SCANNING ELECTRON MICROSCOPIC AIDS FOR IDENTIFICATION OF LARVAL AND POST-LARVAL BIVALVES.
The identification of bivalve larvae and early postlarvae has been important for many ecological research efforts in marine, estuarine, and freshwater environments for over a century (Stafford 1912, Odhner 1914, Lebour 1938, Werner 1939, Jorgensen 1946, Sullivan 1948, Rees 1950, Miyazaki 1962, Loosanoff & Davis 1963, Newell & Newell 1963, Loosanoff et al. 1966, Le Pennec & Lucas 1970, Chanley & Andrews 1971, de Schweinitz & Lutz 1976, Lutz & Jablonski 1978a, 1978b, 1979, 1981, Jablonski & Lutz 1980, Le Pennec 1980, Lutz et al. 1982a, 1982b, Lutz 1988, 2012, Fuller & Lutz 1989, Kennedy etal. 1989,1991, Goodsellet al. 1992,Huetal. 1992,1993,Lutz& Kennish 1992, Baldwin et al. 1994, Hare et al. 2000, Garland & Zimmer 2002, Tiwari & Gallager 2003a. 2003b, Hendriks et al. 2005, Larsen et al. 2005, 2007, Wang et al. 2006, North et al. 2008, Henzler et al. 2010, Thompson et al. 2012a, 2012b, Malchus & Sartori 2013, Goodwin et al. 2014, 2016a, 2016b). As Hendriks et al. (2005) emphasized, "Despite the importance of the planktonic larval stage in intertidal bivalves, our understanding of this stage is still insufficient. A major obstacle in the quantification of planktonic larval distributions is the identification of sampled larvae."
In early efforts to assist with the identification of larval bivalves isolated from plankton samples. Chanley and Andrews (1971) published a key to the larval stages of a number of bivalve species along the east coast of North America based on optical micrographic sequences of articulated shells of laboratory-cultured larval specimens; however, the usefulness of this key is limited because of the great morphological similarity of the imaged articulated shells, particularly at the early (straight-hinge) developmental stages. Although a number of techniques have been refined in recent years and show promise for use in routine identifications of larval and post-larval bivalves (e.g., single-step nested multiplex polymerase chain reaction; in situ hybridization protocols through color coding with taxon-specific, dye-labeled DNA probes; coupled fluorescence in situ hybridization and cell sorting; and image analysis techniques using species-specific shell birefringence patterns under polarized light; Larsen et al. 2005, 2007, Henzler et al. 2010, Thompson et al. 2012a, 2012b, Goodwin et al. 2014, 2016a, 2016b, 2018), no adequate comprehensive reference source exists that accurately depicts the morphology and morphometry of the shells of larval and post-larval stages of target bivalve species in a consistent format to assist in identification of such stages.
With the aforementioned as background, in this monograph scanning electron micrograph (SEM) sequences of the disarticulated shell valves of laboratory-reared larval and post-larval stages of 56 species of bivalve molluscs from a wide spectrum of aquatic habitats are presented (Table 1). Emphasis is placed on the usefulness of the morphology and morphometries of consistently-oriented, disarticulated shell valves and associated hinge structures in discriminating the early life-history stages of these species. Most of these species are from environments along the east coast of North America and include most of the commercially important species in this region.
Taxonomic nomenclature was assigned according to the latest "accepted name" (or acceptable "alternate representation") and associated classification hierarchy in the World Register of Marine Species (www.marinespecies.org).
Over the years, various workers have used both optical and scanning electron microscopy to describe in detail the larval and/or post-larval hinge structures of a number of bivalves and have suggested that such structures may be diagnostic at the generic, or even specific, level (Chanley 1965, 1969, Turner & Johnson 1970. Pascual 1971. 1972, Scheltcma 1971, Le Pennec 1973, 1978. 1980. LaBarbera 1975, Boyle & Turner 1976. Culliney & Turner 1976, Dinamani 1976, Le Pennec & Masson 1976, Booth 1977, 1979a, 1979b, Siddall 1977, 1978, Lutz & Jablonski 1978a, 1978b, 1981, Carriker & Palmer 1979, Lutz & Hidu 1979, Chanley & Dinamani 1980, Jablonski & Lutz 1980, Lutz et al. 1982a, 1982b, Redfearn 1982, 1987, Jablonski & Lutz 1983. Ramorino & Campos 1983, Redfearn et al. 1986, Tremblay et al. 1987, Fuller & Lutz 1989, Fuller et al. 1989a, Kennedy et al. 1989, 1991, Hu et al. 1993, Paugam et al. 2006, Wassnig & Southgate 2012). Despite these efforts, much of the morphologic and morphometric data obtained over the years has not been presented in an adequate or sufficiently consistent format to permit unambiguous identification of early life-history stages of bivalves at various taxonomic levels. In recognition of this shortcoming, the SEM sequences of the larval and post-larval stages of a number of the species depicted in this monograph that have been previously published in various journals are presented here in a consistent format, together with other pertinent details related to procurement, preparation, and descriptions of the specimens comprising these sequences (Lutz et al. 1982b, Fuller & Lutz 1989, Fuller et al. 1989b, Kennedy et al. 1989, 1991, Goodsell et al. 1992, Gustafson & Lutz 1992, Hu et al. 1993, Tan et al. 1993). These sequences and pertinent details are included herein, together with a large number of unpublished sequences and associated pertinent details of most of the species in Table 1. The goal is to present a coherent compilation of all 56 species in one single publication to assist in the discrimination of larval and post-larval stages of these species isolated from a variety of aquatic environments.
MATERIALS AND METHODS
Sexually mature adults of 56 species of bivalves were obtained from the sources indicated in Table 1. These adults were induced to spawn (or larvae/juveniles were obtained from spontaneous spawning/release events in the case of certain species with nonplanktotrophic modes of development) using a variety of protocols described by various workers (Loosanoff & Davis 1963, Morse etal. 1977, Lutz et al. 1982a, 1982b, Gibbons & Castagna 1984, Fuller & Lutz 1989, Fuller et al. 1989a, Kennedy et al. 1991, Gustafson & Lutz 1992). The larvae and postlarvae of most of these species were reared using standard hatchery techniques (e.g., techniques described by Loosanoff & Davis 1963. Turner & Johnson 1970, Chanley & Andrews 1971, Castagna 1975, Castagna & Kraeuter 1977). Further specific details concerning the culture of the larvae and postlarvae of a number of these species may be found in Lutz et al. (1982b), Fuller and Lutz (1989), Fuller et al. (1989a), Kennedy et al. (1989). (1991), Goodsell et al. (1992), Gustafson and Lutz (1992). Hu et al. (1993). and Tan et al. (1993).
Cultured larval and post-larval specimens were sampled at frequent intervals (frequency dependent on the growth of organisms since the previous sampling period) and placed in distilled water for 30 min (Calloway & Turner 1979). Immediately following this treatment, specimens were preserved in 95% ethanol. After various lengths of time (up to 2 months), specimens were removed from the ethanol, rinsed in distilled water, and immersed in a 5% solution of sodium hypochlorite (Rees 1950) for approximately 10 min to facilitate separation of shell valves and removal of soft tissue. After rinsing in distilled water, disarticulated valves were mounted on copper or silver tape (or double-sided sticky tape), coated (under vacuum) with approximately 400 [Angstrom] of gold-palladium or a combination of gold and carbon, and examined under an ETEC Autoscan scanning electron microscope [or, in the case of Ostrea stentina (= Ostrea equestris), Solemya velum and Mercenaria campechiensis specimens, under Hitachi S-540 and JEOL 848 scanning electron microscopes].
Procedures used for accurate documentation of shapes and dimensions of the larval and post-larval shells using scanning electron microscopy were those of Fuller et al. (1989b) and are outlined below.
Before imaging individual larval or post-larval specimens under the scanning electron microscope, great care was taken to adjust the microscope so that x and y dimensions were equal on a calibration sphere that was approximately the same size as the specimen being photographed. In turn, these adjustments were made at a magnification close to that at which the specimen was to be photographed. The calibration spheres were sand-blasting beads that were selected for roundness by comparing measurements of the diameter on electron micrographs taken at rotations of 0, 45, and 90 degrees (see Fuller et al. 1989b for further details).
The method used for consistent orientation of the disarticulated shell valves, in which each larval or post-larval valve is positioned with points of the hinge and shell margin aligned in a plane normal to the axis of the electron optical system, is described by Fuller et al. (1989b, p. 59) as follows. "Specifically, a disarticulated valve with the interior shell surface visible on the microscope screen is rotated until the anterior and posterior margins are at equal working distances. A digital voltmeter (monitoring the reference voltage of the lens control) is used to measure carefully the differences in working distance when opposite margins of the shell are successively focused at 30,000x. A difference of 1 mV on the meter is equal to a change in working distance of about 0.34 [micro]m. Subsequently, the specimen is tilted perpendicularly to the first axis until the dorsal and ventral margins of the valve also are at the same working distances. A photomicrograph of the shell in this position documents its characteristic shape."
It should be noted that it was necessary to modify the aforementioned procedure to obtain consistent orientations for later post-larval stages of a number of species because of the irregular contours of the shell margins of these stages. In the case of post-larval Teredo navalis, for example, orienting the specimens in a manner similar to that described previously was impossible because points along the post-larval shell margin do not lie in a single plane. As articulated by Fuller et al. (1989a, p. 25) concerning T. navalis: Throughout the post-larval developmental period, however, points along the dorsoventral margin of the anterior slope (except those at the extreme ventral region) comprise a plane. Thus, consistent orientation of post-larval shells was achieved by positioning specimens such that this plane was perpendicular (Fuller et al. 1989a mistakenly indicated "parallel") to the electron optical axis. Additional adjustments were made so that dorsal and ventral condyles were at an equal working distance. Similar adjustments were also used to obtain consistent orientations of post-larval specimens of Bankia gouldi (Tan et al. 1993).
The dimensions of the larval and post-larval shells were determined by positioning a flat 400-mesh copper transmission electron microscope grid (on the same specimen mount, near the shell valve) normal to the electron optical axis and photographing this grid at the identical magnification at which the shell valve was photographed. Measurements of the shell dimensions are based on the 63.5-[micro]m grid spacings of the 400-mesh grid, rather than on magnification or scale bar displays on the microscope screen (for further details, see Fuller et al. 1989b).
The numbers depicted above each of the micrographs in Figures 1-195 indicate the maximum linear distance in micrometers measured along any axis of the shell, with a few exceptions. In most cases, this maximum distance represents "shell length" as defined by numerous authors (Fuller & Lutz 1989, Kennedy et al. 1989, 1991, Goodsell et al. 1992, Gustafson & Lutz 1992, Hu et al. 1993), although in the case of the larval stages of certain pholads and teredinids [e.g., Bankia gouldi (Tan et al. 1993) (Figs. 69 and 70) and Teredo navalis (Fuller et al. 1989a) (Figs. 73 and 74)], this maximum distance represents "shell height." Shell nomenclature of the teredinids is taken from Turner (1966, 1971). The numbers depicted above the larval stages of Crassostrea gigas, Crassostrea virginica, Ostrea edulis, and Ostrea stentina (= Ostrea equestris) (Figs. 105-123) represent the maximum anteroposterior dimension ("shell length", which in some larval stages of these species is less than or equal to "shell height"; see Hu et al. 1993).
We use the term "provinculum" in the sense of Bernard (1898) and Rees (1950). Provinculum length represents the linear distance between the lateral extremes of the hinge apparatus in larval and early post-larval shells (see Bayne 1976, p. 87, for a diagrammatic illustration of this dimension).
RESULTS AND DISCUSSION
An inability to identify bivalve larvae and postlarvae within planktonic and benthic samples has long hampered both applied and basic research efforts in marine, estuarine, and freshwater environments (Hu et al. 1992). For example, assessing the impact of natural or anthropogenic disturbances (e.g., chemical pollutants, thermal discharges, oil spills, dredge spoil dumping, ocean acidification, discharges of ships' ballast water containing entrained meroplanktonic organisms) on marine ecosystems; predicting recruitment for fisheries management; optimizing the timing of substrate placement for aquaculture; assessing the impact of fluctuations in climatic conditions; and conducting basic environmental surveys all depend on identifying the temporal and spatial abundance of meroplanktonic and/or early post-larval stages of various species. Along these lines, the discovery of planktonic stages of a myriad of invertebrate organisms within the ballast waters of ships traveling between countries has led to concerns regarding potentially severe and irreversible biological conservation impacts of the introduction of invasive aquatic species via the discharge of such ballast waters in areas where the entrained species are not indigenous (Cohen & Carlton 1998, Carlton 1999, Ruiz et al. 2000, Gollasch 2002, Murphy et al. 2002). As Hayes and Hewitt (1998) point out: "The assessment of risk associated with ballast water discharge depends on reliable knowledge of the identity, viability and quantity of its inhabitants." Hence, while this monograph consists predominantly of species from environments along the east coast of North America, a number of bivalve species (e.g., Area noae, Pecten maximus, and Ruditapes philippinarum) that various individuals collected from waters in the eastern Atlantic and cultured under controlled laboratory conditions have been included in this treatise.
For over a century, workers have attempted to define larval and post-larval morphological characters diagnostic at various systematic levels (for discussions, see Stafford 1912, Odhner 1914, Werner 1939, Rees 1950, Loosanoff & Davis 1963, Chanley & Andrews 1971, Lutz & Hidu 1979, Fuller & Lutz 1989, Hu et al. 1993, Hendriks et al. 2005, Larsen et al. 2007, Goodwin et al. 2014, 2016a, 2016b, 2018). Historically, the larval and early post-larval characteristics generally used in routine plankton and benthic identifications have been shell length, height, and depth, as well as the length of the "straight-hinge line"; differences in larval shell shape, color, and texture; provinculum length; number and configuration of hinge teeth; and presence or absence of a byssal notch, eyespot, or apical cilia ("apical flagellum") (Loosanoff et al. 1966, Chanley & Andrews 1971, Turner & Boyle 1975, Chanley & Chanley 1980, Lutz et al. 1982b, Fuller & Lutz 1989, Hu et al. 1993, Tan et al. 1993, Larsen et al. 2007). More recently, a spectrum of image analysis, cell sorting, and genetic techniques has proven useful for the identification of a myriad of larval and post-larval bivalves (Tiwari & Gallager 2003a, 2003b, Hendriks et al. 2005, Larsen et al. 2005, 2007, Henzler et al. 2010, Thompson et al. 2012a, 2012b, Goodwin et al. 2014, 2016a, 2016b, 2018).
The present monograph is designed to provide additional tools for the identification of larval and post-larval bivalve species isolated from planktonic and benthic samples from aquatic environments. To this end, Figures 1-195 depict scanning electron micrographic sequences of the gross shell morphologies, morphometries, and details of the hinge regions of disarticulated shell valves of a spectrum of larval and post-larval bivalves at various stages of development. These sequences are presented in a manner (i.e., a consistent orientation of imaged specimens) that will facilitate comparison of the shell morphologies and morphometries of the early life-history stages of these species. The micrographic sequences depict larval and post-larval shell features of species in 47 genera from 25 bivalve families. The morphologies of the larval hinges range from distinctly taxodont dentition in the case of the Arcoidea, Mytiloidea, and Pectinoidea to a lack of prominent denticular structures in the Arcticoidea, Hiatelloidea, Myoidea, and Veneroidea (with the exception of the venerid Ruditapes philippinarum that has fairly well-defined hinge teeth along the larval shell provinculum). The morphological features of various ontogenetic stages of the disarticulated larval and post-larval shell valves of many of the species are quite distinct, permitting discrimination at the specific level. Although differences in morphological features among many other taxa are subtle, it is believed that they can be defined, permitting discrimination of bivalve larvae and postlarvae at the levels of subfamily and genus, respectively.
The following sections summarize the utility (as articulated by various authors) of comparing scanning electron micrographs of early ontogenetic stages of species from a spectrum of select families for discrimination of larval and post-larval bivalves at various taxonomic levels.
Veneridae and Arcticidae
From detailed examination and analyses of the SEM sequences of the early life-history stages of five venerids (Chione cancellata--Figs. 168-170; Mercenaria mercenaria--Figs. 173-175; Mercenaria campechiensis--Figs. 176-178; Mercenaria campechiensis texana--Figs. 179-181; and Pilar morrhuanus--Figs. 186-189), Goodsell et al. (1992) concluded that "documentation and comparison of scanning electron photomicrographs of larval and post-larval venerids would appear to be a successful aid for identification at the levels of subfamily and genus, respectively." As articulated by Lutz et al. (1982b), the sequence of ontogenetic changes in the morphology of the larval hinge apparatus of Arctica islandica (Figs. 149 and 150) is remarkably similar to that described and illustrated by Le Pennec (1978, 1980) for various species of venerids. Throughout larval development, the provinculum of A. islandica is slightly wedge shaped with the narrower portion toward the posterior region of the shell (Fig. 150). "Denticles," analogous to those described by Le Pennec (1980) along the provinculum of certain venerids, are absent. During the early straight-hinge stage, an elongated ridge ("fold"; Le Pennec, 1980) develops in the left valve along almost the entire length of the anterior half of the provinculum (Figs. 149 and 150). At the anterior extremity of the provinculum of the left valve, there is a slight depression in shells of specimens greater than approximately 170 [micro]m in length. In the right valve, a relatively short projection develops during the early straight-hinge stage along the central region of the provinculum (Fig. 150). This latter projection subsequently develops into the first cardinal tooth (according to the nomenclature of Le Pennec, 1978, 1980), which is readily apparent in the post-larval stages (Figs. 151 and 152) (Lutz et al. 1982b).
No "primary" (after Trueman 1950; "primitive" of Le Pennec 1980) ligament pits ("fossettes ligamentaire" of Bernard 1896a) were observed in Arctica islandica specimens with shell lengths <200 [micro]m (Fig. 150) or in numerous specimens ranging in length from 200 to 230 [micro]m. Ligament pits, although often very much reduced in size, appeared to be present in all specimens examined with shell lengths >230 [micro]m. Since the classic studies of Bernard in the late nineteenth century (Bernard 1895, 1896a, 1896b, 1897, 1898), numerous workers have commented on the presence of ligaments or ligament pits in "larval" specimens (Rees 1950, Ansell 1962, Loosanoff et al. 1966, Chanley & Andrews 1971, Bayne 1976; for further discussion concerning the significance of the presence or absence of ligament pits in the shells of early ontogenetic stages of bivalves, see Lutz et al. 1982b). Lutz and Hidu (1979) suggested that "primary" (after Trueman 1950) ligament pits do not form until metamorphosis has been initiated (see also Lutz 1979). It has been further suggested that changes associated with metamorphosis proceed in an orderly fashion (Bayne 1965, 1971,Turner 1976b)and that "any interruption... in the normal sequence of events affects the ability of the larvae to progress to the next step whether that be the loss of a larval organ or the acquisition of a post-larval one" (Turner 1976a).
In specimens of Arctica islandica examined during the course of preparation of the present monograph,,ligament pits were observed in shells of a number of specimens with larval lengths ranging between 200 and 230 [micro]m. If, as suggested by Lutz and Hidu (1979, pp. 117-118), development of the primary ligament pit is "one of the first morphological changes that occurs during metamorphosis," it is reasonable to conclude that larvae within this size range are at least capable of metamorphosis (Lutz et al. 1979). Ligament pits were not observed in any A. islandica specimens with shell lengths <200 [micro]m (Figure 150) (Lutz et al. 1982b). This observation, when coupled with the fact that no pediveliger larva with a length below 200 [micro]m was found in any of the cultures, strongly suggests that the larvae of this species are not capable of metamorphosing at shell lengths below this size and has implications for determining the size at which most of the species in this monograph are capable of metamorphosing.
After metamorphosis, relatively dramatic changes take place in the morphology of the hinge apparatus of Arctica islandica. The sequence of ontogenetic changes photographically illustrated in Figures 151 and 152 is similar to that diagrammatically illustrated by Le Pennec (1978, 1980) in his detailed summary of the development of the heterodont hinge. A close similarity exists between the early post-larval hinge apparatus of Venus verrucosa figured by Le Pennec (1980, p. 617) and that of A. islandica. At a shell length of approximately 1-2 mm (Figs. 151, 152), the A. islandica hinge has acquired many adult characteristics, although it does not attain its definitive form until specimens have reached a shell length of approximately 4 mm (Fig. 12 in Lutz et al. 1982b, p. 761).
Fuller and Lutz (1989) published scanning electron micrograph sequences of the larvae and postlarvae of six mytilids from the northwestern Atlantic: Arcuatula papyria (= Amygdalum papyrium) (Figs. 77-80); Brachidontes exustus (Figs. 81-84); Geukensia demissa (Figs. 85-88); Ischadium recurvum (Figs. 89-92); Modiolus modiolus (Figs. 97-100); and Mytilus edulis (Figs. 101-104). All six species have a long provinculum with taxodont dentition, with provinculum length and number of teeth increasing steadily during the larval period. The bold, comparatively few provincular teeth of A. papyria (Fig. 78) and the small, numerous provincular teeth of M. edulis (Fig. 102) clearly differentiate these two species. Most of these mytilid species have a low umbo, round posterior margin, and more pointed anterior margin, although A. papyria is distinguished by a high, prominent umbo. The larval shells of G. demissa and I. recurvum are difficult to differentiate because of the similarity in their shapes and hinge dentition; however, as articulated by Fuller and Lutz (1989), "discriminant analysis using larval shell length, shell height, provinculum length, and number of teeth aided in classification of these and other sympatric species."
By contrast, post-larval stages of these six mytilid species are plainly distinguished by the presence and type of lateral teeth seen in the SEM sequences published by Fuller and Lutz (1989). Brachidontes exustus has all three types of mytilid lateral teeth, including (1) primary lateral teeth, which form immediately posterior to provincular teeth; (2) secondary lateral teeth, which are posterior to the primary lateral teeth and are part of the dissoconch; and (3) dysodont teeth, which form on the anterior margin of the dissoconch (Figs. 83 and 84). Modiolus modiolus has primary lateral teeth (Figs. 99 and 100), Ischadium recurvum has dysodont teeth (Figs. 91 and 92), Mytilus edulis has secondary lateral and dysodont teeth (Figs. 103 and 104), and there are no lateral teeth in Geukensia demissa (Figs. 87 and 88) or Arcuatula papyria (Figs. 79 and 80) during early post-larval development. The provinculum increases in size and complexity during post-larval development in A. papyria, B. exustus, G. demissa, I. recurvum, and M. modiolus, but not in M. edulis (Fuller & Lutz 1989).
Hu et al. (1993) published scanning electron micrograph sequences to elucidate species-specific shell features in larval and post-larval stages of four Ostreidae species: Crassostrea gigas, Crassostrea virginica, Ostrea edulis, and Ostrea stentina (= Ostrea equestris) (Figs. 105-123). Useful features for distinguishing larvae of the Ostreidae from those within other families of bivalves include asymmetry of left and right valves associated with a pronounced umbo of the left valve; one or two provincular teeth on each side of the provinculum; and a fasciole with a corresponding notch on the left valve. Qualitative characters, such as hinge dentition and shell shape of the larvae and postlarvae of the four ostreids, distinguish noncongeneric species, whereas species in the same genus generally are differentiated by quantitative features such as the dimensions of the provinculum and shell.
The results of the study by Hu et al. (1993), when combined with those of previous studies on Crassostrea angulata (Pascual 1971), Crassostrea ariakensis (Tanaka 1980), Crassostrea glomerata (Dinamani 1973), Crassostrea iredalei (Ver 1986), Ostrea denselamellosa (Tanaka 1980), Ostrea lurida (Loosanoff et al. 1966), Ostrea permollis (Forbes 1967), Ostrea puelchana (Castro & Le Pennec 1988), Ostrea stentina (Pascual 1972), and Ostrea spp. (Chanley & Dinamani 1980), illustrate that the morphological differences between early ontogenetic stages of species within the genus Crassostrea and those within the genus Ostrea are quite striking. The diagnostic characters for larval and post-larval shells of species within these two genera are summarized as follows.
Morphological larval features of species within the genus Crassostrea include (1) a knobby or beak-shaped umbo directed posteriorly; (2) shell height that is generally greater than shell length; (3) provinculum length that ranges from 68 to 88 [micro]m; (4) the presence of two rectangular teeth on each side of the provinculum; (5) a well-defined central apparatus; and (6) posterior obscured teeth. By contrast, morphological features of species within the genus Ostrea include (1) a round shell with a large, moderately prominent umbo directed dorsally; (2) shell length that is greater than shell height; (3) provinculum length that ranges from 70 to 100 [micro]m; (4) the presence of one anterior and one posterior tooth on the left valve and two anterior and two posterior teeth on the right valve, with teeth that are square or triangular; (5) a partially developed central apparatus; and (6) anterior obscured teeth.
Post-larval features of species within the genus Crassostrea include (1) a left valve umbo that is beak shaped and skews backward; (2) two posterior hinge teeth in shells up to 500 [micro]m in length; and (3) a ligament that is 40 [micro]m from the anterior end of the provinculum. By contrast, post-larval features of species within the genus Ostrea include (1) an umbo that is round and prominent dorsally; (2) one or two anterior hinge teeth in shells up to 500 [micro]m in length; and (3) a ligament that is close to the anterior end of the provinculum (Hu et al. 1993).
The shell morphological features that are most useful for discriminating larval and post-larval specimens of the four ostreids that are included in this monograph are articulated by Hu et al. (1993) and may be summarized as follows:
(1) The species-specific characteristics of the provinculum remain relatively constant throughout the developmental stages. These diagnostic characters include the dimensions of the provinculum, the shape of the hinge teeth, and the modification pattern of the hinge teeth during the late larval stages. For example, the provinculum of larval and post-larval (up to a shell length of 550 [micro]m) specimens of Crassostrea virginica is shorter but wider (50 [micro]m long and 14 [micro]m wide) than that of Crassostrea gigas (56 [micro]m long and 10 [micro]m wide). The transverse ridges are less defined and fewer in C. gigas than in C. virginica. The posterior hinge teeth of C. virginica are well defined even in late-stage larvae. These observations agree with the pattern of larval hinge modification in the genus Crassostrea postulated by Dinamani (1976). The range of provincular lengths of Ostrea stentina is 70-79 [micro]m, which is much shorter than the 80-95 [micro]m provincular length range of Ostrea edulis (Hu et al. 1993).
(2) The four species have different shell shapes, including prominence of umbos, shapes of anterior and posterior ends, rotation of the longitudinal axis, and shell length-height ratio. The shell shape of larval specimens of Crassostrea virginica is relatively compressed dorsoventrally, with a narrow and pointed anterior end, whereas that of Crassostrea gigas larvae is more extended in dorsoventral directions and its anterior end appears broad and blunt. A knobby umbo with an elongated anterior end is present in late-stage larval specimens of Ostrea stentina, whereas the umbo of Ostrea edulis is always flat and the anterior and posterior are nearly equally developed. Shell shape is also useful for identification of post-larval specimens. For example, two shoulders (anterior and posterior dorsal margins) extend at an angle of approximately 90 deg in C. gigas and at an angle of 140 deg in C. virginica, and were well defined in O. stentina but irregular in O. edulis (Hu et al. 1993).
In summary, generic and species diagnostic shell characters have been identified for early life-history stages of two species within the genus Crassostrea (C. gigas and C. virginica) and two species within the genus Ostrea (O. edulis and O. stentina). A key has been presented by Hu et al. (1993) summarizing these morphological characteristics that should provide a practical tool for the identification of larval and post-larval specimens of these ostreids isolated from planktonic and benthic samples.
Scanning electron micrograph sequences published by Fuller et al. (1989a) and Tan et al. (1993) of disarticulated shell valves of Teredo navalis and Bankia gouldi, respectively, revealed qualitative and quantitative differences in larval and post-larval morphological features (Figs. 69-76). These sequences provide useful aids for the identification of larval and post-larval specimens of these two species isolated from plankton and benthic samples. In particular, the following features are important in distinguishing the larvae of B. gouldi from those of T. navalis: slope of the shoulders [the dorsal shell margin on the anterior and posterior ends of the hinge (Chanley & Dinamani 1980)]; length of the provinculum/hinge-line; and length of the posterior tooth of the left valve provinculum.
Tan et al. (1993) point out that, in larval shells of similar size, the shoulders of Teredo navalis (Fig. 73) are considerably steeper (less rounded) than those of Bankia gouldi (Fig. 69). This difference in shell shape is useful in distinguishing the two species. The initial size of the planktonic stage of a B. gouldi larva is much smaller than that of the larviparous T. navalis because of the difference in their development. Despite the difference in initial larval shell size, both species metamorphose at a shell height of about 230 [micro]m, as indicated by the appearance of the ligament pit.
The larvae of Bankia gouldi and Teredo navalis can be distinguished on the basis of the length of the provinculum of the left valve. The average length of the provinculum/hinge-line measured by Tan et al. (1993) from micrographs of the left valves of B. gouldi (38.1 [+ or -] 1.8 [micro]m, n = 9) (Fig. 70) is significantly smaller than that of T. navalis (46.7 [+ or -] 1.3 [micro]m, n = 9) (Fig. 74). The provincular length measurements of the left valve are more useful than those of the right valve because the latter varies significantly with growth.
Larvae of Bankia gouldi can also be distinguished from those of Teredo navalis by the length of the posterior provincular tooth of the left valve (6.4 [+ or -] 0.6 [micro]m, n = 7) (Fig. 70) which is significantly shorter than that of T. navalis (8.2 [+ or -] 0.3 [micro]m) (Fig. 74) (Tan et al. 1993).
Practical Use of this Monograph
Although the SEM sequences presented in Figures 1-195 accurately depict the gross morphologies/morphometrics and hinge structures of the disarticulated shell valves of the larvae and/or postlarvae of 56 species of bivalves, it is important to emphasize that a scanning electron microscope is not necessary to observe even fine hinge structures associated with the ontogenetic stages of these species. Such structures are readily visible using a wide range of optical compound microscopes equipped with high-intensity reflected light sources, although the disarticulated shell valves must be viewed in several planes of focus to discern the often subtle details seen clearly in the scanning electron micrographs.
The comparisons of the larval and early post-larval stages of two mytilids (Mytilus edulis and Modiolus modiolus) published by Lutz and Hidu (1979) and depicted in Figures 97-104 dramatically illustrate this point. In their studies, measurements of larval dimensions (shell length, height, and provinculum length) and dentition counts were made under a standard petrographic microscope equipped with a high-intensity reflected light source. Using this procedure, Lutz and Hidu (1979) were able to obtain quantitative measurements and counts on approximately 20 specimens per hour. Numerous specimens were also examined and measured under a scanning electron microscope to confirm the accuracy of these optical microscopic measurements and counts. Their results indicated that, for a given number of provincular teeth, there was no overlap between the range extremes in either provinculum length or total shell length of the two species. There were no quantitative differences between left and right valve of individual specimens other than occasional minor discrepancies in dentition counts due to the interlocking nature of the hinge teeth. Lutz and Hidu (1979, p. 115) concluded that "careful examination of the hinge morphology of prodissoconchs and early dissoconchs using routine optical microscopic techniques should facilitate unambiguous differentiation of all early life-history stages of these two species."
In conclusion, scanning electron microscopy provides a powerful tool for photographically depicting the accurate gross shell morphology morphometry and details of the hinge structure of the disarticulated shell valves of the larvae and early postlarvae of known species of bivalves cultured under controlled laboratory conditions. These morphological characters, in turn, provide researchers with invaluable aids for identifying (using routine optical microscopic techniques) the early life-history stages of these species isolated from plankton and benthic samples.
The authors thank William Arnold, Robert Bisker, Thomas Bright, Bernardita Campos, Judy McDowell Capuzzo, Melbourne Carriker, Paul Chanley, Alison Craig, Greg Debrosse, William Foster, Mary Gibbons, John Grazul, Victor Greenhut, Harold Haskin, Kathleen Huntington, John Kraeuter, Dale Leavitt, Robert Loveland, William Mook, James Moore, Steven Murawski, Elizabeth W. North, Hugh Porter, Robert Prezant, Robert Robertson, John Ropes, William Sacco, Neil Savage, Rudolf Scheltema, Ruth Turner, Rosa Van Dessel, Eric Wagner. Randy Walker, Thomas Waller, Jean Watkinson, and John Whitcomb for their advice, assistance, and helpful comments. This research was supported by the New Jersey and South Carolina Agricultural Experiment Stations; the School of Environmental and Biological Sciences of Rutgers University; the South Carolina Sea Grant Consortium; Steinetz Awards from the Department of Biological Sciences, Rutgers University; a Grant-in-Aid of Research from the Society of Sigma Xi (SCF); the Maryland Power Plant Research Program; the Center for Environmental and Estuarine Studies, University of Maryland; NSF Grants EAR 81-21212 and EAR-84-17011 (RAL); and various NOAA Sea Grants (RAL).
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RICHARD A. LUTZ, (1*) JACOB D. GOODWIN, (1) BRAD S. BALDWIN, (2) GAVIN BURNELL, (3) MICHAEL CASTAGNA, (4) SAMUEL CHAPMAN, (5) AL CHESTNUT, (6) PATRICK DABINETT, (7) CHRIS DAVIS, (5) ARNOLD G. EVERSOLE, (8) S. CYNTHIA FULLER, (1) SCOTT M. GALLAGER, (9) RONALD GOLDBERG, (10) JOY GOODSELL, (1) JUDITH GRASSLE, (1) RICHARD G. GUSTAFSON, (11) HERBERT HIDU, (5) YA-PING HU, (1) DAVID JABLONSKI, (12) SHANNON JOHNSON, (13) VICTOR S. KENNEDY, (14) MARCEL LE PENNEC, (15) ROGER MANN, (16) CARTER NEWELL, (5) ALAN S. POOLEY, (1) ANTONIETO S. TAN, (1) ROBERT C. VRIJENHOEK (13) AND A. PARTRIDGE (17)
(1) Department of Marine and Coastal Sciences, Rutgers University, 71 Dudley Road, New Brunswick, NJ 08901; (2) Department of Biology, 23 Romoda Drive, St. Lawrence University, Canton, NY 13617; (3) School of Biological, Earth and Environmental Sciences, University College Cork, North Mall Campus, Distillery Fields, Cork, Ireland T23 N73K; (4) Virginia Institute of Marine Science, College of William and Mary, 40 Atlantic Avenue, Wachapreague, VA 23480; (5) Ira C. Darling Center, University of Maine, 193 Clarks Cove Road, Walpole, ME 04573; (6) Department of Biology, Belhaven College, 1701 N. State Street, Jackson, MS 39202; (7) Marine Sciences Research Laboratory, Memorial University of Newfoundland, 230 Elizaabeth Avenue, St. John's, NF A1C 5S7, Canada; (8) Department of Aquaculture, Fisheries and Wildlife, 261 Lehotsky Hall, Clemson University, Clemson, SC 29634; (9) Woods Hole Oceanographic Institution, 266 Woods Hole Road, Woods Hole, MA 02543; (10) Northeast Fisheries Center, National Marine Fisheries Service, Milford Laboratory, 212 Rogers Avenue, Milford, CT06460; (11) Northwest Fisheries Science Center, National Marine Fisheries Service, National Oceanic and Atmospheric Administration, 2725 Montlake Blvd. E., Seattle, WA 98112; (12) Department of Geophysical Sciences, University of Chicago, 5734 S. Ellis Ave., Chicago, IL 60637; (13) Monterey Bay Aquarium Research Institute, 7700 Sandholdt Road, Moss Landing, CA 95039; (14) Chesapeake Biological Laboratory, University of Maryland Center for Environmental Science, 146 Williams Street, Solomons, MD 20688; (15) Laboratoire des Sciences de l'Environnement Marin (LEMAR), Institut Universitaire Europeen de la Mer, Place Nicolas Copernic, Plouzane 29280, France; (16) Virginia Institute of Marine Science, College of William and Mary, 1375 Greate Road, Gloucester Point, VA 23062; (17) in a pear tree
(*) Corresponding author. E-mail: email@example.com
TABLE 1. The 56 species of bivalves for which larval and/or post-larval SEM sequences are depicted in Figures 1-195. Figure Order Family Genus Species numbers 1-4 Adapedonta Hiatellidae Hiatella arctica 5-8 Adapedonta Pharidae Ensis leei 9-12 Arcida Arcidae Area none 13-16 Arcida Arcidae Lunarca ovaiis 17-19 Arcida Noetiidae Noetia ponderosa 20-23 Cardiida Cardiidae Dinocardium robustum 24-27 Cardiida Cardiidae Laevicardium mortoni 28-31 Cardiida Solecurtidae Tagelus plebeius 32-35 Cardiida Tellinidae Ameritella agilis 36-39 Cardiida Tellinidae Ameritella mitchelli 40-43 Cardiida 1 ellinidae Limecola balthica 44 Carditida Carditidae Cyclocardia borealis 45 Carditida Astartidae Astarte castanea 46-49 Myida Dreissenidac Dreissena bugensis 50-53 Myida Dreissenidae Dreissena polymorpha 54-57 Myida Dreissenidae Mytilopsis leucophaeata 58-61 Myida Myidae My a arenaria 62-64 Myida Myidae My a truncal a 65, 66 Myida Pholadidae Cyrtopleura costata 67, 68 Myida Pholadidae Diplothyra carta 69-72 Myida Teredinidae Bankia gouldi 73-76 Myida Teredinidae Teredo navalis 77-80 Mytilida Mytilidae Arcuatula papyriu 81-84 Mytilida Mytilidae Brachidontes exustus 85-88 Mytilida Mytilidae Geukensia demissa 89-92 Mytilida Mytilidae Ischadium recurvum 93, 94 Mytilida Mytilidae Leiosolenus bisulcatus 95, 96 Mytilida Mytilidae M odiolus americanus 97-100 Mytilida Mytilidae Modiolus modiolus 101-104 Mytilida Mytilidae Mytilus edulis 105-109 Ostreida Ostreidae Crassostrea gigas 110-114 Ostreida Ostreidae Crassostrea virginica 115-118 Ostreida Ostreidae Ostrea edulis 119-123 Ostreida Ostreidae Ostrea stentina 124-127 Pectinida Anomndae Anomia simplex 128-130 Pectinida Pectinidae Argopecten irradians 131-134 Pectinida Pectinidae Argopecten irradians concentricus 135-138 Pectinida Pectinidae Pecten maximus 139-143 Pectinida Pectinidae Placopecten magellanicus 144. 145 Pholadomyida Lyonsiidae Lyonsia hyalina 146 Pholadomyida Periplomatidae Periploma leanum 147, 148 Solemyida Solemyidae Solemya velum 149-152 Venerida Arcticidae Arctica islandica 153-156 Venerida Mactridae Mulinia Lateralis 157-159 Venerida Mactridae Rangia cuneata 160-163 Venerida Mactridae Spisula solidissima 164-167 Venerida Mesodesmatidae Mesodcsma arctatum 168-170 Venerida Veneridae Chione cancellata 171, 172 Venerida Veneridae Gemma gemma 173-175 Venerida Veneridae Mercenaria mercenaria 176-178 Venerida Veneridae Mercenaria campechiensis 179-181 Venerida Veneridae Mercenaria campechiensis 182-185 Vcnerida Veneridae Petricolaria tC'XWltl pholadiformis 186-189 Venerida Veneridae Pitar morrhuanus 190-193 Venerida Veneridae Ruditapes philippinarum 194. 195 Venerida Cyrenidae Corbicula fluminea Figure Authority Source of sexually mature adult numbers bivalves 1-4 (Linnaeus, 1767) Pemaquid. ME 5-8 M. Huber, 2015 Damariscotta River, ME 9-12 Linnaeus, 1758 Istrian Peninsula Coast, Rovinj, Croatia 13-16 (Bruguiere, 1789) Wachapreague, VA 17-19 (Say, 1822) Wachapreague, VA 20-23 (Lightfoot. 1786) Indian River near Grant, FL 24-27 (Conrad, 1831) Coastal bay near Wachapreague, VA 28-31 (Lightfoot, 1786) Delaware Bay, NJ 32-35 (Stimpson. 1857) Wachapreague Inlet, Wachapreague, VA 36-39 (Dall, 1895) Choptank River, MD 40-43 (Linnaeus, 1758) Lowe's Cove, Walpole, ME 44 (Conrad, 1832) Continental Shelf off NJ 45 (Say, 1822) Continental Shelf off NJ 46-49 (Andrusov, 1897) Lake Ontario near Rochester, NY 50-53 (Pallas. 1771) St. Lawrence River near Cape Vincent, NY 54-57 (Conrad. 1831) Hudson River near Piermont, NY 58-61 Linnaeus, 1758 Damariscotta River, Walpole, ME 62-64 Linnaeus, 1758 Gulf of Maine off Boothbay Harbor, ME 65, 66 (Linnaeus, 1758) Indian River, FL 67, 68 (G. B. Sowerby 1, 1834) Mississippi Sound. MS 69-72 (Bartsch, 1908) York River, Gloucester Point, VA 73-76 Linnaeus, 1758 Coastal bay near Wachapreague Inlet, VA 77-80 (Conrad. 1846) Indian River, FL 81-84 (Linnaeus, 1758) Cabbage Island and Wilmington Island, GA 85-88 (Dillwyn, 1817) Maurice River, NJ 89-92 (Rafinesque, 1820) James River, VA 93, 94 (d'Orbigny, 1853) Jamaica. West Indies 95, 96 (Leach, 1815) West coast of Florida 97-100 (Linnaeus, 1758) Cape Newagen, ME 101-104 Linnaeus, 1758 Continental Shelf off NJ 105-109 (Thunberg, 1793) Coast Oyster Company, Quilcene, WA 110-114 (Gmelin, 1791) Delaware Bay, NJ 115-118 Linnaeus. 1758 Walpole, ME 119-123 Payraudeau, 1826 Newport River Estuary, NC 124-127 d'Orbigny. 1853 Wachapreague Inlet, Wachapreague, VA 128-130 (Lamarck. 1819) Cape Cod Bay, MA 131-134 (Say, 1822) Coast of North Carolina near Morehead City 135-138 (Linnaeus, 1758) Coast of Brest, France 139-143 Gmelin, 1791 St. John's, Newfoundland (larvae); Damariscotta River, ME (postlarvae) 144. 145 (Conrad. 1831) Mason's Beach, VA 146 (Conrad, 1831) Continental Shelf off New Jersey 147, 148 Say, 1822 Upper Buttermilk Bay, MA and Shark River, NJ 149-152 (Linnaeus, 1767) Continental Shelf, off New Jersey and Rhode Island 153-156 (Say, 1822) Wachapreague, VA 157-159 (G. B. Sowerby I, Cohansey River near Bridgcton, NJ 1832) 160-163 (Dillwyn, 1817) Continental Shelf off Rhode Island 164-167 (Conrad, 1831) Sable Island Bank (depth 42 m) off Nova Scotia, Canada 168-170 (Linnaeus, 1767) Indian River, FL 171, 172 (Totten, 1834) Delaware Bay, NJ 173-175 (Linnaeus, 1758) Milford, CT 176-178 (Gmelin, 1791) Gulf of Mexico near Apalachicola, FL 179-181 (Dall, 1902) Gulf of Mexico near Galveston, TX 182-185 (Lamarck, 1818) Continental Shelf off southern New Jersey 186-189 (Dall. 1902) Continental Shelf off Wachapreague, VA 190-193 (Adams & Reeve. Obtained from the Carna Research 1850) Station, Galway, Ireland 194. 195 (O. F. Miiller, 1774) Nassawango Creek, MD Taxonomic nomenclature was assigned according to the latest "accepted name" (or acceptable "alternate representation") and associated classification hierarchy in the World Register of Marine Species (www.marinespecies.org). Sexually mature adult bivalves were obtained from the indicated sources and used as broodstock to obtain the larvae and postlarvae depicted in this monograph.
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|Author:||Lutz, Richard A.; Goodwin, Jacob D.; Baldwin, Brad S.; Burnell, Gavin; Castagna, Michael; Chapman, S|
|Publication:||Journal of Shellfish Research|
|Date:||Jun 1, 2018|
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