Abstract

This study describes the processes of embryogenesis, shell formation and larval development of the winged pearl oyster, Pteria penguin. Broodstock were induced to spawn using the standard method of thermal stimulation and fertilized eggs were incubated at a density of 50 ml−1 and a temperature of 27 ± 1°C. After an incubation period of 24 h, shelled larvae were stocked at 3 ml−1 and fed a mixed microalgae diet until reaching settlement age. Embryos and larvae were sampled periodically for examination by scanning electron microscope. The resulting high-resolution images were used to record the timing of developmental stages including the first cleavage (1 h postfertilization, hpf), morula (2.5 hpf), blastula (4.5 hpf), gastrula (5.5 > hpf), trochophore (7 > hpf), D-stage (20–22 hpf), prodissoconch II (3–6 days posthatching, dph), umbone (10–12 dph) and pediveliger (22 dph). Comparison with other oviparous oyster species revealed a similar sequence of key events, with differences occurring in the timing of developmental stages, shell structure and shell shape. This study is the first to describe early shell formation for a species belonging to the family Pteriidae. Shell development begins with formation a shell-field invagination (sfi) at the dorsoposterior end of the embryo (7 hpf), indicating the creation of a shell gland. The sfi stretches laterally to create a deep crevice (9 hpf) before eventually everting to form a flat central hinge linking the two expanding shell valves (12 hpf).

INTRODUCTION

Knowledge of early development of pearl oysters has become increasingly important as more commercial pearl farms choose to propagate juveniles for future pearl production, rather than rely on collection of either juveniles or adults from the wild. Hatchery propagation is now a necessity in regions with a prolonged history of pearl oyster exploitation (Southgate, 2008) and provides a basis for selective breeding programmes for genetically desirable traits (Kvingedal et al., 2010).

The majority of literature documenting early developmental stages in pearl oysters has focused on species belonging to the genus Pinctada (Wada, 1942; Minaur, 1969; Alagarswami et al., 1982, 1989; Rose & Baker, 1994; Doroudi & Southgate, 2003), while little information is available regarding embryonic and larval development of Pteria species. Pteria penguin (Röding, 1798) is traditionally used for the production of half-pearls (or ‘mabé’) in southern Japan, but the relatively simple methods required for pearl culture from this species have supported expansion of the industry to other localities in the Indo-Pacific (Arjarasirikoon et al., 2004; Strack, 2006; Southgate et al., 2008). Hatchery culture of P. penguin has recently been undertaken in areas with a low natural spat fall, with the aim of providing an adequate supply of oysters to permit large-scale commercial half-pearl production (e.g. Teitelbaum & Ngaluafe, 2008). An understanding of the processes of embryonic and larval development, particularly the timing of developmental stages, is central to the advancement of hatchery techniques for this commercially important species.

Knowledge regarding early development of P. penguin is also necessary to discover morphological characteristics that can be used to identify the species within plankton communities. This in turn will facilitate the investigation of patterns of dispersal, settlement and recruitment in wild populations (Gribben and Hay, 2003; Da Costa, Darriba & Martinez-Patino, 2008). A chronological description of early developmental stages of a bivalve can only be achieved by culturing the species.

This study describes embryonic and larval development in P. penguin with the aid of high-resolution imagery achieved using scanning electron microscopy (SEM). The images of embryogenesis and of early shell formation add to existing knowledge of these processes in bivalves. The findings of this study will form the basis for future research on hatchery culture of P. penguin and the monitoring of recruitment in wild populations.

MATERIAL AND METHODS

Thirty adult Pteria penguin with anteroposterior shell sizes of 155–237 mm were collected from Orpheus Island, off the north Queensland coast of Australia (18°36′24′′S, 146°29′10′′E). They were induced to spawn using the standard method of thermal stimulation (e.g. Rose & Baker, 1994; Southgate & Beer, 1997). Fertilized eggs were stocked at a density of 50 ml−1 in three replicate 500-l incubation tanks filled with UV treated, 1-µm filtered sea water (FSW) at a temperature of 27 ± 1°C (Southgate et al., 2008). The broad-spectrum antibiotic streptomycin sulphate was added to the incubation tanks at a concentration of 5 mg l−1 to reduce the proliferation of harmful bacteria (Wassnig & Southgate, 2011).

After 24 h of incubation, D-stage larvae were collected from each tank on a 25-µm nylon sieve mesh and transferred to 500-l larval rearing tanks containing FSW at a density of 3 larvae ml−1 (Southgate & Beer, 1997; Doroudi & Southgate, 2000). The static culture systems were drained, cleaned and refilled with FSW on day 4 and then every 2 days until larval settlement on day 22. Each tank received constant gentle aeration, while salinity was kept at 34–35 PSU and pH at 8.15–8.19. A temperature of 27 ± 1°C was maintained, which is within the range experienced by wild larvae during the peak spawning period in northern Australia (Beer & Southgate, 2000).

D-stage larvae were fed a 1:1 mixture of the golden-brown flagellates Isochrysis aff. galbana Tahitian and Pavlova sp. until day 12, when the diatom Chaetoceros muelleri was incorporated into the diet to provide a 1:1:1 ratio of the three species on the basis of cell numbers (Southgate, 2008). The daily feeding rate was 1,800 cells ml−1 on day 1 of larval culture, 3,000 ml−1 from days 2–4 and 5,000 ml−1 on day 3, after which the ration was increased by c. 1,000 cells ml−1 day−1 (Doroudi, Southgate & Mayer, 1999a; Doroudi & Southgate, 2000).

Embryos were sampled every 15 min during the first 3 h and then every half hour until D-stage larvae developed. Once reaching D-stage, larvae were sampled daily until settlement occurred. Samples were preserved in 2.5% glutaraldehyde in seawater at pH 8.15–8.19. Preserved embryos and larvae were prepared for examination using SEM by washing in a phosphate buffer, rinsing with distilled water and then dehydration through a graduated series of ethanol (Turner & Boyle, 1974). After dehydration, the samples were placed briefly in chloroform solution before being allowed to dry in a sealed glass Petri dish. The Petri dish contained filter paper soaked in chloroform so the samples could dry within a vaporous environment. Once dry, samples were mounted on aluminium stubs and coated with gold.

Standard descriptive terminology for embryogenesis and larval development of bivalve molluscs was used (e.g. Wada, 1942; Loosanoff & Davis, 1963; Kniprath, 1979, 1980; Eyster & Morse, 1984; Weiss et al., 2002). Growth during the larval period was monitored by measuring the shell length (anteroposterior measurement, APM) and height (dorsoventral measurement, DVM) of 20 individuals every 2 days.

RESULTS

Embryogenesis

Figure 1A shows a single sperm, with a round nucleus and broad acrosome, resting on the contoured surface of an egg. Newly fertilized eggs had a mean diameter of 45 ± 1.3 µm. Polar bodies became evident within 28 min of fertilization, indicating meiotic division (Fig. 1B). The first cleavage occurred at c. 1 h postfertilization (hpf), with the second cleavage following rapidly, resulting in four blastomeres (Fig. 1C). The blastomeres divided unequally, creating cells of various sizes, including macromeres and micromeres. The large macromere at the vegetal pole of eight-cell embryos (Fig. 1D) was roughly twice the diameter of neighbouring micromeres, which measured on average approximately 10 µm in diameter. This large macromere underwent a series of unequal divisions, eventually producing a daughter cell that divided bilaterally to create the two largest cells present in the 28-cell embryo (Fig. 1E). The morula stage occurred within 2.5 hpf and more than 70% of embryos had reached the blastula stage by 4.5 hpf (Fig. 1F). Embryos were oval at 5–6 hpf (Fig. 1G), before beginning to extend along their longitudinal axis to become conical with a broad anterior region and narrower posterior region (Fig. 1H). A large blastopore could be seen by 7 hpf, indicating that gastrulation had commenced (Fig. 1H). Cilia first appeared on the surface of young trochophores by 7 hpf and posterior cilia at the apical plate continued to elongate (15–20 µm long) and thicken during the following 2 h to form an apical tuft (Fig. 1I). A summary of the timing of developmental stages is given in Table 1.

Table 1.

Timing of the stages in embryonic development of five pearl oyster species (Pteriidae).

 Pinctada fucataPinctada maxima Pinctada margaritifera Pteria sterna§ Pteria penguin 
Temperature (°C) 27–29 27–29 28 24 26–28 
Polar body — — 24 min  28 min 
1st division 45 40 min — 90 min 60 min 
2nd division 60 min 60 min 2 h 4 h 75 min 
4th division 108 min — — 4 h 90 min 
Morula — 3 h — 4–5 h 2.5 h 
Blastula 6 h – — 5–6 h 4.5 h 
To gastrula 6.75 h 5 h 5 h 6–12 h 5–6 h 
Trochophore 11 h 7 h 8–12 h 12–16 h 7–14 h 
D-stage 20–21 h 18–24 h 24 h 21–24 h 18–22 h 
 Pinctada fucataPinctada maxima Pinctada margaritifera Pteria sterna§ Pteria penguin 
Temperature (°C) 27–29 27–29 28 24 26–28 
Polar body — — 24 min  28 min 
1st division 45 40 min — 90 min 60 min 
2nd division 60 min 60 min 2 h 4 h 75 min 
4th division 108 min — — 4 h 90 min 
Morula — 3 h — 4–5 h 2.5 h 
Blastula 6 h – — 5–6 h 4.5 h 
To gastrula 6.75 h 5 h 5 h 6–12 h 5–6 h 
Trochophore 11 h 7 h 8–12 h 12–16 h 7–14 h 
D-stage 20–21 h 18–24 h 24 h 21–24 h 18–22 h 
Figure 1.

Embryonic development of Pteria penguin. A. Single sperm on egg surface. B. Fertilized egg displaying polar body (0.5 hpf). C. Embryos after first and second divisions (1.25 hpf). D. Embryo after fourth division (1.5 hpf). E. Morula (2.5 hpf). F. Blastula (4.5 hpf). G. Gastrula (6 hpf). H. Early trochophore (7 hpf). I. Trochophore prior to shell formation (7.5 hpf). Abbreviations: at, apical tuft; b, blastopore; pb, polar body; pl, polar lobe.

Figure 1.

Embryonic development of Pteria penguin. A. Single sperm on egg surface. B. Fertilized egg displaying polar body (0.5 hpf). C. Embryos after first and second divisions (1.25 hpf). D. Embryo after fourth division (1.5 hpf). E. Morula (2.5 hpf). F. Blastula (4.5 hpf). G. Gastrula (6 hpf). H. Early trochophore (7 hpf). I. Trochophore prior to shell formation (7.5 hpf). Abbreviations: at, apical tuft; b, blastopore; pb, polar body; pl, polar lobe.

Early shell formation

Soon after the blastopore had begun to enlarge at the anterior end of the trochophore (7 hpf), the formation of a shell gland resulted in another depression at the dorsoposterior end, termed the shell-field invagination (sfi). At 7.5 hpf, the sfi was concave in shape and two to three times larger than the round blastopore (Fig. 2A). At 8–9 hpf the sfi began to extend along the dorsal surface of the posterior end of the trochophore, creating a deep, narrow crevice (Fig. 2B). Organic shell material (pellicle) extended across the aperture of the sfi and began to accumulate on either side of the central crevice, forming the beginnings of what would later develop into the two valves of the larval shell (Fig. 2C). The early shell material continued to expand forming a ‘saddle’ shape over the trochophore (11–12 hpf), during which time the central hinge depression everted and flattened (Fig. 2D–F). The central region between the two valves broadened and the shell valves thickened over the following 5–8 h, covering an increasing proportion of the trochophore and compressing it laterally (Fig. 2G, H). The previously conical trochophore became heart-shaped 14–16 hpf, before assuming the D shape common to newly hatched bivalve larvae. Early organic shell material appeared wrinkled at 17 hpf, suggesting that the process of calcification had not yet begun (Fig. 2I). Shell mineralization commenced once the periostracum covered the whole epithelial surface, giving rise to D-stage larvae with a calcified shell by 20–22 hpf.

Figure 2.

Early shell formation in Pteria penguin. A. Trochophore displaying initial sfi (8 hpf). B. Trochophore with elongated sfi (8.5 hpf). C. Early shell material (circled) on either side of the narrow crevice that will form the hinge (9 hf). D. Partially everted hinge region (10 hpf). E. Shell extended over the trochophore surface to form a ‘saddle’ shape (12 hpf). F. Developing shell and flat surface of hinge region (13 hpf). G. Spread of shell material over the surface of heart-shaped trochophore (14 hpf). H. Lateral compression of trochophore approaching D-stage (15 hpf). I. Trochophore completely covered with shell material before calcification. Abbreviations: b, blastopore; h, hinge; sfi, shell-field invagination.

Figure 2.

Early shell formation in Pteria penguin. A. Trochophore displaying initial sfi (8 hpf). B. Trochophore with elongated sfi (8.5 hpf). C. Early shell material (circled) on either side of the narrow crevice that will form the hinge (9 hf). D. Partially everted hinge region (10 hpf). E. Shell extended over the trochophore surface to form a ‘saddle’ shape (12 hpf). F. Developing shell and flat surface of hinge region (13 hpf). G. Spread of shell material over the surface of heart-shaped trochophore (14 hpf). H. Lateral compression of trochophore approaching D-stage (15 hpf). I. Trochophore completely covered with shell material before calcification. Abbreviations: b, blastopore; h, hinge; sfi, shell-field invagination.

Larval development

Within 1 h of hatching, D-stage larvae (Fig. 3A) were capable of utilizing their velum to swim actively through the water column and achieve a relatively even distribution within the tank. Velar retractor muscles attached to the shell at points near to the hinge. Larvae possessed a simple digestive tract, consisting of an oesophagus, stomach, digestive gland and intestine, all of which became more distinct with increasing shell size. Initial comarginal growth lines in the larval shell were visible by 3 days posthatching (dph) and prodissoconch I and II were clearly identifiable during the late D-stage (Fig. 3B). By 7 dph, D-stage larvae displayed hinge dentition with lateral tooth and socket joints present on the inner surface of both valves, adjacent to the central region of the hinge (Fig. 3C). A broken section of a D-stage shell is magnified in Figure 3D, to show the thin inner and outer prismatic shell layers and thicker homogenous granular layer in between. By 12 dph, more than 80% of larvae had become umbonate (Fig. 3E), possessing a cardinal tooth and adjacent socket directly below the umbo (Fig. 3F). Umbonate larvae developed an asymmetrical shell shape (Fig. 3G) as a result of skewed growth towards the posterior end of the deepest valve, which became increasingly pronounced as the larvae approached settlement (Fig. 3H). A red pigmented ‘eye spot’ could be seen through the partially transparent shell at c. 18 dph in fast-growing individuals. Figure 3I shows the outer shell edge of a pediveliger larva, where the innermost shell layer has peeled back during sample preparation to reveal the amorphous calcium carbonate matrix that makes up the bulk of the shell. Pediveligers were capable of using their foot to crawl actively on the tank surface, during which time slight gill ciliation became evident. A summary of larval development is provided in Table 2.

Table 2.

Timing (T) of the stages of larval development of five pearl oyster species (Pteriidae) and their mean size (S, in µm) at each stage.

 Pinctada fucata*
 
Pinctada maxima
 
Pinctada margaritifera
 
Pteria sterna§
 
Pteria penguin
 
Temperature (°C) 25–30 25–29.5 27–29 23 26–28 
Stage 
D-stage 21 h 67.5 18–24 h 85 24 h 80 21–24 h 48 18–22 h 83 
Early umbone 10 days — 8 days 110 8 days 110 14 days 82 8 days 107 
Umbone 12 days 135 10 days 114 12 days 141 18 days 192 12 days 129 
Eye spot 15 days 210 20 days 205 22 days 231 32 days – 20 days 226 
Pediveliger 20 days 230 22–24 days 230 22+ days — 29–35 days 259 22 days 233 
 Pinctada fucata*
 
Pinctada maxima
 
Pinctada margaritifera
 
Pteria sterna§
 
Pteria penguin
 
Temperature (°C) 25–30 25–29.5 27–29 23 26–28 
Stage 
D-stage 21 h 67.5 18–24 h 85 24 h 80 21–24 h 48 18–22 h 83 
Early umbone 10 days — 8 days 110 8 days 110 14 days 82 8 days 107 
Umbone 12 days 135 10 days 114 12 days 141 18 days 192 12 days 129 
Eye spot 15 days 210 20 days 205 22 days 231 32 days – 20 days 226 
Pediveliger 20 days 230 22–24 days 230 22+ days — 29–35 days 259 22 days 233 
Figure 3.

Larval development of Pteria penguin. A. Early D-stage larvae with calcified shell (1 dph). B. D-stage with prodissoconch I and II (6 dph). C. D-stage hinge of a single valve and a section of broken shell shown magnified in D (7 dph). D. Section of broken shell showing three shell layers. E. Early umbone stage (8 ph). F. Umbone hinge (12 dph). G. Late umbone stage (14 dph). H. Pediveliger (21 dph). I. Edge of pediveliger shell showing the peeling back of the inner prismatic layer to reveal the internal shell matrix. Abbreviations: acc, amorphous calcium carbonate; bs, section of broken shell; ct, central cardinal tooth; g, granular shell layer; ip, inner prismatic shell layer; op, outer prismatic shell layer; p, periostracum; s, socket opposing hinge tooth; t, hinge tooth.

Figure 3.

Larval development of Pteria penguin. A. Early D-stage larvae with calcified shell (1 dph). B. D-stage with prodissoconch I and II (6 dph). C. D-stage hinge of a single valve and a section of broken shell shown magnified in D (7 dph). D. Section of broken shell showing three shell layers. E. Early umbone stage (8 ph). F. Umbone hinge (12 dph). G. Late umbone stage (14 dph). H. Pediveliger (21 dph). I. Edge of pediveliger shell showing the peeling back of the inner prismatic layer to reveal the internal shell matrix. Abbreviations: acc, amorphous calcium carbonate; bs, section of broken shell; ct, central cardinal tooth; g, granular shell layer; ip, inner prismatic shell layer; op, outer prismatic shell layer; p, periostracum; s, socket opposing hinge tooth; t, hinge tooth.

Larval growth

Twenty-four hours after fertilization, D-stage larvae had a shell length (APM) (mean ± SE) of 83 ± 3.7 µm (Fig. 4) and height (DVM) of 63 ± 5.1 µm, of which c. 3.5% was attributed to prodissoconch II. Larvae grew in an approximately linear fashion during D-stage (APM: y = 8.42x + 74.6, R2 = 0.97; DVM: 4.80x + 59.9, R2 = 0.98) until an umbone was developed at 12 dph, when mean APM (±SE) and DVM were 129 ± 6.4 and 120 ± 4.5 µm, respectively (Fig. 4). Shell size increased rapidly between days 12 and 20, with a mean growth rate of 12.3 µm day−1 in APM and 11.2 µm day−1 in DVM (Fig. 4). Growth slowed after day 20 and by day 22 more than 70% of surviving larvae had reached the eyed pediveliger stage, with a mean APM and DVM of 233 ± 31.3 and 211 ± 25.1 µm, respectively (Fig. 4). Larvae showed a mean increase in APM of 7.2 µm day−1 and DVM of 7.0 µm day−1 during the 22-day larval period. Growth of Pteria penguin larvae in terms of shell size from initial D-stage to settlement is best modelled by the polynomial equations: APM = 0.39x2 − 2.09x + 92.3 (R2 = 0.97) and DVM = 0.27x2 − 0.14x + 72.0 (R2 = 0.96) (Fig. 4). The data for growth were heteroscedastic, with size becoming more variable with age (Fig. 4).

Figure 4.

Growth models for the shell of Pteria penguin larvae from D-stage on day 1 to settlement on day 22. Anteroposterior measurement (APM) (solid line) described by 0.39x2 − 2.09x + 92.3 (R2 = 0.97) and dorsoventral measurement (DVM) (dashed line) described by 0.27x2 − 0.14x + 72.0 (R2 = 0.96). Vertical lines represent the standard error at each sampling time.

Figure 4.

Growth models for the shell of Pteria penguin larvae from D-stage on day 1 to settlement on day 22. Anteroposterior measurement (APM) (solid line) described by 0.39x2 − 2.09x + 92.3 (R2 = 0.97) and dorsoventral measurement (DVM) (dashed line) described by 0.27x2 − 0.14x + 72.0 (R2 = 0.96). Vertical lines represent the standard error at each sampling time.

DISCUSSION

The high quality of images generated using SEM is partly attributed to the novel method used for drying samples prior to gold coating, i.e. rinsing samples with chloroform and allowing them to dry within a vaporous chloroform environment. The method was effective in eliminating nonevaporable deposits from the surface of samples and avoiding the shrinkage commonly encountered when using critical-point or air-drying techniques (Bozzola & Russell, 1999). Critical-point drying is widely used for preparation of fine structures for SEM (Turner & Boyle, 1974; Bozzola & Russell, 1999), but this study demonstrates the successful application of an alternative technique.

Pteria penguin eggs are roughly spherical and possess an uneven and highly contoured surface. This study provides the first high-magnification image of a sperm from a pteriid species. The sperm nucleus is ovoid in shape with a short broad acrosome situated at the tip. The morphology is similar to that of sperm from Crassostrea virginica (Ostreidae), but differs from the arched head and elongated acrosome seen in the families Dreissenidae, Mytilidae and Veneridae (Niijima & Dan, 1965; Mouëza, Gros & Frenkiel, 1999, 2006; McAnlis, Lynn & Misamore, 2010). The spiralian pattern of cleavage described for P. penguin is typical of that observed in other bivalve species (Henry & Martindale, 1999; Kin, Kakoi & Wada, 2009).

The timing of developmental stages during embryogenesis varies between pteriid species, although all complete the transition from fertilized egg to D-stage larva within 18–24 h of fertilization. This relatively rapid rate of embryogenesis is also seen in other tropical and subtropical oviparous oysters (Southgate & Lee, 1998; Kakoi et al., 2008). In contrast, temperate species generally require 32–48 h to reach D-stage (Fujita, 1929; Roughley, 1933; Loosanoff & Davis, 1963; Galstoff, 1964; Dinamani, 1973). This is in keeping with the importance of water temperature in explaining the decrease in developmental rates of marine invertebrate embryos with increasing latitude (Hoegh-Guldberg & Pearse, 1995). While it may be beneficial for oyster hatcheries to maximize water temperature and therefore rate of development, the threshold for normal development should not be exceeded (Dos Santo & Nascimento, 1985; Doroudi, Southgate & Mayer, 1999b). It is particularly important to maintain optimal water temperature during the early cleavage stages when bivalve embryos are most sensitive to mortality caused by temperature shock (Wright et al., 1983).

This study provides the first high-resolution record of early shell formation in a pteriid. The initial stages of shell formation during bivalve embryogenesis, whereby an sfi is formed when ectodermal cells migrate inwards to form the shell gland, are well understood (Kniprath, 1980; Waller, 1981; Eyster & Morse, 1984). Casse, Devauchelle & Le Pennec (1998) found that the microvilli-bearing cells that form the pore of the invagination in Pecten maximus are the cells responsible for pellicle secretion. The initial sfi can occur at various developmental stages in molluscs (Gros, Frenkiel & Mouëza, 1997) and there is conjecture as to whether the shell gland is typically first present before, during or after gastrulation (Waller, 1981). This study shows that in P. penguin, the sfi first becomes evident during gastrulation, following the appearance of a blastopore. As expected, the sfi appears sooner after fertilization in P. penguin than in bivalve species with a slower rate of embryonic development (Casse et al., 1998).

The concave shape of the initial sfi resembles that first observed in Spisula solidissima (Mactridae) (Eyster & Morse, 1984) although, rather than expanding, the sfi in P. penguin stretches laterally, creating a deep, narrow crevice similar to that described in Pecten maximus (Casse et al., 1998) and Saccostrea kegaki (Kakoi et al., 2008). Observations of shell secretion in S. solidissima suggest that bivalves develop two opposing shells by expanding their shell field over both sides of the trochophore in a saddle- or ribbon-shape (Eyster & Morse, 1984). This pattern has also been observed in the families Lucinidae (Gros et al., 1997), Veneridae (Mouëza et al., 1999, 2006) and Ostreidae (Kakoi et al., 2008) and this study confirms its occurrence in at least one member of the Pteriidae. SEM images show that the early central hinge region of P. penguin broadens and then folds outward to form a flat surface. The inner region of the shell gland in bivalves is thought to evert to form the mantle epithelium (Neff, 1972), followed by the onset of shell mineralization and calcification during the late trochophore stage (Eyster, 1986; Mouëza et al., 2006). In P. penguin shell calcification does not take place until the periostracum spans the entire epithelial surface. The ripples on the outer surface of the newly calcified D-stage shell recall the punctate–stellate pattern common on young bivalve shells (Waller, 1981; Eyster & Morse, 1984; Hayakaze & Tanabe, 1999).

This study is the second to observe a change in shape during the trochophore stage of a pearl oyster, suggesting it is a common phenomenon in the Pteriidae. Doroudi & Southgate (2003) observed that Pinctada margaritifera trochophores extend along their longitudinal axis, resulting in the anterior region becoming broader than the posterior region. There is a similar transformation from ovoid to conical in P. penguin, with the blastopore present at the centre of the broad anterior region while shell formation initiates on the dorsoposterior surface. Early shell material compresses the trochophore laterally, giving rise to a distinctive heart shape for the few hours preceding D-stage.

A larval period of 18–24 days is common in many oviparous oyster species, regardless of latitudinal distribution (Loosanoff & Davis, 1963; Dinamani, 1973; Gerdes, 1983; Buroker, 1985; Southgate & Lee, 1998; Kakoi et al., 2008). Oviparous oysters are thought to have evolved over a relatively narrow range of ecological conditions (Buroker, 1985), which may account for this similarity. The long planktonic stage enables gene flow over a wide geographical range (Grassle, 1972), particularly important where habitats are patchy and adults are sedentary (Strathmann, 1974; Jablonski & Lutz, 1983; Buroker, 1985).

Food supply during hatchery rearing of pearl oyster species is based on previous research determining optimal diet and feed ration (Southgate, 2008), leaving water temperature as the primary determinant of development time (Alagarswami et al., 1983; Araya-Nunez & Ganning, 1995; Doroudi et al., 1999b). Water temperatures in excess of 25°C result in an optimal larval period of 18–24 days in pearl oysters (Table 2), but a proportion of larvae will fail to maintain the rapid rate of development and effectively become trapped in early larval stages before eventually dying (Rose & Baker, 1994). This may contribute to the greater variation in size of P. penguin larvae approaching settlement age. Some hatchery operations utilize lower temperatures to maximize survival while prolonging time to settlement, exemplified by the comparatively slow rate of development shown by P. sterna larvae when cultured at a suboptimal water temperature of 23°C (Table 2) (Araya-Nunez & Ganning, 1995).

Shell structure, with daily growth lines and lateral hinge dentition, in D-stage P. penguin is consistent with patterns observed for other pteriid species (Rose & Baker, 1994; Doroudi & Southgate, 2003). This study shows a prominent cardinal tooth and socket in umbonate larvae. A similar hinge structure is present in juvenile and adult Pteria, described by Morton (1995) as a single tooth that interlocks between two smaller teeth on the opposing valve. Larvae of P. penguin possess the three shell layers common to many molluscs (Carriker & Palmer, 1979; Waller, 1981; Weiss et al., 2002): an outer prismatic layer below the periostracum, an inner prismatic layer adjacent to the mantle and a thick homogenous granular layer in between. Doroudi & Southgate (2003) did not distinguish between the inner prismatic and granular layers during their study on embryonic and early larval development of Pinctada margaritifera. The lack of a clear-cut boundary between these layers has also been observed in larvae of the temperate oyster Crassostrea gigas (Weiss et al., 2002). SEM analysis suggests that an inner prismatic layer is relatively prominent in P. penguin, but the outer prismatic layer is almost nonexistent in some parts.

Pteria penguin larvae possess a large flat umbo, rather than the conical umbo typical of Pinctada species (Rose & Baker, 1994; Doroudi & Southgate, 2003), and show skewed shell growth towards the posterior end of the deepest valve. This species typically settles in areas of strong current (Ito, 1999), where an asymmetrical shell shape may decrease resistance of the shell to fast-flowing water, therefore reducing susceptibility to detachment. A pronounced hinge is present in the adult to aid in supporting the asymmetrical shell shape, but this does not begin to develop until after larval settlement.

ACKNOWLEDGEMENTS

This project was funded by the Australian Centre for International Agricultural Research (ACIAR) as part of Project FIS/2006/172 ‘Winged Oyster Pearl Industry Development in Tonga’. We thank the staff at James Cook University's Marine and Aquaculture Research Facilities Unit (MARFU) and Orpheus Island Research Station (OIRS) for their help in facilitating hatchery operations, the staff at the Advanced Analytical Centre (AAC) for their assistance in operating the scanning electron microscope and the late Prof. Christopher Alexander for his expertise in sample preparation and particularly for his development of the drying technique used here.

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