. pusio, N. solidula, N. verrilliana, Orobitella floridana, Ostreola equestris, Pandora bushiana, P. inflata, Papyridea semisulcata, PF. soleniformis, Parastarte triquetra, Parvilucina multilineata, Periglypta listeri, Periploma anguliferum, P. tenerum, Petricola lapicida, Petricolaria pholadiformis, Pinctada imbricata, P. longisquamosa, Pinna carnea, P. rudis, Pitar cordatus, P. fulminatus, P. simpsoni, Plectodon granulatus, Pleuromeris tridentata, Plicatula gibbosa, Polymesoda maritima, Poromya granulata, P. rostrata, Propeamussium pourtalesianum, P. sayanum, Protothaca granulata, Pseudochama тегае, P. radians, Репа colymbus, Pteromeris perplana, Puberella intapurpurea, P. pubera, Raeta plicatella, Rangia flexuosa, Semele bellastriata, S. proficua, S. purpurascens, Semelina nuculoides, Semierycina sp., Solecurtus cumingianus, Solemya occidentalis, Spathochlamys benedicti, Spengleria rostrata, Sphenia antillensis, Spisula raveneli, Spondylus americanus, $. gussoni, $. ictericus, Strigilla gabbi, $. mirabilis, $. pisiformis, Tagelus divisus, T. plebeius, Tellidora cristata, Tellina aequistriata, Т. agilis, T. alternata, T. americana, T. angulosa, T. candeana, T. consobrina, T. fausta, T. CRITICAL CATALOG AND BIBLIOGRAPHY 373 gouldii, T. iris, T. laevigata, T. lineata, Т. listeri, T. magna, T. martinicensis, T. mera, T. nitens, Т. рагатега, T. persica, Т. probrina, T. punicea, T. radiata, Т. similis, T. здиатйега, T. sybaritica, T. tampaensis, T. texana, T. versicolor, Teredo clappi, Thracia corbuloides, T. phaseolina, T. stimpsoni, Thyasira trisinuata, Timoclea grus, T. pygmaea, Tivela floridana, Trachycardium egmontianum, Т. magnum, T. muricatum, Transenella (sic) conradina, T. (sic) cubaniana, T. (sic) culebrana, T. (sic) stimpsoni, Trigonulina ornata, Varicorbula limatula, V. philippii, Verticordia acuticostata. MIKKELSEN, Р. М. & К. BIELER, 2001, Varicorbula (Bivalvia: Corbulidae) of the western Atlantic: taxonomy, anatomy, life habits, and distribution. The Veliger, 44(3): 271-293. With Florida Keys specimens cited [includ- ing material from this study]: Varicorbula disparilis, V. philippii. MIKKELSEN, Р. М. & R. BIELER, 2003, Sys- tematic revision of the western Atlantic file clams, Lima and Ctenoides (Bivalvia: Limoida: Limidae). Invertebrate Systematics, 17: 667-710, cover. With Florida Keys specimens cited [includ- ing material from this study]: Ctenoides mi- tis, C. planulatus, C. sanctipauli, C. scaber, C. miamiensis sp. nov., Lima caribaea. MIKKELSEN, Р. M. |. TEMKIN, В. BIELER & W. G. LYONS, 2004, Pinctada longisqua- mosa (Dunker, 1852) (Bivalvia: Pteriidae), an unrecognized pearl oyster in the western Atlantic. In: В. BIELER 8 P. М. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceed- ings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 473-501. Pinctada longisquamosa is redescribed based on living populations from the Florida Keys [material from this study]; Pinctada imbricata and Pteria colymbus also listed. SMILLER, J., 2001, Euvola ziczac (Linnaeus, 1758). American Conchologist, 29(1): back cover. Euvola ziczac figured from Key West. *MOORE, D. R., 1977, Small species of Nuculidae (Bivalvia) from the tropical west- ern Atlantic. The Nautilus, 91(4): 119-128. Nucula calcicola n. sp., from Key Largo. MOORE, Н. В. & В. N. LÓPEZ, 1970, A con- tribution to the ecology of the lamellibranch Tellina alternata. Bulletin of Marine Science, 20(4): 971-979. With distributions including Florida Keys (or specific localities): Tellina alternata, T. ra- diata. MORRIS, P. A., 1947, A field guide to the shells of our Atlantic coast. Houghton Mifflin Com- pany, Boston. xvii + 190 pp., 40 pls. With distributions including Florida Keys: Chama sarda, Codakia orbicularis. MORRIS, P.A., 1951, A field guide to the shells of our Atlantic and Gulf coasts, rev. and en- larged ed. Houghton Mifflin Company, Bos- ton. xix + 236 pp., 45 pls. With distributions including Florida Keys: Chama sarda, Codakia orbicularis, Tellina anguilosa (sic). MORRIS, P.A., 1973, A field guide to shells of the Atlantic and Gulf coasts and the West Indies, 3" ed. Houghton Mifflin Company, Boston. xxviii + 330 pp., 76 pls. With distributions including Florida Keys: Nuculana verrilliana, Pitar cordata. *MORRISON, J. P. E., 1958, Ellobiid and other ecology in Florida. The Nautilus, 71(4): 118— 124. Account of collecting at Bahia Honda Key and Plantation Key, Nov. 1955: Brachidontes sp., Crenella sp., Laemodonta cubensis, Pseudocyrena maritima. SMORRISON, J. P. E., 1970, East Florida Donax. Seafari [Palm Beach County Shell Club Newsletter], 12(7): 1-2, 5. “Unsolved problems” are noted regarding Key West Donax. MORTON, B., 2000, The pallial eyes of Ctenoides floridanus (Bivalvia: Limoidea). Journal of Molluscan Studies, 66(4): 449- 455. Using specimens from Stirrup Key [material from this study]. MORTON, B. & M. KNAPP, 2004, Predator- prey interactions between Chione elevata (Bivalvia: Chioninae) and Naticarius canrena (Gastropoda: Naticidae) in the Florida Keys, U.S.A. In: R. BIELER & P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceed- ings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 295-307. From Long Key and Lower Matecumbe Key [material from this study]: Pleuromeris tridentata, Chione elevata, Ctena orbiculata, Laevicardium mortoni, Lucinisca nassula, Pitar simpsoni, Tellina iris, T. тега, Т. similis, Tucetona pectinata. MPITSOS, G. J., 1973, Physiology of vision in the mollusk Lima scabra. Journal of Neuro- physiology, 36(2): 371-383. 574 MIKKELSEN & BIELER Lima scabra and L. $. tenera collected from the Florida Keys for visual physiological stud- ies. NUTTING, C. C., 1895, Narrative and prelimi- nary report of Bahama Expedition. Bulletin from the Laboratories of Natural History of the State University of lowa, 3(1-2): i-vi + 1-252. The 1893 S. U. I. Bahama Biological Expe- dition narrative contains reference to mol- lusks collected from the Dry Tortugas and Pourtales Plateau off Key West: Arca velata, Arca sp., Avicula margaritifera, Cardium isocardium, Chione cigenda, Lucina tigrina, Pecten ornatus. The author noted that “There is no place, probably, on our Atlantic coast where Mollusca are more abundant and more conspicuous than at the Tortugas” (p. 127). The mollusks were reported on more completely by Dall (1896a). S*ODE, H., 1975, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Part 1. Texas Conchologist, 12(2): 40-56. With distribution including Key West (citing earlier literature): Strigilla gabbi. S*ODE, H., 1976a, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Part I. Texas Conchologist, 12(3): 79-94. With distributions including Florida Keys (cit- ing earlier literature): Pseudocyrena mar- itima, Ventricolaria rigida. SODÉ, H., 1976b, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Texas Conchologist, 12(4): 108-124. With distributions including Florida Keys (cit- ing earlier literature): Pitar cordatus, Transennella cubaniana. SODÉ, H., 1977a, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Texas Conchologist, 13(3): 74-81, 84-88. With distribution including Florida Keys (or specific localities): Plectodon granulatus. SODE, H., 1977b, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Texas Conchologist, 13(4): 106-107, 114-122. With distribution including Florida Keys (cit- ing earlier literature): Myrtea sagrinata. SODE, H., 1979a, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Texas Conchologist, 15(3): 69-80. With distribution including Florida Keys (cit- ing earlier literature): Isognomon bicolor. SODE, H., 1979b, Distribution and records of the marine Mollusca in the northwest Gulf of Mexico (a continuing monograph). Texas Conchologist, 16(1): 14-32. With distribution including Florida Keys (cit- ing earlier literature): Divarilima albicoma. SODE, H., 1984, Additions to monographic list of bivalves of the northwest Gulf of Mexico. Texas Conchologist, 20(3): 76-83. With distributions including Florida Keys (cit- ing earlier literature): Cymatioa sp. D. OLIVER, Р. С. & J. JÁRNEGREN, 2004, How reliable is morphology based species tax- onomy in the Bivalvia? A case study on Arcopsis adamsi (Bivalvia: Arcoidea) from the Florida Keys. In: В. BIELER & P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the Interna- tional Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 327- 330: Acar domingensis, Arca imbricata, Arca sp., Arcopsis adamsi, Brachidontes sp., Chama sp., /5одпотоп sp., oysters [material from this study]. “OLSSON, A. A., 1951, New Floridan species of Ostrea and Vermicularia. The Nautilus, 65(1): 6-8, pl. 1. Ostrea weberi n. sp. described from Grassy Key and Key West. OLSSON, А. А. 8 А. HARBISON, 1952 [re- print 1979], Pliocene Mollusca of southern Florida with special reference to those from North Saint Petersburg. Academy of Natu- ral Sciences of Philadelphia Monograph 8. v + 457 pp., 65 pls. With Recent distributions including Florida Keys (or specific localities): Cumingia coarctata, Hemimetis (Florimetis) intastriata, Papyridea semisulcatum, Tellina (Scissula) candeana, T. (S.) similis. SORLIN, Z., 2003, A shelling trip to Florida and the Bahamas. La Conchiglia, 34(305): 36-40. While based at Grassy Key, the author col- lected on the Atlantic beaches of Key Vaca, Missouri Key, Ohio Key, and Boca Chica. The largest number of species (44) was found on Ohio Key; 34 species were not found on the Gulf coast of Florida. An additional 28 species were collected as microshells. On Boca Chica beach, Atlantic side: Codakia orbicularis, Lucina pensylvanica. Also from unclear localities: Arca zebra, Chama macerophylla, Pinctada imbricata, Pteria CRITICAL CATALOG AND BIBLIOGRAPHY 575 colymbus. In all, 45 species were found that were not found the weeks before on Florida's Gulf coast. PALMER, K. V. W., 1927-1929, The Veneridae of eastern America, Cenozoic and Recent. Palaeontographica Americana, 1(5): 209- 522 (1927), pls. 32-76 (1929). From Florida Keys: Antigona (Dosina) listeri. *PALMER, К. V. W., 1947, Notes on Costacallista eucymata (Dall). The Nautilus, 61(2): 44-47, pl. 4. Stations cited from off Key Largo to Looe Key. PARKER, R. H. 8 J. R. CURRAY, 1956, Fauna and bathymetry of banks on continental shelf, northwest Gulf of Mexico. Bulletin of the American Association of Petroleum Geologists, 40(10): 2428-2439. With distributions including Florida Keys (or specific localities): Corbula cymella, Trachycardium magnum. PEARSE, A. 5., 1929, Observations on certain littoral and terrestrial animals at Tortugas, Florida, with special reference to migrations from marine to terrestrial habitats. Papers from Tortugas Laboratory of the Carnegie Institution of Washington, 26(6) (Carnegie Institution of Washington Publication 391): 205-223. From Loggerhead Key and Garden Key, Dry Tortugas: Arca saccharina, Pteria vitrea. S*PEASE, R., 1980, Don’t underestimate tourist spots. Conchologists of America Bulletin, (20): 14. Account of diving at Sand Key in May 1979: Chlamys sentis. PETERSEN, D. W., 1989, Assessing environmental parameters and transport from the spatial distribution of a mollusc- dominated modern shell concentration in a restricted subtropical lagoon, Long Key Lake, Florida Keys, USA. The Compass [Earth Science Journal of Sigma Gamma Epsilon], 67(1): 15-29. An assessment of the malacofauna of ап oceanside lagoon on Long Key includes Acropsi (sic) adamsi, Anomalocardia auberiana, Argopectin (sic) nucleus, Brachiodonta (sic) exustus [listed under Gastropoda], Carditamera floridana, Chione cancellata, Codakia costata, Laevicardium mortoni, Ostrea frons, Pseudocyrena maritima, Tellina mera, T. texana. This is a published version of a Master's thesis (Petersen, 1988). *PETUCH, E. J., 1987, New Caribbean molluscan faunas. Coastal Education é Research Foundation (CERF), Carlottesville, Virginia. 154 + 4 pp., incl. 29 pls. Argopecten irradians taylorae n. ssp. from Rabbit Key Basin, Upper Florida Keys. *PETUCH, E. J., 1988, Neogene history of tropical American mollusks. Coastal Estuarine & Research Foundation, Charlottesville, Virginia. 217 pp. From Florida Keys localities: Brachidontes modiolus. Holotype of Argopecten irradians taylorae mentioned and refigured. *PILSBRY, H. А., ed., 1890a, American Association of Conchologists, December 10, 1890. The Nautilus, 4(8): 91-95. Among donations to the collection of the society is Pinna carnea from Key West. *PILSBRY, H. A., ed., 1890b, American Association of Conchologists, December 31, 1890. The Nautilus, 4(9): 104-107. Among donations to the collection of the society is Cardium magnum from Key West and Asaphis deflorata from Elbow Key [= Reef]. PILSBRY, H. А. & Т. E. MEGINTY 1938, Review of Florida Chamidae. The Nautilus, 518) 73-79 plat Chama sarda not observed but earlier reported from Florida Keys by earlier authors. *PILSBRY, Н. А. & К. A. MCLEAN, 1939, A new Агса from the West Indian region. Notula Naturae, по. 39: 1-2. Arca (Barbatia) balesi n. sp., from Missouri Key and Key West. *PILSBRY, Н. А. & A. А. OLSSON, 1946, Condylocardia in Florida and middle America. The Nautilus, 60(1): 6-7, pl. 1. Condylocardia floridensis n. sp., from Ohio Key. SPLOCKELMAN, C., 1968a, Chione pygmaea. Seafari [Palm Beach County Shell Club Newsletter], 10(12): 5. Chione pygmaea as uncommonly found in the Florida Keys. SPLOCKELMAN, C., 1968b, Do you collect chiones? Seafari [Palm Beach County Shell Club Newsletter], 10(8): 8-10. Chione pygmaea occurs uncommonly under rocks and on old conchs, in generally silty spots in the Florida Keys. SPLOCKELMAN, С. H., ed., 1968c, Entries for the Whopper Club. Seafari [Palm Beach County Shell Club Newsletter], 10(9): 7. Among large specimens collected by club members is Papyridea soleniformis (50 mm) from Molasses Keys. 576 MIKKELSEN & BIELER GPLOCKELMAN, C., 1968d, Mid-summer review of shelling. Seafari [Palm Beach County Shell Club Newsletter], 10(8): 1-3. From the Florida Keys, June—July 1968: Americardia sp., Codakia orbicularis, Tellina sp., Trachycardium sp. SPLOCKELMAN, C. H., 1968e, Tiny chiton on Codakia. Seafari [Palm Beach County Shell Club Newsletter], 10(9): 3. Codakia orbicularis abundant on Little Duck Key. SPLOCKELMAN, C. [H.], 1969a, A Florida prize. Seafari [Palm Beach County Shell Club Newsletter], 11(11): 5-6. With type locality of Pseudochama inezae given as Carysfort Reef. SPLOCKELMAN, C. [H.], 1969b, November in the Keys. Seafari [Palm Beach County Shell Club Newsletter], 11(1): 12. From Key West, mid-November 1969: Chlamys sentis among very little variety in bivalves. From Grassy Key: Barbatia cancellaria, Lima scabra tenera. Of special note on south Key West beach: large-sized Brachidontes citrinus. SPLOCKELMAN, C. H., ed., 1969c, Whopper Club. Seafari [Palm Beach County Shell Club Newsletter], 11(2): 3. From Crawl Key: Rupellaria typica (35.7 mm). SPLOCKELMAN, C. H., ed., 1970a, Eleventh annual shell show. Seafari [Palm Beach County Shell Club Newsletter], 12(3): 1-3. An award winning display by Corinne Edwards featured a Strombus gigas from the Florida Keys with a juvenile Spondylus ictericus lodged in the suture of the body whorl. SPLOCKELMAN, C. [H.], 1970b, Maybe you have one? Seafari [Palm Beach County Shell Club Newsletter|, 12(5): 5-6. Aequipecten acanthodes from west of Sombrero Light is noted in a display of deep water species at Elsie Malone’s shell shop (Sanibel Island). SPLOCKELEMAN, "С. [НН], 19/706, Re: Brachidontes citrinus Roding (sic). Seafari [Palm Beach County Shell Club Newsletter], 12(3): 14. Brachidontes citrinus is found on beaches in the Florida Keys, also living embedded in sili among seagrass and algae (patchy in distribution). SPLOCKELMAN, C. [H.], 1970d, Shells of Palm Beach County #23. Seafari [Palm Beach County Shell Club Newsletter], 12(6): 2-3. Cooperella atlantica from Little Duck Key. SPOLAND, P., 2001, Reef encounters of the first kind. Shell-o-Gram (Jacksonville [Florida] Shell Club), 42(3): 1,6. Reproduced at http://www.jaxshells.org/looe.htm; last accessed May 29, 2001. Caribachlamys sentis from Looe Key in the early 1970s. SPOMPEY, 5. L., 1974, Introducing the pretty pecten. Of Sea & Shore, 5(4): 161-164, 166. With distribution including Florida Keys: Aequipecten lineolaris. SPULLEY, T. E., 1952, A zoogeographic study based on the Bivalves of the Gulf of Mexico. Ph.D. Dissertation, Harvard University, Cambridge, Massachusetts. 215 pp., 19 pls. With distributions or figure captions including Florida Keys (or specific localities): Aequipecten gibbus nucleus, Aloidis aequivalvis, A. operculata, Amusium papyraceum, Amygdalum arborescens, A. papyrium, Anadara baughmani, А. transversa, Antigona listeri, Astarte nana, Botula castanea, B. fusca, Brachidontes citrinus, Cardiomya perrostrata, Cardita domingensis (sic), C. floridana, Chlamys imbricatus, C. muscosus, C. phrygius, C. sentis, Codakia orbicularis, Coralliophaga coralliophaga, Costacallista eucymata, Cumingia antillarum, Cyrenoida floridana, Diplodonta simiaspera (sic), Eucrassatella speciosa, Euvola raveneli, Glycymeris decussata, С. pectinata, G. undata, Isognomon alata, |. bicolor, |. listeri, Lima caribaea, L. scabra, L. tenera, Lithophaga antillarum, Lucina amiantus, L. leucocyma, L. sombrerensis, Lyrodus pedicellata, Lyropecten antillarum, L. nodosus, Macoma cerina, M. extenuata, M. limula, M. pseudomera, Musculus lateralis, Nucula proxima, Nuculana carpenteri (cotype figured), Papyridea semisulcata, Pinctada radiata, Pseudocyrena floridana, Tellina candeana, T. lintea, T. тега, Т. здиатйега, T. versicolor, Teredo bartschi, Venericardia tridentata. S*PURTYMUN, B., 1997, Echoes of the past. American Conchologist, 25(4): 28. From vicinity of Key West, 1944: Chama macerophylla. S*REDLA, М. T., 1990, An excursion to the fabled land of Florida. Texas Conchologist, 26(2): 60-61. Codakia orbicularis and Linga pensylvanica from Keys localities. *REHDER, Н. A., 1939, New marine mollusks from the west Atlantic. The Nautilus, 53(1): 16-21, pl. 6. CRITICAL CATALOG AND BIBLIOGRAPHY 577 New species from Keys localities: Cymatoica orientalis hendersoni, Cumingia tellinoides vanhyningi. *REHDER, H.A.,1943a, New marine mollusks from the Antillean region. Proceedings of the United States National Museum, 93(3161): 187-203, pls. 19-20. Asthenothaerus balesi n. sp., from Missouri Key. REHDER, H.A., 1943b, Corrections and eco- logical notes on some recently described Florida marine shells. The Nautilus, 57: 32- 33 From Missouri Key: Asthenothaerus balesi. REHDER, H. A., 1981, The Audubon Society field guide to North American seashells. Alfred A. Knopf, New York. 894 pp., 705 figs. With distributions including Florida Keys (or specific localities): Chama sarda, Chlamys sentis, Glans dominguensis, Isognomon bi- color, Lucina dentata, L. leucocyma, Parvilucina amianta, P. multilineata, Phacoides muricata, Pleuromeris tridentata. *REHDER, H.A. & К. T. ABBOTT, 1951, Some new and interesting mollusks from the deeper waters of the Gulf of Mexico. Revista de la Sociedad Malacologica “Carlos de la Torre”, 8(2): 53-66, pls. 8-9. Anadara springeri п. sp., from the Dry Tortugas. Also cited from Florida Keys locali- ties are Aequipecten glyptus, Amusium papyraceum, Pitar (Pitarenus) cordata. RICHARDS, H. G., 1936, Some shells from the North Carolina “banks”. The Nautilus, 49(4): 130-134. Arca auriculata from the Keys. SRING, Е. M., 1980, Of coral reefs, intertidal shores and sand flats in the Florida Keys. Of Sea & Shore, 11(2): 129-135. An ecological account of the marine habi- tats of the Keys, based on a March [presum- ably 1980] vacation trip. Specific localities visited were Sand Key Light, bridge channel between Boca Chica Key and Key West, and seaward sand flats off Bahia Honda Key, Marathon and Key Largo. Among the few mollusks specifically mentioned or figured are: Lima scabra, Ostrea frons, mussels. RIOS, E. DE C., 1994, Seashells of Brazil, 2" ed. Museu Oceanográfico, Rio Grande, Bra- zil. 368 pp., 113 pls. With distributions including Florida Keys (or specific localities): Chama sarda, Corbula (Caryocorbula) cymella, Limaria albicoma, Lucina (Lucinisca) muricata, Nuculana (Jupitaria) solidula, Pitar (Pitarenus) cordatus, Strigilla (Strigilla) gabbi, Trachycardium (Acrosterigma) magnum, Ventricolaria (Ventricolaria) rigida. RIPPLE, J. [with photographs by B. KEOGH 8 J. RIPPLE], 1995, The Florida Keys - the natural wonders of an island paradise. Voyageur Press, Stillwater, Minnesota. 128 pp. Includes a photograph of a living Ctenoides scaber (as “rough fileclam”) from an unspeci- fied locality. ROGERS, G. F., 1941, Wreck of the Janthina janthina. The Nautilus, 54(3): 75-77. Janthina janthina washed ashore on Key Largo; also “a good Venus clam, two nice live cowries and several live discus clams”. ROGERS, J. E., 1908, The shell book, a popu- lar guide to a knowledge of the families of living mollusks, and an aid to the identifica- tion of shells native and foreign. Doubleday, Page & Company, New York. хх! + 485 pp. With distributions including Florida Keys (or specific localities): Arca transversa, Lithodomus lithophagus, Lucina floridana. *ROMBOUTS, A., 1991, Guidebook to pecten shells; Recent Pectinidae and Propea- mussiidae of the world. Universal Book Ser- vices/Dr. W. Backhuys, Oegstgeest. 157 pp. Figured from Keys localities: Aequipecten glyptus, A. muscosus, Euvola ziczac, Nodipecten nodosus. ROMASHKO, S., 1984, The complete collector's guide to shells 8. shelling. Wind- ward Publishing Company, Miami, Florida. 112. рр: With distributions including Florida Keys (ог specific localities): Aequipecten lineolaris, Chlamys sentis, Trachycardium magnum. ROOPNARINE, Р. D. 8 С. J. VERMEIJ, 2000, One species becomes two: the case of Chione cancellata, the resurrected C. elevata, and a phylogenetic analysis of Chione. Journal of Molluscan Studies, 66(4): 517-534. Among the material used to distinguish the two species is Chione elevata from Key Largo (ANSP 264071). SROSS, B., 1969, Field trip to the Keys. Seafari [Palm Beach County Shell Club Newsletter, 11(7): 8-10. Collected in the Marathon area, during Me- morial Day weekend 1969: Anadara notabilis, Anomalocardia cuneimeris, Antigona listeri, Arca imbricata, А. zebra, Arcopagia fausta, Barbatia cancellaria, B. domingensis, Brachidontes citrinus, B. exustus, Chama congregata, C. macero- phylla, Chione cancellata, Chlamys sentis, Codakia obicularis (sic), C. orbiculata, 578 MIKKELSEN & BIELER Glycymeris pectinata, Isognomon alatus, |. radiatus, Laevicardium laevigatum, L. mortoni, Lima pellucida, L. scabra, L. $. tenera, Lima sp., Lucina pensylvanica, Mactra fragilis, Modiolus americanus, Papyridea soleniformis, Pinctada radiata, Pinna carnea, Tellina mera, T. paramera, T. similis, Trachycardium egmontianum, T. muricatum. SROSS, B., 1971, The Keys — what's become of Melongena bicolor? Seafari [Palm Beach County Shell Club Newsletter], 13(5): 14-15. Collecting out of Marathon, May [1971]. A few bivalves in mud at East Sister Rock; Chlamys sentis and Lyropecten antillarum at Looe Key; Botula fusca and Tellina similis from Little Duck/Ohio/Missouri Keys; several Tellina from further north up the Keys. From the more productive gulf side [locale unspeci- fied], Mr. Bennett (at Palm's Motel) found Lima scabra tenera by snorkeling. “While shelling in the Keys is not what it used to be, it is still better than any other place in Florida, on a day-to-day basis.” S*SAGE, W., 1987, A day's collecting in the Florida Keys. American Conchologist, 15(3): 12. Collecting Бу scuba near Boca Grande off Key West at 15-18 ft deep: Chlamys imbricata, C. sentis, Lima scabra. À second station 5 miles off Key West, less than 6 ft deep: Chlamys imbricata, C. sentis. *SALAS, С. & $. GOFAS, 1997, Brooding and non-brooding Dacrydium (Bivalvia: Mytilidae): a review of the Atlantic species. Journal of Molluscan Studies, 63(2): 261- 283. Dacrydium elegantulum hendersoni n. ssp. described from Eolis specimens from Sand Key; other specimens from Key West and Western Dry Rocks. Reports of D. vitreum (Holbgll in Moller, 1842) are attributed to this species. SALISBURY, A. E., 1952, Mollusca of the Uni- versity of Oxford Expedition to the Cayman Islands in 1938. Proceedings of the Malaco- logical Society of London, 30(1-2): 39-54, piso: With distribution including Florida Keys (cit- ing previous authors): Chlamys nucleus. SSCHOMER, М. 5. & R. D. DREW, 1982, Ап ecological characterization of the lower Ev- erglades, Florida Bay and the Florida Keys. United States Fish and Wildlife Service, Of- fice of Biological Services, Washington, D.C., FWS/OBS-82/58.1, xv + 246 pp. Chapter 9 lists Florida Keys mollusks, in great detail, by habitat (presumably based on Stephensen & Stephensen, 1950): Anadara sp., Anomalocardia cuneiveis (sic), Агса зр., Atrina rigida, А. seminuda, Barbatia sp., Chione cancellata, Codakia orbicularis, C. orbiculata, Crassostrea virginica, Isognomon alatus, Laevicardium laevigatum, Lithophaga sp., Mytilus exustus, Tellina sp. SSCHROEDER, R. E., 1964, Photographing the night creatures of Alligator Reef. National Geographic, 125(1): 128-154. Color photographs of living Dendostrea frons [as “coon oysters ... on dead догдотап”], and Ctenoides floridanus [as “flame scallop”]. *SCHWENGEL, J. S., 1951, New marine mol- lusks from British West Indies and Florida Keys. The Nautilus, 64(4): 116-119, pl. 8. Pitaria cordata n. sp. from Key West. S*SEDLAK, R., 1986, A perfect “10”. The Busycon [Broward Shell Club, Ft. Lauder- dale, Florida], 21(9): 5-7. From Money Key [near Little Duck Key at 7- Mile bridge, from a boat out of mile marker 50], August 1986: ВаграНа cancellaria, Codakia orbicularis, Papyridea soleniformis, Pinctada imbricata, Pinna carnea. *SHIRAI, S., 1994, Pearls and pearl oysters of the world. Marine Planning Company, Okinawa, Japan. 108 pp. Pinctada imbricata figured from Key West, as Akoya Pearl Oyster. SHOEMAKER, A. H., 1973, Thermal and sa- linity effects on ciliary activity of excised gill tissue from bivalves of North and South Carolina. The Veliger, 15(3): 215-222. Arca zebra shells litter beaches of the Florida Keys after storms, attributed to preferred shallow depths in this part of its distribution. SIEKMAN [as SEIKMAN (sic)], L. [with color illustrations by Е. MALONE], 1965, The great outdoors book of shells, 1‘ ed. Great Out- doors Publishing Company, St. Petersburg, Florida. 99 pp. With distributions including Florida Keys (il- lustrated by poor black-and-white line draw- ings and color photos): Anomia simplex, Chione а га, Noetia ponderosa, Pseudocyrena floridana. *SIEKMAN, L., 1981, Handbook of shells, rev. ed. Great Outdoors Publishing Company, St. Petersburg, Florida. 46 [+ ii] pp. With distribution including Key West: Noetia ponderosa. SIEKMAN, L. [with color photographs by R. VAN DE GOHM], 1982, Handbook of shells — sea shells of the Gulf and Atlantic Coast, rev. ed. Great Outdoors Publishing Company, St. Petersburg, Florida. 48 pp., 16 color pls. CRITICAL CATALOG AND BIBLIOGRAPHY DE) With distribution including Key West: Noetia ponderosa. SIMONE, L.R.L. 8 A. CHICHVARKHIN, 2004, Comparative morphological study of four species of Barbatia occurring on the south- ern Florida coast (Arcoidea, Arcidae). In: R. BIELER 8 P. M. MIKKELSEN, eds., Bivalve stud- ies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 355-379. From Florida Keys localities: Arca zebra, Barbatia candida, B. cancellaria, B. domingensis, B. tenera [material from this study]. SIMONE, (. R. L. & J. DOUGHERTY, 2004, Anatomy and systematics of northwestern Atlantic Donax (Bivalvia, Veneroidea, Donacidae). In: R. BIELER & P. M. MIKKELSEN, eds., Bivalve studies in the Florida Keys, Proceedings of the International Marine Bi- valve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 459-472. Donax variabilis from the Florida Keys. SIMPSON, С. T., 1887-1889, Contributions to the Mollusca of Florida. Proceedings of the Davenport Academy of Natural Sciences, 5: 45-72, 63-72 [“read 31 December 1986”; exact dates of pages unclear: p. 49 dated 04 November 1887; p. 57 dated 04 February 1889; p. 65 dated 19 February 1889; second р. 63 dated 01 March 1889]. 98 Florida Keys bivalve names (= 86 spe- cies here considered valid) are listed with specific localities: Arca barbadensis var., A. candida, A. deshayesii, A. domingensis, А. fusca, A. gradata, A. imbricata, A. incongrua, A. noae var. americana, A. transversa, Asaphis deflorata, Avicula ala-perdicis, A. radiata, Botula semen, Cardita floridana, Cardium medium, C. petitianum, Chama macerophylla, Choristodon typicum, Coralliophaga hornbeckiana, Corbula swiftiana, Cumingia tellinoides, Cypricardia coralliophaga, Cyrenoida floridana, Cytherea circinata, С. conradina, С. convexa, С. dione, C. hebraea, C. (Trigona) incerta, Diplodonta candeana, D. semiaspera, D. soror, Donax denticulatus, Ervilia concentrica, E. nitens, Heterodonax bimaculatus, Laevicardium serratum, Lasea (sic) rubra, Lepton bowmani, Lima scabra, L. squamosa, L. tenera, Lithodomus bisulcatus, L. forficatus, L. niger, Lucina costata, L. lintea, L. muricata, L. pecten yellow var., L. pennsylvanica (sic), L. quadrisulcata, L. squamosa, L. tigrina, Lutricola gruneri, Macoma anomala, M. fausta, Martesia cuneiformis, Modiolaria cinnamomea, Mytilus exustus, M. lavalleanus, Ostrea parasitica, Pecten antillarum, P. dis- locates (sic), P. hemicyclica, P. imbricatus, P. ornatus, P. ornatus purplish var., Pectunculus castaneus, Periploma angulifera, Perna ephippium, P. obliqua, Petricola divaricata, Pinna carnea, P. muricata, Rocellaria ovata, R. rostrata, Semele obliqua, S. reticulata, Spondylus croceus, S. spathuliferus, Strigilla pisum (sic), Tellina decora, T. decora white var., T. gouldii, T. interrupta, T. lineata, T lineata var. albida, T. mera, T. radiata var., T. tayloriana, Thracia rugosa, Venus beaui, V. granulata, V. listeri, V. mortoni, V. paphia, V. pygmaea, V. pygmaea var. inaequivalvia. “Strigillas, Mytilus exustas, the pernas” are among species considered characteristic of the Florida Keys, scarcely ever found on Florida west coast, largely due to the trajec- tory of the Gulf Stream. The author further noted (р. 47) that “The molluscan fauna of the Bermudas, though these islands lie north of the northern limit of Florida, is much more like that of the lower Keys than that of the west coast.” SIMPSON, С. Т., 1897, The ianthinas. The Nautilus, 10(12): 133-134. From Key West, January 1893: An account of “untold millions of lanthina, which had washed up in the night”. Also collected “bright Tellinas ... along the south shore”. SMITH, J. T., 1991, Cenozoic giant pectinids from California and the Tertiary Caribbean Province: Lyropecten, “Macrochlamis”, Vertipecten, and Nodipecten species. United States Geological Survey Professional Pa- per 1391, vi + 155 pp., 38 pls. Nodipecten fragosus common in the Florida Keys; this paper distinguishes continental М. fragosus from Caribbean N. nodosus. SMITH, M., 1937, East coast marine shells — descriptions of shore mollusks together with many living below tide mark, from Maine to Texas inclusive, especially Florida. Edwards Brothers, Ann Arbor, Michigan. vii + 308 pp., 74 pls. From Florida Keys localities: Antigona listeri, Arca (Acar) adamsi, A. auriculata, A. (Barbatia) barbata, A. (Acar) reticulata, A. transversa, A. (Navicula) umbonata, Barnea truncata, Chama sarda, Chione (Timoclea) granulata, C. intapurpurea, C. pubera, C. (Timoclea) pygmaea, Corbula cymella, Cumingia coarctata, Donax denticulata, D. fossor, Dosinia concentrica, Ensis directus, Ervilia concentrica, E. nitens, Glycymeris 580 pectinatus, Lucina (Cavilucina, Lucinisca) muricata, Parastarte triquetra, Pecten (Lyropecten) antillarum, P. (Chlamys) imbricatus, P. (Aequipecten, Plagioctenium) nucleus, Pedalion alata, P. listeri, Periploma angulifera, Petricola lapicida, Pinna carnea, Polymesoda floridana, Semele bellastriata, Spondylus americanus, Strigilla flexuosa, S. pisiformis, Tellina (Acropagia, Eurytellina) angulosa, T. (Angulus, Scissula) candeana, T. (Acropagia, Cyclotellina) fausta, T. (An- gulus, Scissula) iris, Tivela mactroides, Transennella conradina, Тидопосага!а (Americardia) medium. SMITH, M. [together with two articles by Dr. JOSHUA L. BAILY], 1940, World-wide sea shells — illustrations, geographical range and other data covering more than sixteen hun- dred species and sub-species of molluscs. Tropical Photographic Laboratory, Lantana, Florida. xviii + 139 pp. With distributions including Florida Keys (or specific localities): Arca auriculata, Barnea truncata, Donax fossor, Ensis directus, Lucina muricatus, Tivela mactroides. SMITH, M., 1945, East coast marine shells — descriptions of shore mollusks together with many living below tide mark, from Maine to Texas inclusive, especially Florida, 3"* ed. Edwards Brothers, Ann Arbor, Michigan. vii + 314 pp., 77 pls. From Florida Keys localities: Arca (Acar) adamsi, A. auriculata, A. (Barbatia) barbata, A. (Acar) reticulata, A. transversa, A. (Nav- icula) umbonata, Barnea truncata, Chama sarda, Chione (Timoclea) granulata, C. intapurpurea, C. pubera, C. (Timoclea) pygmaea, Corbula cymella, Cumingia coarctata, Donax denticulatus, D. fossor, Dosinia concentrica, Ensis directus, Ervilia concentrica, E. nitens, Glycymeris pectinatus, Lucina (Cavilucina, Lucinisca) muricata, Parastarte triquetra, Pecten (Lyropecten) antillarum, P. (Chlamys) imbricatus, P. mildredae, P. (Aquipecten, Plagioctenium) nucleus, Pedalion alata, P. listeri, Periploma angulifera, Petricola lapicida, Pinna carnea, Polymesoda floridana, Pseudochama тегае, Semele bellastriata, Spondylus americanus, Strigilla pisiformis, Tellina (Acropagia, Eurytellina) angulosa, T. (Angulus, Scissula) candeana, T. (Acropagia, Cyclotellina) fausta, Т. (Angulus, Scissula) iris, Tivela mactroides, Transennella conradina, Trigonocardia (Americardia) medium. SMITH, M., 1961, Universal shells — marine- fresh water-land. Alpine Press, Asheville, MIKKELSEN & BIELER North Carolina. 254 pp. + unnumbered color pls. From Florida Keys: Eucrassatella floridana. *SOLEM, A., 1955, Living species of the pele- cypod family Trapeziidae. Proceedings of the Malacological Society of London, 31 (2 “1954”): 64-84, pls. 5-7. Coralliophaga coralliophaga, with an exten- sive synonymy and a distribution map show- ing records in the Lower Keys and Dry Tortugas. Plate 6, figs. 14-15, shows a pho- tograph of a specimen from “Solem Coll. Garden Keys, Dry Tortugas” (verified as FMNH 99831). STEPHENSON, T. A. & A. STEPHENSON, 1950, Life between tide-marks in North America. |. The Florida Keys. The Journal of Ecology, 38: 354—402, pls. 9-15. Clearly datable, well-identified material, based on fieldwork in 1947-48. Mostly cov- ering region from Key Largo to Key West (“North of Key Largo we visited two keys which can only be reached by boat”; p. 362). Areas visited/collected: Plantation Key, Crawl Key, Vaca Key, West Summerland Key, sea-wall at Key West, Little Duck Key, points of the coast of Key Largo, Knights Key, Pigeon Key, Missouri Key, Ohio Key, East Summerland Key and Big Pine Key. Detailed locality informantion given on p. 362. Mol- lusks were identified by Clench, Keen, Test, Salisbury, Hubendick, A.G. Smith, Hertlein, and F.M. Bayer. Includes detailed discussion of zonation patterns and individual habitats. From the Florida Keys (some with specific localities): Arca barbata, A. occidentalis, A. umbonata, Isognomon (Pedalion) alata, |. chemnitziana, Mytilus (Brachidontes) exustus. STEVENSON, G. B., 1970, Keyguide to Key West and the Florida Keys. Banyan Books, Miami, Florida. 64 pp. Identifiable species from line drawings are labelled with common names only: Anadara notabilis (ark shell), Arca zebra (turkey wing), Chama sp. (jewelbox), Codakia orbicularis (lucine), Dinocardium sp. (great heart cockle), Mytilidae (mussel), Pinnidae (pen shell), Pteria colymbus (Atlantic wing oys- ter), Teredinidae (shipworm). STEVENSON, G. B., 1993, Keyguide to the Florida Keys and Key West. Blue Water Pub- lishing, Key Largo, Florida. 64 pp. Identifiable species from line drawings la- belled with common names only: Anadara notabilis (ark shell), Arca zebra (turkey wing), Chama sp. (jewelbox), Codakia orbicularis CRITICAL CATALOG AND BIBLIOGRAPHY 581 (lucine), Dinocardium sp. (great heart cockle), Mytilidae (mussel), Pinnidae (pen shell), Pteria colymbus (Atlantic wing oys- ter), Teredinidae (shipworm). SSTUARDO, J. R., 1968, On the phylogeny, taxonomy and distribution of the Limidae (Mollusca: Bivalvia). Ph.D. dissertation, Harvard University, Cambridge, Massachu- setts. 327 pp., 37 pls., 24 maps, 44 figs. Two new species are described from the Florida Keys, although never formally pub- lished: Limaria (Limatulella) sp., Limea (Limea) sp. STUARDO, J. [R.], 1982, A new species of Ctenoides from the central Atlantic (Bivalvia: Limidae). Boletin de la Sociedad de Biologia de Concepción, Chile, 53: 145-149. Ctenoides sanctipauli n.sp., described in part from Sand Key Reef. S*SUNDERLAND, K., 1988, Exploring Pickles Reef. American Conchologist, 16(3): 12-13. From south end of Key Largo (with black- and-white photographs): Nodipecten nodosus, Spondylus americanus. S*SUNDERLAND, K. & M. CAHILL, 1990, Caribbean Pectinidae and Propeamussidae. American Conchologist, 18(2): 14-15. From the Keys (with black-and-white photo- graph): Aequipecten acanthodes. SUTTY, L., 1990, Seashells of the Caribbean. MacMillan Press Ltd., London, etc. vi + 106 pp. With distribution including Florida Keys (or specific localities): Chione paphia. SWEENEY, М. J. & М. С. HARASEWYCH, 1999, Harald А. Rehder (1907-1996): bio- graphical sketch and malacological contri- butions. The Nautilus, 113(4): 127-150. Rehder's taxa originally described from Florida Keys localities: Anadara springeri, Asthenothaerus balesi, Cumingia tellinoides vanhyningi. [TABB, D. С. & К. В. MANNING, 1961, А check- list of the flora and fauna of northern Florida Bay and adjacent brackish waters of the Florida mainland collected during the period July, 1957 through September, 1960. Bulle- tin of Marine Science of the Gulf and Carib- bean, 11(4): 552-649. Although frequently cited as a source of Florida Keys biotic records, it is here ex- cluded as outside our defined area, which extends roughly half-way between the Florida Keys island chain and the tip of pen- insular Florida. The southernmost sampling site, Sandy Key Basin, is north of this limit. The list includes species more typical of the brackish Everglades fringe [e. g., Tagelus plebeius, Crassostrea virginica] than of the more saline, yet still estuarine, Florida Bay. TAYLOR, J. D., Е. GLOVER, M. PEHARDA, С. BIGATTI & А. BALL, 2004, Extraordinary flexible shell sculpture: the structure and for- mation of calcified periostracal lamellae in Lucina pensylvanica (Bivalvia: Lucinidae). In: В. BIELER & P. M. MIKKELSEN, eds., Bivalve stud- ies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 277-294. Lucina pensylvanica [material from this study]. STEARE, М. M., 1949, The Key Largo trip. The American Malacological Union, News Bulle- tin and Annual Report 1949, pp. 16-17. Field trip “near an inlet” [called “a shoal at the north end of Key Largo” in annual meet- ing report on p. 15] during 1949 annual meet- ing in Miami, 19 June 1949: Arca barbata. STESKEY, M. C., ed., 1969, American Mala- cological Union thirty-fifth annual meeting. The American Malacological Union, Inc., Annual Reports for 1969, Bulletin 36: 1-2. On display throughout the meeting were a dozen specimens of Spondylus americanus from the steel hull of an experimental Navy ship, sunk off Key West in 165 ft of water. STHEROUX, Б. В. & К. L. WIGLEY, 1983, Dis- tribution and abundance ofeast coast bivalve mollusks based on specimens in the National Marine Fisheries Service Woods Hole Col- lection. NOAA Technical Report NMFS SSRF-768, 174 pp. From distribution maps of the east coast of the US (extending to 82°W, approximately midway between Key West and Marquesas Keys): Abra sp., Astarte crenata subequilatera, Cuspidariidae, Cyclocardia borealis, Glycymeris pectinata, Glycymeris sp., Limopsidae, Nemocardium peramabile, Nuculana carpenteri, Nuculana sp., Nuculanidae, Pectinidae, Periglypta listeri, Pitar sp., Pleuromeris tridentata, Tellina sp. THIELE, J., 1910, Molluskenfauna West- indiens. Zoologische Jahrbúcher, Suppl. 11: 109-132, pl. 9. Pre-1910 data for mollusks from the Dry Tortugas: Arca listeri, A. plicata, A. umbonata, Cardium mortoni, Chama macrophylla, Glycymeris pectinatus, Lima lima, L. scabra, Lithophaga nigra, Melina lata, M. listeri, Modiolaria lateralis, Pecten (Chlamys) imbricatus, P. (C.) pusio, Pinna carnea, Репа radiata. 582 STHOMAS, БК. D. K., 1970, Functional mor- phology, ecology and evolution in the genus Glycymeris. Ph.D. Dissertation, Harvard University, Cambridge, Massachusetts. xxii + 397 DD. 11 pls: Locality data are provided for living and dead Glycymeris pectinata (supplementing those provided in the published version; Thomas 1975) and С. decussata from the Middle and Lower Florida Keys. THOMAS, R. D. K., 1975, Functional morphol- ogy, ecology, and evolutionary conservatism in the Glycymerididae (Bivalvia). Paleontol- ogy, 18(2): 217-254, pl. 38. [published ver- sion of Thomas, 1970] The distribution of living and dead Glycymeris pectinata [assumed 1960s] is indicated on a map of the Lower and Middle Keys, with symbols indicating relative abun- dance. Occasional specimens were widely distributed on offshore, unstable, poorly sorted, skeletal sands in 2—5 т; the species was more common on very shallow subtidal gravel banks and most abundant in sheltered bayside bays, in quiet water at 1—4 т. THOMPSON, А. R., Р. L. MCTINTY & Т. L. МСТИМТУ, 1951, Dredging from the cruiser Triton. The Nautilus, 65(2): 37-43. General account of dredging for mollusks from the Triton which included the Florida Keys. |ТКАРРЕ, С. А. & С. L. BREWSTER- WINGARD, 2001, Molluscan fauna from Core 25B, Whipray Basin, Central Florida Bay, Everglades National Park. United States Geological Survey Open-File Report 01-143, 21 pp. Electronic version available at http://pubs.usgs.gov/of/of01-143/; last accessed 12 September 2003. This work is here excluded as outside our defined area, which extends roughly half-way between the Florida Keys island chain and the tip of peninsular Florida. Whipray Basin is north of this limit. S*TREMOR, М. E., JR., 1998, A shelling trip to Key West. Shell-o-gram [Jacksonville Shell Club, Jacksonville, Florida], 39(5): 7-9. An account of a shelling trip with Peggy Wil- liams. From snorkeling at 3 ft in the back country, north of Key West: Codakia orbicu- laris, Linga pensylvanica, Lithopoma americanum, Periglypta listeri. West of Key West: Tellina fausta, T. radiala (sic). Origi- nally published in Tidelines [St. Petersburg Shell Club], September 1998 [not seen]. TRYON, С. W., JR., 1873, American marine conchology: descriptions of the shells of the MIKKELSEN & BIELER Atlantic coast of the United States from Maine to Florida. G. W. Tryon Jr., Philadel- phia, Pennsylvania. 208 pp., 44 pls. From Florida Keys localities: Cardita (Carditamera) floridana. TURGEON, D: D., J. Е. QUINN, JR., A. Е. ВОСАМ, E. V. COAN, Е. С. НОСНВЕКС, W. С. LYONS, Р. М. MIKKELSEN, R. J. NEVES, C.F.E.ROPER, G. ROSENBERG, В. КОТЫ, A. SCHELTEMA, F. G. THOMPSON, M. VECCHIONE & J. D. WILLIAMS, 1998, Com- mon and scientific names of aquatic inver- tebrates from the United States and Canada: mollusks, 2" ed. American Fisheries Soci- ety, Special Publication 26, Bethesda, Mary- land. 526 pp. + CD. From the Florida Keys: Argopecten irradians taylorae. *TURNER, К. D., 1955, The family Pholadidae in the western Atlantic and the eastern Pa- cific. Part | - Martesiinae, Jouannetiinae and Xylophaginae. Johnsonia, 3(34): 65-160. Museum lots (ANSP, Charleston Museum, USNM) from the Florida Keys cited for: Martesia cuneiformis, M. striata. TURNER, R. D., 1966, A survey and illustrated catalogue of the Teredinidae (Mollusca: Bivalvia). Museum of Comparative Zoology, Harvard University, Cambridge, Massa- chusettes. 265 pp., 64 pls. Reiteration of original description of Teredo (Zopoteredo) clappi Bartsch, from Key West. *TURNER, К. D. & К. J. BOSS, 1962, The genus Lithophaga in the western Atlantic. Johnsonia, 4(41): 81-116. Museum lots (MCZ, USNM) from the Florida Keys cited for: Lithophaga antillarum, L. aristata, L. bisulcata, L. nigra. *TURNER, R. D. & J. ROSEWATER, 1958, The family Pinnidae in the western Atlantic. Johnsonia, 3(38): 285-326. Museum lots (ANSP, CAS, MCZ, USNM, collections of Flipse, Kline, Merrill, Schwengel, Schmidt, Yale) from the Florida Keys cited for: Atrina rigida, A. serrata, Pinna carnea. TURNEY, W. J., 1977, Molluscan distribution in Florida Bay. P. 85, in: H. G. MULTER, Field guide to some carbonate rock environments — Florida Keys and western Bahamas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp., 10 maps. No specific molluscan taxa are mentioned, but four subenvironments are described in Florida Bay. “The fauna of Florida Bay 1$ pre- dominantly molluscan, principally gastropods and bivalves which are represented by ap- CRITICAL CATALOG AND BIBLIOGRAPHY 583 proximately 100 genera and 140 recognized species. [...] Molluscan debris comprises 58 to 95 percent of the sediment particles greater than 1/8 тт.” STURNEY, W. J. & В.Е. PERKINS, 1972, Mol- luscan distribution in Florida Bay. Sedimentia Ill: 37 pp. Comparative Sedi- mentology Laboratory, Rosenstiel School of Marine and Atmospheric Science, University of Miami, Florida. These results first appeared as a Shell De- velopment Company report in 1958. Each station included a 2-gallon sample of sedi- ment down to an average depth of seven inches [estimate age of deposition = 300- 400 yrs]; 100 genera and 140 species are tabulated, “based chiefly on death assem- blages”. Identifications were verified using the collections and type specimens at USNM. Florida Bay was divided into four hydrological “subenvironments”: Northern (freshwater from mainland/Everglades), Interior (re- stricted circulation — includes shoreline of Upper Keys above Snake Creek on Planta- tion Key), Atlantic (tidal flow through Florida Keys — the border meets the Keys chain at Plantation Key), and Gulf (continuous with Gulf of Mexico). To assimilate these data into this survey, we drew our mid-line through Florida Bay (as described above), and found that all stations of the Northern subenviron- ment, and part of the stations of the Interior and Gulf subenvironments were outside of our Florida Keys definition; all included sta- tions fell either within our Upper or Middle Keys zones. Collections from each station were qualitatively recorded by the authors as abundant, common, few, or rare, and as in- cluding living specimens or dead only. Recorded from Florida Keys stations: Abra lioica, Anadara notabilis, Anodontia alba, A. philippiana, Anomalocardia auberiana, A. cuneimeris, Arca umbonata, A. zebra, Arcopagia Таиз, Arcopsis adamsi, Atrina sp., Barbatia cancellaria, Barnea costata, Brachidontes exustus, Cardiomya costellata, Cardita floridana, Chione cancellata, C. pygmaea, Codakia orbicularis, C. orbiculata, Corbula sp., Crenella divaricata, Cumingia tellinoides, Diplodonta punctata, Divaricella quadrisulcata, Glycymeris pectinata, Laevi- cardium laevigatum, L. mortoni, Lima pellu- cida, Linga trisulcata, Lucina multilineata, L. pensylvanica, Mactra fragilis, Noetia ponde- rosa, Nucula proxima, Nuculana acuta, Parastarte triquetra, P. sp., Pectinidae, Phacoides nassula, Pinctada radiata, Pitar fulminata, Plicatula gibbosa, Pseudochama radians, Pseudocyrena maritima, Semele proficua, Strigilla mirabilis, Tagelus divisus, Tellidora cristata, Tellina alternata, T. mera, T. similis, T. texana, Trachycardium muricatum, Transennela (sic) sp., Trigonocardia medium, Venericardia tridentata, Volsella americana. SUNITED STATES GEOLOGICAL SURVEY [USGS], 2003 [ver. 24 January 2003], South Florida Ecosystem History Project: Florida Bay. http://sofia.usgs.gov/flaecohist/ floridabay.html; last accessed 12 September 2003. Twenty-eight of 39 USGS field sites in Florida Bay lie within our definition of the Florida Keys; to our knowledge only sites 1-25 have had data published in some form. The online database includes living molluscan species occurrences from 160 specific USGS sta- tions sampled between 1994 and 2001 in Florida Bay: Anodontia alba, Anomalocardia auberiana, Anomia simplex, Arcoidea, Arcopsis adamsi, Argopecten irradians, Brachidontes exustus, Carditamera floridana, Chione cancellata, Codakia spp., Cumingia sp. or spp., Cyrenoida floridana, Diplodonta spp., Laevicardium тойоту, Limaria sp. cf. L. pellucida, Lucina pectinata, Lucinidae, Mactridae, Mercenaria spp., Modiolus squamosus, Nucula proxima, Ostreidae, Parastarte triquetra, Pinnidae, Pitar simpsoni, Polymesoda maritima, Pteria longisquamosa, Tagelus spp., Tellina sp. or Spp., Trachycardium muricatum, Transennella spp., and unknown pelecypod. Empty shell records dated to the mid-1800s are derived from six sediment cores: An- odontia ?alba, Anomalocardia auberiana, Arcopsis adamsi, Argopecten irradians, Brachidontes exustus, Chione cancellata, Codakia orbicularis, Codakia spp., Cumingia tellinoides, Cyrenoida floridana, Laevi- cardium laevigatum, L. mortoni, L. spp., Lima spp., Limaria sp. cf. L. pellucida, Lucinisca nassula, Mysella planulata, Nucula proxima, Ostreea equestris, Parastarte triquetra, Parvilucina multilineata, pectinid, Pitar simpsoni, Pteria longisquamosa, Semele bellastriata, Tellina spp., Transennella spp., and unidentified pelecypod. VALENTICH-SCOTT, P. 8 G. E. DINESEN, 2004, Rock and coral boring Bivalvia (Mol- lusca) of the Middle Florida Keys, U.S.A. In: R. BIELER 8 P. M. MIKKELSEN, eds., Bivalve stud- 584 MIKKELSEN & BIELER ies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 339-354. From Florida Keys localities: Botula fusca, Choristodon robustum, С. sp. А, Gastrochaena hians, Lithophaga antillarum, L. aristata, L. bisulcata, Petricola lapicida [material from this study]. VERRILL, А. E., 1882, Catalogue of marine Mollusca added to the fauna of the New England region, during the past ten years. Transactions of the Connecticut Academy, 5: 447-587, pls. 42-44, 57, 58. Neaera rostrata from Sand Key as per Dall's Blake report (Dall, 1881). *VILAS, С. М. & М. К. VILAS, 1945, Florida marine shells — a guide to [sic] amateur col- lectors of Florida marine shells which 1$ com- pletely illustrated with colored photo- engravings. Aberdeen Press, Chicago. iv + 152 pp., incl. 12 color pls. With distribution including Florida Keys (or specific localities): Barnea truncata, Codakia orbicularis, Pecten muscosus, Pedalion alata, Pinctada radiata, Semele bellastriata, Spondylus americanus, Tellina radiata, T. r. unimaculata. VILAS, С. М. & М. К. VILAS, 1970, Florida marine shells. Charles E. Tuttle, Rutland, Vermont. 170 pp., 14 pls. With distribution including Florida Keys (or specific localities): Aequipecten muscosus, Barnea truncata, Codakia orbicularis, Pedalion alata, Pinctada imbricata, Semele bellastriata, Spondylus americanus, Tellina radiata, T. r. var. unimaculata. [BARRY A.] VITTOR 8 ASSOCIATES, INC., 1997a [March], Florida Bay Benthic Commu- nity Assessment. Report to NOAA, Silver Spring, Maryland, 42 pp. From bayside localities in Upper, Middle and Lower Florida Keys: Lucinidae, Nucula aegeenis, Solemya occidentalis, Tellinidae. [BARRY A.] VITTOR 8 ASSOCIATES, INC., 1997b [June], Florida Bay and adjacent wa- ters benthic community assessment. Report to NOAA, Silver Spring, Maryland, unpaginated. From bayside localities in Lower Florida Keys: Crassinella martinicensis, Diplodonta semiaspera, Tellina sp. [BARRY A.] VITTOR 8 ASSOCIATES, INC., 1997c [August], Biscayne Bay, Florida benthic community assessment. Report to NOAA, Silver Spring, Maryland, unpagi- nated. From Card Sound: Americardia media, Arcopsis adamsi, Brachidontes exustus, Chione cancellata, Diplodonta semiaspera, Laevicardium топот! Lucina multilineata, L. radians, L. sp., Lucinidae, Modiolus modiolus squamosus, Musculus lateralis, Mysella planulata, Mytilidae, Nucula aegensis, Pelecypoda, Pitar sp., Semelidae, Tellina similis, T. sybaritica, T. sp., Tellinidae, Transennella conradina, Veneridae. [BARRY A.] VITTOR 8 ASSOCIATES, INC., 1998 [April], Florida Keys to Dry Tortugas benthic community assessment. Report to NOAA, Silver Spring, Maryland, un- paginated. From localities from Lower Florida Keys to Dry Tortugas: Abra aequalis, Amygdalum sagittatum, Anadara sp., Anomalocardia auberiana, Arca zebra, Arcidae, Arcinella cornuta, Argopecten gibbus, Asthenothaerus hemphilli, Bivalvia, Bushia sp., Carditidae, Cardiomya perrostrata, Cardiomya sp., Chama congregata, Chione cancellata, C. grus, C. sp., Codakia orbicularis, C. sp., Corbula contracta, Corbulidae, Crassatellidae, Crassinella lunulata, С. martinicensis, C. sp., Crenella divaricata, Cumingia coarctata, С. tellinoides, Diplodonta semiaspera, D. sp., Dosinia dis- cus, Ervilia concentrica, Gastrochaena hians, Glans dominguensis, Glycymerididae, Glycymeris pectinata, G. sp., Gouldia cerina, Hiatella arctica, Lima pellucida, Linga ami- antus, L. pensylvanica, L. sp., Lioberus castaneus, Lucina blanda, L. multilineata, L. nassula, L. sp., Lucinidae, Lyonsia beana, L. hyalina floridana, Macoma brevifrons, M. sp., Macrocallista maculata, Mactridae, Modiolus modiolus squamosus, Montacutidae, Musculus lateralis, Mytilidae, Naeromya floridana, Nucula aegeenis, Nuculana acuta, N. concentrica, М. sp., Pandora sp., Pectinidae, Pitar fulminatus, P. simpsoni, P. sp., Pleuromeris tridentata, Semele bellastriata, S. nuculoides, S. proficua, Semelidae, Solemya occidentalis, S. sp., Tellidora cristata, Tellina aequistriata, T. iris, T. listeri, T. mera, T. similis, T. sybaritica, T. texana, T. versicolor, T. sp., Tellinidae, Thraciidae, Trachycardium egmontianum, Transennella stimpsoni, Ungulinidae, Varicorbula operculata, Veneridae. S[BARRY А.] VITTOR 8 ASSOCIATES, INC., 1999a [June], Florida Keys (including outer Florida Keys National Marine Sanctuary Boundary) — Dry Tortugas benthic commu- CRITICAL CATALOG AND BIBLIOGRAPHY 585 nity assessment, July 1998. Report to NOAA, Silver Spring, Maryland, unpaginated. Twenty stations from off the oceanside ofthe Upper and Lower Keys and Dry Tortugas include collections of: Abra aequalis, Amygdalum sagittatum, A. sp., Anadara notabilis, Arca zebra, Argopecten sp., Asthenothaerus hemphilli, Bivalvia, Cardiidae, Cardiomya perrostrata, Carditidae, Chione grus, Corbula contracta, Crassinella lunulata, C. martini- censis, Crenella divaricata, Dosinia discus, Ervilia concentrica, Gastrochaena hians, Glans dominguensis, Glycymeris decussata, Gouldia cerina, Hiatella arctica, Laevicardium laevigatum, Lima locklini, L. pellucida, L. sp., Lucina multilineata, L. muricata, L. nassula, L. pectinata, L. radians, L. sp., Lucinidae, Lyonsia hyalina floridana, Macoma tenta, Macrocallista maculata, M. nimbosa, Montacutidae, Musculus lateralis, Musculus sp., Mysella sp., Nucula aegeenis (sic), Nuculana acuta, N. concentrica, Pandora arenosa, P. sp., Pitar simpsoni, P. sp., Pleuromeris tridentata, Semele bellastriata, S. nuculoides, S. proficua, S. purpurascens, S. sp., Semelidae, Tellina sybaritica, Т. texana, T. sp., Tellinidae, Thraciidae, Varicorbula operculata, Veneridae. S[BARRY A.] VITTOR 8 ASSOCIATES, INC., 1999b [November], Florida Keys National Marine Sanctuary benthic community as- sessment, August-September 1998. Report to NOAA, Silver Spring, Maryland, unpaginated. Thirteen stations from off the oceanside of the Upper, Middle and Lower Keys include collections of: Abra lioica, А. sp., Anadara notabilis, Arca zebra, Arcidae, Asthenothaerus hemphilli, A. sp., Bivalvia, Bushia elegans, Cardiidae, Carditidae, Chione cancellata, C. grus, C. sp., Codakia orbicularis, Crassinella lunulata, C. sp., Crenella divaricata, Cumingia tellinoides, Diplodonta punctata, D. sp., Dosinia discus, Ervilia concentrica, E. nitens, E. sp., Glans dominguensis, Glycymerididae, Gouldia cerina, Hiatella arctica, Laevicardium laevigatum, L. sybariticum, L. sp., Lima pel- lucida, L. sp., Limidae, Lioberus castaneus, Lucina muricata, L. nassula, L. radians, L. sp., Lucinidae, Macoma sp., Modiolus modiolus squamosus, Montacutidae, Mus- culus lateralis, Mytilidae, Neaeromya floridana, Nucula aegeenis (sic), Pectinidae, Pitar fulminatus, P. simpsoni, P. sp., Pleuromeris tridentata, Pteriidae, Semele bellastriata, $. nuculoides, $5. sp., Semelidae, Tagelus divisus, Tellina gouldii, Т. iris, T. тега, T. similis, T. tampaensis, T. tenella, T. sp., Tellinidae, Trachycardium muricatum, T. sp., Veneridae. Five stations (98FKNMSO09, -10, -13, -14, -15) had no mollusks listed; two additional stations (98FKNMS01, -02) lie north of Keys north- eastern boundary, as defined by this study. *VOKES, Н. E., 1969, The anadarid subge- nus Caloosarca in the western Atlantic re- gion. Tulane Studies in Geology and Paleontology, 7(1): 1-40, incl. pls. 1-6. Anadara (Caloosarca) notabilis from Long Key. SVOSS, С. L., 1948, A trip to the outer reef. Shell Notes, 2(5): 58-72. Collecting with Frank Lyman and Don Moore, from reefs out of Garden Cove [Key Largo]: Pecten imbricatus, P. sentis. SVOSS, G. L., 1949, Notes from the log of the Junonia. Shell Notes, 2(7-9): 112-120. From reefs out of Garden Cove [Key Largo]: Arca sp., Glycimeris americana lineata, Lima spp., Pecten antillarum, P. sentis. The au- thors notes of Conch Reef: “We had very little luck here ... as the reef in this section is dead as far as molluscan life is concerned.” SVOSS, С. L., 1983, Final report: an environ- mental assessment of the Key Largo Na- tional Marine Sanctuary. Rosenstiel School of Marine and Atmospheric Science, Univer- sity of Miami. 517 pp. Requested by the U.S. Office of Coastal Zone Management, “to provide a base-line study of the fauna and flora for use in man- agement, planning and future studies”. Sur- veying 10 reef sites and 7 shallow grass and hard-bottom sites shoreward of 18 m isobath, using rotenone, visual transect lines, and collection of unknown organisms. Identifications of mollusks were achieved using Voss (1980). In sand areas, the sand was removed to a depth of 10 cm in 4 т? quadrats, then screened; all specimens 1 cm or longer in length were retained for identification. Voucher specimens are de- posited at the Rosenstiel School of Marine and Atmospheric Sciences. The molluscan species list includes Americardia media, Anadara notabilis, Arca imbricata, A. zebra, Argopecten gibbus, Barbatia cancellaria, В. candida, B. domingensis, Chama macero- phylla, Chione cancellata, С. paphia, Codakia orbicularis, Diplodonta punctata, Glycimeris 586 (sic) pectinata, Laevicardium laevigatum, Lima lima, L. pellucida, Linga pensylvanica, Lithophaga sp., Lopha frons, Lyropecten antillarum, Modiolus americanus, M. modio- lus squamosus, Musculus lateralis, Ostrea frons, Рарупаеа semisulcata, Pecten sp., Periglypta listeri, Pinctada radiata, Plicatula gibbosa, Tagelus sp., Tellina listeri. SVOSS, С. L., М. А. VOSS, А. Y. CANTILLO 8 М. J. BELLO, 1983, An environmental as- sessment of the John Pennekamp Coral Reef State Park and the Key Largo Coral Reef Marine Sanctuary. Joint NOAA/Univer- sity of Miami Report. NOAA Technical Memo- randum NOS NCCOS CCMA 161. NOAA LISD Current References 2002-6. Rosenstiel School of Marine and Atmospheric Science, University of Miami, Miami, Florida. 452 pp. 2002 edited version available at http:// www.aoml.noaa.gov/general/lib/ cedardoc.html; last accessed 04 April 2003. Qualitative and quantitative studies were conducted to determine the state of marine communities after approximately 10 years post-establishment of John Pennekamp Coral Reef State Park and Key Largo Coral Reef Marine Sanctuary. Transects of 400 m length (3 per site) perpendicular to the reef or shoreline were surveyed by scuba divers. Some voucher specimens were located by PMM in February 2003 in the Invertebrate Museum, Rosenstiel School of Marine and Atmospheric Science. Americardia media, Anadara floridana, A. notabilis, Arca imbricata, A. zebra, Arca sp., ark shells, Argopecten gibbus, Barbatia cancellaria, B. candida, B. domingensis, bivalve, Chama congregata, C. macerophylla, Chione cancellata, C. paphia, Codakia orbicular (sic), C. orbicularis, Diplodonta punctata, Glycymeris pectinata, Glycymeris sp., Isognomon sp., Laevicardium laevigatum, Lima lima, L. pellucida, L. scabra, Lima spp. (with “red and yellow tentacles”), Linga pensylvanica, Lithophaga antillarum, L. ni- gra, Lithophaga spp., Lopha frons, Lyro- pecten antillarum, Modiolus americanus, M. modiolus squamosus, Musculus lateralis, Ostrea frons, O. (Lopha) frons, Papyridea semisulcata, Pecten antillarum, P. sp., Periglyphus (sic) listeri, Periglypta listeri, Pinctada radiata, Plicatula gibbosa, Tagelus sp., Tellina listeri. |VOSS, С. |. 8 М. А. VOSS, 1955, An ecologi- cal survey of Soldier Key, Biscayne Bay, Florida. Bulletin of Marine Science of the Gulf and Caribbean, 5(3): 203-229. MIKKELSEN 8 BIELER Although cited as a source of Florida Keys biotic records (Levy et al., 1996), this work is here excluded as being outside our de- fined area, which extends south of Broad Creek at the northern end of Key Largo. Soldier Key is north of this point, at the east- ern extent of Biscayne Bay. WAGNER, Б. J. L. & R. T. ABBOTT, 1990, Wagner and Abbott's world size records. Standard Catalog of Shells, Suppl. 4. Ameri- can Malacologists, Melbourne, Florida. ii + 80-001-80-080. World size records from Florida Keys: Aequipecten acanthodes, A. muscosus, Arca zebra, Chlamys sentis, Isognomon alatus, Lyropecten antillarum, Mercenaria campechiensis, Psammotreta intastriata, Tellina fausta, T. listeri, T. magna, T. radiata, Т. r. unimaculata. WALLER, T. R., 1969, The evolution of the Argopecten gibbus stock (Mollusca: Bivalvia), with emphasis on the Tertiary and Quaternary species of eastern North America. The Paleontological Society, Mem- oir 3. Journal of Paleontology, 43(Suppl. to no. 5): v + 125 pp., 3 fold-outs. From Florida Keys or Keys localities: Argopecten gibbus, A. irradians concen- tricus, A. nucleus. *WALLER, T. R., 1993, The evolution of “Chlamys” (Mollusca: Bivalvia: Pectinidae) in the tropical western Atlantic and eastern Pacific. American Malacological Bulletin, 10(2): 195-249. From Florida Keys localities: Caribachlamys imbricata, C. mildredae, C. ornata, C. sentis. “WALLER, Т. R. & I. G MACINTYRE, 1982, Larval settlement behavior and shell morphol- ogy of Malleus candeanus (d'Orbigny) (Mol- lusca: Bivalvia). Pp. 489-497, in: K. RUTZLER 8.1. С. MACINTYRE, eds., The Atlantic barrier reef ecosystem at Carrie Bow Cay, Belize, 1. Structure & Communities, Smithsonian Con- tributions to Marine Science 12, 539 pp. Malleus candeanus from the Dry Tortugas. *WARMKE, С. L. 8 К. T. ABBOTT, 1961, Car- ibbean seashells. Dover Publications, New York. 348 pp., 44 pls. With distribution including Florida Keys (or specific localities): Aequipecten lineolaris, Antigona rigida, Chama sarda, Chione paphia, Cumingia antillarum, Isognomon bi- color, Phacoides muricata, Strigilla pisiformis, Tellina punicea, Trachycardium magnum, Transennella cubaniana. WATTERS, G. T., 2002, The status and iden- tity of Papyridea soleniformis (Bruguière, CRITICAL CATALOG AND BIBLIOGRAPHY 987 1789) (Bivalvia: Cardiidae). The Nautilus, 116(4): 118-128. А systematic study of Papyridea soleniformis reveals involvement of a second species, P lata. Florida Keys localities are cited for each, from University of Florida, University of Michigan, ANSP, and H. G. Lee collections. WEBB, W. F., 1937, Shells and other inverte- brates of the United States. Privately pub- lished, Rochester, New York. xiv + 80 pp. From Florida Keys (or specific localities): Cytherea hebraea, Dosinia discus, Mytilus perna, Ostrea foliata, Transenella (sic) conradina. WEBB, W. F., 1939, A Catalogue of Recent Mollusca for sale by Walter F Webb. W. Е. Webb, Rochester, New York. [iv] + 148 pp. incl. 34 pls. From Florida Keys (or specific localities): Cytherea hebraea, Mytilus perna, Transenella (sic) conradina. WEBB, W. F., 1942, United States Mollusca: a descriptive manual of many of the marine, land and fresh water shells of North America, north of Mexico, 1* ed. Privately published, Rochester, New York. 220 pp., 63 pls. “The Pourtales Plateau which lies just off the S. E. Florida coast at the Tropic of Cancer fairly swarms with oceanic life. The Gulf Stream sweeps over it constantly, bringing warm water literally swarming with minute life. The larger Pelagic life like Janthinas and the smaller Hyalaeas, Creseis, Cuverias and others sweep along by the millions. The minute pelagic animals are constantly dying, and there is always a gengle rain of food falling over the bottom of the ocean. A veri- table free soup kitchen for the myriads of shell life. The food literally drops into their mouth without any effort to obtain same. It is no wonder that the dredge bridge brings up unbelievable quantities of shells which are seldom found on the shore lines.” (p. 37). From Florida Keys (or specific localities): Chama variegata, Chione latilirata, C. paphia, Cytherea hebraea, Dosinia discus, Glycimeris (sic) americana, Lima fragilis, L. tenera, Lithophaga antillarum, Lucina pensylvanica, Macoma constricta, Modiola duplicata, Mytilus perna, Ostrea foliata, Paphridea (sic) spinosum, Pecten antillarum, P. sentis, Pitar fulminata, Plicatula mantilla, Strigilla pisiformis, Tellina candeana, Transenella (sic) conradina. WEBB, W. F., 1951, United States Mollusca — a descriptive manual of many of the marine, land and fresh water shells of North America, north of Mexico. Privately published, St. Pe- tersburg, Florida. 224 pp., incl. 67 pls. With distribution including Florida Keys (or specific localities): Arca candida, Chama variegata, Chione latilirata, С. paphia, Cyrena floridana, Cytherea hebraea, Dosinia discus, Glycimeris americana, Lima fragilis, L. tenera, Lithophaga antilarum (sic), Lucina pensylvanica, Macoma constricta, Modiola duplicata, Mytilus perna, Ostrea foliata, Paphridea spinosum, Pecten antillarum, Р imbricatus, P. muscosus, P. nucleus, P. ornatus, P. sentis, Pitar fulminata, Plicatula mantilla, Strigilla pisiformis, Tellina braziliana, T. candeana, T. crystallina, T. laevigata, Transenella (sic) conradina. WEBSTER, R., 1978, Gems: their sources, descriptions and identification, 3% ed. Ar- chon Books, Handon, Connecticut. 931 pp. Reference to pearling (undoubtedly from Pinctada imbricata) is made: “The Gulf of Mexico is often mentioned as an area for pearl fishery, and admittedly there is an un- important fishery off the Marquesas ...”. WHEATLEY, C. M., 1845, Catalogue of the shells of the United States, with their locali- ties, 2" ed. John Towers, New York. 35 pp. Chama lazerus from Key West. SWIENER, J., 1988b, More Mystery Island ... The Busycon [Broward Shell Club, Ft. Lau- derdale, Florida], 23(10): 2-4. Account of a July 1988 shelling trip to Saw- yer Island, out of Little Torch Key lists only “assorted bi-valves”. SWILLIAMS, P., 1990, Scallops | have known. American Conchologist, 18(2): 3—4. Chlamys sentis, common in shallow reefs in the Keys. *WILLIAMS, W., 1988, Florida’s fabulous sea- shells and other seashore life, 2" ed. World Publications, Tampa, Florida. 112 pp. From Florida Keys (with color photographs labelled by common names): Chlamys imbricatus, C. sentis, Codakia orbicularis, Ctenoides floridana [photographed, as “file clam”], Lima lima, Lyropecten nodosus, Spondylus americanus. Also notes on com- mercial use of bivalve species. SWINGARD, С. L., $. ISHMAN, T. CRONIN, L. E. EDWARDS, D. A. WILLARD & R. B. HALLEY, 1995, Preliminary analysis of down- core biotic assemblages: Bob Allen Keys, Everglades National Park, Florida Bay. United States Geological Survey Open-File Report 95-628, 35 pp. Electronic version available at http://131.247.143.93/publications/ofr/95-628/; last accessed 12 September 2003. 588 From Core 6A, a sediment core dated to the mid-1800s, at the Bob Allen Keys, Florida Bay: Anomalocardia cuneimeris, Arcopsis adamsi, Brachiodontes (sic) sp., Chione cancellata, Cumingia tellinoidea, Laevi- cardium spp., Lima sp., Mysella sp., Nucula proxima, Parastarte triquetra, pectinid, Pinctada radiata, Pitar sp., Semele bellastriata, Tellina spp., Transenella (sic) spp., unidentified pelecypod fragments. A summary of these data was published by Brewster-Wingard et al. (1998) and Brewster-Wingard & Ishman (1999). WOODS, E., 1970, June Keys field trip. Seafari [Palm Beach County Shell Club Newsletter], 12(10): 2-4. From Missouri Key [June 1970], among rubble, seagrass, and mud: Antigona listeri, Arca imbricata, A. zebra, Arcopagia fausta, arks, Codakia orbicularis, Glycymeris pectinata, Laevicardium laevigatum, Lucina pensylvanica, Modiolus americanus, Pinctada radiata; from along shore: Pecten antillarum; from an evening trip to an un- specified area: Arcopsis adamsi, Lima scabra tenera. “WOODS, E., 1971, Grassy Key enclosure — gone! Seafari [Palm Beach County Shell Club Newsletter], 13(1): 1-2. The author is lamenting closure of a “man- made pool” on the Gulf side of Grassy Key that was formerly a popular shelling site. From among rocks at that site are Barbatia cancellaria, Lima pellucida, Lima scabra tenera, and “little mussels and oysters”. WORK, R.C., 1969, Systematics, ecology, and distribution of the mollusks of Los Roques, Venezuela. Bulletin of Marine Science, 19(3): 614-711. From Florida Keys localities (citing literature and author's personal records): Americardia media, Arca imbricata, Arca zebra, Barbatia cancellaria, Barbatia candida, Brachidontes exustus, Chama congregata, С. macerophylla, Chlamys imbricata, С. ornata, Codakia orbicularis, Isognomon alatus, |. radiatus, Laevicardium laevigatum, Lima lima, L. pellucida, L. scabra, L. scabra form tenera, Modiolus americanus, Pinctada imbricata, Pinna carnea, Pteria colymbus, Spondylus americanus, $. ictericus, Tellina fausta, T. laevigata, T. listeri, T. radiata. SZIEMAN, J. C., 1982, The ecology of the seagrasses of south Florida: a community profile. United States Fish and Wildlife Ser- vices, FWS/OBS - 82/25, 158 pp. Reprinted September 1985. MIKKELSEN 8 BIELER Referring to a paper by Chan (1977), report- ing on the effects of a 1975 tanker discharge SW of the Marqueses: “The author attributed mass mortalities of the pearl oyster (Pinctada radiata) a grass bed inhabitant, to some soluble fraction of petroleum” (p. 88). SZISCHKE, J. A., 1973, An ecological guide to the shallow-water marine communities of Pigeon Key, Florida. St. Olaf College, Northfield, Minnesota. [vi +] 44 pp. The text refers to some molluscan species’ particular zones and habitats. “This list does not include all species present, nor does it include species that are exclusively found in mangrove and coral communities or forms generally restricted to deeper water” (p. 26). The listing indicates habitats for each spe- cies [intertidal, Echinometra zone, loose rock, Alcyonaria-sponge zone, grass beds]. Included are: Americardia media, Anadara notabilis, Antigona listeri, Arca imbricata, A. zebra, Arcopsis adamsi, Atrina rigida, Barbatia cancellaria, Brachidontes exustus, Chione cancellata, Chlamys sentis, Codakia orbicularis, Isognomon alatus, |. bicolor, 1. radiatus, Lima scabra, Lithophaga antillarum, L. nigra, Lucina pensylvanica, Modiolus americanus, Ostrea equestris, Petricola lapicida, Pinctada imbricata, Sanguinolaria sanquinolenta. ZISCHKE, J. A., 1977a, Checklist of macro- flora, invertebrates and fishes of Pigeon Key. Pp. 27-30, in: H. с. MULTER, Field guide to some carbonate rock environments — Florida Keys and western Bahamas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp., 10 maps. From Pigeon Key: Americardia media, Anadara notabilis, Antigona listeri, Arca imbricata, A. zebra, Arcopsis adamsi. Atrina rigida, Barbatia cancellaria, Brachidontes exustus, Chione cancellata, Chlamys sentis, Codakia orbicularis, Isognomon alatus, |. bicolor, I. radiatus, Laevicardium laevigatum, Lima scabra, Lithophaga antillarum, L. nigra, Lucina pensylvanica, Modiolus americanus, Ostrea equestris, Petricola lapicida, Pinctada imbricata, Pteria colymbus, Sanguinolaria sanguinolenta. ZISCHKE, J. A., 1977b, An ecological guide to the shallow-water marine communities of Pigeon Key, Florida. Pp. 23-27, in: H. 6. MULTER, Field guide to some carbonate rock environments — Florida Keys and western Bahamas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp., 10 maps. Mentions mollusks (some with sketched fig- CRITICAL CATALOG AND BIBLIOGRAPHY 589 ures) and other organisms and their occur- rence in the various ecological zones of Pi- geon Key, including: Anadara sp., Arca sp., Arcopsis adamsi, Atrina rigida, Barbatia sp., Brachidontes exustus, Chione cancellata, Isognomon alatus, |. bicolor, Ostrea equestris. ZISCHKE, J. A., 1977c, Some common inver- tebrates of Pigeon Key. Figure A.14, in: H. с. MULTER, Field guide to some carbonate rock environments — Florida Keys and western Bahamas, new ed. Kendall/Hunt Publishing Company, Dubuque, lowa. 415 pp. + 10 maps. Illustrated from Pigeon Key: Anadara notabilis, Antigona listeri, Arca zebra, Arcopsis adamsi, Atrina rigida, Barbatia cancellaria, Brachidontes exustus, Chione cancellata, Chlamys sentis, Codakia orbicu- laris, Isognomon alatus, Lima scabra. CRITICAL CATALOG OF FLORIDA KEYS BIVALVES (Those species listed without references occur in the Florida Keys according to our sur- vey of original and museum collections, but have not been previously recorded as such in the literature.) Anomiidae Anomia simplex Orbigny, 1842: Dall, 1889a, 1903b; Lermond, 1936; Siekman, 1965; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, DT]; USGS, 2003 [UFK, MFK]. Pododesmus rudis (Broderip, 1834): Dall, 1889a [as Placunanomia]; Lermond, 1936 [as Polodesmus (sic) decipiens Philippi, 1837]. Arcidae Edwards, 1970 [LFK; as ark shells]; Woods, 1970 [LFK; as arks]; Voss et al., 1983 [UFK; as ark shells]; Clampit, 1987 [LFK; as ark shells]; Vittor 8 Associates, 1998 [DT], 1999b [MFK, LFK]. Acar domingensis (Lamarck, 1819): Dall, 1883 [LFK; as Arca (Barbatia) dominguensis and misidentified as Arca (Barbatia) gradata Broderip 8 С. В. Sowerby |, 1829, a recog- nized eastern Pacific species], 1885 [LFK; as Arca (Barbatia) dominguensis and as Arca (Barbatia) gradata], 1889а [as Arca (Byssoarca) reticulata auctt. non Gmelin, 1792; also misidentified as Arca (Byssoarca) nodulosa Múller, 1776, a synonym of Barbatia scabra (Poli, 1795), a recognized European species], 1903b [as Arca (Byssoarca) reticulata]; Simpson, 1887- 1889 [DT; as Arca domingensis and as Arca gradata]; Thiele, 1910 [DT; misidentified as Arca plicata Dillwyn, 1817, a recognized Indo-Pacific species]; Lermond, 1936 [as Arca reticulata]; M. Smith, 1937, 1945 [as Arca (Acar) reticulata]; Edwards, 1968b [LFK; as Ваграйа]; Ross, 1969 [MFK; as Barbatia]; Godcharles & Jaap, 1973 [UFK; as Barbatia]; Mikkelsen, 1981 [UFK; as Barbatia (Acar)]; Voss, 1983 [UFK; as Barbatia]; Voss et al., 1983 [UFK; as Barbatia]; Lyons & Quinn, 1995 [as Barbatia]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT; as Barbatia]; Oliver 8 Járnegren, 2004 [LFK]; Simone 8 Chichvarkhin, 2004 [MFK, ЕЕК; as Barbatia]. Anadara baughmani Hertlein, 1951: Rehder 8 Abbott, 1951 [DT; as A. springeri Rehder 8 Abbott, 1951]; Pulley, 1952 [DT]; Sweeney 8 Harasewych, 1999 [DT; as A. springeril; Mikkelsen & Bieler, 2000 [DT]. |Anadara brasiliana (Lamarck, 1819) — see under Scapharca. |Anadara chemnitzii (Philippi, 1851) — see under Scapharca. Anadara floridana (Conrad, 1869): Dall, 1889a [misidentified as Arca (Scapharca) lienosa Say, 1832, which is fossil]; Pearse, 1929 [DT; as Arca “saccharina”, probably a misspell- ing for Arca (Anadara) floridana var. secernenda Lamy, 1907]; Lermond, 1936 [misidentified as Arca secticostata Reeve, 1844, from indeterminate locality]; Magnotte, 1970-1979 [as A. lienosa floridana]; Voss et al., 1983 [UFK]; Mikkelsen & Bieler, 2000 [LK Anadara notabilis (Róding, 1798): Simpson, 1887-1889 [LFK; as Arca deshayesii Hanley, 1843]; Dall, 1889a, 1903b [LFK; misidentified as Arca (Scapharca) auriculata Lamarck, 1819, a recognized Red Sea species], 1898 [LFK; as Scapharca (Scapharca, section Scapharca) auriculata]; Maury, 1920 [LFK; as S. (S.) auriculata], 1925 [as S. (S.) auriculata]; Johnson, 1934 [as Arca auriculata and A. deshayesii]; Lermond, 1936 [as Arca auriculata]; Richards, 1936; M. Smith, 1937, 1940, 1945 [as Arca auriculata]; Eubanks, 1964 [as A. nobilis]; Ross, 1969 [MFK]; Vokes, 1969 [MFK]; 590 MIKKELSEN & BIELER Magnotte, 1970-1979; Stevenson, 1970, 1993 [both as ark shell]; Turney & Perkins, 1972 [UFK, МЕК]; Godcharles & Jaap, 1973 [UFK]; Zischke, 1973, 1977a, с [MFK]; Antonius et al., 1978 [ЕЕК]; Voss, 1983 [ЧЕК]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1999a [ЧЕК], b [ЕЕК]; Mikkelsen & Bieler, 2000 IUEK MEK, ВЕК, DIT: |Ападага ovalis (Bruguière, 1789) — see ип- der Lunarca. Anadara transversa (Say, 1822): Simpson, 1887-1889 [LFK; as Arca]; Dall, 1889a, 1903b [LFK; as Arca (Scapharca)], 1898 [LFK; as Scapharca (Scapharca, section Scapharca)]; Rogers, 1908 [as Arca]; Aldrich 8 Snyder, 1936 [as Arca]; Lermond, 1936 [as Arca]; M. Smith, 1937, 1945 [LFK; as Arca]; Pulley, 1952 [DT]; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, DT]. Anadara sp.: Zischke, 19776 [МЕК]; Schomer & Drew, 1982; Vittor 8 Associates, 1998 [ЕЕК]. Arca imbricata Bruguière, 1789: Simpson, 1887-1889 [ОТ; also as Агса поае var. americana Orbigny, 1846]; Nutting, 1895 [DT; as Arca velata “Sowerby”, [date unknown]. Although the taxonomic status of Sowerby's name could not be verified, Nutting's Tortugas material was later identified as the recognized synonym, A. umbonata Lamarck, 1819, by Dall, 1896a]; Dall, 1889a [as A. (Arca)], 1896a [ОТ; as A. umbonata], 1903b [as A. (Arca)], 1898 [Pleistocene; as A. (Lunarca) umbonata]; Thiele, 1910 [DT; as A. umbonata]; Lermond, 1936 [also as А. umbonata]; М. Smith, 1937 [as A. (Navicula) umbonata], 1945 [as A. (Navicula) umbonata]; Bippus, 1950 [UFK; as A. umbonata]; T. A. Stephenson & A. Stephenson, 1950 [LFK; as A. umbonata]; Kissling, 1965 [LFK], 1977a [UFK, LFK], 1977b [UFK; all as A. umbonata]; Edwards, 1968b [ЕЕК]; Iversen & Roessler, 1969 [UFK; also as A. umbonata]; Jindrich, 1969 [LFK; as A. umbonata]; Lee, 1969 [LFK]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Edwards, 1970 [LFK]; Magnotte, 1970-1979; Woods, 1970 [LFK]; Turney € Perkins, 1972 [UFK, МЕК; as A. umbonata]; Zischke, 1973, 1977a [MFK]; Antonius et al., 1978 [LFK]; Mikkelsen, 1981 [UFK]; Voss, 1983 [UFK]; Voss et al., 1983 [ЧЕК]; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]; Oliver 8 Járnegren, 2004 [LFK]. Arca zebra (Swainson, 1833): Melvill, 1880 [LFK; as Scapharca occidentalis (Philippi, 1847) and misidentified as Arca noae (Linnaeus, 1758), a recognized eastern At- lantic species]; Simpson, 1887-1889 [DT; as var. of А. barbadensis “Petiver” Orbigny, 1846]; Dall, 1889a [as A. (4.) noae], 1896a [LFK; as A. noae], 1898 [Pleistocene; as А. (Lunarca) occidentalis], 1903b [as A. (A.) noae]; Lermond, 1936 [as A. occidentalis]; Lyman, 1947b [UFK; as A. occidentalis]; T. A. Stephenson & A. Stephenson, 1950 [LFK; as A. occidentalis]; Edwards, 19686 [ЕЕК]; lversen & Roessler, 1969 [UFK]; Ross, 1969 [MFK]; Work, 1969 [ЕЕК]; Magnotte, 1970- 1979; Stevenson, 1970, 1993 [both as tur- key wing]; Woods, 1970 [ЕЕК]; Turney € Perkins, 1972 [UFK, MFK]; Godcharles & Jaap, 1973 [UFK]; Shoemaker, 1973; Zischke, 1973, 1977a, c [MFK]; Kissling, 1977a; Goldberg, 1978 [ЕЕК]; Edwards, 1980 [LFK]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Wagner & Abbott, 1990; Andrews, 1994: Lyons & Quinn, 1995; Hutsell et al., 1997; Vittor & Associates, 1998 [DT], 1999a [UFK], b [LFK]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]; Orlin, 2003; Simone 8 Chichvarkhin, 2004 [ЕЕК]. Arca (s. |.) sp.: Nutting, 1895 [ЕЕК]; T. Е. McGinty, 1939 [LFK], 1942; Lyman, 1944b [UFK]; Voss, 1949 [UFK; as “some Агсаз"]; Bender, 1965 [MFK, LFK; as Arcas]; Zischke, 1977b [MFK]; Gaertner, 1978 [LFK, as turkey wings]; Schomer & Drew, 1982; Voss et al., 1983 [ЧЕК]; Oliver & Járnegren, 2004 [ЕЕК]. Barbatia cancellaria (Lamarck, 1819): Simpson, 1887-1889 [DT; misidentified as Arca fusca Bruguière, 1789, synonym of Barbatia amygdalumtostum (Róding, 1798) a recognized Indian Ocean species]; Вай, 1889а, 1903b [misidentified as Arca (Barbatia) barbata Linnaeus, 1758, a recog- nized Mediterranean species]; Thiele, 1910 [DT; as Arca listeri Philippi, 1849, newly syn- onymized herein]; Lermond, 1936 [as Arca barbata]; M. Smith, 1937, 1945 [UFK; as Arca (B.) barbata]; Lyman, 1949b [as A. barbata]; Teare, 1949 [UFK; as A. barbata]; Bippus, 1950 [UFK; as A. barbata]; Т. A. Stephenson & A. Stephenson, 1950 [LFK; as A. barbata]; Ginsburg, 1952 [UFK, MFK; as A. barbata]; Eubanks, 1964; Iversen & Roessler, 1969 [UFK]; Plockelman, 1969b [МЕК]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Woods, 1971 [MFK]; Turney 8 Perkins, 1972 [UFK, MFK]; Zischke, 1973, 1977a, c [MFK]; Kissling, 1977b [UFK]; Antonius et al., 1978 [LFK]; Edwards, 1980 [LFK; as Arca]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Sedlak, 1986 CRITICAL CATALOG AND BIBLIOGRAPHY 591 [LFK]; Lyons 8 Quinn, 1995; Campbell et al., 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT]; Simone & Chichvarkhin, 2004 [UFK, MFK, ЕЕК]. |Barbatia candida (Helbling, 1779) — see un- der Cucullaearca. |Barbatia domingensis (Lamarck, 1819) — see under Acar. |Barbatia tenera (С. В. Adams, 1845) — see under Fugleria. Barbatia sp.: Melvill, 1880 [LFK]; Kissling, 1977a; Zischke, 1977b [MFK]; Schomer & Drew, 1982. Bathyarca glomerula (Dall, 1881): Dall, 1889a, 1903b [as Arca (Вуззоагса)]; Bartsch, 1937 [LFK; as Arca]; Mikkelsen & Bieler, 2000 [DT]. Bathyarca inaequalis (Dall, 1927). Bentharca sagrinata (Dall, 1886): Dall, 1889a [as Arca (Macrodon)]. Bentharca sp.: Dall, (Macrodon)]. Cucullaearca candida (Helbling, 1779): Simpson, 1887-1889 [DT; as Arca]; Lermond, 1936 [as Arca]; Webb, 1951 [as Arca]; Work, 1969 [LFK; as Barbatia]; Magnotte, 1970-1979 [as Barbatia]; Antonius et al., 1978 [LFK; as Barbatia]; Voss, 1983 [UFK; as Barbatia]; Voss et al., 1983 [UFK; as Barbatia]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Barbatia]; Simone & Chichvarkhin, 2004 [UFK, МЕК, ЕЕК; as Barbatia]. Fugleria tenera (C. B. Adams, 1845): Pilsbry & McLean, 1939 [LFK; as Arca (Barbatia) balesi Pilsbry & McLean, 1939]; Aguayo & Jaume, 1947b [LFK; as Arca (B.) balesi]; Abbott, 1958 [LFK; as Barbatia and as Arca balesi]; Magnotte, 1970-1979 [as Barbatia]; Lyons & Quinn, 1995 [as Barbatia]; Simone & Chichvarkhin, 2004 [MFK, LFK; as Barbatia]. Lunarca ovalis (Bruguiere, 1789): Dall, 1889а [as Arca (Argina) Americana “Gray” Wood, 1828]; Lermond, 1936 [аз Arca campechiensis americana]; Mikkelsen & Bieler, 2000 [MFK]. Scapharca brasiliana (Lamarck, 1819): Melvill, 1880 [LFK; misidentified as S. inaequivalvis (Bruguiére, 1789), a recognized Indian Ocean species]; Simpson, 1887-1889 [as A. incongrua Say, 1822]; Dall, 1889a, 1903b [both as А. (S.) incongrua]; Lermond, 1936 [as A. incongrua]. Scapharca chemnitzii (Philippi, 1851): Dall, 1889a [as Arca (Noetia) Orbignyi (err. pro d’orbignyi) Kobelt, 1891]; Lermond, 1936 [as Arca]. 1889a [as Arca Astartidae Astarte crenata subequilatera G. B. Sowerby Il, 1854: Dall, 1889a [as A. lens “Stimpson” Verrill, 1872]; Theroux & Wigley, 1983 [UFK, ЕЕК]. Astarte globula Dall, 1886. Astarte nana Jeffreys in Dall, 1886: Dall, 1889a, 1903b, c; Lermond, 1936; Pulley, 1952 [ЧЕК]; Mikkelsen & Bieler, 2000 [UFK, Din]: Astarte smithii Dall, 1886: Dall, 1889a, 1903b [аз A. Smithii]. Cardiidae Foster, 1945; Vittor & Associates, 1999a [ЧЕК], 1999b [UFK, МЕК]. Acrosterigma magnum (Linnaeus, 1758): Dall, 1889a, 1903b [as Cardium]; Pilsbry, 1890b [LFK; as Cardium]; Lermond, 1936 [as Cardium]; Clench & Smith, 1944 [DT; as Trachycardium]; Abbott, 1954 [LFK; as Trachycardium], 1958 [LFK; as Trachy- cardium], 1961 [LFK; as Trachycardium], 1974 [as Trachycardium (Acrosterigma)]; Parker & Curray, 1956 [LFK; as Trachy- cardium]; Warmke & Abbott, 1961 [LFK; as Trachycardium]; Barrett & Patterson, 1967 [LFK; as Trachycardium]; Humfrey, 1975 [LFK; as Trachycardium]; Romashko, 1984 [LFK; as Trachycardium]; Lawson, 1993 [as Trachycardium]; Rios, 1994 [аз 7. (Acrosterigma)]; Abbott & Morris, 1995 [LFK; as Trachycardium]; Lyons & Quinn, 1995 [as Trachycardium]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Trachycardium]. Americardia guppyi (Thiele, 1910): Abbott, 1958 (MFK, LFK], 1974; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, ВРК Dit}: Americardia media (Linnaeus, 1758): Melvill, 1880 [LFK; as Hemicardium medium]; Simpson, 1887-1889 [LFK; as Cardium]; Dall, 1889a [as C. medium], 1896a [LFK; as C. medium], 1903b [as C. medium]; Lermond, 1936 [as C. (Hemicardium) me- dium]; М. Smith, 1937, 1945 [both UFK; as Trigoniocardia (A.) medium]; Clench & Smith, 1944 [UFK, MFK, LFK; as Trigoniocardia (A.)]; Brooks, 1968b [MFK]; Iversen & Roessler, 1969 [UFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, MFK; as T. medium]; Zischke, 1973, 1977a [both MFK]; Voss, 1983 [UFK]; 592 MIKKELSEN & BIELER Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor 8 Associates, 1997c [ЧЕК]; Mikkelsen € Bieler, 2000 [UFK, MFK, LFK, ОТ]. Americardia sp.: Plockelman, 19684. Dinocardium robustum (Lightfoot, 1786): Magnotte, 1970-1979. Dinocardium sp.: Stevenson, 1970, 1993 [both as great heart cockle]. Laevicardium laevigatum (Linnaeus, 1758): Melvill, 1880 [LFK; as L. laevigatum and misidentified as L. serratum (Linnaeus, 1758), a recognized Indo-Pacific species]; Dall, 1886 [LFK; as Cardium], 1889a [as Papyridea (Liocardium) laevigatum and P. (L.) serratum], 1896a [LFK; as С. serratum], 1903b [as P (Liocardium) laevigatum and P. (L.) serratum]; Simpson, 18871889 [LFK; as L. serratum]; Lermond, 1936 [as Cardium (Laevicardium) laevigatum and C. (L.) serratum]; Clench & Smith, 1944 [LFK, DT]; Bippus, 1950 [UFK; as L. serratum]; Brooks, 1968b [MFK]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Woods, 1970 [LFK]; Turney & Perkins, 1972 [UFK, MFK]; Godcharles & Jaap, 1973 [UFK]; Zischke, 1977a [MFK]; Goldberg, 1978c [LFK]; Schomer & Drew, 1982; Voss, 1983 [ЧЕК]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1999a [DT], b [MFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT]; USGS, 2002 [ЧЕК]. Laevicardium mortoni (Conrad, 1830): Thiele, 1910 [DT; as Cardium]; Bartsch, 1937; Clench & Smith, 1944 [UFK, MFK]; Bippus, 1950 [UFK]; Ross, 1969 [MFK]; Howard et al., 1970 [LFK]; Hudson et al., 1970 [UFK]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, MFK]; Lineback, 1977 [ЕЕК]; Petersen, 1989 [MFK]; Lyons & Quinn, 1995; Wingard et al., 1995 [UFK; as L. spp.]; Brewster-Wingard et al., 1996, 1997 [both as L. spp.], 1998 [as L. sp.] 2001 [also as L. spp.] [all ЧЕК]; Vittor & Associates, 1997c [UFK]; Brewster-Wingard & Ishman, 1999 [UFK; as L. spp.]; Mikkelsen & Bieler, 2000 МРК; МЕК, “LEK DT]; USGS; 2003 ШЕК, МЕК]; Morton & Knapp, 2004 [UFK, MFK]. Laevicardium pictum (Ravenel, 1861): Mikkelsen & Bieler, 2000 [UFK, LFK, DT]. Laevicardium sybariticum (Dall, 1886): Lyons 8 Quinn, 1995; Vittor & Associates, 1999b [LFK]; Mikkelsen & Bieler, 2000. Laevicardium sp.: Gaertner, 1978 [LFK; as egg cockles]; Bielsa & Labisky, 1987 [ЕЕК]; Vittor 8 Associates, 1999b [LFK]; USGS, 2003 [UFK; as L. spp.]. Nemocardium peramabile (Рай, 1881): Бай, 1881 [DT; as Cardium (Fulvia) peramabilis], 1886 [LFK; as С. (F.) peramabilis], 1889a, 1903b [as С. peramabilis]; Lermond, 1936 [as С. (Protocardia) peramabilis]; Bartsch, 1937 [LFK; as P. peramabilis]; Clench & Smith, 1944 [LFK; as Microcardium]; Theroux & Wigley, 1983 [LFK]; Mikkelsen & Bieler, 2000 [UFK, LFK]. Nemocardium tinctum (Dall, 1881): Dall, 1881 [DT; as Cardium (Fulvia) peramabilis var. tinctum], 1889a [LFK; as C. peramabilis var. tinctum], 1903b [LFK; as C. peramabilis var. tinctum], 1900b [LFK; as Protocardia tincta Dall, 1886]; Johnson, 1934 [LFK; as P. tincta]; Mikkelsen & Bieler, 2000 [LFK, DT]. Papyridea lata (Born, 1778): Webb, 1942, 1951 [in part, figured as Paphridea spinosum (Meuschen, 1787)]; Clench & Smith, 1944 [in part, as P. hiatus (Meuschen, 1787)]; Barfield, 1990 [LFK; as Р. soleniformis (Bruguiére, 1789)]; Watters, 2002 [LFK]. Papyridea semisulcata (Gray, 1825): Simpson, 1887-1889 [ОТ; as Cardium petitianum Orbigny, 1842]; Dall, 1889a [as P Petitiana], 1900a [UFK; as Cardium (Papyridea) semisulcatum]; Lermond, 1936 [as Cardium (Papyridea) semisulcatum]; Clench & Smith, 1944 [LFK]; Olsson & Harbison, 1952 [LFK; as P. semisulcatum]; Pulley, 1952 [ЕЕК]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, (ЕК, DT]. Papyridea soleniformis (Bruguiére, 1789): Dall, 1889a [misidentified as P. bullata (Linnaeus, 1758)]; Lermond, 1936 [as Cardium (Papyridea) зртозит]; Webb, 1942, 1951 [in part, figured as Paphridea spinosum (Meuschen, 1787)]; Clench & Smith, 1944 [МЕК, LFK; in part, as P hiatus (Meuschen, 1787)]; Brooks, 1968b [МЕК]; Plockelman, 1968c [MFK]; Ross, 1969 [MFK]; Sedlak, 1986 [LFK]; Krisberg, 1993 [LFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Watters, 2002 [LFK, DT]. Trachycardium egmontianum (Shuttleworth, 1856): Dall, 1889a [misidentified as Cardium isocardia Linnaeus, 1758, a recognized southern Caribbean species]; Nutting, 1895 [ОТ; as С. isocardium]; Lermond, 1936 [as С. isocardia]; Clench & Smith, 1944 [DT ]; Brooks, 1968b [MFK]; Ross, 1969 [MFK]; Magnotte, 1970-1979; Godcharles & Jaap, 1973 [UFK]; Antonius et al., 1978 [LFK; as T. isocardia]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. CRITICAL CATALOG AND BIBLIOGRAPHY 993 |Trachycardium magnum (Linnaeus, 1758) — see under Acrosterigma. Trachycardium muricatum (Linnaeus, 1758): Melvill, 1880 [LFK]; Dall, 1889a, 1903b [as Cardium]; Lermond, 1936 [as Cardium]; Clench & Smith, 1944 [MFK, LFK]; Brooks, 1968b [MFK]; Ross, 1969 [MFK]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, МЕК]; Godcharles & Jaap, 1973 [ЧЕК]; Lyons & Quinn, 1995; Vittor & Associates, 1999b [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК]; USGS, 2003 [МЕК]. Trachycardium sp.: Plockelman, 1968а; Iversen & Roessler, 1969 [UFK]; Vittor & Associates, 1999b [LFK]. Trigoniocardia antillarum (Orbigny, 1842): Dall, 1889a [as Cardium]; Lermond, 1936 [as Cardium (Trigoniocardia)]. Carditidae Vittor & Associates, 1998 [LFK, DT], 1999a [DT], 1999b [ЧЕК]. Carditamera floridana Conrad, 1838: Tryon, 1873 [LFK; as Cardita (Carditamera)]; Melvill, 1880 [LFK; Cardita (Mytilicardia) floridana “(Sowerby)”]; Simpson, 1887-1889 [MFK; as Cardita]; Dall, 1889a [LFK; also as “2” Cardita Conradii (err. pro conrad!) Shuttleworth, 1856], 1903b [LFK; as Cardita], 1903a [as С. (Carditamera)]; Lermond, 1936 [as Cardita]; Bartsch, 1937 [as Cardita]; Pul- ley, 1952 [as Cardita]; Hudson et al., 1970 [UFK; as Cardita]; Turney & Perkins, 1972 [UFK, MFK; as Cardita]; Petersen, 1989 [MFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK, МЕК; also as Cardita]. Glans dominguensis (Orbigny, 1842): Dall, 1889a [as Cardita domingensis (sic)]; Pul- ley, 1952 [UFK; as Cardita domingensis (sic)]; Rehder, 1981; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK, DT], 1999a [UFK], b [MFK, LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Pleuromeris tridentata (Say, 1826): Henderson, 1913 [UFK]; Pulley, 1952 [as Venericardia]; Turney & Perkins, 1972 [UFK; as Venericardia]; Rehder, 1981; Theroux & Wigley, 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK, DT], 1999a ОЕ, ВЕК DIT BU ER MEK, ERK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Morton & Knapp, 2004 [UFK, MFK]. Pteromeris perplana (Conrad, 1841): Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MEK EEK DIT]: Chamidae Dent, 1998. Arcinella cornuta Conrad, 1866: Calkins, 1878 [DT; misidentified as Chama arcinella Linnaeus, 1767, a recognized Caribbean species of Arcinella]; Dall, 1889a [as С. arcinella], 1896a [as С. (Echinochama) arcinella], 1903b [as С. arcinella]; Lermond, 1936 [as Cardium arcinella, but corrected to Chama arcinella in copy signed by Lermond]; Magnotte, 1970-1979; Dalton, 1991 [LFK; as A. arcinella]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT]; Campbell et al., 2004. Chama congregata Conrad, 1833: Lermond, 1936; Bartsch, 1937 [LFK, DT]; Eubanks, 1964; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Godcharles & Jaap, 1973 [UFK]; Antonius et al., 1978 [LFK]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Campbell et al., 2004 [MFK, ЕЕК]. Chama florida Lamarck, 1819: Bayer, 1943b [UFK, DT]; Edwards, 1968a [LFK], 1969 [UFK]; Antonius et al., 1978 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Campbell et al., 2004 [MFK]. Chama inezae (F. M. Bayer, 1943): Bayer, 1943b [UFK; as Pseudochama]; M. Smith, 1945 [UFK; as Pseudochama]; Aguayo & Jaume, 19506 [UFK; as Pseudochama]; Plockelman, 1969a [UFK; as Pseudochama]; Mikkelsen & Bieler, 2000 [UFK; as Pseudochama]; Campbell et al., 2004 [MFK]. Chama lactuca Dall, 1886: Р. L. McGinty & T. L. McGinty, 1957 [MFK, LFK]; Mikkelsen, 1981 [UFK; as C. sp.]; Lyons & Quinn, 1995; Mikkelsen € Bieler, 2000 [UFK, MFK, LFK]; Campbell et al., 2004. Chama macerophylla Gmelin, 1791: Wheatley, 1845 [misidentified as C. lazarus Linnaeus, 1758, a recognized Indo-Pacific species]; Calkins, 1878; Melvill, 1880 [LFK]; Simpson, 1887-1889 [DT]; Dall, 1889a, 1903b; Thiele, 1910 [DT]; Lermond, 1936; Bayer, 1943b [LFK, DT]; Eubanks, 1964; Lee, 1969 [LFK]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Close, 1974 [ЕЕК]; Goldberg, 1978 [MFK]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Clampit, 1987 [ЕЕК]; Clampit 1988 [LFK]; Lyons 8 Quinn, 1995; Purtymun, 1997 [ЕЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Orlin 2003; Campbell et al., 2004 [MFK, LFK]. 594 MIKKELSEN & BIELER Chama radians Lamarck, 1819: Webb, 1942, 1951 [as Chama variegata Reeve, 1847, a small corrugated growth form from eastern Florida]; Eubanks, 1964 [as Pseudochama]; Lee, 1969 [LFK; as Pseudochama radians variegata]; Turney & Perkins, 1972 [MFK; as Pseudochama]; Godcharles & Jaap, 1973 [UFK; as Pseudochama]; Antonius et al., 1978 [LFK; as Pseudochama]; Lyons & Quinn, 1995 [аз Pseudochama]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Pseudochama]; Campbell et al., 2004 [MFK, ЕЕК]. Chama sarda Reeve, 1847: Dall, 1889a, 1903b; Johnson 1934; Lermond, 1936; Bartsch, 1937; M. Smith, 1937, 1945; Pilsbry 8 McGinty, 1938; Bayer, 1943b [UFK]; Mor- ris, 1947, 1951; Warmke 8 Abbott, 1961; Magnotte, 1970-1979 [as С. sardo (sic)]; Abbott, 1974; Humfrey, 1975; Rehder, 1981; Díaz Merlano & Puyana Hegedus, 1994; Rios, 1994; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT]; Campbell et al., 2004 [UFK, МЕК, ЕЕК]. Chama sinuosa Broderip, 1835: Bayer, 1943b [ОТ; as С. sinuosa bermudensis Heilprin, 1889, & as С. sinuosa firma Pilsbry & McGinty, 1938]; Antonius et al., 1978 [LFK; as C. sinosa (sic)]; Lyons & Quinn, 1995; Cantillo et al., 1997 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Campbell et al., 2004 [MFK]. Chama sp.: T. L. McGinty, 1939 [LFK], 1942; Brooks, 1968b [MFK]; Edwards, 1970 [LFK, as large and small Chama]; Stevenson, 1970, 1993 [both as Jewelbox]; Oliver & Jarnegren, 2004 [LFK]. |Pseudochama inezae F. М. Bayer, 1943 — see under Chama. |Pseudochama radians (Lamarck, 1819) — see under Chama. Condylocardiidae Carditopsis smithii (Dall, 1896): Pilsbry & Olsson, 1946 [LFK; as Condylocardia floridensis Pilsbry & Olsson, 1946]; Aguayo & Jaume, 1950c [LFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Corbiculidae Polymesoda maritima (Orbigny, 1842): Бай, 1883, 1885 [LFK; misidentified as Cyrena carolinensis; err. pro caroliniana Bosc, 1801, a recognized western Atlantic species], 1889a, 1903b [as Cyrena (Leptosiphon) carolinensis (sic)], 1903a [аз С. (Pseudocyrena) floridana Conrad, 1846]; M. Smith, 1937, 1945 [LFK; as Polymesoda floridana]; Webb, 1951 [as Cyrena floridana]; Pulley, 1952 [as Pseudocyrena floridana]; Abbott, 1954, 1961 [LFK, as Pseudocyrena], 1970 [LFK; as Pseudocyrena floridana]; Morrison, 1958 [UFK, LFK; as Pseudocyrena]; Siekman, 1965 [as Pseudocyrena floridana]; Bender, 1968 [probably misidentified as Rangia flexuosa (Conrad, 1840), a recognized species of Rangianella from Gulf of Mexico marshes]; Howard et al., 1970 [LFK; as Polymesoda floridana]; Andrews, 1971 [LFK; as Polycyrena floridana]; Turney & Perkins, 1972 [MFK]; Emerson & Jacobson, 1976 [LFK; as Pseudocyrena floridana]; Ode, 1976a [LFK; as Pseudocyrena]; Andrews, 1977, 19814 b 1992, 1994 [LEK asta (Pseudocyrena)]; Lineback, 1977 [LFK; as P. floridana]; Petersen, 1989 [MFK; as Pseudocyrena]; Lyons & Quinn, 1995; Brewster-Wingard et al., 1996 [UFK; as Polymesoda sp.]; Brewster-Wingard & Ishman, 1999 [UFK; as Polymesoda sp.]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK; also as Rangia flexuosa, based on Bender, 1968 (see previous)]; USGS, 2003 [UFK, MFK]. Corbulidae Foster, 1945 [as Aloididae]; Vittor & Associ- ates, 1998 [LFK, ОТ]. Caryocorbula caribaea Orbigny, 1842: Simpson, 1887-1889 [DT; as Corbula swiftiana (C. B. Adams, 1852)]; Dall, 1889a [as Corbula barrattiana (C. B. Adams, 1852) and Corbula swiftiana], 1903b [as Corbula Barrattiana and Corbula Swiftiana]; Lermond, 1936 [as Corbula barrattiana and Corbula swiftiana]; Antonius et al., 1978 [LFK; as Corbula swiftiana]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Corbula, also as Corbula barrattiana and Corbula swiftiana]. Caryocorbula chittyana (C. B. Adams, 1852): Dall, 1889a [as Corbula nasuta Say, 1833]; Lermond, 1936 [as Corbula nasutal. Caryocorbula contracta (Say, 1822): Lermond, 1936 [as Corbula]; Vittor & Associates, 1998 [DT; as Corbula], 1999a [LFK, DT; as Corbula}; Mikkelsen & Bieler, 2000 [DT; as Corbula]. CRITICAL CATALOG AND BIBLIOGRAPHY 595 Caryocorbula cymella (Dall, 1881): Dall, 1881, 1889a, 1903b [all Gordon Key; as Corbula]; Johnson, 1934 [Gordon Key; as Corbula (Caryocorbula)]; Lermond, 1936 [as Corbula]; M. Smith, 1937, 1945 [both Gor- don Key; as Corbula]; Aguayo & Jaume, 1950a [Gordon Key; as Corbula]; Parker € Curray, 1956 [Gordon Key; as Corbula]; Boss et al., 1968 [Gordon Key; as Corbula]; Abbott, 1974 [Gordon Key; as Corbula (Caryocorbula)]; Rios, 1994 [as Corbula (Caryocorbula)]. Caryocorbula dietziana (C. B. Adams, 1852): Dall, 1886 [Gordon Key; as Corbula], 1889a [as Corbula], 1903b [as Corbula Dietziana]; Lermond, 1936 [as Corbula]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT; as Corbula]. Corbula (s. |.) sp.: Turney & Perkins, 1972 [UFK, MFK]. Juliacorbula aequivalvis Philippi, 1836: Dall, 1889a [as Corbula Cubaniana (Orbigny, 1842)]; Pulley, 1952 [as Aloidis]. Varicorbula disparilis (Orbigny, 1842): Dall, 1881 [DT]; Lermond, 1936 [as Corbula disparillis (sic)]; Pulley, 1952 [as Aloidis operculata (Philippi, 1848)]; Vittor & Associ- ates, 1998 [LFK, DT; as V. operculatal, 1999а [LFK, DT; as М. operculata]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT; as V. limatula (Conrad, 1846)], 2001 [UFK, LFK, DT]. Varicorbula krebsiana (C. B. Adams, 1852): Dall, 1889a [as Corbula Krebsiana]. Varicorbula philippii (E. A. Smith, 1885): Mikkelsen & Bieler, 2000 [LFK, DT], 2001 ЕК, РТ]. Crassatellidae Vittor & Associates, 1998 [ЕЕК]. Crassinella dupliniana (Dall, 1903): Mikkelsen & Bieler, 2000 [МЕК]. Crassinella lunulata (Conrad, 1834): Dall, 1889a [as Crassatella (Eriphyla), also as Crassatella (Eriphyla) lunulata var. parva (C. B. Adams, 1845)], 1903b [as Crassatella (Eriphyla)l; Johnson, 1934 [as Gouldia parva (С. В. Adams, 1845)]; Lermond, 1936 [as Gouldia mactracea (Linsley, 1845) and as С. parva]; Lyons & Quinn, 1995; Vittor & As- sociates, 1998 [LFK, DT], 1999a [LFK, ОТ], b [UFK, МЕК, LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Crassinella martinicensis (Orbigny, 1842): Lyons & Quinn, 1995; Vittor 8 Associates, 1997b [LFK], 1998 [LFK, DT], 1999a [LFK, DT]; Mikkelsen & Bieler, 2000 [LFK, DT]. Crassinella sp.: Vittor & Associates, 1998 [LFK, DT], 1999b [МЕК]. Eucrassatella speciosa (A. Adams, 1852): Dall, 1889a [as Crassatella floridana Dall, 1881], 1890 [as Crassatella floridana], 1903b [as Crassaella floridana]; Lermond, 1936 [as Crassatellites gibbsii err. pro gibbesii (Tuomey 8 Holmes, 1856)]; Pulley, 1952; M. Smith, 1961 [as Eucrassatella floridana]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [UFK, LFK, DT]. Cuspidariidae Theroux & Wigley, 1983 [UFK, ЕЕК]. Cardiomya alternata (Orbigny, 1842): Dall, 1881 [LFK; as Neaera]; Knudsen, 1982 [ЕЕК Cardiomya costellata (Deshayes, 1830): Dall, 1889a, 1903b [as Cuspidaria (Cardiomya)]; Lermond, 1936 [as Cuspidaria (Cardiomya)]; Turney & Perkins, 1972 [МЕК]; Mikkelsen & Bieler, 2000 [LFK]. Cardiomya glypta (Bush, 1885): Mikkelsen & Bieler, 2000 [ОТ]. Cardiomya ornatissima (Orbigny, 1842): Mikkelsen & Bieler, 2000 [DT]. Cardiomya perrostrata (Dall, 1881): Dall, 1886 [ОТ; as Cuspidaria (Cardiomya)], 1889a, 1903b [DT; as Cuspidaria (Cardiomya)]; Pulley, 1952 [DT]; Vittor & Associates, 1998 [LFK, DT], 1999a [LFK]; Mikkelsen & Bieler, 2000 [LFK, DT]. Cardiomya striata (Jeffreys, 1876): Dall, 1889a [as Cuspidaria (Cardiomya)]. Cardiomya sp.: Vittor & Associates, 1998 [DT]. Cuspidaria gigantea (A. E. Verrill, 1884): Mikkelsen & Bieler, 2000 [LFK]. Cuspidaria obesa (Lovén, 1846): Dall, 1889a [as Cuspidaria (Cuspidaria)]. Cuspidaria rostrata (Spengler, 1793): Dall, 1881 [LFK; as Neaera], 1889a, 1903b [as С. (Cuspidaria)]; A. E. Verrill, 1882 [LFK; as Neara]; Aguayo 8 Jaume, 1950f [ЕЕК]; Mikkelsen € Bieler, 2000 [DT]. |Leiomya claviculata (Dall, 1881) — all records based on Blake sta. 44 (here excluded; see entry for Dall, 1881). Myonera lamellifera (Dall, 1881): Dall, 1889a. |Myonera limatula (Dall, 1881) — all records based on Blake sta. 44 (here excluded; see entry for Dall, 1881). Myonera paucistriata Dall, 1886. Plectodon granulatus (Dall, 1881): Dall, 1889a, 1903b [as both Cuspidaria (Liomya, Plectodon) granulata and С. (L., Р) 9. var. 596 MIKKELSEN & BIELER velvetina Dall, 1881]; Johnson, 1934 [as both Leiomya (P.) granulata granulata and L. (P) granulata velvetinal; Aguayo & Jaume, 1950d [as Leiomya (P.) granulata 1. velvetina]; Abbott, 1974; Odé, 1977a; Mikkelsen & Bieler, 2000 [ЧЕК]. Cyrenoididae Cyrenoida floridana (Dall, 1896): Simpson, 1887-1889 [MFK; nomen nudum]; Dall, 1889a [nomen nudum, as Cyrenoidea], 1903b [as Cyrenoidea]; Pulley, 1952; Mikkelsen & Bieler, 2000 [UFK, LFK]; USGS, 2003 [MFK]. Donacidae Donax variabilis Say, 1822: Simpson, 1887- 1889 [LFK; misidentified as D. denticulatus Linnaeus, 1758, a recognized Caribbean species]; Dall, 1889a, 1903b [also misidentified as D. denticulatus and D. fossor Say, 1822, a recognized Atlantic U.S. spe- cies], 1900a [as D. fossor]; Maury, 1920 [as D. fossor]; Johnson, 1934 [as D. denticulata]; Lermond, 1936 [also as D. roemeri Philippi, 1849, and D. fossor protractus Conrad, 1849, and misidentified as D. denticulata, D. fossor, and D. tumidus Philippi, 1848, a synonym of D. texasianus Philippi, 1847, a recognized Gulf of Mexico species]; Bartsch, 1937; M. Smith, 1937, 1945 [аз D. denticulata and D. fossor], 1940 [as D. fossor]; Mikkelsen 8 Bieler, 2000 [UFK, MFK]; Simone & Dougherty, 2004 [ЧЕК]. Donax sp.: Bippus, 1950 [UFK]; Morrison, 1970 [ЕЕК]. Iphigenia brasiliana (Lamarck, 1818): Бай, 1889a [as /. braziliana], 1903b; Lermond, 1936; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [UFK]. Dreissenidae Mytilopsis leucophaeata (Conrad, 1831): Mikkelsen & Bieler, 2000 [UFK]. Mytilopsis sallei (Récluz, 1849): Johnson, 1934 [as Congeria rossmássleri Dunker, 1853]; Lermond, 1936 [as С. rossmassleri]; Mikkelsen & Bieler, 2000 [UFK]. Entoliidae Pectinella sigsbeei (Dall, 1886): Dall, 1889a [as Pecten (Pecten, Pseudamusium) Sigsbeel]. Galeommatoidea Hendler et al., 1995 [as “commensal bi- valve”]; Vittor 8 Associates, 1998 [LFK, DT; as Montacutidae], 1999а [LFK; as Montacutidae], b [LFK; as Montacutidae]. Cymatioa sp.: Odé, 1984; Mikkelsen 8 Bieler, 2000 [ЧЕК]. Kellia suborbicularis (Montagu, 1803): Mikkelsen, 1981 [UFK; misidentified as Diplodonta (Diplodonta) ?punctata (Say, 1822); reidentification of voucher specimen]; Mikkelsen & Bieler, 2000 [UFK]. Lasaea adansoni (Gmelin, 1791): Simpson, 1887-1889 [DT; as Lasea (sic) rubra (Montagu, 1803)]; Johnson, 1934 [DT; as Kellia rubra]; Aguayo & Jaume, 1949a [DT]; Mikkelsen & Bieler, 2000 [MFK, DT]. Mysella planulata (Krause, 1885): Wingard et al., 1995 [UFK; as М. sp.]; Vittor 8 Asso- ciates, 1997c [UFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Brewster- Wingard et al., 2001 [UFK]; USGS, 2003 [UFK]. Mysella sp.: Hendler et al., 1995 [as M. sp. С]; Vittor & Associates, 1999a [DT]. Orobitella floridana (Dall, 1899): Hendler et al., 1995 [LFK; as Naeromya sp.]; Vittor & As- sociates, 1998 [DT; as Naeromya], 1999b [UFK, LFK; as Naeromya]; Mikkelsen & Bieler, 2000 [MFK, LFK, DT]. Semierycina sp.: Mikkelsen & Bieler, 2000 [UFK, МЕК]. Gastrochaenidae Gastrochaena hians (Gmelin, 1791): Dall, 1889a [misidentified as С. cuneiformis Spengler, 1783], a recognized Indo-Pacific species; Bales, 1944 [as С. cueniformis (sic)]; Lermond, 1936 [аз С. cuneiformis]; Edwards, 1968a [ЕЕК]; Crovo, 1970 [LFK]; Andrews, 1971 [as Rocellaria]; Carter, 1978 [аз С. (Gastrochaena)]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK], 1999a [UFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Valentich-Scott & Dinesen, 2004 [MFK, LFK]. Gastrochaena ovata С. В. Sowerby |, 1834: Simpson, 1887-1889 [LFK; as Rocellaria]; Dall, 1889a, 1903b; Lermond, 1936; Bales, 1940, 1944; Carter, 1978 [as С. (Rocellaria)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. |Rocellaria ovata (С. В. Sowerby |, 1834) — see under Gastrochaena. CRITICAL CATALOG AND BIBLIOGRAPHY 597 Spengleria rostrata (Spengler, 1783): Dall, 1886 [LFK; as Cuspidaria], Dall, 1889a, 1903b [аз Gastrochaena (Spengleria)]; Simpson, 1887- 1889 [LFK; as Rocellaria]; Lermond, 1936 [as Gastrochaena]; Bales, 1940, 1944 [as Gastrochaena]; Boss, 1968a; Carter, 1978; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Glycymerididae Vittor & Associates, 1998 [LFK], 1999b [UFK]. Glycymeris americana (DeFrance, 1829): Lermond, 1936; Bartsch, 1937; Webb, 1942, 1951 [as Glycimeris]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [UFK, ОТ]. Glycymeris decussata (Linnaeus, 1758): Calkins, 1878 [as Pectunculus pennaceus Lamarck, 1819]; Lermond, 1936 [as G. pennacea]; Pulley, 1952 [LFK]; Magnotte, 1970-1979; Thomas, 1970 [MFK]; Lyons & Quinn, 1995; Vittor 8 Associates, 1999a [UFK, LFK, DT]; Mikkelsen 8 Bieler, 2000 [UFK, МЕК, LFK, ОТ]. |Glycymeris pectinata (Gmelin, 1791) — see under Tucetona. Glycymeris spectralis (Nicol, 1952). Glycymeris undata (Linnaeus, 1758): Dall, 1889а, 1903b [as Pectunculus undatus]; Lermond, 1936 [as G. lineata Reeve, 1847]; Voss, 1949 [UFK; as Glycimeris americana lineata]; Pulley, 1952; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000. Glycymeris (s. |.) sp.: Theroux & Wigley, 1983 [UFK, MFK]; Voss et al., 1983 [UFK]; Vittor & Associates, 1998 [LFK, DT]. Tucetona pectinata (Gmelin, 1791): Melvill, 1880 [LFK; misidentified as Pectunculus pectiniformis Lamarck, 1819, a junior syn- onym of Tucetona pectunculus (Linnaeus, 1758), a recognized Indo-Pacific species]; Dall, 1889a, 1903b [as Pectunculus pectinatus]; Thiele, 1910 [DT; as Glycymeris pectinatus]; Lermond, 1936 [as Glycymeris]; М. Smith, 1937, 1945 [as С. pectinatus]; Lyman, 1947b [UFK; as Glycimeris (sic) pectinatus]; Pulley, 1952 [as Glycymeris]; Eubanks, 1964 [as Glycymeris]; Brooks, 1968b [MFK; as Glycymeris]; Burggraf, 1969 [LFK; as Glycymeris]; Ross, 1969 [MFK; as Glycymeris]; Magnotte, 1970-1979 [as Glycymeris]; Thomas, 1970 [MFK, LFK; as Glycymeris]; Woods, 1970 [LFK; as Glycymeris]; Turney & Perkins, 1972 [UFK, МЕК; as Glycymeris]; Godcharles & Jaap, 1973 [UFK; as Glycymeris]; Thomas, 1975 [MFK, LFK; as Glycymeris]; Hertweck, 1977 [LFK; as Glycymeris]; Theroux & Wigley, 1983 [UFK; as Glycymeris]; Voss, 1983 [UFK; as Glycimeris (sic)]; Voss et al., 1983 [UFK; as Glycymeris]; Lyons & Quinn, 1995 [as Glycymeris]; Vittor & Associates, 1998 [LFK, DT; as Glycymeris]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Glycymeris]; Morton & Knapp, 2004 [UFK, МЕК]. Gryphaeidae Hyotissa mcgintyi (Harry, 1985): Kirkendale et al., 2004 [MFK]. Neopycnodonte cochlear (Poli, 1795): Mikkelsen & Bieler, 2000 [UFK, LFK]. Hiatellidae Hiatella arctica (Linnaeus, 1767): Dall, 1889a, 1903b [as Saxicava]; Vittor & Associates, 1998 [LFK, DT], 1999a [DT], b [UFK, MFK]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT]. Hiatella azaria (Dall, 1881): Dall, 1889a [as Saxicava]. Isognomonidae Isognomon alatus (Gmelin, 1791): Calkins, 1878 [ОТ; misidentified as Рета perna “Wood”, err. pro (Linnaeus, 1767) (non Linnaeus, 1758 = Mytilidae) with synonym Perna ephippium “Sowerby” err. pro (Linnaeus, 1758), a recognized Indo-Pacific species of Isognomon]; Dall, 1883, 1885 [LFK; misidentified as P. ephippium], 1889a [as Р ephippium “Lamarck” and Р. obliqua Lamarck, 1819], 1896a [as Perna oblique (sic)]; 1903b [as P. ephippium “Lamarck’”]; Simpson, 1887-1889 [LFK, DT, as Perna obliqua; also DT, as P. ephippium]; Thiele, 1910 [DT; as Melina lata (sic)]; Lermond, 1936 [as Pedalion (Perna)]; Bartsch, 1937 [as Melina]; M. Smith, 1937 [UFK; as Pedalion alata], 1945 [UFK; as Pedalion alata]; С. М. Vilas & М. К. Vilas, 1945, 1970 [as Pedalion alata]; Т. A. Stephenson & A. Stephenson, 1950 [LFK; as /. (Pedalion) alata]; Pulley, 1952 [UFK; as /. alata]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Zischke, 1973, 1977a, b, с [MFK]; Chan, 1977b [LFK]; Kissling, 1977a [as Isognomen (sic)]; Goldberg, 1978c [МЕК]; Schomer 8 Drew, 1982; Wagner & Abbott, 1990; Lyons 8 Quinn, 1995; Hutsell et al., 1997; Campbell et al., 1998 [LFK]; 598 Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]; Domaneschi & Mantovani, 2002 [MFK, LFK]. Isognomon bicolor (С. В. Adams, 1845): Johnson, 1934 [as Pedalion semiaurita (Linnaeus, 1758)]; Lermond, 1936 [as Pedalion, also as P. semiaurita]; Т. A. Stephenson & A. Stephenson, 1950 [LFK; as /. chemnitziana (Orbigny, 1846)]; Pulley, 1952; Abbott, 1954 [ЕЕК]; Warmke & Abbott, 1961; Andrews, 1971, 1977, 1981a, b, 1992, 1994; Zischke, 1973, 1977a, b [MFK]; Abbott, 1974; Humfrey, 1975; Emerson & Jacobson, 1976; Ingham 8 Zischke, 1977 [MFK]; Odé, 1979a; Rehder, 1981; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]; Domaneschi 8 Mantovani, 2002. Isognomon radiatus (Anton, 1839): Thiele, 1910 [ОТ; as Melina listeri Hanley, 1846]; Johnson 1934 [as Pedalion listeri]; Lermond, 1936 [as Р listeri]; M. Smith, 1937, 1945 [as Р listeri); Lyman, 1943 [LFK; as Р listeri]; Pulley, 1952 [as /sognomon listeri]; Edwards, 1968b [LFK]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Zischke, 1973, 1977a [MFK]; Ingham 8 Zischke, 1977 [MFK]; Antonius et al., 1978 [LFK]; Mikkelsen, 1981 [ЧЕК]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, DT; also as Neopycnodonte cochlear from MFK, based on misidentified specimen]. Isognomon sp.: Voss et al., 1983 [UFK]; Oliver & Jarnegren, 2004 [LFK]. Limidae Henderson, 1911 [DT; as Limas]; Vittor & Associates, 1999b [UFK]. |Ctenoides floridanus (Olsson & Harbison, 1953) — see Ctenoides mitis. Ctenoides miamiensis Mikkelsen & Bieler, 2003: Mikkelsen & Bieler, 2003 [LFK, DT]. Ctenoides mitis (Lamarck, 1807): Simpson, 1887-1889 [DT; as Lima tenera “Chemnitz” G. B. Sowerby II, 1843]; Dall, 1889a, 1903b [both as L. tenera]; Lermond, 1936 [as L. [епега]; Webb, 1942, 1951 [both as L. tenera]; Pulley, 1952 [as L. tenera]; Schroeder, 1964 [UFK; as flame scallop]; Brooks, 1968b [MFK; as L. scabra form tenera]; Plockelman, 1969b [MFK; as L. scabra tenera]; Ross, 1969 [MFK; as L. scabra tenera], 1971 [MFK; as L. scabra tenera]; Work, 1969 [LFK; as L. scabra form {епега]; Woods, 1970, 1971 [МЕК] [both as L. scabra tenera]; Mpitsos, 1973 [as L. scabra tenera]; Voss et al., 1983 [UFK; as MIKKELSEN & BIELER Lima sp. with “red and yellow tentacles”); Williams, 1988 [as “file clam”]; Cohen & Cohen, 1991 [UFK; unnamed, with light-or- ange tentacles]; Lyons & Quinn, 1995 [as L. scabra tenera]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as C. floridanus (Olsson & Harbison, 1953)], 2003 [UFK, MFK, ЕЕК, DT]; Morton, 2000 [MFK; as С. floridanus]. Ctenoides planulatus (Dall, 1886): Mikkelsen & Bieler, 2000 [LFK; as С. planulatatus (sic)]; 2003 [ЕЕК]. Ctenoides sanctipauli Stuardo, 1982: Stuardo, 1982 [LFK]; Mikkelsen & Bieler, 2000 [MFK, ГЕК, DT], 2003 [MFK, ЕЕК, DT]. Ctenoides scaber (Born, 1778): Calkins, 1878 [DT; as Lima scabra]; Melvill, 1880 [LFK; as L. scabra]; Simpson, 1887-1889 [DT; as L. scabra]; Dall, 1889a, 1903b [as L. scabra]; Thiele, 1910 [DT; as L. scabra]; Lermond, 1936 [as L. scabra]; Bartsch, 1937 [LFK, DT; as L. scabra]; Webb, 1942 [misidentified as L. tenera, synonym of Ctenoides floridanus (see above)]; Pulley, 1952 [as L. scabra]; Ross, 1969 [MFK; as L. scabra]; Work, 1969 [LFK; as L. scabra]; Magnotte, 1970-1979 [as L. scabra]; Jacobson & Hernandez, 1973 [DT; as L. scabra]; Mpitsos, 1973 [as L. scabra]; Zischke, 1973, 1977a, c [MFK; as L. scabra]; Antonius et al., 1978 [LFK; as L. scabra]; Ring, 1980 [LFK; as L. scabra]; Voss et al., 1983 [UFK; as L. scabra and as L. sp. with “red and yellow tentacles”]; Sage, 1987 [LFK; as L. scabra]; Gilmour, 1990 [LFK; as С. scabra]; Lyons & Quinn, 1995 [as EL. scabra scabra]; Ripple, 1995 [as rough fileclam]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT], 2003 [UFK, MFK, LFK, DT]. Divarilima albicoma (Dall, 1886): Dall, 1889a, 1903b [as Lima]; Abbott, 1974 [as Lima]; Odé, 1979b; Diaz Merlano & Puyana Hegedus, 1994; Rios, 1994 [as Limaria]; Mikkelsen & Bieler, 2000. Lima caribaea Orbigny, 1842: Calkins, 1878 [DT; misidentified as Lima squamosa Lamarck, 1801, a synonym of L. lima Linnaeus, 1758, a recognized eastern Atlan- tic species]; Simpson, 1887-1889 [DT; as L. squamosa]; Dall, 1889a [as L. squamosal, 1903b [as L. squamosa]; Thiele, 1910 [DT; as L. lima (Linnaeus, 1758)]; Lermond, 1936 [as L. lima]; Lyman, 1948b [as L. lima]; Pul- ley, 1952; Eubanks, 1964 [as L. lima]; Brooks, 1968a [МЕК; as L. /ima]; Work, 1969 [LFK; as L. lima]; Magnotte, 1970-1979 [as L. lima]; Godcharles & Jaap, 1973 [UFK; as L. lima]; Voss, 1983 [UFK; as L. lima]; Voss CRITICAL CATALOG AND BIBLIOGRAPHY 999 et al., 1983 [UFK; as L. lima]; Williams, 1988 [as “spiny lima Lima lima”]; Krisberg, 1993 [LFK; as L. lima]; Lyons & Quinn, 1995 [as L. lima]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT], 2003 [UFK, MFK, LFK, DT]. |Lima lima Linnaeus, 1758 — see Lima caribaea. |Lima scabra (Born, 1778) — see Ctenoides scaber. |Lima tenera “Chemnitz” С. В. Sowerby ll, 1843 — see Ctenoides floridanus. Lima (s. |.) sp.: Henderson, 1911 [LFK]; Lyman, 1943; Voss, 1949 [UFK]; Ross, 1969 [МЕК]; Wingard et al., 1995 [UFK]; Brewster- Wingard et al., 1996 [UFK]; Vittor & Associ- ates, 1999а [ЧЕК], b [UFK, MFK]; USGS, 2003 [UFK; as Lima spp.]. Limaria locklini (T. L. McGinty, 1955): Vittor & Associates, 1999a [LFK; as Lima]. Limaria pellucida (C. B. Adams, 1846): Dall, 1886 [LFK; misidentified as Lima inflata Link, 1807, arecognized eastern Atlantic species], 1889a [misidentified as Lima inflata and Lima hians (Gmelin, 1791), a recognized eastern Atlantic species], 1903b [as Lima inflata]; Lermond, 1936 [as Lima inflata and Lima hians]; Webb, 1942, 1951 [both misidentified as Lima fragilis “Conrad” (?)]; Eubanks, 1964; Brooks, 1968b [MFK; as Lima]; Edwards, 1968b [LFK; as Lima]; lversen & Roessler, 1969 [UFK; as Lima]; Lee, 1969 [LFK; as Lima]; Ross, 1969 [MFK; as Lima]; Work, 1969 [LFK; as Lima]; Hudson et al., 1970 [UFK; as Lima]; Woods, 1971 [MFK; as Lima]; Turney & Perkins, 1972 [UFK, МЕК]; Antonius et al., 1978 [LFK; as Lima]; Voss, 1983 [UFK; as Lima]; Voss et al., 1983 [UFK; as Lima]; Lyons & Quinn, 1995 [as Lima]; Vittor & Associates, 1998 [LFK; as Lima], 1999a [UFK, DT; as Lima], b [UFK, MFK, LFK; as Lima]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK, MFK; as Limaria sp. cf. L. pellucida); Brewster-Wingard et al., 2001 [UFK; as Limaria cf. pellucida]. Limaria sp.: Stuardo, 1968 [UFK; as Limaria (Limatulella) sp. nov., but never published]. Limatula confusa (E. A. Smith, 1885): Lermond, 1936. Limatula setifera Dall, 1886: Dall, 1889a [as Lima (Limatula)]. Limatula subauriculata (Montagu, 1808): Dall, 1889a [as Lima (Limatula)]. Limea bronniana Dall, 1886. Limea sp.: Stuardo, 1968 [LFK; as Limea (Limea) sp. nov., but never published]. Limopsidae Theroux & Wigley, 1983 [ЕЕК]. Limopsis aurita (Brocchi, 1814): Dall, 1889a, 1903b; Mikkelsen & Bieler, 2000 [DT]. Limopsis cristata Jeffreys, 1876: Dall, 1889a, 1903b; Mikkelsen & Bieler, 2000 [DT]. Limopsis minuta Philippi, 1836: Dall, 1889a, 1903b; Mikkelsen & Bieler, 2000 [DT]. Limopsis sulcata A. E. Verrill & Bush, 1898: Mikkelsen & Bieler, 2000 [UFK, DT]. |Limopsis tenella Jeffreys, 1876 — all records based on Blake sta. 44 (here excluded; see entry for Dall, 1881). Lucinidae Vittor & Associates, 1997a [UFK, MFK, LFK], 1997с [ЧЕК], 1998 [LFK, DT], 1999a [UFK, ЕЕК, DT], b [UFK, MFK, ЕК]; USGS, 2003 [MFK]. Anodontia alba Link, 1807: Melvill, 1880 [LFK; as Loripes chrysostoma (“Meuschen” Philippi, 1847) and misidentified as L. edentula (Linnaeus, 1758), a recognized Micronesian species]; Dall, 1889a [as Loripes edentula and as L. e. var. chrysostoma]; 1903a [Pleistocene; as Lucina chrysostoma], b [as Loripes edentula and as L. e. var. chrysostoma “Mörch”]; Lermond, 1936 [as Lucina (Loripinus) edentula and as L. (L.) e. chrysostoma]; Bartsch, 1937; Britton, 1970 [МЕК, LFK, DT; as A. (Anodon- tia)]; Magnotte, 1970-1979; Turney & Perkins, 1972 [ЧЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK, МЕК]. Anodontia schrammi (Crosse, 1876): Lermond, 1936 [as Lucina (Loripinus)]; Britton, 1975 [LFK, DT; as A. (Anodontia)]; Turney & Perkins, 1972 [MFK; as A. philippiana (Reeve, 1850), a recognized Indo-Pacific species]; Mikkelsen & Bieler, 2000 [LFK; as A. philippianal. Callucina keenae Chavan in Cox et al., 1971: Britton, 1975 [LFK, DT; as Callucina (Callucina) radians Conrad, 1841, non Bory de St. Vincent, 1824]; Vittor 8 Associates, 1997с [UFK; as Lucina radians], 1999a [UFK, LFK, DT; as L. radians], b [UFK, MFK, ЕЕК; as L. radians]; Mikkelsen & Bieler, 2000 [UFK, DT; as L. radians]. Cavilinga blanda (Dall & Simpson, 1901): Dall, 1889a [misidentified as Lucina (Lucina) 600 trisulcata Conrad, 1841, and L. (L.) crenulata Conrad, 1845, two recognized Miocene- Pliocene fossil species from the eastern United States], 1903b [as L. (Lucina) trisulcata]; Lermond, 1936 [as L. (Anodon- tia) trisulcata]; Britton, 1975 [UFK, MFK, LFK, DT]; Turney & Perkins, 1972 [UFK, MFK; as Linga trisulcata]; Lyons & Quinn, 1995 [as Parvilucina]; Vittor 8 Associates, 1998 [LFK; as Lucina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; Lucina trisulcata]. Codakia orbicularis (Linnaeus, 1758): Melvill, 1880 [LFK; as Lucina tigerina (Linnaeus, 1758)]; Dall, 1883, 1885 [LFK; as L. tigrina (sic)], 1889a, 1903b [as L. (L.) tigrina (sic)], 1901, 1903a [Pleistocene]; Simpson, 1887- 1889 [LFK; as L. tigrina (sic)]; Nutting, 1895 [DT, as L. tigrina (sic)]; С. М. Vilas & М. В. Vilas, 1945, 1970; Morris, 1947, 1951; Pul- ley, 1952 [MFK]; Eubanks, 1964; Bender, 1965 [MFK, LFK]; Kissling, 1965 [LFK], 1977a [UFK], 1977b [UFK]; Brooks, 1968b [MFK]; Plockelman, 1968d, 1968e [МЕК]; Edwards, 1968b, 1980 [both LFK]; Jindrich, 1969 [LFK]; Ross, 1969 [MFK; as C. obicularis (sic)]; Work, 1969 [ЕЕК]; Britton, 1975 [UFK, MFK, LFK, DT; as С. (Codakia)]; Magnotte, 1970-1979; Stevenson, 1970, 1993 [both as lucine]; Woods, 1970 [LFK]; Turney & Perkins, 1972 [UFK, МЕК]; Godcharles & Jaap, 1973 [UFK]; Zischke, 1973, 1977a, с [МЕК]; Antonius et al., 1978 [LFK]; Schomer & Drew, 1982; Voss, 1983 [UFK]; Voss et al., 1983 [UFK; also as C. orbicular (sic)]; Sedlak, 1986 [LFK]; Clampit, 1987 [LFK]; Williams, 1988; Redla 1990 [UFK, LFK]; Lyons & Quinn, 1995; Tremor, 1998 [LFK]; Vittor & Associates, 1998 [LFK], 1999b [UFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK]; Orlin, 2003 [ЕЕК]; Bigatti et al., 2004 [UFK, MFK, ЕЕК]. Codakia (5. |.) sp.: Lyman, 19446 [ЧЕК]; Brewster-Wingard et al., 1996, 2001 [as Codakia spp.] [both ЧЕК]; Vittor & Associ- ates, 1998 [LFK, ОТ]; USGS, 2003 [UFK, МЕК; as Codakia spp.]. Ctena orbiculata (Montagu, 1808): Simpson, 1887-1889 [МЕК, ЕЕК; as Lucina squamosa Lamarck, 1806, and L. pecten Lamarck, 1818, yellow var.]; Dall, 1889a [as L. (Lucina) pecten and L. (L.) squamosa]; Dall, 1901 [as Чадота orbiculata var. filiata п. var. and J. O. var. recurvata n. var.]; Henderson, 1913 [UFK; as Codakia]; Johnson 1934 [as Codakia (Jagonia) orbiculata filiata and C. (Jagonia) orbiculata recurvata]; Lermond, MIKKELSEN 8 BIELER 1936 [as Lucina (J.), also as L. (J.) o. filiata and L. (J.) o. recurvata]; Aguayo 8 Jaume, 1949d [as Codakia]; lversen & Roessler, 1969 [UFK; as Codakia]; Lee, 1969 [LFK; as Codakia orbiculata and as С. о. ?form filiata]; Ross, 1969 [MFK; as Codakia]; Britton, 1975 [UFK, MFK, LFK, DT; as Codakia (Ctena)); Howard et al., 1970 [LFK; as Codakia]; Hudson et al., 1970 [UFK; as Codakia]; Turney & Perkins, 1972 [UFK, MFK; as Codakia]; Lineback, 1977 [LFK; as Codakia]; Schomer & Drew, 1982 [as Codakia]; Lyons 8 Quinn, 1995 [as Codakia]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Codakia]; Bigatti et al., 2004 [UFK, MFK, ЕЕК]; Morton 8 Knapp, 2004 [UFK, MFK]. Ctena pectinella (С. В. Adams, 1852): Dall, 1889a [as Lucina], 1903b [as Lucina (Lucina)]; Johnson 1934 [as Codakia (Jagonia)]; Lermond, 1936 [as L. (Jagonia)]; Britton, 1975 [UFK, LFK, DT; as Parvilucina (Parvilucina)]; Abbott, 1974 [as Codakia (Ctena)]; Hemmen 8 Hemmen, 1979 [as Codakia (Ctena)]; Lyons 8 Quinn, 1995 [as Codakia]; Mikkelsen 8 Bieler, 2000 [DT; as Codakia]. Divalinga quadrisulcata (Orbigny, 1842): Simpson, 1887-1889 [ОТ; as Lucina]; Dall, 1889a, 1903b [as Lucina (Divaricella)]; Britton, 1975 [UFK, MFK, LFK, DT; as Divaricella (Divalinga)]; Turney & Perkins, 1972 [UFK; as Divaricella]; Lyons & Quinn, 1995 [as Divaricella]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]. Divaricella dentata (Wood, 1815): Lermond, 1936 [as Lucina (D.)]; Britton, 1975 [UFK, МЕК, LFK, ОТ; as О. (Divaricella)]; Rehder, 1981 [as Lucina]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK]. Linga sp.: Vittor 8 Associates, 1998 [ЕЕК]. |Lucina leucocyma (Dall, 1886) — see under Pleurolucina. |Lucina pectinata (Gmelin, 1791) — see under Phacoides. Lucina pensylvanica (Linnaeus, 1758): Melvill, 1880 [LFK; as Lucina]; Simpson, 1887-1889 [LFK; as L. pennsylvanica (sic)]; Dall, 1889a, 1903b [as L. (L.) pennsylvanica (sic)]; Lermond, 1936 [as L. pennsylvanica (sic)]; Webb, 1942, 1951 [MFK]; Eubanks, 1964; Brooks, 1968b [MFK]; Ross, 1969 [MFK]; Britton, 1975 [UFK, MFK, LFK, DT; as Lucina (Lucina)]; Magnotte, 1970-1979; Woods, 1970 [LFK]; Turney 8 Perkins, 1972 [UFK, МЕК]; Godcharles & Jaap, 1973 [ЧЕК]; Zischke, 1973, 1977a [MFK]; Voss, 1983 [UFK; as Linga]; Voss et al., 1983 [UFK; as CRITICAL CATALOG AND BIBLIOGRAPHY 601 Linga]; Clampit, 1987 [LFK]; Redla 1990 [UFK, LFK]; Lyons & Quinn, 1995 [as Linga]; Tremor, 1998 [LFK]; Vittor & Associates, 1998 [LFK, DT]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]; Orlin, 2003 [LFK]; Bigatti et al., 2004 [UFK, МЕК, LFK]; Taylor et al., 2004 [UFK, MFK]. |Lucina radians Conrad, 1841 — see Callucina keenae. |Lucina sombrerensis (Dall, 1886) — see un- der Pleurolucina. Lucina sp.: Vittor & Associates, 1997c [UFK], 1998 [LFK, DT], 1999a [UFK, ЕЕК], b [MFK, ЕЕК]. Lucinisca muricata (Spengler, 1798): Simpson, 1887-1889 [DT; as [иста]; Dall, 1889a [as Lucina (Lucina) scabra Lamarck, 1819], 1901, 1903a [as Phacoides (Lucinisca) muricatus]; Johnson, 1934 [as L. (Lucinisca) muricata and L. scabra]; Lermond, 1936 [as Lucina (Lucinisca)]; M. Smith, 1937, 1945 [as Lucina (Cavilucina, Lucinisca)], 1940 [as Lucina muricatus]; Warmke 8 Abbott, 1961 [as Phacoides]; Aguayo & Jaume, 19490; Abbott, 1974 [LFK; as Lucina (Lucinisca)]; Humfrey, 1975 [as Phacoides]; Rehder, 1981 [LFK; as Phacoides]; Díaz Merlano 8 Puyana Hegedus, 1994 [as Lucina (Lucinisca)]; Rios, 1994 [as Lucina (Lucinisca)]; Lyons 8 Quinn, 1995 [as Lucina]; Vittor & Associates, 1999a [LFK; as Lucina], b [UFK, LFK; as Lucina]; Mikkelsen & Bieler, 2000 [LFK]. Lucinisca nassula (Conrad, 1846): Simpson, 1887-1889 [МЕК; as Lucina lintea Conrad, 1866]; Dall, 1889a [as Lucina (Lucina) lintea]; Howard et al., 1970 [LFK; as Lucina]; Britton, 1975 [UFK, MFK, LFK, DT; as Parvilucina (Lucinisca)]; Turney & Perkins, 1972 [UFK, MFK; as Phacoides]; Lineback, 1977 [LFK; as Lucina]; Lyons & Quinn, 1995 [as Lucina]; Vittor & Associates, 1998 [LFK, DT; as Lucina], 1999a [LFK; as Lucina], b [MFK, ЕЕК; as Lucina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Brewster-Wingard et al., 2001 [UFK]; USGS, 2003 [UFK]; Morton & Knapp, 2004 [UFK, MFK]. Lucinoma filosa (Stimpson, 1851): Dall, 1889a, 1903b [as Lucina (Lucina)]; Britton, 1975 [LFK, DT; as Phacoides (Lucinoma) filosus]; Mikkelsen & Bieler, 2000 [DT]. |Myrtea lens (A. Е. Verrill & Smith, 1880) — see under Myrteopsis. Myrtea sagrinata (Dall, 1886): Dall, 1889a, 1903b [as Lucina (Lucina)], 1901 [as M. (Eulopia)]; Johnson, 1934 [as М. (Eulopia)]; Abbott, 1974 [as M. (Eulopia)]; Ode, 1977b; Mikkelsen & Bieler, 2000. Myrteopsis lens (A. E. Verrill & Smith, 1880): Dall, 1889a, 1903b [both as Loripes]. Parvilucina costata (Orbigny, 1842): Simpson, 1887-1889 [DT; as Lucina]; Lermond, 1936 [as Lucina (Jagonia)]; Britton, 1975 [UFK, ЕЕК, DT; as P. (Parvilucina)]; Turney 8 Perkins, 1972 [UFK, МЕК; as Barnea]; Petersen, 1989 [МЕК; as Codakia]; Lyons & Quinn, 1995 [as Codakia]; Mikkelsen 8 Bieler, 2000 [as Codakia]. Parvilucina crenella (Dall, 1901): Dall, 1889a [as Lucina (L.) multilineata (“Conrad” Tuomey & Holmes, 1857)], 1903b [as L. (L.) multilineata]; Lermond, 1936 [as L. (Parvilucina)]; Britton, 1975 [UFK, MFK, LFK; as Р (Parvilucina) multilineata]; Turney & Perkins, 1972 [UFK; as L. multilineata]; Rehder, 1981 [as P multilineata]; Lyons & Quinn, 1995 [as P. multilineata]; Vittor & As- sociates, 1997с [UFK; as L. multilineata], 1998 [LFK, DT; as L. multilineata], 1999a [LFK, DT; as L. multilineata]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as P. multilineata]; Brewster-Wingard et al., 2001 [UFK; as Р multilineata]; USGS, 2003 [UFK; as Р multilineata]. Phacoides pectinata (Gmelin, 1791): Melvill, 1880 [LFK; as Lucina jamaicensis (Lamarck, 1801)]; Lermond, 1936 [as Lucina (Anodon- tia) jamaicensis]; Britton, 1975 [LFK; as Phacoides (Phacoides) pectinatus]; Magnotte, 1970-1979; Antonius et al., 1978 [LFK; as Phacoides pectinatus]; Lyons & Quinn, 1995 [as Lucina]; Vittor 8 Associates, 1999a [LFK; as Lucina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK; as Lucina]; USGS, 2003 [MFK; as Lucinal. Pleurolucina leucocyma (Dall, 1886): Dall, 1889a [as Lucina], 1903b [as Lucina (Lucina)]; Pulley, 1952 [UFK; as Lucina]; Britton, 1975 [UFK, MFK, LFK, DT; as Lucina (Pleurolucina)]; Rehder, 1981 [as Lucina]; Lyons & Quinn, 1995 [as Linga]; Mikkelsen & Bieler, 2000 [UFK, DT; as Lucina]. Pleurolucina sombrerensis (Dall, 1886): Dall, 1889а [as Lucina], 1903b [as Lucina (Lucina)]; Pulley, 1952 [MFK; as Lucina]; Britton, 1975 [UFK, MFK, LFK; as Lucina (Pleurolucina)]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Lucina]. |Pseudomiltha floridana (Conrad, 1833) — see under Stewartia. Radiolucina amianta (Dall, 1901): Dall, 1889a [as Lucina (L.) costata Tuomey & Holmes, 1857, non Orbigny, 1842]; Lermond, 1936 [as Lucina (Bellucina) amiantus]; Pulley, 1952 [LFK; as Lucina amiantus]; Britton, 602 MIKKELSEN & BIELER 1975 [UFK, LFK, DT; as Parvilucina (Bellucina) amiantus]; Rehder, 1981 [as Parvilucina]; Lyons & Quinn, 1995 [as Linga amiantus]; Vittor 8 Associates, 1998 [LFK, DT; as Lucina amiantus]; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT; as Lucina amiantus]. Stewartia floridana (Conrad, 1833): Dall, 1889a, 1903b [both ЕЕК; as Lucina (Lucina)]; Rogers, 1908 [as Lucina Floridana]; Aldrich 8 Snyder, 1936 [as Lucina]; Britton, 1975 [LFK; as Megaxinus floridanus]; Mikkelsen & Bieler, 2000 [LFK; as Lucina]. Lyonsiidae Entodesma beana (Orbigny, 1842): Dall, 1889a, 1903b [as Lyonsia Веапа]; Lermond, 1936 [as Lyonsia]; Lyons 8 Quinn, 1995; Vittor 8 Associates, 1998 [UFK; as Lyonsia]; Mikkelsen 8 Bieler, 2000 [UFK, МЕК, LFK, BT]: Lyonsia floridana Conrad, 1849: Hudson et al., 1970 [UFK; as L. hyalina floridana]; Vittor 8 Associates, 1998 [LFK, DT; as L. hyalina floridana], 1999a [LFK, DT; as L. hyalina floridana]; Mikkelsen & Bieler, 2000 [UFK, LEK; ЭТ]. Mactridae Vittor & Associates, 1998 [LFK]; USGS, 2003 [UFK, МЕК]. Anatina anatina (Spengler, 1802): Lermond, 1936 [as A. lineata (Say, 1822)]; Mikkelsen & Bieler, 2000 [MFK]. Mactrotoma fragilis (Gmelin, 1791): Dall, 1889a [as Mactra brasiliana Lamarck, 1818]; Lermond, 1936 [as Mactra]; Ross, 1969 [MFK; as Mactra]; Turney & Perkins, 1972 [MFK; as Mactra]; Lyons & Quinn, 1995 [as Mactra]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Mulinia lateralis (Say, 1822): Dall, 1889a [as Mactra]; Lermond, 1936; Magnotte, 1970- 1979. Raeta plicatella (Lamarck, 1818): Dall, 1889a [as Labiosa canaliculata Say, 1822]; Lermond, 1936 [as Anatina (Raeta) canaliculata]; Magnotte, 1970-1979 [as Anatina]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК}. Spisula raveneli (Conrad, 1831): Dall, 1889a, 1903b [аз Mactra solidissima var. similis Say, 1822]; Lermond, 1936 [as Spisula solidissima similis]; Mikkelsen & Bieler, 2000 [UFK, MFK]. Malleidae Malleus candeanus (Orbigny, 1842): Bales, 1944 [as Fundella candeana]; Boss & Moore, 1967 [UFK, ЕЕК, DT; аз М. (Parimalleus)]; Waller & Mcintyre, 1982 [DT]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, LEK; BT]. Myidae Sphenia fragilis (H. & A. Adams, 1854): Mikkelsen & Bieler, 2000 [UFK; as Sphenia antillensis Dall & Simpson, 1901]. Mytilidae Stevenson, 1970, 1993 [both as mussel]: Woods, 1971 [MFK; as little mussels]; Gaertner, 1978 [LFK; as mussels]; Ring, 1980 [LFK; as mussels]; Edwards, 1987 [LFK; as Ribbed Mussel]; Vittor & Associates, 1997c [UFK], 1998 [DT], 1999b [MFK, LFK]. Amygdalum papyrium (Conrad, 1846): Lermond, 1936 [as Modiolaria arborescens (Dillwyn, 1817); this species name is vari- ously considered as a synonym of A. dendriticum Muhlfeld, 1811, or (as auctt., non Dillwyn) a synonym of A. papyrium; it is con- servatively listed here]; Pulley, 1952 [also as A. arborescens]; Mikkelsen & Bieler, 2000 [UFK, MFK]. Amygdalum politum (A. E. Verrill & Smith, 1880): Dall, 1881, 1886 [DT; as Modiola polita], 1889a, 1903b [as M. (Amygdalum) polita]; Bartsch, 1937 [LFK; as Modiola polita]; Mikkelsen 8 Bieler, 2000 [DT]. Amygdalum sagittatum (Rehder, 1935): Dall, 1889a, 1903b [as Modiola (Amygdalum) polita var. sagittata (nomen nudum)]; Fos- ter, 1945 [as Modiolus politus sagittatus]; Vittor & Associates, 1998 [LFK, DT], 1999a [LFK]; Mikkelsen & Bieler, 2000 [LFK, DT]. Amygdalum sp.: Мог & Associates, 1999a [DT]. Botula fusca (Gmelin, 1791): Simpson, 1887- 1889 [DT; as Modiolaria cinnamomea Lamarck, 1819]; Dall, 1889a [as Modiola (Botula) cinnamomea]; Bales, 1940, 1944; Lermond, 1936; Pulley, 1952; Edwards, 1968a [LFK]; Brooks, 1969 [MFK]; Ross, 1971 [LFK]; Emerson & Jacobson, 1976; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Valentich-Scott & Dinesen, 2004 [MFK, LFK]. |Brachidontes domingensis (Lamarck, 1819) — see B. exustus. CRITICAL CATALOG AND BIBLIOGRAPHY 603 Brachidontes exustus (Linnaeus, 1758): Dall, 1883, 1885 [LFK; as Mytilus], 1889a, 1903b [as Mytilus]; Simpson, 1887-1889 [LFK; as Mytilus]; Simpson, 1887-1889 [LFK; as Mytilus lavalleanus Orbigny, 1842]; Lermond, 1936 [as Mytilus]; T. A. Stephenson & A. Stephenson, 1950 [MFK, LFK; as M. (Brachidontes)]; Ginsburg, 1952 [UFK, MFK]; Iversen 8 Roessler, 1969 [UFK; also as Trachidontes ($/с)]; Ross, 1969 [MFK]; Work, 1969 [UFK]; Hudson et al., 1970 [UFK]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, MFK]; Zischke, 1973, 1977a, b, с [MFK]; Ingham & Zischke, 1977 [MFK]; Antonius et al., 1978 [LFK; as Brachiodontes (sic)]; Schomer & Drew, 1982 [аз Mytilus]; Petersen, 1989 [MFK; as Brachiodonta (sic)]; Lyons & Quinn, 1995 [as B. domingensis (Lamarck, 1819)]; Wingard etal., 1995 [UFK; as Brachiodontes (sic) sp.]; Brewster-Wingard et al., 1996, 1997, 1998 [as Brachiodontes (sic) sp.], 2001 [all ЧЕК]; Vittor & Associates, 1997c [ЧЕК]; Lyons, 1998 [UFK, MFK]; Brewster-Wingard & Ishman, 1999 [as Brachiodontes (sic) sp.], 2001 [both UFK]; Lyons, 1999; Mikkelsen € Bieler, 2000 [UFK, MFK, LFK; also as B. domingensis]; USGS, 2003 [UFK, MFK]. Brachidontes modiolus (Linnaeus, 1767): Calkins, 1878 [LFK; as Modiola sulcata Lamarck, 1819]; Melvill, 1880 [LFK; as Mytilus cubitus Say, 1822]; Dall, 1889a, 1903b [both as Modiola (Brachydontes) sulcata]; Lermond, 1936 [as Modiolaria sulcatus]; Webb, 1937, 1939, 1942, 1951 [LFK; figured, as Mytilus perna “Dall” (Linnaeus, 1758)] Pulley, 1952 [as Brachidontes citrinus (Róding, 1798)]; Lee, 1969 [LFK; as B. citrinus]; Plockelman, 1969b [LFK, as В. citrinus]; Ross, 1969 [MFK; as B. citrinus]; Plockelman, 1970c [as В. citrinus]; Edwards, 1980 [LFK; as Brachidonta (sic)]; Petuch, 1988 [ЧЕК]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Brachidontes sp.: Morrison, 1958 [UFK]; Oliver 8 Járnegren, 2004 [ЕЕК]. Crenella decussata (Montagu, 1808): Dall, 1889a, 1903b [as C. divaricata (Orbigny, 1845)]; Lermond, 1936 [as С. divaricata]; Turney 8 Perkins, 1972 [UFK, MFK; as C. divaricata]; Lyons & Quinn, 1995 [as С. divaricata]; Vittor 8 Associates, 1998 [LFK, ОТ; as С. divaricata], 1999a [LFK, ОТ; as С. divaricata], 1999b [UFK, MFK, LFK; as C. divaricata]; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, ОТ]. Crenella sp.: Morrison, 1958 [ЧЕК]. Dacrydium elegantulum hendersoni Salas and Gofas, 1997: Dall, 1889a, 1903b [misidentified as D. vitreum (Moller, 1842, ex Holbgll ms), a recognized North Atlantic species]; Lermond, 1936 [as D. vitreum]; Salas and Gofas, 1997 [LFK]; Mikkelsen € Bieler, 2000 [ЕЕК]. Geukensia granosissima (С. В. Sowerby Ш, 1914): Melvill, 1880 [LFK; as Modiola plicatula Lamarck, 1819]; Lermond, 1936 [as Modiolaria demissus (Dillwyn, 1817) and M. d. granosissimus]; Mikkelsen & Bieler, 2000 [MFK]. Gregariella coralliophaga (Gmelin, 1791): Simpson, 1887-1889 [LFK, ОТ; as Botula semen (“Reeve” err. pro Lamarck, 1819)]; Dall, 1889a [as Modiola (Botulina) opifex Say, 1825]; Lermond, 1936 [as Modiolaria opifex]; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK]. Ischadium recurvum (Rafinesque, 1820): Dall, 1889а, 1903b [as Mytilus hamatus Say, 1822]; Lermond, 1936 [as Mytilus recurvus]; Magnotte, 1970-1979 [as Brachidontes recurvus]; Kissling, 1977a [as Brachiodontes (sic) recurvus]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК]. Lioberus castaneus (Say, 1822): Simpson, 1887—1889 [DT; as Pectunculus]; Dall, 1889a [as Modiola (Amygdalum) lignea Reeve, 1858]; Lermond, 1936 [as Botula castanea]; Pulley, 1952 [as Botula]; Andrews, 1971, 1977, 1981a, b, 1992, 1994 [as Lioberis (sic)]; Antonius et al., 1978 [ЕЕК]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [DT], 1999b [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Lithophaga antillarum (Orbigny, 1842): Calkins, 1878 [DT; as Lithodomus]; Dall, 1889a [as Lithophagus], 1896a [DT; as Lithophagus], 1903b [as Lithophagus]; Lermond, 1936 [as Lithodomus]; Bales, 1940, 1944; Webb, 1942 [LFK], 1951 [LFK; as L. antilarum (sic)]; Lyman, 1948b; Pulley, 1952; Turner & Boss, 1962 [UFK, МЕК, ЕЕК]; Brooks, 1968а [MFK]; Magnotte, 1970- 1979; Zischke, 1973, 1977a [MFK]; Emerson & Jacobson, 1976; Kissling, 1977a; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Valentich-Scott & Dinesen, 2004 [UFK, МЕК, LFK]. Lithophaga aristata (Dillwyn, 1817): Melvill, 1880 [LFK; as Lithodomus candigerus, err. pro L. caudigerus (Lamarck, 1819)]; Simpson, 1887-1889 [DT; as Lithodomus 604 forficatus Ravenel, 1861]; Dall, 1889a [as Lithophagus forficatus]; Lermond, 1936 [as Lithodomus]; Bales, 1944; Turner 8 Boss, 1962 [LFK, DT]; Emerson & Jacobson, 1976; Antonius et al., 1978 [ЕЕК]; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [LFK, DT]; Valentich-Scott & Dinesen, 2004 [LFK]. Lithophaga bisulcata (Orbigny, 1842): Simpson, 1887-1889 [MFK; as Lithodomus bisulcatus]; Dall, 1889a, 1903b [аз Lithophagus bisulcatus]; Aldrich & Snyder, 1936; Lermond, 1936 [as Lithodomus]; Bales, 1940, 1944; Turner & Boss, 1962 [MFK]; Antonius et al., 1978 [LFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]; Valentich-Scott & Dinesen, 2004 [MFK, LFK]. Lithophaga nigra (Orbigny, 1842): Calkins, 1878 [DT; misidentified as Lithodomus lithophagus (Linnaeus, 1758), a recognized eastern Atlantic species]; Simpson, 1887- 1889 [DT; as Lithodomus niger]; Dall, 1889a [as Lithophagus caribaeus Philippi, 1847]; Rogers, 1908 [as Lithodomus lithophagus]; Thiele, 1910 [DT]; Lermond, 1936 [as Lithodomus]; Bales, 1940, 1944; Turner & Boss, 1962 [UFK, MFK, LFK, DT]; Edwards, 1968a [LFK; as Lithophagis (sic) nigra]; Zischke, 1973, 1977a [MFK]; Emerson & Jacobson, 1976; Antonius et al., 1978 [LFK]; Kleemann, 1983; Voss et al., 1983 [ЧЕК]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT]. Lithophaga sp.: Henderson, 1911 [LFK; as Lithodomus sp.]; Lyman, 1943; Schomer & Drew, 1982; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]. Modiolus americanus (Leach in Leach & Nodder, 1815): Calkins, 1878 [LFK, as Modiola tulipa Lamarck, 1819; MFK, as М. t. var. nigra п. уаг.]; Lermond, 1936 [as Modiolaria tulipus]; Lyman, 1947a [МЕК; as Modiolus tulipus]; lversen & Roessler, 1969 [UFK]; Ross, 1969 [MFK]; Work, 1969 [LFK]; Magnotte, 1970-1979; Woods, 1970 [ЕЕК]; Turney & Perkins, 1972 [UFK, MFK; as Volsella americana]; Godcharles & Jaap, 1973 [UFK]; Zischke, 1973, 1977a [MFK]; Antonius et al., 1978 [LFK]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT]. Modiolus squamosus Beauperthuy, 1967: Beauperthuy, 1967 [LFK]; Voss, 1983 [UFK; as М. modiolus squamosus]; Voss et al., 1983 [UFK; аз М. modiolus squamosus]; Lyons 8 Quinn, 1995 [as M. modiolus MIKKELSEN 8 BIELER squamosus]; Vittor 8 Associates, 1997c [UFK], 1998 [LFK], 1999b [LFK] [all as M. modiolus squamosus]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as M. modiolus squamosus]; USGS, 2003 [UFK, МЕК]. Musculus lateralis (Say, 1822): Dall, 1889a [as Modiolaria], 1903b [as Modiolaria]; Thiele, 1910 [DT; as Modiolaria]; Lermond, 1936 [as Modiolaria]; Webb, 1942, 1951 [as Modiola duplicata (Say, [date unknown)); although this name has not been verified as a synonym and Webb's (1942: 74, pl. 25, fig. 30) figure is poor, the figure and brief text suggest M. lateralis]; Pulley, 1952; Voss, 1983 [ЧЕК]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1997с [UFK], 1998 [LFK, ОТ], 1999a [DT], b [UFK, MFK, ЕЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Musculus sp.: Vittor & Associates, 1999а [ЧЕК]. |Mytilus californianus Conrad, 1837: Abbott, 1961 [LFK; in error]. |Mytilus perna (Linnaeus, 1758) — see under Brachidontes Modiolus. |Perna viridis (Linnaeus, 1758): Benson et al. 2001 (predicted introduction). |Neilonellidae |Neilonella pusio (Philippi, 1844) — Florida Keys records (Abbott, 1974 [LFK; as Nuculana]; Mikkelsen & Bieler, 2000 [LFK; as Nuculana]) based on Dall’s (1889a, 1903b [as Leda (Leda)]) archibenthal category (50- 800 fms); however species’ minimum depth range given by Dall (1889a) is 856 [fms], and it is here excluded as beyond depth limit. Noetiidae Arcopsis adamsi (“Shuttleworth” Dall, 1886): Dall, 1889a [as Arca (Byssoarca) Adamsi]; Lermond, 1936 [as Arca admsi (sic)]; M. Smith, 1937, 1945 [as Arca (Acar)]; Edwards, 1968b [LFK]; Hudson et al., 1970 [UFK]; Woods, 1970; Turney & Perkins, 1972 [UFK, MFK]; Zischke, 1973, 1977a, b, с [MFK]; Antonius et al., 1978 [LFK; as Acropsis (sic)]; Mikkelsen, 1981 [UFK]; Petersen, 1989 [MFK; as Acropsi (sic)]; Lyons & Quinn, 1995; Wingard et al., 1995 [UFK]; Brewster- Wingard et al., 1996, 1997, 2001 [all UFK]; Vittor & Associates, 1997c [UFK]; Brewster- Wingard & Ishman, 1999, 2001 [ЧЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK, МЕК]; Oliver & Jarnegren, 2004 [МЕК, ЕЕК]. CRITICAL CATALOG AND BIBLIOGRAPHY 605 Noetia ponderosa (Say, 1822): Dall, 1889a, 1903b [as Arca (Noetia)]; Lermond, 1936 [as Arca]; Abbott, 1954 [LFK; as N. (Eontia)], 1961 [LFK], 1968, 1970 [LFK]; Siekman, 1965, 1981, 1982 [ЕЕК]; Magnotte, 1970- 1979; Andrews, 1971, 1977, 1981a, b, 1992, 1994 [LFK; as N. (Eontia)]; Turney & Perkins, 1972 [UFK, MFK]; Emerson & Jacobson, 1976 [LFK]; Long Island Shell Club, 1988; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Nuculanidae Theroux & Wigley, 1983 [UFK] |Ledella solidula (E. A. Smith, 1885): Florida Keys records (Lermond, 1936, as Nuculana]; Rios, 1994 [as N. (Jupitaria)]; Mikkelsen & Bieler, 2000 [as Nuculana]) probably based on Dall (1889a, 1903b [as Leda (Leda)]), categorized as archibenthal (50-800 fms) but with minimum species depth as 640 [fms]; here excluded as beyond depth limit. Ledella sublevis A. E. Verrill & Bush, 1898: Dall, 1889a [misidentified as Leda (Leda) messanensis Seguenza, 1877, a recognized eastern Atlantic species of Yoldiella; some material identified as Y. messanensis was subsequently described as the new species L. bushae Warén, 1978, but all cited mate- rial is post-1889; L. sublevis was originally described as a western Atlantic variety of Y. messanensis]. Nuculana acuta (Conrad, 1832): Dall, 1886 [LFK; as Leda], 1889a, 1903b [as Leda (Leda)]; Turney & Perkins, 1972 [UFK, MFK]; Vittor & Associates, 1998 [LFK, DT], 1999a [LFK, DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Nuculana concentrica (Say, 1824): Vittor & Associates, 1998 [LFK, DT], 1999a [DT]; Mikkelsen & Bieler, 2000 [MFK, LFK, DT]. Nuculana jamaicensis (Orbigny, 1842): Dall, 1881 [LFK; as Leda]. Nuculana solidifacta (Dall, 1886). Nuculana verrilliana (Dall, 1886): Johnson, 1934; Lermond, 1936; Morris, 1973; Abbott, 1974; Mikkelsen & Bieler, 2000. Nuculana vitrea (Orbigny, 1842): Dall, 1889a [as Leda (Leda)]. Nuculana sp.: Theroux & Wigley, 1983 [МЕК, LFK]; Bielsa & Labisky, 1987 [LFK]; Vittor & Associates, 1998 [LFK, DT]. Propeleda carpenteri (Dall, 1881): Dall, 1881 [LFK; as Leda], 1889a [as L. (L.) Carpenter; Lermond, 1936 [as Nuculana]; Pulley, 1952 [LFK; as Nuculana]; Theroux & Wigley, 1983 [UFK; as Nuculana]. Nuculidae Ennucula aegeensis (Forbes, 1844): Dall, 1889а, 1903b [both as Nucula aegeénsis]; Lermond, 1936 [as Nucula]; Vittor & Associ- ates, 1997a [UFK, МЕК, ЕЕК; as N. aegeenis (sic)], 1997c [UFK; as N. aegensis (sic)], 1998 [LFK, DT; as N. aegeenis (sic)], 1999a [DT; аз N. aegeenis (sic)], 1999b [LFK; as N. aegeenis (sic)]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Nucula]. Ennucula tenuis Montagu, 1808: Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Nucula calcicola Moore, 1977: Moore, 1977 [UFK]; Diaz Merlano & Puyana Hegedus, 1994; Mikkelsen & Bieler, 2000 [UFK, МЕК, LEKI: Nucula crenulata A. Adams, 1856: Abbott, 1954 [LFK]; Mikkelsen & Bieler, 2000 [DT]; other “Tortugas” records are based on Blake sta. 44, here excluded (see entry for Dall, 1881). Nucula proxima Say, 1822: Pulley, 1952; Howard et al., 1970 [LFK]; Turney & Perkins, 1972 [UFK, MFK]; Lineback, 1977 [LFK]; Wingard et al., 1995 [UFK]; Brewster- Wingard et al., 1996, 1997, 2001 [all UFK]; Brewster-Wingard & Ishman, 1999 [ЧЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, ОТ]; USGS, 2003 [UFK]. Ostreidae Woods, 1971 [MFK; as little oysters]; USGS, 2003 [UFK, MFK]; Oliver & Järnegren, 2004 [LFK; as oysters]. Crassostrea rhizophorae (Guilding, 1828): Melvill, 1880 [LFK; as Ostrea]; Simpson, 1887-1889 [DT; misidentified as O. parasitica “Lamarck” err. pro Gmelin, 1791, a recognized Indo-Pacific species of Striostrea]; Mikkelsen & Bieler, 2000 [LFK]. Crassostrea virginica (Gmelin, 1791): Dall, 1889a, 1903b [as Ostrea]; Lermond, 1936 [as O. verginica (sic)]; Bartsch, 1937 [as Ostrea floridensis С. В. Sowerby Il, 1871]; Chan, 1977b [LFK]; Schomer & Drew, 1982; Mikkelsen & Bieler, 2000 [UFK, LFK]. |Cryptostrea permollis (С. В. Sowerby Il, 1871) — see Teskeyostrea weber. Dendostrea frons (Linnaeus, 1758): Melvill, 1880 [LFK; as Ostrea]; Dall, 1889a, 1903b [as Ostrea]; Lermond, 1936 [as Ostrea]; Webb, 1937, 1942, 1951 [all as Ostrea foliata Lamarck [date unknown]; although the taxo- nomic status of this name 1$ unverified, Webb's illustrations strongly suggest D. frons; ?err. pro Ostrea folium Linnaeus, 1758, 606 MIKKELSEN & BIELER a recognized Indo-West Pacific species of Dendostrea; Schroeder, 1964 [UFK; as “coon oysters”]; Magnotte, 1970-1979 [as Ostrea]; Voss, 1983 [UFK; as Lopha and as Ostrea]; Voss et al., 1983 [UFK; as Lopha, Ostrea and as Ostrea (Lopha)]; Boone, 1986 [MFK]; Petersen, 1989 [MFK; as Ostrea]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Kirkendale et al., 2004. Ostrea equestris (Say, 1834): Dall, 1889a [misidentified as Ostrea cristata Born, 1778, a recognized South American species]; Lermond, 1936 [as Ostrea cristata]; Magnotte, 1970-1979; Zischke, 1973, 1977a, b [MFK]; Lyons & Quinn, 1995 [as Ostreola]; Campbell et al., 1998 [МЕК; as Ostreola]; Jozefowicz & O Foighil, 1998 [LFK; misidentied as Teskeyostrea weberi, fide Kirkendale et al., 2004]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK; as Ostreola]; USGS, 2002 [UFK]; Kirkendale et al., 2004 [MFK, LFK; as Ostreola]. Teskeyostrea weberi (Olsson, 1951): Olsson, 1951 [MFK, LFK; as Ostrea]; Forbes, 1964 [UFK; misidentified as Ostrea permollis С. В. Sowerby Il, 1871, a recognized obligate sponge commensal of Cryptostrea from the western Atlantic]; Kraeuter, 1973 [UFK; as Ostrea permollis]; Harry, 1985 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК; as Ostrea permollis]; Kirkendale et al., 2004 [МЕК]. Pandoridae Pandora arenosa Conrad, 1834: Vittor & As- sociates, 1999a [LFK, DT]. Pandora bushiana Dall, 1886: Boss & Merrill, 1965 [UFK, DT]; Mikkelsen & Bieler, 2000 ЕК РТ Pandora glacialis Leach, 1819: Dall, 1889a, 1903b [аз Р. (Kennerlia)]. Pandora inflata Boss & Merrill, 1965: Boss & Merrill 1965 [УЕК, MER ЕЕК, ЭТ] Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT]. Pandora sp.: Vittor 8 Associates, 1998 [LFK, DT], 1999a [ЧЕК]. Pectinidae Henderson, 1911 [LFK; as Pectens]; Lyman, 19446 [UFK; as pectens], 1951 [UFK, as black pecten]; Foster, 1945; Turney & Perkins, 1972 [UFK, MFK]; Edwards, 1980 [LFK; as Pecten pairs]; Theroux 8 Wigley, 1983 [MFK]; Wingard et al., 1995 [UFK; as pectinid]; Brewster-Wingard et al., 1997 [UFK; as pectinid]; Vittor & Associates, 1998 [DT], 19996 [MFK, ЕЕК]; USGS, 2002 [ЧЕК]. |Aequipecten acanthodes Dall, 1925 — see Lindapecten muscosus. Aequipecten glyptus (A. E. Verrill, 1882): Rehder & Abbott, 1951 [DT]; Rombouts, 1991 [DT]; Mikkelsen & Bieler, 2000 [LFK, BE Aequipecten heliacus (Dall, 1925): Lermond, 1936 [as Рецепт]. Aequipecten lineolaris (Lamarck, 1819): Abbott, 1954; Р. L. McGinty & T. Е. McGinty, 1957 [MFK, LFK]; Warmke 8 Abbott, 1961; Barrett & Patterson, 1967; Magnotte, 1970- 1979; Pompey, 1974; Humfrey, 1975; Romashko, 1984; Mikkelsen & Bieler, 2000 [МЕК, ЕЕК, DT; as Argopecten]. |Aequipecten exasperatus (С. В. Sowerby Il, 1847) — see Lindapecten muscosus. |Aequipecten muscosus (Wood, 1828) — see under Lindapecten. |Aequipecten phrygius (Dall, 1886) — see Cryptopecten phrygium. |Amusium laurentii (Gmelin, 1791) — see un- der Euvola. |Amusium papyraceum (Gabb, 1873) — see Euvola cf. papyracea. Argopecten gibbus (Linnaeus, 1758): Dall, 1886 [LFK; as Pecten dislocatus Say, 1822], 1889а, 1903b [as Р (Pecten) irradians var. dislocatus]; Simpson, 1887-1889 [MFK; as P. dislocates (sic)]; Lermond, 1936 [as Pecten]; Waller, 1969; Magnotte, 1970-1979 [as Aequipecten]; Godcharles 8 Jaap, 1973 [UFK]; Allen, 1979 [LFK]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Krause et al., 1994 [LFK]; Lyons & Quinn, 1995; Vittor & Asso- ciates, 1998 [DT]; Mikkelsen & Bieler, 2000 [ОРКИ MEK. ERK ВТ]: Argopecten irradians (Lamarck, 1819): lversen & Roessler, 1969 [UFK; also as Aequipecten or Aeguipecten (sic)]; Waller, 1969 [MFK; as A. i. concentricus (Say, 1822)]; Hudson et al., 1970 [UFK; as A. i. concentricus]; Magnotte, 1970-1979 [as Aequipecten]; Petuch, 1987 [MFK; as A. i. taylorae Petuch, 1987], 1988 [MFK; as A. i. taylorae]; Marelli et al., 1997 [MFK; as A. i. concentricus and A. i. taylorae]; Turgeon et al., 1988; Abbott, 1974; Lyons & Quinn, 1995; DeMaria, 1996 [UFK, MFK; as scallops]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]; Brewster-Wingard et al., 2001 [UFK]; USGS, 2003 [UFK, MFK]. CRITICAL CATALOG AND BIBLIOGRAPHY 607 Argopecten nucleus (Born, 1778): Dall, 1889a, 1903b [as Pecten (Pecten)], 1898 [as P. (Chlamys, section Plagioctenium) gibbus var. nucleus]; 1925 [as P. (Plagioctenium)]; Johnson 1934 [as Pecten (Plagioctenium)]; Е егтопа, 1936 [as Pecten]; M. Smith, 1937, 1945 [as Pecten (Aequipecten, Plagio- ctenium)]; Lyman, 1944a [as Pecten]; Webb, 1951 [as Pecten]; Salisbury, 1952 (as Chlamys); Abbott, 1954 [as Aequipecten (Plagioctenium) gibbus nucleus]; Bender, 1968 [LFK]; Waller, 1969; Godcharles 8 Jaap, 1973 [UFK]; Petersen, 1989 [MFK; as Argopectin (sic)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Argopecten sp.: Vittor & Associates, 1999a [LFK]. Brachtechlamys antillarum (Récluz, 1853): Simpson, 1887—1889 [LFK, DT; as Pecten]; Dall, 1886 [as Pecten], 1889a, 1903b [LFK; as Рецепт (Pecten)], 1898 [as Р (Chlamys, section Nodipecten)]; Johnson, 1934 [as Pecten (L.)]; Lermond, 1936 [as Pecten]; М. Smith, 1937, 1945 [as Pecten (Lyropecten)|; Webb, 1942, 1951 [LFK; as Pecten]; Voss, 1949 [UFK; as Pecten], 1983 [UFK; as Lyropecten]; Pulley, 1952 [as Lyropecten]; lversen & Roessler, 1969 [UFK; as Lyropecten]; Magnotte, 1970-1979 [as Lyropecten]; Woods, 1970 [LFK; аз Рецепт]; Ross, 1971 [LFK; as Lyropecten]; Voss et al., 1983 [UFK; as Lyropecten and as Pecten]; Wagner 8 Abbott, 1990 [as Lyropecten]; Lyons & Quinn, 1995; Hutsell et al., 1997 [as Lyropecten]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, ОТ]. Caribachlamys imbricata (Gmelin, 1791): Simpson, 1887-1889 [DT; as Pecten imbricatus]; Dall, 1889a, 1903b [DT; as Р (P.) imbricatus]; Thiele, 1910 [DT; as P. (Chlamys) imbricatus]; Johnson 1934 [as P. (Chlamys) imbricatus]; Lermond, 1936 [as P. imbricatus]; M. Smith, 1937, 1945 [as P (Chlamys)]; Lyman, 1944a, 1948a, b [as P imbricatus]; Voss, 1948 [UFK; as P. imbricatus]; Bippus, 1950 [UFK; as P. imbricatus]; Webb, 1951 [as P. imbricatus]; Pulley, 1952 [as Chlamys]; Edwards, 1969 [UFK; as Chlamys]; Work, 1969 [UFK, ЕЕК; as Chlamys]; Magnotte, 1970-1979 [as Chlamys]; Antonius et al., 1978 [LFK; as Chlamys]; Sage, 1987 [LFK; as Chlamys]; Williams, 1988; Waller, 1993 [UFK]; Lyons & Quinn, 1995 [as Chlamys]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Caribachlamys mildredae (F. M. Bayer, 1941): Bayer, 1941 [DT, as Pecten (Chlamys) imbricatus var. mildredae], 1942 [UFK, LFK, DT; as Pecten imbricatus mildredae]; Lyman, 1945 [UFK; as P (Chlamys) Mildredaea (sic)]; М. Smith, 1945 [as Pecten]; Magnotte, 1970- 1979 [as Chlamys]; Waller, 1993 [DT]; Lyons 8 Quinn, 1995 [as Chlamys]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, ОТ]. Caribachlamys ornata (Lamarck, 1819): Simpson, 1887-1889 [LFK, DT; as Pecten ornatus and as purplish var.]; Dall, 1889a [as P. (P.) ornatus], 1896a [DT; as P. ornatus]; 1903b [as Р (P.) ornatus]; 1898 [Pleistocene; as P. (Chlamys, section Chlamys) ornatus]; Nutting, 1895 [DT; as Р ornatus]; Lermond, 1936 [as Р ornatus]; Lyman, 1944a [as P ornatus]; Webb, 1951 [as Pecten]; Edwards, 1969 [UFK; as Chlamys]; Work, 1969 [UFK; as Chlamys]; Waller, 1993 [ЧЕК]; Lyons & Quinn, 1995 [as Chlamys]; Gundersen, 1997; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Caribachlamys sentis (Reeve, 1853): Lermond, 1936 [LFK; as Pecten]; Webb, 1942 [LFK; as Pecten], 1951 [as Pecten]; Lyman, 1943 [as Pecten]; Lyman, 1944a, 1948b, 1949a [as Pecten], 1950 [UFK; as Pecten]; Voss, 1948, 1949 [UFK; as Pecten]; Bippus, 1950 [UFK; as Pecten]; Pulley, 1952 [as Chlamys]; Eubanks, 1964 [as Chlamys]; Bender, 1968 [LFK]; Brooks, 1968b [MFK; as Chlamys]; Edwards, 1968a, b [both ЕЕК], 1969 [UFK] [all as Chlamys]; Brooks, 1969 [МЕК; as Chlamys]; Burggraf, 1969 [LFK; as Chlamys]; Magnotte, 1970-1979 [as Chlamys]; Plockelman, 1969b [LFK, as Chlamys]; Ross, 1969 [MFK; as Chlamys], 1971 [LFK; as Chlamys]; Zischke, 1973, 1977a, с [МЕК; as Chlamys]; Hughes, 1976 [LFK; as Chlamys]; Kissling, 1977b [UFK; as Chlamys]; Antonius et al., 1978 [LFK; as Chlamys]; Pease, 1980 [LFK; as Chlamys]; Rehder, 1981 [as Chlamys]; Romashko, 1984 [as Chlamys]; Sage, 1987 [LFK; as Chlamys]; Williams, 1988 [as Chlamys]; Wagner 8 Abbott, 1990 [as Chlamys]; Will- iams, 1990 [as Chlamys]; Krisberg, 1993 [LFK; as Chlamys]; Waller, 1993 [UFK]; Lyons & Quinn, 1995 [as Chlamys]; Gundersen, 1997; Hutsell et al., 1997; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT]; Poland, 2001 [ЕЕК]. |Chlamys benedicti A. E. Verrill 8 Bush, 1897 — see under Spathochlamys. |Chlamys imbricata (Gmelin, 1791) — see un- der Caribachlamys. |Chlamys multisquamata (Dunker, 1864) — see under Laevichlamys. 608 MIKKELSEN & BIELER |Chlamys mildredae (Е. M. Bayer, 1941) — see under Caribachlamys. |Chlamys ornata (Lamarck, 1819) — see un- der Caribachlamys. |Chlamys sentis (Reeve, 1853) — see under Caribachlamys. Chlamys (s. |.) sp.: Lyman, 1943 [as Pecten (Chlamys)]. Cryptopecten phrygium (Dall, 1886): Dall, 1889a, 1903b [as Pecten (Pecten)]; Pulley, 1952 [UFK, ОТ; as Chlamys phrygius]; Abbott, 1954 [LFK; as Aequipecten phrygius]; P. Е. McGinty & T. Е. McGinty, 1957 [MFK, LFK; as Pecten]; Abbott, 1974 [as Aequipecten]; Hayami, 1984 [LFK]; Mikkelsen & Bieler, 2000 [MFK, ЕЕК, DT]. Euvola chazaliei (Dautzenberg, 1900): Dall, 1925 [LFK; as Pecten (E.) tereinus Dall, 1925]; Johnson, 1934 [LFK; as P. (E.) tereinus]; Lermond, 1936 [LFK; as P. tereinus]; Foster, 1945 [as Pecten tereinus]; Aguayo 8 Jaume, 1950h [LFK; as P. (E.) tereinus]; Grau, 1955 [LFK; as P. (E.) tereinus]; Р. L. McGinty & Т.Е. McGinty, 1957 [MFK,LFK; as Р chazaliei (tereinus)]; Boss et al., 1968 [LFK; as P (E.) tereinus]; Lyons & Quinn, 1995 [as Pecten]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Euvola laurentii (Gmelin, 1791): Frumar, 2000 [ОТ; as Amusium]; Mikkelsen & Bieler, 2000 [LFK, DT; as Amusium]. Euvola cf. papyracea (Gabb, 1873) [= Euvola sp. Aof Waller (1991); Amusium papyraceum, a Dominican Republic fossil, is a true Amusium and not conspecific]: Rehder & Abbott, 1951 [DT; as Amusium раругасеит]; Pulley, 1952 [DT; as A. раругасеит]; Mikkelsen & Bieler, 2000 [LFK, DT; as А. раругасеит]. Euvola raveneli (Dall, 1898): Simpson, 1887- 1889 [misidentified as Pecten “hemicyclica”, err. pro Р hemicyclicus “Ravenel” Tuomey & Holmes, 1855, a recognized Pliocene Euvola from South Carolina]; Dall, 1889a [as Pecten (Janira) hemicyclica]; Lermond, 1936 [as Pecten]; Pulley, 1952; Mikkelsen & Bieler, 2000 [MFK, LFK, DT]. Euvola ziczac (Linnaeus, 1758): Dall, 1889a, 1903b [as Pecten (Janira)]; Lermond, 1936 [as Pecten]; Magnotte, 1970-1979 [as Pecten]; Godcharles & Jaap, 1973 [UFK; as Pecten]; Rombouts, 1991 [LFK]; Lyons & Quinn, 1995 [as Pecten]; Mikkelsen & Bieler, 2000 [LFK, DT]; Miller, 2001 [LFK]. Laevichlamys multisquamata (Dunker, 1864): Dall, 1889a [as Pecten (Pecten) effluens Dall, 1886]; Lyons & Quinn, 1995 [as Chlamys]; Mikkelsen & Bieler, 2000. Lindapecten muscosus (Wood, 1828): Dall, 1889a, 1903b [both misidentified as Pecten (Pecten) exasperatus С. В. Sowerby Il, 1847, a recognized Caribbean species of Lindapecten]; Lermond, 1936 [as P. exasperatus and P. acanthodes Dall, 1925]; С. М. Vilas & N. R. Vilas, 1945 [as Pecten], 1970 [as Aequipecten]; Webb, 1951 [as Pecten]; Pulley, 1952 [as Chlamys]; lversen & Roessler, 1969 [UFK; as Aequipecten]; Magnotte, 1970-1979 [as Aequipecten]; Plockelman, 1970b [MFK; as Aequipecten acanthodes]; Andrews, 1971, 1977, 1994 [as Aequipecten]; Sunderland & Cahill, 1990 [MFK; as A. acanthodes]; Wagner & Abbott, 1990 [as Aequipecten, also as A. acanthodes]; Rombouts, 1991 [LFK; as Aequipecten]; Abbott & Morris, 1995 [as A. acanthodes]; Lyons & Quinn, 1995 [as A. acanthodes exasperatus]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK; also as L. exasperatus]. |Lyropecten antillarum (Récluz, 1853) — see under Brachtechlamys. Lyropecten kallinubilosus (Е. М. Bayer, 1943): Mikkelsen & Bieler, 2000 [LFK]. Nodipecten fragosus (Conrad, 1849): Dall, 1889a, 1903b [both misidentified as Pecten (Pecten) nodosus (Linnaeus, 1758), a rec- ognized Caribbean species of Nodipecten]; Lermond, 1936 [as P. nodosus fragosus, also as Р nodosus]; T. L. McGinty, 1942 [as P nodosus]; Pulley, 1952 [as Lyropecten nodosus]; Magnotte, 1970-1979 [DT; as Lyropecten nodosus]; Andrews, 1971, 1977, 1981a, b, 1992, 1994 [all as Lyropecten (Nodipecten) nodosus]; Gaertner, 1978 [LFK, as Lion’s Paws]; Sunderland, 1988 [UFK; as N. nodosus]; Williams, 1988 [as N. nodosus]; Rombouts, 1991 [DT; as N. nodosus]; Lyons & Quinn, 1995 [as N. nodosus]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК, DT; as N. nodosus]. |Nodipecten nodosus (Linnaeus, 1758) — see Nodipecten fragosus. |Ресеп chazaliei (Dautzenberg, 1900) — see under Euvola. |Pecten raveneli Dall, 1898 — see under Euvola. |Pecten ziczac (Linnaeus, 1758) — see under Euvola. Pecten (s. |.) sp.: Henderson, 1911 [ЕЕК]; Voss, 1983 [UFK]; Voss et al., 1983 [ЧЕК]. Spathochlamys benedicti (A. E. Verrill 8 Bush, 1897): Lyons & Quinn, 1995 [as Chlamys]; Frumar, 2000 [DT; as Chlamys]; Mikkelsen & Bieler, 2000 [LFK, DT]. CRITICAL CATALOG AND BIBLIOGRAPHY 609 Periplomatidae Cochlodesma pyramidatum Stimpson, 1860: Aguayo & Jaume, 1949 f. Periploma margaritaceum (Lamarck, 1801): Simpson, 1887-1889 [DT; as Р angulifera (Philippi, 1847)]; Dall, 1889a, 1903a, b [as P. angulifera]; Johnson 1934 [as angu- liferum]; Lermond, 1936 [as P. angulifera (sic)]; M. Smith, 1937, 1945 [both as P. angulifera]; Abbott, 1974 [as P anguliferum]; Abbott & Morris, 1995 [as P. anguliferum]; Mikkelsen & Bieler, 2000 [LFK; as P. anguliferum]. Periploma tenerum P. Fischer, 1882: Dall, 1889a [as Р. tenera “Jeffreys”]; Johnson 1934 [as P tenera]; Lermond, 1936 [as P tenera]; Mikkelsen & Bieler, 2000 [МЕК]. Petricolidae Choristodon robustum (С. В. Sowerby |, 1834): Simpson, 1887-1889 [LFK, DT; as C. typicum Jonas, 1844]; Dall, 1889a, 1903b [as Petricola (Choristodon) robusta]; Bales, 1940 [as Rupellaria typical], 1944 [as R. typica]; Plockelman, 1969c [MFK; as К. typical; Lyons & Quinn, 1995 [аз К. typical; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Valentich-Scott & Dinesen, 2004 [ЕЕК]. Choristodon sp.: Valentich-Scott & Dinesen, 2004 [LFK; as С. sp. А]. Cooperella atlantica Rehder, Plockelman, 1970d [ЕЕК]. Petricola lapicida (Gmelin, 1791): Dall, 1886 [Gordon Key; as P. divaricata Chemnitz in Orbigny, 1842], 1889a, 1903b [as P. (Naranaio)]; Simpson, 1887-1889 [DT; as P. divaricata]; Johnson, 1934; Lermond, 1936; M. Smith, 1937, 1945; Bales, 1940, 1944; Edwards, 1968a [LFK]; Zischke, 1973, 1977a [MFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Valentich-Scott & Dinesen, 2004 [LFK]. Petricolaria pholadiformis (Lamarck, 1818): Рай, 1889a, 1903b [as Petricola]; Lermond, 1936 [as Petricola]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [ЕЕК]. Petricolaria sp.: Dall, 1889a, 1903b [as Petricola]. |Rupellaria typica (Jonas, 1844) — see Choristodon robustum. 1943: Pharidae Ensis minor Dall, 1900: Dall, 1889a [misidentified as E. americana (Gould, 1870), a synonym of the recognized but much larger E. directus Conrad, 1843, from the eastern United States], 1899b [UFK; as Е. directus], 1900a [UFK; as Е. directus]; Maury, 1920 [as E. directus]; M. Smith, 1937, 1940, 1945 [as E. directus]; Mikkelsen & Bieler, 2000 [ЕЕК]. Philobryidae Cratis antillensis (Dall, 1881): Dall, 1889a, 1903b (both as Limopsis). Pholadidae Barnea truncata (Say, 1822): Dall, 1889a, 1903b [as Pholas (Ватеа)]; Lermond, 1936 [as Pholas (Barnea)]; M. Smith, 1937, 1940, 1945; С. М. Vilas & М. R. Vilas, 1945, 1970; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000. Cyrtopleura costata (Linnaeus, 1758): Dall, 1889a, 1903b [as Pholas (Barnea)]; Lermond, 1936 [as Pholas (Barnea)]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [ЕЕК]. Martesia cuneiformis (Say, 1822): Simpson, 1887-1889 [MFK, DT]; Dall, 1889a, 1903b; Lermond, 1936 [also as M. caribaea (Orbigny, 1842)]; Turner, 1955 [LFK]; Mikkelsen & Bieler, 2000 [ЕЕК]. Martesia striata (Linnaeus, 1758): Simpson, 1887—1889 [DT; as Coralliophaga horn- beckiana, err. pro Pholas hornbeckii Orbigny, 1842]; Dall, 1889a [also as M. corticata Adams, err. pro corticaria “Gray” С. В. Sowerby Il, 1849], 1903b; Lermond, 1936; Turner, 1955 [LFK]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК]. Pholas campechiensis Gmelin, 1791: Dall, 1889a, 1903b; Lermond, 1936. IXylopholas altenai Turner, 1972 — Florida Keys records (Abbott, 1974; Mikkelsen & Bieler, 2000 [as X. altanai (sic)]) based on original description, here excluded as be- yond depth limit). Pinnidae Stevenson, 1970, 1993 [both as pen shell]; Clampit, 1987 [LFK; as “two types of Pen Shells”]; USGS, 2003 [UFK, MFK]. Atrina rigida (Lightfoot, 1786): Lermond, 1936 [as Pinna]; Turner & Rosewater, 1958 [MFK, LFK]; Kissling, 1965 [LFK], 1977b [UFK]; Iversen & Roessler, 1969 [UFK]; Magnotte, 610 MIKKELSEN & BIELER 1970-1979; Zischke, 1973, 1977a, b, c [МЕК]; Antonius et al., 1978 [ЕЕК]; Schomer & Drew, 1982; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК]. Atrina seminuda (Lamarck, 1819): Dall, 1889a, 1903b [as Pinna]; Schomer & Drew, 1982; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК]. Atrina serrata (G. B. Sowerby |, 1825): Simpson, 1887-1889 [as Pinna muricata Linnaeus, 1758]; Dall, 1889a [as P. muricata]; Lermond, 1936 [as Pinna]; Turner & Rosewater, 1958 [UFK, MFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Atrina sp.: Turney & Perkins, 1972 [MFK]. Pinna сатеа Gmelin, 1791: Simpson, 1887— 1889 [LFK, DT]; Dall, 1889a, 1896a [DT; as Pinna pernula Chemnitz, 1785], 1898 [post- Pleistocene], 1903b; Pilsbry, 1890a [LFK]; Dall, 1897; Thiele, 1910 [DT]; Lermond, 1936; M. Smith, 1937, 1945 [DT]; Lyman, 1944c, 1946; Turner & Rosewater, 1958 [UFK, MFK, LFK, DT]; Edwards, 1968a [LFK]; Ross, 1969 [MFK]; Work, 1969 [LFK, DT]; Magnotte, 1970-1979; Sedlak, 1986 [LFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; also misidentified as Pinna rudis Linnaeus, 1758, a recognized Mediterranean species]. Pinna sp.: Kirkendale et al., 2004 [MFK]. Plicatulidae Plicatula gibbosa Lamarck, 1801: Dall, 1889a [аз P. ramosa Lamarck, 1819]; Lermond, 1936; Webb, 1942, 1951 [as Plicatula mantilla Conrad [date unknown]) (not veri- fied as a зупопут)]; Magnotte, 1970-1979; Turney & Perkins, 1972 [MFK]; Antonius et al., 1978 [LFK; as P. spondyloidea Meuschen, 1781]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, Dm] Poromyidae |Cetoconcha margarita (Dall, 1886) - all records based on Blake sta. 44 (here ex- cluded; see entry for Dall, 1881), including Mikkelsen & Bieler, 2000. Poromya albida Dall, 1886: Dall, 1889a [as Poromya (Cetomya)]. |Poromya elongata Dall, 1886 — only Florida Keys record (Dall, 1889a) probably based on Gulf of Mexico, 199 fms (Abbott, 1974; here excluded as beyond depth limit. Poromya granulata (Nyst & Westendorp, 1839): Dall, 1881 [LFK], 1889a, 1903b, 1927; Johnson, 1934 [as Р. д. granulata]; Lermond, 1936; Mikkelsen & Bieler, 2000 [DT]. Poromya rostrata Rehder, 1943: Mikkelsen & Bieler, 2000 [MFK]. Propeamussiidae Cyclopecten nanus A. E. Verrill & Bush, 1897. Cyclopecten strigillatus (Dall, 1889): Dall, 1889b [as Pseudamusium]. Cyclopecten sp.: Mikkelsen & Bieler, 2000 [DT]. Parvamussium cancellatum (E. A. Smith, 1885): Dall, 1889a [as Pecten (Amusium, Propeamussium)]. Parvamussium thalassinum (Dall, 1886): Dall, 1889a [as Pecten (Pecten, Pseudamusium)]. |Propeamussium dalli (Е. A. Smith, 1885) — only Florida Keys record (Mikkelsen & Bieler, 2000) based on specimens beyond depth limit). Propeamussium pourtalesianum (Dall, 1886): Dall, 1881 [LFK; as Amussium lucidum (Jeffreys in Thompson, 1873)], 1889a [as Pecten (Amusium, Propeamussium) Pourtalesianum and as var. marmoratum (Dall, 1881)], 1903b [as Pecten (Amusium, Propeamussium) Pourtalesianum]; Mikkelsen & Bieler, 2000 [DT]. Propeamussium sayanum (Dall, 1886): Dall, 1889a, 1903b [as Pecten (Amusium, Propeamussium) Sayanum]; Mikkelsen & Bieler, 2000 [DT]. Psammobiidae Asaphis deflorata (Linnaeus, 1758): Melvill, 1880 [LFK; as Asaphis dichotoma (Anton, 1839), synonym of A. violascens (Forsskäl, 1775), a recognized Indo-Pacific species]; Simpson, 1887-1889 [DT]; Dall, 1889a, 1903b; Pilsbry, 1890b [UFK]; Lermond, 1936; Magnotte, 1970-1979; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК}; Domaneschi & Shea, 2004 [LFK]. Gari circe (Mórch, 1876): Simpson, 1887-1889 [MFK; misidentified as Macoma anomala (Deshayes, 1855), a recognized Indo-Pacific species of Сам. Heterodonax bimaculatus (Linnaeus, 1758): Simpson, 1887-1889 [DT]; Dall, 1889a, 1903b [as H. bimaculata]; Lermond, 1936 [as H. bimaculata]; Mikkelsen & Bieler, 2000. Sanguinolaria sanquinolenta (Gmelin, 1791): Calkins, 1878 [DT]; Zischke, 1973, 1977a [MFK]. CRITICAL CATALOG AND BIBLIOGRAPHY 611 Pteriidae Vittor 8 Associates, 1999b [UFK, МЕК]. Pinctada imbricata Róding, 1798: Simpson, 1887-1889 [DT; as Avicula radiata Leach, 1814, also as A. ala-perdicis Reeve, 1857]; Dall, 1889a [as Margaritiphora radiata], 1896a [DT; as A. radiata and as A. crocata (Swainson, 1831), a possible synonym de- scribed from Ceylon], 1903b [as Margaritiphora radiata]; Thiele, 1910 [DT; as Pteria radiata]; Lermond, 1936 [as Margaritifera radiata]; Webb, 1937 [misidentified as Pedalion alata Gmelin, 1791, now Isognomon alatus (see above)]; C.N. Vilas & N. R. Vilas, 1945 [as Pinctada radiata], 1970; Bippus, 1950 [UFK; as Pteria radiata (variety?)]; Pulley, 1952 [as Pinctada radiata]; lversen & Roessler, 1969 [ЧЕК]; Ross, 1969 [MFK; as Pinctada radiata]; Work, 1969 [ЕЕК]; Magnotte, 1970-1979 [as Pinctada radiata]; Woods, 1970 [LFK; as Pinctada radiata]; Hayes, 1972 [UFK, MFK, ЧЕК, DT]; Zischke, 1973, 1977a [both МЕК]; Chan, 1977a, b [LFK; as Pinctada radiata]; Kissling, 1977b [UFK; as Pinctada radiata]; Lineback, 1977 [LFK; as P radiata]; Antonius et al., 1978 [LFK; as Pinctada radiata]; Webster, 1978 [LFK]; Voss, 1983 [UFK; as Pinctada radiata]; Voss et al., 1983 [UFK; as Pinctada radiatal; Sedlak, 1986 [ЕЕК]; Shirai, 1994 [LFK]; Lyons & Quinn, 1995; Campbell et al., 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]; Orlin 2003; Mikkelsen et al., 2004. Pinctada longisquamosa (Dunker, 1852): Hudson et al., 1970 [UFK; misidentified as Pinctada radiata (Leach, 1814), a synonym of P. imbricata (above)]; Hayes, 1972 [UFK, MFK, ЕЕК, DT; as Репа]; Turney & Perkins, 1972 [UFK, MFK; misidentified as Pinctada radiata, fide Brewster-Wingard et al., 2001]; Abbott, 1974; Wingard et al., 1995 [UFK; as Pinctada radiata]; Brewster-Wingard et al., 1996, 1997, 1998 [all as Pinctada radiatal, 2001 [as Репа] [all ЧЕК]; Brewster-Wingard 8 Ishman, 1999 [UFK; as Pinctada radiata]; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT]; USGS, 2003 [UFK, MFK; as Репа]; Mikkelsen et al., 2004 [UFK, MFK, LFK, DT]. Pinctada margaritifera (Linnaeus, 1758): Nut- ting, 1895 [DT; as Avicula] [This distinctive Indo-Pacific species, widely appreciated for its natural and cultured pearls, has been re- cently and irrefutably recorded as introduced into the western Atlantic (Chesler, 1994; Carlton, 1996), but is not believed to have become established; if it can be believed, Nutting’s record would establish a consider- ably earlier date of first introduction.]. Репа colymbus (Róding, 1798): Calkins, 1878 [as Avicula atlantica Lamarck, 1819]; Dall, 1889a [as A. atlantica], 1896a [DT; as A. atlantica], 1903b [as A. atlantica]; Lermond, 1936 [as A. atlantica]; Eubanks, 1964; Iversen & Roessler, 1969 [UFK]; Work, 1969 [LFK]; Hayes, 1972 [UFK, MFK, LFK, DT]; Magnotte, 1970-1979; Stevenson, 1970, 1993 [аз Atlantic wing oyster], 1993; Zischke, 1977a [MFK]; Lyons 8 Quinn, 1995; Hutsell et al., 1997; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Orlin, 2003; Mikkelsen et al., 2004. |Репа longisquamosa (Dunker, 1852) — see under Pinctada. Pteria vitrea (Reeve, 1857): Dall, 1889a [as Avicula nitida A. E. Verrill, 1880]; Maury, 1920 [DT]; Pearse, 1929 [DT]; Johnson, 1934 [as P. hirundo vitrea]; Hayes, 1972 [UFK, DT; as Pteria hirundo vitrea]; Abbott, 1974. Репа (3. |.) sp.: Calkins, 1878 [DT; as Avicula]; Dall, 1881 [DT; as Avicula]; Henderson, 1911 [LFK; as Avicula]. Semelidae Vittor & Associates, 1997c [ЧЕК], 1998 [LFK, DT], 1999a [LFK, DT], 1999b [UFK, ЕЕК]. Abra aequalis (Say, 1822): Lermond, 1936; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK, DT], 1999a [LFK]; Mikkelsen 8 Bieler, 2000 [UFK, LFK, DT]. Abra lioica (Dall, 1881): Dall, 1881 [LFK; as Syndosmya], 1886 [LFK], 1889a, 1903b; Lermond, 1936; Turney & Perkins, 1972 [UFK, MFK]; Vittor & Associates, 1999b [LFK]; Mikkelsen & Bieler, 2000 [MFK, LFK]. Abra longicallis americana A. E. Verrill & Bush, 1898: Dall, 1889a [as A. longicallus, err. pro A. longicallis (Scacchi, 1836)]. Abra sp.: Theroux & Wigley, 1983 [UFK, MFK, LFK]; Vittor & Associates, 1999b [UFK]. Cumingia coarctata G. B. Sowerby |, 1833: Dall, 1900a; Lermond, 1936; M. Smith, 1937, 1945; Olsson & Harbison, 1952 [LFK]; Pul- ley, 1952 [LFK; as C. antillarum (Orbigny, 1842)]; Abbott, 1954 [ЕЕК], 1958 [LFK]; Warmke & Abbott, 1961 [LFK; as C. antillarum]; Edwards, 1968a [LFK; as Cummingia (sic) antillarum]; Emerson & Jacobson, 1976 [as С. antillarum, a “form” of С. tellinoides (Conrad, 1831)]; Lyons & 612 MIKKELSEN & BIELER Quinn, 1995; Vittor & Associates, 1998 [DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, Dill: Cumingia vanhyningi Rehder, 1939: Simpson, 1887—1889 [MFK, DT; as С. tellinoides (Conrad, 1831)]; Dall, 1889a, 1903b [both as С. tellinoides]; Lermond, 1936 [аз С. tellinoides]; Rehder, 1939 [UFK, LFK; as C. tellinoides vanhyningil; Aguayo & Jaume, 1948а [UFK; as С. vanhyningil; Turney & Perkins, 1972 [UFK, MFK; as С. tellinoides]; Wingard et al., 1995 [UFK; as С. tellinoidea (sic)]; Brewster-Wingard et al., 1996 [as С. tellinoidea (sic)], 1997, 2001 [both as C. tellinoides] [all ЧЕК]; Vittor & Associates, 1998 [LFK; as С. tellinoides], 1999b [LFK; as С. tellinoides]; Sweeney 8 Harasewych, 1999 [UFK; as С. tellinoides vanhyningi]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK; as С. tellinoides vanhyningi]; USGS, 2003 [UFK; as С. tellinoides]. Cumingia sp.: USGS, 2003 [UFK; as Cumingia sp. or spp.]. Ervilia concentrica (Holmes, 1860): Simpson, 1887-1889 [MFK, DT]; Dall, 1896b, 1889a, 1903b [LFK]; Johnson, 1934 [as Gould, 1862]; Lermond, 1936; M. Smith, 1937, 1945; Davis, 1973 [LFK, ОТ]; Lyons & Quinn, 1995; Vittor 8 Associates, 1998 [LFK, ОТ], 1999a [LFK, DT], b [UFK, MFK]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, ЕЕК, DT]. Ervilia nitens (Montagu, 1806): Simpson, 1887-1889 [DT]; Dall, 1896b, 1889a, 1903b [DT]; Johnson, 1934; Lermond, 1936; M. Smith, 1937, 1945; Davis, 1973 [MFK, LFK]; Lyons & Quinn, 1995; Vittor & Associates, 1999b [UFK, MFK, LFK]; Mikkelsen & Bieler, 2000 [MFK, LFK]. Ervilia subcancellata E. A. Smith, 1885: Davis, 1973 [UFK, MFK, LFK, DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Ervilia sp.: Vittor & Associates, 1999b [MFK]. Semele bellastriata (Conrad, 1837): Dall, 1889a [аз $. cancellata (Orbigny, 1842)]; Lermond, 1936; M. Smith, 1937, 1945 [LFK]; С. М. Vilas & М. В. Vilas, 1945, 1970 [ЕЕК]; Lyman, 1949c; T. L. McGinty, 1955; Boss, 1972 [UFK, LFK, DT; as $. bellestriata (sic)]; Andrews, 1994; Lyons & Quinn, 1995; Wingard et al., 1995 [UFK]; Vittor & Associ- ates, 1998 [LFK], 1999a [UFK, DT], b [UFK, ЕЕК]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; Brewster-Wingard et al., 2001 [UFK]; USGS, 2003 [ЧЕК]. Semele proficua (Pulteney, 1799): Simpson, 1887—1889 [as $. reticulata “Lamarck” Spengler, 1798]; Dall, 1889a [as S. reticulata “Gmelin” Spengler, 1798]; Lermond, 1936; Lyman, 1949c [also as S. radiata (Say, 1826) and $. radiata “dark form”]; Boss, 1972 [UFK, MFK, LFK, DT]; Turney & Perkins, 1972 [MFK]; Lyons & Quinn, 1995; Vittor & Associ- ates, 1998 [DT], 1999a [LFK, DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Semele purpurascens (Gmelin, 1791): Dall, 1886 [DT], 1889a, 1903b [as S. obliqua (Wood, 1815, non J. Sowerby, 1817)]; Simpson, 1887-1889 [DT; as $. obliqua]; Lermond, 1936; Lyman, 1949c; Boss, 1972 [LFK, DT]; Lyons & Quinn, 1995; Vittor € Associates, 1999a [LFK, DT]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Semele sp.: Vittor 8 Associates, 1999a [UFK, LFK], b [UFK, MFK]. Semelina nuculoides (Conrad in Hodge, 1841): Lermond, 1936 [as Semele]; Boss, 1972 [LFK; as Semele (Semelina)]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK, ОТ; as Semele], 1999a [UFK, ЕЕК, DT; as Semele], b [MFK, LFK; as Semele]; Mikkelsen & Bieler, 2000 [LFK, DT]. Solecurtidae Solecurtus cumingianus (Dunker, 1861): Mikkelsen & Bieler, 2000 [MFK, LFK, DT]. Tagelus divisus (Spengler, 1794): Dall, 1889a, 1895a [LFK], 1903b; Lermond, 1936; Magnotte, 1970-1979; Turney & Perkins, 1972 [MFK]; Krisberg, 1993 [LFK]; Vittor & Associates, 1999b [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Tagelus plebeius (Lightfoot, 1786): Bartsch, 1937 [as T. gibbus (Spengler, 1794)]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [ЧЕК]. Tagelus зр.: Voss, 1983 [ЧЕК]; Voss et al., 1983 [ЧЕК]; USGS, 2003 [UFK, MFK; as Tagelus spp.]. Solemyidae Solemya occidentalis Deshayes, 1857: Dall, 1889a, 1903b [as Solenomya]; Lermond, 1936; Lyons & Quinn, 1995; Vittor 8 Associ- ates, 1997a [MFK, LFK], 1998 [LFK, ОТ]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Solemya velum Say, 1822. Solemya sp.: Vittor 8 Associates, 1998 [DT]. Spondylidae ?Thiele, 1910 [DT; misidentified as Pecten (Chlamys) pusio Linnaeus, 1758, a recog- CRITICAL CATALOG AND BIBLIOGRAPHY 613 nized eastern Atlantic species of Hinnites; possibly a Spondylus sp.]. Spondylus americanus Hermann, 1781: Lermond, 1936 [also as S. echinatus Martyn, 1784]; М. Smith, 1937, 1945; С. М. Vilas & М. К. Vilas, 1945, 1970 [ЕЕК]; Bender, 1968 [MFK]; Mason, 1969 [ЕЕК]; Teskey, 1969 [LFK]; Work, 1969; Magnotte, 1970-1979; Artman, 1974 [LFK]; Colin, 1978; Sunderland, 1988 [UFK]; Williams, 1988; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; also as $5. gussoni O. G. Costa, 1829, based on misidentified specimens]. Spondylus ictericus Reeve, 1856: Calkins, 1878 [DT; misidentified as S. gaederopus Linnaeus, 1758, a recognized Mediterranean species]; Simpson, 1887-1889 [misidentified as $. croceus “Chemnitz”, err. pro Lamarck, 1819, a recognized Indo-Pacific species; and DT, misidentified as S. spathuliferus Lamarck, 1819, a recognized Indo-Pacific species]; Melvill, 1880 [LFK; as S. ramosus Reeve, 1856; Dall, 1889а [as 5. spathuliferus]; Edwards, 1968a [LFK]; Work, 1969 [UFK, LFK]; Plockelman, 1970a; Goldberg, 1978с [ЕЕК]; Magnotte, 1970- 1979 [misidentified аз $. gussoni]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, ОТ]. Spondylus зр.: Т. Е. McGinty, 1939 [ЕЕК], 1942; Jaap, 1984; Boone, 1986 [MFK]. Sportellidae Basterotia elliptica (Récluz, 1850): Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000. Basterotia quadrata (Hanley, 1843): Dall, 1889а, 1903b [as В. quadrata “Hinds”]; Lermond, 1936 [also as В. q. granatina (Dall, 1881)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000. Ensitellops protexta (Conrad, 1841): Lermond, 1936 [UFK; as Egeta]. Tellinidae Brooks, 1968a [МЕК; as tellins], b [MFK; as tellins]; Lee, 1969 [LFK]; Vittor & Associates, 1997а [UFK, МЕК, ЕЕК], 1997с [ЧЕК], 1998 [LFK, DT], 1999а [ЕЕК], b [UFK, MFK]; Brewster-Wingard et al., 2001 [UFK; as tellinid]. Acorylus gouldii (Hanley, 1846): Simpson, 1887-1889 [DT; as Tellina]; Dall, 1889a [as Tellina], 1903b [as T. Gouldií]; Lermond, 1936 [as Т. gouldi]; Boss, 1966 [UFK, MFK, LFK, DT; as Tellina (Acorylus)]; Lyons 8 Quinn, 1995 [as Tellina]; Vittor 8 Associates, 1999b [UFK, MFK; as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, ОТ; as Tellina]. Angulus agilis (Stimpson, 1857): Calkins, 1878 [as Tellina tenera Say, 1822]; Melvill, 1880 [LFK; as 7. tenera]; Dall, 1886 [LFK; as Tellina], 1889a [as Tellina], 1903b [as 7. tenera]; Lermond, 1936 [as T. tenera]; Mikkelsen & Bieler, 2000 [LFK; as Tellina]. Angulus merus (Say, 1834): Dall, 1883, 1885 [LFK; as Tellina], 1889a, 1903b [both as Tellina]; Simpson, 1887-1889 [LFK, DT; as Tellina]; Lermond, 1936 [as Tellina, also as Macoma leptonoides, err. pro M. leptonoidea Dall, 1895, a synonym of Macoma carlotten- sis Whiteaves, 1880, a recognized Califor- nian species, superficially similar to A. mera]; Pulley, 1952 [MFK; as Tellina]; Boss, 1968b [UFK, MFK, LFK, DT; as Tellina (Angulus)]; Ross, 1969 [МЕК; as Tellina]; Howard et al., 1970 [LFK; as 7. cf. mera]; Magnotte, 1970- 1979 [as Tellina]; Turney 8 Perkins, 1972 [UFK, MFK; as Tellina]; Lineback, 1977 [LFK; as T. cf. mera]; Petersen, 1989 [MFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Vittor & Associates, 1998 [LFK, ОТ; as Tellina], 1999b [LFK; as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT; as Tellina]; Morton & Knapp, 2004 [UFK, MFK; as Tellina]. Angulus paramerus (Boss, 1964): Boss, 1964 [LFK, DT; as Tellina (Angulus)]; Ross, 1969 [МЕК; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT; as Tellina]. Angulus probrinus (Boss, 1964): Boss, 1964 [МЕК, LFK, DT; as Tellina (Angulus)]; Boss, 1968b [MFK, LFK, DT; as Tellina (Angulus)]; Lyons & Quinn, 1995 [as T. probina (sic)]; Mikkelsen & Bieler, 2000 [MFK, LFK, DT; as Tellina]. Angulus sybariticus (Dall, 1881): Boss, 1968b [UFK, LFK, DT; as Те/та (Angulus)]; Lyons & Quinn, 1995 [as Tellina]; Vittor 8 Associ- ates, 1997c [UFK; as Tellina], 1998 [LFK, DT; as Tellina], 1999a [LFK, ОТ; as Tellina]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT; as Tellina]. Angulus tampaensis (Conrad, 1866): Henderson, 1913 [UFK]; Boss, 1968b [UFK, LFK; as Tellina (Angulus)]; Hudson et al., 1970 [UFK; as Tellina]; Magnotte, 1970- 1979 [as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Lyons, 1998 [UFK, MFK; as Tellina]; 614 MIKKELSEN & BIELER Vittor & Associates, 1999b [UFK; as Tellina]; Mikkelsen & Bieler, 2000 [UFK, LFK; as Те!та]. Angulus tenellus (А. E. Verrill, 1874): Dall, 1889a [as Те/та modesta А. Е. Verrill, 1872, non Carpenter 1864]; Lermond, 1936 [as Т. modesta]; Vittor & Associates, 1999b [МЕК; as Tellina]. Angulus texanus (Dall, 1900): Lermond, 1936 [as Tellina sayi“Deshayes” Dall, 1900]; Boss, 1968b [UFK, MFK, LFK; as Tellina (Angu- lus); Turney & Perkins, 1972 [UFK, MFK; as Tellina]; Petersen, 1989 [MFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Vittor 8 Associates, 1998, 1999а [both ОТ, as Tellina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Tellina]. Angulus versicolor (“Cozzens” DeKay, 1843): Lermond, 1936 [as Tellina]; Pulley, 1952 [MFK; as Tellina]; Boss, 1968b [UFK, MFK, ЕЕК, DT; as Tellina (Angulus)]; Abbott, 1974 [LFK; as Tellina (Angulus)]; Lyons & Quinn, 1995 [as Tellina]; Vittor & Associates, 1998 [LFK; as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, ОТ; as Tellina]. Arcopagia fausta (Pulteney, 1799): Melvill, 1880 [LFK; as Tellina, also as Tellina robusta Hanley, 1844, a recognized Indo-Pacific, pos- sibly misidentified juvenile 7. fausta; Simpson, 1887-1889 [LFK; as Macoma fausta “Dillwyn”]; Dall, 1889a, 1903b [as Tellina]; Lermond, 1936 [as Tellina]; M. Smith, 1937, 1945 [LFK; as Т. (Acropagia, Cyclotellina)]; Eubanks, 1964; Bender, 1965 [MFK, LFK; as Tellina]; Boss, 1966 [UFK, MFK, LFK, DT; as Tellina (Arcopagia)]; Bender, 1968 [MFK, LFK]; Brooks, 1968a [MFK], b [MFK; as Tellina]; Ross, 1969 [MFK]; Work, 1969 [DT: as Tellina]; Woods, 1970 [ЕЕК]; Turney & Perkins, 1972 [MFK]; Emerson & Jacobson, 1976 [as Tellina]; Goldberg, 1978c [LFK; as Tellina]; Antonius et al., 1978 [ЕЕК]; Edwards, 1980 [LFK; as Tellina]; Wagner & Abbott, 1990 [as Tellina]; Krisberg, 1993 [LFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Tremor, 1998 [LFK; as Tellina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Tellina]. Cymatoica orientalis forma hendersoni Rehder, 1939: Rehder, 1939 [DT; as subspe- cies]; Mikkelsen & Bieler, 2000 [MFK, DT; as subspecies]. Elliptotellina americana (Dall, 1900): Boss, 1966 [LFK; as Tellina (Elliptotellina)]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [LFK; as Tellina]. Eurytellina alternata (Say, 1822): Simpson, 1887—1889 [DT; as Tellina tayloriana С. В. Sowerby Il, 1867]; Dall, 1889a [as Tellina], 1903b [as Tellina]; Bartsch, 1937 [as Tellina]; Lermond, 1936 [as Tellina]; Boss, 1968b [UFK, LFK, DT; as Tellina (Eurytellina)]; Magnotte, 1970-1979 [as Tellina]; Moore & Lopez, 1970 [as Tellina]; Turney & Perkins, 1972 [UFK, MFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT; as Tellina]. Eurytellina angulosa (Gmelin, 1791): Dall, 1889a [as Tellina striata “Hanley”, err. pro Spengler, 1798, 1900c [as T. (Eurytellina)]; Johnson, 1934 [аз T. (Arcopagia, Eurytellina)]; Lermond, 1936 [as Tellina; also as T. striata]; M. Smith, 1937, 1945 [as T. (Acropagia (sic), Eurytellina)]; Morris, 1951 [as T. апдийоза (sic)]; Abbott, 1954 [as 7. (Eurytellina)]; Boss, 1968b [LFK; as Tellina (Eurytellina)]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [LFK; as Tellina]. Eurytellina lineata (Turton, 1819): Calkins, 1878 [LFK; as Tellina braziliana, err. pro brasiliana Lamarck, 1819]; Melvill, 1880 [LFK; as Tellina]; Simpson, 1887-1889 [MFK, DT; also as 7. lineata var. albida Hanley [date unknown]]; Dall, 1889a, 1903b [as Tellina]; Hudson et al., 1970 [UFK; as Tellina]; Lermond, 1936 [as Tellina]; Webb, 1951 [as Т. braziliana, err. pro brasiliana]; Magnotte, 1970-1979 [as Tellina]; Kissling, 1977b [UFK; as Tellina]; Krisberg, 1993 [LFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [UFK, LFK; as Tellina]. Eurytellina nitens (C. B. Adams, 1845): Lermond, 1936 [as Tellina georgiana Dall, 1900]; Boss, 1968b [LFK; as Tellina (Eurytellina)]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [LFK; as Tellina]. Eurytellina punicea (Born, 1778): Abbott, 1954 [as Tellina (Eurytellina)]; Warmke & Abbott, 1961 [as Tellina]; Humfrey, 1975 [as Tellina]; Mikkelsen & Bieler, 2000 [as Tellina]. Laciolina laevigata (Linnaeus, 1758): Webb, 1951 [as Tellina]; Boss, 1966 [UFK, LFK; as Tellina (Laciolina)]; Work, 1969 [as Tellina]; Magnotte, 1970-1979 [as Tellina]; Antonius etal., 1978 [LFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK; as Tellina]. Laciolina magna (Spengler, 1798): Melvill, 1880 [LFK; as Tellina (Phylloda) sol (Hanley, 1844)]; Dall, 1889a, 1903b [as Tellina]; Boss, 1966 [MFK, LFK, DT; as Tellina (Laciolina)]; Magnotte, 1970-1979 [as Tellina]; Wagner CRITICAL CATALOG AND BIBLIOGRAPHY 615 & Abbott, 1990 [as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Hutsell et al., 1997 [as Tellina]; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, ОТ; as Tellina]. Leporimetis intastriata (Say, 1827): Simpson, 1887-1889 [LFK; as Lutricola gruneri (Philippi, 1845)]; Dall, 1889a [as Lutricola interstriata (sic)], 1900a [as Metis]; Lermond, 1936 [as Apolymetis intasriata (sic)]; Olsson & Harbison, 1952 [LFK; as Hemimetis (Florimetis)]; Wagner 8 Abbott, 1990 [as Psammotreta]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK]. Macoma brevifrons (Say, 1834): Dall, 1889a, 1903b; Lermond, 1936; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. |Macoma calcarea (Gmelin, 1791): Abbott, 1961 [LFK; in error]. Macoma cerina (C. B. Adams, 1845): Dall, 1889a, 1903b; Lermond, 1936; Pulley, 1952 [MFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LAS IDA: Macoma constricta (Bruguiére, 1792): Lermond, 1936; Webb, 1942, 1951 [LFK]; Mikkelsen & Bieler, 2000 [ЧЕК]. Macoma extenuata Dall, 1900: Pulley, 1952 [MFK]. Macoma limula Dall, 1895: Lermond, 1936 [as М. cimula (sic)]; Pulley, 1952 [MFK]. Macoma mitchelli Dall, 1895: Mikkelsen & Bieler, 2000 [ЧЕК]. Macoma pseudomera Dall & Simpson, 1901: Pulley, 1952 [MFK]. Macoma tageliformis Dall, 1900: Mikkelsen & Bieler, 2000 [ЕЕК]. Macoma tenta (Say, 1834): Dall, 1889a [also as М. t. var. Souleyetiana Récluz, 1852], 1903b; Lermond, 1936 [also as M. t. souleyetiana]; Vittor 8 Associates, 1999а [DT]; Mikkelsen & Bieler, 2000 [UFK, LFK, ОТ]. Масота sp.: Dall, 1896a [ЕЕК]; Vittor & As- sociates, 1998 [LFK, DT], 1999b [ЕЕК]. Merisca aequistriata (Say, 1824): Lermond, 1936 [as Tellina; also as Т. lintea Conrad, 1837]; Bartsch, 1937 [as Т. lintea]; Pulley, 1952 [MFK; as Т. lintea]; Boss, 1966 [UFK, ЕЕК; as Tellina (Merisca)]; Lyons & Quinn, 1995 [as Tellina]; Vittor & Associates, 1998 [LFK, DT; as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, LFK, DT; as Tellina]. Merisca cristallina (Spengler, 1798): Lermond, 1936 [as Tellina crystallina Wood, 1815]; Webb, 1951 [as T. crystallina]. Merisca martinicensis (Orbigny, 1842): Lermond, 1936 [LFK; as Tellina]; Boss, 1966 [LFK, DT; as Tellina (Merisca)]; Lyons 8 Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [LFK, DT; as Tellina]. |Psammotreta intastriata (Say, 1827) — see under Leporimetis. Scissula candeana (Orbigny, 1842): Dall, 1900c [as Tellina (Scissula)]; Johnson, 1934 [as Т. (Angulus, Scissula)]; Lermond, 1936 [as Tellina]; M. Smith, 1937, 1945 [as T. (An- gulus, Scissula)]; Webb, 1942, 1951 [as Tellina]; Olsson & Harbison, 1952 [as 7. (Scissula)]; Pulley, 1952 [MFK; as Tellina]; Abbott, 1954 [LFK; as 7. (Scissula)], 1968 [LFK; as Tellina]; Boss, 1968b [MFK, LFK; as Tellin (Scissula)]; Howard et al., 1970 [LFK; as Tellina]; Lineback, 1977 [LFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Mikkelsen & Bieler, 2000 [МЕК, LFK; as Tellina]. Scissula consobrina (Orbigny, 1842): Boss, 1968b [UFK, MFK, LFK, DT; as Tellina (Scissula)]; Lyons & Quinn, 1995 [as 7. consorbrina (sic)]; Mikkelsen 8 Bieler, 2000 [UFK, МЕК, ЕЕК, DT; as Tellina]. Scissula iris (Say, 1822): Calkins, 1878 [as Те!та]; Dall, 1900c [as 7. (Scissula); also as Tellina (Scissula) exilis Lamarck, 1818, non Meuschen, 1787, nec Link, 1808] for which Boss (1968b) proposed 7. (S.) sandix as a replacement name. Scissula sandix is Caribbean and South American in distribu- tion and Floridian records were listed by Boss, who gave Jamaica as its northernmost record. Boss (1968b: 335) said S. sandix “has often been confused” and “is very closely allied” with $. iris]; Johnson, 1934 [as T. (Angulus, Scissula)]; Lermond, 1936 [as Tellina]; М. Smith, 1937, 1945 [as Т. (An- gulus, Scissula)]; Boss, 1968b [LFK, DT; as Tellina (Scissula)]; Vittor 8 Associates, 1998 [LFK, DT; as Tellina], 1999b [UFK, MFK, LFK; as Tellina]; Mikkelsen & Bieler, 2000 [UFK, МЕК, LFK, DT; as Tellina]; Morton & Knapp, 2004 [UFK, MFK; as Tellina]. Scissula similis (J. Sowerby, 1806): Calkins, 1878 [as Tellina decora Say, 1826]; Melvill, 1880 [LFK; as Tellina]; Simpson, 1887-1889 [MFK, LFK; as 7. decora and Т. decora white var.]; Dall, 1889a, 1903b [as Т. decora]; Lermond, 1936 [as Т. decora]; Olsson 8 Harbison, 1952 [as T. (Scissula)]; Boss, 1968b [UFK, MFK, LFK, DT; as Tellina (Scissula)]; Brooks, 1968a, b [MFK; as Tellina]; Ross, 1969 [МЕК; as Tellina], 1971 [LFK; as Tellina]; Hudson et al., 1970 [UFK; as Tellina]; Magnotte, 1970-1979 [as Tellina]; Turney & Perkins, 1972 [UFK, MFK; as Tellina]; Clampit, 1987 [LFK; as “some small 616 MIKKELSEN & BIELER rose-striped tellin-like shells”]; Haviland, 1994 [LFK; as Tellina]; Lyons & Quinn, 1995 [as Tellina]; Vittor 8 Associates, 1997c [UFK; as Tellina], 1998 [LFK, DT; as Tellina], 1999b [LFK; as Tellina]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Tellina]; Morton & Knapp, 2004 [UFK, MFK; as Tellina]. Strigilla carnaria (Linnaeus, 1758): Melvill, 1880 [LFK]; Dall, 1889a, 1900c [as S. rombergii Mórch, 1853]; 1903b; Lermond, 1936 [also as $. rombergiil; Boss, 1969 [LFK]; Magnotte, 1970-1979 [as $. romgergi (sic)]; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [ЕЕК]. Strigilla gabbi Olsson & McGinty, 1958: Boss, 1969 [LFK]; Abbott, 1974 [LFK; as S. (Strigilla)]; Ode, 1975 [ЕЕК]; Rios, 1994 [LFK; as S. (Strigilla)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [LFK]. Strigilla mirabilis (Philippi, 1841): Calkins, 1878 [DT; as S. flexuosa (Say, 1822), non Montagu, 1803]; Dall, 1889a, 1903b [as S. flexuosa]; Lermond, 1936 [аз $. flexuosa]; Bartsch, 1937 [as S. flexuosa]; M. Smith, 1937 [LFK; as S. flexuosa]; Boss, 1969 [MFK, LFK]; Turney & Perkins, 1972 [UFK]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Strigilla pisiformis (Linnaeus, 1758): Simpson, 1887-1889 [LFK, DT; аз $. pisum (sic)]; Calkins, 1878; Melvill, 1880 [LFK]; Dall, 1889a, 1903b [LFK], 1900c; Johnson, 1934; Lermond, 1936; M. Smith, 1937, 1945; Webb, 1942, 1951; Abbott, 1954, 1968; Warmke & Abbott, 1961; Mikkelsen & Bieler, 2000 [LFK]. Tellidora cristata (Récluz, 1842): Dall, 1889a, 1903b; Boss, 1968b [UFK]; Lermond, 1936; Magnotte, 1970-1979; Turney & Perkins, 1972 [MFK]; Lyons & Quinn, 1995; Vittor 8 Associates, 1998 [ЕЕК]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК]. | Tellina aequistriata (Say, 1824) — see under Merisca. | Tellina agilis (Stimpson, 1857) — see under Angulus. |Tellina alternata (Say, 1822) — see under Eurytellina. |Tellina americana (Dall, 1900) — see under Elliptotellina. | Tellina angulosa (Gmelin, 1791) — see under Eurytellina. |Tellina candeana (Orbigny, 1842) — see un- der Scissula. | Tellina consobrina (Orbigny, 1842) — see un- der Scissula. | Tellina cristallina (Spengler, 1798) — see un- der Merisca. |Tellina fausta (Pulteney, 1799) — see under Arcopagia. |Tellina gouldii (Hanley т С. В. Sowerby |, 1846) — see under Acorylus. | Tellina iris (Say, 1822) — see under Scissula. |Tellina laevigata (Linnaeus, 1758) — see un- der Laciolina. |Tellina lineata (Turton, 1819) — see under Eurytellina. |Tellina listeri (Róding, 1798) — see under Tellinella. |Tellina magna (Spengler, 1798) — see under Laciolina. |Tellina martinicensis (Orbigny, 1842) — see under Merisca. | Tellina mera (Say, 1834) — see under Angulus. |Tellina nitens (С. В. Adams, 1845) — see un- der Eurytellina. |Tellina paramera (Boss, 1964) — see under Angulus. Tellina persica Dall & Simpson, 1901: Mikkelsen & Bieler, 2000 [LFK]. |Tellina probrina (Boss, 1964) — see under Angulus. |Tellina punicea (Born, 1778) — see under Eurytellina. Tellina radiata Linnaeus, 1758: Calkins, 1878 [DT]; Melvill, 1880 [LFK; also 7. г. var. unimaculata Lamarck, 1818]; Simpson, 1887—1889 [DT; as T. radiata var.]; Dall, 1889a, 1903b; Aldrich 8 Snyder, 1936 [ЕЕК]; Lermond, 1936; С. М. Vilas 8 М. К. Vilas, 1945, 1970 [also as Т. г. var. unimaculata; Boss, 1966 [UFK, MFK, LFK, ОТ; as Т. (Те/та)]; Burggraf, 1969 [ЕЕК]; Work, 1969 [UFK, ОТ]; Magnotte, 1970-1979; Moore & López, 1970; Gaertner, 1978 [LFK; as sun- rise clams]; Edwards, 1980 [ЕЕК]; Wagner & Abbott, 1990 [as T. radiata and T. r. unimaculata]; Krisberg, 1993 [ЕЕК]; Haviland, 1994 [LFK]; Lyons 8 Quinn, 1995; Tremor, 1998 [LFK; as radiala (sic)]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, ОТ]. |Tellina sandix (Boss, 1968) — see under Scissula iris. | Tellina similis (J. Sowerby, 1806) — see under Scissula. Tellina squamifera Deshayes, 1855: Dall, 1889a, 1903b; Lermond, 1936; Pulley, 1952 [MFK]; Boss, 1967 [as 7. (Phyllodina)]; Boss, 1966 [UFK, MFK, LFK, DT; as 7. (Phyllodina)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, ОТ]. |Tellina sybaritica (Рай, 1881) — see under Angulus. |Tellina tampaensis (Conrad, 1866) — see un- der Angulus. CRITICAL CATALOG AND BIBLIOGRAPHY 617 |Tellina tenella (А. Е. Мег, 1874) — see un- der Angulus. |Tellina texana (Dall, 1900) — see under Angu- lus. |Tellina versicolor (“Cozzens” DeKay, 1843) — see under Angulus. Tellina ($5. |.) sp.: Dall, 1889a [ЕЕК], 1895а [ЕЕК], 1903b [ЕЕК]; Simpson, 1897 [LFK; as “bright Tellinas”]; Plockelman, 1968d; Ross, 1971 [UFK]; Schomer 8 Drew, 1982; Theroux & Wigley, 1983 [МЕК, LFK]; Wingard et al., 1995 [UFK; as 7. spp.]; Brewster-Wingard et al., 1996, 1997, 2001 [all ЧЕК; as 7. spp.]; Vittor 8 Associates, 1997b [LFK], 1997с [ЧЕК], 1998 [LFK, DT], 1999a [UFK, LFK, DT], b [UFK, MFK, LFK]; Dent, 1998; USGS, 2003 [UFK, MFK; as Tellina sp. or spp.]. Tellinella listeri (Róding, 1798): Melvill, 1880 [LFK; as Tellina interrupta Wood, 1815]; Simpson, 1887-1889 [LFK, ОТ; as 7. interrupta]; Dall, 1889a, 1903b [as Т. interrupta]; Lermond, 1936 [as 7. interrupta]; Boss, 1966 [UFK, MFK, LFK, DT; as Tellina (Tellinella)]; Work, 1969 [LFK; as Tellina]; Magnotte, 1970-1979 [as Tellina]; Voss, 1983 [UFK; as Tellina]; Voss et al., 1983 [UFK; as Tellina]; Wagner & Abbott, 1990 [as Tellina]; Krisberg, 1993 [LFK; as Tellina]; Lyons 8 Quinn, 1995 [as Tellina]; Vittor 8 Associates, 1998 [LFK; as Tellina]; Mikkelsen 8 Bieler, 2000 [UFK, MFK, LFK, DT; as Tellina]. Teredinidae Stevenson, 1970, 1993 [both as shipworm]. Bankia carinata (Gray, 1827): Mikkelsen & Bieler, 2000 [МЕК]. Bankia fimbriatula (Moll & Roch, 1931): Calkins, 1878 [LFK; as Xylotrya fimbiata, err. pro X. fimbriata (Jeffreys, 1860)]; Dall, 1889a [as X. fimbriata]; Lermond, 1936 [as X. fimbriata]. Lyrodus pedicellatus (de Quatrefages, 1849): Pulley, 1952 [LFK; as L. pedicellata]. Nototeredo Кпох! (Bartsch, 1917): Mikkelsen 8 Bieler, 2000 [UFK]. Teredo bartschi Clapp, 1923: Pulley, 1952. Teredo clappi Bartsch, 1923: Bartsch, 1923 [LFK; as Т. (Zopoteredo)]; Johnson, 1934 [LFK; as T. (Zopoteredo)]; Lermond, 1936 [LFK]; Aguayo & Jaume, 19504 [ЕЕК]; Turner, 1966 [LFK; as 7. (Zopoteredo)]; Mikkelsen & Bieler, 2000 [UFK, ЕЕК]. Teredora malleolus (Turton, 1822): Dall, 1889a [as Teredo Thomsoni Tryon, 1863]; Lermond, 1936 [as T. thomsoni]. Thraciidae Vittor & Associates, 1998 [LFK, DT], 1999a [DT]. Asthenothaerus hemphillii Dall, 1886: Dall, 1889a, 1903b; Lermond, 1936 [as A. hemphilli; Rehder, 1943a, b [LFK; as A. balesi Rehder, 1943]; Aguayo & Jaume, 1947a [LFK; as A. (Asthenothaerus) balesi]; Abbott, 1974 [LFK; as A. balesi]; Lyons & Quinn, 1995 [as A. balesi]; Vittor & Associ- ates, 1998 [LFK, DT], 1999a [UFK, LFK, DT], b [MFK, LFK] [all as A. hemphillil; Sweeney & Harasewych, 1999 [LFK; as A. balesil; Mikkelsen & Bieler, 2000 [UFK, LFK, DT; as А. hemphilli and A. balesi. Asthenothaerus sp.: Vittor & Associates, 1999b [LFK]. Bushia elegans (Dall, 1886): Dall, 1889a [as Asthenothaerus (Bushia)]; Vittor & Associ- ates, 1999b [UFK]. Bushia sp.: Vittor & Associates, 1998 [DT]. Thracia distorta (Montagu, 1803): Simpson, 1887—1889 [MFK; as T. rugosa Lamarck, 1818]. Thracia morrisoni Petit, 1964: Dall, 1886 [LFK; misidentified as Thracia corbuloides (sic, err. pro corbuloidea) Blainville, 1827, a recog- nized Mediterranean species], 1889a [LFK; аз Т. corbuloidea], 1903b [LFK; as T. corbuloideal; Johnson 1934 [as Т. corbuloides (sic)]; Lermond, 1936 [as Т. corbuloides (sic)]; Abbott, 1974 [as Т. corbuloides (sic)]; Mikkelsen 8 Bieler, 2000 [LFK; as 7. corbuloides (sic)]. “Thracia phaseolina Lamarck, 1822” [a Euro- pean species; western Atlantic specimens appear to represent a new species (Coan, 1990)]: Рай, 1889a, 1903b; Mikkelsen & Bieler, 2000. Thracia stimpsoni Dall, 1886: Dall, 1889a, 1903b [DT]; Maury, 1920 [DT]; Johnson, 1934 [DT]; Aguayo & Jaume, 1950е [DT]; Boss et al., 1968 [DT]; Vittor & Associates, 1998 [LFK]; Mikkelsen & Bieler, 2000 [LFK, ОТ]. Thyasiridae Thyasira grandis (А. E. Verrill & Smith, in: A. E. Verrill, 1885): Dall, 1889a [as Cryptodon pyriformis Dall, 1886, and misidentified as С. obesus А. Е. Verrill, 1872 (fide Maury, 1920)]. Thyasira trisinuata (Orbigny, 1842): Mikkelsen & Bieler, 2000 [DT]. 618 MIKKELSEN & BIELER Anomalocardia auberiana (Orbigny, 1842): Melvill, 1880 [LFK; misidentified as Anomalocardia impressa (Anton, 1837), syn- Trapezidae Coralliophaga coralliophaga (Gmelin, 1791): Simpson, 1887-1889 [DT; as Cypricardia]; Lermond, 1936; Bales, 1944; Pulley, 1952; Solem, 1955 [LFK, DT]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [MFK, LFK, ОТ]. Ungulinidae Vittor & Associates, 1998 [LFK]. Diplodonta notata Dall & Simpson, 1901: Lermond, 1936 [LFK; as Taras]. Diplodonta nucleiformis Wagner, 1838: Lermond, 1936 [as Taras]. |Diplodonta pilula Dall, 1881 — all records based on Blake sta. 43 (here excluded; see entry for Dall, 1881). Diplodonta punctata (Say, 1822): Dall, 1889a [as Diplodonta subglobosa (C. B. Adams, 1852)]; Lermond, 1936 [as Taras]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, МЕК]; Voss, 1983 [ЧЕК]; Voss et al., 1983 [UFK]; Lyons & Quinn, 1995; Vittor & Asso- ciates, 1999b [UFK, MFK, LFK]; Mikkelsen & Bieler, 2000 [MFK, LFK, DT]. Diplodonta sp.: Vittor & Associates, 1998 [LFK, DT], 1999b [UFK, MFK, LFK]; USGS, 2003 [MFK; as Diplodonta spp.]. Felaniella candeana (Orbigny, 1842): Simpson, 1887-1889 [DT; as Diplodonta]. Phlyctiderma semiaspera (Philippi, 1836): Simpson, 1887-1889 [DT; as Diplodonta]; Dall, 1889a, 1903b [both as Diplodonta]; Lyons & Quinn, 1995 [as Diplodonta]; Vittor 8 Asso- ciates, 1997c [UFK], 1997b [LFK], 1998 [LFK, DT] [all as Diplodonta]; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT; as Diplodonta]. Phlyctiderma soror (C. B. Adams, 1852): Simpson, 1887-1889 [DT; as Diplodonta]; Dall, 1889a [ОТ; as Diplodonta], 1899a [as Diplodonta (Phlyctiderma)]; Lermond, 1936 [as Taras]. Veneridae Dall, 1889a [as Cytherea sp.; also as Venus Lamarckii Gray, 1838, synonym of Antigona lamellaris Schumacher, 1817, a recognized Indo-Pacific species, similar to several Chione s.l. spp. (so cannot be assigned to a recognized Florida Keys species)], 1896a [LFK; as Venus sp.]; Rogers, 1941 [UFK; as Venus clam]; Vittor 8 Associates, 1997с [UFK], 1998 [LFK, DT], 1999a [ЕЕК], b [UFK, МЕК]. опут of А. producta Kuroda & Habe, 1951, a recognized Chinese/Japanese species]; Dall, 1883, 1885 [both LFK; as А. flexuosa (Linnaeus, 1767), a recognized Caribbean/ South American species, probably misidentified A. auberiana], 1889a [as Ve- nus (Anomalocardia) rostrata G. B. Sowerby |, 1853]; Lermond, 1936 [misidentified as Animalocardia (sic) brasiliana, err. pro А. brasiliana (Gmelin, 1791), a recognized spe- cies of the Caribbean, Central and South America, and as À. cuneimeris (Conrad, 1846)]; Lee, 1969 [LFK; as A. brasiliensis (sic)]; Ross, 1969 [МЕК; as A. cuneimeris]; Howard et al., 1970 [LFK; as A. cuneimeris]; Hudson et al., 1970 [UFK; as A. cuneimeris]; Magnotte, 1970-1979 [as А. cuneimeris]; Turney & Perkins, 1972 [UFK, MFK; also as A. cuneimeris]; Lineback, 1977 [LFK; as A. cuneimeris]; Schomer & Drew, 1982 [as А. cuneiveis (sic)]; Petersen, 1989 [MFK]; Lyons & Quinn, 1995; Wingard et al., 1995 [UFK; as A. cuneimeris]; Brewster-Wingard et al., 1996, 1997, 1998 [all as A. sp.], 2001 [all ЧЕК]; Lyons, 1998 [UFK, МЕК]; Vittor & Associates, 1998 [LFK]; Brewster-Wingard 8 Ishman, 1999 [UFK; as A. sp.]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]; USGS, 2003 [UFK, МЕК]. Callista eucymata (Dall, 1890): Lermond, 1936 [as Pitar encymata (sic)]; Palmer, 1947 [UFK, MFK, LFK; as Costacallista]; Pulley, 1952 [UFK; as Costacallista]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. |Chione cancellata (Linnaeus, 1767) - see С. elevata. |Chione clenchi Pulley, 1952 — see under Lirophora. Chione elevata (Say, 1822): Melvill, 1880 [LFK; as С. cancellata (Linnaeus, 1767)]; Dall, 1886 [Gordon Key; as C. cancellata], 1889a [as Venus cancellata; also LFK, as V. Beaui Récluz, 1852; also DT, misidentified as V. granulata Gmelin, 1791, a recognized Car- ibbean species of Protothaca], 1896a [LFK, DT; as V. (C.) cancellata], 1902b [mis- identified as C. (C.) subrostrata (Lamarck, 1818), a recognized Brazilian species], 1903b [as V. cancellata; also DT, as V. granulata]; Simpson, 1887-1889 [DT, as V. реаи!; also DT, as V. granulata]; Nutting, 1895 [LFK, DT; as C. cigenda (Dillwyn, 1817)]; Johnson, 1934 [as C. (Timoclea) granulata]; Lermond, 1936 [as V. (C.) CRITICAL CATALOG AND BIBLIOGRAPHY 619 cancellatus, V. (C.) subrostrata, and V. (C.) granulatus]; M. Smith, 1937, 1945 [аз С. (Timoclea) granulata]; Bippus, 1950 [UFK; as С. cancellata]; lversen & Roessler, 1969 [UFK; as С. cancellata]; Jindrich, 1969 [LFK; as С. cancellata]; Ross, 1969 [MFK; as С. cancellata]; Howard et al., 1970 [LFK; as C. cancellata]; Hudson et al., 1970 [UFK; as C. cancellata]; Magnotte, 1970-1979 [as C. cancellata]; Turney 8 Perkins, 1972 [UFK, MFK; as C. cancellata]; Godcharles € Jaap, 1973 [UFK; as С. cancellata]; Zischke, 1973, 1977a, b, с [MFK; as С. cancellata]; Lineback, 1977 [LFK; as С. cancellata]; Schomer & Drew, 1982 [as as С. cancellata]; Voss, 1983 [UFK; as С. cancellata]; Voss et al., 1983 [UFK; as C. cancellata]; Petersen, 1989 [МЕК; as С. cancellata]; Krisberg, 1993 [LFK; as С. cancellata]; Lyons & Quinn, 1995 [as С. cancellata]; Wingard et al., 1995 [UFK; аз С. cancellata]; Brewster-Wingard et al. 1996, 1997, 1998 [all ЧЕК; as С. cancellata]; Vittor & Associates, 1997с [UFK; as С. cancellata], 1998 [LFK, ОТ; as С. cancellata], 1999b [UFK, MFK, LFK; as C. cancellata]; Lyons, 1998 [UFK, MFK; as C. cancellata]; Brewster-Wingard & Ishman, 1999, 2001 [UFK; as С. cancellata]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT, as C. cancellata; also as Protothaca granulata]; Roopnarine & Vermeij, 2000 [UFK]; USGS, 2003 [UFK, MFK; as C. cancellata]; Morton & Knapp, 2004 [UFK, MFK]. |Chione grus (Holmes, 1858) — see under Timoclea. |Chione intapurpurea (Conrad, 1849) — see under Puberella. |Chione latilirata (Conrad, 1841) — see under Lirophora. Chione mazyckii Dall, 1902: Lermond, 1936 [as Venus (Chione)]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. |Chione paphia (Linnaeus, 1767) — see under Lirophora. |Chione pubera (Bory Saint-Vincent, 1827) — see under Puberella. |Chione pygmaea (Lamarck, 1818) — see un- der Timoclea. Chione (s. |.) sp.: Vittor & Associates, 1998 [LFK, DT], 1999b [ЕЕК]. Circomphalus strigillinus (Dall, 1902): Бай, 1902, 1903b [LFK; as Cytherea (Ventricola) strigillina]; Johnson, 1934 [as Antigona (Circomphalus, Ventricola)]; Lermond, 1936 [аз Antigona strigillina]; Aguayo & Jaume, 1949е; P. L. McGinty & Т. Е. McGinty, 1957 [MFK, LFK; as Antigona strigillina]; Boss et al., 1968 [LFK; as Cytherea (Ventricola) strigillina]; Mikkelsen & Bieler, 2000 [MFK, LEK, DT]: Cyclinella tenuis (Récluz, 1852): Dall, 1889a, 1903b [as Lucinopsis]; Lermond, 1936; Boss & Wass, 1970 [MFK, LFK, DT]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, МЕК ВЕК, Dit: Dosinia discus (Reeve, 1850): Lermond, 1936; Bartsch, 1937; Webb, 1937, 1942, 1951; Rogers, 1941 [UFK; as discus clam]; Magnotte, 1970-1979; Vittor 8 Associates, 1998 [DT], 1999а (LEK; DIT], 5 [MEK]; Mikkelsen & Bieler, 2000 [МЕК, ЕЕК, DT]. Dosinia elegans (Conrad, 1846): Dall, 1889a, 1903b, 1902a, b [DT; as Dosinia (Dosinidia) and misidentified as D. (Dosinidia) concentrica (Born, 1780), a recognized Car- ibbean to South American species “not found in Florida” (Abbott, 1974: 533)]; Conrad, 1866 [as D. floridana Conrad, 1866]; Johnson 1934 [as D. concentrica]; Lermond, 1936 [also as Dosinia concentrica]; Bartsch, 1937; M. Smith, 1937, 1945 [as D. concentrica]; Clench, 1942a [as О. floridana]; Magnotte, 1970-1979; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Gemma gemma (Totten, 1834): Lermond, 1936 [as Gemma purpurea (H. C. Lea, 1842)]. Globivenus rigida (Dillwyn, 1817): Dall, 1889a [as Venus rugosa Gmelin, 1791, non Linnaeus, 1758], 1902b [as Cytherea (Ventricola)]; Johnson, 1934 [as Antigona (Circomphalus, Ventricola)]; Lermond, 1936 [as Antigona]; Abbott, 1974; Humfrey, 1975 [as Antigona (Ventricolaria)|; Odé, 1976a; Warmke & Abbott, 1961 [as Antigona]; Diaz Merlano & Puyana Hegedus, 1994 [as Ventricolaria]; Rios, 1994 [as Ventricolaria (Ventricolaria)]; Lyons & Quinn, 1995 [as Ventricolaria]; Mikkelsen & Bieler, 2000 [DT]. Globivenus rugatina (Heilprin, 1886): Dall, 1889a, 1903a [as Cytherea (Cytherea, sec- tion Ventricola)], 1903b [as Venus rugosa var. rugatina]; Goldberg, 1978c [LFK]; Mikkelsen & Bieler, 2000 [LFK, DT]. Globivenus sp.: Dall, 1889a [misidentified as Venus pilula Reeve, 1863, a recognized Okinawan species of Globivenus, similar to С. rigida/rugatina]. Gouldia cerina (C. B. Adams, 1845): Dall, 1889a, 1903b [as Circe (Gouldia)]; Lermond, 1936 [also as Circe]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK, DT], 1999a [LFK], b [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. 620 MIKKELSEN & BIELER Lirophora clenchi (Pulley, 1952). Lirophora latilirata (Conrad, 1841): Lermond, 1936 [as Venus (Chione) latiliratus]; Webb, 1942, 1951 [as Chione]; Siekman, 1965 [as Chione]; Lyons & Quinn, 1995 [as Chione]; Mikkelsen & Bieler, 2000 [LFK, ОТ]. Lirophora paphia (Linnaeus, 1767): Simpson, 1887-1889 [as Chione]; Lermond, 1936 [as Venus (Chione)]; Webb, 1942, 1951[as Chione]; Abbott, 1954 [LFK; as Chione (Lirophora)]; Warmke 8 Abbott, 1961 [LFK; as Chione]; Barrett & Patterson, 1967 [LFK; as Chione]; Magnotte, 1970-1979 [as Chione]; Humfrey, 1975 [LFK; as Chione]; Voss, 1983 [UFK; as Chione]; Voss et al., 1983 [UFK; as Chione]; Sutty, 1990 [LFK; as Chione]; Mikkelsen 8 Bieler, 2000 [LFK, DT; as Chione]. Macrocallista maculata (Linnaeus, 1758): Dall, 1889a, 1903b [as Cytherea (Callista)], 1902b [as M. (Chionella)]; Lermond, 1936; Magnotte, 1970-1979; Lyons & Quinn, 1995; Vittor & Associates, 1998 [DT], 1999a [DT]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT]. Macrocallista nimbosa (Lightfoot, 1786): Melvill, 1880 [LFK; as Callista (Dione) gigantea “(Chemnitz)” (Gmelin, 1790)]; Dall, 1889a [as Cytherea (Callista) gigantea], 1903a; Lermond, 1936 [as M. (Callista) gigantea]; Abbott, 1961 [LFK], 1970 [LFK]; Vittor & Associates, 1999a [DT]; Mikkelsen & Bieler, 2000 [UFK, LFK, DT]. Mercenaria campechiensis (Gmelin, 1791): Simpson, 1887-1889 [as Venus топот Conrad, 1837]; Dall, 1889a [as V. mercenaria var. Mortoni]; Lermond, 1936 [as Venus]; Magnotte, 1970-1979; Wagner & Abbott, 1990; Hutsell et al., 1997; Mikkelsen & Bieler, 2000 [LFK]. Mercenaria mercenaria (Linnaeus, 1758): Dall, 1889a [as Venus], 1903b [as Venus and as V. т. var. notata (Say, 1822)], 1902b [as Venus]; Lermond, 1936 [as Venus]; Mikkelsen & Bieler, 2000 [MFK; as M. m. forma notata]. Mercenaria sp.: USGS, 2003 [MFK; as Mercenaria spp.]. Parastarte triquetra (Conrad, 1846): Dall, 1902b, 1903a; M. Smith, 1937, 1945; Howard et al., 1970 [LFK]; Turney & Perkins, 1972 [UFK, MFK; also as Parastarte sp.]; Lineback, 1977 [LFK]; Wingard et al., 1995 [UFK]; Brewster-Wingard et al., 1996, 2001 [UFK]; Brewster-Wingard & Ishman, 1999 [UFK]; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, DT]; USGS, 2003 [ЧЕК]. Periglypta listeri (Gray, 1838): Simpson, 1887- 1889 [LFK; as Venus]; Dall, 1889a, [misidentified as Venus crispata Deshayes, 1853, an Indo-Pacific form of unresolved sta- tus in Periglypta], 1902b [as Cytherea (Cytherea)l; Palmer, 1927-1929 [as Antigona (Dosina)]; Lermond, 1936 [as Antigonal; M. Smith, 1937 [LFK; as Antigona]; Pulley, 1952 [LFK; as Antigona]; Abbott, 1961, 1970 [both as Antigona]; Brooks, 1968а [MFK; as Antigona]; Ross, 1969 [MFK; as Antigona]; Magnotte, 1970- 1979 [as Antigona]; Woods, 1970 [LFK; as Antigona]; Godcharles & Jaap, 1973 [UFK; as Antigona]; Zischke, 1973, 1977a, с [MFK; as Antigona]; Theroux & Wigley, 1983 [MFK]; Tremor, 1998 [LFK]; Edwards, 1980 [LFK]; Voss, 1983 [UFK]; Voss et al., 1983 [UFK; also as Periglyphus (sic)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, ЕЕК, DT]; Bieler et al., 2004 [UFK, МЕК, LFK, DITE Pitar albidus (Gmelin, 1791): Dall, 1889a [as Cytherea albida], 1896a [DT; as С. albida]; Lermond, 1936 [as Cytherea]. Pitar circinatus (Born, 1778): Simpson, 1887- 1889 [DT; as Cytherea circinata]. Pitar cordatus (Schwengel, 1951): Schwengel, 1951 [DT; as Pitaria cordata]; Rehder & Abbott, 1951 [DT; as Р (Pitarenus) cordata], 1954 [as P (Pitarenus) cordata], 1974 [as Р. (Pitarenus)]; Morris, 1973 [LFK; as Pitar cordata]; Abbott & Morris, 1995 [LFK]; Odé, 1976b; Rios, 1994 [as P. (Pitarenus)]; Mikkelsen & Bieler, 2000 [LFK, DT]. Pitar dione (Linnaeus, 1758): Calkins, 1878 [DT; as Cytherea dione]; Simpson, 1887-1889 [DT; as Cytherea]; Dall, 1889a, 1903b [as C. (Dione) Dione]; Lermond, 1936 [аз С. (D.)]. Pitar fulminatus (Menke, 1828): Simpson, 1887-1889 [DT; as Cytherea hebraea Lamarck, 1818]; Dall, 1886 [Gordon Key; as С. (Dione) hebraea], 1889a [as С. hebraeal, 1903b [as C. hebraea]; Lermond, 1936 [also as С. hebraea]; Webb, 1937, 1939 [both as С. hebraea], 1942, 1951 [both as Р fulminata and C. hebraea]; Howard et al., 1970 [LFK; as P cf. fulminata]; Magnotte, 1970-1979; Turney & Perkins, 1972 [UFK, MFK; as Р fulminata]; Lineback, 1977 [LFK; as P cf. fulminata]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [DT], 1999b [UFK, ЕЕК}; Mikkelsen & Bieler, 2000 [LFK, DT]. Pitar simpsoni (Dall, 1895): Dall, 1889a [as Cytherea Simpsoni]; Lermond, 1936 [as Cytherea]; Lyons & Quinn, 1995; Wingard et al., 1995 [UFK; as P sp.]; Vittor & Associ- ates, 1998 [LFK, DT], 1999a [UFK, LFK, DT], b [UFK, MFK, LFK]; Mikkelsen 8 Bieler, 2000 CRITICAL CATALOG AND BIBLIOGRAPHY 621 [UFK, MFK, LFK, ОТ]; Brewster-Wingard et al., 2001 [ЧЕК]; USGS, 2003 [UFK, МЕК]; Morton & Knapp, 2004 [UFK, МЕК]. Pitar sp.: Theroux & Wigley, 1983 [MFK, ЕЕК]; Vittor 8 Associates, 1997c [ЧЕК], 1998 [LFK, DT], 1999a [ЕЕК], b [ЧЕК]. Puberella intapurpurea (Conrad, 1849): Бай, 1902b [as Chione (Chione)]; M. Smith, 1937, 1945 [as Chione]; Godcharles € Jaap, 1973 [UFK; as Chione]; Antonius et al., 1978 [LFK; as Chione]; Lyons & Quinn, 1995 [as Chione]; Mikkelsen 8 Bieler, 2000 [UFK, МЕК, LFK, ОТ]. Puberella pubera (Вогу Saint-Vincent, 1827): Johnson, 1934 [as Valenciennes, 1827]; Lermond, 1936 [as Venus (Chione)]; M. Smith, 1937, 1945 [as Chione]; Abbott, 1974 [as Chione (Chione)]; Lyons & Quinn, 1995 [as Chione puber]; Mikkelsen & Bieler, 2000. Timoclea grus (Holmes, 1858): Henderson, 1913 [UFK; as Chione]; Lermond, 1936 [as Venus (Chione)]; Abbott, 1954 [LFK; as Chione (Tellina)]; Magnotte, 1970-1979 [as Chione]; Andrews, 1971 [LFK; as Chione], 1977, 1981a, b, 1992, 1994 [LFK; as Chione (Timoclea)]; Emerson 8 Jacobson, 1976 [LFK; as Chione]; Vittor 8 Associates, 1998 [LFK, ОТ; as Chione], Vittor & Associates, 1999a [DT; as Chione], b [LFK; as Chione]; Mikkelsen & Bieler, 2000 [UFK, МЕК, ЕЕК, ОТ]. Timoclea рудтаеа (Lamarck, 1818): Simpson, 1887-1889 [DT; as Venus, also as У. pygmaea var. inaequivalvia Orbigny, 1846]; Dall, 1889a, 1903b [as Venus], 1902b [as Chione (Timoclea)]; Johnson, 1934 [as Chione (Timoclea)]; Lermond, 1936 [as Ve- nus (Chione) pygmaeus]; M. Smith, 1937, 1945 [as Chione (Timoclea)]; Plockelman, 1968a, b [both as Chione]; Turney & Perkins, 1972 [UFK, MFK; as Chione]; Lyons 8 Quinn, 1995 [as Chione]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT]. Tivela abaconis Dall, 1902. Tivela floridana Rehder, 1939: Mikkelsen & Bieler, 2000 [UFK]. Tivela mactroides Born, 1778: Dall, 1889a [as Cytherea (T.)]; Johnson, 1934; Lermond, 1936 Ча TC: 475) *mactroides and Г. mactroides]; М. Smith, 1937, 1940, 1945; Aguayo & Jaume, 1948c; Barrett & Patterson, 1967. Tivela trigonella (Lamarck, 1818): Simpson, 1887—1889 [DT; as С. (Trigona) incerta “Römer” err. pro С. В. Sowerby Il, 1851]. Transennella conradina (Dall, 1884): Simpson, 1887-1889 [МЕК; as Cytherea]; Dall, 1889a, 1903b [LFK; as Cytherea (T.) Conradina]; 1902a [LFK; as Meretrix (T.)]; Lermond, 1936 [аз С. (Т.) conradiana (sic) and Transenella (sic) conradina]; M. Smith, 1937, 1945; Webb, 1937, 1939, 1942, 1951 [all as Transenella (sic)]; Vittor & Associates, 1997c [UFK]; Johnson, 1934 [as Transenella (sic)]; Lyons & Quinn, 1995; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK, DT; as Transenella (sic)]. Transennella cubaniana (Orbigny, 1842): Dall, 1881 [LFK; as Gouldia], 1889a, 1903b [as Cytherea (T.)]; Johnson, 1934 [as Transenella (sic)]; Lermond, 1936 [аз С. (T.) cubaniana and Transenella (sic) cubaniana]; Hudson et al., 1970 [UFK]; Abbott, 1974; Emerson & Jacobson, 1976; Odé, 1976b; Warmke 8 Abbott, 1961; Díaz Merlano & Puyana Hegedus, 1994 [as Transenella (sic)]; Lyons & Quinn, 1995; Mikkelsen 8 Bieler, 2000 [MFK; as Transenella (sic)]. Transennella culebrana (Dall & Simpson, 1901): Aguayo & Jaume, 1948b [ЕЕК]; Mikkelsen & Bieler, 2000 [as Transenella (sic)]. Transennella stimpsoni Dall, 1902: Dall, 1902b [LFK]; Henderson, 1913 [UFK; as Transenella (sic)]; Johnson, 1934 [LFK; as Transenella (sic)]; Lermond, 1936 [as Transenella (sic)]; Hudson et al., 1970 [UFK]; Lyons & Quinn, 1995; Vittor & Associates, 1998 [LFK]; Mikkelsen & Bieler, 2000 [UFK, MFK, LFK; as Transenella (sic)]. Transennella sp.: Turney & Perkins, 1972 [UFK, MFK; as Transennela (sic)]; Wingard et al., 1995 [UFK; as Transenella (sic) spp.]; Brewster-Wingard et al., 1996, 1997 [both as Transenella (sic) spp.], 1998 [as Т. spp.], 2001 [all ЧЕК]; Brewster-Wingard & Ishman, 1999 [UFK; as 7. spp. and Т. sp.]; USGS, 2003 [UFK, MFK; as Т. spp.]. |Ventricolaria spp. — see Globivenus spp. Verticordiidae Euciroa elegantissima (Dall, 1881): Bartsch, 1937. Haliris fischeriana (Dall, 1881): Dall, 1889a, 1903b [as Verticordia (H.) Fischeriana]; Mikkelsen & Bieler, 2000 [DT]. Spinosipella acuticostata (Philippi, 1844): Dall, 1881 [LFK], 1889a, 1903b; Mikkelsen & Bieler, 2000 [LFK] [all as Verticordia]. Trigonulina ornata Orbigny, 1842: Dall, 1889a, 1903b [as Verticordia (Trigonulina)]; Lermond, 1936 [as Verticordia]; Mikkelsen & Bieler, 2000 [UFK, DT]. 622 MIKKELSEN & BIELER |Verticordia acuticostata Philippi, 1844 — see under Spinosipella. |Verticordia elegantissima (Dall, 1881) — see under Euciroa. |Verticordia fischeriana (Dall, 1881) — see un- der Haliris. |Verticordia ornata (Orbigny, 1842) — see un- der Trigonulina. Vesicomyidae |Vesicomya venusta (Dall, 1886) — only Florida Keys record (Dall, 1889a) probably based on Florida Straits, 801 fms (Abbott, 1974); here excluded as beyond depth limit. Vesicomya vesica (Dall, 1886): Dall, 1889a [as Cytherea (Veneriglossa)]. Yoldiidae Yoldia liorhina Dall, 1881: Dall, 1889a [as Leda (Yoldia)]. Species of Uncertain Taxonomic Status Callucina bermudensis (Dall, 1901): Dall, 1889a [UFK; as Lucina (Lucina) lenticula Reeve, 1850, non Gould, 1850]. Dall (1889a) tabulated L. lenticula as a deep-water spe- cies from the Florida Keys, with Turtle Har- bor (a problematic locality, see above) as its northern limit. However, Dall (1901: 810) stated in the original description of C. bermudensis that “К is not the species cata- logued by me in 1889 as L. lenticula”, with- out any indication of what the latter might be. Dall's (1889a) column listing might be suspected as derived from his Blake report material (Dall, 1886), all of which is deep water, but none from the Florida Keys. In his revision of western Atlantic Lucinidae, Britton (1970) did not include Dall (1889a) in his synonymy of С. bermudensis; neither did Britton list Turtle Harbor as the locality for any L. lenticula examined, stating that C. bermudensis occurs only in Bermuda. Dall (1886: 265) said that his Blake material (again, none from the Florida Keys) was “too poor and insufficient for a satisfactory deter- mination”; Dall (1901) reidentified part of this material as Codakia cubana Dall, 1901, a species otherwise known from the Caribbean and Gulf of Mexico, but not from the Florida Keys [Dall's (1886) “L. lenticula” material also included specimens of Myrtea pristiphora Dall 8 Simpson, 1901 (Britton, 1970)]. Dall's (1889a) “Turtle Harbor” record must then de- rive from another source, possibly the USNM collections; Britton (1970) saw Turtle Harbor material in the USNM of Cavilinga blanda, Codakia orbicularis, Ctena orbiculata, C. pectinella, Divalinga quadrisulcata, Lucinisca nassula, Pleurolucina leucocyma, and P. sombrerensis, mainly from the Eolis expedi- tions (see Bieler & Mikkelsen, 2003). Cyclocardia borealis (Conrad, 1831): Theroux 8 Wigley, 1983 [UFK]. This species” Florida Keys record is based on an unverified speci- men lot at Woods Hole Oceanographic In- stitution; itis distributed from eastern Canada to North Carolina, and is thus probably misidentified. Lepton bowmani (Holmes, 1860): Simpson, 1887-1889 [DT]. Lepton bowmani was origi- nally described from the Pleistocene of South Carolina. To our knowledge, it does not ap- pear in any current work on galeommato- idean bivalves, but it cannot be attributed to another species due to rampant taxonomic uncertainties in this group. Pitar morrhuanus (“Linsley” Dall, 1902): Simpson, 1887-1889 [MFK; as Cytherea convexa “Say,” err. pro Conrad, 1830]. Pitar morrhuanus is typically a cold-water species, ranging from eastern Canada to Cape Hatteras. Sometimes called the “false qua- hog”, it is superficially similar to a cherry- stone-sized Mercenaria spp., but it's true identity in the Florida Keys (from Long Key, a problematic locality, see above) cannot be determined with any degree of certainty. Both Р fulminatus and Agriopoma texasiana Dall, 1892, have been called similar to P. morrhuanus, but are distributed farther south. ACKNOWLEDGMENTS We are indebted to Don Cameron (Univer- sity of Michigan) for linguistic advice concern- ing the gender of certain Greek-derived names, to Jim Clupper (Marathon Public Li- brary) for interpreting old Florida Keys place names, and to Richard E. Petit (Myrtle Beach, South Carolina) for tracking down enigmatic original descriptions. Eugene Coan (Palo Alto, California) suggested numerous improve- ments during the editorial process. This work was supported in part by a grant from the Comer Research and Education Foundation and by NSF award DEB/PEET-9978119. CRITICAL CATALOG AND BIBLIOGRAPHY 623 LITERATURE CITED BIELER, R. & P. M. MIKKELSEN, 2003, The cruises of the Eolis — John В. Henderson's mollusk collections off the Florida Keys, 1910- 1916. American Malacological Bulletin, 17(1/ 2) “2002”: 125-140. [Supplemental tables (day- by-day cruise summaries; revised SI tag num- bers) at http://fm1.fieldmuseum.org/aa/Files/ bieler/Eolis_stations.html.] BIELER, R. & P. М. MIKKELSEN, 2004, Marine bivalves of the Florida Keys: a qualitative fau- nal analysis based on original collections, mu- seum holdings and literature data. In: R. BIELER & P.M. MIKKELSEN, eds., Bivalves studies in the Florida Keys, Proceedings of the International Marine Bivalve Workshop, Long Key, Florida, July 2002. Malacologia, 46(2): 503-544. CARLTON, J. Т., 1996, Marine bioinvasions: the alteration of marine ecosystems by non-indig- enous species. Oceanography, 9(1): 36-43. CHESLER, J., 1994, Not just bilge water. Ameri- can Conchologist, 22(2): 13. CLENCH, W. J. & L. С. SMITH, 1944, The family Cardiidae in the western Atlantic. Johnsonia, 13, 32 pp. CLUPPER, J., 2003, Key names: a gazetteer of the islands ofthe Florida Keys. Monroe County Public Library, http://keys.fiu.edu/gazetteer/ key_search.htm; last accessed 28 October 2003. СОАМ, Е. V., 1990, The Recent eastern Pacific species of the bivalve family Thraciidae. The Veliger, 33(1): 20-55. JOHNSON, R. 1., 1989, Molluscan taxa of Addison Етегу Verrill and Katharine Jeannette Bush, including those introduced by Sanderson Smith and Alpheus Hyatt Verrill. Occasional Papers on Mollusks, Harvard University, 5(67): 1-143. PEIRCE, B. & C. P. PATTERSON, 1880, List of dredging stations occupied by the United States Coast Survey Steamers “Corwin”, “Bibb”, “Hassler”, and “Blake” from 1867 to 1879. Bul- letin of the Museum of Comparative Zoölogy, 6(1): 1-15. PETERSEN, D. W., 1988, Taphonomy and com- munity analysis of a restricted, subtropical la- goon: Long Key Lake, Long Key, U.S.A. MS Thesis, University of Cincinnati, Cincinnati, Ohio. 156 pp. POURTALES, L. F., 1871, Report of Assistant L. F. Pourtales on dredgings made in the sea near the Florida reefs. Report ofthe Superintendent of the United States Coast Survey, Showing the Progress of the Survey During the Year 1868, Appendix 12; House of Representatives, Fortieth Congress, Third Session, H. Ex. Doc. 71: 168-170. SCHUCHERT, C., W. H. DALL, T. W. STANTON & К. $. BASSLER, 1905, Catalogue of the type and figured specimens of fossils, minerals, rocks and ores in the Department of Geology, United States National Museum. Part I. Fossil Invertebrates. Section |. Catalogue of the type specimens of fossil invertebrates in the depart- ment of geology, United States National Mu- seum. Bulletin of the United States National Museum, 53(1): 1-704. SMITH, S., 1889 [*1888”], List of dredging sta- tions occupied by the U. S. Coast Survey Steam- ers Corwin, Bibb, Hassler, and Blake, from 1867 to 1880. Pp. 85-100, in: Lists of dredging sta- tions in North American waters from 1867 to 1887 [extracted from the Annual Report of the Commissioner of Fish and Fisheries for 1886, pp. 957-972]. United States Government Print- ing Office, Washington, DC. 145 pp., charts. [date as per Johnson, 1989: 96]. TOWNSEND, C. H., 1901, Dredging and other records of the U.S. Fish Commission Steamer Albatross, with bibliography relative to the work of the vessel. Report of the Commissioner for the year ending June 30, 1900, United States Commission of Fish and Fisheries, Part XXVI, Appendix: 387-562, pls. 1-7. VOSS, С. M., 1980, Seashore Life of Florida and the Caribbean: a guide to the common marine invertebrates of the Atlantic from Bermuda to the West Indies and of the Gulf of Mexico. Е. A. Seamann Publishing, Miami, Florida. 168 pp. + [12] pls. WALLER, T. R., 1969, The evolution of the Argopecten gibbus stock (Mollusca: Bivalvia), with emphasis on the Tertiary and Quaternary species of eastern North America. Journal of Paleontology, 43(5, Suppl.): v + 125 pp., 3 fold- outs. WALLER, T. R., 1991, Evolutionary relationships of commercial scallops (Mollusca: Bivalvia: Pectinidae). Pp. 1-73, in: S. SHUMWAY, ed., Scallops: biology, ecology and aquaculture. Elsevier, Amsterdam, etc. WAREN, A., 1978, The taxonomy of some north Atlantic species referred to Ledella and Yoldiella (Bivalvia). Sarsia, 63: 213-219. Revised ms. accepted 23 January 2004 HD "EN = E ME 7 MALACOLOGIA, 2004, 46(1-2): 625-677 INDEX Таха in bold are new; pages т italic indicate figures of taxa. abaconis, Tivela 525, 621 abaliena, Cribrarula cribraria 140 Cribrarula cf. cribraria 138, 154 Abra 272, 581, 585, 611 aequalis 520, 567, 569, 571, 584-585, 611 kanamarui 158 lioica 520, 533, 549, 558, 560, 567, A 583, 585, 611 longicallis 611 longicallis americana 520, 539, 611 longicallus 559, 611 profundorum 271-273 Abrina 157-158, 162, 164, 165, 166-167 cunelpyga 157 declivis 157, 162, 166 hainanensis 157, 164, 167 inanis 162, 166 kinoshitai 157, 162, 166-168 lunella 157, 162, 165, 166-167 magna 157, 164, 165, 167 sachalinica 157 scarlatoi 157-158, 159, 161-162, 162-163, 166-167 shiashkotanika 157 рода! 162, 166 tatarica 157 weberi 162, 166 abrolhensis, Cribrarula cribraria 138, 140, 154 acanthodes, Aequipecten 550, 569, 576, 581, 586, 606, 608 Pecten 568, 608 Acanthopleura granulata 563 Acar 355, 372, 378, 591 domingensis 328, 510, 533, 574, 589 reticulata 363 accrescens, Aegista 109 Aegista (Aegista) 89, 94, 111 Acetabularia 497 achatidea, Schilderia 132, 135, 137-138, 152 acicularis, Erosaria 133, 149 Acmaeidae 169, 179-180 Acmaeoidea 180 Acorylus 616 gouldii 521,613 acrilla, Lippistes 547 Acropsi adamsi 575 Acropsis 604 625 adamsi 551 Acrosterigma 593 magnum 511, 591 aculeata, Crepidula 200 aculeatus, Bostrycapulus 198-200 Acusta 81, 84, 96, 103-104, 106-107, 109 ravida 89, 98, 99, 111 acuta, Leda 558 Leda (Leda) 559 Nuculana 517, 572, 583-585, 605 acuticostata, Spinosipella 525, 621 Verticordia 558, 560, 573, 622 Adamantia florida 133, 147 adamsi, Acropsi 575 Acropsis 551 Arca 567 Arca (Acar) 579-580 Arca (Byssoarca) 559, 604 Arcopsis 327-331, 332-335, 336, 516, 530-535, 551, 554-555, 562, 565, 569, 571-572, 574, 583-584, 588-589, 604 adamsonii, Pseudocypraea 133, 147 adansoni, Lasaea 513, 572, 596 Lucina 277, 288, 290-291, 291-293 admirabilis, Leporicypraea 136 Leporicypraea mappa 135-136, 149 admsi, Arca 567, 604 Adusta 127, 131, 140, 142 adusta, Erronea 141 Erronea (Adusta) adusta 155 aegeenis, Nucula 584-585, 605 aegeénsis, Nucula 559-560, 605 aegeensis, Ennucula 517, 605 Nucula 567-568, 572 aegensis, Nucula 584, 605 Aegista 81, 106-108, 114 accrescens 109 (Aegista) 92 (Aegista) accrescens 89, 94, 111 (Plectotropis) 92 (Plectotropis) gerlachi 89, 95, 111 Aegistinae 107, 108-109 Aegistohadra 79, 81, 85, 89, 106-108, 112, 118 delavayana 87, 92, 112-113, 114-117, 124 seraphinica 112 aegrota, Venus 441 Aeguipecten 565, 606 aenigma, Nesiocypraea 132, 143 aequalis, Abra 520, 567, 569, 571, 584-585, 611 Neosimnia 133 626 Simnia 147 Aequipecten 565, 606, 608 acanthodes 550, 569, 576, 581, 586, 606, 608 acanthodes exasperatus 608 exasperatus 606 gibbus 570 gibbus nucleus 576 ОУ O17, oF 1,577,006 heliacus 517, 606 irradians 570 lineolaris 517, 533, 549, 552, 565, 570-571, 576-577, 586, 606 muscosus 551, 565, 570, 577, 584, 586, 606 phrygium 550 phrygius 549, 606, 608 (Plagioctenium) gibbus nucleus 549, 607 aequistriata, Merisca 522, 615 Tellina 568, 570, 572, 584, 616 Tellina (Merisca) 553 aequivalvis, Aloidis 576 Juliacorbula 512, 595 afra, Arcopsis 327, 332, 336 agglutinans, Aspergillum 39 agilis, Angulus 521, 613 Tellina 572, 616 Agriopoma texasiana 622 Agrolimax columbianus 215 reticulatus 215 ala-perdicis, Avicula 579, 611 alata, Isognomon 576, 597 Isognomon (Pedalion) 580, 597 Melina 552 Pedalion 580, 584, 597, 611 Pedalion (Perna) 568 Schasichella 224 alatus, Isognomen 557, 566 Isognomon 316, 501, 514, 535, 556, 562, 564—565, 570, 572, 578, 586, 588-589, 597, 611 alba, Anodontia 292, 514, 552, 570-571, 583, 599 Anodontia (Anodontia) 556 albicoma, Divarilima 514, 562, 572, 574, 598 Lima 550, 559 Limaria 577 albida, Cytherea 559, 567, 620 Poromya 519, 539, 610 Poromya (Cetomya) 560 albidus, Pitar 524, 620 Albinaria caerulea 23 albuginosa, Erosaria 133-134, 148 Alcadia boeckeleri 217, 220, 223 INDEX hojarasca 217, 220, 223 hollandi 224 jamaicensis 224 major 224 rotunda 224 alderi, Polinices 296 Alectryonella plicatula 316-317 alexhuberti, Austrasiatica 143 alfredensis, Cypraeovula 137-138, 153 algoensis, Cypraeovula 137-139, 153 Ostrea 316-317 aliwalensis, Leporicypraea mappa 134-135, 149 Aloididae 563, 594 Aloidis 595 aequivalvis 576 operculata 576, 595 altaicum, Deroceras (Deroceras) 125 altanai, Xylopholas 609 altenai, Xylopholas 609 alternata, Cardiomya 512, 566, 595 Eurytellina 521, 614 Neaera 558 Tellina 552, 559, 568, 570, 572-573, 583, 616 Tellina (Eurytellina) 554 Alveinus ojianus 1-3, 3, 5, 12-13, 15 alveolus, Talostolida teres 139 Alviniconcha hessleri 178, 178 amekiensis, Bonellitia 296 americana, Abra longicallis 520, 539, 611 Arca (Argina) 559, 591 Arca campechiensis 567, 591 Elliptotellina 521,614 (Elliptotellina) Tellina 553 Ensis 559, 609 Glycimeris 587 Glycymeris 513, 552, 567, 570, 572, 597 Tellina 570, 572, 616 Volsella 583, 604 americanus, Lithopoma 582 Modiolus 516, 531-535, 548, 551, 564-565, 570, 572, 578, 586, 588, 604 Spondylus 520, 530, 533, 552-553, 557, 568, 570-572, 580-581, 584, 587-588, 613 Americardia 576, 592 guppyi 511, 549-550, 569, 571, 591 media 511, 533, 565, 569-571, 584-586, 588, 591 Americardium media 556 amianta, Lucina 572 Parvilucina 577 Radiolucina 515, 533, 601 amiantus, Linga 570, 584, 602 Lucina 576, 601-602 Lucina (Bellucina) 567, 601 Parvilucina (Bellucina) 556, 602 amiges, Ovatipsa chinensis 138-139, 154 amoena, Helicina 224 amorimi, Barycypraea fultoni 136 Amphidesma laeta 548 variegata 548 Amphidromus 217 Amphioplus sepultus 564 Amphipholis gracillima 564 Amusium 608 laurenti 563 laurentii 571, 606 papyraceum 571, 576-577, 606, 608 Amussium lucidum 558, 610 Amygdalum 585, 602 arborescens 576, 602 dendriticum 602 papyrium 516, 571, 576, 602 politum 516, 571, 602 sagittatum 516, 571, 584-585, 602 amygdalumtostum, Barbatia 590 Anadara 369, 372, 378, 578, 584, 589-590 baughmani 510, 548, 571, 576, 589 brasiliana 369, 589 (Caloosarca) notabilis 585 chemnitzii 589 floridana 510, 571, 586, 589 granosa 295 lienosa floridana 570 nobilis 563 notabilis 356, 369, 377, 510, 530-531, 933. 555, 551, 563, 569-571, 577. 580, 583, 585-586, 588-589 ovalis 571, 590 springeri 548, 577, 581, 589 transversa 510, 571, 576, 590 trapezia 372, 374 analoga, Xylodiscula 178, 178, 179 Anatina 602 anatina 515, 571, 602 lineata 567, 602 plicatella 570 (Raeta) canaliculata 567, 602 anatina, Anatina 515, 571, 602 Anodonta 208 androyensis, Palmadusta 143 angasi, Ostrea 316-317, 322 angelicae, Pseudozonaria 137 Zonaria 135, 138, 143 Zonaria pyrum 152 angioyorum, Erronea 143 angolensis, Pseudozonaria pyrum 137 Zonaria pyrum 152 INDEX 627 anguilosa, Tellina 573,614 Angulata brasiliensis 224 angulifera, Periploma 559, 561, 566, 568, 579-580, 609 anguliferum, Periploma 550, 572, 609 angulosa, Eurytellina 521,614 Tellina 568, 570, 572, 616 Tellina (Acropagia) 580 Tellina (Arcopagia) 566 Tellina (Eurytellina) 549, 554, 561, 566, 580 Angulus 616-617 agilis 521,613 mera 613 merus 521, 530-532, 534-535, 613 paramera 549 paramerus 521,613 probrina 549 probrinus 521,613 sybariticus 521,613 tampaensis 521, 613 tenellus 521,614 texanus 521, 614 versicolor 521, 614 angusta, Notocypraea 137 angustata, Notocypraea 138, 152 Animalocardia brasiliana 567, 618 cuneimeris 567 Anisotremus virginicus 423 Annepona 131 mariae 135, 151 annettae, Pseudozonaria 135, 138, 152 annulus, Monetaria 133, 148 Anodonta 205, 208 anatina 208 cygnea 208 grandis 208 piscinalis 208 subcircularis 208 woodiana 205-208, 206-207 Anodontia alba 292, 514, 552, 570-571, 583, 599 (Anodontia) 599 (Anodontia) alba 556 (Anodontia) schrammi 556 philippiana 572, 583, 599 schrammi 514, 599 anomala, Macoma 579, 610 Anomalocardia 427, 554-555 auberiana 523, 531, 534-535, 555, 569, 572, 575, 583-584, 618 brasiliana 618 brasiliensis 567, 618 cuneimeris 565, 568, 570, 577, 583, 588, 618 628 cuneiveis 578, 618 Пехиоза 558, 618 impressa 571, 618 producta 618 Anomia simplex 510, 559, 567, 570, 572, 578, 583, 589 Anomiidae 510, 589 Antigona 429, 444, 619-620 (Antigona) listeri 430 caribbeana 432 (Circomphalus) 619 (Circomphalus) rigida 566 (Circomphalus) strigillina 566 dominica 432 (Dosina) 620 (Dosina) listeri 430, 575 lamellaris 618 listeri 430, 441, 549-550, 556, 563, 567, 570, 576-577, 579, 588-589 (Periglypta) listeri 430 rigida 567, 586 strigillina 550, 567, 571, 619 (Ventricola) 619 (Ventricola) rigida 566 (Ventricola) strigillina 566 (Ventricolaria) 619 (Ventricolaria) rigida 565 antilarum, Lithophaga 587, 603 antillarum, Brachtechlamys 518, 572, 607 Bractechlamys 569-570 Cardium 559 Cardium (Trigoniocardia) 567 Cumingia 563, 576, 586, 611 Cummingia 562, 611 Lithodomus 556, 567 Lithophaga 339, 342, 343, 516, 552, 556, 563, 566, 569-570, 572, 576, 582, 584, 586-588, 603 Lithophagus 559-560 Lyropecten 565, 570, 576, 578, 586, 608 Pecten 558, 568, 579, 585-588 Pecten (Chlamys) 560 Pecten (Lyropecten) 566, 580 Pecten (Pecten) 559 Trigoniocardia 511, 593 antillensis, Cratis 519, 539, 609 Limopsis 559 Sphenia 572, 602 antiqua, Venus 305 aplysioides, Crepidula 185-186, 198-201 Apolymetis intasriata 567, 615 appressa, Chama 401-403 arabica, Mauritia 135-136 Mauritia arabica 135, 150 arabicula, Pseudozonaria 135, 137-138, 152 arborescens, Amygdalum 576, 602 Modiolaria 567, 602 Агса 328, 372, 393, 563: 568, 571: 574 578, 585-586, 589-591, 605 (Acar) 604 (Acar) adamsi 579-580 (Acar) reticulata 579-580, 589 adamsi 567 admsi 567, 604 (Anadara) floridana маг. secernenda 589 (Arca) 590 (Arca) imbricata 559 (Arca) noae 559, 590 (Arca) zebra 551 (Argina) americana 559, 591 auriculata 566-567, 577, 579-580, 589 balesi 591 barbata 355, 553, 563, 567, 569, 580-581, 590 barbadensis 579, 590 (Barbatia) balesi 548, 550, 575, 591 (Barbatia) barbata 559, 579-580, 590 (Barbatia) dominguensis 558, 589 (Barbatia) gradata 558, 589 (Byssoarca) 591 (Byssoarca) adamsi 559, 604 (Byssoarca) glomerula 559 (Byssoarca) nodulosa 560, 589 (Byssoarca) reticulata 559, 589 campechiensis americana 567, 591 cancellaria 562 candida 567, 579, 587 chemnitzi 567 deshayesii 579, 589 domingensis 579, 589 fusca 579, 590 glomerula 552 gradata 355, 579, 589 imbricata 328, 378, 510, 530-535, 551, 562, 565, 567, 569-572, 574 5771579; 585-586, 588, 590 incongrua 567, 579, 591 lienosa floridana 589 listeri 581, 590 (Lunarca) occidentalis 560, 590 (Lunarca) umbonata 560, 590 (Macrodon) 559, 591 (Macrodon) sagrinata 559 (Navicula) umbonata 579-580, 590 noae 560, 571, 590 поае var. americana 579, 590 nobilis 589 (Noetia) 605 (Noetia) orbignyi 559, 591 (Noetia) ponderosa 559 notabilis 372 occidentalis 567, 569, 580, 590 plicata 581, 589 ponderosa 567 reticulata 567, 589 saccharina 575, 589 (Scapharca) 590 (Scapharca) auriculata 559, 589 (Scapharca) incongrua 559, 591 (Scapharca) lienosa 559, 589 (Scapharca) transversa 559 secticostata 567, 589 transversa 551, 567, 577, 579-580 umbonata 553, 560, 565-567, 580-581, 583, 590 velata 574, 590 zebra 356, 369, 377, 510, 530-533, 535, 562—566, 569-570, 572, 574, 577-580, 583-586, 588-590 arcana, Chama 391 Arcas 590 Archaeoxesta 83 Arcidae 247, 355, 510, 557, 584-585, 589 Arcinella 381, 385-387, 390, 408, 593 arcinella 387, 408-409, 561, 593 brasiliana 408 соти 382, 408—409, 409, 411, 511, 556, 569-570, 572, 584, 593 (Nicolia) cornuta 410 arcinella, Arcinella 387, 408-409, 561, 593 Cardium 567, 593 Chama 408, 556, 559, 567, 593 Chama (Echinochama) 560, 593 Echinochama 408 Pseudochama (Echinochama) 408 Arcinelloidea 385 Arcoidea 327, 336, 355, 583 Arcopagia 616 fausta 521, 530-531, 533, 535, 551, 553, 556, 563, 577, 583, 588, 614 Arcopsis 327-328, 331, 332, 336-337 adamsi 327-331, 332-335, 336, 516, 530-535, 551, 554-555, 562, 565, 569; 571-572, 574, 583-584, 588-589, 604 afra 327, 332, 336 solida 327, 336 arctica, Hiatella 514, 572, 584-585, 597 Saxicava 559 Arcticidae 12 arenaria, Mya 208, 296 arenosa, Pandora 517, 585, 606 Arestoides 131 argus 136 argus argus 135-136 629 argus contrastriata 135-136 argentea, Pteria 491 argentina, Crepidula 198-201 Argopecten 585, 606-607 gibbus 518, 530, 533, 563, 567, 569, 572, 584-586, 606 irradians 518, 531, 534, 548, 550, 555, 562,565, 569 572: 583, 606 irradians concentricus 548, 565, 570, 586, 606 irradians taylorae 548, 570, 575, 582, 606 lineolaris 572 nucleus 518, 553, 563, 569, 572, 586, 607 Argopectin 607 nucleus 575 argus, Arestoides 136 Arestoides argus 135-136 Lyncina (Arestoides) argus 151 ariakensis, Crassostrea 316 aristata, Lithodomus 567 Lithophaga 339, 343, 344, 516, 551, 563, 570, 572, 582, 584, 603 Armandiella 83 armatum, Ascaulocardium 50 armeniaca, Umbilia 133, 135, 149 artuffeli, Palmadusta 138-139, 153 Asaphis 249-252, 255, 269-272 deflorata 249-255, 252-253, 256-261, 258-262, 263-267, 264-265, 268-273, 519, 559, 562, 567, 569-570, 572, 575, 579, 610 dichotoma 249-250, 571, 610 violascens 249-251, 253, 253-255, 259-261, 263-265, 268-272, 610 Ascaulocardium armatum 50 asellus, Palmadusta 139 Palmadusta asellus 138-139 Palmadusta cf. asellus 153 asiatica, Mauritia arabica 135, 150 asperella, Chama 398 Aspergillum agglutinans 39 dichotomum 50 novaehollandiae 39, 39 aspersa, Helix 73, 211-212, 215-216 aspersum, Cornu 73 aspersus, Cantareus 73 Cryptomphalus 73 Astarte crenata subequilatera 510, 581, 591 globula 510, 591 lens 559, 591 nana 510, 559, 561, 567, 572, 576, 591 smithii 510, 559, 591 630 Astartidae 510, 591 astaryi, Cribrarula 128, 128, 138, 154 Asthenothaerus 585, 617 (Asthenothaerus) balesi 550, 617 balesi 548, 550, 569, 572, 577, 581,617 (Bushia) 617 (Bushia) elegans 55 hemphilli 523, 559, 567, 572, 584-585, 617 athlia, Bruceiella 172, 173, 179 atlantica, Avicula 556, 560, 567, 611 Cooperella 518, 576, 609 atra, Aulacomya 186-187, 187 Atrina 583, 610 rigida 519, 551, 565-566, 569-570, 572, 578, 582, 588-589, 609 seminuda 186-187, 187, 519, 563, 569, 572.578, 610 serrata 519, 572, 582, 610 auberiana, Anomalocardia 523, 531, 534-535, 555, 569, 572, 575, 583-584, 618 Aulacomya atra 186-187, 187 aupouria, Ostrea 309, 311-312, 314-318, 318,321, 222 323 aurantia, Lucina 293 aurantiaca, Melanodrymia 169, 174-175, 174 aurantium, Lyncina 135 Lyncina (Callistocypraea) 151 aureola, Lucidella 224 auriculata, Arca 566-567, 577, 579-580, 589 Arca (Scapharca) 559, 589 Scapharca 571, 589 Scapharca (Scapharca) 560, 571, 589 Aurinia schmitti 571 aurita, Limopsis 514, 559, 572, 599 australiensis, Cribrarula cribraria 140, 154 Cribrarula cf. cribraria 138, 140 Austrasiatica 127, 131-132, 140 alexhuberti 143 deforgesi 143 hirasei 132, 138, 140-141, 155 langfordi 132, 140, 154 langfordi cavatoensis 138, 141 sakurai 132, 138, 140-141, 155 Austrocypraea 131 reevei 135 Austrocypraeini 136 Austroginella 295 Austrovenus 443 Avicula 473, 475, 556, 558, 564, 611 ala-perdicis 579, 611 atlantica 556, 560, 567, 611 crocata 491, 560, 611 guadalupensis 491 longisquamosa 476 longisquamosa (Meleagrina) 476 longisqvamosa 476 margaritifera 574 nitida 559, 611 radiata 560, 579,611 Aviculidae 475 azaria, Hiatella 514, 597 Saxicava 559 balesi, Arca 591 Arca (Barbatia) 548, 550, 575, 591 Asthenothaerus 548, 550, 569, 572, 577, ASIA Asthenothaerus (Asthenothaerus) 550, 617 balthica, Macoma 167 Bankia carinata 522, 572, 617 fimbriatula 523, 617 barbadensis, Arca 579, 590 barbata, Arca 355, 553, 563, 567, 569, 580-581, 590 Arca (Barbatia) 559, 579-580, 590 Barbatia 356 Barbatia 355-356, 366-367, 369, 372, 374, 377-378, 566, 571, 578, 589, 591 (Acar) 589 (Acar) domingensis 571 (Acar) dominguensis 363 amygdalumtostum 590 barbata 356 (Barbatia) cancellaria 356 (Barbatia) candida 360 cancellaria 355-356, 357, 361-363, 365, 366-367, 368-369, 370-371, 372, 374-375, 377-378, 510, 527, 530-535, 551, 556, 563, 565-566, 569-570, 572, 576-579, 583, 585-586, 588-590 candida 355, 360, 361, 363, 366-367, 369, 372, 373, 374-375, 377-378, 551, 570, 572, 579, 585-586, 588, 591 (Cucullaearca) candida 360 domingensis 562-563, 569, 572, 577, 579, 585-586, 591 dominguensis 355, 363, 364, 368-369, 372, 374, 377-378 (Fugleria) tenera 365 tenera 355, 365, 366-367, 369, 372, 374, 376, 377-378, 549, 569-570, 579, 591 barbieri, Purpuradusta 143 barclayi, Contradusta 141-142, 156 Erronea 142 Barnea costata 583 truncata 519, 570, 572, 579-580, 584, 609 barrattiana, Corbula 559, 567, 572, 594 bartschi, Teredo 523, 576, 617 Barycypraea 127, 130-132, 134 fultoni 132, 136, 150 fultoni amorimi 136 fultoni massieri 135 teulerei 132, 135-136, 150 Bassina 296, 305, 427 Basterotia elliptica 521, 569, 572, 613 quadrata 521, 559, 567, 569, 572, 613 quadrata granatina 567, 613 Bathyarca glomerula 510, 572, 591 inaequalis 510, 591 Bathymargarites symplector 173, 174, 180 Bathymodiolus 52 Bathynerita 181 naticoidea 176-177, 176, 181 naticoides 169 Bathyxylophila excelsa 176 baughmani, Anadara 510, 548, 571, 576, 589 beana, Entodesma 515, 570, 572, 602 Lyonsia 559, 567, 584, 602 beatrix, Helicina 217 Helicina beatrix 220, 222 beaui, Venus 559, 579, 618 beckii, Erosaria 133-134, 148 belizana, Lucina 292 bellastriata, Semele 520, 530, 547, 551, 554-555, 568-572, 580, 583-585, 588, 612 bellestriata, Semele 612 benedicti, Chlamys 563, 570, 607 Spathochlamys 518, 572, 608 Bentharca 591 sagrinata 510, 539, 591 berinii, Notadusta punctata 141, 156 berjadinensis, Chama 387, 411 bermudensis, Callucina 622 Chama 395-396, 398, 401 Chama sinuosa 398, 400, 553, 594 Bernayinae127, 131, 134, 136 bicolor, Isognomon 356, 377, 514, 533, 549-551, 562-563, 565, 570, 572, 574, 576-577, 586, 588-589, 598 Pedalion 568 bicornis, Chama 395 bimaculata, Heterodonax 559, 567, 610 bimaculatus, Heterodonax 258-259, 261, 203 272. 51915721579. 610 binodata, Pseudaspasita 92, 102, 112 Bistolida 131, 139 brevidentata 143 INDEX 631 erythraeensis 138-139, 141, 153 goodallii 138, 141, 153 hirundo 138, 141, 153 kien depriesteri 138, 141 kieneri 139 kieneri depriesteri 154 kieneri kieneri 138, 141, 154 owenii 138-139, 153 owenii vasta 141 stolida clavicola 138, 141, 153 stolida diagues 138-139, 141, 153 stolida rubiginosa 138-139, 141, 153 stolida stolida 138, 141, 153 ursellus 138-139, 141, 153 bistrinotata, Pustularia 137 Pustularia bistrinotata 135, 138, 152 bisulcata, Lithophaga 339, 344, 345, 516, 551-552, 570, 572, 582, 584, 604 bisulcatus, Lithodomus 567, 579, 604 Lithophagus 559, 604 bitaeniata, Palmadusta asellus 138-139 Palmadusta cf. asellus 153 blanda, Cavilinga 515, 556, 599, 622 Lucina 584 Parvilucina 570 Blasicrura 131, 140, 143 interrupta 141, 156 pallidula 143 pallidula pallidula 141, 156 pallidula rhinoceros 141, 143, 156 pallidula cf. vivia 141 summersi 141, 143, 156 blauneri, Hyalinia var. cloacarum 21 boeckeleri, Alcadia 217, 220, 223 boivinii, Егозапа 133, 149 Bombylidae 429 Bonellitia amekiensis 296 borealis, Cyclocardia 581, 622 Bostrycapulus aculeatus 198-200 Botula 339, 603 castanea 567, 576, 603 cinnamonea 342 fusca 339-341, 341, 353, 516, 552, 556, 562-563, 567, 569, 572, 576, 578, 584, 602 semen 579, 603 boucheti, Palmulacypraea 143 bowmani, Lepton 579, 622 Brachidonta 603 Brachidontes 278, 328, 555, 573-574, 603 citrinus 567, 576-577, 603 domingensis 569, 572, 602 exustus 516, 527, 530-532, 534-535, 551, 555, 565, 569-570, 572, 577, 583-584, 588-589, 602-603 632 granulatus 292 modiolus 516, 562, 569, 572, 575, 603-604 pharaonis 208 recurvus 570, 603 Brachiodonta exustus 575 Brachiodontes 554-555, 588, 603 domingensis 603 recurvus 566, 603 Brachtechlamys 608 antillarum 518, 572, 607 Bractechlamys antillarum 569-570 Bradybaena 81, 85, 106-108 similaris 83, 92, 111 Bradybaenidae 79-80, 84, 98, 109 Bradybaeninae 79, 107, 109 brasiliana, Anadara 369, 589 Animalocardia 567, 618 Anomalocardia 618 Arcinella 408 Iphigenia 513, 567, 570, 572, 596 Mactra 559, 602 Scapharca 510, 591 Tellina 614 brasiliensis, Angulata 224 Anomalocardia 567, 618 braziliana, Iphigenia 559, 596 Tellina 556, 587, 614 Brechites 37-38, 50, 53 (Foegia) novaezelandiae 38-39 (Foegia) veitchi 38 (Penicillus) philippinensis 38 penis 50 vaginiferus 37-38, 41—42, 45, 47—48, 50, 52 bregeriana, Contradusta 141, 156 brevidentata, Bistolida 143 brevifrons, Macoma 521, 559, 567, 570, 572, 584, 615 brevirostris, Pustularia globulus 135, 138, 151 brevis, Prionovolva 133, 147 briandi, Shinkailepas 177, 177, 180-181 broderipii, Lyncina 135-136 Lyncina (Callistocypraea) 151 bronniana, Limea 514, 539, 599 Bruceiella athlia 172, 173, 179 Bryopa 37, 50 buchivacoana, Pseudochama 387, 411 bullata, Papyridea 559, 592 bushae, Leda 605 Bushia 584, 617 elegans 523, 585, 617 bushiana, Pandora 517, 554, 572, 606 Byssoarca lima 355 Bythiospeum 229-230 INDEX cachimilla, Crepidula 185-186, 187-188, 190, 192-194, 196-197, 198-201 caerulea, Albinaria 23 caimitica, Chama 387, 411 calcarea, Macoma 167, 549-550, 615 calcicola, Nucula 517, 548, 562, 572-573, 605 californianus, Mytilus 208, 549-550, 604 Callista 444 (Dione) gigantea 571,620 eucymata 523, 530, 533, 572, 618 callista, Erosaria helvola 134 Erosaria cf. helvola 133-134, 148 Callistocypraea 131, 136 Callocardia 277 Callucina bermudensis 622 (Callucina) radians 556, 599 keenae 515, 599, 601 Calocochlea 81, 106-109 coccomelos 85, 89, 89, 112 caloosana, Pseudochama 387, 411 calophyllum, Placamen 288, 292, 295-296, 303, 305 Calpurnus lacteus 147 verrucosus 133, 147 Calyptogena 13, 52 Calyptraeidae 200 Calyptraeoidea 185 Camaena 79, 81, 106-107 platyodon 105, 112 Camaenidae 79, 84, 112, 217 camelopardis, Lyncina 143 campechiensis, Mercenaria 428, 441, 524, 565, 570, 572, 586, 620 Pholas 519, 559, 568, 609 Venus 568 canaliculata, Anatina (Raeta) 567, 602 Labiosa 559, 602 Cancellaria 409 cancellaria, Arca 562 Barbatia 355-356, 357, 361-363, 365, 366-367, 368-369, 370-371, 372, 374-375, 377-378, 510, 527, 530-535, 551, 556, 563, 565-566, 569-570, 572, 576-579, 583, 585-586, 588-590 Barbatia (Barbatia) 356 cancellata, Chione 296, 299, 301, 553-555, 563-572, 575, 577-578, 583-587, 589, 618-619 Semele 559, 612 Venus 558-559, 618 Venus (Chione) 560, 618 cancellatum, Parvamussium 519, 610 Pecten (Amusium) 560 Pecten (Propeamussium) 560 Propeamussium 539 cancellatus, Venus (Chione) 568, 618-619 candeana, Diplodonta 579 Felaniella 523, 618 Fundella 552, 602 Scissula 522, 615 Пета! 550, 565, 568, 570, 572, 576, 587, 616 Tellina (Angulus) 566, 580 Tellina (Scissula) 549, 554, 561, 566, 574, 580 candeanus, Malleus 489, 516, 554, 570, 572, 586, 602 candida, Arca 567, 579, 587 Barbatia 355, 360, 361, 363, 366-367, 369, 372, 373, 374-375, 377-378, 551, 570, 572, 579, 585-586, 588 Barbatia (Barbatia) 360 Barbatia (Cucullaearca) 360 Cucullaearca 510, 533, 591 Palmadusta clandestina 138-139 Palmadusta cf. clandestina 153 candigerus, Lithodomus 603 canrena, Naticarius 295, 297, 302, 303, 305 Cantareus aspersus 73 capensis, Cypraeovula 137-138, 152 capricornica, Umbilia 133-135, 149 caputdraconis, Monetaria 133, 148 caputophidii, Monetaria 133 Monetaria caputophidii 148 Monetaria caputserpentis 133 caputserpentis, Monetaria 133 Monetaria caputserpentis 148 Cardiidae 511, 563, 585, 591 Cardiomya 584, 595 alternata 512, 566, 595 costellata 512, 572, 583, 595 glypta 512, 572, 595 ornatissima 512, 538, 572, 595 perrostrata 512, 572, 576, 584-585, 595 Striata 512, 533, 595 Cardita 593 (Carditamera) floridana 561, 582 conradi 593 conradii 559, 593 domingensis 559, 576, 593 floridana 552, 559, 565, 567, 576, 579, 583 (Mytilicardia) floridana 571, 593 Carditachama 385 Carditamera (Carditamera) 593 floridana 511, 530-535, 570, 572, 575, 583, 593 Carditidae 12, 511, 584-585, 593 INDEX 633 Carditinae 1 Carditopsis smithii 512, 548, 570, 572, 594 Cardium 591-593 antillarum 559 arcinella 567, 593 (Fulvia) peramabilis 548, 558, 592 (Fulvia) peramabilis var. tinctum 558, 592 (Hemicardium) medium 567, 591 isocardia 559, 567, 592 isocardium 574, 592 (Laevicardium) laevigatum 567, 592 (Laevicardium) serratum 567, 592 laevigatum 558 magnum 559, 567, 575 medium 559-560, 579, 591 mortoni 581 muricatum 559, 567, 571 (Papyridea) semisulcatum 561, 567, 592 (Papyridea) spinosum 567, 592 peramabilis 559, 592 peramabilis var. tinctum 560 petitianum 579, 592 (Protocardia) peramabilis 567, 592 serratum 560, 592 (Trigoniocardia) 593 (Trigoniocardia) antillarum 567 Caribachlamys 607-608 imbricata 518, 572, 586, 607 mildredae 518, 549, 572, 586, 607 ornata 518, 564, 572, 586, 607 sentis 518, 531, 533, 553, 564-565, 572, 576, 586, 607 caribaea, Caryocorbula 512, 594 Corbula 572 Lima 514, 530-531, 533, 572-573, 576, 598-599 Martesia 567, 609 caribbea, Cercaria 423 caribbeana, Antigona 432 carinata, Bankia 522, 572, 617 caribaeus, Lithophagus 559, 604 carinifera, Thais 295 carlottensis, Macoma 613 carnaria, Strigilla 522, 554, 559, 568, 570-571, 616 carnea, Pinna 519, 559-560, 562, 568-570, 572, 575, 578-582, 588, 610 carneola, Lyncina 135, 151 carnosa, Lucina 291, 293 carolinensis, Cyrena 558, 594 Cyrena (Leptosiphon) 559, 594 caroliniana, Cyrena 594 carpenteri, Leda 558 Leda (Leda) 560, 605 Nuculana 568, 576, 581 634 Propeleda 517, 605 Caryocorbula caribaea 512, 594 chittyana 512, 594 contracta 512, 594 cymella 512, 595 dietziana 512, 595 casina, Circomphalus 428, 443-444 caspari, Nesiohelix 103, 112 castanea, Botula 567, 576, 603 Cypraeovula 138-139, 152 Ervilia 258, 296 castaneus, Lioberis 551 Lioberus 516, 551, 570, 572, 584-585, 603 Pectunculus 579 Cataegis meroglypta 180 catena, Natica 296 Polinices 296 Cathaica 81, 85, 106-109 (Cathaica) fasciola 92, 102, 111 (Pliocathaica) 79, 109 (Pliocathaica) gansuica 89, 93, 111 catholicorum, Cribrarula 128, 128, 138, 140, 154 caudigerus, Lithodomus 603 Caulerpa verticillata 499 caurica, Етопеа 141-142, 155 Erronea caurica 141-142, 155 cavatoensis, Austrasiatica langfordi 138, 141 Cavilinga blanda 515, 556, 599, 622 caymanana, Lucina podagrina 293 Cepaea 217 hortensis 73 nemoralis 73 Cephalaspidea 13 Cercaria caribbea 423 cerina, Circe 567 Circe (Gouldia) 559 Gouldia 524, 533, 567, 570, 572, 584-585, 619 Macoma 522, 559, 567, 572, 576, 615 cernica, Erosaria 133, 149 cervinetta, Macrocypraea 135, 149 cervus, Macrocypraea 135, 149 Cetoconcha margarita 610 Chama 328, 381-390, 392, 393, 394, 397, 401, 403, 404, 405—406, 408, 410-411, 556, 5621567. 571. 574. 590.594 appressa 401—403 агсапа 391 arcinella 408, 556, 559, 567, 593 asperella 398 berjadinensis 387, 411 bermudensis 395-396, 398, 401 INDEX bicornis 395 caimitica 387, 411 chinensis 398 chipolana 387, 394, 411 cistula 398, 401, 403 citrea 395 congregata 381-389, 391-392, 393, 394-396, 410—411, 511, 530, 533, 591-932, 556, 563, 567, 570 572457070 584, 586, 588, 593 congregatoides 392 coralliophaga 410 cornuta 398 crassa 410 cristella 388-391, 410 damaecornis 396 (Echinochama) arcinella 560, 593 emmonsi 387-388, 405, 411 ferruginea 401—402 florida 382-385, 388-389, 392, 396, 397, 398, 403, 405, 410-411, 511, 551, 553, 556, 962, 5727595 foliacea 392, 394 gardnerae 405 gryphina 403 gryphoides 386, 395 heilprini 387, 398, 410-411 imbricata 395-396 inezae 382-386, 388, 405-406, 408, 410-412, 511, 549, 556, 593 involuta 387, 411 iudicai 389, 395, 411 lactuca 382, 386, 388, 403, 404, 408, 411, 511, 556, 570-572, 593 lamarckiana 398, 401 lamellosa 392, 394 lazarus 389, 395-396, 410, 593 linguafelis 405, 410 lobata 410 macerophylla 382-389, 393, 395-396, 397, 398, 401-402, 405, 408, 410-411, 512, 531, 533, 553, 090=99/ 099 563-564, 567, 570-572, 574, 576-577, 579, 585-586, 588, 593 macerophylla var. purpurascens 395 macerophylla var. sulphurea 395 macrophylla 395, 556, 581 расйса 398 paschauli 387 paschuali 411 pellucida 387, 391 prætexta 401 producta 410 pulchella 390 radians 382-384, 386, 388-392, 400, INDEX 401-403, 402, 405, 410-412, 512, 556, 594 radians ferruginea 402 radians radians 402 radians variegata 402 reevana 390 rotunda 402 rubea 410 ruderalis 401, 408 rugosa 392, 394 ruppelli 390 sarda 381-384, 386, 388-389, 396, 398, 403, 404, 405, 408, 409, 410-412, 512, 550, 552-553, 556, 560, 562, 565-567, 570, 572-573, 575, 577, 579-580, 586, 594 sarda lutea 403 sardo 570, 594 sinistrorsa 390, 403 sinosa 551, 594 sinuosa 382, 388-389, 392, 396, 398, 399, 401-403, 411, 512, 556-557, 570, 572, 594 sinuosa bermudensis 398, 400, 553, 594 sinuosa firma 398, 399, 553, 594 spinosa 398 squamosa 392, 394 strepta 387, 411 tumulosa 398 variegata 401-402, 587, 594 willcoxii 386-387, 396, 411 Chamelea 427, 443 Chamidae 247, 381-382, 385, 388-389, 409, 511, 557, 562, 593 chazaliei, Euvola 518, 549, 572, 608 Pecten 564, 570, 608 Pecten (tereinus) 571,608 Chelycypraea 131 testudinaria 135, 151 chemnitzi, Arca 567 chemnitziana, Isognomon 580, 598 chemnitzii, Anadara 589 Scapharca 510, 591 Venus 441 chentingensis, Pseudiberus (Platypetasus) 104, 112 Chesapecten 394 chiapponii, Pustularia 143 childreni, Ipsa 130, 133, 148 chilensis, Ostrea 314, 316-317 chinensis, Chama 398 Ovatipsa 139 Ovatipsa chinensis 138-139, 154 Chione 295-296, 305, 427, 443, 584-585, 618-621 635 cancellata 296, 299, 301, 553-555, 563-572, 575, 577-578, 583-587, 589, 618-619 (Chione) 621 (Chione) pubera 550 (Chione) intapurpurea 561 (Chione) subrostrata 561, 618 cigenda 574, 618 clenchi 618 elevata 295-299, 298-300, 301-305, 303-304, 523, 527, 530-535, 573, 577, 618 erosa 296 grus 551, 563-564, 570, 584-585, 619 intapurpurea 551, 563, 570, 579-580, 619 latilirata 570, 578, 587, 619 (Lirophora) 620 (Lirophora) paphia 549 listeri 441 mazyckii 523, 531, 572, 619 paphia 552, 565, 570, 572, 581, 585-587, 619 puber 570, 621 pubera 566, 579-580, 619 pygmaea 570, 575, 583, 619 (Tellina) 621 (Timoclea) 621 (Timoclea) granulata 566, 579-580, 618-619 (Timoclea) grus 549, 551 (Timoclea) pygmaea 561, 566, 579-580 undatella 301 Chioninae 295-296, 427 chipolana, Chama 387, 394, 411 chiquitica, Helicina 217-218, 220, 223 chittyana, Caryocorbula 512, 594 Chlamys 607-608 benedicti 563, 570, 607 imbricata 551, 562, 570, 578, 588, 607 imbricatus 570, 576, 587 mildredae 570, 608 multisquamata 570, 607 muscosus 576 nucleus 578 ornata 562, 570, 588, 608 phrygius 576, 608 sentis 551, 556, 562-563, 565-567, 570, 575-578, 586-589, 608 Chondria 486 Choristodon 339, 347-348, 349, 350, 584, 609 robustum 339, 346, 346-347, 350-351, 518, 572, 584, 609 typica 350 636 typicum 579, 609 chlorizans, Erosaria cf. erosa 149 chrysalis, Purpuradusta microdon 138, 141-142, 156 chrysostoma, Erronea ovum 141, 155 Loripes 571, 599 Lucina 561, 599 Lucina (Loripinus) edentula 567, 599 cicercula, Pustularia 135, 138, 152 cigenda, Chione 574, 618 cimula, Macoma 567, 615 cinerea, Luria 133, 135, 151 cinnamomea, Modiola (Botula) 559, 602 Modiolaria 579, 602 cinnamonea, Botula 342 Circe 619 cerina 567 (Gouldia) 619 (Gouldia) cerina 559 circe, Gari 519, 538, 610 Circinae 1, 427, 444 circinata, Cytherea 579, 620 circinatus, Pitar 524, 620 Circomphalus 443 casina 428, 443-444 strigillinus 441, 524, 548, 572, 619 Cirridae 180 cistula, Chama 398, 401, 403 citrea, Chama 395 citrina, Erosaria 133-134, 148 citrinus, Brachidontes 567, 576-577, 603 clandestina, Palmadusta 139 Palmadusta clandestina 138-139 Palmadusta cf. clandestina 153 clappi, Teredo 523, 551, 568, 573, 617 Teredo (Zopoteredo) 549, 552, 582 clarus, Oxychilus 33 clathrata, Venus 441 Clausina 428 Clausinella 427, 443-444 Clavagella 37, 50, 53 Clavagellidae 53 Clavagelloidea 37-38 clavicola, Bistolida stolida 138, 141, 153 claviculata, Leiomya 595 clenchi, Chione 618 Lirophora 524, 620 coarctata, Cumingia 520, 549, 561, 567, 570, 572, 574, 579-580, 584, 611 Coccoglypta 83 coccomelos, Calocochlea 85, 89, 89, 112 cochlear, Neopycnodonte 310, 312, 315-316, 514, 572, 597-598 Cochliolepis parasitica 566 Cochlodesma pyramidatum 518, 550, 609 Codakia 420, 423, 554-555, 568-569, 583-584, 600-601 (Codakia) 600 (Codakia) orbicularis 556 costata 570, 572, 575 (Ctena) 600 (Ctena) orbiculata 556 (Ctena) pectinella 550, 564 cubana 622 (Jagonia) 600 (Jagonia) orbiculata filiata 566, 600 (Jagonia) orbiculata recurvata 566, 600 (Jagonia) pectinella 566 obicularis 577, 600 orbicular 586, 600 orbicularis 279, 288, 292, 417-418, 420, 420-421, 422, 423-424, 515, 527, 530-535, 551, 553, 556-557, 561, 563, 565-567, 570, 572-574, 576, 578, 580, 582-589, 600, 622 orbiculata 564-565, 567-568, 570, 572, 577-578, 583, 600 orbiculata form filiata 600 pectinella 570, 572 punctata 424 colligata, Cypraeovula 143 coloba, Ovatipsa 138, 154 columbianus, Agrolimax 215 colymbus, Pteria 473-474, 476, 489-493, 520, 533, 547, 563-565, 570, 572-575, 580-581, 588, 611 comma, Cribrarula cribraria 138, 154 comandorica, Kellia 57-59, 60-62, 62, 64-66, 71 commercialis, Saccostrea 316 compressa, Myrtea 547 comptoni, Notocypraea 138, 152 concentrica, Dosinia 566-567, 579-580, 619 Dosinia (Dosinidia) 561, 619 Ervilia 520, 559-560, 562, 566-567, 570, 572, 579-580, 584-585, 612 Nuculana 517, 572, 584-585, 605 concentricus, Argopecten irradians 548, 565, 570, 586, 606 Pecten irradians 549 conchaphila, Ostrea 316 Ostreola 317, 322 conchyliophora, Xenophora 564 concinna, Primovula 132-133, 147 Condylocardia floridensis 548, 550, 575, 594 Condylocardiidae 1, 512, 594 Coneulota 83 confusa, Helicina beatrix 220-221 Limatula 514, 567, 599 Congeria rossmassleri 567, 596 rossmässleri 566, 596 congregata, Chama 381-389, 391-392, 393, 394-396, 410-411, 511, 530, 533, 55.552.556, 563, 567, 570, 572. 577, 584, 586, 588, 593 congregatoides, Chama 392 connelli, Cypraeovula 138, 152 conradi, Cardita 593 conradiana, Cytherea (Transennella) 567, 621 conradii, Cardita 559, 593 conradina, Cytherea 579 Cytherea (Transennella) 559, 621 Meretrix (Transennella) 561 Transenella 566, 568, 573, 587, 621 Transennella 525, 580, 584, 621 consobrina, Scissula 522, 615 Tellina 572, 616 consorbrina, Tellina 570, 615 constricta, Macoma 522, 567, 572, 587, 615 contaminata, Palmadusta 139 Palmadusta contaminata 138, 153 contracta, Caryocorbula 512, 594 Corbula 567, 572, 584-585 Contradusta 131, 140, 142 barclayi 141-142, 156 bregeriana 141, 156 pulchella 141-142, 156 walkeri 141, 156 contrastriata, Arestoides argus 135-136 Lyncina (Arestoides) argus 151 controversa, Luria 143 convexa, Cytherea 579, 622 Cooperella atlantica 518, 576, 609 coquimbensis, Crepidula 198-199, 201 Coralliophaga coralliophaga 410, 523, 552, 567 570, 572, 576, 580; 618 hornbeckiana 579, 609 coralliophaga, Chama 410 Coralliophaga 410, 523, 552, 567, 570, 572,576, 580, 618 Cypricardia 579 Gregariella 516, 570, 572, 603 Corbicula fluminea 48 Corbiculidae 512, 594 Corbula 583, 594-595 barrattiana 559, 567, 572, 594 caribaea 572 (Caryocorbula) 595 (Caryocorbula) cymella 566, 577 contracta 567, 572, 584-585 crassa 305 INDEX 637 cubaniana 559, 595 cymella 547-548, 550, 554, 558-559, 567, 575, 579-580 dietziana 558-559, 567, 572, 595 disparilis 558, 567, 595 krebsiana 559, 595 nasuta 559, 567, 594 swiftiana 551, 560, 567, 572, 579, 594 Corbulidae 512, 584, 594 corbuloidea, Thracia 558-559, 617 corbuloides, Thracia 550, 566, 568, 573, 617 cordata, Pitar 573, 620 Pitar (Pitarenus) 549, 577, 620 Pitaria 578 cordatus, Pitar 524, 549-550, 572, 574, 620 Pitaria 549 Pitar (Pitarenus) 550, 577 Cornu aspersum 73 cornuta, Arcinella 382, 408-409, 409, 411, 511, 556, 569-570, 572, 584, 593 Arcinella (Nicolia) 410 Chama 398 Echinochama 408 Echinochama arcinella 408 coronata, Cypraeovula 138, 152 corticaria, Martesia 559, 609 corticata, Martesia 609 corticosa, Pseudochama 386-387, 411 corticosaformis, Pseudochama 387, 411 Costacallista 618 eucymata 576 costata, Barnea 583 Codakia 570, 572, 575 Cyrtopleura 519, 570, 572, 609 Lucina 579 Lucina (Jagonia) 567 Lucina (Lucina) 559, 601 Parvilucina 515, 601 Parvilucina (Parvilucina) 556 Pholas 571 Pholas (Barnea) 559, 568 costellata, Cardiomya 512, 572, 583, 595 Cuspidaria (Cardiomya) 560, 567 Lirapex 176 costulifera, Xyloskenea 176 coxeni, Eclogavena 136, 141, 156 crassa, Chama 410 Corbula 305 Crassatella (Eriphyla) 595 (Eriphyla) lunulata 559 (Eriphyla) lunulata var. parva 559, 595 floridana 559-560, 595 Crassatellidae 12, 512, 584, 595 638 Crassatellites gibbesii 595 gibbsii 567, 595 Crassinella 584-585, 595 dupliniana 512, 572, 595 lunulata 512, 570, 572, 584-585, 595 martinicensis 512, 570, 572, 584-585, 595 Crassostrea ariakensis 316 gigas 316 rhizophorae 310, 316, 517, 572, 605 virginica 310, 312, 314-316, 318, 319, 322-323, 517, 557, 572, 578, 581, 605 Crassostreinae 310, 312, 320 crassula, Macoma 167 Cratis antillensis 519, 539, 609 crenata, Patella 170, 180 Crenavolva rosewateri 133, 147 cf. rosewateri 133, 147 tokuoi 133, 147 Crenella 573, 603 decussata 516, 572, 603 divaricata 559, 567, 570, 583-585, 603 crenella, Lucina (Parvilucina) 567 Parvilucina 515, 601 Crenellidae 1 crenulata, Lucina (Lucina) 559 Nucula 517, 549, 572, 605 Crepidula 192, 198, 200 aculeata 200 aplysioides 185-186, 198—201 argentina 198-201 cachimilla 185-186, 187-188, 190, 192-194, 196-197, 198-201 coquimbensis 198-199, 201 dilatata 198-199, 201 fecunda 198-199, 201 onyx 185-186, 198-200 philippiana 198-199, 201 plana 185, 200-201 protea 198-201 Creseis 587 cribellum, Cribrarula 154 Cribrarula esontropia 140 cribraria, Cribrarula 140 Cribrarula cribraria 138, 140, 154 Cribrarula 128, 131, 139-140 astaryi 128, 128, 138, 154 catholicorum 128, 128, 138, 140, 154 cribellum 154 cribraria 140 cribraria abaliena 140 cribraria cf. abaliena 138, 154 cribraria abrolhensis 138, 140, 154 cribraria australiensis 140, 154 cribraria cf. australiensis 138, 140 cribraria cribraria 138, 140, 154 cribraria comma 138, 154 cribraria melwardi 138, 154 cribraria rottnestensis 138, 140, 154 cumingii 128, 128, 138, 140, 154 esontropia 154 esontropia cribellum 140 esontropia esontropia 138, 140 esontropia francescoi 138, 140, 154 exmouthensis 154 exmouthensis exmouthensis 138, 140 exmouthensis magnifica 138, 140, 154 fallax 138, 154 garciai 128, 128, 138, 140, 143, 154 gaskoini 128, 128, 138, 154 gaspardi 138, 154 melwardi 140 pellisserpentis 138, 140, 154 taitae 128, 128, 138, 140, 143, 154 crispata, Venus 429, 441, 559, 620 cristagalli, Lopha 316-317 cristallina, Merisca 522, 615 Tellina 616 cristata, Limopsis 514, 559, 572, 599 Ostrea 559, 568, 606 Tellidora 522, 554, 559, 568, 570, 572, 583-584, 616 cristella, Chama 388-391, 410 Pseudochama 390-392, 403, 407, 412 crocata, Avicula 491, 560, 611 croceus, Spondylus 579, 613 cruickshanki, Cypraeovula 143 Cryptocypraea 130-132 dillwyni 133, 135, 148 Cryptodon obesus 559, 617 pyriformis 559, 617 Cryptomphalus aspersus 73 Cryptopecten phrygium 518, 530, 533, 564, 572, 606, 608 Cryptostrea 606 permollis 309-310, 312-314, 316-317, 320, 321, 3229/2009 crystallina, Tellina 568, 587, 615 Ctena orbiculata 302, 417-418, 419, 420-424, 422, 515, 530-535, 548, 553, 572-573, 600, 622 orbiculata forma recurvata 550 pectinella 423, 515, 600, 622 Ctenoides floridana 587 floridanus 557, 572, 578, 598-599 miamiensis 514, 573, 598 mitis 533, 573, 598 planulatatus 572, 598 planulatus 514, 573, 598 sanctipauli 514, 548, 572-573, 581, 598 scaber 514, 572-573, 577, 598-599 scabra 563 Ctenopelta porifera 175, 176, 181 cubana, Codakia 622 cubaniana, Corbula 559, 595 Cytherea (Transennella) 559, 567, 621 Gouldia 558 Juliacorbula 539 Transenella 562, 566, 568, 573, 621 Transennella 525, 550, 563, 565, 570, 574, 586, 621 cubensis, Laemodonta 328, 573 cubitus, Mytilus 571, 603 Cucullaearca 355, 378, 591 candida 510, 533, 591 cucullata, Saccostrea 316 Cucurbitula 50 cueniformis, Gastrochaena 552, 596 cuericiensis, Helicina punctisulcata 217, 219 culebrana, Transenella 573 Transennella 525, 550, 621 Cumingia 583, 612 antillarum 563, 576, 586, 611 coarctata 520, 549, 561, 567, 570, 572, 574, 579-580, 584, 611 tellinoidea 554-555, 587, 612 tellinoides 555, 559, 567, 579, 583-585, 611-612 tellinoides vanhyningi 548, 572, 577, 581, 612 vanhyningi 520, 531, 534-535, 550, 612 cumingianus, Solecurtus 520, 572, 612 cumingii, Cribrarula 128, 128, 138, 140, 154 Cummingia antillarum 562, 611 cuneata, Grateloupea 440 cuneatus, Donax 459 cuneiformis, Gastrochaena 559, 567, 596 Martesia 519, 559, 567, 572, 579, 582, 609 cuneimeris, Animalocardia 567 Anomalocardia 565, 568, 570, 577, 583, 588, 618 cuneipyga, Abrina 157 cuneiveis, Anomalocardia 578, 618 Cuspidaria 597 (Cardiomya) 595 (Cardiomya) costellata 560, 567 (Cardiomya) perrostrata 559 (Cardiomya) striata 559 (Cuspidaria) 595 (Cuspidaria) obesa 559 gigantea 572, 595 (Liomya) granulata 559, 595 (Liomya) granulata var. velvetina 559, INDEX 639 595-596 obesa 513, 595 (Plectodon) granulata 559, 595 (Plectodon) granulata var. velvetina 559, 595-596 rostrata 513, 551, 558, 572, 595 Cuspidariidae 13, 512, 581, 595 Cuverias 587 Cyathermia naticoides 175, 175 Cyclinella tenuis 524, 567, 570, 572, 619 Cyclininae 427 Cyclocardia borealis 581, 622 Cyclopecten 572, 610 nanus 519, 610 strigillatus 519, 610 thalassinus 539 cygnea, Anodonta 208 cylindrica, Erronea 142 Erronea cylindrica 141, 155 Cymatioa 548, 572, 574, 596 Cymatoica hendersoni 521 orientalis 521 orientalis hendersoni 572, 577 orientalis forma hendersoni 614 cymella, Caryocorbula 512, 595 Corbula 547-548, 550, 554, 558-559, 567, 575, 579-580 Corbula (Caryocorbula) 566, 577 Cyphoma gibbosum 132-133, 147, 242 Cypraea 127, 131 pantherina 133, 135, 150 tigris 133, 135, 150 Cypraeidae 127, 131-133 Cypraeinae 131-132, 134 Cypraeovula 130-131, 137, 139 alfredensis 137-138, 153 algoensis 137-139, 153 capensis 137-138, 152 castanea 138-139, 152 colligata 143 connelli 138, 152 coronata 138, 152 cruickshanki 143 edentula 137-138, 153 fuscodentata 137-138, 152 fuscorubra 137-138, 152 immelmani 143 iutsui 138-139, 152 mikeharti 137-139, 143, 153 Cypraeovulinae 130-131, 137 Cypricardia 618 coralliophaga 579 Cyrena carolinensis 558, 594 caroliniana 594 floridana 587, 594 640 (Leptosiphon) carolinensis 559, 594 (Pseudocyrena) floridana 561, 594 Cyrenoida floridana 513, 554, 559, 572, 576, 579, 5853, 596 Cyrenoidea 596 Cyrenoididae 513, 596 Cyrtopleura costata 519, 570, 572, 609 Cytherea 429, 559, 618, 620-621 albida 559, 567, 620 (Callista) 620 (Callista) gigantea 559, 620 (Callista) maculata 559 circinata 579, 620 conradina 579 convexa 579, 622 (Cytherea) 619-620 (Cytherea) listeri 429, 561 (Cytherea) rugatina 561 dione 556, 579, 620 (Dione) 620 (Dione) dione 559, 567, 620 (Dione) hebraea 620 hebraea 558, 560, 567, 579, 587, 620 listeri 429, 441 simpsoni 559, 567, 620 (Tivela) 621 (Tivela) mactroides 559, 567, 621 (Transennella) 621 (Transennella) conradiana 567, 621 (Transennella) conradina 559, 621 (Transennella) cubaniana 559, 567, 621 (Trigona) incerta 579, 621 (Veneriglossa) 622 (Veneriglossa) vesica 559 (Ventricola) 619 (Ventricola) rigida 561 (Ventricola) strigillina 554, 561, 619 (Ventricola) strigillinus 548 dacostae, Trishoplita 89, 94, 98, 100, 112 Dacrydium elegantulum hendersoni 516, 099 272, 516, 603 vitreum 559, 567, 578, 603 dalli, Propeamussium 610 damaecornis, Chama 396 Dasycladus 497 dayritiana, Eclogavena 141, 156 decipiens, Polodesmus 568, 589 Zoila 135-136, 150 declivis, Abrina 157, 162, 166 Notocypraea 138, 152 decora, Tellina 556, 559, 568, 579, 615 decussata, Crenella 516, 572, 603 Glycymeris 513, 570, 572, 576, 582, 585, 597 INDEX deflorata, Asaphis 249-255, 252-253, 256-261, 258-262, 263-267, 264-265, 268-273, 519, 559, 562, 567, 569-570, 572, 5705. 579.610 deforgesi, Austrasiatica 143 delavayana, Aegistohadra 87, 92, 112—113, 114-117, 124 Nanina 79, 112, 114 demissus, Modiolaria 567, 603 Dendostrea 606 folium 316-317 frons 310, 316-317, 517, 530-531, 538: 553, 566, 500, 972, 578,605 dendriticum, Amygdalum 602 denselamellosa, Ostrea 316-317 dentata, Divaricella 515, 570, 572, 600 Divaricella (Divaricella) 556 Lucina 577 Lucina (Divaricella) 567 denticulata, Donax 566-567, 579, 596 denticulatus, Donax 559, 579-580, 596 dentifera, Lamellolucina 291 Dentiovula 132 takeoi 132-133, 147 depressa, depressa Mauritia 135-136, 150 depriesteri, Bistolida kien 138, 141 Bistolida kieneri 154 Deroceras (Deroceras) altaicum 125 derosa, Erronea caurica 141 Erronea cf. caurica 142, 155 deshayesii, Arca 579, 589 diagues, Bistolida stolida 138-139, 141, 153 Dianadema 37, 50, 53 multangularis 42, 45, 48, 50 dichotoma, Asaphis 249-250, 571, 610 dichotomum, Aspergillum 50 diductus, Oxychilus 32-33 dietziana, Caryocorbula 512, 595 Corbula 558-559, 567, 572, 595 dilatata, Crepidula 198-199, 201 dillwyni, Cryptocypraea 133, 135, 148 diluculum, Palmadusta 138-139, 153 Dinocardium 580-581, 592 robustum 511, 570, 592 dione, Cytherea 556, 579, 620 (Dione) dione 559, 567, 620 Pitar 524, 620 Diplodonta 583-585, 618 candeana 579 (Diplodonta) punctata 571, 596 notata 523, 618 nucleiformis 523, 618 (Phlyctiderma) 618 pilula 618 punctata 523, 570, 572, 583, 585-586, 618 semiaspera 559, 570, 572, 579, 584 simiaspera 576 soror 559-560, 579 subglobosa 559, 618 diploura, Retiskenea 174, 174, 179 directus, Ensis 560-561, 571, 579-580, 609 Discus rotundatus 215 discus, Dosinia 524, 552, 567, 570, 572, 584-585, 587, 619 dislocates, Pecten 579, 606 dislocatus, Pecten 558, 606 disparilis, Corbula 558, 567, 595 Varicorbula 512, 573, 595 dispersa, Mauritia depressa 135-136, 150 distans, Palmadusta contaminata 138, 153 distorta, Thracia 523,617 Divalinga quadrisulcata 515, 533, 572, 600, 622 divaricata, Crenella 559, 567, 570, 583-585, 603 Petricola 558, 579, 609 Divaricella 600 dentata 515, 570, 572, 600 (Divalinga) 600 (Divaricella) 600 (Divaricella) dentata 556 (Divaricella) quadrisulcata 556 quadrisulcata 292, 570, 583 Divarilima albicoma 514, 562, 572, 574, 598 divisus, Tagelus 264, 520, 559-560, 567. 568. 570, 572, 583, 585, 612 Dolicheulota 83 dombeii, Tagelus 264 domingensis, Acar 328, 510, 533, 574, 579, 589 Arca 579, 589 Barbatia 562-563, 569, 572, 577, 579, 585-586, 591 Barbatia (Acar) 571 Brachidontes 569, 572, 602 Brachiodontes 603 Cardita 559, 576, 593 dominguensis, Arca (Barbatia) 558, 589 Barbatia 355, 363, 364, 368-369, 372, 374, 375, 377-378 Barbatia (Acar) 363 Glans 511, 570, 572, 577, 584-585, 593 dominica, Antigona 432 Donacidae 247, 459, 513, 596 Donax 459-460, 462-463, 469, 553, 573, 596 cuneatus 459 INDEX 641 denticulata 566-567, 579, 596 denticulatus 559, 579-580, 596 dorotheae 459 fossor 459-461, 462-464, 466, 467-470, 559, 561, 567, 571, 579-580, 596 fossor protractus 567 gouldii 469-470 parvula 459 protracta 459 roemeri 459, 567, 596 roemeri protacta 459 roemeri protracta 459 roemeri roemeri 459 texasianus 459, 596 trunculus 208, 469 tumidus 567, 596 variabilis 459-461, 462-463, 467-470, 468. 913.592, 999) 567. 3725797596 venustus 469 vittatus 296 dondani, Serratovolva 133, 147 dorotheae, Donax 459 dorsalis, Erronea subviridis 141 Erronea (Adusta) subviridis 155 Dosina 440, 444 listeri 429, 440 veerrucosa 440 zelandica 444 Dosinia 443-444 concentrica 566-567, 579-580 discus 524, 552, 567, 570, 572, 584-585, 587,619 (Dosinidia) 619 (Dosinidia) concentrica 561, 619 (Dosinidia) elegans 561 elegans 524, 548, 552, 559, 561, 567, 570, 572,619 floridana 548, 557, 619 listeri 441 Dosiniinae 427, 444 draceana, Erronea caurica 141-142, 155 draconis, Pseudochama 387, 411 draparnaudi, Oxychilus 19-23, 30-34 Dreissena polymorpha 208 Dreissenidae 513, 596 duplicata, Modiola 587, 604 duplicatus, Polinices 296, 305 dupliniana, Crassinella 512, 572, 595 dysoni, Helicina 224 eburnea, Erosaria 133-134, 143, 149 echandiensis, Helicina 219 echinatus, Spondylus 568, 613 Echinochama 408 arcinella 408 642 arcinella cornuta 408 cornuta 408 Eclogavena 131, 140 coxeni 136, 141, 156 dayritiana 141, 156 luchuana 143 quadrimaculata quadrimaculata 141, 156 quadrimaculata thielei 141, 156 Ecphora 409 edentula, Cypraeovula 137-138, 153 Loripes 559, 571, 599 Loripes var. chrysostoma 559, 599 Lucina (Loripinus) 567, 599 Psammotreta (Tellinimactra) 167 edulis, Ostrea 316-317, 322 effluens, Pecten (Pecten) 560, 608 Едепа 469 radiata 469 Egeta protexta 567 eglantina, Mauritia 135, 150 egmontianum, Trachycardium 511, 956-557, 564, 570, 573, 578, 584, 592 Electroma 473 elegans, Asthenothaerus (Bushia) 559 Bushia 523, 585, 617 Dosinia 524, 548, 552, 559, 561, 567, 910; 572 619 Dosinia (Dosinidia) 561 elegantissima, Euciroa 525, 552, 621 Verticordia 622 elevata, Chione 295-299, 298-300, 301-305, 303-304, 523, 527, 530-535, 57.3, 577, 613 elevatus, Lepetodrilus 169, 173 elliptica, Basterotia 521, 569, 572, 613 Laternula 424 Elliptotellina 616 americana 521, 614 elongata, Erronea caurica 141-142, 155 Poromya 610 eludens, Zoila 135, 150 emmonsi, Chama 387-388, 405, 411 encymata, Pitar 568, 618 englerti, Erosaria 133-134, 148 Ennucula aegeensis 517, 605 tenuis 517, 538, 572, 605 Ensis americana 559, 609 directus 560-561, 571, 579-580, 609 minor 519, 572, 609 Ensitellops protexta 521, 613 entochilus, Stilpnodiscus 86, 97, 112 Entodesma beana 515, 570, 572, 602 Entoliidae 513, 596 Eocypraeinae 132 Eopsuma 385 INDEX ephippium, Perna 558-559, 579, 597 equestris, Ostrea 556, 570, 588-589, 606 Ostreola 309-314, 317-319, 318-319, 320, 327, 322-323, 517, 566, 570; 572 Eratoidae 132 erosa, Chione 296 Erosaria 133, 149 Erosaria 130-132, 134 acicularis 133, 149 albuginosa 133-134, 148 beckii 133-134, 148 boivinii 133, 149 cernica 133, 149 citrina 133-134, 148 eburnea 133-134, 143, 149 englerti 133-134, 148 erosa 133, 149 erosa cf. chlorizans 149 gangranosa 133, 149 helvola 134 helvola callista 134 helvola cf. callista 133-134, 148 helvola hawaliensis 133-134, 148 helvola helvola 133-134, 148 irrorata 133-134, 148 kingae 133, 149 labrolineata 133, 149 lamarckii lamarckii 133-134, 149 lamarckii cf. redimita 133-134, 149 тасапагем 133-134, 148 marginalis 133-134, 148 miliaris 133-134, 143, 149 nebrites 133, 149 ocellata 133, 149 ostergaardi 143 poraria 133-134, 148 spurca 133, 149 thomasi 133, 149 turdus 133-134, 148 Erosariinae 130-131, 134 Erronea 127, 130-131, 140, 142-143 adusta 141 (Adusta) adusta 155 (Adusta) onyx 155 (Adusta) onyx melanesiae 155 (Adusta) subviridis dorsalis 155 (Adusta) subviridis subviridis 155 angioyorum 143 barclayi 142 caurica 141-142, 155 caurica caurica 141-142, 155 caurica derosa 141 caurica cf. derosa 142, 155 caurica draceana 141-142, 155 caurica elongata 141-142, 155 caurica palauensis 141 caurica quinquefasciata 141-142, 155 caurica samoensis 141-142, 155 cylindrica 142 cylindrica cylindrica 141, 155 cylindrica lenella 141-142, 155 errones 141-142, 155 fernandoi 141—142, 155 hungerfordi 142 nymphae 143 onyx 141 onyx melanesiae 141 ovum 142 ovum chrysostoma 141, 155 ovum оуит 141, 155 ovum palauensis 142, 155 pallida 141-142, 155 pulchella 142 pyriformis 141-142, 155 rabaulensis 141-142, 155 subviridis dorsalis 141 subviridis subviridis 141 vredenburgi 141-142, 155 xanthodon 141-142, 155 Erroneinae 131, 137, 139 errones, Erronea 141-142, 155 erubescens, Leptaxis 73, 75-76, 75-77 Ervilia 585, 612 castanea 258, 296 concentrica 520, 559-560, 562, 566-567, 570, 572, 579-580, 584-585, 612 nitens 520, 559-560, 562, 566-567, 570, 572, 579-580, 585, 612 subcancellata 520, 562, 572, 612 Erycinidae 15 erythraeensis, Bistolida 138-139, 141, 153 escondida, Helicina 217, 220, 222-223 esontropia, Cribrarula 154 Cribrarula esontropia 138, 140 Euciroa 622 elegantissima 525, 552, 621 Eucrassatella floridana 580, 595 speciosa 512, 533, 570, 572, 576, 595 eucymata, Callista 523, 530, 533, 572, 618 Costacallista 576 Eueuhadra 79, 81, 85, 89, 103, 106-108, 119 gonggashanensis 79, 88, 112, 114, 118—120, 119, 121-125 Eufistulana 50 Euhadra 81, 106-109, 114, 119 herklotsi 89, 94, 98, 101, 112 Euhadrinae 107, 108 Eulepetopsis vitrea 170-171, 171, 180 Eurytellina 616 INDEX 643 alternata 521, 614 angulosa 521, 614 lineata 521, 614 nitens 521, 614 punicea 521, 614 Eutrochatella pulchella 217, 224 Euvola 606, 608 chazaliei 518, 549, 572, 608 laurentii 518, 608 cf. papyracea 518, 606, 608 raveneli 518, 572, 576, 608 ziczac 518, 572-573, 577, 608 exasperatus, Aequipecten 606 Aequipecten acanthodes 608 Lindapecten 572, 608 Pecten 568, 608 Pecten (Pecten) 559, 608 excelsa, Bathyxylophila 176 exilis, Tellina (Scissula) 561, 615 exmouthensis, Cribrarula 154 Cribrarula exmouthensis 138, 140 exogyra, Pseudochama 391 exquisita, Pseudocypraea 132-133, 147 extenuata, Macoma 522, 576, 615 exusta, Talparia 133, 135-136, 151 exustas, Mytilus 579 exustus, Brachidontes 516, 527, 530-532, 534—535, 551, 555, 565, 569-570, 572, 577, 583-584, 588-589, 602-603 Brachiodonta 575 Mytilus 558-559, 567, 578-579 Mytilus (Brachidontes) 563, 580 fabula, Melicerona felina 143 fallax, Cribrarula 138, 154 Falsimargarita 180 fasciola, Cathaica (Cathaica) 92, 102, 111 fausta, Arcopagia 521, 530-531, 533, 535, 5511553006, 563: Ol 583, 588,614 Масота 579, 614 Tellina 556, 559, 562-564, 567-568, 570-572, 582, 586, 588, 614, 616 Tellina (Acropagia) 580 Tellina (Arcopagia) 553 Tellina (Cyclotellina) 580 fecunda, Crepidula 198-199, 201 Felaniella candeana 523, 618 felina, Melicerona 141, 143, 156 Melicerona felina 143 fernandoi, Erronea 141-142, 155 ferruginea, Chama 401-402 Chama radians 402 Pseudochama 386, 401 ferrugivora, Paralepetopsis 170-171, 171, 180 644 festivus, Nassarius 295 filiata, Codakia (Jagonia) orbiculata 566, 600 Lucina (Jagonia) orbiculata 567, 600 filiforme, Syringodeum 474, 487 filippina, Laeocathaica (Laeocathaica) 98 filosa, Lucina (Lucina) 559 Lucinoma 515, 530, 533, 572, 601 filosum, Lucinoma 572 filosus, Phacoides (Lucinoma) 556, 601 fimbiata, Xylotrya 556, 617 fimbriata, Purpuradusta 142 Purpuradusta fimbriata 138, 141, 156 Xylotrya 559, 568, 617 fimbriatula, Bankia 523,617 firma, Chama sinuosa 398, 399, 553, 594 fischeriana, Haliris 525, 572, 621 Venus (Haliris) 560 Verticordia 622 Verticordia (Haliris) 621 Fissurellidae 169, 180 flexuosa, Anomalocardia 558, 618 Rangia 553, 572, 594 Strigilla 522, 556, 559, 568, 580, 616 flindersi, Lepsiella 296 florida, Adamantia 133, 147 Chama 382-385, 388-389, 392, 396, 397, 398, 403, 405, 410-411, 511, 551, 559#990, 902101207093 floridana, Anadara 510, 571, 586, 589 Anadara lienosa 570 Arca lienosa 589 Arca (Anadara) var. secernenda 589 Cardita 552, 559,565, 567, 576, 579, 583 Cardita (Carditamera) 561, 582 Cardita (Mytilicardia) 571, 593 Carditamera 511, 530-535, 570, 572, 575,583, 593 Crassatella 559-560, 595 Ctenoides 587 Cyrena 587, 594 Cyrena (Pseudocyrena) 561, 594 Cyrenoida 513, 554, 559, 572, 576, 579, 583, 596 Dosinia 548, 557, 619 Eucrassatella 580, 595 Lucina 55: 5725771602 Lucina (Lucina) 559 Lyonsia 515, 572, 602 Lyonsia hyalina 565, 584-585, 602 Naeromya 584 Neaeromya 585 Orobitella 513, 572, 596 Polycyrena 551, 594 INDEX Polymesoda 565, 568, 580, 594 Pseudocyrena 549-550, 563, 576, 578, 594 Pseudomiltha 601 Stewartia 515, 602 Tivela 525, 538, 573, 621 floridanus, Ctenoides 557, 572, 578, 598-599 Megaxinus 556, 602 floridensis, Condylocardia 548, 550, 575, 594 Ostrea 552, 605 fluminea, Corbicula 48 Foegia 37-39, 53 novaezelandiae 37—43, 39-51, 45, 47—48, 50, 52 foliacea, Chama 392, 394 Tellina 263 foliata, Ostrea 587, 605 folium, Dendostrea 316-317 Ostrea 605 forcartianus, Oxychilus (Ortizius) 21 forficatus, Lithodomus 579, 603-604 Lithophagus 559, 604 fossor, Donax 459-461, 462-464, 466, 467-470, 559, 561, 567, 571, 579-580, 596 fragilis, Lima 587, 599 Mactra 567, 570, 578, 583 Mactrotoma 515, 572, 602 Sphenia 516, 602 fragosus, Nodipecten 518, 527-528, 530, 533, 579, 608 Pecten nodosus 568, 608 francescoi, Cribrarula esontropia 138, 140, 154 fretterae, Neomphalus 169, 181 friendii, Zoila 136 Zoila friendii 150 frons, Dendostrea 310, 316-317, 517, 530-531, 533, 553, 566, 570; 572; 576, 605 Lopha 586 Ostrea 559, 568, 570-571, 575, 577, 586 Ostrea (Lopha) 586 Fruticicola 81, 85, 106-108 fruticum 82, 86, 111 fruticum, Fruticicola 82, 86, 111 Fugleria 355, 372, 378, 591 pseudoillota 355 tenera 510, 548, 591 fulminata, Pitar 568, 583, 587, 620 Pitar cf. 565, 568, 620 fulminatus, Pitar 524, 531, 535, 570, 572, 584—585, 620, 622 fultoni, Вагусургаеа 132, 136, 150 funcki, Helicina 217-218, 219, 221 Fundella candeana 552, 602 fusca, Arca 579, 590 Botula 339-341, 341, 353, 516, 552, 556, 562-563, 567, 569, 572, 576, 578, 584, 602 fuscodentata, Cypraeovula 137-138, 152 fuscorubra, Cypraeovula 137-138, 152 gabbi, Strigilla 522, 554, 570, 572, 574, 616 Strigilla (Strigilla) 550, 577 gaederopus, Spondylus 556, 613 Gafrarium 443-444 Galeommatoidea 513, 596 gangranosa, Erosaria 133, 149 gansuica, Cathaica (Pliocathaica) 89, 93, UA garciai, Cribrarula 128, 128, 138, 140, 143, 154 gardnerae, Chama 405 Gari 258-259, 261, 263, 610 circe 519, 538, 610 solida 258-260, 262, 272 tellinella 258 vespertina 262 gaskoini, Cribrarula 128, 128, 138, 154 gaspardi, Cribrarula 138, 154 Gastrochaena 339, 352, 596-597 cueniformis 592, 596 cuneiformis 559, 567, 596 (Gastrochaena) 596 (Gastrochaena) hians 557 hians 339, 351-352, 352, 513, 557, 562, 570, 572, 584-585, 596 оу=:352: 513, 552, 559, 567, 570, 572, 596 (Rocellaria) ovata 557 rostrata 552, 567 (Spengleria) 597 (Spengleria) rostrata 559 Gastrochaenidae 247, 278, 339, 351, 513, 596 Gemma gemma 305, 524, 619 purpurea 567, 619 gemma, Gemma 305, 524, 619 Helicina 217, 220, 222 Gemminae 427 geographica, Leporicypraea 131, 149 georgiana, Tellina 568, 614 gerlachi, Aegista (Plectotropis) 89, 95, 111 Geukensia granosissima 516, 572, 603 gibbesii, Crassatellites 595 gibbosa, Plicatula 519, 530, 532-533, 568, 570, 572, 583, 586, 610 INDEX 645 gibbosum, Cyphoma 132-133, 147, 242 gibbsii, Crassatellites 567, 595 gibbus, Aequipecten 570 Argopecten 518, 530, 533, 563, 567, 569, 572, 584-586, 606 Pecten 568 Pecten (Chlamys) var. nucleus 560, 607 Pecten (Plagioctenium) var. amplicostatus 548 Tagelus 552, 612 gigantea, Callista (Dione) 571, 620 Cytherea (Callista) 559, 620 Cuspidaria 572, 595 Macrocallista (Callista) 567, 620 gigas, Crassostrea 316 Strombus 242, 576 gilvella, Luria 143 glabra, Hyalogyrina 176, 179 glacialis, Pandora 517, 539, 606 Pandora (Kennerlia) 560 Glans dominguensis 511, 570, 572, 577, 584-585, 593 Globivenus 427, 619, 621 rigida 441, 524, 572, 619 rugatina 441, 524, 572, 619 toreuma 428, 443 globula, Astarte 510, 591 globulus, Pustularia globulus 135, 138, 151 glomerula, Arca 552 Arca (Byssoarca) 559 Bathyarca 510, 572, 591 Glossidae 12 Glycimeris 997 americana 587 americana lineata 585, 597 pectinata 585-586 pectinatus 597 Glycimerus pectinatus 569 Glycymerididae 513, 584-585, 597 Glycymeris 581,584, 586, 597 americana 513, 552, 567, 570, 572, 597 decussata 513, 570, 572, 576, 582, 585, 597 lineata 567, 597 pectinata 556, 563-564, 567, 570, 572, 576, 578, 581-584, 586, 588, 597 pectinatus 567, 579-581, 597 pennacea 567, 597 spectralis 513, 597 undata 513, 570, 572, 576, 597 glypta, Cardiomya 512, 572, 595 glyptus, Aequipecten 517, 571, 577, 606 gonggashanensis, Eueuhadra 79, 88, 112, 114, 118-120, 119, 121-125 goodallii, Bistolida 138, 141, 153 646 gordensis, Neolepetopsis cf. 170, 170-171, 180 gouldi, Tellina 568, 613 Gouldia 444, 621 cerina 524, 533, 567, 570, 572, 584-585, 619 cubaniana, 558 mactracea 567, 595 parva 566-567, 595 gouldii, Acorylus 521, 613 Donax 469-470 Tellina 559, 570, 572-573, 579, 585, 613, 616 Tellina (Acorylus) 553, 613 Gouldiinae 427, 444 Gracilaria 499 gracilis, Purpuradusta gracilis 138, 141, 156 gracillima, Amphipholis 564 gradata, Arca 355, 579, 589 Arca (Barbatia) 558, 589 granatina, Basterotia quadrata 567, 613 grandis, Anodonta 208 Thyasira 523, 539, 617 Granicorium 277 granosa, Anadara 295 granosissima, Geukensia 516, 572, 603 granosissimus, Modiolaria demissus 567, 603 granulata, Acanthopleura 563 Chione (Timoclea) 566, 579-580, 618-619 Cuspidaria (Liomya) 559, 595 Cuspidaria (Liomya) var. velvetina 559, 595-596 Cuspidaria (Plectodon) 559, 595 Cuspidaria (Plectodon) var. velvetina 559, 595-596 Leiomya 551 Leiomya (Plectodon) granulata 566, 596 Nucleolaria 133, 135, 143, 148 Poromya 519, 558, 560-561, 568, 572, 610 Poromya granulata 566, 610 Protothaca 572, 619 Venus 560, 579, 618 granulatus, Brachidontes 292 Plectodon 513, 550, 572, 574, 595 Venus (Chione) 568, 619 Grateloupea cuneata 440 grayana, Mauritia 135, 150 Gregariella coralliophaga 516, 570, 572, 603 gruneri, Lutricola 579, 615 grus, Chione 551, 563-564, 570, 584-585, 619 INDEX Chione (Timoclea) 549, 551 Timoclea 525, 573, 621 Venus (Chione) 568 Gryphaeidae 247, 309-310, 312, 316, 513, 597 gryphina, Chama 403 gryphoides, Chama 386, 395 guadalupensis, Avicula 491 guidoni, Hyalinia 20 guppyi, Americardia 511, 549-550, 569, 591 gussoni, Spondylus 570, 572, 613 guttata, Perisserosa 133, 148 haemastoma, Thais 296 hainanensis, Abrina 157, 164, 167 Halimeda 245-247, 278, 452—454, 457, 486 Haliotidae 180 Haliotis 410 Haliris 622 fischeriana 525, 572, 621 Halodule 245-246, 278, 418, 454, 498—499 Halolimnohelix 84 Halophila 418 hamatus, Mytilus 559, 603 hammondae, Purpuradusta 138, 141, 156 Haplohelix 84 hartsmithi, Notocypraea 137-138, 152 hawaiiensis, Erosaria helvola 133-134, 148 hebraea, Cytherea 558, 560, 567, 579, 587, 620 Cytherea (Dione) 620 heilprini, Chama 387, 398, 410—411 heliacus, Aequipecten 517, 606 Pecten 568 Helicidae 79, 84, 99, 112, 217 Helicina 217 amoena 224 beatrix 217 beatrix beatrix 220, 222 beatrix confusa 220-221 beatrix riopejensis 220-221 chiquitica 217-218, 220, 223 dysoni 224 echandiensis 219 escondida 217, 220, 222-223 funcki 217-218, 219, 221 gemma 217, 220, 222 monteverdensis 217-218, 220, 222 neritella 224 orbiculata 224 pitalensis 217, 219, 221 platychila 224 punctisulcata cuericiensis 217, 219 зепсеа 224 talamancensis 217-218, 220, 222 tenuis 217-218, 219, 221 turbinata 224 Helicinidae 217-218 Helicoidea 73, 79 Helicostylidae 79, 107, 108-109 Helicostylinae 79, 107, 108-109 Helix 73, 81, 85, 106-107, 211, 215 aspersa 73, 211-212, 215-216 pomatia 85, 91, 92, 99, 112, 212, 215-216 seraphinica 114, 118, 118-119 Helminthoglypidae 84, 109 helvetica, Hyalina (Polita) 33 helvola, Erosaria 134 Erosaria helvola 133-134, 148 Hemicardium medium 571, 591 hemicyclica, Pecten 579, 608 Pecten (Janira) 560, 608 hemicyclicus, Pecten 608 Hemimetis (Florimetis) 615 intastriata 574 hemphilli, Asthenothaerus 523, 559, 567, 572, 584-585, 617 hendersoni, Cymatoica 521 Cymatoica orientalis 572, 577 Dacrydium elegantulum 516, 533, 572, 578, 603 herklotsi, Euhadra 89, 94, 98, 101, 112 hesitata, Umbilia 133, 135, 149 hessleri, Alviniconcha 178, 178 Heterodonax bimaculata 559, 567, 610 Heterodonax bimaculatus 258-259, 261, 263 272, 519, 572, 579, 610 hians, Gastrochaena 339, 351-352, 352, 513, 557, 562, 570, 572, 584-585, 596 Gastrochaena (Gastrochaena) 557 Lima 559, 567, 599 Rocellaria 551 Hiatella arctica 514, 572, 584-585, 597 azaria 514, 597 Hiatellidae 12, 514, 597 hiatus, Paphridea 592 Рарупаеа 557 Hinnites 613 hirasei, Austrasiatica 132, 138, 140-141, 155 Nesiocypraea 130 hirsuta, Trichomya 292 hirundo, Bistolida 138, 141, 153 Pteria 473, 491 histrio, Mauritia 135, 150 hojarasca, Alcadia 217, 220, 223 hollandi, Alcadia 224 INDEX 647 hornbeckiana, Coralliophaga 579, 609 hornbeckii, Pholas 609 hortensis, Cepaea 73 Humboldtiana 73 Humphreyia 37 strangei 42, 48, 50 humphreysii, Palmadusta 143, 153 hungerfordi, Erronea 142 Notadusta 141, 156 Hyalaeas 587 Hyalina scotophila 20 (Polita) helvetica 33 scotophila var. dilatata 21 hyalina, Lyonsia 42 Hyalinia blauneri var. cloacarum 21 guidoni 20 isseliana 20-21 meridionalis 20-21 nitidula var. amiatae 21 paulucciae 20-21, 31 scotophila var. notha 20 sylvicola 21 Hyalogyrina glabra 176, 179 umbellifera 179, 179 Hyalogyrinidae 176, 179, 181 Hydrobia 273 Hygromia 73-74, 77 Hygromiidae 73, 77 hyotis, Hyotissa 309, 312, 315-316, 320 Hyotissa 309-310, 320 hyotis 309, 312, 315-316, 320 mcgintyi 513, 566, 597 Hypophthalmichthys molitrix 205 ictericus, Spondylus 520, 530, 533, 562, 564, 570, 572, 576, 588, 613 imbricata, Arca 328, 378, 510, 530-535, 551, 562, 565, 567, 569-572, 574, 577, 579, 585-586, 588, 590 Arca (Arca) 559 Caribachlamys 518, 572, 586, 607 Chama 395-396 Chlamys 551, 562, 570, 578, 588, 607 Parahyotissa (Parahyotissa) 320 Pinctada 316, 473-474, 489, 489-493, 520, 530-531, 533, 556, 564-565, 570, 572-574, 578, 584, 587-588, 611 imbricatus, Chlamys 570, 576, 587 Pecten 553, 568-569, 579, 585, 587, 607 Pecten (Chlamys) 566, 580-581, 607 Pecten (Chlamys) var. mildredae 607 Pecten (Pecten) 559, 607 immanis, Mauritia arabica 135, 150 immelmani, Cypraeovula 143 impressa, Anomalocardia 571, 618 648 inanis, Abrina 162, 166 inaequalis, Bathyarca 510, 591 inaequivalvis, Scapharca 571, 591 incerta, Cytherea (Trigona) 579, 621 incongrua, Arca 567, 579, 591 Arca (Scapharca) 559, 591 Macoma 167 inconstans, Xenostrobus 296 indica, Mauritia scurra 135, 149 inezae, Chama 382-386, 388, 405-406, 408, 410-412, 511, 549, 556, 593 Pseudochama 381, 405, 411, 549-550, 553, 572, 575, 580, 594 inflata, Lima 558-559, 567, 599 Pandora 517, 530, 533, 554, 572, 606 intapurpurea, Chione 551, 563, 570, 579-580, 619 Chione (Chione) 561 Puberella 524, 533, 572, 621 intasriata, Apolymetis 567, 615 intastriata, Hemimetis (Florimetis) 574 Leporimetis 521, 570, 572, 615 Metis 561 Psammotreta 586, 615 interrupta, Blasicrura 141, 156 Pseudochama 381 Tellina 559, 568, 571, 579, 617 interstincta, Staphylaea limacina 133, 135, 148 interstriata, Lutricola 559, 615 involuta, Chama 387, 411 Iphigenia 469 brasiliana 513, 567, 570, 572, 596 braziliana 559, 596 Ipsa 130-132, 134 childreni 130, 133, 148 iridescens, Ostrea 320 iris, Scissula 522, 615-616 Tellina 302, 556, 568, 573, 584-585, 616 Tellina (Angulus) 566, 580 Tellina (Scissula) 561, 566, 580 irradians, Aequipecten 570 Argopecten 518, 531, 534, 548, 550, 555, 562, 565, 569,572, 983, 606 Pecten (Pecten) var. dislocatus 559, 606 irrorata, Erosaria 133-134, 148 Irus 444 isabella, Luria 133, 135, 137, 151 isabellamexicana, Luria 133, 135, 151 Ischadium recurvum 516, 572, 603 isocardia, Cardium 559, 567, 592 Trachycardium 552, 592 isocardium, Cardium 574, 592 Isognomen 597 alatus 557, 566 Isognomon 316, 328, 574, 586, 597-598 alata 576, 597 alatus 316, 501, 514, 535, 556, 562, 564-565, 570, 572, 578, 586, 588-589, 597,611 bicolor 356, 377, 514, 533, 549-551, 562-563, 565, 570, 572, 574, 576-577, 586, 588-589, 598 chemnitziana 580, 598 listeri 576, 598 (Pedalion) alata 580, 597 radiatus 514, 551, 562, 565, 570-572, 578, 588, 598 Isognomonidae 514, 597 Isorropodon 13 isseliana, Hyalinia 20-21 iudicai, Chama 389, 395, 411 iutsui, Cypraeovula 138-139, 152 Jagonia orbiculata var. filiata 548, 561, 600 orbiculata var. recurvata 548, 561, 600 jamaicensis, Alcadia 224 Leda 558 Lucina 571, 601 Lucina (Anodontia) 567, 601 Nuculana 517, 605 janae, Talostolida teres 139 Janthina janthina 577 janthina, Janthina 577 Janthinas 587 japonica, Kellia 58, 63, 64-66, 71 jeaniana, Zoila friendii 135 Zoila jeaniana 150 Jenneria 132 pustulata 133, 147 johnsonorum, Palmadusta 143 joycae, Lyncina 143 Juliacorbula aequivalvis 512, 595 cubaniana 539 kallinubilosus, Lyropecten 518, 572, 608 kanamarui, Abra 158 Karaftohelix 81, 106-109 weyrichii 84, 86, 94, 111 Katelysia 444 katsuae, Palmulacypraea 138, 141, 155 Pamulacypraea 140 keelingensis, Pustularia bistrinotata 135, 137-138, 152 keenae, Callucina 515, 599, 601 Kellia 57-59, 63, 64, 66-67 comandorica 57-59, 60-62, 62, 64-66, 71 japonica 58, 63, 64-66, 71 kussakini 57, 64-67, 68-70, 70-71 laperousii 66-67 porculus 58, 63, 64-66, 71 rubra 566, 596 suborbicularis 57-59, 63, 64-67, 71, 513, 572, 596 subrotundata 58, 63, 64-66, 71 Kelliella 1 Kelliellidae 1-2, 12-15 Kelliidae 57 ketyana, Zoila marginata 135-136, 150 kiangsinensis, Mastigeulota 92, 111 kieneri, Bistolida 139, 141 Bistolida kieneri 138 kingae, Erosaria 133, 149 kinoshitai, Abrina 157, 162, 166-168 knoxi, Nototeredo 523, 572, 617 krebsiana, Corbula 559, 595 Varicorbula 512, 539, 595 krynickii, Xeropicta 74 Kuia vellicata 444 kuroharai, Lyncina 135-137, 151 kussakini, Kellia 57, 64-67, 68-70, 70-71 Labiosa canaliculata 559, 602 labrolineata, Erosaria 133, 149 lacerata, Venus 441 Laciolina 616 laevigata 521, 614 magna 521, 614 lactea, Striarca 327, 332, 334, 336 lacteus, Calpurnus 147 Procalpurnus 133 lactuca, Chama 382, 386, 388, 403, 404, 408, 411, 511, 556, 570-572, 593 laeta, Amphidesma 548 Laemodonta cubensis 328, 573 Laeocathaica 81, 106-107 (Laeocathaica) filippina 98 (Laeocathaica) subsimilis 86, 89, 112 Laevicardium 553-555, 583, 585, 588, 592 laevigatum 511, 530-535, 556-557, 563-564, 570-572, 578, 583, 585-586, 588, 592 mortoni 302, 511, 530-534, 552-553, 555, 557, 564-565, 568, 570, 572-573, 575, 578, 583-584, 592 pictum 511, 572, 592 serratum 553, 571, 579, 592 sybariticum 511, 570, 572, 585, 592 Laevichlamys 607 multisquamata 518, 572, 608 laevigata, Laciolina 521,614 Staphylaea staphylaea 133, 135, 148 Tellina 552, 570, 573, 587-588, 616 Tellina (Laciolina) 553 INDEX 649 laevigatum, Cardium 558 Cardium (Laevicardium) 567, 592 Laevicardium 511, 530-535, 556-557, 563-564, 570-572, 578, 583, 585-586, 588, 592 Papyridea (Liocardium) 560, 592 laevis, Pachydermia 175, 175, 181 lama, Macoma 167 lamarckiana, Chama 398, 401 lamarckii, Erosaria lamarckii 133-134, 149 Venus 560, 618 Lamellaridae 132 lamellaris, Antigona 618 lamellifera, Myonera 513, 539, 559, 595 Lamellolucina 291 dentifera 291 gemma 291 lamellosa, Chama 392, 394 langfordi, Austrasiatica 132, 140, 154 Nesiocypraea 130 lanzai, Oxychilus 19, 21-22, 26, 29-31 Oxychilus (Ortizius) 19, 21, 31 laperousii, Kellia 66—67 lapicida, Petricola 339, 347-348, 350, 350-351, 353, 518, 552, 562, 566, 568, 570, 572, 580, 584, 588, 609 Petricola (Naranaio) 560 laqueata, Venus 441 Lasaea adansoni 513, 572, 596 rubra 5, 550 Lasaeidae 1, 59 Lasea rubra 579, 596 lata, Melina 581, 597 Papyridea 511, 587, 592 lateralis, Mactra 559 Modiolaria 559, 567, 581, 604 Mulinia 515, 567, 570, 602 Musculus 516, 570, 572, 576, 584-586, 604 Laternula 278 elliptica 424 truncata 48 latilirata, Chione 570, 578, 587,619 Lirophora 524, 572, 620 latiliratus, Venus (Chione) 568, 620 latior, Talostolida 138-139, 154 Latona 459 variabilis 459 Laurencia 486 laurenti, Amusium 563 laurentii, Amusium 571, 606 Euvola 518, 608 lavalleanus, Mytilus 579, 603 lazai, Pseudochama 387, 411 lazarus, Chama 389, 395-396, 410, 593 650 [еда 605 acuta 558 bushae 605 carpenteri 558 jamaicensis 558 (Leda) 604, 605 (Leda) acuta 559 (Leda) carpenteri 560, 605 (Leda) messanensis 559, 605 (Leda) vitrea 559 sublevis 605 (Yoldia) 622 (Yoldia) liorhina 559 Ledella solidula 605 sublevis 517, 605 Leiomya claviculata 595 granulata 551 (Plectodon) granulata granulata 566, 596 (Plectodon) granulata velvetina 566, 596 lenella, Erronea cylindrica 141-142, 155 lens, Astarte 559, 591 Loripes 559 Myrtea 601 Myrteopsis 515, 601 lenticula, Lucina 622 Lucina (Lucina) 559, 622 lentiginosa, Palmadusta 138, 153 Lepetodrilidae 173, 180 Lepetodriloidea 169 Lepetodrilus elevatus 169, 173 pustulosus 173, 173 Leporicypraea 131 admirabilis 136 geographica 135, 149 mappa 136 mappa admirabilis 135-136, 149 mappa aliwalensis 134-135, 149 mappa geographica 136 mappa mappa 135-136, 149 mappa panerythra 136 mappa rewa 136 mappa rosea 135-136, 149 mappa viridis 135-136, 149 valentia 135, 149 Leporimetis 615 intastriata 521, 570, 572, 615 Lepsiella flindersi 296 vinosa 296 Leptaxis 73-77, 74 erubescens 73, 75-76, 75-77 nivosa 73, 75, 75-76 undata 73, 75, 75-76 Lepton bowmani 579, 622 Leptonacea 564 leptonoidea, Macoma 613 INDEX leptonoides, Macoma 567, 613 leucocyma, Linga 570 Lucina 547, 572, 576-577, 600 Lucina (Lucina) 559 Pleurolucina 515, 530, 533, 601, 622 (Pleurolucina) Lucina 556 leucodon, Lyncina 135 Lyncina (Callistocypraea) 151 leucophaeata, Mytilopsis 538, 554, 572, 596 Mytilus 513 leviathan, Lyncina 135, 151 lewisi, Polinices 216 lewisii, Polinices 305 lienosa, Arca (Scapharca) 559, 589 lignea, Modiola (Amygdalum) 559, 603 Liguus 217 lima, Byssoarca 355 Lima 556, 563, 567, 569-570, 581, 586-588, 598-599 Lima 554, 564, 568, 578, 583, 585-586, 588, 598-599 albicoma 550, 559 caribaea 514, 530-531, 533, 572-573, 576, 598-599 fragilis 587, 599 hians 559, 567, 599 inflata 558-559, 567, 599 lima 556, 563, 567, 569-570, 581, 586-588, 598-599 (Limatula) 599 (Limatula) setifera 559 (Limatula) subauriculata 559 locklini 585 pellucida 551, 556, 562-563, 565, 567, 570, 578, 583-586, 588 scabra 551-552, 556, 559, 566-567, 570-571, 574, 576-579, 581, 586, 589, 598 scabra scabra 570, 598 scabra tenera 570, 574, 576, 578, 588, 598 scabra form tenera 556, 588, 598 squamosa 556, 559, 579, 598 tenera 559, 567, 576, 579, 587, 598-599 limacina, Staphylaea limacina 133, 135, 148 Limaria 583, 598-599 albicoma 577 (Limatulella) 581, 599 locklini 514, 599 pellucida 514, 531-532, 534, 555, 572, 583, 599 cf. pellucida 599 Limatula confusa 514, 567, 599 setifera 514, 539, 599 subauriculata 514, 539, 599 limatula, Myonera 595 Varicorbula 573, 595 Limax maximus 212, 214 Limea 599 bronniana 514, 539, 599 (Limea) 581, 599 Limidae 514, 585, 598 Limopsidae 581, 599 Limopsis 609 antillensis 559 aurita 514, 559, 572, 599 cristata 514, 559, 572, 599 minuta 514, 559, 572, 599 sulcata 514, 572, 599 tenella 599 limula, Macoma 522, 576, 615 Lindapecten 606, 608 exasperatus 572, 608 muscosus 518, 533, 572, 606, 608 Lindholmomneme 77 lineata, Anatina 567, 602 Eurytellina 521, 614 Glycimeris americana 585, 597 Glycymeris 567, 597 Tellina 559, 565-568, 570-571, 573, 579, 616 Tellina var. albida 579, 614 lineolaris, Aequipecten 517, 533, 549, 552, 565, 570-571, 576-577, 586, 606 Argopecten 572 Linga 278, 584, 600-601 amiantus 570, 584, 602 leucocyma 570 pensylvanica 557, 570, 576, 582, 584, 586 trisulcata 583, 600 linguafelis, Chama 405, 410 lintea, Lucina 579, 601 Lucina (Lucina) 560, 601 Tellina 552, 568, 576, 615 Lioberis 603 castaneus 551 Lioberus castaneus 516, 551, 570, 572, 584-585, 603 Lioconcha 277 lioica, Abra 520, 533, 549, 558, 560, 567, 51.1583, 585, 611 Syndosmya 549, 558 liorhina, Leda (Yoldia) 559 Yoldia 525, 539, 622 Lippistes acrilla 547 Lirapex costellata 176 lirata, Lucidella 217, 220, 223 651 Lirophora 427, 618-619 clenchi 524, 620 latilirata 524, 572, 620 paphia 524, 620 lisetae, Nesiocypraea 132, 143 listeri, Antigona 429, 441, 549-550, 556, 563, 567, 570, 576-577, 579, 588-589 Antigona (Antigona) 430 Antigona (Dosina) 430, 575 Antigona (Periglypta) 430 Arca 581, 590 Chione 441 Cytherea 429, 441 Cytherea (Cytherea) 429, 561 Dosina 429, 440 Dosinia 441 Isognomon 576, 598 Melicerona 141, 156 Melina 581, 598 Omphaloclathrum 429 Pedalion 566, 568, 580, 598 Periglyphus 430, 586 Periglypta 427-430, 431, 432, 433-438, 439-444, 524, 531, 533, 535, 553, 562, 570, 572, 581-582, 586, 620 Tellina 567, 570, 573, 584, 586, 588, 616 Tellina (Tellinella) 553 Tellinella 522, 617 Venus 429, 441, 579 Venus (Chione [Omphaloclathrum]) 429 Venus (Periglypta) 429 Lithodomus 564, 603-604 antillarum 556, 567 aristata 567 bisulcatus 567, 579, 604 candigerus 603 caudigerus 603 forficatus 579, 603-604 lithophagus 556, 577, 604 niger 579, 604 nigra 567 Lithophaga 339, 568, 578, 586, 604 antilarum 587, 603 antillarum 339, 342, 343, 516, 552, 556, 563, 566, 569-570, 572, 576, 582, 584, 586-588, 603 aristata 339, 343, 344, 516, 551, 563, 570, 572, 582, 584, 603 bisulcata 339, 344, 345, 516, 551-552, 570, 572, 582, 584, 604 nigra 516, 551-552, 563, 566, 570, 572, 581-582, 586, 588, 604 Lithophaginae 278 Lithophagis nigra 562, 604 Lithophagus 603 652 INDEX antillarum 559-560 bisulcatus 559, 604 caribaeus 559, 604 forficatus 559, 604 lithophagus, Lithodomus 556, 577, 604 Lithopoma americanum 582 lobata, Chama 410 locklini, Lima 585 Limaria 514, 599 longicallis, Abra 611 longicallus, Abra 559, 611 longisquamosa, Avicula 476 Avicula (Meleagrina) 476 Meleagrina 476 Pinctada 473-474, 474, 476, 477-484, 486-487, 486-489, 489-493, 520, 530-532, 534-535, 572-573, 611 Pteria 476, 550, 555, 564, 583, 611 longisqvamosa, Avicula 476 Lopha 606 cristagalli 316-317 frons 586 Lophinae 310, 320 Loripes 601 chrysostoma 571, 599 edentula 559, 571, 599 edentula var. chrysostoma 559, 599 lens 559 lucinalis 424 loveni, Macoma 167 luchuana, Eclogavena 143 Lucidella aureola 224 lirata 217, 220, 223 lucidum, Amussium 558, 610 Lucina 277-278, 292-293, 584-585, 600-602 adansoni 277, 288, 290-291, 291-293 amianta 572 amiantus 576, 601-602 (Anodontia) jamaicensis 567, 601 (Anodontia) trisulcata 567, 600 aurantia 293 belizana 292 (Bellucina) amiantus 567, 601 blanda 584 carnosa 291, 293 (Cavilucina) 601 (Cavilucina) muricata 580 chrysostoma 561, 599 costata 579 dentata 577 (Divaricella) 600 (Divaricella) dentata 567 (Divaricella) quadrisulcata 559 floridana 551, 572, 577, 602 (Jagonia) 600-601 (Jagonia) costata 567 (Jagonia) orbiculata 567 (Jagonia) orbiculata filiata 567, 600 (Jagonia) orbiculata recurvata 567, 600 (Jagonia) pectinella 567 jamaicensis 571, 601 lenticula 622 leucocyma 547, 572, 576-577, 600 lintea 579, 601 (Loripinus) 599 (Loripinus) edentula 567, 599 (Loripinus) edentula chrysostoma 567, 599 (Loripinus) schrammi 567 (Lucina) 600-602 (Lucina) costata 559, 601 (Lucina) crenulata 559 (Lucina) filosa 559 (Lucina) floridana 559 (Lucina) lenticula 559, 622 (Lucina) leucocyma 559 (Lucina) lintea 560, 601 (Lucina) multilineata 559, 601 (Lucina) muricata 559 (Lucinisca) 601 (Lucinisca) muricata 550, 562, 566-567, 577, 580, 601 (Lucina) pecten 559, 600 (Lucina) pectinella 559 (Lucina) pensylvanica 556 (Lucina) pensylvannica 559, 600 (Lucina) sagrinata 559 (Lucina) scabra 559, 601 (Lucina) sombrerensis 559 (Lucina) squamosa 559, 600 (Lucina) tigrina 559, 600 (Lucina) trisulcata 559, 599-600 multilineata 583-585, 601 muricata 570, 579, 585 muricatus 580, 601 nassula 565, 568, 570, 584-585 (Parvilucina) 601 (Parvilucina) crenella 567 pecten 579, 600 pectinata 570, 572, 583, 585, 600 pensylvanica 277-278, 279-288, 284, 288-293, 417-418, 421-422, 422-425, 515, 530-535, 553, 556, 563, 5704572. 574, 578, 581, 583, 587-588, 600 pensylvannica 567, 579, 600 (Pleurolucina) 601 (Pleurolucina) leucocyma 556 (Pleurolucina) sombrerensis 556 podagrina caymanana 293 podagrina podagrina 293 quadrisulcata 579 radians 572, 584-585, 599, 601 roquesana 293 rosceorum 293 roscoeorum 291 scabra 601 sombrerensis 547, 572, 576, 601 squamosa 579, 600 tigerina 558, 571, 600 tigrina 574, 579, 600 trisulcata 572, 600 lucinalis, Loripes 424 Lucinidae 247, 277-279, 291, 417, 514, 527, 583-585, 599, 622 Lucinisca 292 muricata 515, 572, 601 nassula 302, 515, 530-535, 555, 572-573, 583, 601, 622 Lucinoidea 52 Lucinoma filosa 515, 530, 533, 601 filosum 572 Lucinopsis 619 tenuis 559 Lunarca 590 ovalis 510, 591 lunella, Abrina 157, 162, 165, 166-167 Macoma 158 lunulata, Crassatella (Eriphyla) 559 Crassatella (Eriphyla) var. parva 559, 595 Crassinella 512, 570, 572, 584-585, 595 Luria 131, 136 cinerea 133, 135, 151 controversa 143 gilvella 143 isabella 133, 135, 137, 151 isabellamexicana 133, 135, 151 lurida 133, 135, 151 pulchra 133, 135, 151 tessellata 133, 135, 151 lurida, Luria 133, 135, 151 Luriinae 131, 134, 136 lutea, Chama sarda 403 Palmadusta 138, 153 Lutricola gruneri 579, 615 interstriata 559, 615 Lyncina 131 (Arestoides) argus argus 151 (Arestoides) argus contrastriata 151 aurantium 135 (Austrocypraea) reevei 151 broderipii 135-136 (Callistocypraea) aurantium 151 (Callistocypraea) broderipii 151 (Callistocypraea) leucodon 151 INDEX 653 (Callistocypraea) nivosa 151 camelopardis 143 carneola 135, 151 joycae 143 kuroharai 135-137, 151 leucodon 135 leviathan 135, 151 [упх 135, 151 (cf. Miolyncina) ропеп 151 nivosa 135-136 porteri 135 propinqua 135, 151 schilderorum 135, 151 sulcidentata 135, 137, 151 ventriculus 135-137, 151 vitellus 135, 151 lynx, Lyncina 135, 151 Lyonsia 278, 602 beana 559, 567, 584, 602 floridana 515, 572, 602 hyalina 42 hyalina floridana 565, 584-585, 602 Lyonsiidae 515, 602 Lyrodus pedicellata 576, 617 pedicellatus 523,617 Lyropecten 607 antillarum 565, 570, 576, 578, 586, 608 kallinubilosus 518, 572, 608 (Nodipecten) nodosus 551, 608 nodosus 570, 576, 587, 608 macandrewi, Erosaria 133-134, 148 Macerophylla 395 macerophylla, Chama 382-389, 393, 395-396, 397, 398, 401-402, 405, 408, 410—411, 512; 531, 533, 553, 556—557, 559, 563-564, 567, 570-572, 574, 576-577, 579, 585-586, 588, 593 Chama var. purpurascens 395 Chama var. sulphurea 395 Macoma 157, 167, 273, 560, 584-585, 615 anomala 579, 610 balthica 167 brevifrons 521, 559, 567, 570, 572, 584, 615 calcarea 167, 549-550, 615 carlottensis 613 cerina 522, 559, 567, 572, 576, 615 cimula 567, 615 constricta 522, 567, 572, 587, 615 crassula 167 extenuata 522, 576, 615 fausta 579, 614 incongrua 167 lama 167 654 leptonoidea 613 leptonoides 567, 613 limula 522, 576, 615 loveni 167 lunella 158 mitchelli 522, 572, 615 pseudomera 522, 576, 615 tageliformis 522, 572, 615 tenta 522,559 567, 572: 585, 615 tenta souleyetiana 567, 615 tenta var. souleyetiana 559, 615 Macrocallista 444 (Callista) gigantea 567, 620 (Chionella) 620 (Chionella) maculata 561 maculata 524, 567, 570, 572, 584-585, 620 nimbosa 524, 549-550, 561, 572, 585, 620 Macrocypraea 131 cervinetta 135, 149 cervus 135, 149 zebra 135, 149 macrophylla, Chama 395, 556, 581 Mactra 602 brasiliana 559, 602 fragilis 567, 570, 578, 583 lateralis 599 solidissima var. similis 559, 602 mactracea, Gouldia 567, 595 Mactridae 515, 583-584, 602 mactroides, Cytherea (Tivela) 559, 567, 621 Tivela 525, 550, 552, 566, 568, 580, 621 Mactrotoma fragilis 515, 572, 602 maculata, Cytherea (Callista) 559 Macrocallista 524, 567, 570, 572, 584-585, 620 Macrocallista (Chionella) 561 maculifera, Mauritia maculifera 135, 150 maculosa, Natica 295-296 magna, Abrina 157, 164, 165, 167 Laciolina 521, 614 Те!та 559, 565, 570, 573, 586, 616 Tellina (Laciolina) 553 magnifica, Cribrarula exmouthensis 138, 140, 154 Venus 440-441 magnum, Acrosterigma 511, 591 Cardium 559, 567, 575 Trachycardium 549-550, 552, 557, 565, 567, 570% 57315715 511156804598 Trachycardium (Acrosterigma) 550, 577 major, Alcadia 224 majori, Oxychilus 19, 22-23, 30-34 INDEX Malleidae 489, 516, 602 malleolus, Teredora 523, 617 Malleus candeanus 489, 516, 554, 570, 572, 586, 602 (Parimalleus) 602 mangle, Rhizophora 501 mantilla, Plicatula 587, 610 Mantissa 440 mappa, Leporicypraea 136 Leporicypraea mappa 135-136, 149 Margarita 475 margarita, Cetoconcha 610 Pustularia 135, 138, 152 margaritacea, Striostrea 316 margaritaceum, Periploma 518, 609 Margaritifera radiata 567, 611 margaritifera, Avicula 574 Pinctada 473, 491, 520, 528, 611 margaritiferus, Mytilus 475 Margaritiphora 475 radiata 559, 611 marginalis, Erosaria 133-134, 148 marginata, Zoila 136 Zoila marginata 135-136, 150 Marginellidae 295 mariae, Annepona 135, 151 mariellae, Zoila 135-136, 150 marina, Zostera 486 maritima, Polymesoda 512, 535, 570, 572, 583, 594 Polymesoda (Pseudocyrena) 551 Pseudocyrena 573-575, 583 marquesana, Purpuradusta fimbriata 138, 141-142, 156 martensii, Pinctada 492 Pinctada fucata 479 Martesia caribaea 567, 609 corticaria 559, 609 corticata 609 cuneiformis 519, 559, 567, 572, 579, 582, 609 striata 519, 559, 567, 572, 582, 609 martini, Notadusta 140-142, 156 martinicensis, Crassinella 512, 570, 572, 584-585, 595 Merisca 522, 615 Tellina 568, 570, 573, 616 Tellina (Merisca) 553 martybealsi, Mauritia maculifera 135, 150 massieri, Barycypraea fultoni 135 Mastigeulota 81, 85, 90, 106-108 kiangsinensis 92, 111 mauiensis, Pustularia 135, 138, 152 Mauritia 131, 136 arabica 135-136 arabica arabica 135, 150 arabica asiatica 135, 150 arabica immanis 135, 150 depressa depressa 135-136, 150 depressa dispersa 135-136, 150 eglantina 135, 150 grayana 135, 150 histrio 135, 150 maculifera maculifera 135, 150 maculifera martybealsi 135, 150 maculifera scindata 135, 150 таипйапа 135, 150 scurra indica 135, 149 scurra scurra 135, 149 mauritiana, Mauritia 135, 150 mauryae, Periglypta 432 maximus, Limax 212, 214 mazyckii, Chione 523, 531, 572, 619 Venus (Chione) 568 mcgintyi, Hyotissa 513, 566, 597 Parahyotissa 309-310, 312, 313, 315-316, 320, 566 Parahyotissa (Parahyotissa) 310 media, Americardia 511, 533, 565, 569-571, 584-585, 588, 591 Americardium 556 Trigoniocardia (Americardia) 557 medium, Cardium 559-560, 579, 591 Cardium (Hemicardium) 567, 591 Hemicardium 571, 591 Trigoniocardia 591 Trigoniocardia (Americardia) 591 Trigonocardia 583 Trigonocardia (Americardia) 580 Megaxinus floridanus 556, 602 melanesiae, Erronea onyx 141 Erronea (Adusta) onyx 155 Melanodrymia aurantiaca 169, 174-175, 174 Meleagrina 475 longisquamosa 476 Melicerona 131, 140, 143 felina 141, 143, 156 felina fabula 143 felina felina 143 listeri 141, 156 listeri melvilli 141, 143, 156 Melina 597 alata 552 lata 581, 597 listeri 581, 598 melvilli, Melicerona listeri 141, 143, 156 melwardi, Cribrarula 140 Cribrarula cribraria 138, 154 Meoma ventricosa 564 INDEX 655 mera, Angulus 613 Tellina 302, 558-559, 568, 570, 573, 575-576, 578-579, 583-586, 616 Tellina (Angulus) 554 Tellina cf. 565, 568, 613 Mercenaria 394, 427, 440, 443, 583, 620, 622 campechiensis 428, 441, 524, 565, 570, 572, 586, 620 mercenaria 524, 620 mercenaria forma notata 572, 620 mercenaria, Mercenaria 524, 620 Mercenaria forma notata 572, 620 Venus 559, 561, 568 Venus var. mortoni 559, 620 Venus var. notata 561, 620 Meretricinae 428, 444 Meretrix 444 (Transennella) 621 (Transennella) conradina 561 meridionalis, Hyalinia 20-21 Oxychilus 19-24, 30-34 Merisca 616 aequistriata 522, 615 cristallina 522, 615 martinicensis 522, 615 meroglypta, Cataegis 180 merus, Angulus 521, 530-532, 534-535, 613 Mesodesmatidae 258 : messanensis, Leda (Leda) 559, 605 Yoldiella 605 Metis intastriata 561 Metodontia 79, 81, 84, 106-108 yantaiensis 85, 92, 111 miamiensis, Ctenoides 514, 573, 598 micans, Pfeifferia 85, 89, 90, 104, 108, 112 Microcardium 592 peramabile 557 microdinus, Pyrgodomus 217, 220, 223 microdon, Purpuradusta 142 Purpuradusta microdon 138, 141, 156 midatlantica, Pseudorimula 173 midwayensis, Nesiocypraea 132, 143 mikeharti, Cypraeovula 137-139, 143, 153 mildredae, Caribachlamys 518, 549, 572, 586, 607 Chlamys 570, 608 Pecten 580 Pecten imbricatus 607 Pecten (Chlamys) 568 Pecten (Chlamys) imbricatus 549, 552 mildredaea, Pecten (Chlamys) 607 miliaris, Егозапа 133-134, 143, 149 minor, Ensis 519, 572, 609 656 minoridens, Purpuradusta 138, 141-142, 155 minuta, Limopsis 514, 559, 572, 599 Типота 5, 12 Miolyncina 131 mirabilis, Strigilla 522, 554, 570, 572, 583, 616 misella, Palmadusta ziczac 138, 153 mitchelli, Масота 522, 572, 615 mitis, Ctenoides 533, 573, 598 modesta, Tellina 559, 568, 614 Modiola (Amygdalum) lignea 559, 603 (Amygdalum) polita 559, 602 (Amygdalum) polita Var. sagittata 559, 602 (Botula) cinnamomea 559, 602 (Botulina) opifex 559, 603 (Brachydontes) sulcata 559, 603 duplicata 587, 604 plicatula 571, 603 polita 552, 602 sulcata 556, 603 tulipa 556, 604 tulipa var. nigra 548, 556, 604 Modiolaria 604 arborescens 567, 602 cinnamomea 579, 602 demissus 567, 603 demissus granosissimus 567, 603 lateralis 559, 567, 581, 604 opifex 567, 603 sulcatus 567, 603 tulipus 567, 604 Modiolus americanus 516, 531-535, 548, 551, 5642565, 570, 572. 578.586, 588, 604 modiolus squamosus 570, 572, 584-586, 604 politus sagittatus 563, 602 squamosus 516, 535, 553, 583, 604 tulipus 568, 604 modiolus, Brachidontes 516, 562, 569, 572, 575, 603-604 moellendorffi, Stilonodiscus 86, 97, 112 molitrix, Hypophthalmichthys 205 Monachoides 73, 77 moneta, Monetaria 133, 148 Monetaria 130-132 annulus 133, 148 caputdraconis 133, 148 caputophidii 133 caputserpentis 133 caputserpentis caputophidii 148 caputserpentis caputserpentis 148 moneta 133, 148 INDEX obvelata 133, 143, 148 monilifera, Venus 441 Monorchiidae 420 Montacutidae 1, 584—585, 596 monteverdensis, Helicina 217-218, 220, 222 moreletiana, Nesiohelix 103 morrhuanus, Pitar 622 morrisoni, Thracia 523, 617 mortilleti, Oxychilus 32-33 mortoni, Cardium 581 Laevicardium 302, 511, 530-534, 552-553, 555, 557, 564-565, 568, 570, 572-573, 575, 578, 583-584, 592 Venus 579, 620 Morula musiva 296 Mulinia lateralis 515, 567, 570, 602 multangularis, Dianadema 42, 45, 48, 50 multicostata, Periglypta 432, 441-442, 442, 444 Venus 441 multilineata, Lucina 583-585, 601 Lucina (Lucina) 559, 601 Parvilucina 555, 570, 572, 577, 583, 601 Parvilucina (Parvilucina) 556, 601 multisquamata, Chlamys 570, 607 Laevichlamys 518, 572, 608 Muracypraea 131 mus 133, 135; 150 muricata, Lucina 570, 579, 585 Lucina (Cavilucina) 580 Lucina (Lucina) 559 Lucina (Lucinisca) 550, 562, 566-567, 5775801601 Lucinisca 515, 572, 601 Phacoides 577, 586 Pinna 559, 579, 610 muricatum, Cardium 559, 567, 571 Trachycardium 511, 531, 533-535, 556-557, 564, 570, 573, 578, 583,585: 593 muricatus, Lucina 580, 601 Phacoides 550, 565 Phacoides (Lucinisca) 561, 601 Muricidae 295 mus, Muracypraea 133, 135, 150 musaica, Pyropelta 171, 172, 179-180 muscosus, Aequipecten 551, 565, 570, 577, 584, 586, 606 Chlamys 576 Lindapecten 518, 533, 572, 606, 608 Pecten 584, 587 Musculus 585, 604 lateralis 516, 570, 572, 576, 584-586, 604 musiva, Morula 296 musumea, Notadusta 143 Palmulacypraea 138, 141, 155 Pamulacypraea 140 Mya arenaria 208, 296 suborbicularis 59 Myidae 516, 602 myojinensis, Shinkailepas 180 Myonera lamellifera 513, 539, 559, 595 limatula 595 paucistriata 513, 595 Myrtaea (Eulopia) sagrinata 561 Myrtea compressa 547 (Eulopia) 601 (Eulopia) sagrinata 550, 566 lens 601 pristiphora 622 sagrinata 515, 572, 574, 601 Myrteopsis 601 lens 515, 601 Mysella 564, 585, 588, 596 planulata 513, 555, 572, 583-584, 596 Mytilidae 247, 278, 339-340, 516, 527, 563, 580-581, 584-585, 597, 602 Mytilopsis 528 leucophaeata 538, 554, 572, 596 sallei 596 Mytilus 603 (Brachidontes) 603 (Brachidontes) exustus 563, 580 californianus 208, 549-550, 604 cubitus 571, 603 exustas 579 exustus 558-559, 567, 578-579 hamatus 559, 603 lavalleanus 579, 603 leucophaeata 513 margaritiferus 475 perna 587, 603-604 recurvus 567, 603 sallei 513, 572 trossulus 208 Naeromya 596 floridana 584 nana, Astarte 510, 559, 561, 567, 572, 576, 591 Nanina delavayana 79, 112, 114 nanus, Cyclopecten 519, 610 Маззатаае 295 Nassarius festivus 295 nassula, Lucina 565, 568, 570, 584-585 Lucinisca 302, 515, 530-535, 555, 572-573, 583, 601, 622 Parvilucina (Lucinisca) 556 INDEX 657 Phacoides 583 nasuta, Corbula 559, 567, 594 Natica 305 catena 296 maculosa 295-296 Naticarius 295, 305 canrena 295, 297, 302, 303, 305 Naticidae 295 naticoidea, Bathynerita 176-177, 176, 181 naticoides, Bathynerita 169 Cyathermia 175, 175 Neaera alternata 558 rostrata 558, 584 Neaeromya 564 floridana 585 nebrites, Erosaria 133, 149 Neilonella pusio 604 Neilonellidae 604 Nemocardium peramabile 511, 530, 532-533, 547-548, 572, 581, 592 tinctum 511, 572, 592 nemoralis, Cepaea 73 Neobernaya 131-132, 137 spadicea 135, 138, 152 neocaledonica, Nesiocypraea teramachii 130, 133, 148 Neolepetopsidae 169-170, 179-180 Neolepetopsis cf. gordensis 170, 170-171, 180 Neomphalidae 169, 174, 180-181 Neomphalus fretterae 169, 181 Neopycnodonte cochlear 310, 312, 315-316, 514, 572, 597-598 Neosimnia aequalis 133 Neoteredo reynei 273 neritella, Helicina 224 Neritidae 176-177, 181 Neseulota 83 Nesiocypraea 127, 130-132, 134, 140 aenigma 132, 143 hirasei 130 langfordi 130 lisetae 132, 143 midwayensis 132, 143 sakurai 130 teramachii 130, 132, 140 teramachii neocaledonica 130, 133, 148 Nesiohelix 81, 85, 96, 103, 106-108 caspari 103, 112 moreletiana 103 samarangae 103 swinhoei 85, 86, 86, 89, 94, 98, 103, 112 newtoni, Tivelina 296 niger, Lithodomus 579, 604 nigra, Lithodomus 567 658 Lithophaga 516, 551-552, 563, 566, 570, 572, 581-582, 586, 588, 604 Lithophagis 562, 604 nigropunctata, Pseudozonaria 135, 137-138, 152 nimbosa, Macrocallista 524, 549-550, 561, 572. 585, 020 nitens, Ervilia 520, 559-560, 562, 566-567, 570, 572, 579-580, 585, 612 Eurytellina 521,614 Tellina 570, 573, 616 Tellina (Eurytellina) 554 nitida, Avicula 559, 611 nitidula, Hyalinia var. amiatae 21 nivosa, Leptaxis 73, 75, 75-76 Lyncina 135-136 Lyncina (Callistocypraea) 151 noae, Arca 560, 571, 590 Arca (Arca) 559, 590 Arca var. americana 579, 590 nobilis, Anadara 563 Arca 589 Nodipecten 608 fragosus 518, 527-528, 530, 533, 579, 608 nodosus 570, 572, 577, 581, 608 nodosus, Lyropecten 570, 576, 587, 608 Lyropecten (Nodipecten) 551, 608 Nodipecten 570, 572, 577, 581, 608 Pecten 568, 571, 608 Pecten (Pecten) 559, 608 nodulosa, Arca (Byssoarca) 560, 589 Ricinula 571 Noetia (Eontia) 605 (Eontia) ponderosa 549, 551 ponderosa 516, 549-550, 563, 568, 570, 572, 578—579, 583, 605 Noetiidae 516, 604 notabilis, Anadara 356, 369, 377, 510, 530—531, 593, 585,551, 563, 569-571, 577, 580, 583, 585-586, 588-589 Anadara (Caloosarca) 585 Arca 372 Notadusta 131, 140, 142-143 hungerfordi 141, 156 martini 140-142, 156 musumea 143 punctata 140-141, 156 punctata berinii 141, 156 punctata punctata 141, 143, 156 punctata trizonata 141, 143, 156 rabaulensis 143 notata, Diplodonta 523, 618 Purpuradusta gracilis 138, 141, 156 Taras 568 INDEX Notocypraea 130-131, 137, 139 angusta 137 angustata 138, 152 comptoni 138, 152 declivis 138, 152 hartsmithi 137-138, 152 occidentalis 143 piperita 138, 152 pulicaria 138, 152 Nototeredo knoxi 523, 572, 617 novaehollandiae, Aspergillum 39, 39 novaezelandiae, Brechites (Foegia) 38-39 Foegia 37-43, 39-51, 45, 47-48, 50, 52 Nucinellidae 13 nucleiformis, Diplodonta 523, 618 Taras 568 Nucleolaria 130-132 granulata 133, 135, 143, 148 nucleus 133, 135, 148 nucleus, Aequipecten gibbus 576 Aequipecten (Plagioctenium) gibbus 549, 607 Argopecten 518, 553, 563, 569, 572, 586, 607 Argopectin 575 Chlamys 578 Nucleolaria 133, 135, 148 Pecten 568, 587 Pecten (Aequipecten) 580 Pecten (Pecten) 559 Pecten (Plagioctenium) 561, 566, 580 Nucula 605 aegeenis 584-585, 605 aegeënsis 559-560, 605 aegeensis 567-568, 572 aegensis 584, 605 calcicola 517, 548, 562, 572-573, 605 crenulata 517, 549, 572, 605 proxima 517, 531, 535, 554-555, 565, 568, 572, 576, 583, 588, 605 Nuculana 553, 581, 584, 604-605 acuta 517, 572, 583-585, 605 carpenteri 568, 576, 581 concentrica 517, 572, 584-585, 605 jamaicensis 517, 605 (Jupitaria) 605 pusio 550, 572 solidifacta 517, 605 solidula 568, 572, 577 verrilliana 517, 550, 566, 568, 572-573, 605 vitrea 517, 539, 605 Nuculanidae 517, 581, 605 Nuculidae 13, 517, 605 nuculoides, Semele 568, 584-585 Semele (Semelina) 554 Semelina 520, 570, 572, 612 numisma, Parahyotissa 309, 312, 315-316, 320 Parahyotissa (Numismoida) 310 Nutricola tantilla 444 nymphae, Erronea 143 obesa, Cuspidaria 513, 595 Cuspidaria (Cuspidaria) 559 obesus, Cryptodon 559, 617 obicularis, Codakia 577, 600 obliqua, Perna 559, 579, 597 Semele 558-559, 579, 612 Tellina 549 oblique, Perna 560, 597 obvelata, Monetaria 133, 143, 148 occidentalis, Arca 567, 569, 580, 590 Arca (Lunarca) 560, 590 Notocypraea 143 Scapharca 571, 590 Solemya 520, 568, 570, 572, 584, 612 Solenomya 559 ocellata, Erosaria 133, 149 Odantella 8 oglasicola, Oxychilus 32-33 ojianus, Alveinus 1-3, 3, 5, 12-13, 15 olivetorum, Retinella 23, 32-33 отй, Palmulacypraea 143 Omphaloclathrum listeri 429 onyx, Crepidula 185-186, 198-200 Erronea 141 Erronea (Adusta) 155 operculata, Aloidis 576, 595 Varicorbula 584-585, 595 Ophiophragmus septus 564 opifex, Modiola (Botulina) 559, 603 Modiolaria 567, 603 oppressus, Oxychilus 32-33 orbicular, Codakia 586, 600 orbicularis, Codakia 279, 288, 292, 417-418, 420, 420-421, 422, 423-424, 515-527, 530-535, 551, 553, 556-557, 561, 563, 565-567, 570, 572-574, 576, 578, 580, 582-589, 600, 622 Codakia (Codakia) 556 orbiculata, Ctena 302, 417-418, 419, 420-424, 422, 515, 530-535, 548, 553, 572-573, 600, 622 Ctena forma recurvata 550 Codakia 564-565, 567-568, 570, 577-578, 583, 600 Codakia form filiata 600 Codakia (Ctena) 556 Helicina 224 INDEX 659 Jagonia var. filiata 548, 561, 600 Jagonia var. recurvata 548, 561, 600 Lucina (Jagonia) 567 orbignyi, Arca (Noetia) 559, 591 orientalis, Cymatoica 521 Cymatoica forma hendersoni 614 Zoila 143 ornata, Caribachlamys 518, 564, 572, 586, 607 Chlamys 562, 570, 588, 608 Trigonulina 525, 573, 621 Verticordia 568, 622 Verticordia (Trigonulina) 559 ornatissima, Cardiomya 512, 538, 572, 595 ornatus, Pecten 560, 568, 574, 579, 587, 607 Pecten (Chlamys) 560, 607 Pecten (Pecten) 559, 607 Orobitella floridana 513, 572, 596 Orthalicidae 217 Ortizius 19, 33 oryzaeformis, Purpuradusta 138, 141-142, 155 ostergaardi, Erosaria 143 Ostrea 322, 394, 424, 605-606 algoensis 316-317 angasi 316-317, 322 aupouria 309, 311-312, 314-319, 378, 321, 322-323 chilensis 314, 316-317 conchaphila 316, 322 cristata 559, 568, 606 denselamellosa 316-317 edulis 316-317, 322 equestris 556, 570, 588-589, 606 floridensis 552, 605 foliata 587, 605 folium 605 frons 559, 568, 570-571, 575, 577, 586 iridescens 320 (Lopha) 606 parasitica 579, 605 permollis 563, 566, 606 puelchana 316-317, 322 rhizophorae 571 verginica 568, 605 virginica 559 weberi 310, 548, 550, 574 Ostreea equestris 583 Ostreidae 247, 309-310, 312, 316, 517, 583, 605 Ostreinae 310, 312, 320 Ostreola 322, 606 conchaphila 317 equestris 309-314, 317-318, 318-319, 3205321322323 517, 566, 570,572 660 stentina 322 ovalis, Anadara 571, 590 Lunarca 510, 591 ovata, Gastrochaena 352, 513, 552, 559, 567, 570, 572, 596 Gastrochaena (Rocellaria) 557 Rocellaria 552, 579, 596 Ovatipsa 131, 139 chinensis 139 chinensis amiges 138-139, 154 chinensis chinensis 138—139, 154 coloba 138, 154 Ovula ovum 133, 147 Ovulidae 127, 131-133 Ovulinae 132 ovum, Ovula 133, 147 Erronea 142 Erronea ovum 141, 155 owenii, Bistolida 138-139, 153 Oxychilus 19-21, 23, 26, 31-32, 34 clarus 33 diductus 32-33 draparnaudi 19-23, 30-34 lanzai 19, 21-22, 26, 29-31 majori 19, 22-23, 30-34 meridionalis 19-24, 30-34 mortilleti 32-33 oglasicola 32-33 oppressus 32-33 (Ortizius) forcartianus 21 (Ortizius) lanzai 19, 21, 31 (Ortizius) paulucciae 19, 31 (Ortizius) tongiorgii 21 paulucciae 19-23, 24-25, 27-29, 30, 31, 31-34 pilula 19, 22-23, 31-34 uzielli 19, 22 uziellii 19-20, 23, 30-34 Pachydermia laevis 175, 175, 181 pacifica, Chama 398 Pedicularia 133, 147 palauensis, Erronea caurica 141 Erronea ovum 142, 155 pallida, Erronea 141-142, 155 pallidula, Blasicrura 143 Blasicrura pallidula 141, 156 Palliolum strigillatum 549 Palmadusta 131, 139 androyensis 143 artuffeli 138-139, 153 asellus 139 asellus asellus 138-139 asellus cf. asellus 153 asellus bitaeniata 138-139 INDEX asellus cf. bitaeniata 153 asellus vespacea 138-139 asellus cf. vespacea 153 clandestina 139 clandestina candida 138-139 clandestina cf. candida 153 clandestina clandestina 138-139 clandestina cf. clandestina 153 clandestina passerina 138, 153 contaminata 139 contaminata contaminata 138, 153 contaminata distans 138, 153 diluculum 138-139, 153 humphreysii 143, 153 johnsonorum 143 lentiginosa 138, 153 lutea 138, 153 saulae 138, 153 ziczac 139 ziczac misella 138, 153 ZIeZae ziczac 138, 155 Palmulacypraea 140, 143 boucheti 143 katsuae 138, 141, 155 musumea 138, 141, 155 omii 143 Pamulacypraea 131, 140 katsuae 140 musumea 140 Pandora 584-585, 606 arenosa 517, 585, 606 bushiana 517, 554, 572, 606 glacialis 517, 539, 606 inflata 517, 530, 533, 554, 572, 606 (Kennerlia) 606 (Kennerlia) glacialis 560 Pandoridae 517, 606 panerythra, Leporicypraea mappa 136 pantherina, Cypraea 133, 135, 150 Paphia 443 paphia, Chione 552, 565, 570, 572, 581, 585-587, 619 Chione (Lirophora) 549 Lirophora 524, 620 Venus 579 Venus (Chione) 568 Paphridea hiatus 592 soleniformis 592 spinosum 587, 592 papyracea, Euvola cf. 518, 606, 608 papyraceum, Amusium 571, 576-577, 608, 606 Papyridea bullata 559, 592 hiatus 557 lata 511, 587, 592 (Liocardium) laevigatum 560, 592 (Liocardium) serratum 559, 592 petitiana 559, 592 semisulcata 511, 557, 570, 572, 576, 585-586, 592 semisulcatum 574, 592 soleniformis 511, 533, 552, 556, 563, 567, 570, 572, 575, 578, 587, 592 papyrium, Amygdalum 516, 571, 576, 602 Parahyotissa 309-310, 315, 320 mcgintyi 309-310, 312, 313, 315-316, 320, 566 numisma 309, 312, 315-316, 320 (Numismoida) numisma 310 (Parahyotissa) imbricata 320 (Parahyotissa) mcgintyi 310 (Pliohyotissa) quercinus 320 Paralepetopsis ferrugivora 170-171, 171, 180 paramera, Angulus 549 Tellina 553, 570, 573, 578, 616 Tellina (Angulus) 549 paramerus, Angulus 521, 613 parasitica, Cochliolepis 566 Ostrea 579, 605 Parastarte 583, 620 triquetra 524, 554-555, 561, 565, 568, 572, 580, 583, 588, 620 parva, Gouldia 566-567, 595 Parvamussium cancellatum 519, 610 thalassinum 519, 610 Parvilucina 600, 602 amianta 572 (Bellucina) amiantus 556, 602 blanda 570 costata 515, 601 crenella 515, 601 (Lucinisca) 601 (Lucinisca) nassula 556 multilineata 555, 570, 572, 577, 583, 601 (Parvilucina) 600-601 (Parvilucina) costata 556 (Parvilucina) multilineata 556, 601 (Parvilucina) pectinella 556 parvula, Donax 459 paschauli, Chama 387 paschuali, Chama 411 passerina, Palmadusta clandestina 138, 1153 Patella crenata 170, 180 vulgata 212, 216 paucistriata, Муопега 513, 595 Paulonaria 134 paulucciae, Hyalinia 20-21, 31 Oxychilus 19-23, 24-25, 27-29, 30, 31, 31-34 661 Oxychilus (Ortizius) 19, 31 ресет, Lucina 579, 600 Lucina (Lucina) 559, 600 Pecten 564, 569, 586, 606, 608 acanthodes 568, 608 (Aequipecten) 607 (Aequipecten) nucleus 580 (Amusium) 610 (Amusium) cancellatum 560 (Amusium) pourtalesianum 560, 610 (Amusium) pourtalesianum var. marmoratum 560, 610 (Amusium) sayanum 560, 610 antillarum 558, 568, 579, 585-588 (Chlamys) 607-608 (Chlamys) antillarum 560 (Chlamys) gibbus var. nucleus 560, 607 (Chlamys) imbricatus 566, 580-581, 607 (Chlamys) imbricatus mildredae 549, 552 (Chlamys) imbricatus var. mildredae 607 (Chlamys) mildredae 568 (Chlamys) mildredaea 607 (Chlamys) ornatus 560, 607 (Chlamys) pusio 581, 612 chazaliei 564, 570, 608 chazaliei (tereinus) 571, 608 dislocates 579, 606 dislocatus 558, 606 (Euvola) tereinus 549, 554, 561, 566, 608 exasperatus 568, 608 gibbus 568 heliacus 568 hemicyclica 579, 608 hemicyclicus 608 imbricatus 553, 568-569, 579, 585, 587, 607 imbricatus mildredae 607 irradians concentricus 549 (Janira) 608 (Janira) hemicyclica 560, 608 (Janira) ziczac 559 (Lyropecten) 607 (Lyropecten) antillarum 566, 580 mildredae 580 muscosus 584, 587 nodosus 568, 571, 608 nodosus fragosus 568, 608 nucleus 568, 587 ornatus 560, 568, 574, 579, 587, 607 (Pecten) 607-608 (Pecten) antillarum 559 (Pecten) effluens 560, 608 (Pecten) exasperatus 559, 608 (Pecten) imbricatus 559, 607 (Pecten) irradians var. dislocatus 559, 606 662 (Рецепт) nodosus 559, 608 (Pecten) nucleus 559 (Pecten) ornatus 559, 607 (Pecten) phrygium 560 (Pecten) sigsbeei 560, 596 (Pecten) thalassinus 560 phrygium 571 (Plagioctenium) 607 (Plagioctenium) gibbus var. amplicostatus 548 (Plagioctenium) nucleus 561, 566, 580 (Propeamussium) 610 (Propeamussium) cancellatum 560 (Propeamussium) pourtalesianum 560, 610 (Propeamussium) pourtalesianum var. marmoratum 560, 610 (Propeamussium) sayanum 560, 610 (Pseudamusium) sigsbeei 560, 596 (Pseudamusium) thalassinus 560 гауепе! 568, 608 sentis 568-569, 585, 587 tereinus 551, 563-564, 568, 608 ziczac 564, 568, 570, 608 pectinata, Glycimeris 585-586 Glycymeris 556, 563-564, 567, 570, 572, 576, 578, 581-584, 586, 588 Lucina 570, 572, 583, 585, 600 Phacoides 418, 423-424, 515, 570, 601 Tucetona 302, 513, 530-535, 573, 597 pectinatus, Glycimerus 569 Glycimirus 597 Glycymeris 567, 579-581, 597 Pectunculus 559, 597 Phacoides 551, 601 Phacoides (Phacoides) 556, 601 Tucetona 597 Pectinella sigsbeei 513, 539, 596 pectinella, Codakia 570, 572 Codakia (Ctena) 550, 564 Codakia (Jagonia) 566 Ctena 423, 515, 600, 622 Lucina (Lucina) 559 Lucina (Jagonia) 567 Parvilucina (Parvilucina) 556 Pectinidae 517, 527, 563, 581, 583-585, 606 pectiniformis, Pectunculus 571, 597 Pectunculus 603 castaneus 579 pectinatus 559 pectiniformis 571, 597 pennaceus 556, 597 undatus 559, 597 pectunculus, Tucetona 597 Pedalion 597-598 INDEX alata 580, 584, 597, 611 bicolor 568 listeri 566, 568, 580, 598 (Perna) 597 (Perna) alata 568 semiaurita 566, 568, 598 pedicellata, Lyrodus 576, 617 pedicellatus, Lyrodus 523, 617 Pedicularia 132 pacifica 133, 147 pellisserpentis, Cribrarula 138, 140, 154 pellucens, Talostolida 138-139, 154 pellucida, Chama 387, 391 Lima 551, 556, 562-563, 565, 567, 570, 578, 583-586, 588 Limaria 514, 531-532, 534, 555, 572, 583, 599 Limaria cf. 599 Peltospira smaragdina 175, 175, 181 Peltospiridae 169, 175-176, 180-181 Penicillidae 53 Penicillus 38, 246, 278, 452, 498 поуае zelandiae 39 penis, Brechites 50 pennacea, Glycymeris 567, 597 pennaceus, Pectunculus 556, 597 pennsylvanica, Lucina 567, 579, 600 Lucina (Lucina) 559, 600 pensylvanica, Linga 557, 570, 576, 582, 584, 586 Lucina 277-278, 279-288, 284, 288-293, 417-418, 421-422, 422-425, 515, 530-535, 553, 556, 563, 570, 572, 574, 578, 581, 583, 587-588, 600 Lucina (Lucina) 556 peramabile, Microcardium 557 Nemocardium 511, 530, 532-533, 547-548, 572, 581, 592 peramabilis, Cardium 559, 592 Cardium var. tinctum 560, 592 Cardium (Fulvia) 548, 558, 592 Cardium (Fulvia) var. tinctum 558, 592 Cardium (Protocardia) 567, 592 Protocardia 552, 592 Periglyphus 620 listeri 430, 586 Periglypta 427-429, 432, 440, 442, 443-444, 620 listeri 427-430, 431, 432, 433-438, 439-444, 524, 531, 533, 535, 553, 562, 570, 572, 581-582, 586, 620 таигуае 432 multicostata 432, 441—442, 442, 444 риегрега 441—442, 442, 444 reticulata 444 tamiamensis 432 tarquinia 432 Periploma angulifera 559, 561, 566, 568, 579-580, 609 anguliferum 550, 572, 609 margaritaceum 518, 609 tenera 560, 566, 568, 609 tenerum 518, 550, 572, 609 Periplomatidae 518, 609 Perisserosa 130-132 guttata 133, 148 Perkinsus 383 perlae, Zoila 143 Perlamater 475 permollis, Cryptostrea 309-310, 312-314, 316-317, 320, 321, 322, 572, 605 Ostrea 563, 566, 606 Perna ephippium 558-559, 579, 597 obliqua 559, 579, 597 oblique 560, 597 perna 556, 597 viridis 528, 553, 604 perna, Mytilus 587, 603-604 Perna 556, 997 pernula, Pinna 560, 610 perplana, Pteromeris 511,570, 572, 593 perrostrata, Cardiomya 512, 572, 576, 584-585, 595 Cuspidaria (Cardiomya) 559 persica, Tellina 522, 573, 616 petilirostris, Umbilia 134-135 Umbilia cf. 133-135, 149 petitiana, Papyridea 559, 592 Zonaria 143 petitianum, Cardium 579, 592 Petricola 339, 350, 559, 609 (Choristodon) robusta 559, 609 divaricata 558, 579, 609 lapicida 339, 347-348, 350, 350-351, 353, 518, 552, 562, 566, 568, 570, 572, 580, 584, 588, 609 (Naranaio) 609 (Naranaio) lapicida 560 pholadiformis 351, 559, 568, 570 stellae 350 Petricolaria pholadiformis 519, 572, 609 Petricolidae 247, 339, 347, 518, 609 Pfeifferia 81, 85, 106-109 micans 85, 89, 90, 104, 108, 112 Phacoides 600 (Lucinisca) muricatus 561, 601 (Lucinoma) filosus 556, 601 muricata 577, 586 muricatus 550, 565 nassula 583 INDEX 663 pectinata 418, 423-424, 515, 570, 601 pectinatus 551, 601 (Phacoides) pectinatus 556, 601 pharaonis, Brachidontes 208 Pharidae 519, 609 phaseolina, Thracia 523, 559, 573, 617 Phenacolepas pulchellus 177 Phenacolepidae 177, 181 Phenacovolva tokioi 133, 147 weaveri 133, 147 philippiana, Anodontia 572, 583, 599 Crepidula 198-199, 201 philippii, Varicorbula 512, 573, 595 Philippinae 429 philippinensis, Brechites (Penicillus) 38 Philobryidae 1, 519, 609 Phlyctiderma semiaspera 523, 618 soror 523, 618 Pholadidae 273, 519, 609 pholadiformis, Petricola 351, 559, 568, 570 Petricolaria 519, 559, 568, 572, 609 Pholadoidea 273 Pholas (Barnea) 608 (Barnea) costata 559, 568 (Ватеа) truncata 559, 568 campechiensis 519, 559, 568, 609 costata 571 hornbeckii 609 phrygium, Aequipecten 550 Cryptopecten 518, 530, 533, 564, 572, 606, 608 Pecten 571 Pecten (Pecten) 560 phrygius, Aequipecten 549, 606, 608 Chlamys 576, 608 picta, Zonaria 135, 137-138, 152 pictum, Laevicardium 511, 572, 592 piligerus, Pseudobuliminus (Pseudobuliminus) 96, 112 pilula, Diplodonta 618 Oxychilus 19, 22-23, 31-34 Venus 560, 619 Pinctada 316, 473, 475-476, 489, 611 fucata martensii 479 imbricata 316, 473-474, 489, 489-493, 520, 530-531, 533, 556, 564-565, 570, 572-574, 578, 584, 587-588, 611 longisquamosa 473-474, 474, 476, 477-484, 486—487, 486-489, 489-493, 520, 530-532, 534-535, 572-573, 611 margaritifera 473, 491, 520, 528, 611 martensii 492 radiata 476, 489, 492, 552, 554-555, 557, 565-568, 570, 576, 578, 583-584, 586, 588,611 664 vulgaris 485, 491—492 xanthia 476, 486 Pinna 566, 609-610 carnea 519, 559-560, 562, 568-570, 572, 575, 578-582, 588, 610 типса 559, 579, 610 pernula 560, 610 rigida 568 rudis 572, 610 seminuda 559 serrata 568 Pinnidae 519, 557, 580-581, 583, 609 piperita, Notocypraea 138, 152 piscinalis, Anodonta 208 pisiformis, Strigilla 522, 549-550, 556, 559, 561, 566, 568, 571-572, 580, 586-587, 616 Pisulina 181 pisum, Strigilla 579, 616 pitalensis, Helicina 217, 219, 221 Pitar 581, 584-585, 588, 620-621 albidus 524, 620 circinatus 524, 620 cordata 573, 620 cordatus 524, 549-550, 572, 574, 620 dione 524, 620 encymata 568, 618 fulminata 568, 583, 587, 620 cf. fulminata 565, 568, 620 fulminatus 524, 531, 535, 570, 572, 584-585, 620, 622 morrhuanus 622 (Pitarenus) 620 (Pitarenus) cordata 549, 577, 620 (Pitarenus) cordatus 550, 577 simpsoni 302, 524, 531, 535, 555, 570, 572-573, 583-585, 620 Pitaria cordata 578 cordatus 549 Pitarinae 428, 444 Placamen 296, 305, 444 calophyllum 288, 292, 295-296, 303, 305 tiara 443 Placunanomia 589 rudis 559 plana, Crepidula 185, 200-201 Scrobicularia 424 planispira, Planorbidella 175 Planorbidella planispira 175 planulata, Mysella 513, 555, 572, 583-584, 596 planulatatus, Ctenoides 572, 598 planulatus, Ctenoides 514, 573, 598 platychila, Helicina 224 platyodon, Camaena 105, 112 INDEX Platypetasus 81, 106-108 plebeius, Tagelus 520, 570, 572, 581, 612 Plecteulota 83 Plectodon granulatus 513, 550, 572, 574, 595 Plectotropis 81, 106-108 Pleurolucina 532, 600-601 leucocyma 515, 530, 533, 601, 622 sombrerensis 515, 530, 533, 601, 622 Pleuromeris tridentata 302, 511, 564, 570, 572-573, 577, 581, 584-585, 593 Pleurotomariidae 180 plicata, Arca 581, 589 plicatella, Anatina 570 Raeta 516, 572, 602 Pliocathaica 81, 106-109 Plicatula gibbosa 519, 530, 532-533, 568, 570: 572, 583, 586, 610 mantilla 587, 610 ramosa 559, 610 spondyloidea 552, 610 plicatula, Alectryonella 316-317 Modiola 571, 603 Plicatulidae 519, 610 podagrina, Lucina podagrina 293 Pododesmus rudis 510, 589 Polinices 295, 305 alderi 296 catena 296 duplicatus 296, 305 lewisi 216 lewisii 305 tumidus 296, 305 polita, Modiola 552, 602 Modiola (Amygdalum) 559, 602 Modiola (Amygdalum) var. sagittata 559, 602 politum, Amygdalum 516, 571, 602 Polodesmus decipiens 568, 589 rudis 589 Polycyrena floridana 551, 594 Polygridae 84 Polymesoda 554, 594 floridana 565, 568, 580, 594 maritima 512, 535, 570, 572, 583, 594 (Pseudocyrena) maritima 551 polymorpha, Dreissena 208 pomatia, Helix 85, 91, 92, 99, 112, 212, 215-216 ponderosa, Arca 567 Arca (Noetia) 559 Noetia 516, 549-550, 563, 568, 570, 572, 578-579, 583, 605 Noetia (Eontia) 549, 551 Ponsadenia 83 рогапа, Erosaria 133-134, 148 Porcellidae 180 porculus, Kellia 58, 63, 64-66, 71 porifera, Ctenopelta 175, 176, 181 Porites 245-246, 452-454 Poromya albida 519, 539, 610 (Cetomya) 610 (Cetomya) albida 560 elongata 610 granulata 519, 558, 560-561, 568, 572, 610 granulata granulata 566, 610 rostrata 519, 572, 610 Poromyidae 13, 519, 610 porteri, Lyncina 135 Lyncina (cf. Miolyncina) 151 pourtalesianum, Pecten (Amusium) 560, 610 Pecten (Propeamussium) 560, 610 Pecten (Amusium) var. marmoratum 560, 610 Pecten (Propeamussium) var. marmoratum 560, 610 Propeamussium 519, 572, 610 prætexta, Chama 401 Primovula 132 concinna 132-133, 147 Prionovolva brevis 133, 147 prismatica, Striostrea 320 pristiphora, Myrtea 622 probina, Tellina 613 probrina, Angulus 549 Tellina 553, 570, 573, 616 Tellina (Angulus) 549, 554 probrinus, Angulus 521, 613 Procalpurnus lacteus 133 Proctotrema 423 producta, Anomalocardia 618 Chama 410 proficua, Semele 520, 554, 568-570, 572, 583-585, 612 profundorum, Abra 271-273 Propeamussiidae 1, 519, 610 Propeamussium cancellatum 539 dalli 610 pourtalesianum 519, 572, 610 sayanum 519, 572,610 Propeleda carpenteri 517, 605 propinqua, Lyncina 135, 151 Propustularia 127, 130-132, 134 surinamensis 133, 148 Prosimnia 132 semperi 132-133, 147 protacta, Donax roemeri 459 protea, Crepidula 198-201 protexta, Egeta 567 INDEX 665 Ensitellops 521, 613 Protocardia peramabilis 552, 592 tincta 561, 566, 592 Protolira valvatoides 172, 173, 179 Protothaca 427, 618 granulata 572, 619 staminea 305 protracta, Donax 459 Donax roemeri 459 protractus, Donax fossor 567 Provanna variabilis 177, 178 Provannidae 178, 181 proxima, Nucula 517, 531, 535, 554-555, 565, 568, 572, 576, 583, 588, 605 Psammobiidae 247, 249, 258-259, 262, 519, 610 Psammotreta 164, 167, 615 intastriata 586, 615 (Tellinimactra) edentula 167 Pseudamusium 610 strigillatum 549, 560 Pseudaspasita 81, 85, 106-108 binodata 92, 102, 112 Pseudiberus chentingensis 104, 112 (Platypetasus) 79 Pseudobuliminus 81, 106-108 (Pseudobuliminus) piligerus 96, 112 Pseudochama 381, 385-392, 407, 408, 410-412, 593-594 buchivacoana 387, 411 caloosana 387, 411 corticosa 386-387, 411 corticosaformis 387, 411 cristella 390-392, 403, 407, 412 draconis 387, 411 (Echinochama) arcinella 408 exogyra 391 ferruginea 386, 401 inezae 381, 405, 411, 549-550, 553, 572, 575, 580, 594 lazai 387, 411 quirosana 387, 411 radians 381, 392, 401, 552, 563-564, 570, 572, 583, 594 radians variegata 401, 567, 594 riocanica 387, 411 scheibei 387, 411 similis 390-391 Pseudocypraea 132 adamsonii 133, 147 exquisita 132-133, 147 Pseudocyrena 594 floridana 549-550, 563, 576, 578, 594 maritima 573-575, 583 pseudoillota, Fugleria 355 666 рзеиаотега, Масота 522, 576, 615 Pseudomiltha floridana 601 Pseudorimula midatlantica 173 Pseudozonaria 131-132, 137 angelicae 137 annettae 135, 138, 152 arabicula 135, 137-138, 152 nigropunctata 135, 137-138, 152 pyrum 137 pyrum angolensis 137 pyrum senegalensis 137 robertsi 135, 138, 152 Репа 473, 475, 487, 489, 555, 611 argentea 491 colymbus 473-474, 476, 489-493, 520, 533, 547, 563-565, 570, 572-575, 580-581, 588, 611 hirundo 473, 491 hirundo vitrea 564, 566, 611 longisquamosa 476, 550, 555, 564, 583, 611 radiata 553, 581, 611 sterna 491 viridizona 476, 491 viridozona 476 vitrea 473, 491, 520, 550, 571, 575, 611 xanthia 476, 491 Pteriadae 475 Pteriidae 247, 473, 475, 490, 520, 585, 611 Pterioidea 475, 489 Pteromeris perplana 511, 570, 572, 593 puber, Chione 570, 621 pubera, Chione 566, 579-580, 619 Chione (Chione) 550 Puberella 524, 572, 621 Venus (Chione) 568 Puberella 619 intapurpurea 524, 533, 572, 621 pubera 524, 572, 621 puelchana, Ostrea 316-317, 322 puerpera, Periglypta 441-442, 442, 444 Venus 429, 440-441 pulchella, Chama 390 Contradusta 141-142, 156 Erronea 142 Eutrochatella 217, 224 Truncatella 328 pulchellus, Phenacolepas 177 pulchra, Luria 133, 135, 151 pulex, Xenostrobus 296 pulicaria, Notocypraea 138, 152 pullastra, Venerupis 444 punctata, Codakia 424 Diplodonta 523, 570, 572, 583, 585-586, 618 INDEX Diplodonta (Diplodonta) 571, 596 Notadusta 140-141, 156 Notadusta punctata 141, 143, 156 Taras 568 punicea, Eurytellina 521, 614 Tellina 565, 573, 586, 616 Tellina (Eurytellina) 549 Purpuradusta 130-131, 140, 142 barbieri 143 fimbriata 142 fimbriata fimbriata 138, 141, 156 fimbriata marquesana 138, 141-142, 156 fimbriata unifasciata 142, 156 fimbriata cf. unifasciata 138, 141 fimbriata waikikiensis 138, 141-142, 156 gracilis gracilis 138, 141, 156 gracilis notata 138, 141, 156 hammondae 138, 141, 156 microdon 142 microdon chrysalis 138, 141-142, 156 microdon microdon 138, 141, 156 minoridens 138, 141-142, 155 oryzaeformis 138, 141-142, 155 serrulifera 138, 141, 155 purpurascens, Semele 520, 548-549, 554, 568-570, 572, 585, 612 Venus 549 purpurea, Gemma 567, 619 pusio, Neilonella 604 Nuculana 550, 572 Pecten (Chlamys) 581, 612 Pustularia 131, 134, 137 bistrinotata 137 bistrinotata bistrinotata 135, 137, 152 bistrinotata keelingensis 135, 137-138, 152 bistrinotata sublaevis 135, 137-138, 152 chiapponii 143 cicercula 135, 138, 152 globulus brevirostris 135, 138, 151 globulus globulus 135, 138, 151 margarita 135, 138, 152 mauiensis 135, 138, 152 spadicea 138 Pustulariinae131 pustulata, Jenneria 133, 147 pustulosus, Lepetodrilus 173, 173 Pycnodonteinae 310, 312 pygmaea, Chione 570, 575, 583, 619 Chione (Timoclea) 561, 566, 579-580 Timoclea 525, 573, 621 Venus 559, 579 Venus var. inaequivalvia 579, 621 pygmaeus, Venus (Chione) 568, 621 pyramidatum, Cochlodesma 518, 550, 609 Pyrgodomus microdinus 217, 220, 223 pyriformis, Cryptodon 559, 617 Erronea 141-142, 155 Pyropelta musaica 171, 172, 179-180 Pyropeltidae 171, 180 pyrum, Pseudozonaria 137 Zonaria 135, 138 Pyxidicula 8 quadrata, Basterotia 521, 559, 567, 569, 572.613 quadrimaculata, Eclogavena quadrimaculata 141, 156 quadrisulcata, Divalinga 515, 533, 572, 600, 622 Divaricella 292, 570, 583 Divaricella (Divaricella) 556 Lucina 579 Lucina (Divaricella) 559 quercinus, Parahyotissa (Pliohyotissa) 320 quinquefasciata, Erronea caurica 141-142, 155 quirosana, Pseudochama 387, 411 rabaulensis, Erronea 141-142, 155 Notadusta 143 radiala, Tellina 582, 616 radians, Callucina (Callucina) 556, 599 Chama 382-384, 386, 388-392, 400, 401-403, 402, 405, 410-412, 512, 556, 594 Chama radians 402 Lucina 572, 584-585, 599, 601 Pseudochama 381, 392, 401, 552, 563-564, 570, 572, 583, 594 radiata, Avicula 560, 579, 611 Egeria 469 Margaritifera 567, 611 Margaritiphora 559, 611 Pinctada 476, 489, 492, 552, 554-555, 557, 565-568, 570, 576, 578, 583-584, 586, 588, 611 Pteria 553, 581, 611 Semele 569, 612 Tellina 522, 551, 556, 559, 562, 564, 567-568, 570-571, 573, 579, 584, 586, 588, 616 Tellina (Tellina) 553 Tellina var. unimaculata 571, 584, 586, 616 radiatus, Isognomon 514, 551, 562, 565, 570-572, 578, 588, 598 Radiolucina amianta 515, 533, 601 Raeta plicatella 516, 572, 602 ramosa, Plicatula 559, 610 ramosus, Spondylus 571 INDEX 667 Rangia flexuosa 553, 572, 594 Rangianella 594 rashleighana, Talostolida 143 Rasta 277 raveneli, Euvola 518, 572, 576, 608 Pecten 568, 608 Spisula 516, 572, 602 ravida, Acusta 89, 98, 99, 111 recurvata, Codakia (Jagonia) orbiculata 566, 600 Lucina (Jagonia) orbiculata 567, 600 recurvum, Ischadium 516, 572, 603 recurvus, Brachidontes 570, 603 Brachiodontes 566, 603 Mytilus 567, 603 redimita, Erosaria lamarckii cf. 133-134, 149 reevana, Chama 390 reevei, Austrocypraea 135 Lyncina (Austrocypraea) 151 resticulata, Venus 441 reticulata, Acar 363 Acar (Byssoarca) 559, 589 Arca 567, 589 Arca (Acar) 579-580, 589 Periglypta 444 Semele 559, 579, 612 Venus 440-441 reticulatus, Agrolimax 215 Retinella olivetorum 23, 32-33 Retiskenea diploura 174, 174, 179 rewa, Leporicypraea mappa 136 reynei, Neoteredo 273 rhinoceros, Blasicrura pallidula 141, 143, 156 Rhizophora mangle 501 rhizophorae, Crassostrea 310, 316, 517, 572, 605 Ostrea 571 Ricinula nodulosa 571 rigida, Antigona 567, 586 Antigona (Circomphalus) 566 Antigona (Ventricola) 566 Antigona (Ventricolaria) 565 Atrina 519, 551, 565-566, 569-570, 572, 578, 582, 588-589, 609 Cytherea (Ventricola) 561 Globivenus 441, 524, 572, 619 Pinna 568 Ventricolaria 550, 562, 570, 574 Ventricolaria Ventricolaria 577 riocanica, Pseudochama 387, 411 riopejensis, Helicina beatrix 220-221 robertsi, Pseudozonaria 135, 138, 152 robusta, Petricola (Choristodon) 559, 609 668 INDEX Tellina 571 saccharina, Arca 575, 589 robustum, Choristodon 339, 346, 346-347, Saccostrea commercialis 316 350-351, 518, 572, 584, 609 cucullata 316 Dinocardium 511, 570, 592 sachalinica, Abrina 157 Rocellaria 596-597 sagittatum, Amygdalum 516, 571, 584-585, hians 551 602 ovata 552, 579, 596 sagittatus, Modiolus politus 563, 602 rostrata 579 sagrinata, Arca (Macrodon) 559 roemeri, Donax 459, 567, 596 Bentharca 510, 539, 591 Donax roemeri 459 Lucina (Lucina) 559 rombergii, Strigilla 561, 568, 616 Myrtaea (Eulopia) 561 romgergi, Strigilla 570, 616 Myrtea 515, 572, 574, 601 roquesana, Lucina 293 Myrtea (Eulopia) 550, 566 rosceorum, Lucina 293 Sahlingia 180 roscoeorum, Lucina 291 xandaros 171, 172, 180 rosea, Leporicypraea mappa 135-136, 149 sakurai, Austrasiatica 132, 138, 140-141, rosewateri, Crenavolva 133, 147 155 Crenavolva cf. 133, 147 Nesiocypraea 130 rosselli, Zoila 135, 150 sallei, Mytilopsis 596 rossmassleri, Congeria 567, 596 Mytilus 513, 572 rossmässleri, Сопдепа 566, 596 samarangae, Nesiohelix 103 rostrata, Cuspidaria 513, 551, 558, 572, 595 Samarangia 277 Gastrochaena 552, 567 samoensis, Erronea саипса 141-142, 155 Gastrochaena (Spengleria) 559 sanctipauli, Ctenoides 514, 548, 572-573, Neaera 558, 584 581, 598 Poromya 519, 572, 610 sandix, Scissula 615 Rocellaria 579 Tellina 616 Spengleria 278, 513, 552, 557, 570, 572, Tellina (Scissula) 615 3% Sanguinolaria sanguinolenta 556, 588 Venus (Anomalocardia) 559, 618 sanquinolenta 519, 610 rottnestensis, Cribrarula cribraria 138, 140, sanguinolenta, Sanguinolaria 556, 588 154 Zonaria 135, 138, 152 rotunda, Alcadia 224 sanquinolenta, Sanguinolaria 519, 610 Chama 402 sarda, Chama 381-384, 386, 388-389, rotundatus, Discus 215 rubea, Chama 410 rubiginosa, Bistolida stolida 138-139, 141, 153 rubra, Kellia 566, 596 Lasaea 5, 550 Lasea 579, 596 ruderalis, Chama 401, 408 rudis, Pinna 572, 610 Placunanomia 559 Pododesmus 510, 589 Polodesmus 589 rugatina, Cytherea (Cytherea) 561 Globivenus 441, 524, 572, 619 Ventricolaria 564 rugosa, Chama 392, 394 Thracia 579, 617 Venus 560, 619 Venus var. rugatina 560, 619 Ruppellaria typica 552, 570, 576, 609 ruppelli, Chama 390 396, 398, 403, 404, 405, 408, 409, 410-412, 512, 550, 552-553, 556, 560, 562, 565-567, 570, 572-573, 575, 577, 579-580, 586, 594 sardo, Chama 570, 594 Sargassum 453, 486, 500 saulae, Palmadusta 138, 153 Saxicava 597 arctica 559 azaria 559 Saxidomus 443 sayanum, Pecten (Amusium) 560, 610 Pecten (Propeamussium) 560, 610 Propeamussium 519, 572, 610 sayi, Tellina 568, 614 scaber, Ctenoides 514, 572-573, 577, 598-599 scabra, Ctenoides 563 Lima 551-552, 556, 559, 566-567, 570-571, 574, 576-579, 581, 586, 589, 598-599 Lima scabra 570, 598 Lima form tenera 556, 588, 598 Lucina 601 Lucina (Lucina) 559, 601 Scalaria venosa 571 Scapharca 589 auriculata 571, 589 brasiliana 510, 591 chemnitzii 510, 591 inaequivalvis 571, 591 occidentalis 571, 590 (Scapharca) 590 (Scapharca) auriculata 560, 571, 589 (Scapharca) transversa 560 scarlatoi, Abrina 157-158, 159, 161-162, 162-163, 166-167 Schasichella alata 224 scheibei, Pseudochama 387, 411 Schilderia 127, 131-132, 137 achatidea 132, 135, 137-138, 152 schilderorum, Lyncina 135, 151 schmitti, Aurinia 571 schrammi, Anodontia 514, 599 Anodontia (Anodontia) 556 Lucina (Loripinus) 567 scindata, Mauritia maculifera 135, 150 Scissula 616 candeana 522, 615 consobrina 522, 615 iris 522, 615-616 sandix 615 similis 522, 530-535, 615 Scissurellidae 180 scotophila, Hyalina 20 Hyalina var. dilatata 21 Hyalinia var. попа 20 Scrobicularia plana 424 scurra, Mauritia scurra 135, 149 secticostata, Arca 567, 589 Seguenziidae 180 Semele 259, 263-264, 585, 612 bellastriata 520, 530, 547, 551, 554-555, 568-572, 580, 583-585, 588, 612 bellestriata 612 cancellata 559, 612 nuculoides 568, 584-585 obliqua 558-559, 579, 612 proficua 520, 554, 568-570, 572, 583-585, 612 purpurascens 520, 548-549, 554, 568-570, 572, 585, 612 radiata 569, 612 reticulata 559, 579, 612 (Semelina) 612 (Semelina) nuculoides 554 INDEX 669 Semelidae 157-158, 258, 520, 584-585, 611 Semelina nuculoides 520, 570, 572, 612 semen, Botula 579, 603 semiaspera, Diplodonta 559, 570, 572, 579, 584, 618 Phlyctiderma 523 semiaurita, Pedalion 566, 568, 598 Semibuliminus 84 Semierycina 548, 572, 596 semigranatus, Trichomusculus 296 seminuda, Atrina 186-187, 187, 519, 563, 569, 572, 578,610 Pinna 559 semiplota, Staphylaea 133, 135, 148 semisulcata, Papyridea 511, 557,570, 572, 576, 585-586, 592 semisulcatum, Cardium (Papyridea) 561, 567. 592 Papyridea 574, 592 semperi, Prosimnia 132-133, 147 senegalensis, Pseudozonaria pyrum 137 Zonaria pyrum 152 sentis, Caribachlamys 518, 531, 533, 553, 564-565, 572, 576, 586, 607 Chlamys 551, 556, 562-563, 565-567, 570, 575-578, 586-589, 608 Pecten 568-569, 585, 587 septus, Ophiophragmus 564 sepultus, Amphioplus 564 seraphinica, Aegistohadra 112 Helix 114, 118, 118-119 sericea, Helicina 224 serrata, Atrina 519, 572, 582, 610 Pinna 568 Serratovolva dondani 133, 147 serratum, Cardium 560, 592 Cardium (Laevicardium) 567, 592 Laevicardium 553, 571, 579, 592 Papyridea (Liocardium) 559, 592 serrulifera, Purpuradusta 138, 141, 155 setifera, Lima (Limatula) 559 Limatula 514, 539, 599 sherlyae, Zoila jeaniana 150 shiashkotanika, Abrina 157 Shinkailepas briandi 177, 177, 180-181 myojinensis 180 sibogai, Abrina 162, 166 sigsbeei, Pecten (Pecten) 560, 596 Pecten (Pseudamusium) 560, 596 Pectinella 513, 539, 596 simiaspera, Diplodonta 576 similaris, Bradybaena 83, 92, 111 similis, Pseudochama 390-391 Scissula 522, 530-535, 615 Spisula solidissima 568, 602 670 Tellina 302, 556, 564-565, 570-571, 573, 578, 583-585, 616 Tellina (Scissula) 554, 574 Simnia aequalis 147 simplex, Anomia 510, 559, 567, 570, 572, 578,565, 589 simpsoni, Cytherea 559, 567, 620 Pitar 302, 524, 531, 535, 339, 370, 572-573, 583-585, 620 sinistrorsa, Chama 390, 403 sinosa, Chama 551, 594 sinuosa, Chama 382, 388-389, 392, 396, 398, 399, 401-403, 411, 512, 556-557, 570, 572, 594 Skeneidae 173, 176, 180 smaragdina, Peltospira 175, 175, 181 smithii, Astarte 510, 559, 591 Carditopsis 512, 548, 570, 572, 594 sol, Tellina 571 Tellina (Phylloda) 614 Solecurtidae 520, 612 Solecurtus cumingianus 520, 572, 612 Solemya 584, 612 occidentalis 520, 568, 572, 584, 612 velum 520, 612 Solemyidae 520, 612 Solemyiidae 13 Solemyoidea 52 soleniformis, Paphridea 592 Papyridea 511, 533, 552, 556, 563, 567, 570, 572,575, 576, 007, 092 Solenomya occidentalis 559, 570 solida, Arcopsis 327, 336 Gari 258-260, 262, 272 solidifacta, Nuculana 517, 605 solidissima, Mactra var. similis 559, 602 solidula, Ledella 605 Nuculana 568, 572 Nuculana (Jupitaria) 577 sombrerensis, Lucina 547, 572, 576, 601 Lucina (Lucina) 559 Lucina (Pleurolucina) 556 Pleurolucina 515, 530, 533, 601, 622 soror, Diplodonta 559-560, 579, 618 Phlyctiderma 523 Taras 568 souleyetiana, Macoma tenta 567, 615 sowerbyi, Venus 441 spadicea, Neobernaya 135, 138, 152 Pustularia 138 Spathochlamys 607 benedicti 518, 572, 608 spathuliferus, Spondylus 559, 579, 613 speciosa, Eucrassatella 512, 533, 570, 572, 5767095 spectralis, Glycymeris 513, 597 Spengleria rostrata 278, 513, 552, 557, 570, 572, 597 Sphenia antillensis 572, 602 fragilis 515, 602 spinosa, Chama 398 Spinosipella 622 acuticostata 525, 621 spinosum, Cardium (Papyridea) 567, 592 Paphridea 587, 592 Spisula raveneli 516, 572, 602 solidissima similis 568, 602 Spondylidae 520, 612 spondyloidea, Plicatula 552, 610 Spondylus 553, 566, 571, 613 americanus 520, 530, 533, 552-553, 557, 568, 570-572, 580-581, 584, 587-588, 613 croceus 579, 613 echinatus 568, 613 gaederopus 556, 613 gussoni 570, 572, 613 ictericus 520, 530, 533, 562, 564, 570, 572. 576, 588,613 ramosus 571 spathuliferus 559, 579, 613 Sportellidae 521, 613 springeri, Anadara 548, 577, 581, 589 spurca, Erosaria 133, 149 squamifera, Tellina 522, 533, 559, 568, 570, 5753, 576; 616 Tellina (Phyllodina) 553 squamosa, Chama 392, 394 Lima 556, 559, 579, 598 Lucina 579, 600 Lucina (Lucina) 559, 600 squamosus, Modiolus 516, 535, 553, 583, 604 Modiolus modiolus 570, 572, 584-586, 604 staminea, Protothaca 305 Staphylaea 130-132 limacina interstincta 133, 135, 148 limacina limacina 133, 135, 148 semiplota 133, 135, 148 staphylaea laevigata 133, 135, 148 staphylaea staphylaea 133, 135, 148 staphylaea, Staphylaea staphylaea 133, 135, 148 stellae, Petricola 350 stentina, Ostreola 322 stercoraria, Trona 135, 151 sterna, Pteria 491 Stewartia 601 floridana 515, 602 Stilpnodiscus 81, 87, 97, 106-108 entochilus 86, 97, 112 moellendorffi 86, 97, 112 yeni 86 stimpsoni, Thracia 523, 551, 554, 559, 566, SAP 53, 617. Transenella 561, 564, 566, 573 Transennella 525, 549, 565, 568, 570, 584, 621 Stirpulina 37 stolida, Bistolida stolida 138, 141, 153 strangei, Humphreyia 42, 48, 50 strepta, Chama 387, 411 Striaciinae 327 Striarca 327, 332 lactea 327, 332, 334, 336 symmetrica 334 striata, Cardiomya 512, 533, 595 Cuspidaria (Cardiomya) 559 Martesia 519, 559, 567, 572, 582, 609 Tellina 559, 568, 614 striatula, Venus 296 Strigilla carnaria 522, 554, 559, 568, 570-571, 616 flexuosa 552, 556, 559, 568, 580, 616 gabbi 522, 554, 570, 572, 574, 616 mirabilis 522, 554, 570, 572, 583, 616 pisiformis 522, 549-550, 556, 559, 561, 566, 568, 571-572, 580, 586-587, 616 pisum 579, 616 rombergii 561, 568, 616 romgergi 570, 616 (Strigilla) 616 (Strigilla) gabbi 550, 577 strigillatum, Palliolum 549 Pseudamusium 549, 560 strigillatus, Cyclopecten 519, 610 strigillina, Antigona 550, 567, 571, 619 Antigona (Circomphalus) 566 Antigona (Ventricola) 566 Cytherea (Ventricola) 554, 561, 619 strigillinus, Circomphalus 441, 524, 548, 572, 619 Cytherea (Ventricola) 548 Striostrea 605 margaritacea 316 prismatica 320 Strombus gigas 242, 576 subauriculata, Lima (Limatula) 559 Limatula 514, 539, 599 subcancellata, Ervilia 520, 562, 572, 612 subcircularis, Anodonta 208 subequilatera, Astarte crenata 510, 581, 591 subglobosa, Diplodonta 559, 618 INDEX 671 sublaevis, Pustularia bistrinotata 135, 137-138, 152 sublevis, Leda 605 Ledella 517, 605 submissa, Trichobradybaena 103, 112 suborbicularis, Kellia 57-59, 63, 64-67, 71, 513, 572, 596 Муа 59 subrostrata, Chione (Chione) 561, 618 Venus (Chione) 568, 619 subrotundata, Kellia 58, 63, 64-66, 71 subsimilis, Laeocathaica (Laeocathaica) 86, 89, 112 subteres, Talostolida 138-139, 154 subviridis, Erronea subviridis 141 Erronea (Adusta) subviridis 155 sulcata, Limopsis 514, 572, 599 Modiola 556, 603 Modiola (Brachydontes) 559, 603 sulcatus, Modiolaria 567, 603 sulcidentata, Lyncina 135, 137, 151 summersi, Blasicrura 141, 143, 156 Sunetta 444 Sunettinae 444 surinamensis, Propustularia 133, 148 Sutilizona theca 173, 173, 180 Sutilizonidae 173, 180 swiftiana, Corbula 551, 560, 567, 572, 579, 594 swinhoei, Nesiohelix 85, 86, 86, 89, 94, 98, 108112 sybaritica, Tellina 570, 573, 584-585, 616 Tellina (Angulus) 554 sybariticum, Laevicardium 511, 570, 572, 585, 592 sybariticus, Angulus 521, 613 sylvicola, Hyalinia 21 symmetrica, Striarca 334 symplector, Bathymargarites 173, 174, 180 Syndosmya 611 lioica 549, 558 Syringodeum 245-246, 278, 418, 452-454, 498 filiforme 474, 487 tageliformis, Macoma 522, 572, 615 Tagelus 583, 586, 612 divisus 264, 520, 559-560, 567-568, 570, 572, 583, 585, 612 dombeii 264 gibbus 552, 612 plebeius 520, 570, 572, 581, 612 taitae, Cribrarula 128, 138, 140, 143, 154 takeoi, Dentiovula 132-133, 147 talamancensis, Helicina 217-218, 220, 222 672 Talostolida 131, 139 latior 138-139, 154 pellucens 138-139, 154 rashleighana 143 subteres 138-139, 154 teres 138-139, 154 teres alveolus 139 teres janae 139 teres teres 139 talpa, Talparia 133, 135-136, 151 Talparia 131, 136 exusta 133, 135-136, 151 talpa 133, 135-136, 151 tamiamensis, Periglypta 432 tampaensis, Angulus 521, 613 Tellina 565, 569-570, 573, 585, 616 Tellina (Angulus) 554 tantilla, Nutricola 444 Tapetinae 428 Taras 618 notata 568 nucleiformis 568 punctata 568 soror 568 tarquinia, Periglypta 432 tatarica, Abrina 157 Tawera 427 taylorae, Argopecten irradians 548, 570, 575, 582, 606 tayloriana, Tellina 579, 614 Tellidora cristata 522, 554, 559, 568, 570, 572, 583-584, 616 Tellin (Scissula) 615 Tellina 554-555, 559-560, 562, 576, 578, 581, 583-585, 588, 613-617 (Acorylus) gouldii 553, 613 (Acropagia) 614 (Acropagia) angulosa 580 (Acropagia) fausta 580 aequistriata 568, 570, 572, 584, 616 agilis 572,616 alternata 552, 559, 568, 570, 572-573, 583, 616 americana 570, 572, 616 anguilosa 573,614 angulosa 568, 570, 572,616 (Angulus) 613-615 (Angulus) candeana 566, 580 (Angulus) iris 566, 580 (Angulus) mera 554 (Angulus) paramera 549 (Angulus) probrina 549, 554 (Angulus) sybaritica 554 (Angulus) tampaensis 554 (Angulus) texana 554 INDEX (Angulus) versicolor 550, 554 (Arcopagia) 614 (Arcopagia) angulosa 566 (Arcopagia) Таиз 553 brasiliana 614 braziliana 556, 587, 614 candeana 550, 565, 568, 570, 572, 576, 587, 616 consobrina 572, 616 consorbrina 570, 615 cristallina 616 crystallina 568, 587, 615 (Cyclotellina) 614 (Cyclotellina) fausta 580 decora 556, 559, 568, 579, 615 (Elliptotellina) 614 (Elliptotellina) americana 553 (Eurytellina) 614 (Eurytellina) alternata 554 (Eurytellina) angulosa 549, 554, 561, 566, 580 (Eurytellina) nitens 554 (Eurytellina) punicea 549 fausta 556, 559, 562-564, 567-568, 570-572, 582, 586, 588, 614, 616 foliacea 263 georgiana 568, 614 gouldi 568, 613 gouldii 559, 570, 572-573, 579, 585, 613, 616 interrupta 559, 568, 571, 579, 617 iris 302, 556, 568, 573, 584-585, 616 (Laciolina) 614 (Laciolina) laevigata 553 (Laciolina) magna 553 laevigata 552, 570, 573, 587-588, 616 lineata 559, 565-568, 570-571, 573, 579, 616 lineata var. albida 579, 614 lintea 552, 568, 576, 615 listeri 567, 570, 573, 584, 586, 588, 616 magna 559, 565, 570, 573, 586, 616 martinicensis 568, 570, 573, 616 mera 302, 558-559, 568, 570, 573, 575-576, 578-579, 583-585, 616 cf. mera 565, 568, 613 (Merisca) 615 (Merisca) aequistriata 553 (Merisca) martinicensis 553 modesta 559, 568, 614 nitens 570, 573, 616 obliqua 549 paramera 553, 570, 573, 578, 616 persica 522, 573, 616 (Phylloda) sol 614 (Phyllodina) 616 (Phyllodina) squamifera 553 probina 613 probrina 553, 570, 573, 616 punicea 565, 573, 586, 616 radiala 582, 616 radiata 522, 551, 556, 559, 562, 564, 567-568, 570-571, 573, 579, 584, 586, 588, 616 radiata unimaculata 616 radiata var. unimaculata 571, 584, 586, 616 robusta 571 sandix 616 sayi 568, 614 (Scissula) 615 (Scissula) candeana 549, 554, 561, 566, 574, 580 (Scissula) exilis 561, 615 (Scissula) iris 561, 566, 580 (Scissula) sandix 615 (Scissula) similis 554, 574 similis 302, 556, 564-565, 567, 570-571, 573, 578, 583-585, 616 sol 571 squamifera 522, 533, 559, 568, 570, 573, 576, 616 striata 559, 568, 614 sybaritica 570, 573, 584-585, 616 tampaensis 565, 569-570, 573, 585, 616 tayloriana 579, 614 (Tellina) 616 (Tellina) radiata 553 (Tellinella) 617 (Tellinella) listeri 553 tenella 585, 617 tenera 556, 558-559, 568, 571, 613 tenuis 296 texana 570, 573, 575, 583-585, 617 versicolor 568, 570, 573, 576, 584, 617 Tellinella 616-617 listeri 522, 617 tellinella, Gari 258 Tellinidae 157, 164, 258, 521, 527, 557, 584-585, 613 Tellinoidea 258-259, 263, 271-272 tellinoidea, Cumingia 554-555, 588, 612 tellinoides, Cumingia 555, 559, 567, 579, 583-585, 611-612 tenella, Limopsis 599 Tellina 585, 617 tenellus, Angulus 521, 614 tenera, Barbatia 355, 365, 366-367, 369, 372, 374, 376, 377-378, 549, 569-570, 579, 591 INDEX 673 Barbatia (Fugleria) 365 Fugleria 510, 548, 591 Lima 559, 567, 576, 579, 587, 598-599 Lima scabra 570, 574, 576, 578, 588, 598 Periploma 560, 566, 568, 609 Tellina 556, 558-559, 568, 571, 613 tenerum, Periploma 518, 550, 572, 609 tenta, Macoma 522, 559, 567, 572, 585, 615 Macoma var. souleyetiana 559, 615 tenuis, Cyclinella 524, 567, 570, 572, 619 Ennucula 517, 538, 572, 605 Helicina 217-218, 219, 221 Lucinopsis 559 Tellina 296 teramachii, Nesiocypraea 130, 132, 140 Teredinidae 273, 522, 539, 580-581, 617 Teredo bartschi 523, 576, 617 clappi 523, 551, 568, 573, 617 thomsoni 559, 568, 617 (Zopoteredo) 617 (Zopoteredo) clappi 549, 552, 582 Teredora malleolus 523, 617 tereinus, Pecten 551, 563-564, 568, 608 Pecten (Euvola) 549, 554, 561, 566, 608 teres, Talostolida 138-139, 154 Talostolida teres 139 Teskeyostrea 310 weberi 309-312, 315, 317, 320, 321, 517, 548, 564, 566, 605-606 tessellata, Luria 133, 135, 151 testudinaria, Chelycypraea 135, 151 testudinum, Thalassia 439, 474, 486-487 teulerei, Barycypraea 132, 135-136, 150 texana, Tellina 570, 573, 575, 583-585, 617 Tellina (Angulus) 554 texanus, Angulus 521, 614 texasiana, Agriopoma 622 texasianus, Donax 459, 596 Thais carinifera 295 haemastoma 296 Thalassia 245-246, 278, 292, 418, 451-455, 457-458, 486-487, 492-493, 497-500, 540, 555 testudinum 439, 474, 486-487 thalassinum, Parvamussium 519, 610 thalassinus, Cyclopecten 539 Pecten (Pecten) 560 Pecten (Pseudamusium) 560 Thalassiosira 8 theca, Sutilizona 173, 173, 180 thersites, Zoila 143, 150 Zoila friendii 135 674 thielei, Eclogavena quadrimaculata 141, 156 thomasi, Erosaria 133, 149 thomsoni, Teredo 559, 568, 617 Thracia corbuloidea 558-559, 617 corbuloides 550, 566, 568, 573,617 distorta 523, 617 morrisoni 523, 617 phaseolina 523, 559, 573, 617 rugosa 579, 617 stimpsoni 523, 551, 554, 559, 566, 571, 573, 617 Тргасйдае 523, 584-585, 617 Thyasira grandis 523, 539, 617 trisinuata 523, 573, 617 Thyasiridae 12, 523, 617 tiara, Placamen 443 tigerina, Lucina 558, 571, 600 Lucina (Lucina) 559, 600 tigrina, Lucina 574, 579, 600 (Lucina) pecten 559 tigris, Cypraea 133, 135, 150 Timoclea 427, 443, 619 grus 525, 573, 621 pygmaea 525, 573, 621 tincta, Protocardia 561, 566, 592 tinctum, Nemocardium 511, 572, 592 Tishoplita 109 Tivela 443—444 abaconis 525, 621 floridana 525, 538, 573, 621 mactroides 525, 550, 552, 566, 568, 580, 621 trigonella 525, 621 Tivelina newtoni 296 tokioi, Phenacovolva 133, 147 tokuoi, Crenavolva 133, 147 tongiorgii, Oxychilus (Ortizius) 21 toreuma, Globivenus 428, 443 Venus 443 Trachidontes 565, 603 Trachycardium 565, 576, 585, 591, 593 (Acrosterigma) 591 (Acrosterigma) magnum 550, 577 egmontianum 511, 556-557, 564, 570, 573 578.584. 592 isocardia 552, 592 magnum 549-550, 552, 557, 565, 567, 510; 573. 575.577. 566: 593 muricatum 511, 531, 533-535, 556-557, 564, 570; 573, 578,583, 585, 593 Transenella 555, 588, 621 conradina 566, 568, 573, 587, 621 cubaniana 562, 566, 568, 573, 621 culebrana 573 INDEX stimpsoni 561, 564, 566, 573 Transennela 621 Transennella 554, 570, 583, 621 conradina 525, 580, 584, 621 cubaniana 525, 550, 563, 565, 570, 574, 586, 621 culebrana 525, 550, 621 stimpsoni 525, 549, 565, 568, 570, 584, 621 transversa, Anadara 510, 571, 576, 590 Arca 551, 567, 577, 579-580 Arca (Scapharca) 559 Scapharca (Scapharca) 560 trapezia, Anadara 372, 374 Trapezidae 12, 523, 618 Tricheulota 83 Trichia 73-74, 77 Trichiinae 77 Trichobradybaena 81, 85, 90, 106-109 submissa 103, 112 Trichomusculus semigranatus 296 Trichomya 278 hirsuta 292 tridentata, Pleuromeris 302, 511, 564, 570, 572-573, 577, 581, 584-585, 593 Venericardia 576, 583 trigonella, Tivela 525, 621 Trigoniocardia (Americardia) 591 (Americardia) media 557 (Americardia) medium 591 antillarum 511, 593 medium 591 Trigonocardia (Americardia) medium 580 medium 583 Trigonulina 622 ornata 525, 573, 621 triquetra, Parastarte 524, 554-555, 561, 565. 568, 572, 580, 583, 5881620 Trishoplita 81, 106-109 dacostae 89, 94, 98, 100, 112 trisinuata, Thyasira 523, 573, 617 trisulcata, Linga 583, 600 Lucina 572, 600 Lucina (Anodontia) 567, 600 Lucina (Lucina) 559, 599-600 Triviidae 132 trizonata, Notadusta punctata 141, 143, 156 Trochidae 173, 180 Trona 131 stercoraria 135, 151 trossulus, Mytilus 208 truncata, Barnea 519, 570, 572, 579-580, 584, 609 Laternula 48 Pholas (Barnea) 559, 568 Truncatella pulchella 328 trunculus, Donax 208, 469 Tucetona 597 pectinata 302, 513, 530-535, 573, 597 pectinatus 597 pectunculus 597 tulipa, Modiola 556, 604 Modiola var. nigra 548, 556, 604 tulipus, Modiolaria 567, 604 Modiolus 568, 604 tumidus, Donax 567, 596 Polinices 296, 305 tumulosa, Chama 398 turbinata, Helicina 224 turdus, Erosaria 133-134, 148 Turtonia minuta 5, 12 typica, Choristodon 350 Ruppellaria 552, 570, 576, 609 typicum, Choristodon 579, 609 umbellifera, Hyalogyrina 179, 179 Umbilia 130-131, 134, 139 armeniaca 133, 135, 149 capricornica 133-135, 149 hesitata 133, 135, 149 petilirostris 134-135 cf. petilirostris 133-135, 149 umbonata, Arca 553, 560, 565-567, 580-581, 583, 590 Arca (Lunarca) 560, 590 Arca (Navicula) 579-580, 590 Umbonium vestiarium 296 undata, Glycymeris 513, 570, 572, 576, 597 Leptaxis 73, 75, 75-76 undatella, Chione 301 undatus, Pectunculus 559, 597 Ungulinidae 523, 584, 618 unifasciata, Purpuradusta fimbriata 142, 156 Purpuradusta cf. fimbriata 138, 141 unimaculata, Tellina radiata 616 Unionidae 205, 208 Urguessella 84 ursellus, Bistolida 138-139, 141, 153 uzielli, Oxychilus 19, 22 uziellii, Oxychilus 19-20, 23, 30-34 ‚ Zonites 20-21 vaginiferus, Brechites 37-38, 41-42, 45, 47-48, 50, 52 valentia, Leporicypraea 135, 149 valvatoides, Protolira 172, 173, 179 vanhyningi, Cumingia 520, 531, 534-535, 550, 612 Cumingia tellinoides 548, 572, 577, 581, 612 INDEX 675 variabilis, Donax 459-461, 462-463, 467-470, 468, 513, 552, 559, 567, 572, 579, 596 Latona 459 Provanna 177, 178 Varicorbula disparilis 512, 573, 595 krebsiana 512, 539, 595 limatula 573, 595 operculata 584-585, 595 philippii 512, 573, 595 variegata, Amphidesma 548 Chama 401-402, 587, 594 Chama radians 401-402 Pseudochama radians 401, 567, 594 vasta, Bistolida owenii 141 veerrucosa, Dosina 440 veerruicosa, Venus 440 veitchi, Brechites (Foegia) 38 velata, Arca 574, 590 vellicata, Kuia 444 velum, Solemya 520, 612 velvetina, Leiomya (Plectodon) granulata 566, 596 Venericardia 593 tridentata 576, 583 Veneridae 12, 15, 247, 258, 277, 295, 427, 429, 523, 527, 584-585, 618 Venerinae 427-429 Veneroidea 429, 459 Venerupis 443 pullastra 444 venosa, Scalaria 571 Ventricolaria 441,619, 621 rigida 550, 562, 570, 574 rugatina 564 (Ventricolaria) 619 (Ventricolaria) rigida 577 Ventricoloidea 427 ventricosa, Meoma 564 ventriculus, Lyncina 135-137, 151 Venus 427-428, 441, 560, 618-621 aegrota 441 (Anomalocardia) rostrata 559, 618 antiqua 305 beaui 559, 579, 618 campechiensis 568 cancellata 558-559, 618 chemnitzii 441 (Chione) 620-621 (Chione) cancellata 560, 618 (Chione) cancellatus 568, 618-619 (Chione) granulatus 568, 619 (Chione) grus 568 (Chione) latiliratus 568, 620 (Chione) mazyckii 568 676 (Chione) paphia 568 (Chione) pubera 568 (Chione) pygmaeus 568, 621 (Chione) subrostrata 568, 619 (Chione [Omphaloclathrum]) listeri 429 clathrata 441 crispata 429, 441, 559, 620 granulata 560, 579, 618 (Haliris) fischeriana 560 lacerata 441 lamarckii 560, 618 laqueata 441 listeri 429, 441, 579 magnifica 440-441 mercenaria 559, 561, 568 тегсепапа var. mortoni 559, 620 mercenaria var. notata 561, 620 monilifera 441 mortoni 579, 620 multicostata 441 paphia 579 (Periglypta) listeri 429 pilula 560, 619 puerpera 429, 440-441 purpurascens 549 pygmaea 559, 579 pygmaea var. inaequivalvia 579, 621 resticulata 441 reticulata 440-441 rugosa 560, 619 rugosa var. rugatina 560, 619 sowerbyi 441 striatula 296 toreuma 443 veerruicosa 440 verrucosa 428, 440, 443 venusta, Vesicomya 622 Zoila 135, 150 venustus, Donax 469 verginica, Ostrea 568, 605 verrilliana, Nuculana 517, 550, 566, 568, 572-573, 605 verrucosa, Venus 428, 440, 443 verrucosus, Calpurnus 133, 147 versicolor, Angulus 521, 614 Tellina 568, 570, 573, 576, 584, 617 Tellina (Angulus) 550, 554 verticillata, Caulerpa 499 Verticordia 621 acuticostata 558, 560, 573, 622 elegantissima 622 fischeriana 622 (Haliris) fischeriana 621 ornata 568, 622 INDEX (Trigonulina) 621 (Trigonulina) ornata 559 Verticordiidae 1, 12-15, 525, 621 vesica, Vesicomya 525, 622 Cytherea (Veneriglossa) 559 Vesicomya venusta 622 vesica 525, 622 Vesicomyidae 13, 525, 622 vespacea, Palmadusta asellus 138-139 Palmadusta cf. asellus 153 vespertina, Gari 262 vestiarium, Umbonium 296 Vicarihelix 84 vinosa, Lepsiella 296 violascens, Asaphis 249-251, 253, 253-255, 259-261, 263-265, 268-272, 610 virginica, Crassostrea 310, 312, 314-316, 318, 319, 322-323, 517, 557 9729518! 581, 605 Ostrea 559 virginicus, Anisotremus 423 viridis, Leporicypraea mappa 135-136, 149 Perna 528, 553, 604 viridizona, Pteria 476, 491 viridozona, Pteria 476 vitellus, Lyncina 135, 151 vitrea, Eulepetopsis 170-171, 171, 180 Leda (Leda) 559 Nuculana 517, 539, 605 Pteria 473, 491, 520, 550, 571, 575,611 Pteria hirundo 564, 566, 611 vitreum, Dacrydium 559, 567, 578, 603 vittatus, Donax 296 vivia, Blasicrura cf. pallidula 141 Volsella americana 583, 604 Volva 132 volva 132-133, 147 volva, Volva 132-133, 147 vredenburgi, Erronea 141-142, 155 vulgaris, Pinctada 485, 491-492 vulgata, Patella 212, 216 waikikiensis, Purpuradusta fimbriata 138, 141-142, 156 walkeri, Contradusta 141, 156 weaveri, Phenacovolva 133, 147 weberi, Abrina 162, 166 Ostrea 310, 548, 550, 574 Teskeyostrea 309-312, 315, 317, 320, 321, 517, 548, 564, 566, 605-606 weyrichii, Karaftohelix 84, 86, 94, 111 willcoxii, Chama 386-387, 396, 411 woodiana, Anodonta 205-208, 206-207 xandaros, Sahlingia 171, 172 xanthia, Pinctada 476, 486 Pteria 476, 491 xanthodon, Erronea 141-142, 155 Xanthonychidae 84, 109 Xenophora conchyliophora 564 Xenostrobus inconstans 296 pulex 296 Xeropicta krynickii 74 Xylodiscula analoga 178, 178, 179 Xylodisculidae 178-179, 181 Xylophaginidae 273 Xylopholas altanai 609 altenai 609 Xyloskenea costulifera 176 Xylotrya fimbiata 556, 617 Xylotrya fimbriata 559, 568, 617 yantaiensis, Metodontia 85, 92, 111 yeni, Stilonodiscus 86 Yoldia liorhina 525, 539, 622 Yoldiella 605 messanensis 605 Yoldiidae 525, 622 zebra, Arca 356, 369, 377, 510, 530-533, 535, 562-566, 569-570, 572, 574, 577-580, 583-586, 588-590 Arca (Arca) 551 Macrocypraea 135, 149 zelandiae, Penicillus novae 39 zelandica, Dosina 444 ziczac, Euvola 518, 572-573, 577, 608 Palmadusta 139 Palmadusta ziczac 138, 153 INDEX 677 Pecten 564, 568, 570, 608 Pecten (Janira) 559 Zoila 127, 130-132, 134, 139 decipiens 135-136, 150 eludens 135, 150 тепай 136 friendii friendii 136 friendii jeaniana 135 friendii thersites 135 jeaniana jeaniana 150 jeaniana sherlyae 150 marginata 136 marginata ketyana 135-136, 150 marginata marginata 135-136, 150 mariellae 135-136, 150 orientalis 143 perlae 143 rosselli 135, 150 thersites 143, 150 venusta 135, 150 Zonaria 131-132, 137 angelicae 135, 138, 143 petitiana 143 picta 135, 137-138, 152 pyrum 135, 138 pyrum angelicae 152 pyrum angolensis 152 pyrum senegalensis 152 sanguinolenta 135, 138, 152 zonaria 135, 138, 152 zonaria, Zonaria 135, 138, 152 Zonitidae 19 Zonites uziellii 20-21 Zostera marina 486 и ож Е; MALACOLOGIA International Journal of Malacology Bivalve Studies n the Florida Keys Proceedings of the International Marine Bivalve Workshop | Long Key, Florida, July 2002 — Edited by Rudiger Bieler and Paula М. Mikkelsen Vol. 46(2) | [à | 2004 > if | | | Vol. Vol. Vol. Vol. Vol. Vol. Vol Vol. Vol. Vol. Vol Vol. Vol. Vol. Publication dates 37, No. 37, No. 38, No. 39, No. 40, No. 41, No. . 41, No. 42, No. 43, No. 44, No. . 44, No. 45, No. 45, No. 46, No. 13 Nov 8 Mar. 17 Dec. 13 May 17 Dec. 22 Sep. 31 Dec. 18 Oct. 20 Aug. 8 Feb. 30 Aug. 29 Aug. 22 Mar. 23 Aug. 1995 1996 1996 1998 1998 1999 1999 2000 2001 2002 2002 2003 2004 2004 VOL. 46, МО. 2 MALACOLOGIA CONTENTS RÜDIGER BIELER, ISABELLA KAPPNER & PAULA M. MIKKELSEN Periglypta listeri (J. E. Gray, 1838) (Bivalvia: Veneridae) in the Western Atlantic: Taxonomy, Anatomy, Life Habits, and Distribution ......................... RÜDIGER BIELER & PAULA M. MIKKELSEN Marine Bivalves of the Florida Keys: À Qualitative Faunal Analysis Based on Origi- nal Collections, Museum Holdings and Literature Data ..................... GREGORIO BIGATTI, MELITA PEHARDA & JOHN TAYLOR Size at First Maturity, Oocyte Envelopes and External Morphology of Sperm in Three Species of Lucinidae (Mollusca: Bivalvia) from Florida Keys, U.S.A....... MATTHEW R. CAMPBELL, GERHARD STEINER, LYLE D. CAMPBELL & HERMANN DREYER Recent Chamidae (Bivalvia) from the Western Atlantic Ocean ................ OSMAR DOMANESCHI & ELIZABETH K. SHEA Shell Morphometry of Western Atlantic and Indo-West Pacific Asaphis; Functional Morphology and Ecological Aspects of А. deflorata from Florida Keys, U.S.A. (Вмамазеъатитовнаае): Etes MP Ada Aoi NE LISA KIRKENDALE, TAEHWAN LEE, PATRICK BAKER & DIARMAID О FOIGHIL Oysters of the Conch Republic (Florida Keys): А Molecular Phylogenetic Study of Parahyotissa mcgintyi, Teskeyostrea weberi and Ostreola equestris........... PAULA M. MIKKELSEN & RÜDIGER BIELER Critical Catalog and Annotated Bibliography of Marine Bivalve Records for the ОО ЕО о Re re eee car ce a PAULA M. MIKKELSEN & RÜDIGER BIELER International Marine Bivalve Workshop 2002: Introduction and Summary ...... PAULA M. MIKKELSEN, ILYA TÉMKIN, RÜDIGER BIELER & WILLIAM G. LYONS Pinctada longisquamosa (Dunker, 1852) (Bivalvia: Pteriidae), an Unrecognized PearlOysterinithe WesternvAtlantic: ao ER ее: BRIAN MORTON & MARTINA KNAPP Predator-Prey Interactions between Chione elevata (Bivalvia: Chioninae) and Naticarius canrena (Gastropoda: Naticidae) in the Florida Keys, U.S.A. ........ P. GRAHAM OLIVER & JOHANNA JARNEGREN How Reliable is Morphology Based Species Taxonomy in the Bivalvia? A Case Study on Arcopsis adamsi (Bivalvia: Arcoidea) from the Florida Keys .... LUIZ RICARDO L. SIMONE 8 ANTON CHICHVARKHIN Comparative Morphological Study of Four Species of Barbatia Occurring on the Southern Florida Coast (Arcoidea, Arcidae) ............................. LUIZ RICARDO L. SIMONE 8 JOANNE R. DOUGHERTY Anatomy and Systematics of Northwestern Atlantic Donax (Bivalvia, Veneroidea, DONA o oca Al de OOO AE UTA ee oe JOHN D. TAYLOR, ЕМШУ GLOVER, МЕНТА PEHARDA, GREGORIO BIGATTI & ALEX BALL Extraordinary Flexible Shell Sculpture: the Structure and Formation of Calcified Periostracal Lamellae in Lucina pensylvanica (Bivalvia: Lucinidae) ........... PAUL VALENTICH-SCOTT 8 GRETE ELISABETH DINESEN Rock and Coral Boring Bivalvia (Mollusca) of the Middle Florida Keys, U.S.A. ... 2004 427 503 417 249 473 295 327, 459 339 MALACOLOGIA SUBSCRIPTION AND PAST ISSUE ORDER FORM Name: Address: Personal rates: Per volume Subscription $56.00 Single volumes $56.00 Institutional rates: Subscription $75.00 Single volumes $75.00 Postage & handling per volume for single/past volumes $5.00 ¢ Subscriptions begin with the current volume. Past volumes are available at single volume rate with the exception of volumes 17(1) and 18 which are out of print. + Prepayment is required. 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Students will receive a dis- count and be charged U.S. $30.00. Institu- tional, agency or dealership subscriptions are U.S. $75.00. Back issues and single volumes: $56.00 for individual purchas- ers; $75.00 for institutional or agency pur- chasers. There is a $5.00 handling charge per volume for all purchases of past vol- umes. Address inquiries to the Subscrip- tion Office. Electronic submission is desired. Regular subscribers are those who have paid- up subscriptions for the current issue and the following issue. Students (including individuals submitting dissertations) must identify themselves at the time of manuscript submission and also provide the e-mail address of their advisor. VOL. 46, МО. 2 MALACOLOGIA CONTENTS PAULA M. MIKKELSEN & RÜDIGER BIELER International Marine Bivalve Workshop 2002: Introduction and Summary . ..... OSMAR DOMANESCHI & ELIZABETH K. SHEA Shell Morphometry of Western Atlantic and Indo-West Pacific Asaphis; Functional Morphology and Ecological Aspects of А. deflorata from Florida Keys, U.S.A. (Bivalvia: Psammobiidae): 02m. as RER OR PE PEER JOHN D. TAYLOR, EMILY GLOVER, MELITA PEHARDA, GREGORIO BIGATTI & ALEX BALL Extraordinary Flexible Shell Sculpture: the Structure and Formation of Calcified Periostracal Lamellae in Lucina pensylvanica (Bivalvia: Lucinidae) ........... BRIAN MORTON & MARTINA KNAPP Predator-Prey Interactions between Chione elevata (Bivalvia: Chioninae) and Naticarius canrena (Gastropoda: Naticidae) in the Florida Keys, U.S.A......... LISA KIRKENDALE, TAEHWAN LEE, PATRICK BAKER & DIARMAID O FOIGHIL Oysters of the Conch Republic (Florida Keys): A Molecular Phylogenetic Study of Parahyotissa mcgintyi, Teskeyostrea weberi and Ostreola equestris........... P. GRAHAM OLIVER & JOHANNA JARNEGREN How Reliable is Morphology Based Species Taxonomy in the Bivalvia? A Case Study on Arcopsis adamsi (Bivalvia: Arcoidea) from the Florida Keys .... PAUL VALENTICH-SCOTT & GRETE ELISABETH DINESEN Rock and Coral Boring Bivalvia (Mollusca) of the Middle Florida Keys, U.S.A. ... LUIZ RICARDO L. SIMONE & ANTON CHICHVARKHIN Comparative Morphological Study of Four Species of Barbatia Occurring on the Southern Florida Coast (Arcoidea, Arcidae) ............................. MATTHEW R. CAMPBELL, GERHARD STEINER, LYLE D. CAMPBELL & HERMANN DREYER Recent Chamidae (Bivalvia) from the Western Atlantic Ocean ............... GREGORIO BIGATTI, MELITA PEHARDA & JOHN TAYLOR Size at First Maturity, Oocyte Envelopes and External Morphology of Sperm in Three Species of Lucinidae (Mollusca: Bivalvia) from Florida Keys, U.S.A....... RUDIGER BIELER, ISABELLA KAPPNER & PAULA M. MIKKELSEN Periglypta listeri (J. E. Gray, 1838) (Bivalvia: Veneridae) in the Western Atlantic: Taxonomy, Anatomy, Life Habits, and Distribution ......................... LUIZ RICARDO L. SIMONE & JOANNE R. DOUGHERTY Anatomy and Systematics of Northwestern Atlantic Donax (Bivalvia, Veneroidea, Donacidaë). 3. ds 000% sure sen ee hee a a ee PAULA M. MIKKELSEN, ILYA TEMKIN, RUDIGER BIELER & WILLIAM G. LYONS Pinctada longisquamosa (Dunker, 1852) (Bivalvia: Pteriidae), an Unrecognized Pearl Oyster in the Western Atlantic ......--- ооо ne oes see ee RUDIGER BIELER & PAULA M. MIKKELSEN Marine Bivalves of the Florida Keys: A Qualitative Faunal Analysis Based on Origi- nal Collections, Museum Holdings and Literature Data ..................... PAULA M. MIKKELSEN & RÜDIGER BIELER Critical Catalog and Annotated Bibliography of Marine Bivalve Records for the Florida Keys... cs 302000 bale 94 nr OR SR EE 2004 249 295 417 427 459 473