Ca²⁺ in Specification of Vegetal Cell Fate in Early sea Urchin Embryos

It is well-known that during the early development of sea urchin embryos, the fate of a particular cell depends on its position in the embryo, rather than on the cytoplasm it has acquired from the egg. It was shown (Driesch, 1900) that, if isolated, every blastomere up to the 4-cell stage was able to form a small but complete pluteus larva (Fig. 1I, J). The third division occurs at right angles to the first and second cleavage planes (Fig. 1A–C). If an embryo is bisected along the first or second cleavage plane, each half forms a small but complete larva. If the embryo is bisected along the third cleavage plane, however, the result is different; one half forms a dwarf larva but the other develops into a hollow, ciliated blastula-like embryo (Fig. 1E) (Horstadius, 1939). This means that the four blastomeres at the 4-cell stage are equivalent and regulative, but by the next division the blastomeres are segregated into cells with different potentialities. This differential ability has been recognized in unfertilized eggs. There is no visible marker to indicate the polarity of sea urchin eggs but such a marker is present in the extracellular structure surrounding the eggs, the jelly layer. The position of the jelly canal in this transparent jelly coat marks the future animal pole of the embryo (Boveri, 1901). When the jelly canal was visualized using carbon particles and the unfertilized egg bisected into animal and vegetal halves before insemination, it was found that the animal half developed into a blastula-like larva and the vegetal half became an almost normal larva (Maruyama et al., 1985).

At the fourth division, the cells occupying the animal half of the embryo divide equally, while in the vegetal half unequal divisions occur, so that three different sizes and qualities of blastomere are formed: the mesomeres, the macromeres and the micromeres (Fig. 1D). The tiers of blastomeres located along the animal–vegetal (A–V) axis are fated to give rise to the ectoderm, the endoderm and the mesoderm, and are specified and patterned by the coordinated action of localized, maternally derived determinants and cell–cell signals (Horstadius, 1939). The A–V axis is further patterned to produce five distinct territories by the 60-cell stage that are specified by a signal transduction cascade initiatiated by the micromeres (Davidson, 1989).

Signal molecules or determinant molecules involved in the specification of sea urchin embryonic cells have recently been found. Transcripts of bone morphogenic protein (BMP) gene of sea urchin, which first appear in the oral ectoderm region at about the 200-cell stage, suppress the commitment of cells to vegetal fates (Angerer et al., 2000). The oral ectoderm occupies an epithelial region surrounded by four arms of a pluteus including a mouth, and is distinct from the aboral ectoderm, the dorsal and the ventral epithelium (Fig. 1I,J). Specification of the fate of secondary mesenchyme cells (SMC in Fig. 1G) in the endoderm involves the Notch signaling pathway in which the Notch receptor, a membrane protein, is expressed from the 120-cell stage and decreases the number of endodermal cells formed by causing the production of SMC (Sherwood and McClay, 1999). β-catenin, which is involved in the Wnt pathway, initially increases in the micromeres at the 16-cell stage and then accumulates in the vegetal nuclei to specify the endoderm (Logan et al., 1999). The β-catenin is regulated by a ternary complex factor (TCF), a binding partner of nuclear β-catenin, until the 60-cell stage (Vonica et al., 2000). While these facts are very interesting it is still not known how the signals are activated, or what physiological conditions in cells regulate these signal pathways. In this paper I discuss morphological and physiological cell functions that may be involve in the autonomous specification of the micromeres.

The first specification of blastomeres occurs autonomously in the micromeres at the 16-cell stage, and the first signals in embryos are sent out from these micromeres to the neighbouring cells, the macromeres, to induce the archenteron (Fig. 1G). The micromeres self-differentiate to the primary mesenchyme cells (PMC, Fig. 1F), and form the larval spicules (Fig. 1G) and skeletal rods (Fig. 1H–J). If the micromeres are removed from 16-cell stage embryos and cultured in isolation, they form mesenchymal aggregates, which subsequently differentiate into skeletogenic cells, as in normal embryos (Okazaki, 1975). The inductive ability of the micromeres has been demonstrated by transplantation experiments (Horstadius, 1939; Ransick and Davidson, 1993) showing that if extra micromeres were transplanted onto the mesomeres, a second archenteron invaginated from the animal pole, in addition to the original archenteron, which invaginates from the vegetal pole of the host embryo (Fig. 2). Experiments in which micromeres were removed at various stages suggested that contact with micromeres during the 16-to 60-cell stage was necessary for normal induction of the archenteron (Ransick and Davidson, 1995).

Electrical coupling is initially established between the micromeres and macromeres at the 16-cell stage. The maximum junctional conductance of 26 nS decreases to 13 nS in the latter half of the cell cycle and coupling is finally lost before the next division (2 in Fig. 3). In most cases, some electrical coupling remains after each cleavage via the cytoplasmic bridges that connect newly divided sister cells, but this later disappears (1, 3, 4 in Fig. 3). The electrical coupling is reestablished between the micromeres and the neighbouring macromeres after the fifth and sixth divisions, although the cells of these pairs are not sisters (5 in Fig. 3). There is no significant coupling between other pairs of blastomeres up to the 60-cell stage. Intercellular current intensity is independent of transjunctional voltage, nondirectional and sensitive to 1-octanol, indicating that it is gated through gap-junction channels. However, when a fluorescent dye, Lucifer Yellow, is injected into individual blastomeres it does not diffuse between any of the blastomeres. Diffusion of the dye is only recognized after the seventh division (120-cell stage) and is independent of cell type. Since a significant delay in the formation of the archenteron is induced in embryos treated with 1-octanol during the 16-to 60-cell stages, electrical communication between blastomeres may play a role in embryonic induction by the micromeres (Yazaki et al., 1999). If the micromeres are deleted from the embryos, invagination of the archenteron is markedly delayed, although the descendants of macromeres finally form not only the archenteron but also the skeleton, instead of the deleted micromeres (Horstadius, 1939).

The plasma membrane of the micromeres is distinguished from that of other blastomeres by its smooth surface (Fig. 4) and its specific electrophysiological characteristics (Fig. 7, Table 1). The surface membrane of the zygote is microvillous. When the zygote divides, the resulting blastomeres have two distinct membrane regions: one is the egg-derived villous region (the apical membrane furthest away from the cleavage furrow) and the other a newly formed smooth region (the basal membrane adjoining the cleavage furrow). The blastomeres never adhere to other cells at the apical membrane area. When two blastomeres are adjacent at the apical membrane region, the two cells independently develop into separate larvae, whereas if they are adjacent at the basal membrane region, they form one larva by association of the two cells (Kuraishi, 1989). An egg surface-specific antibody purified by immunological absorption with eggs from different species has been shown to stain the apical membrane, but does not stain the basal membrane (Yazaki, 1984; Yazaki and Uemura, 1989). After furrowing is completed, shrinkage of the cortical network of actin filaments lining the original egg membrane occurs so that the basal membrane in the cleavage plane is an area of smooth membrane without either microvilli or a cortical actin network (Yazaki, 1991). The appearance of the micromere membrane and the fact that only a small area of it binds the egg surface-specific antibody shows that it is mainly composed of basal membrane, which differs from both the apical membrane and its underlying cortical structures.

The micromeres of sea urchins Arbacia punctulata and Hemicentrotus pulcherrimus lack the cortical pigment granules that occur in other blastomeres (Dan, 1954; Dan, 1960). In both mesomeres and macromeres of the 16-cell stage embryos, cortical pigment granules line the outward-facing surfaces but do not occur on the interblastomeric surfaces. Movements of pigment granules from the vegetal pole are initiated at the 2-to 4-cell stages and can be arrested by microfilament inhibitors (Belanger and Rustad, 1972). This observation suggests that the properties of the micromere membranes are specified by cortical movements from the vegetal to the animal pole, in addition to the formation of new membrane at each cleavage, as described above.

Using the whole-cell clamp technique, it was found that L-type Ca²⁺ channels are present in early sea urchin blastomeres, and that these are activated during M-phase and subsequently inactivated in S-phase of the mitotic cycle (Fig. 5). Fragmentation experiments, in which blastomeres were divided into apical and basal regions before patch clamping, show that the calcium channels are clustered in the apical membrane (Fig. 6). Meanwhile, the specific steady-state conductance, detected as K⁺ efflux, is approximately twofold higher in the basal membrane than in the apical membrane (Yazaki et al., 1995). The finding that cytochalasin B inhibits cyclic activity in both the M- and S-phases, while colchicine inactivates the L-type current in M-phase and activates a large T-type calcium current in the S-phase, suggests that channel behaviour is regulated by cytoskeletal elements (Yazaki et al., 1995). In parallel with the increase in basal membrane area as division progresses, the calcium currents in M-phase decrease from 8.5 μA cm⁻² at the 4-cell stage to 5.4 μA cm⁻² at the 8-cell stage. In 16-cell stage embryos, calcium currents are 7.4 μA cm⁻² in the mesomeres, 2.3 μA cm⁻² in the macromeres, and are not detected in the micromeres, in keeping with the fact that this plasma membrane is derived from basal membrane (Fig. 7, Table 1). The micromeres do, however, have a two- to threefold higher specific steady state conductance than either the mesomeres or the macromeres, suggesting that many more K⁺ channels are present in the micromeres than in the other blastomeres (Table 1).

Unequal division is preceded by migration of the nucleus towards the cell periphery at the vegetal pole at the 8-cell stage, when a hemicentric mitotic apparatus is formed. This gives rise to unequal blastomeres, the macromere and the micromere (Dan, 1979; Dan et al., 1983). If embryos are exposed to a 10-minute pulse of the surfactant, sodium dodecyl sulphate (SDS) during the 4-cell stage, both nuclear migration and micromere formation are inhibited. The SDS-treated embryos, which have 16 equal-sized blastomeres, are capable of development but show a decrease in the number of mesenchyme cells, have defective skeletons and show retardation or inhibition of gastrulation (Tanaka, 1976; Tanaka, 1979).

After the 8-cell stage in Ascidians, there are three successive unequal divisions in the posterior region, which always produce smaller cells posteriorly. These unequal divisions are preceded by migration of the nucleus, led by one centrosome, in the posterior direction, where a unique structure designated the centrosome-attracting body (CAB) is present (Hibino et al., 1998). The CAB, which is visible after ooplasmic segregation and is inhibited by treatment with SDS, is essential for nuclear migration, which is accomplished by the shortening of microtuble bundles formed between the centrosome and the CAB (Nishikata et al., 1999).

Neither a CAB nor CAB-like structures have been found in sea urchin embryos. We stained the endoplasmic reticulum (ER) by microinjecting a saturated solution of the fluorescent dye, dicarbocyanine DiIC₁₆ in soybean oil according to the published method (Terasaki and Jaffe, 1991). The dye spreads from the oil drop into ER membranes throughout the egg, but not into other organelles. The stained ER has the appearance of a tubular membrane network in the cortex of sea urchin blastomeres (Fig. 8A2′). In 16-cell stage embryos, there is a considerable accumulation of ER in the micromere, although a meshwork of ER is visible in the cortical layer of every blastomere (Fig. 8A1, A2, A2′). The accumulation of ER is intially observed in the vegetal half of 4-cell stage blastomeres (Fig. 8B2, B3), and subsequently accumulates at the vegetal pole of the 8-cell blastomere (arrowheads) although it is not present in another adjacent blastomere (arrows in Fig. 8C2, C3). If the embryo is treated with SDS during the 4-cell stage to inhibit the unequal cleavage at the fourth division, the accumulation of ER is also inhibited. Fig. 8D shows the equal size of blastomeres and equal distribution of ER in a quarter embryo at the 16-cell stage. This suggests that the local accumulation of ER in the vegetal half of the 4-cell stage blastomeres is the first step in the process that gives rise to the micromeres.

The cortical endoplasmic reticulum of sea urchin eggs can release and take up Ca²⁺ (Terasaki and Sardet, 1991). Using the acetoxymethyl esters of Fura-2 (Fura-2/AM), Ca²⁺ concentrations in blastomeres from the 2-to 16-cell stages were compared. Fura-2/AM is a cell-permeant dye that becomes impermeant after being hydrolysed by esterase within the cell. The Ca²⁺ concentration can be obtained as a relative value by calculating the ratio of fluorescence intensity at 510 nm emitted from dye activated at two wavelengths, 340 and 380 nm. Thus the size and thickness of cells do not affect the measurements.

Embryos were removed from their fertilization envelopes and blastomeres were separated at the 4-cell stage by placing embryos in Ca²⁺-deficient sea water and then allowing them to develop in normal sea water. This procedure results in the formation of many ‘quarter-embryos’ consisting of two mesomeres, one macromere and one micromere (Fig. 4). Fura-2/AM was loaded into the 4-cell stage blastomeres and ratio-imaging measurements were carried out at 30 s intervals over a period of 40 min (one division cycle takes 35–40 min) (Fig. 9A,B). Since the fluorescent ratios increase in every blastomere within 40 min of loading, measurements were usually carried out 1 h or more after loading.

In Fig. 9, the ratio in the area of the micromere that was measured, blastomere 1 (indicated by a white line) increases as furrowing begins (Fig. 9A3, 3′) and is maintained at a higher level than that of the other blastomeres, i.e. blastomere 2 for the macromere, and 3 and 4 for the mesomeres during the 16-cell stage (Fig. 9B). Relative Ca²⁺ concentrations were measured as the ratio between two blastomeres so that micromere was compared with macromere, mesomere with macromere and mesomere with mesomere. In the case of natural embryos, i.e. unloaded embryos, the ratios between all blastomere pairs measured was close to 1.0, which means there is no difference between the three types of blastomeres in the ratio of fluorescence intensity excited at two wavelengths. In embryos loaded with Fura-2/AM, the fluorescence intensity ratio of micromere to macromere is significantly higher than 1.0 (1.14±0.018 for Hemicentrotus pulcherrimus and 1.10±0.010 for Pseudocentrotus depressus; these values are significantly different from the ratios in mesomere/macromere and mesomere/mesomere pairs (P<<0.05), but all other blastomeres have almost the same intracellular calcium concentration (Table 2).

Just before furrowing at the fourth division, there is an elevation of [Ca²⁺]_i in the small area at the vegetal pole of the 8-cell blastomere, from which micromeres are liberated (Fig. 9A2, 2′ and A3, 3′). As previously stated, SDS-treated embryos divide equally at the fourth division and fail to make either a larval skeleton or an archenteron. In the blastomeres of such embryos, no accumulation of ER is seen (Fig. 8D) and [Ca²⁺]_i in all 16-cell stage blastomeres are similar (Table 2). It is interesting that in normal embryos, ER accumulation is detected from the 4-cell stage, while a high [Ca^2+]i is detected later, just as furrowing is initiated at the fourth division (Fig. 9A3, 3′).

Repeated elevations of [Ca²⁺]_i are measured after the fourth cleavage (Fig. 9A4, 4′–A5, 5′) and then the levels stabilize in the later stages of the mitotic cycle (Fig. 9B). The intervals between peaks of [Ca²⁺]_i range between approximately 30 s and 60 s. Oscillations do not occur in Ca²⁺-deficient sea water and are evoked by the addition of Ca²⁺ ions into the bathing medium (data not shown), suggesting that Ca²⁺ influx may initiate the Ca²⁺ oscillations. Since L-type calcium channels are absent from the micromeres (Fig. 7, Table 1), other types of Ca²⁺ channels may be involved in the initiation of these Ca²⁺ oscillations.

The rise in [Ca²⁺]_i during fertilization of sea urchin eggs is caused by an influx of Ca²⁺ (Paul and Johnston, 1978) and release from intracellular stores (Shen and Buck, 1993). Two intracellular Ca²⁺ release mechanisms regulated by inositol triphosphate (IP₃) and ryanodine have been identified in many somatic cell types (Berridge, 1993). Pharmacological studies (Sardet et al., 1992; Livingston and Wilt, 1992; Livingston and Wilt, 1995) and immunohistochemical studies using antibodies against ryanodine receptors (McPherson et al., 1992) and IP₃ receptors (Parys et al., 1994) have shown that sea urchin eggs contain both mechanisms. Transients in [Ca²⁺]_i control cell cycle progression by translational and post-translational regulation of the cell cycle control proteins pp34 and cyclin (Whitaker and Patel, 1990).

Ca²⁺ oscillations accompany fertilization in many animals including mammals, amphibia, ascidians and nemertean worms (Miyazaki et al., 1993; Kubota et al., 1993; Speksnijder et al., 1989; Stricker, 1996; Stricker et al., 1998), although distinct repetitive calcium waves after fertilization have not been observed in sea urchins. Two intracellular calcium channels have been implicated in different roles during fertilization of ascidians: ryanodine receptors cause capacitance changes, implying a net insertion of membrane in the oocyte surface, while IP₃ receptors trigger sustained [Ca²⁺]_i oscillations (Albrieux et al., 1997). In sea urchins, the ryanodine receptors localized in the cortical region of mature eggs are lost after the cortical exocytosis at fertilization, suggesting that the ryanodine receptors are linked with insertion of membrane at the egg surface (McPherson et al., 1992).

In nemerteans, microdomains of ER occur during oocyte maturation. These are closely related to Ca²⁺ oscillations after fertilization but are independent of the egg axis (Stricker et al., 1998). In ascidian eggs, in contrast, the calcium oscillations are initiated near the vegetal pole by a cortical pacemaker (Speksnijder, 1992). The calcium oscillations may influence various events after fertilization, including meiotic events and ooplasmic segregation in ascidian eggs, and mitotic events in sea urchin eggs; however, the physiological role of Ca²⁺ oscillations in embryonic cells is unclear, although in mammalian cells in culture, [Ca²⁺]_i oscillations and their frequency have been shown to be specific for gene activation (Dolmetsch et al., 1998; Li et al., 1998).

Lithium chloride vegetalizes sea urchin embryos, i.e. transforms many more cells to endoderm than occurs in normal embryos (Nocente-McGrath et al., 1991), by blocking signalling through the IP₃-pathway (Livingston and Wilt, 1990; Livingston and Wilt, 1992; Livingston and Wilt, 1995). This action is effective not only on the A–V axis of sea urchin but also on the ventral–dorsal axis of amphibian embryos. Lithium chloride is known to interfere with the intracellular Ca²⁺ concentration via the IP₃-signalling pathway and to regulate gene expression in the nucleus (Berridge, 1993), although in Xenopus laevis embryos it has recently been proposed that lithium chloride acts through inhibition of glycogen synthetase kinase-3β (GSK-3β), a member of the Wnt signalling pathway, which regulates cell fate determination in a number of diverse organisms including Dictyostelium discoideum, Drosophila melanogaster and Xenopus laevis (Klein and Melton, 1996). In the absence of a Wnt signal, GSK-3 negatively regulates the Wnt signalling pathway by phosphorylating β-catenin, which causes the breakdown of β-catenin. Overexpression of β-catenin causes vegetalization of sea urchin embryos and the lithium chloride treatment enhances the accumulation of β-catenin in the nuclei of vegetal cells (Wikramanayake et al., 1998). The function of β-catenin is thought to be mediated through direct association with transcription factors and the accumulation of this complex in the nucleus (Cavallo et al., 1997). The activity of TCF, a β-catenin binding transcriptional regulator, affects specification along the A–V axis between fertilization and the 60-cell stage (Vonica et al., 2000). According to Logan et al., 1999, β-catenin is initially recognized in the cytoplasm of the 16-cell stage micromeres of sea urchin embryos, and then, after the fifth division, high levels of cytoplasmic and nuclear β-catenin are detected in both the small and large micromeres while lower levels of β-catenin are present in the nuclei of the macromere descendants. It is not known, however, how the β-catenin autonomously increases and localizes in the vegetal cells.

It has been suggested that two functions of the micromeres in Hemicentrotus pulcherrimus, skeletogenesis and inductive ability to give rise to the endoderm, are acquired by unequal division at the fourth cleavage (Tanaka, 1976). We have found high [Ca²⁺]_i and the presence of Ca²⁺ oscillations associated with the micromeres (Table 2, Fig. 9). It is, however, pointless to seek the causes of gastrulation and skeletogenesis in the micromeres, since it is a lengthy and complex process to specify the endodermal cells (gastrulation) and mesodermal cells (skeletogenesis), including complicated regulative cellular interactions (Ettensohn and McClay, 1988; Ettensohn, 1990). For example, a gene for spicule matrix protein SM30 is transcribed in cultured micromeres as well as in intact embryos but the transcripts of SM30 are first detectable at around the onset of spicule formation (Kitajima et al., 1996). In the case of Endo 16, which encodes the surface protein of the archenteron cells, the gene is activated at the early blastula stage, before gastrulation, but not at the 16-cell stage (Nocente-McGrath et al., 1989). To my knowledge, the micromere-specific gene expression has not been identified (Giudice, 1995; Giudice, 1999).

A novel tyrosine kinase, which phosphorylates and activates GSK3β to direct cell fate specification, is found in Dictyostelium discoidium (Kim et al., 1999). In the sea urchin, β-catenin initially appears in the cytoplasm of micromeres at the 16-cell stage and accumulates in the nuclei at the 32-cell stage (Logan et al., 1999); however, transcripts of sea urchin homolog of GSK3β are expressed from unfertilized eggs to blastula in every embryonic cell (Emily-Fenouil et al., 1998). High [Ca²⁺]_i is first detectable at the time when the micromeres are about to constrict. At present we have no evidence to link the behaviour of β-catenin and [Ca²⁺]_i; however, high [Ca²⁺]_i may regulate the localization of β-catenin through phosphorylation of GSK3β by a Ca²⁺-sensitive protein kinase or tyrosine kinase, which has not been identified in sea urchin embryos.

High [Ca²⁺]_i and [Ca²⁺]_i oscillations are first detectable when the micromeres are about to constrict, though a Ca²⁺ reservoir, in the form of the ER, accumulates earlier. Since the high [Ca²⁺]_i and [Ca²⁺]_i oscillations are inhibited by 2-aminoethoxy diphenylborate (2APB), a modulator of IP₃-induced Ca²⁺ (Maruyama et al., 1997), both may be attenuated by release from the ER Ca²⁺ reservoir. The [Ca²⁺]_i oscillations at fertilization are initiated by Ca²⁺ influx following sperm entry. The present study suggest that voltage-dependent L-type Ca²⁺ channels are absent in micromeres (Fig. 7, Table 1). When the 8-cell stage blastomeres are exposed to a blocker of Ca²⁺ influx, SKF 96356 (Cabello and Shilling, 1993), the blastomeres cannot initiate furrowing, [Ca²⁺]_i increases or [Ca²⁺]_i oscillations. When applied after the elevation of [Ca²⁺]_i in the micromeres, SKF 96356 does not affect on the level of [Ca²⁺]_i, but [Ca²⁺]_i oscillations are not detected (data not shown). The initial influx of Ca²⁺ may result from activation of a calcium channel that is sensitive to stretch-stress produced by the cleavage furrowing (Lee et al., 1999; Mitsuyama et al., 1999). In this way, the micromeres are specified to have high [Ca²⁺]_i or [Ca²⁺]_i oscillations subsequent to autonomous specification of their surface membrane and cytoplasmic components, which are derived from eggs by autonomous cortical movements and cleavages.

I thank Drs Y. Kudo and K. Mikoshiba for their valuable suggestions on calcium measurements and also Ms Y. Koyama and Dr M. Abe for their collaborations on calcium data in the present paper. I greatly appreciate Dr R. Hill for critically reading the manuscript and for improving the English. My research on calcium was supported by the Fund for Special Research Projects 1997 at Tokyo Metropolitan University.