Printer Friendly

On fertilization in Chaetopleura apiculata and selected Chitonida.


Sirenko (1993) presented evidence for classifying chitons into two Orders, Lepidopleurida and Chitonida (which includes most extant chitons), and subdivided Chitonida into two Suborders, Chitonina and Acanthochitonina; this classification has been supported by more recent studies (Buck-land-Nicks, 1995, 2006, 2008; and Sirenko, 2006), with some reservations on the monophyly of Acanthochitonina (Okusu et al., 2003). Lepidopleurida exhibit a series of plesiomorphic characters for shell morphology (van Belle, 1983; Kaas and van Belle 2003), egg and gill morphology (Eernisse, 1984; Sirenko, 1993), sperm and fertilization biology (Buckland-Nicks, 1995, 2006, 2008), and molecular sequences (Okusu et al., 2003; Lieb et al., 2006) and are therefore considered to be the basal Order of chitons.

The sperm of Chitonida, including Callochitonidae, are quite different from the typical aquasperm of Lepidopleurida, having an acrosome that is just a small vesicle at the tip of an elongated nuclear filament (Buckland-Nicks et al., 1988; Buckland-Nicks and Hodgson, 2000; Buckland-Nicks, 2006), and an acentric basal body comprising the two fused centrioles (Hodgson et al., 1988; Buckland-Nicks, 2008).

The eggs of Chitonida differ from those of Lepidopleurida, with the exception of Callochitonidae, in that the plesiomorphic jelly coat is replaced by a thinner, more resilient hull that usually is elaborated into a series of projections (Eernisse, 1984; Sirenko, 1993, 2006; Buckland-Nicks, 2008). Experiments have shown that these projections improve flotation and direct sperm to specific areas on the egg surface (Buckland-Nicks, 1993, 1995, 2006). The hulls of Chitonina eggs are characteristically elaborated into long spines with narrow bases (Sirenko, 1993). These spines themselves are closed, forcing the sperm to penetrate the egg between their polygonal bases (Buckland-Nicks, 1995, 2006, 2008). During fertilization, the chromatin is injected into the egg via a narrow tubular nuclear filament that appears to exclude other sperm organelles, including centrioles, mitochondria, and flagellum (Buckland-Nicks and Hodgson, 2000; Buckland-Nicks, 2006). This mechanism of fertilization is defined by a series of apomorphic characters that unify the order Chitonida, which recently has been shown to include the controversial family Callochitonidae as sister taxon to the rest of Chitonida (Buckland-Nicks, 2008). Implications for embryonic development in these chitons are discussed in light of new evidence from studies of fertilization in Mopalia muscosa that support the hypothesis that sperm centrioles and mitochondria are abandoned on the egg surface.

This study examines, for the first time, fertilization in Chaetopleura apiculata, bringing to light differences in sperm and egg structure compared to other selected Chitonina; these differences distinguish at least two groups within this Suborder.

Materials and Methods

Fertilization has been studied in representative species from two families of suborder Chitonina (classification by Sirenko, 2006): Ischnochitonidae and Chitonidae. The following three minor families have not been investigated: Callistoplacidae, Loricidae, and Schizochitonidae. Specimens of the following species were obtained: Acanthopleura granulata (Gmelin, 1791), collected off the Bahamas (25[degrees]02'N, 77[degrees]16'W) in June 1993; Chaetopleura apiculata (Say, in Conrad 1834), purchased from Gulf Specimen Co. and collected off the Florida coast, near Panacea, in May 1993 and June 2004; Rhyssoplax tulipa Quoy and Gaimard, 1835 (= Chiton tulipa Quoy and Gaimard, 1835), collected from East London, South Africa (32[degrees]54'S, 28[degrees]04'E), in September 1999; Radsia nigrovirescens de Blainville, 1825 (= Chiton nigrovirescens de Blainville, 1825), collected from the west coast of the Cape Peninsula, South Africa (34[degrees]16'S, 18[degrees]40'E), in September 1999; Mopalia muscosa (Gould, 1846), collected on San Juan Island, Washington State (48[degrees]2'N, 122[degrees]54'W), in May 2004. All animals were placed individually in small petri dishes containing seawater passed through a 0.45-[micro]m Millipore filter (MFSW). Males often spawned first, and if this occurred the sperm were removed and added to the other dishes. Confirmed males were dissected, and small pieces of testis were removed with fine forceps and placed in primary fixative. The presence of sperm in the water often stimulated females or other males to spawn. When this occurred the water was changed and the animal was washed thoroughly with seawater. Any fertilized eggs were saved for observation of normal cleavage. Females usually continued to spawn after washing, and these unfertilized eggs were collected in filtered seawater and fertilized separately. Sperm of the same species were obtained by dissecting out testes in filtered seawater and making a milky sperm suspension. Several drops of this suspension were added to each batch of eggs in 50 ml of MFSW. Fertilization was observed with differential interference contrast light microscopy. Eggs were fixed unfertilized and at intervals following fertilization of 1 min, 5 min, 1 h, and 2 h.

The primary fixative contained 50 ml of 0.2 mol [l.sup.-1] sodium cacodylate buffer (adjusted to pH 7.4) with 40 ml of MFSW containing 3 g of sucrose and 10 ml of 25% glutaraldehyde. After fixation on ice for 1 h, samples were allowed to come to room temperature and fixation was continued overnight. The next day, after two rinses in 0.1 mol [l.sup.-1] sodium cacodylate in seawater and 3% sucrose, secondary fixation was performed with 2% osmium tetroxide in the same buffer for 1 h, followed by dehydration in an ethanol series. The samples for scanning electron microscopy (SEM) were dehydrated in an ethanol series up to 100%, and then pipetted into Teflon flow-through vials (Cedar Lane Inc.) in 100% ethanol under a dissecting scope. The vials were capped and underwent critical-point-drying in a Samdri PVT-3B (Tousimis). Each vial was then uncapped and inverted on an SEM stub covered with a double-sided sticky carbon tab, such that the eggs adhered to the stub. Under a dissecting scope, some eggs were then rolled--using a stainless steel insect pin (000 gauge, Fine Science Tools Inc.) glued to a toothpick with 5-min epoxy cement--on the sticky tape to remove spines and expose fertilizing sperm. Finally, the eggs were coated with gold in a Polaron SC502 sputter coater and photographed in a JSM 5300 JEOL scanning electron microscope.

Samples for transmission electron microscopy (TEM) were processed through three changes of 100% ethanol and then exchanged through propylene oxide before infiltrating and embedding in a 1:1 mixture of Spurr and Eponate 812 resins. Polymerization in embedding molds was completed in a 60 [degrees]C oven for up to 2 days. In the case of Acanthopleura granulata, three fertilized eggs had been mounted and examined with SEM some years previously. A fine insect pin was used to gently flick these eggs off the stub into 100% ethanol. The eggs were then embedded in Spurr/Eponate resin mixture, as above, and sectioned. Except for the layer of gold, both thick and thin sections appeared relatively normal.

An RMC MT2C ultramicrotome was used to cut 1-[micro]m sections for light microscopy and silver-gold thin sections for TEM. Thick sections were stained for 15 s with 1% toluidine blue at pH 9. Thin sections were stained with aqueous uranyl acetate for 30 min, then with lead citrate for 5 min, before being washed in cold, boiled distilled water.


On fertilization in Chaetopleura apiculata

The egg hull of Chaetopleura apiculata (Ischnochitonidae: Chaetopleurinae) consists of highly branched hollow spines closed at the tip, extending from polygonal bases (Figs. 1, 2). The diameter of each egg is approximately 290 [micro]m, with spines that extend about 35 [micro]m from the egg surface, making a total diameter, including spines, of 360 [micro]m. The hull is perforated around the perimeter of the base of each spine by a series of open pores, ranging in diameter from 0.1 [micro]m to 0.5 [micro]m, which provide sperm with direct access to the vitelline layer(Figs. 3-5).


The sperm have a bullet-shaped nucleus, 5 [micro]m in length, with a filamentous nuclear extension also 5 [micro]m in length and 0.08 [micro]m in width, at the tip (Fig. 6). The acrosome vesicle is 0.25 [micro]m in length and 0.1 [micro]m in width and is homogeneous, without any evidence of stratification. The base of the nucleus has a flattened area on top of the basal body and a small posterior lobe that leads into a shallow nuclear fossa where the outer membrane is thickened (Fig. 6). Usually four mitochondria together with numerous glycogen granules are contained in a sac-like extension of the plasma membrane below the nucleus, forming a collar next to the flagellum. The membrane closest to the flagellum is thickened by deposits extending posteriorly from the annulus (Fig. 6). During spermiogenesis the proximal centriole fuses laterally and orthogonally to the distal centriole to form the basal body, which produces an acentric flagellum (Fig. 6). The annulus, as is typical for aquasperm, binds the distal centriole (of the basal body) to the plasma membrane (Fig. 6).

Released sperm must find a route between the complex egg spines to locate a pore in the hull (Figs. 3, 4). The nuclear filament is extended through the pore and down to the vitelline layer, where the acrosome digests a hole for it to enter and fuse with the egg. Some eggs had a few large pores up to 2 [micro]m in diameter that gave sperm access to the space beneath the hull, enabling direct penetration of the vitelline layer (Figs. 4, 5). Penetration of the sperm nuclear filament and subsequent fusion of sperm and egg membranes in C. apiculata causes a cortical granule reaction that is followed by the elevation of a fertilization membrane a few minutes later.

On fertilization in selected Chitonina

Rhyssoplax tulipa (= Chiton tulipa) (Chitonidae: Chitoninae). The eggs of Rhyssoplax tulipa have long spines terminating in petalloid tips (Figs. 7, 8). The egg is smaller than that of Chaetopleura apiculata, being about 160 [micro]m in diameter, with spines extending 20 [micro]m from the surface, making a total diameter including spines of 200 [micro]m. The hull is composed of an inner thick (0.7-1.0 [micro]m) amorphous layer and an outer thin (0.1 [micro]m) dense fibrous layer, which is continuous but thickened over the spines (Figs. 9, 11). The vitelline layer covers the egg membrane, which is formed into a series of microvilli.

Sperm, released in the water near to the eggs, swim down between spines and penetrate the hull, often near the junction between the polygonal bases of two adjacent spines (Figs. 8, 10). The acrosome reaction occurs on contact with the hull, resulting in the digestion of a quite large area of the dense layer, around the narrow nuclear filament (Fig. 11). A small pore is created in the vitelline layer, through which the sperm nuclear filament is extended to meet and fuse with an egg microvillus. Penetration of the egg by one sperm induces the cortical granule reaction, and a fertilization membrane is raised after a few minutes.

Radsia nigrovirescens (= Chiton nigrovirescens) (Chitonidae: Chitoninae). The egg hull of Radsia nigrovirescens is elaborated into long simple spines terminating in hooks (Figs. 12-14), quite different from those of Rhyssoplax tulipa and other species of Chitoninae. The diameter of each egg of R. nigrovirescens is larger than that of Chaetopleura apiculata, being about 404 [micro]m, with spines that are 38 [micro]m long, extending from the surface, making a total diameter including spines of 480 [micro]m. As in other Chitonina, the spines are closed so that sperm must penetrate the egg between them, although fertilization has not been observed in this species. This species differs from Rhyssoplax tulipa in that the outer dense layer of the hull appears to be absent, and a scattered series of pores, ranging in diameter from 0.1 [micro]m to 0.5 [micro]m, perforate the hull and may provide sperm access to the vitelline layer (Fig. 15).

The eggs are stored in the pallial grooves of mature females, where their hooked spines may interlock and cause them to become attached to each other, which happens readily in vitro (Fig. 13). Radsia nigrovirescens usually broods its young to the crawl-away stage inside the pallial grooves. However, one female was observed spawning individual eggs into a petri dish in the laboratory, suggesting that this species also has the ability to free-spawn.

The sperm nucleus and nuclear filament together are about 6 [micro]m in length, terminating in a small homogeneous acrosome vesicle 0.27 [micro]m long and 0.11 [micro]m wide (Fig. 16). In other respects, sperm morphology in R. nigrovirescens is similar to that of other species of Chitoninae: a long posterior collar surrounds the flagellum and contains the mitochondria and numerous glycogen granules. The proximal centriole is fused laterally to the distal centriole, creating an acentric basal body (Figs. 17, 18).

Acanthopleura granulata (Chitonidae: Acanthopleurinae). Acanthopleura granulata is unique in that it has three types of spinous projections of the egg hull. Long spines, intermediate spines, and short bifurcating spines can be found (Figs. 19, 20). The longer spines have scale-like structures on the surface (Fig. 20), perhaps similar to those at the tips of Chaetopleura spines. Sperm fertilize the eggs near junctions of the bases (Fig. 21), although sperm penetration does not happen exclusively in this location. In spite of a delay for several years in embedding and sectioning the fertilized eggs for TEM, fixation and cell continuity was quite good, revealing a continuous outer dense fibrous layer (clearly visible below the gold coating for the SEM), and an inner amorphous layer (Fig. 22). Sperm appeared to penetrate the hull directly without access to pores, as the dense layer was continuous over any pore-like depressions (Fig. 22).




Fate of sperm organelles during fertilization in Chitonida

At fertilization in all Chitonida examined (including Callochitonidae, Chitonina, and Acanthochitonina), the sperm injects the nuclear chromatin from the surface of the egg and does not appear to be engulfed itself (Buckland-Nicks, 2008). Examination of serial 1-[micro]m sections of fertilized eggs of Mopalia muscosa (Acanthochitonina: Mopaliidae) 2 h after fertilization repeatedly revealed a bag of membrane located on the egg surface and containing particles corresponding in size and number to the mitochondria of this species (Fig. 23).


Sperm morphology and phylogeny

The sperm of Chitonina, like that of all Chitonida, is bullet-shaped with an elongate nuclear filament capped by a minute acrosome (Buckland-Nicks et al., 1990). An evagination of the plasma membrane forms a collar, which contains the mitochondria and extends posteriorly, surrounding the flagellum in most species (see reviews by Buckland-Nicks, 1995, 2008). Hodgson et al. (1988) examined mature sperm of six species of Chitonina, including Rhyssoplax tulipa and Radsia nigrovirescens. It was not known at the time that chitons had a highly reduced acrosome, but results confirm this for Radsia nigrovirescens (this study) and for Chiton tuberculatus and Rhyssoplax tulipa (Buckland-Nicks, unpubl. obs.). Mature sperm of Chitonina can be distinguished from those of Lepidopleurida and Acanthochitonina by the arrangement of centrioles, position of mitochondria, and reinforcement of the flagellum. These and other shared apomorphies suggest that Chitonina is a monophyletic taxon (Buckland-Nicks, 2008), which is supported also by molecular studies (Okusu et al., 2003; Lieb et al., 2006).

Egg morphology and fertilization

The egg hulls of Chaetopleura apiculata, as well as those of Stenosemus albus (Buckland-Nicks, 2008), contain a series of permanent micropores around the bases of the spines. Sperm locate these micropores in seconds and thereby gain direct access to the vitelline layer. Permanent pores like these are absent from hulls of Rhyssoplax tulipa, Acanthopleura granulata, and Stenoplax conspicua (Buckland-Nicks, 1995, 2006), even though retraction of projecting microvilli during oogenesis may leave pathways of least resistance for sperm (Buckland-Nicks, 1995, 2006), or sperm may penetrate at junctions of spine bases. Nevertheless, these sperm retain the ability to digest all layers of the hull and vitelline layer (Buckland-Nicks, 2008). Micropores have been found in some species of Acanthochitonina, including Cyanoplax fernaldi (= Lepidochitona fernaldi) and Cyanoplax dentiens (= Lepidochitona dentiens) (Buckland-Nicks and Eernisse, 1992; Buckland-Nicks, 1993). Apparent differences in acrosomal contents suggest that species with open hull pores have simpler acrosomes than those with continuous hulls (Buckland-Nicks et al., 1990; Pash-chenko and Drozdov, 1998; Buckland-Nicks, 2008). When the hull presents a continuous barrier, as was shown in Tonicella lineata, the acrosome may contain more than one granule, and these are used up sequentially during penetration (Buckland-Nicks et al., 1988). It could not be determined whether this applies to all species with continuous hulls lacking pores; to be able to make this distinction, more precise methods of examining sperm acrosomes, such as freeze substitution fixation or labeled antibodies, will have to be employed.

Eggs of Radsia nigrovirescens have long spines ending in prominent hooks. These spines are more similar to those of Stenosemus albus (= Ischnochiton albus), Stenosemus pectinatus (= Ischnochiton pectinatus), and Stenosemus sp. (= Chondropleura sp.) than to spines of other Chitoninae, although this simple form could have arisen by convergence. However, unlike other Chitoninae, Radsia nigrovirescens has an egg hull that is perforated by pores, although we do not know if sperm use these pores to gain access to the vitelline layer during fertilization, as they do in S. albus and Chaetopleura apiculata (Buckland-Nicks, 2008). A molecular analysis of these genera in comparison with known Chitonina would be useful for clarifying their phylogenetic relationships.

In many brooding chitons, spines or cupules are reduced, which is thought to allow for improved packing and storage of larger numbers of eggs in the pallial grooves (Eernisse, 1984, 1988; Buckland-Nicks and Eernisse, 1992). Radsia nigrovirescens is an exception as it has retained long spines. Once these hooked spines come in contact, they simply interlock, which causes eggs to stick together where they are brooded in the pallial grooves (Buckland-Nicks, 2006). In vitro, eggs form clumps whenever they touch, but Eernisse (unpubl. data) suggests that since juveniles of this species form a monolayer on each side of the foot after they emerge, eggs are probably brooded in layers inside the pallial grooves, rather than in clumps. One female was observed releasing individual eggs into a petri dish, which suggests that the animals can also free-spawn. One selective advantage of retaining the long spines is that they dramatically slow sinking rates and keep eggs suspended for longer in the water column, which likely improves chances of fertilization (Buckland-Nicks, 1993, and unpubl. data). Eernisse (1984, 1988) has also suggested that spines may catch on floating debris or form chains or clumps in the water, achieving a similar purpose.

The egg spines of Rhyssoplax tulipa and Chiton tuberculatus have petalloid tips, the shape of which appears, on the basis of diagrams of their eggs, to characterize this genus and several other genera (Sirenko, 1993). These include some species of Acanthopleura and Ischnochiton, as well as Onithochiton, Tonicia, and Stenochiton. Furthermore, spines with multi-branched tips are found in several genera, including Callistochiton, Stenosemus, Subterenochiton, and Lepidozona, that also exhibit simple spines or spines with bifurcations in other species (Sirenko, 1993). Gaymer et al. (2004) showed clearly that eggs of Acanthopleura echinata have spines with petalloid tips, very like those of Rhyssoplax tulipa and Chiton tuberculatus, C. viridis, and C. marmoratus (Sirenko, 1993). Other species of Acanthopleura have been reported to have uniform spines with points (A. japonica, A. brevispinosa, A. gemmata, A. granulata) or recurved tips (A. miles) (Sirenko, 1993, 2006). However, Acanthopleura granulata is shown here to have polymorphic spines, including long and intermediate scaly ones as well as short bifurcating ones, and not the single spine type reported previously for this species (Sirenko, 1993). This considerable variation in spine form in the genus Acanthopleura may underscore problems with classification, as has been true of the genus Ischnochiton. It also may indicate that spine form is in some respects homoplastic. For this reason, precise character definition must be worked out in relation to the mechanism of spine formation during oogenesis in order to confirm homology. At present these results provide some support for previous reports, based on shell characters, gill placement, and gamete morphology, that placed Acanthopleurinae within Chitonidae (Sirenko, 1993; 2006; Buckland-Nicks, 1995; 2006); this placement is in contrast to recent studies that show them to be more closely related to Chaetopleuridae (Okusu et al., 2003; Buckland-Nicks, 2006). The latter cannot yet be ruled out, however, and represents an interesting example of character conflict.

Importance of the fate of sperm organelles in the zygote

In the process of spermiogenesis in Chitonida, after the distal centriole has produced the flagellum, the proximal centriole fuses with the distal centriole to form the basal body. New evidence presented here supports the hypothesis that, at fertilization in all Chitonida, these organelles, as well as the mitochondria, are abandoned on the egg surface. It is usually the case that paternal mitochondria are eliminated and do not play a role in embryonic development (Gilbert, 2006), although there are exceptions among molluscs, such as some mussels (Hoeh et al., 1990); and in most animals, mitochondria degenerate after entering the egg. However, it is unusual for the proximal centriole of the sperm to be eliminated, as it plays a crucial role in assisting the migration of pronuclei in the uncleaved egg, as well as organizing the mitotic spindle during cleavage in most animals (Schatten et al., 1986). In fact, from brown algae (Motomura and Nagasato 2004) and nematodes (Sathananthan et al., 2006) to echinoderms (Sluder et al., 1993), vertebrates, and most mammals (Kalnins, 1992)--including humans (Palermo et al., 1994)--the sperm proximal centriole regulates mitotic spindle formation and, therefore, cleavage of the embryo. Thus, it is very unusual that chitons might have evolved a mechanism that abandons sperm organelles outside the egg, and if true, it raises a number of developmental questions as to how the egg has evolved to compensate.

Mechanism for injection of sperm chromatin

The mechanism of controlling the injection of sperm chromatin into the egg cortex in Chitonida is also interesting. During spermiogenesis, the chromatin is twisted inside the nucleus in characteristic patterns, with several fine strands extending up into the nuclear filament (Buckland-Nicks et al., 1990), being the first to enter the egg. Exactly how the unraveling of chromatin occurs is not known, but it is clear that the chromatin is injected as a fine thread deep into the egg cortex (Buckland-Nicks and Hodgson, 2000; Buckland-Nicks, 2006, 2008). Possibly the energy required for this process is stored in the twisted chromatin during spermiogenesis, when it is bound in place by nuclear proteins.

It is interesting in this regard that Okusu et al. (2003) found that the gene sequence coding for Histone H3 in Callochiton septemvalvis grouped with Chitonida and not with Lepidopleurida, although their overall conclusion was that Callochiton belonged with Lepidopleurida. Other more recent morphological and molecular analyses have confirmed the placement of Callochiton within Chitonida outside Lepidopleurida (Lieb et al., 2006; Buckland-Nicks, 2008).


Thanks are due to Norma Mitchell for technical assistance in preparing the plates in Photoshop and to Haixin Xu for help with re-embedding and sectioning Acanthopleura eggs. Also we thank Alan Hodgson and Doug Eernisse for help collecting and identifying chitons; and Dennis Willows, Director of the Friday Harbor Laboratories in 2004, for supporting research there. This research was supported by an NSERC Undergraduate Student Research Award to E. Brothers and by the Hugh Kelly Fellowship and an NSERC of Canada Discovery grant, both awarded to J. Buckland-Nicks.

Literature Cited

Buckland-Nicks, J. 1993. Hull cupules of chiton eggs: parachute structures and sperm focusing devices? Biol Bull. 184: 269-276.

Buckland-Nicks, J. 1995. Ultrastructure of sperm and sperm-egg interaction in Aculifera: implications for molluscan phylogeny. Pp. 129-153 in Advances in Spermatozoal Phylogeny and Taxonomy. B. G. M. Jamieson, J. Ausio, and J.-L. Justine, eds. Memoires du Museum National d'Histoire Naturelle 166, Paris.

Buckland-Nicks, J. 2006. Fertilization in chitons: morphological clues to phylogeny. Venus 65: 51-69.

Buckland-Nicks, J. 2008. Fertilization biology and the evolution of chitons. Am. Malacol. Bull. (in press).

Buckland-Nicks, J., and D. J. Eernisse. 1992. Ultrastructure of mature sperm and eggs of the brooding hermaphroditic chiton, Lepidochitona fernaldi Eernisse 1986, with special reference to the mechanism of fertilization. J. Exp. Zool. 265: 567-574.

Buckland-Nicks, J., and A. N. Hodgson. 2000. Fertilization in Callochiton castaneus (Mollusca). Biol. Bull. 199: 59-67.

Buckland-Nicks, J., R. Koss, and F. S. Chia. 1988. Fertilization in a chiton: acrosome-mediated sperm-egg fusion. Gamete Res. 21: 199-212.

Buckland-Nicks, J., F. S. Chia, and R. Koss. 1990. Spermiogenesis in Polyplacophora, with special reference to acrosome formation (Mollusca). Zoomorphology 109: 179-188.

Eernisse, D. J. 1984. Lepidochitona Gray, 1821 (Mollusca: Polyplacophora), from the Pacific Coast of the United States: systematics and reproduction. Ph.D. dissertation, University of California, Santa Cruz.

Eernisse, D. J. 1988. Class Polyplacophora. Pp. 49-73 in Taxonomic Atlas of the Benthic Fauna of the Santa Maria Basin and the Western Santa Barbara Channel, Vol. 8. The Mollusca, Part 1: Aplacophora, Polyplacophora, Scaphopoda, Bivalvia and Cephalopoda, P. Valentich Scott and J. A. Blake, eds. Santa Barbara Museum of Natural History, Santa Barbara, CA.

Gaymer, C. F., C. Guisado, K. B. Brokordt, and J. M. Himmelman. 2004. Gonad structure and gamete morphology of the eastern South Pacific chiton Acanthopleura echinata Barnes, 1824. Veliger 47: 141-152.

Gilbert, S. F. 2006. Developmental Biology. 8th ed. Sinauer Associates, Sunderland, MA.

Hodgson, A. N., J. M. Baxter, M. G. Sturrock, and R. T. F. Bernard. 1988. Comparative spermatology of 11 species of Polyplacophora (Mollusca) from the suborders Lepidopleurina, Chitonina and Acanthochitonina. Proc. R. Soc. Lond. 235: 161-177.

Hoeh, W. R., K. H. Blakely, and W. M. Brown. 1991. Heteroplasmy suggests limited biparental inheritance of Mytilus mitochondrial DNA. Science 251: 1488-1490.

Kaas, P., and R. A. van Belle. 2003. Monograph of Living Chitons (Mollusca: Polyplacophora), Vols. 1-5. Brill, Boston.

Kalnins, V. I. 1992. The Centrosome. Academic Press, New York.

Lieb, B., K. Streit, R. P. Kelly, and D. J. Eernisse. 2006. Hemocyanin meets chitons: phylogeny of polyplacophorans revisited by hemocyanin genes. American Malacological Society 72nd Annual Meeting, Seattle, WA, 29 July-3 August 2006. (Abstract).

Motomura, T., and C. Nagasato. 2004. The first spindle formation in brown algal zygotes. Hydrobiologia 512: 171-176.

Okusu, A., E. Schwabe, D. Eernisse, and G. Giribet. 2003. Towards a phylogeny of chitons (Mollusca, Polyplacophora) based on combined analysis of five molecular loci. Org. Divers. Evol. 3: 281-302.

Palermo, G., S. Munne, and J. Cohen. 1994. The human zygote inherits its mitotic potential from the male gamete. Hum, Reprod. 9: 1220-1225.

Pashchenko, S. V., and A. L. Drozdov. 1998. Morphology of gametes in five species of Far-Eastern chitons. Invertebr. Reprod. Dev. 33: 47-56.

Sathananthan, A. H., W. D. Ratnasooriya, A. de Silva, and P. Randeniya. 2006. Rediscovering Boveri's centrosome in Ascaris (1888): its impact on human fertility and development. Reprod Biomed. Online 12: 254-270.

Schatten, H., G. Schatten, D. Mazia, R. Balczon, and C. Simerly. 1986. Behavior of centrosomes during fertilization and cell division in mouse oocytes and in sea urchin eggs. Proc. Natl. Acad. Sci. USA 83: 105-109.

Sirenko, B. I. 1993. Revision of the system of the order Chitonida (Mollusca: Polyplacophora) on the basis of correlation between the type of gills arrangement and the shape of the chorion processes. Ruthenica 3: 93-117.

Sirenko, B. I. 2006. New outlook on the system of chitons (Mollusca: Polyplacophora). Venus 65: 27-49.

Sluder, G., F. J. Miller, and K. Lewis. 1993. Centrosome inheritance in starfish zygotes. II: Selective suppression of the maternal centrosome during meiosis. Dev. Biol. 155: 58-67.

van Belle, R. A. 1983. The systematic classification of the chitons (Mollusca: Polyplacophora). Inform. Soc. Belge Malacol. 11: 1-179.


(1) St. Francis Xavier University, Antigonish, Nova Scotia B2G 2W5, Canada; and (2) Biology Department, University of Victoria, Victoria, British Columbia V0T 1S0, Canada

Received 11 July 2007; accepted 31 October 2007.

* To whom correspondence should be addressed. E-mail:
COPYRIGHT 2008 University of Chicago Press
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2008 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Buckland-Nicks, John; Brothers, Elizabeth
Publication:The Biological Bulletin
Article Type:Report
Geographic Code:1CANA
Date:Apr 1, 2008
Previous Article:Phylogenetic analysis of caprellid and corophioid amphipods (Crustacea) based on the 18S rRNA gene, with special emphasis on the phylogenetic...
Next Article:Patterns of male reproductive success in Crepidula fornicata provide new insight for sex allocation and optimal sex change.

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