Printer Friendly

Embryogenesis and development of Epimenia babai (Mollusca Neomeniomorpha).

Introduction

Neomenioid aplacophorans (= Solenogastres) are shell-less vermiform molluscs with a ciliated foot and an epidermal cuticle covered by many calcareous spicules. Given that aplacophorans are hypothesized to hold a basal position among molluscs (Gotting, 1980; Salvini-Plawen, 1985; Salvini-Plawen and Steiner, 1996; Ivanov, 1996; Waller, 1998; Haszprunar, 2000), their development may provide the key to answering outstanding questions in the evolution of molluscan larvae. However, little is known about the developmental processes and features of neomenioid aplacophorans. Larval descriptions are based on five neomenioid aplacophoran species: Epimenia babai Salvini-Plawen, 1997 (as E. verrucosa); Halomenia gravida Heath, 1918; Neomenia carinata Tullberg, 1875; Nematomenia (= Dondersia) banyulensis Pruvot, 1890; and Rhopalomnenia (= Proneomenia) aglaopheniae Kowalewsky and Marion, 1887 (Pruvot, 1890, 1892; Heath, 1918; Baba, 1938, 1940, 1951; Thompson, 1959, 1960). Early cleavage pattern is known only for E. babai (B aba, 1940, 1951; and herein). Nothing has been reported on the early embryology of chaetodermomorph aplacophorans (= Caudofoveata), and their larval descriptions are based on a drawing of Scutopus robust us (Salvini-Plawen, 1990), on drawings based on Gustafson's unpublished sketches (Nielsen, 2001), and on scanning electron micrographs of Chaetoderma nitidulum larva (Nielsen, presented at World Congress of Malacology 2002).

Neomenioid aplacophorans are hermaphroditic and either brood juveniles (Heath, 1918; Salvini-Plawen, 1978) or have free-swimming lecithotrophic larvae with an outer locomotory larval test, within which the definitive adult structures develop (Pruvot, 1890, 1892; Thompson, 1960). A similar enveloping test, or "pericalymma," has also been described for the larvae of protobranch bivalves (Drew, 1897, 1899a, b, 1901; Gustafson and Reid, 1986, 1988a, b; Gustafson, 1987; Gustafson and Lutz, 1992; Zardus and Morse, 1998). The homology of larvae with tests among protobranch bivalves and neomenjoid aplacophorans remains uncertain because the phylogenetic relationships of these groups are still debatable, the morphology of these larvae is quite diverse, and the morphology of the neomenioid test is poorly understood (Pruvot, 1890, 1892; Drew, 1899b; Gustafson and Reid, 1986; Thompson, 1959, 1960; Gustafson and Lutz, 1992; Zardus and Morse, 1998).

The neomenioid aplacophoran Epimenia babai from Japan is considered to have a unique larval form among neomenioid aplacophorans in that it lacks the true test structure characteristic of neomenioid pericalymma larvae (Baba, 1938, 1940, 1951, 1999; Thompson, 1960; Nielsen, 2001). Baba (1938, 1940, 1951) was the first to describe the larval development of E. babai, and he suggested that the morphology and fate of the apical cells of the species do not resemble the pericalymma test cells of other neomenioid aplacophorans. Because of its unique development, the larval form of E. babai has been suggested to be an intermediate form between pericalymma and trochophore larvae (Baba, 1938, 1940, 1951, 1999) called "stenocalymma" by Salvini-Plawen (1973, 1980). The descriptions by Baba were based primarily on histological sections, but details of morphology and cell fate during embryogenesis were not studied, and thus such differentiation is not certain.

Adult specimens of E. babai are easily collected and maintained in the laboratory. Whereas most aplacophorans are small (1-5 mm) and inhabit the deep sea where they are not easily obtainable for developmental studies, E. babai can reach 30 cm in length and lives in relatively shallow water from 20 to 70 m in depth (Salvini-Plawen, 1997). The embryos are relatively large (ca. 250 [micro]m) and can be maintained at room temperature. Taking advantage of the accessibility of E. babai, this study reexamines the early and late development of the species to elucidate some of the unknown developmental features of a neomenioid aplacophoran, such as the morphology and fate of the pericalymma test.

Materials and Methods

Animal collection and culture

Adult specimens of E. babai were collected offshore near Amakusa Marine Biological Laboratory (Kumamoto, Japan) in 1999 and 2000 during the most active breeding period between June and September (Baba, 1938, 1940, 1951). These specimens were up to 20 cm in length and 1 cm in diameter. The exemplars were collected from synthetic gill nets (3 m width, 500 m length, 5 X 5 cm mesh size) that were set on a rocky bottom at a depth of 20 to 30 m among beds of the soft coral Alcyonium gracillimum.

The adults were maintained in an aquarium with running seawater at ambient temperature, about 26 [degree]C, and were supplied with fresh Alcyonium gracillimum for food. The animals spawned spontaneously several hours after sunset or were induced to spawn by turning off the light after 12-14 h of constant light. Spawned egg sheets (see Fig. la) were transferred to culture jars containing filtered seawater (0.45-[micro]m mesh) with streptomycin (50 mg/ml) and penicillin (35 mg/ml) to reduce bacterial growth (Strathmann, 1987). Cultures were kept at both ambient temperature and 16 [degrees]C and their development was compared. Developmental descriptions are based on cultures maintained at 16 More than 1000 embryos from three adults were analyzed.

Scanning electron microscopy

Egg capsules were freed from a jelly mucous casing by gentle agitation, and the embryos were dissected out of their capsules with fine forceps. The embryos were fixed in 1% osmium tetroxide (OsO4) in filtered seawater and washed twice in 0.2 M sodium cacodylate buffer solution (pH 7.4). They were then dehydrated through a series of ethanol washes (20%, 50%, 75%) and transported from Amakusa to Woods Hole, Massachusetts, where dehydration in ethanol (100%, 100%) was completed. The samples were criticalpoint-dried (SAMDRI-780), mounted on aluminum stubs with double-sided tape, coated with gold in a sputter-coater (SAMSPU'TTER-2a), and imaged using a JEOL scanning electron microscope (JSM-840).

Semithin sections

Dissected embryos were first fixed in a solution of 3% gluteraldehyde in 0.2 M sodium cacodylate buffer at pH 7.4, washed in 0.2 M sodium cacodylate buffer, postfixed in 1% osmium tetroxide and washed in the buffer again. The embryos were then dehydrated, transported as above, embedded in Araldite/Epon (EMbed-812), and sectioned with glass knives into semithin sections (0.30-1.0 [micro]m). The sections were mounted onto clean slides on a hot plate (60 [degrees]C), stained with a mixture of methylene blue and azure II (Richardson et al., 1960), washed under tap water, and imaged on a Zeiss light microscope.

Hoechst nuclear staining

Larvae were incubated overnight at 4 [degrees]C in Hoechst solution (1 [micro]g/ml) in 4% formalin (in seawater). The larvae were then washed in 1X phosphate buffered saline (PBS), mounted on a glass slide, and examined with fluorescence under a microscope with a UV filter set.

Programmed cell death determination

Cell viability and apoptosis in the larvae were detected in vivo using acridine orange/ethidium bromide staining according to the method described by Martin et al. (1996) with slight modifications. Programmed cell death (PCD) occurs frequently during animal development when larval tissues are degenerated and lost during the transformation into the adult body form. Live larvae were incubated for 10 min in acridine orange (100 [micro]g/ml + ethidium bromide (100 [micro]g/ml) in seawater. The larvae were washed in clean seawater for an hour and, while alive, were examined on a glass slide under a fluorescence microscope with a UV filter set.

The acridine orange intercalates into the DNA of live cells with intact membranes, and the cells fluoresce bright green. As apoptosis progresses, ethidium bromide enters the cells, and they fluoresce bright orange. Late apoptotic cells have a uniform bright orange color (Martin et al., 1996).

Results

Reproduction and spawning

Three individuals of Epimenia babai laid approximately 20,000 eggs over 90 consecutive nights in the laboratory. One individual that was isolated for more than 90 days laid viable eggs. Sperm transfer was not observed.

Fertilized eggs were laid in pairs of square gelatinous sheets (3 x 3 mm) containing a single layer of 20-50 eggs (Fig. 1a). One individual laid one to four pairs of egg-sheets every night, one pair at a time, at intervals of about an hour. The egg sheets were always laid in pairs, presumably one from each of the paired gametoducts that open into the mantle cavity at the posterior end of the animal. At the time of deposition, the fertilized eggs were always at the single-cell stage. No brooding was observed.

Early development

The early development of E. babai was followed using both scanning electron and light microscopy. Development was slower at 16 [degrees]C than at room temperature (26 [degrees]) (Table 1). However, hatching time observed at 16 [degrees]C was the same as at 26 [degrees]C, with larvae reared at 16 [degrees]C hatching at a much earlier stage of development than those at 26 [degrees]. The lower temperature did not cause any abnormalities in further development.

Observations of early embryogenesis are generally consistent with those of Baba (1940). The eggs are oval at first and become spherical within 30 min at a diameter of 250 [micro]m. Each egg is enclosed in a tough transparent egg capsule 300-350 [micro]m in diameter (Fig. lb). The eggs are opaque yellow, presumably from high yolk content, and were surrounded by numerous accessory granules. Formation of two polar bodies, as reported by Baba (1938, 1940, 1951), was not observed.

Cleavage is spiral, unequal, and asynchronous (Figs. 2-9). First cleavage was meridional (= parallel to animal-vegetal axis) and unequal (Fig. 3). Just before the first cleavage was completed, the first polar lobe protruded from the vegetal side of the embryo (Fig. 3). The polar lobe appeared whiter and less yolky than the rest of the contents of the CD cell. The first polar lobe was soon absorbed by one of the two blastomeres, making this blastomere (CD) slightly larger than the other (AB).

Second cleavage was also meridional and unequal, forming blastomeres A-D. Cleavage was slightly asynchronous, with the larger blastomere (CD) dividing slightly later than the AB cell (Figs. 4, 5). The cross-furrow of the four blastomeres was visible but indistinct (Fig. 4). A second polar lobe protruded from the vegetal region during CD division. Within 30 min, it was absorbed by the D blastomere (Fig. 5), making it the largest of the four.

Third cleavage was equatorial (= perpendicular to the animal-vegetal axis), resulting in four micromeres at the animal pole (la-1d); the four macromeres remained at the vegetal pole (lA-1D) (Fig. 6). The chirality of spiral cleavage displaced the four micromeres dextrally (clockwise from the animal pole) relative to the four macromeres. Cell 1D seemed to be slightly larger than the rest of the macromeres, but the difference was not substantial. Cleavage continued to be slightly asynchronous by alternation of chirality: the fourth cleavage was sinistral (anti-clockwise from the animal pole), producing 16 cells (Fig. 7); and the fifth cleavage was dextral, producing 32 cells (Fig. 3).

The embryos reached 64 cells about 15 h after they were laid and became flattened animal-vegetally, forming a blastula (Fig 9). Further cleavage stages until gastrulation were difficult to follow owing to the asynchronous nature of cleavage.

Within 24 h after oviposition, while the embryos were still within the capsule, gastrulation occurred at the vegetal pole by means of both epiboly and invagination (Figs 10, 11). Cells between the blastopore and presumptive prototrochal cells thickened to form a broad lip around the vegetal side of the gastrula (Fig. 10). Patches of short cilia covered the surface of the gastrula. Cells within the blastopore proliferated to form an outgrowth of a definitive ectodermal bud beneath. From this elongation of ectoderm, the blastopore diminished and migrated toward the future ventral side of the larva (Fig. 12).

Two days after oviposition the presumptive apical tuft and prototroch on the apical region, and the telotroch at the caudal end of the ectodermal trunk, became evident (Figs. 13, 27a) The unciliated ectodermal trunk bent slightly towards the ventral side.

Larval development

Three days after oviposition, the larvae hatched and swam forward and spirally, using the prototroch, until they reached metamorphosis 9 to 13 days after oviposition (7- to 11-day-old larvae) (see below). It is not certain whether the telotroch was directly involved in swimming or not The larvae did not seem to be phototactic.

Larvae aged 1 to 3 days. The free-swimming larvae had three distinct body regions: apical cap, trunk, and caudal region Figs. 14a, 27b). The entire pre-oral sphere of the larvae, the apical cap, was completely ciliated and was divided into a pretrochal and a post-trochal region by a prominent row of compound prototroch cilia at its equator (Figs. 14a, b, 16, 17, 27b). The pretrochal hemisphere of the apical cap originated most larval structures, such as the apical tuft, also composed of compound cilia (Figs. 14a, 15), and secretory globules (Figs. 17, 21a, b; morphology discussed below in detail). Cerebral ganglial depressions formed on the pretrochal hemisphere of the apical cap, and they persisted through metamorphosis (Figs. 14a, 17, 19a, 22a, b, 23a, b, 24a, 27, 28; development discussed below in detail).

The trunk region of the larvae was unciliated. The trunk gave rise to definitive ectodermal structures, such as cuticle and epidermis. A ciliated stomodeum formed on the ventral side of the unciliated ectodermal trunk directly below the apical cap (Figs. 14b, 20a, b, 27b, 28a).

The caudal region was covered with short cilia. Globules, similar to those of the pretrochal hemisphere of the apical. cap, were present (Figs. 14b, 18). The cilia at the periphery and the center of the caudal region were composed of compound cilia, and the cilia were longer (ca. 15 [micro]m) than on the fiat bottom surface (ca. 10 [micro]m) (Figs. 14b, 18).

Larvae aged 4 to 6 days. The post-trochal region of the apical cap narrowed and the cerebral depressions became deeper as the trunk grew longer (Figs. 19, 27c). A wide midventral longtitudinal band (15-20 [micro]m across) of the trunk region of the larvae became ciliated, forming a foot (Figs. 19, 27b). At the animal end of the long foot, a pedal pit formed sharp-ended cilia, longer (3-4 [micro]m) than the cilia on the foot, which had blunt ends (1-2 [micro]m) (Fig. 19b). Epidermal papillae formed covering the entire trunk region (Fig. 19a, morphology discussed later in detail).

Larval morphology

Secretory organs. The function of the globules that covered the pretrochal hemisphere of the apical cap and the entire surface of the caudal region (Figs. 14a, b, 17, 18) appears to be secretory, because the cilia-free membranes covering the globules were often ruptured (Fig. 21a, b). The globules appeared iridescent under the light microscope. The globules disappeared at metamorphosis, as the apical cap and the caudal region were withdrawn into the trunk (Fig. 24b).

One to two hundred small epidermal papillae appeared on the trunk (Figs. 19a, 23a, 24b). Like the globules mentioned above, these papillae are suspected to be secretory in function. Unlike the globules, the papillae persisted after metamorphosis (Figs. 24b, 25a). Protonephridia were not observed in the light micrographs of semithin sections.

Nervous system. The cells underneath a pair of depressions on the pretrochal region of the apical cap on the future ventral side of the adult (Figs. 14a, 17, 19a, 22a, 23a, b, 24a, 27, 28) became internalized to differentiate into definitive cerebral ganglia (Fig. 22b). These depressions were free of white globules, but the surface ciliation was not different from that of the rest of the apical area (Figs. 17, 22a). Pedal ganglia arose (Fig. 22b); however, the process by which they were formed is uncertain. Development of terminal sense organs was not apparent, although may have been present.

Metamorphosis

Larvae aged 7 to 8 days: beginning of metamorphosis. In spite of active ciliary movement of the prototroch and apical tuft, larvae at this point were no longer able to swim, and they sank to the bottom of the culture jars. The unciliated trunk grew longer, the epidermal papillae increased in number and size, and presumptive spicules formed (Fig. 23c, d). Spicules seemed still to be under the cuticle just along the foot groove and beneath the apical cap (Fig. 23). Although the post-trochal region of the test narrowed as the trunk grew, lengthening of the trunk appeared to be due to proliferation of definitive ectodermal trunk cells and not to any material supplied by the larval test (Figs. 23a, 27; later discussed in the programmed cell death determination of the test). Larval structures, such as the apical tuft and telotroch, were diminished. The apical cap and the prototroch occasionally became withdrawn into the trunk (Figs. 23b, 27d, e, 28b).

Larvae aged 9 to 11 days: completion of metamorphosis. The trunk was now completely covered with epidermal papillae and spicules that extended well beyond the cuticle (Fig. 24a). The whole caudal region and the cap became enclosed within the trunk and covered by the posterior and anterior extensions of definitive ectoderm (Fig. 24b).

The post-metamorphic juveniles crawled on the ciliated foot located in the midventral pedal groove. A rudiment of the adult anterior atrium developed as an invagination above the mouth. In adults, the invagination was often open, with the anterior end and half the length of the entire body raised above the substrum. The juveniles seemed to be sensitive to vibration, but not to light.

The juveniles grew more transparent as the original yolky yellow color became internally restricted. The animals lived for up to 1 month without any particulate food source, and the gut was not differentiated in histological sections during the period.

Development of hollow spicules. Juvenile spicules are different from adult spicules in shape and distribution. The spicules of adults are solid and bladelike along the foot groove and near the mouth, but the rest of the body is covered by hollow, needlelike spicules forming a crisscrossed meshwork within the cuticle. In juveniles, however, flat bladelike spicules were distributed evenly over the entire body along with upright and needlelike spicules (Fig. 25a, c). The juvenile pedal spicules were flat and bladelike as in the adults (Fig. 25b, c), and the head region also was covered with flat, bladelike spicules (Fig. 25d).

The development of hollow spicules in early juveniles was observed for the first time in aplacophorans (Fig. 26) by observing within the cuticle of live larvae on a glass slide using polarized light microscopy. The first part of the spicules to be secreted was the solid distal tip (Fig. 26a). The spicules continued to grow at the proximal end, becoming hollow with an open end (Fig. 26b). The proximal end finally closed to form a closed-ended hollow spicule with solid base (Fig. 26c). This type of hollow spicule formation is known to continue throughout the growth of adult neomenioids (Hoffman, 1949).

Morphology and fate of apical cells

Hoechst nuclear staining. Hoechst nuclear staining showed that cells of the apical cap were much larger than cells of the trunk region and stayed relatively constant in size (Fig. 27). It also showed that the number of the cells in the apical cap decreased as the apical cap degenerated, while the cells in the trunk region decreased in size and increased in number through proliferation (Fig. 27).

Programmed cell death determination. Cell viability and apoptotic index of apical cells were determined using programmed cell death (PCD) staining. At a stage as early as the first day of hatching, the entire pre-oral region of the embryos, including the prototrochal cells, stained orange (Fig. 28a), thus indicating PCD. The cerebral ganglial depressions, however, remained green and thus viable (Fig. 28a). The definitive ectodermal trunk also remained green and viable (Fig. 28a). The caudal region stained orange (Fig. 28a), indicating that the caudal cells also were degenerating. The pre-oral region of larvae at later stages, 5 to 7 days old, stained in brighter orange to red as it became withdrawn into the definitive ectoderm (Fig. 28b), indicating that these cells entered later stages of apoptosis. The caudal telotroch, partially withdrawn, also stained brighter orange. The elongated ectodermal trunk still remained green, indicating viable cells, but the epidermal papillae stained in orange to red, presumably because they are secretory and undergoing rapid turnover and cell death.

Discussion

Observations and remarks on reproduction and development

As observed in this study, reproduction and spawning in Epimenia babai were mostly consistent with Baba's descriptions (1938, 1940, 1951). E. babai is hermaphroditic, and although the time of sperm transfer is not known, fertilization is assumed to have occurred internally while the posterior ends of animals were entwined. This behavior has also been reported in E. australis (Scheltema and Jebb, 1994). However, an observation that one individual isolated for more than 90 days laid viable fertilized eggs suggests either that E. babai can self-fertilize, or that sperm were held for that period in the seminal receptacles.

Although brooding was not observed in E. babai during this study, the capacity to keep embryos within the posterior mantle cavity in fast water currents cannot be refuted. Brooding of larvae in the posterior mantle cavity was previously reported in E. babai (Baba, 1938, 1951), as well as in E. australis (Scheltema and Jebb, 1994), Halomenia gravida (Heath, 1918), and some perimeniid (= "pruvotinid") neomenioids (Thiele, 1913; Salvini-Plawen, 1978). During this study, E. babai individuals tightly curled their posterior part around the alcyonarian corals, thus pressing closed the mantle cavity opening and keeping their eggs within the mantle cavity. In their natural habitat, E. babai and other species of Epimenia live and feed on the alcyonarian corals that prefer strong swift water currents (Baba, 1938, 1940, 1951; Salvini-Plawen, 1972; Salvini-Plawen and Benayahu, 1991; Scheltema and Jebb, 1994). Therefore, brooding may take place in the natural habitat.

Homology of tests among neomenioid aplacophorans

On the basis of similarity in cell type, ontogeny, ciliation pattern, and cell fate of the apical cap, the larva of E. babai is here proposed to have a test-like apical cap with cells similar to the test cells of larvae of other neomenioid aplacophorans: Neomenia carinata, Nematomenia banyulensis, and Rhopalomenia aglaopheniae (Pruvot, 1890, 1892; Thompson, 1960) (Fig. 29a). The apical cap of E. babai and the tests of the other neomenioids are composed of large yolky cells that are larval in fate and pre-oral in origin, are covered with uniform short cilia, have a row of prototrochal cells and an apical tuft, and have cerebral ganglia depressions that persist through metamorphosis. The "test" of E. babai is less obvious because its cells are much smaller than those of other neomenioid tests, thus leaving the developing definitive ectodermal trunk underneath more exposed (Fig. 29).

The apical cap cells of the E. babai larvae are entirely apoptotic and, like the tests of other neomenioid aplacophorans, presumably do not contribute to development of any definitive adult structures except for the cerebral ganglia. The tests of Nematomenia banyulensis and Rhopalomenia aglaopheniae were said by Pruvot (1890, 1892) to be cast off and discarded at metamorphosis. The test cells of Neomenia carinata are absorbed by the animal at metamorphosis, as are the apical cap cells of E. babai; they degenerate and in N. carinata are known to be the main food reserve of post-larvae (Thompson, 1960). The fate of the cells in the brooded larvae of Halomenia gravida is not clear, although Heath (1918) observed in histological sections that they become reduced in size at later stages, indicating resorption.

There are some differences among neomenioid aplacophoran species in the number of "test" cells and in number of pairs of cerebral ganglia that proliferate from the test (Table 2). Test cells are arranged in six regular tiers of cells in Nematomenia banyulensis and Rhopalomenia aglaopheniae (Pruvot, 1890, 1892) and in five regular tiers of cells in Neomenia carinata (Thompson, 1960), whereas the apical cap cells of E. babai are numerous and irregularly arranged. The number of cerebral ganglia invaginations also varies: there is one pair in E. babai, Nematomenia banyulensis and R. aglaopheniae (Pruvot, 1890, 1892), whereas there are three pairs in Neomenia carinata, the third pair corresponding to the pedal ganglia (Thompson, 1960). In Nematomenia banyulensis and R. aglaopheniae, the pedal ganglia proliferate and split off from the cerebral ganglia internally (Pruvot, 1890, 1892). The process of pedal ganglia formation was not clear for E. babai.

Phylogenetic affinities of pericalymma larvae

Tests or test-like structures also occur in other molluscan taxa. Diverse forms of tests are described from six protobranch species: Acila castrensis (Zardus and Morse, 1998), N. delphinodonta (Drew, 1901), Nucula proxima (Drew, 1899a), Solemya reidi (Gustafson and Reid, 1986), S. velum (Gustafson and Lutz, 1992), and Yoldia limatula (Drew, 1899b). Protobranch bivalve tests are composed of large yolky cells, and the tests engulf the larvae entirely (N. delphinodonta, S. reidi, S. velum, and Y. limatula) or only partly (A. castrensis and N. proxima). The protobranch tests are larval and are cast off and ingested through the mouth at metamorphosis. The ciliation pattern of protobranch bivalve larvae is diverse: A. castrensis, N. proxima, and Y. limatula have an apical tuft and three rows of ciliary bands, whereas N. delphinodonta, S. reidi, and S. velum have no apical tuft and no ciliary bands. The tests of A. castrensis, N. delphinodonta, N. proxima, and Y. limatula are composed of five rows of cells, whereas the tests of S. reidi and S. velum are composed of nine. The caudal organ also seems to be diverse among protobranchs: A. castrensis and S. reidi have a ciliated caudal organ, N. proxima has an unciliated caudal organ, and S. velum and Y. limatula have no caudal organ.

Test-like structures are also found among trochophore larvae of scaphopods, described as having an apical tuft and three or six ciliary bands (Lacaze-Duthiers, 1856; Geilenkirchen et al., 1970; Guerrier et al., 1978; Wanninger and Haszprunar, 2001). No ciliated caudal structure has been reported. The test of scaphopods, in later developmental stages, becomes pushed anteriorly to form a velum (Lacaze-Duthiers, 1856). The scaphopod velum has been homologized with the lamellibranch bivalve velum. However, the scaphopod velum functions solely in locomotion, as in the pericalymma tests, whereas the lamellibranch bivalve velum also functions in feeding, and the homology is not certain.

Polyplacophorans (Grave, 1932; Eernisse and Reynolds, 1994), archaegastropods (Patten, 1886; Kessel, 1964), and chaetoderm aplacophorans (Nielsen, 2001) have trochophore larvae with a ciliated circular velum. Polyplacophoran vela are composed of one to three rows of prototrochal bands (Grave, 1932; pers. obs.). The ventral side of the ciliated apical region of polyplacophorans becomes absorbed at metamorphosis, while the dorsal side of the apical region gives rise to adult features such as the anterior-most shell valve and spicules (Grave, 1932). The archaegastropod vela also have three rows of prototrochal bands, which function in both locomotion and feeding. There seem to be several rows of ciliary bands on the chaetoderm aplacophoran vela; however, lack of complete developmental descriptions of the group limits speculation.

The question of the ancestral larval type among molluscs has been important in many discussions of their origin and diversification. The pericalymma of neomenioid aplacophorans and protobranch bivalves has been regarded as plesiomorphic by some authors (e.g., Drew, 1901; Thompson, 1960; Salvini-Plawen, 1972, 1973, 1980), although the trochophore has been generally considered plesiomorphic (Haszprunar et al., 1995; Rouse, 1999) and the pericalymma as derived. It has also been suggested that the lamellibranch velum could have arisen from the coalescence of pericalymma test (Drew, 1899b, 1901). However, there are as yet neither sufficient morphological data nor any cell lineage data to resolve the question of homology among the tests and homology of the tests to the velum.

Trochophore larvae seem to have gone through many modifications, not only within the clade Mollusca, but also in other spiralian taxa. Test-like structures are also found among lecithotrophic larvae of non-molluscan spiralians: the "serosa" of the sipunculid Sipunculus nudus (Rice, 1988); the "endolarva" of the polychaete Phyllodoce mucosa (Cazaux, 1970) and other Phyllodocidae spp. (Dawydoff, 1959); the "Iwata's larva" of a heteronemertean (Korschelt, 1936; Iwata, 1958; Jagersten, 1968); and the lecithotrophic nemertean larvae of Emplectonema gracile (Delsman, 1915), Geonemertes australiensis (Hickmann, 1963), and Tetrastemma candidum (Maslakova and Malakhov, 1999).

Given that test-like structures occur across various distant taxa within Spiralia that have lecithotrophic larvae, that their tests are composed of yolk cells, and that the tests are merely an enlarged pre-oral region of a trochophore larva, the tests could have arisen independently in each clade by adaptation to lecithotrophy. Homology among larval tests within molluscs and among spiralians and the pattern in which they evolved within the clade can probably be ascertained by more detailed developmental data analyzed within a well-corroborated phylogenetic framework. However, the phylogeny of Mollusca is controversial; cladistic analysis of a limited number of morphological characters (Salvini-Plawen and Steiner, 1996; Waller, 1998; Haszprunar, 2000) and available molecular sequence data on 18S rRNA (Winnepenninckx et al., 1996) have not yet resolved the issue. Larval development of other neomenioids and the Chaetodermomorpha needs to be studied in order to understand the diversity of developmental processes among aplacophorans. In addition, cell lineage tracing would help to ascertain the exact ontogeny of the tests among molluscs and other spiralians. Furthermore, developmental descriptions of monoplacophoran larvae, which are entirely lacking, are desired.
Table 1

Approximate timing of early development of Epimenia babai at 16-18
[degrees]C and ambient temperature (26 [degrees]C)

Developmental stage  16-18 [degrees]C  26 [degrees]C

Spawning                   0 h              0 h
2-cell                     2 h              1 h
4-cell                     4 h             2.5 h
8-cell                     6 h             3.5 h
16-cell                    8 h             4.5 h
32-cell                    10 h           5.5-6 h
Blastula                   15 h            7-9 h
Gastrula                 20-24 h          10-13h
Trochophore                48 h            24 h
Hatching                   3 d              3 d
Metamorphosis            11-13 d           5-7 d
Table 2

Modes of development of neomenioid aplacophorans

                                                              Test
                                                           Morphology


                                  Egg
                               diameter    Mode of larval  Apical
                               ([micro]m)     nutrient      tuft

Nematomenia banyulensis (1)     110-120    lecithotrophic    +

Rhopalomenia aglaopheniae (1)     260      lecithotrophic    +

Halomenia gravida (2)             320      lecithotrophic    ?
                                             (brooded)
Neomenia carinata (3)              ?       lecithotrophic    +

Epimenia babai (4)                250      lecithotrophic    +
                                             (brooded?)

                                            Test Morphology

                                           No. pairs of
                                No. cell    ectodermal     No. rows
                                rows in      cerebral       of test
                               prototroch  depressions       cells

Nematomenia banyulensis (1)        1            ?              6

Rhopalomenia aglaopheniae (1)      1            ?              6

Halomenia gravida (2)              ?            ?              ?

Neomenia carinata (3)              1            3              5

Epimenia babai (4)                 1            1        no regularity


                                  Test
                               Morphology



                               No. cells              Telotroch/
                                in test   Test fate  caudal organ

Nematomenia banyulensis (1)       56      cast off?   ciliated
                                                      telotroch
Rhopalomenia aglaopheniae (1)      ?      cast off?   ciliated
                                                      telotroch
Halomenia gravida (2)              ?          ?       ciliated
                                                      telotroch
Neomenia carinata (3)             ~80     resorbed    ciliated
                                                      telotroch
Epimenia babai (4)               >100     resorbed    ciliated
                                                      telotroch

(1)Pruvot, 1890, 1892

(2)Heath, 1918

(3)Thompson, 1960

(4)Baba, 1938, 1940, 1951, present study.


Acknowledgments

I thank Dr. Ame1ie Scheltema and Kikutaro Baba for guidance in working with aplacophorans. I also thank Dr. Mark Martindale for assistance and support in determining cell fate, and Dr. John Zardus for advice in SEM and sectioning techniques. I thank Dr. Satoshi Nojima and students at Amakusa Marine Biological Laboratories and the fishermen Mr. Kawamoto and Mr. and Mrs. Togawa for their cooperation while collecting in Japan. This work benefited from discussions and comments given by Drs. Amelie Scheltema, Gonzalo Giribet, Damhnait McHugh, Thomas Dahlgren, and Kenneth Halanych. Finally, I thank the reviewers for their helpful comments. The research was supported by a Putnam Grant in animal systematics from the Museum of Comparative Zoology, Harvard University, a National Science Foundation (NSF) Dissertation Improvement Grant in Systematic Biology (#0073312), and an NSF PEET grant (#9521930). This is contribution No. 10754 of the Woods Hole Oceanographic Institution.

Received 25 July 2001; accepted 30 May 2002. Email: aokusu@oeb.harvard.edu

Literature Cited

Baba, K. 1938. The later development of a solenogastre, Epimenia verrucosa. J. Dep. Agric. 6: 21-41.

Baba, K. 1940. The early development of a Solenogastre, Epimenia verrucosa (Nierstrasz). Annot. Zool. Jpn. 19: 107-113.

Baba, K. 1951. General sketch of the development in a Solenogastre, Epimenia verrucosa (Nierstrasz). Misc. Rep. Res. Inst. Nat. Resour. (Tokyo) 19-21: 38-46.

Baba, K. 1999. Solenogastres = Neomeniina, Ventroplicida). Pp. 55- 107 In Mollusca I. (Nakayama Shoten, Tokyo, Japan). [In Japanese.]

Cazaux, C. 1970. Recherches sur l'ecologie et le developpement larvaires des Polychetes de la region d' Arcachon, Vol. 295. These Fac. Sci. Univ. Bordeaux, France.

Dawydoff, C. 1959. Ontogenese des Annelides. Pp. 594-686 In Traite de Zoologie, P. Grasse, ed. Masson, Paris.

Delsman, H. C. 1915. Eifurchung and Gastrulation bei Emplectonema gracile Stimpson. Tijdschr, Ned. Dierk. Ver. 14: 68-109.

Drew, G. A. 1897. Notes on the embryology, anatomy, and habits of Yoldia limatula, Say. Johns Hopkins Univ. Circ. 17: 11-14.

Drew, G. A. 1899a. Some observations on the habits, anatomy and embryology of members of the Protobranchia. Anat. Anz. 15: 493-519.

Drew, G. A. 1899b. The anatomy, habits, and embryology of Yoldia limatula, Say. Mem. Biol. Lab. Johns Hopkins Univ. 4: 1-37.

Drew, G. A. 1901. The life-history of Nucula delphinodonta (Mighels). Quart. J. Microsc. Sci. 44: 313-392.

Eernisse, D. J., and P. D. Reynolds. 1994. Polyplacophora. Pp. 55-110 in Microscopic Anatomy of invertebrates, Vol. 5: Mollusca I, F. W. Harrison and A. J. Kohn, eds, Wiley-Liss, New York.

Geilenkirchen, W. L. M., N. H. Verdonk, and L. P. M. Timmermans. 1970. Experimental studies on morphogenetic factors localized in the first and the second polar lobe of Dentalium eggs. J. Embryol. Exp. Morphol. 23: 237-243.

Gotting, K. J. 1980. Origin and relationships of the Mollusca. Z. Zool. Syst. Evoltionsforsch. 18: 24-27.

Grave, B. H. 1932. Embryology and life history of Chaetopleura apiculata. J. Morphol. 54: 153-160.

Guerrier, P., J. A. M. v. D. Biggelaar, C. A. M. v. Dongen, and N. H. Verdon. 1978. Significance of the polar lobe for the determination of dorsoventral polarity in Dentalium vulgare (da Costa). Dev. Biol. 63: 233-242.

Gustafson, R. G. 1987. Phylogenetic implications of the molluscan pericalymma larva. Am. Zool. 27: 301A.

Gustafson, R. G., and R. A. Lutz. 1992. Larval and early post-larval development of the protobranch bivalve Solemya velum (Mollusca: Bivalvia). J. Mar. Biol. Assoc. UK 72: 383-402.

Gustafson, R. G., and R. G. B. Reid. 1986. Development of the pericalymma larva of Solemya reidi (Bivalvia: Cryptodonta: Solemyidae) as revealed by light and electron microscopy. Mar. Biol. 93: 411-427.

Gustafson, R. G., and R. G. B. Reid. 1988a. Association of bacteria with larvae of the gutless protobranch bivalve Solemya reidi (Cryptodonta: Solemyidae). Mar. Biol. 97: 389-401.

Gustafson, R. G., and R. G. B. Reid. 1988b. Larval and post-larval morphogenesis in the gutless protobranch bivalve Solemya reidi (Cryptodonta: Solemyidae). Mar. Biol. 97: 373-387.

Haszprunar, G., L. v. Salvini-Plawen, and R. M. Rieger. 1995. Larval planktotrophy--a primitive trait in the Bilateria? Acta Zool. 76: 141-154.

Haszprunar, G. 2000. Is the Aplacophora monophyletic? A cladistic point of view. Am. Malacol. Bull. 15: 115-130.

Heath, H. 1918. Solenogastres from the eastern coast of North America. Mem. Mus. Comp. Zool. Harv. 45: 185-263.

Hickman, V. V. 1963. The occurrence in Tasmania of the land nemertine, Geonemertes australiensis Dendy, with some account of its distribution, habits, variations and development. Pap. Proc. R. Soc. Tasmania 97: 63-75.

Hoffman, S. 1949. Studien uber das Integument der Solenogastren, nebst Bemerkungen uber die Verwandtschaft zwischen den Solenogastres und Placophoren. Zool. Bidr. Uppsala 27: 293-427.

Ivanov, D. L. 1996. Origin of Aculifera and problems of monophyly of higher taxa in molluscs. Pp. 59-65 in Origin and Evolutionary Radiation of the Mollusca, J. Taylor, ed. Oxford University Press, Oxford.

Iwata, F. 1958. On the development of the nemertean Micrura akkeshiensis. Embryologia 4: 103-131.

Jagersten, G. 1968. Livcykelns Evolution hos Metazoa. Scandinav. University Books, Lund, Sweden.

Kessel, M. M. 1964. Reproduction and larval development of Acmea testudinalis (Muller). Biol. Bull. 127: 294-303.

Korschelt, E. 1936. Vergleichende Entwicklungsgeschichte der Tiere. G. Fischer, Jena.

Lacaze-Duthiers, F. J. H. 1856. Historie de l'organisation et du developpement du Dentale. Ann. Sci. Nat. Zool. Biol. Anim. 4: 171-255.

Martin, S. J., C. P. M. Reutelingsperger, and D. R. Green. 1996. A specific probe for apoptotic cells. Pp. 107-120 in Techniques in Apoptosis: A User's Guide, T. G. Cotter and S. J. Martin, eds. Portland Press, London.

Maslakova, S. A., and V. V. Malakhov. 1999. Hidden larva in the Hoplonemertini order of nemerteans. Dokl. Akad. Nauk. 366: 849-852.

Nielsen, C. 2001. Animal Evolution: Interrelationships of the Living Phyla. Oxford University Press, Oxford.

Patten, W. 1886. The embryology of Patella. Arb. Zool. Inst. Wien 6: 149-174.

Pruvot, G. 1890. Sur le developpement d'un solenogastre. C. R. Acad. Sci. Paris 3: 689-692.

Pruvot, G. 1892. Sur l'embryogenie d'une Proneomnenia. C. R. Acad. Sci. Paris 114: 1211-1214.

Rice, M. E. 1988. Observations on development and metamorphosis of Siphonosoma cumanense with comparative remarks on Sipunculus nudus (Sipuncula, Sipunculidae). Bull. Mar. Sci. 42: 1-15.

Richardson, K. C., L. Jarrett, and E. H. Finke. 1960. Embedding in epoxy resins for ultrathin sectioning in electron microscopy. Stain Technol. 35: 313-323.

Rouse, G. W. 1999. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biol. J. Linn. Soc. 66: 411-464.

Salvini-Plawen, L. v. 1972. Cnidaria as food-sources for marine invertebrates. Cah. Biol. Mar. 13: 385-400.

Salvini-Plawen, L. v. 1973. Zur Klarung des "Trochophora"-Begriffes. Experientia 29: 1434-1436.

Salvini-Plawen, L. v. 1978. Antarktische und subantarktische Solenogastres (Eine Monographie: 1898-1974). Zoologica 128: 1-315.

Salvini-Plawen, L. v. 1980. Was ist eine Trochophora? Eine Analyse der Larventypen mariner Prostomier. Zool. Jahrb. Abt. Anar. Ont. Tiere 103: 389-423.

Salvini-Plawen, L. v. 1985. Early evolution and the primitive groups. Pp. 59-150 in The Mollusca: Evolution, E. R. Trueman and M. R. Clarke, eds. Academic Press, Orlando, FL.

Salvini-Plawen, L. v. 1990. Origin, phylogeny and classification of the Phylum Mollusca. Iberus 9: 1-33.

Salvini-Plawen, L. v. 1997. Systematic revision of the Epimeniidae (Mollusca: Solenogastres). J. Molluscan Stud. 63: 131-155.

Salvini-Plawen, L. v., and Y. Benayahu. 1991. Epimenia arabica spec. nov. a Solenogaster (Mollusca) feeding on the Alcyonacean Scleronephthya corymnbosa (Cnidaria) from shallow waters of the Red Sea. Mar. Ecol. 12: 139-152.

Salvini-Plawen, L. v., and G. Steiner. 1996. Synapomorphies and plesiomorphies in higher classification of Mollusca. Pp. 29-51 in Origin and Evolutionary Radiation of the Mollusca, J. Taylor, ed. Oxford University Press, Oxford.

Scheltema, A. H., and M. Jebb. 1994. Natural history of a solenogaster mollusc from Papua New Guinea, Epimenia australis (Thiele) (Aplacophora: Neomeniomorpha). J. Nat. Hist. 28: 1297-1318.

Strathmann, M. F. 1987. Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast. Univ. Washington Press, Seattle.

Thiele, J. 1913. Antarktische Solenogastren. Dtsch. Sudpolar-Exp. 1901-1903 14, Zool., Heft 1: 35-65.

Thompson, T. E. 1959. Development of the aplacophorous mollusc Neomenia carinata Tullberg. Nature 184: 122-123.

Thompson, T. E. 1960. The development of Neomenia carinata Tullberg. Proc. R. Soc. B 153: 263-278.

Waller, T. R. 1998. Origin of the molluscan class Bivalvia and a phylogeny of major groups. Pp. 1-45 in Bivalves: An Eon of Evolution--Palaeobiological Studies Honoring Norman D. Newell, P. A. Johnston and J. W. Haggart, eds. University of Calgary Press, Calgary, Alberta.

Wanninger, A., and G. Haszprunar. 2001. The expression of an engrailed protein during embryonic shell formation of the tusk-shell, Antalis entalis (Mollusca, Scaphopoda). Evol. Dev. 3: 312-321.

Winnepenninckx, B., T. Backelijau, and R. D. Wachter. 1996. Investigation of molluscan phylogeny on the basis of 18s rRNA sequences. Mol. Biol. Evol. 13: 1306-1317.

Zardus, J. D., and M. P. Morse. 1998. Embryogenesis, morphology and ultrastructure of the pericalymma larva of Acila castrensis (Bivalvia: Protobranchia: Nuculoida). Invertebr. Biol. 117: 221-244.
COPYRIGHT 2002 University of Chicago Press
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2002 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Okusu, Akiko
Publication:The Biological Bulletin
Geographic Code:1USA
Date:Aug 1, 2002
Words:6499
Previous Article:Embryonic velar structure and function of two sibling species of Crepidula with different modes of development.
Next Article:Undifferentiated cells in the snail myocardium are capable of DNA synthesis and myodifferentiation.
Topics:

Terms of use | Privacy policy | Copyright © 2019 Farlex, Inc. | Feedback | For webmasters