Ultrastructure of hull formation during oogenesis in Rhyssoplax tulipa ...

Invertebrate Reproduction and Development, 53:3 (2009) 165–174 Balaban, Philadelphia/Rehovot 0168-8170/09/$05.00 © 2009 Balaban

165

Ultrastructure of hull formation during oogenesis in Rhyssoplax tulipa (=Chiton tulipa) (Chitonidae: Chitoninae) JOHN BUCKLAND-NICKS1* and ARKADIY REUNOV2 St Francis Xavier University, Antigonish, Nova Scotia, B2G 2W5, Canada email: [email protected] 2 A.V. Zhirmunsky Institute of Marine Biology, Far East Branch of Russian Academy of Sciences, Palchevsky St., 17, 690041 Vladivostok, Russia Tel. +7 (4232) 311143; Fax+7 (4232)310900 1

Received 5 August 2009; Accepted 4 November 2009

Abstract Egg hull formation is described in detail for the first time in Rhyssoplax tulipa from South Africa. Although previous descriptions of hull formation in chitons provide some basis for comparison, the mechanisms we have discovered are totally different. The process begins with the release of clear vacuoles from the oocyte into the intercellular space, which separates it from a layer of follicle cells. Materials contributing to the layers of the hull are formed primarily by micro-apocrine secretion processes, in which extensions of the follicle cells and oocyte are budded off into the intercellular space, rather than by formation and release of vesicles from the Golgi bodies, as previously thought. Later in oogenesis, these secretions become organized into three functional layers, each with a characteristic substructure. Follicle cells mold the shape of the hull spines even down to the petalloid tips that are characteristic of this group. Late in oogenesis the oocyte secretes the vitelline layer beneath the hull and the follicle cells break down and are sloughed off. Differences in the mechanisms of hull formation among chitons are providing a new set of morphological characters that may yield insight into chiton phylogeny. To our knowledge, this is the first description of micro-apocrine secretions during oogenesis in a mollusc, although it has been reported in other phyla. Key words:

Polyplacophora, follicle cells, egg envelopes, micro-apocrine secretion, electron microscopy, chiton phylogeny

Introduction The eggs of chitons are enclosed by an outer hull, which is jelly-like in basal groups, such as Lepidopleurida (Hodgson et al., 1988a; Pashchenko & Drozdov, 1998; Buckland-Nicks & Hodgson, 2000) but is elaborated into complex spines and cupules in most species of the more recent order Chitonida (see reviews

*

Corresponding author.

by Pearse, 1979; Gaymer, et al. 2004; Buckland-Nicks, 2006, 2008). These elaborate spines and cupules are an integral part of a unique mechanism of sperm penetration (Buckland-Nicks et al., 1988) that characterizes all Chitonida (Buckland-Nicks, 2006). Richter (1986) pointed out that the elaborate egg spines are produced from contributions of both oocyte and follicle cells,

166

J. Buckland-Nicks, A. Reunov / IRD 53 (2009) 165–174

which we confirm here, and therefore one should not use the term ‘chorion’ (which is a layer secreted solely by the egg) to properly refer to this layer. Rather, the term ‘hull’ is more appropriate and has replaced the term ‘chorion’ in chiton research for a number of years (Pearse, 1979). Hull spine morphology can provide useful characters for phylogenetic analysis (see: Eernisse, 1984; Sirenko, 1993, 2006; Buckland-Nicks, 1995, 2006, 2008) and moreover, the details of hull formation may provide new insights into these relationships. However, oogenesis in chitons with information on the ultrastructure of hull formation and the relationship between oocyte and follicle cells has been described in only a few species (Anderson, 1969; Selwood, 1968, 1970; Richter, 1976, 1986; Durfort et al., 1982). In the present report egg hull formation is described in detail for the first time in Rhyssoplax tulipa Quoy and Gaimard, 1835. The hull of this species is elaborated into long spines with petalloid tips, quite similar in form to that described for Sypharochiton septentriones (Selwood, 1970) but there are some important differences. The results of this study provide new insight into the mechanism of hull deposition and suggest also that comparing mechanisms of hull formation among different taxa may help to further clarify phylogenetic relationships. Materials and Methods Specimens of Rhyssoplax tulipa Quoy and Gaimard, 1835 (= Chiton tulipa Quoy and Gaimard, 1835) were collected from East London, South Africa (32E54N S, 28E04N E), in September 1999. Chitons were placed singly in small petri dishes containing 0.45 µm Millipore filtered seawater (MFSW). Males often spawned first, releasing a milky white suspension of sperm. When this occurred some sperm were added to the other dishes, which usually stimulated females to spawn. Females were dissected and small pieces of ovary were removed with fine forceps and placed in primary fixative together with some spawned eggs. The primary fixative contained 50 ml of 0.2M sodium cacodylate buffer (adjusted to pH 7.4) with 40 ml 0.45 µm MFSW containing 0.1M sucrose and 10 ml of 25% glutaraldehyde (giving final concentrations of 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer with seawater and 0.1M sucrose at pH 7.4). Fixation was on ice for 1 h, then allowed to come to room temperature (about 20EC) and continued overnight. The next day following two rinses in 0.1M sodium cacodylate in seawater and 0.1M sucrose, secondary fixation was performed with 1.5% osmium tetroxide in the same buffer for one hour at 20EC,

followed by dehydration. 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 microscope. The vials were capped and underwent critical point drying in a Samdri PVT-3B critical point dryer, Tousimis. Each vial was then uncapped and inverted onto an SEM stub covered with a double carbon sticky tab, such that the eggs adhered to the stub. Finally, the eggs were sputter-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 TAAB 812/Araldite. Polymerization in embedding molds was completed in a 60EC oven for up to 2 days. An RMC MT2C ultramicrotome was used to cut 1 µm sections for light microscopy and silver–gold thin sections for TEM using a diamond knife (Diatome). Thick sections were stained for 15 s with 1% toluidine blue adjusted to pH 9 with sodium bicarbonate. Thin sections were picked up on formvar-coated copper slot grids and stained sequentially with aqueous uranyl acetate (10 min) and lead citrate (4 min) with extensive washing with degassed distilled water between stains and after staining. Stained sections were examined and photographed with a Philips TEM 410 operated at 80 kV. Results Hull formation is initiated following the development of numerous ‘areolae’ or clear vacuoles in the oocyte cytoplasm (Fig. 1A), which move to the surface and are exocytosed creating a series of intercellular spaces between the oocyte and follicle cells (Fig. 1B). In more developed oocytes these spaces appear as compartments, comprising invaginations of both the oocyte and follicle cell membranes, and isolated from each other by oocyte and follicle cell projections which meet at junctions in the intercellular space (Fig. 1B,C). In addition, both oocyte and follicle cells extend microvilli into each compartment (Fig. 1C–F). Microvilli extending from follicle cells contain cisternae of rough endoplasmic reticulum, which is highly developed in the follicle cells (Fig. 1D–F). These microvilli elongate and begin rolling up at their tips (Fig. 1E–G), eventually forming membranous aggregates (Fig. 1H), which appear to be released into the intercellular compartments (Fig. 1I). Similar membranous aggregates, presumably detached from the follicle cells, could be seen freely floating in the


167

Fig. 1. Electron micrographs of oogenesis in Rhyssoplax tulipa. A. Portion of a developing oocyte (O), showing numerous ‘areolae’ (A) vacuoles in the cytoplasm and at the surface. The oocyte is covered by a continuous layer of follicle cells (FC). Scale bar = 5 µm. B. Portion of oocyte showing accumulation of ‘areolae’ (asterisks) between follicle cells (FC) and oocyte (O), creating intercellular spaces bridged by cell projections extending from the follicle cells and oocyte which form junctions where they meet (arrowheads). Scale bar = 1 µm. C. Portion of oocyte (O) showing oocyte microvillus (OM). Note junction between projections of oocyte and follicle cell in right part of picture. Scale bar = 0.5 µm. D. Follicle cell microvillus (FCM), containing cisterna of rough endoplasmic reticulum (ER), and extending into the intercellular space. Scale bar = 0.3 µm. E. Follicle cell microvillus (FCM) beginning to roll up at its tip. Oocyte (O), Follicle cell (FC). Scale bar = 0.3 µm. F. Two follicle cell microvilli (FCM) which are rolling up at the tips. Rough endoplasmic reticulum (ER) is abundant in the cytoplasm and can be seen extending into the follicle cell microvilli. Scale bar = 0.3 µm. G. Rolled up follicle cell microvillus (arrow) can be seen in the intercellular space. Follicle cell (FC), oocyte (O). Scale bar = 1 µm. H. Follicle cell microvillus is rolled into a membranous aggregate (MA) just prior to release. Follicle cells (FC), oocyte (O). Scale bar = 0.3 µm. I. Membranous aggregate (MA) breaking away as a secretion in the intercellular space. Follicle cell (FC), oocyte (O). Scale bar = 0.3 µm.

168


Fig. 2. Electron micrographs of oogenesis in Rhyssoplax tulipa. A. Detached membranous aggregate (MA) freely floating in the intercellular space. Follicle cell (FC), oocyte (O). Scale bar = 0.3 µm. B. Intercellular space containing scattered electron opaque substances (asterisks) resulting from the dissemination of follicle cell membranous aggregates (MA). These components contribute mostly to the inner layer of egg spines. Scale bar = 0.3 µm. C. Micro-apocrine secretion vesicle (V) about to be released from the tip of an oocyte microvillus. Scale bar = 0.3 µm. D. Micro-apocrine secretion vesicles (V) of the oocyte are seen freely floating in the intercellular space. Similar vesicle formed spheres (VFS) are seen settling on the follicle cell (FC). Scale bar = 0.3 µm. E–I. Sequence of events following the attachment of micro-apocrine secretion vesicle (V) to the follicle cell membrane and rolling up to form a sphere. Scale bar = 0.1 µm.

intercellular space (Fig. 2A) where they break down, forming a fibrous secretion, together with fragments of membrane that gradually accumulate there (Fig. 2B). Microvilli extending from the oocyte membrane were observed to pinch off vesicles from their tips (Fig. 2C) in a microapocrine secretion process, releasing them into the same space (Fig. 2D). Each membranebound vesicle first attaches to the follicle cell membrane, where it rolls up and forms a small sphere

(Fig. 2E–I). Eventually the outer membrane of the sphere breaks down (Fig. 3A) leaving a layer of closely apposed dense secretions adjacent to the follicle cell membrane, which constitute the “Outer Dense Layer” (Fig. 3B–E). Now, some of the secretions in the intercellular space, which derived from the follicle cells, begin to attach to individual dense spheres of the outer dense layer (Fig. 3A). Once attached, the secretions change


169

Fig. 3. Electron micrographs of oogenesis in Rhyssoplax tulipa. A. The dense spheres (arrowheads) with initial deposit of intermediate layer secretion (ILS) which is presumed to arise by polymerization of scattered substance (asterisk) previously generated by dissemination of membranous aggregates. Follicle cell (FC). Scale bar = 0.3 µm. B. Intermediate layer substance (ILS), now fibrous in structure, is attached to two individual dense spheres (arrowheads). Follicle cell (FC). Scale bar = 0.3 µm. C. Intermediate layer substance (ILS), spreads along adjacent dense spheres (arrowheads) on the follicle cell (FC). Scale bar = 0.3 µm. D. Intermediate Layer substance (ILS) cross connects several adjacent dense spheres (arrowheads) gradually creating a continuous Intermediate Layer. Follicle cell (FC). Scale bar = 0.5 µm. E. Intermediate Layer (IML) is completed between Outer Layer (OL) on the follicle cell membrane (FC) and Inner Layer (IL). Scale bar = 0.5 µm. F. Spines are developing as individual processes, controlled by a single follicle cell (FC). Oocyte microvilli (arrows) project into the intercellular space which forms the inner layer of the spine. Note: follicle cell nucleus (FCN) near apex of spine. Yolk granules (YG), oocyte (O). Scale bar = 3 µm. G. Section of follicle cell and oocyte showing new position of follicle cell nucleus (FCN) at base of spine. Note oocyte microvilli projecting into spine interior (arrow). Scale bar = 3 µm.

from being fibrous amorphous to forming a fibrous network of intricate substructure (Fig. 3B). As this fibrous network increases in size, it spreads outwards and contacts similar networks on adjacent dense spheres (Fig. 3C,D) gradually forming a continuous “Intermediate layer” comprising intricately folded fibrous

elements (Fig. 3E). By the beginning of spine formation all three layers of the hull have been deposited. These are an “Inner Layer” of loose fibers, an “Intermediate Layer” of intricately folded fibers and an “Outer Dense Layer” comprised of electron-dense spheres (Fig. 3E). This phase is also marked by changes in the confor-

170


Fig. 4. Electron micrographs of oogenesis in Rhyssoplax tulipa. A. Almost completed spine with broad tip and follicle cell nucleus (FCN) at base. Note oocyte microvilli projecting into spine interior (arrow). Follicle cell (FC), oocyte (O). Scale bar = 3 µm. B. Cross section below spine tip. Follicle cell (FC). Scale bar = 3 µm. C. Cross section through tip of spine showing development of terminal projections (TP), which form the characteristic petalloid spine tips. Scale bar = 3 µm. D. Scanning electron micrograph of a portion of fully formed ripe egg. Scale bar = 50 µm. E. Oocyte cone (OC) elaborated with microvilli (OM) and projecting into base of spine. Scale bar = 1 µm. F. Secretory stage of oocyte microvilli (OM) with tips swollen with granular secretions. Arrows show detached microvillus. Oocyte cone (OC). Scale bar = 1 µm. G. Section at base of spine showing formation of Outer and Intermediate Layers (arrowheads) between spines. Note follicle cell junction (FCJ) connecting adjacent follicle cells (stars) to each other. Oocyte (O). Scale bar = 3 µm. H. Cross section of follicle cell (FC), showing lobulated nucleus (FCN) Scale bar = 1 µm. I. Section of oocyte and basal part of spine showing absence of follicle cells. Note the vitelline layer (asterisk) newly formed between hull (H) and oocyte (O). Scale bar = 3 µm. J. Cross section through hull spine lacking follicle cell covering and consisting of the Outer Layer (OL), Intermediate Layer (IML), and Inner Layer (IL). Scale bar = 1 µm.


mation of the follicle cells as they begin to mold the shape of the spines beneath them. The follicle cells can be seen to raise up into cell buds (Fig. 3F), incorporating the intercellular spaces now filled with fibrous secretions. Each follicle cell bud is a separate entity, rising up from the oocyte surface (Fig. 3F). Long oocyte microvilli can be seen projecting into the base of each cell bud (Fig. 3F,G) where secretions are added to the Inner Layer of the spines. The next stage in spine formation is marked by the movement of the nucleus from the apex of each bud (Fig. 3F) to its base (Fig. 3G). The follicle cells continue to extend upwards forming the long spines beneath them until the maximum height is reached, then each one begins to widen at its tip (Fig. 4A) and develops a wavy contour (Fig. 4B) that corresponds with the development of six terminal projections (Fig. 4C). These terminal projections become organized to form the “petalloid” spine tip, characteristic of this group (Fig. 4D). During spine formation the oocyte membrane projects into the base of the spine as a pointed cone elaborated into microvilli (Fig. 4E). Later in this process the cones flatten and the microvilli develop bulbous tips containing granular secretions (Fig. 4F). It was frequently seen that microvilli detached from the base and became incorporated in the spine interior (Fig. 4F). Eventually the oocyte microvilli disappear altogether. The later stages of spine formation involve the completion of the outer layer of dense spheres and its accompanying Intermediate Layer between the adjacent follicle cells (Fig. 4G), thus forming a continuous layer over the egg. Follicle cell nuclei are normal in appearance for most of the time that spines are forming but later in this process they develop an irregular outline (Fig. 4H) and the nuclei begin to degenerate. In the last stages of oogenesis, the vitelline layer is secreted between the hull and oocyte membrane (Fig. 4I) and, at about the time the egg is released, the follicle cells are discarded which fully reveals the petalloid-tipped spines (Fig. 4D), completing Outer, Intermediate and Inner Layers (Fig. 4J). Discussion The three layered construction of the hull and the development of spines in Rhyssoplax tulipa is very similar to that described for Sypharochiton septentriones by Selwood (1970). Furthermore, both species have well developed long spines with distinctive broad petalloid tips indicating that R. tulipa and S. septentriones may be quite closely related phylogenetically. The intercellular spaces arising early in oogenesis between follicle cells and oocyte are a result of large clear vacuoles (Richter, 1986) or ‘areolae’ produced

171

inside the oocyte and released into this extracellular space by exocytosis. These secretions are subsequently compartmentalized by the extension of microvillous projections from both follicle cells and oocyte that meet at junctions in the middle (Anderson, 1969; Richter, 1986). These intercellular compartments create the correct spacing and basic pattern of spine distribution on the egg surface, which acts as a template for spine formation by subsequent contributions of follicle cells and oocyte. Mechanism of hull formation Our results on the mechanism of hull formation differ substantially from those previously reported (Anderson, 1969; Selwood, 1970; Richter, 1976, 1986). Selwood, (1970) suggested for S. septentriones that the outer and intermediate hull layers were formed from material derived from Golgi bodies of the follicle cells. In our study we did not find that the Golgi bodies were the main contributor to either of these layers, although they may contribute some material to the inner layer of the hull. Rather, our observations reveal that the majority of the hull is formed by novel mechanisms involving micro-apocrine secretions from both follicle cells and oocyte that have not been fully understood, previously. Specifically there are three key differences in our understanding of hull formation, which relate to individual hull layers. 1. The Outer Dense Layer. This first layer is formed from the dense spheres observed in R. tulipa, which are very similar in construction to those described for S. septentriones. Selwood (1970) suggested that “these dense spheres formed from the accumulation of substances secreted by the Golgi bodies and exocytosed onto the surface”. This statement was based partly on the similarity between the material comprising the dense spheres and the material being released by the Golgi bodies, although micrographs illustrating these points were not provided. We did not observe release of numerous Golgi secretions in this manner, although occasionally active Golgi bodies were seen. According to our findings in R. tulipa the outer layer of dense spheres is the result of a micro-apocrine secretion formed by the oocyte and released from the tips of oocyte microvilli. These vesicles float in the intercellular space and later become attached to the follicle cell membrane, where they roll up and lose their outer membrane, gradually forming the layer of dense spheres. This is the first description of a micro-apocrine secretion in molluscan oogenesis, although a macroapocrine process is responsible for delivering secretions that digest the sperm extracellularly in the bursa copulatrix of some gastropod snails (Buckland-Nicks,

172


1974). Quite similar micro-apocrine secretion processes are well known in the insect literature (Cristofoletti et al., 2001; Bolognesi et al., 2001) and have also been discovered in mice where Hermo and Jacks (2002) remarked on Nature’s “ingenuity in bypassing the classical secretory role via apocrine secretion”. The apocrine mechanism of secretion was described also in embryogenesis of the sea urchin Heliocidaris erythrogramma and sea star Cryptasterina hystera, where lipids are secreted into the blastocoel taking a fragment of cell membrane (Henry et al., 1991; Byrne, 2005). The Outer Dense Layer itself presents a barrier to fertilizing sperm in chiton species like Stenoplax conspicua and R. tulipa (= Chiton tulipa), which upon contact with it, undergo the acrosome reaction and digest it (Buckland-Nicks, 2006, 2008). This process leaves a large circular area around the tip of the penetrating sperm, from where the dense spheres have been eliminated (Buckland-Nicks, 2008). Other genera of Chitonina, such as Stenosemus, Chaetopleura and Dinoplax, lack the outer dense layer. Preliminary studies of Dinoplax suggest that the remaining layers of the hull are formed in other novel ways (Buckland-Nicks and Reunov, unpublished). Such differences in hull formation may, in the future, provide new apomorphic characters for re-analysing phylogenetic relationships of Polyplacophora. 2. Intermediate Layer: Selwood (1970) thought that both outer and intermediate layers were formed from Golgi secretions of the follicle cells when she said: “Secretion of these granules from the follicle cells into the intercellular space was observed, the outer lipid and middle protein layers of the chorion apparently are elaborated from them, possibly by an hydration process”. Raven (1966) and Anderson (1969) studying other chiton species, both noticed that follicle cells actively secrete directly to the egg membranes. Our observations suggest that the follicle cells produce and release a second type of micro-apocrine secretion, by extending long processes that contain cisternae of rough ER, which roll up at the tip and appear to break off into the intercellular space. These micro-apocrine secretions accumulate in the intercellular space to form a loose matrix of amorphous fibres and broken membranes. This may be what Anderson (1969) was referring to when he wrote: “cisternae of the abundant rough endoplasmic reticulum are filled with a dense substance. These extremely dilated cisternae become associated with the plasmalemma, fuse and release their contents onto the basal surface of the follicle cells, thereby forming the secondary coat”. The Outer Dense Layer appears to play a key role in the formation of this Intermediate Layer by inducing attachment and polymerization of the amorphous fibres from the inter-

cellular space. Initially amorphous fibres deposit on individual dense spheres, then develop into a network of complex fibrous elements that gradually spread out to contact and integrate with similar networks of fibres on adjacent dense spheres. Eventually, the Intermediate Layer develops as a continuous layer of intricately folded fibres overlying the dense spheres. We did not observe extensive involvement of the Golgi bodies in the production of this Intermediate Layer, as was observed by Selwood (1970) for S. septentriones. The secretion product that comprises the Intermediate Layer seems to be solely derived from cisternae of the RER of the follicle cells, which is very extensive at this time, and is therefore, most likely, proteinaceous. This agrees with some observations of Richter (1986) as well as the histochemical results of Selwood (1970), who found that this Intermediate Layer was composed of protein whereas the Outer Dense Layer was made up of lipoprotein and phospholipid. 3. Inner Layer. The Inner Layer of the hull and spines is produced partly from contributions of the follicle cells, as well as secretions from the oocyte microvilli, which become swollen with cytoplasmic secretions towards the end of oogenesis. These secretions are once again microapocrine in nature, as the tips of the microvilli can be seen breaking off into the interior compartment of each spine, without following the normal pathway through the Golgi bodies (Hermo & Sacks, 2002). Richter (1986) also noted contributions to this layer by the oocyte microvilli in Lepidochitona but agreed with Selwood (1970) that this inner layer was formed primarily by Golgi body secretions developed in the oocyte. We agree that Golgi vesicles may contribute to the Inner Layer but feel that the majority of it is derived from microapocrine secretions of the oocyte and follicle cells. In conclusion, the mechanisms of hull formation in R. tulipa are quite different than has been realized in previous publications on chitonid oogenesis (Selwood, 1968, 1970; Anderson, 1969; Richter, 1976, 1986). Although the more traditional mechanism of exocytosis of secretory vesicles derived from the Golgi bodies may apply to some extent, the bulk of secretion release is through two different micro-apocrine secretory processes, previously undescribed, one for the follicle cells and the other for the oocyte. These ultrastructural mechanisms of hull formation appear to be specific for different taxa (Buckland-Nicks and Reunov, unpublished) suggesting that it may be possible to establish a new character set for phylogenetic analysis, based on hull formation, which when added to existing data for general morphology (Van Belle, 1983; Kaas & Van Belle, 1994; Kaas et al., 2006; Sirenko, 1993, 1997, 2006; Brooker et al., 2006), sperm structure and


fertilization (Hodgson et al., 1988b; Buckland-Nicks, 1995, 2006, 2008; Buckland-Nicks & Hodgson, 2000) and molecular data (Okusu et al., 2003; Lieb et al., 2006) may further clarify evolutionary relationships among chitons.

Acknowledgements We are grateful to Professor Alan Hodgson for hosting both authors in different years, at Rhodes University, South Africa. This work has been supported by an NSERC Discovery grant and the Hugh Kelly Fellowship (Rhodes University) awarded to John Buckland-Nicks, and by the James Chair Professorship awarded to Arkadiy Reunov, while visiting St Francis Xavier University, Canada.

References Anderson, E. 1969. Oocyte-follicle cell differentiation in two species of Amphineurans (Mollusca), Mopalia muscosa and Chaetopleura apiculata. Journal of Morphology, 129, 89–126. Bolognesi, R., Ribeiro, A.F., Terra, W.F. & Ferreira, C. 2001. The peritrophic membrane of Spodoptera frugiperda: Secretion of peritrophins and role in immobilization and recycling digestive enzymes. Archives of Insect Biochemistry and Physiology, 47(2), 62–75. Brooker, L.R., Lee, A.P., Macey, D.J., Webb, J. & van Bronswijk, W. 2006. In situ studies of biomineral deposition in the radula teeth of chitons of the suborder Chitonina. Venus, 65(1–2), 71–80. Buckland-Nicks, J. 1974. The Functional Morphology of the Reproductive Systems and the Larval Development of Two Intertidal Snails, Littorina sitkana (Philippi, 1845) and Littorina scutulata (Gould, 1849) with Special Reference to the Spermatozoa. Ph.D. Thesis, University of Alberta, pp. 258. Buckland-Nicks, J. 1995. Ultrastructure of sperm and spermegg interaction in Aculifera: implications for molluscan phylogeny. In: Advances in Spermatozoal Phylogeny and Taxonomy, B.G.M. Jamieson, J. Ausio & J.-L. Justine (eds), Mémoires du Muséum National d’Histoire Naturelle, 166, Paris, pp.129–153. 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. American Malacological Bulletin, 25, 97–111. Buckland-Nicks, J. & Hodgson, A.N. 2000. Fertilization in Callochiton castaneus (Mollusca). Biological Bulletin, 199, 59–67. Buckland-Nicks, J., Koss, R. & Chia, F.-S. 1988. Fertilization in a chiton: acrosome-mediated sperm-egg fusion, Gamete Research, 21, 199–212. Byrne, M. 2005. Viviparity in the sea star Cryptasterina

173

hystera (Asterinidae)-conserved and modified features in reproduction and development. Biological Bulletin, 208, 81–91. Cristofoletti, P.T., Ribeiro, A.F. & Terra, W.R. 2001. Apocrine secretion of amylase and exocytosis of trypsin along the midgut of Tenebrio molitor larvae. Journal of Insect Physiology, 47(2), 143–155. Durfort, M., Bargalló, B., Bozzo, M.G., Fontarnau, R. & López-Campos, J. 1982. Relationship between follicular cells and oocytes of Trachydermon cinereus Thiele (Mollusca, Polyplacophora). Malacologia, 22, 211–217. Eckelbarger, K.J. 2005. Oogenesis and oocytes. Hydrobiologia, 535/536, 179–198. Eernisse, D.G. 1988. Reproductive patterns in six species of Lepidochitona (Mollusca: Polyplacophora) from the Pacific Coast of North America. Biological Bulletin, 174, 287–302. Gaymer, C.F., Guisado, C., Brokordt, K.B. & Himmelman, J.H. 2004. Gonad structure and gamete morphology of the eastern South Pacific chiton Acanthopleura echinata Barnes, 1824. Veliger, 47(2), 141–152. Henry, J.J., Wray, G.A., Raff, R.R. 1991. Mechanism of an alternate type of echinoderm blastula formation: the wrinkled blastula of the sea urchin Heliocidaris erythrogramma. Development, Growth and Differentiation, 33, 317–328. Hermo, L. and Sacks, D. 2002. Nature’s ingenuity: Bypassing the classical secretory route via apocrine secretion. Molecular Reproduction and Development, 63(3), 394–410. Hodgson, A.N., Baxter, J.M., Sturrock, M.G. & Bernard, R.T.F. 1988a. The structure of the chorion of the eggs of some Polyplacophora: correlations with sperm morphology and implications for fertilization. European Marine Biology Symposium. Pp.14–15. Hodgson, A.N., Baxter, J.M., Sturrock, M.G. & Bernard, R.T.F. 1988b. Comparative spermatology of 11 species of Polyplacophora (Mollusca) from the suborders Lepidopleurina, Chitonina and Acanthochitonina. Proceedings of the Royal Society of London B, 235, 161– 177. Kaas, P. & Van Belle, R.A. 1994. Monograph of Living Chitons. 4. Suborder Ischnochitonina: Ischnochitonidae: Ischnochitonina (continued). Addition to Vols. 1, 2 and 3. E.J. Brill & W. Backhuys, Leiden. Kaas, P., Van Belle, R.A. & Strack, H. 2006. Monograph of Living Chitons (Mollusca: Polyplacophora). Vols. 1–5. Brill Publishers, Leiden. Lieb, B., Streit, K., Kelly, R.P. & Eernisse, D. 2006. Hemocyanin meets chitons: Phylogeny of polyplacophorans revisited by hemocyanin genes. Abstracts of the 72nd Annual American Malacological Society Meeting, Seattle, Washington, p. 61. Okusu, A., Schwabe, E., Eernisse, D.J. & Giribet, G. 2003. Towards a phylogeny of chitons (Mollusca, Polyplacophora) based on combined analysis of five molecular loci. Organisms, Diversity and Evolution, 3, 281–302. Pashchenko, S.V. & Drozdov, A.L. 1998. Morphology of gametes in five species of Far-Eastern chitons. Invertebrate Reproduction and Development, 33, 47–56.

174


Pearse, J.S. 1979. Polyplacophora. In: Reproduction of Marine Invertebrates, Vol. V. A.C. Giese & J.S. Pearse (eds.), Academic Press, New York, pp. 27–85. Raven, C.P. 1966. Morphogenesis: The Analysis of Molluscan Development. 2nd ed., Pergamon Press, Oxford. Richter, H.-P. 1976. Feinstrukturelle Untersuchingen zur Oogenese der Käferschnecke Lepidochitona cinereus (Mollusca, Polyplacophora). Helgoländer Meeresuntersuchungen, 28, 250–303. Richter, H.-P. 1986. Ultrastructure of follicular epithelia in the ovary of Lepidochitona cinerea L. (Mollusca: Polyplacophora). Development Growth and Differentiation, 28, 7–16. Selwood, L. 1968. Interrelationships between developing oocytes and ovarian tissues in the chiton Sypharochiton septentriones (Ashby) (Mollusca, Polyplacophora). Journal of Morphology, 125, 71–104.

Selwood, L. 1970. The role of follicle cells during oogenesis in the chiton Sypharochiton septentriones (Ashby) (Polyplacophora, Mollusca). Zeitschrift für Zellforschung, 104, 178–192. 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, 93–119. Sirenko, B.I. 1997. The importance of the development of articulamentum for taxonomy of chitons (Mollusca, Polyplacophora). Ruthenica, 7, 1–24. Sirenko, B.I. 2006. New outlook on the system of chitons (Mollusca: Polyplacophora). Venus, 65 (1–2), 27–49. Van Belle, R.A. 1983. The systematic classification of the chitons (Mollusca: Polyplacophora). Addenda 1 (with the description of the genus Incissiochiton gen. n.). Informations de la Société Belge de Malacologie, 11, 1–178.