In insects, egg activation is known to occur in vivo and independently of fertilization, but its mechanisms are poorly understood. To gain understanding of these mechanisms, an attempt was made to activate the egg of Gryllus bimaculatus in vitro. It was found that meiosis resumed and was completed in unfertilized eggs treated with hypotonic buffer. Early developmental processes in activated, unfertilized eggs were investigated and compared with those in fertilized eggs. Mitosis did not progress, resulting in formation of anucleate cytoplasmic islands (pseudoenergids). Development in the activated, unfertilized eggs stopped at this stage and both yolk subdivision and cellularization did not occur. To elucidate the role of the nucleus in the developmental process to the syncytial stage in fertilized eggs, eggs were treated with aphidicolin to inhibit DNA polymerization. It was found that pseudoenergids also formed in these aphidicolin-treated fertilized eggs. These results demonstrate that pseudoenergids can increase in number independently of nuclei, suggesting that the cytoplasm rather than the nucleus plays the primary role in development to the syncytial stage in G. bimaculatus.
In animals, meiosis takes place in both spermatocytes and oocytes. Meiosis is a continuous process in the male germ line. In the female germ line, however, meiosis is usually interrupted once or twice to synchronize the completion of meiosis with the growth of the oocyte and with sperm entry (Page & Orr-Weaver 1997). Oocytes remain arrested until activated by an external signal. Echinoderms and Amphibia are suitable for studies of egg activation, and have been researched in detail. In the case of the insect, analyses of similar events are far more difficult because ovulation, fertilization and egg activation all occur in vivo (Mahowald et al. 1983), and also because the egg has a thick chorion and an abundant yolk mass (Kawamura 2001).
Pterygote insects have been grouped conventionally into two categories, holometabola and hemimetabola, based on the extent of their morphological changes at metamorphosis in postembryonic development. The cricket is one of the model insects of the hemimetabola, and its oviposition process has been investigated to a certain extent (Ai & Ishii 1984; Sugawara 1984, 1987; Sugawara & Loher 1986; Sakai & Taoda 1992; Kumashiro & Sakai 2001a, 2001b). The main female reproductive organs are a pair of ovaries, which connect with a pair of lateral oviducts (Fig. 1A). These join to form a median oviduct opening posteriorly into a genital chamber (Fig. 1D). A spermatheca, used for the storage of sperm from copulation until the time when eggs are fertilized, opens into the genital chamber independently of the oviduct (Fig. 1B). A pair of accessory glands is also present (Fig. 1C). The genital chamber connects posteriorly with an ovipositor (Fig. 1E). The first step in the oviposition process is searching. A female cricket which has mated searches for a place suitable for laying her eggs, and pierces the substrate with her ovipositor. Just after penetration by the ovipositor is completed, the female cricket slightly retracts it. This short lift of the ovipositor accompanies ovulation; that is, progression of an egg from the oviduct to the genital chamber (Sugawara 1984; Sugawara & Loher 1986). After the short lift of the ovipositor, the female cricket stays immobile for approximately 10 s. This rest phase is thought to be the time of fertilization in the genital chamber (Sugawara & Loher 1986). The egg then enters and passes through the ovipositor, and is expelled from the end of the ovipositor into the substrate.
The triggers and events of egg activation in the cricket oviposition process outlined above are not yet known. Activation affects many developmental processes in the egg. The process of embryogenesis in the cricket has been reported (Miyamoto & Shimozawa 1983; Niwa et al. 1997), but the early developmental process just after egg-laying has not been described in detail.
We were interested in activating unfertilized eggs in vitro to study egg activation and early development in the cricket. First, we examined the possibility that unfertilized eggs of Gryllus bimaculatus, the two-spotted cricket, could be activated in vitro. Second, we investigated and compared the early developmental processes of (i) unfertilized eggs activated in vitro and (ii) laid fertilized eggs activated in vivo using staining techniques. To gain a better understanding of mechanisms underlying the early developmental processes in G. bimacullatus, we did cytological experiments using aphidicolin, a specific inhibitor of DNA polymerase-α. We show that the early developmental process in the activated unfertilized egg can be mimicked successfully in the fertilized egg treated with aphidicolin, and we discuss the behavior of the nucleus and cytoplasm in these events.
Materials and Methods
Animals and eggs
Colonies of nymphs of the two-spotted cricket G. bimaculatus were purchased from Scope (Okayama, Japan), and maintained on an artificial diet for insects (Oriental Yeast, Suita, Osaka, Japan) at 28°C.
Fertilized eggs laid on piled moistened tissue paper were washed with 10% commercial bleach for 10 s, rinsed with deionized water for 10 s, washed with 70% ethanol for 10 s and finally rinsed twice for 10 s with deionized water. Incubation in phosphate-buffered saline (PBS) or mineral oil (M-8410; Sigma, St Louis, MO, USA) followed. The eggs were allowed to develop at 28°C.
Unfertilized eggs were dissected from ovaries of mature virgin females of G. bimaculatus, and incubated at 28°C in PBS or mineral oil.
The unfertilized eggs incubated in PBS were transferred to distilled water and incubated for 20 min, or to 10 mm hydrogen peroxide and incubated for 10 min. After activation with distilled water or hydrogen peroxide, the eggs were returned to PBS. All procedures were done at 28°C.
Injection of dextran dye or aphidicolin into eggs
Our method of injection of dextran dye is a modification of that described in Ho et al. (1997). Eggs were lined up on a piece of double-sided adhesive tape on a microscope slide bounded by a corral of piled adhesive vinyl tape. They were left to desiccate for 10 min before they were covered with mineral oil. The eggs were injected using a micromanipulator with approximately 20 nL of 10 mg/mL dextran tetramethylrhodamine 10 000 MW (lysine fixable; D-1817; Molecular Probes, Eugene, OR, USA) or 400 µg/mL aphidicolin (diluted from a 10 mg/mL stock in dimethyl sulfoxide (DMSO)). After injection, eggs were allowed to develop for varying amounts of time at 28°C before fixation or examination under a fluorescence microscope.
Our method of 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI) staining is a modification of that described in Scali & Tinti (1992). Unfertilized eggs were fixed in 4% paraformaldehyde in PBS at 4°C overnight, and dehydrated in 25, 50 and 75% methanol in 0.05% Tween-20 in PBS (PBT), each for 30 min, and in 100% methanol at 4°C overnight. The chorions of eggs were removed mechanically by tungsten needles. The eggs were hydrated again in 75, 50 and 25% methanol in PBT, each for 30 min, and washed with PBS three times, each for 10 min. Eggs were then stained with 0.2 µg/mL DAPI in methanol for 10 min in the dark at room temperature, followed by washing with PBS three times, each for 5 min, in the dark at room temperature. The eggs were examined under a fluorescence microscope.
Fertilized eggs were placed in 50% commercial bleach for 1 min, followed by mechanical removal of chorions by tungsten needles, in PBS, before fixing in 4% paraformaldehyde in PBS at 4°C overnight. The method of DAPI staining was the same as that for unfertilized eggs.
Rhodamine phalloidin staining
The methods of dechorionation and fixation were the same as those for DAPI staining. The fixed eggs were incubated in 1 U/mL rhodamine phalloidin (R-415; Molecular Probes) at 4°C overnight with gentle shaking, washed twice with PBT for 30 min each, and washed with PBS for 30 min.
Indirect immunofluorescence staining for microtubules
Eggs were placed in 50% commercial bleach for 1 min, followed by mechanical removal of chorions by tungsten needles, in PBS. The eggs were then transferred to fresh PBS. We added paclitaxel (T-7402; Sigma) from a stock solution of 2.5 mm in DMSO to the PBS to a final concentration of 10 µm. The eggs were overlaid with an equal volume of heptane and shaken for 90 s. The eggs were then transferred to PBS, overlaid with an equal volume of formalin (37% formaldehyde with 13% methanol) and two volumes of heptane, and shaken for 15 min, followed by washing three times with PBS for 5 min each.
The eggs were incubated in 1% polyoxyethylene(10) octylphenyl ether (Triton X-100) in PBS for 15 min with shaking, followed by washing three times with PBS for 10 min each. The eggs were then incubated in a PEMBAL solution (100 mm PIPES, 1 mm EGTA, 2 mm MgCl2, 1 mg/mL albumin from bovine serum (fatty acid and globulin free; A-0281; Sigma), 0.01% sodium azide and 2.2 mg/mL l-lysine). To detect microtubules, the eggs were incubated with anti-tubulin-α antibody (AB6160-5000; AbCom, Cambridge, UK) at a dilution of 1:20 in PEMBAL solution, for 2 h with shaking at room temperature, followed by washing with PEMBAL solution three times for 10 min each. The eggs were then incubated with secondary antibody (anti-IgG coupled with Alexa-488; A11006; Molecular Probes) at a dilution of 1:200 in PEMBAL solution, for 2 h with shaking at room temperature, followed by washing three times with PEMBAL solution for 10 min each.
Effects of distilled water and hydrogen peroxide on egg activation
To examine whether mature unfertilized eggs of G. bimaculatus could be activated in vitro, we treated them with either hypotonic buffer (Mahowald et al. 1983; Sawa & Oishi 1989) or a solution of hydrogen peroxide (Sato et al. 1998, 2001). Mature oocytes of G. bimaculatus were expected to remain arrested at metaphase of the first meiotic division (metaphase I) from results in Drosophila (Page & Orr-Weaver 1997) and other insects (Sawa & Oishi 1989). This postulation did not contradict the results of DAPI staining: only one nucleus was discerned in every mature G. bimaculatus oocyte just after dissection (Table 1). If the mature oocyte is effectively activated, the meiotic cell cycle should resume and the nucleus in the oocyte should start to divide. We therefore took two (or more) nuclei side by side in one oocyte as the indicator of successful egg activation. Eggs treated with distilled water (hypotonic buffer) or hydrogen peroxide were incubated in PBS for 1 h and were stained with DAPI to estimate activation efficiency. Two nuclei stained with DAPI were discerned in approximately four-fifths of the total number of eggs treated with hypotonic buffer and approximately two-thirds of the total number of eggs treated with a solution of hydrogen peroxide (Table 1).
Table 1. Effects on egg activation of mature unfertilized Gryllus bimaculatus eggs with distilled water (DW) and hydrogen peroxide (H2O2)
Type of egg activation treatment (timing of fixation)
Rate of successful egg activation (%)
One nucleus observed
Two nuclei observed
None (just after dissection)
DW (60 min after activation)
H2O2 (60 min after activation)
None (80 min after dissection)
Early developmental process in fertilized eggs
Freshly laid fertilized eggs of G. bimaculatus were light yellow in color and approximately 2–3 mm long (Fig. 2). One end of their longitudinal axis was slightly pointed and the other was blunt. The pointed end is called the ‘anterior pole’ because the developed embryo hatches out of this end (Miyamoto & Shimozawa 1983). The concave and convex sides are dorsal and ventral, respectively (Miyamoto & Shimozawa 1983; Niwa et al. 1997). At 15 h after oviposition, the eggs developed to the stage that black spots were distributed all over them. At 60 h after oviposition, subdivision of yolk plasmodium was complete (Fig. 2).
A nucleus (chromosomes in a primary oocyte) of a fertilized egg, stained with DAPI, was initially observed just below the dorsal surface approximately in the center of the egg (Fig. 3A,B). Within 1 h of oviposition, two nuclei (chromosomes in a secondary oocyte and a polar body) arranged parallel to the dorsal surface were observed (Fig. 3C), and within 3 h, four (sometimes three) nuclei (a female pronucleus and chromosomes in three (or sometimes two) polar bodies) were observed (Fig. 3D). Images of second meiotic anaphase were observed using indirect immunofluorescence staining for microtubules (Fig. 4K,L). The nuclei subsequently proliferated and migrated posteriorly, and 15 h after oviposition, a number of nuclei were distributed all over the egg (syncytial stage; Fig. 3E). Images of mitotic division were observed using indirect immunofluorescence staining for microtubules (Fig. 4A–J).
A spotted structure stained with dextran tetramethylrhodamine, which we interpreted as an energid, was initially observed just below the dorsal surface at approximately the center of the egg (Fig. 5A). Approximately 3 h after oviposition, the energid started to proliferate somewhat regularly (Fig. 5B), migrated posteriorly, and 15 h after oviposition, energids were distributed all over the egg (Fig. 5C).
The energid has been defined as a unit of nucleus and halo of cytoplasm in the insect egg (Chapman 1998). We double-stained the same fertilized egg 15 h after oviposition with dextran tetramethylrhodamine and DAPI, and found that both stains stained the same positions in the egg (Fig. 6C,D). Because dextran tetramethylrhodamine is known to stain the cytoplasm (Ho et al. 1997) and DAPI has been thought to stain nuclei (Scali & Tinti 1992), we interpreted these spotted structures containing a nucleus and halo of cytoplasm as energids. The results of staining with dextran tetramethylrhodamine 15 h after oviposition also revealed that the black spots observed on the eggs were energids (Fig. 6A,B).
Energids have a characteristic star-like shape and are connected to one another by a cytoplasmic network (Fig. 7A). They increase in number by asynchronous division (Fig. 7E–G). Rhodamine phalloidin staining showed that each superficial energid was surrounded by a patch-like structure rich in fibrous actin: an ‘actin cap’ (Karr & Alberts 1986; Fig. 7D).
Early developmental process in activated unfertilized eggs
The process of meiotic nuclear division in unfertilized eggs activated with distilled water was the same as that in fertilized eggs up to 3 h after activation or oviposition (Fig. 3B–D,G–I). The image of a spindle and chromosomes segregating successfully was observed in an activated unfertilized egg at 2 h after activation treatment, using indirect immunofluorescence staining for microtubules and DAPI staining (Fig. 4M,N). It is possible that the spindle and chromosomes were at telophase of the second meiotic division and that meiosis in the egg was nearly complete. But after that, and different to the situation in normal fertilized eggs, we could not recognize the proliferation of nuclei with DAPI staining (Fig. 3J), whereas the structures stained with dextran tetramethylrhodamine increased in activated unfertilized eggs. These structures had a star-like shape and a cytoplasmic network similar to those in the energids in normal fertilized eggs (Fig. 7B). The expanding pattern of these energid-like structures in the activated unfertilized eggs was also similar to that in the normal fertilized eggs, and the activated unfertilized eggs developed to the stage that energid-like structures were distributed all over them (Fig. 5D–F). The activated unfertilized eggs at this stage resembled normal fertilized eggs at the syncytial stage in the distribution of the energid-like structures and energids, but were different from the normal eggs in the distribution of nuclei. We provisionally call this stage of the activated unfertilized egg the ‘pseudosyncytial stage’, and the energid-like structures in the activated unfertilized egg ‘pseudoenergids’.
While the expanding pattern of anucleate pseudoenergids in the activated unfertilized eggs was similar to that of energids in normal fertilized eggs (Fig. 5), the progression rate to the pseudosyncytial stage in the activated unfertilized eggs was very different from that to the syncytial stage in normal eggs. While it took only approximately 15 h for the fertilized egg to develop to the syncytial stage after oviposition, development of the activated unfertilized egg proceeded more slowly and it took approximately 72 h after in vitro activation to reach the apparent syncytial stage (Fig. 2). The most peculiar feature of the pseudoenergids in the activated unfertilized eggs was that they increased in number not by division but by aggregation of cytoplasm (Fig. 7H–J). We could not confirm any signs of actin caps in the activated unfertilized eggs. The progression of development in the activated unfertilized eggs stopped at the pseudosyncytial stage, and subdivision of yolk plasmodium did not occur. The eggs ultimately deteriorated.
Aberrant developmental process in fertilized eggs when DNA synthesis is inhibited by aphidicolin
To block nuclear division during mitosis, we used aphidicolin, a specific inhibitor of DNA polymerase-α. Aphidicolin and dextran tetramethylrhodamine were co-injected into fertilized eggs 1 h after oviposition, and then the eggs were fixed and stained with DAPI 15 h after oviposition to examine the distribution of nuclei. The distribution of nuclei and energids (and energid-like structures) in control and aphidicolin-injected fertilized eggs at 15 h after oviposition are shown in Figure 8. In control fertilized eggs injected with dextran tetramethylrhodamine alone, both nuclei and energids proliferated all over the egg (Fig. 8C,D). In the eggs injected with aphidicolin, only energid-like structures which lack nuclei increased in number (Fig. 8A,B). The distribution of nuclei and energid-like structures in fertilized eggs injected with aphidicolin was similar to that in unfertilized eggs activated in vitro. The anucleate energid-like structures in the eggs injected with aphidicolin had a star-like shape and a cytoplasmic network (Fig. 7C). In most eggs injected with aphidicolin, the energid-like structures increased in number by aggregation of cytoplasm, but in some eggs injected with aphidicolin, they increased by division, irrespective of the lack of discernible signals of DAPI. There was no sign of actin caps in the eggs injected with aphidicolin.
In insects, egg activation generally occurs in vivo, so analyses of this event are thought to be difficult. It has been feasible only to a limited extent to mimick some of the steps of egg activation in vitro, and unfertilized eggs of some insects have been activated in vitro without fertilization. These insects are Locusta migratoria (Orthoptera; Lanot et al. 1988), Carausius morosus (Phasmida; Pijnacker & Ferwerda 1976), Bombyx mori (Lepidoptera; Kawamura 2001), Drosophila mercatorum, D. melanogaster, D. immigrans (Diptera; Mahowald et al. 1983; Page & Orr-Weaver 1997; Heifetz et al. 2001), Psycoda cinerea (Diptera; Sander 1985; Sander & Feddersen 1985), Athalia rosae (Hymenoptera; Sawa & Oishi 1989), Venturia (Nemeritis) canescens (Hymenoptera; Salt 1965; Sander & Feddersen 1985), Nasonia vitripennis (Hymenoptera; King & Rafai 1973), Campoletis sonorensis (Hymenoptera; Vinson & Jang 1987) and Pimpla turionellae (Hymenoptera; Went & Krause 1973). Almost all of these species in which eggs can be activated in vitro have been reported to have the ability to reproduce parthenogenetically under natural conditions (Stalker 1954; Went 1982; Mahowald et al. 1983; Sawa et al. 1989). G. bimaculatus is not known to reproduce parthenogenetically. We examined the possibility that G. bimaculatus eggs would be able to be activated in vitro like those of the insects which can reproduce parthenogenetically. It was anticipated that G. bimaculatus eggs activated in vitro would not develop normally because of their inability to reproduce parthenogenetically. If this postulation were true, we thought it would be useful to compare the two types of early development – in eggs activated in vivo and in vitro– to obtain information on egg activation.
In Drosophila, mature ovarian eggs have a wrinkled shape, presumably due to dehydration (Mahowald et al. 1979). Because ovulated eggs are expanded, it seems possible that the process of rehydrating the egg as it passes down the oviduct may lead to activation. Mahowald et al. (1983) examined this possibility and showed that unfertilized Drosophila eggs could be activated successfully by immersing unfertilized eggs in hypotonic buffers. In G. bimaculatus, ovarian eggs are expanded rather than wrinkled, but we expected that somewhat similar mechanisms of activation exist as in Drosophila, and we examined the possibility of and succeeded in activating unfertilized eggs of G. bimaculatus by immersion in distilled water. In Xenopus eggs, tyrosine phosphorylation plays an important role in initiating a transient calcium concentration and subsequent egg activation. Hydrogen peroxide is able to induce tyrosine phosphorylation and egg activation-like responses (Sato et al. 1998, 2001). We also examined the ability of hydrogen peroxide to induce tyrosine phosphorylation and egg activation-like responses in the eggs of G. bimaculatus, but the use of distilled water was found to be easier and more effective. The ability to activate unfertilized eggs in vitro with distilled water clearly indicated that activation is independent of fertilization or hormonal interaction, both of which had been previously proposed as triggers of activation (Went 1982; Lanot et al. 1988, 1989).
In the case of unfertilized eggs of A. rosae activated in vitro, arrest at metaphase I was released and nuclear division proceeded to telophase I within 20 min of egg activation. Based on the fact that this 20-min period from metaphase I to telophase I corresponds to the time required for egg activation with distilled water, Sawa & Oishi (1989) suggested that progression from metaphase I to telophase I might represent the actual process in irreversible egg activation in vitro. In contrast, in the case of unfertilized G. bimaculatus eggs activated with distilled water, it took approximately 1 h to proceed from metaphase I to telophase I, while the period of time required to activate unfertilized eggs in vitro is only 20 min. Thus, it was supposed that the actual process of irreversible activation in G. bimaculatus is not the progression from metaphase I to telophase I, and the initial event of irreversible activation might be completed within 20 min.
Within 3 h of activation with distilled water, a nucleus from an unfertilized egg divided into four daughter nuclei. This phenomenon was interpreted as being meiosis. Based on the fact that the dividing pattern and timing of the process in activated unfertilized eggs corresponded to that of normal fertilized eggs, and the image of a spindle at telophase of a second meiotic division was observed, it was supposed that meiosis of activated unfertilized eggs might proceed normally. After meiosis was completed, the progression of mitosis was observed in normal fertilized eggs with DAPI staining, but could not be observed in activated unfertilized eggs. This lack of mitosis in activated unfertilized eggs could be related to the finding that development of activated unfertilized eggs stopped before the subdivision of yolk plasmodium occurred.
The techniques used for in vitro egg activation in this paper could differ from the natural means. Insufficient egg activation is a possible cause of developmental failure in activated unfertilized G. bimaculatus eggs. Unfertilized eggs of V. canescens activated with distilled water revealed a spectrum of developmental anomalies (Sander & Feddersen 1985). In most eggs, meiosis did not progress but the amount of chromatin apparently increased. In some eggs, meiosis resumed but mitosis did not progress. In yet other eggs, mitosis progressed and cleavage nuclei spread all over the egg, but not a single egg established a cellular blastoderm. These findings suggest that the activating stimulus may not exert an all-or-nothing effect, but rather, may pull several developmental triggers independently. Insufficient egg activation then may trigger some developmental processes. In the case of insects that are unable to reproduce parthenogenetically, developmental failure in activated unfertilized eggs could occur, even if egg activation is complete. A lack of sperm contribution is another possible cause of developmental failure in activated unfertilized G. bimaculatus eggs. Advocated materials conveyed by a spermatozoon in insects include the paternal genome, centrosomes (Callaini et al. 1999) and some materials contained in the sperm tail (Hennig 1988).
Ho et al. (1997) reported that, in early development in the locust egg, energids could be stained with dextran tetramethylrhodamine before cellularization occurred, but not after, because the dye could not incorporate into the energids through cell membranes. The development of the activated unfertilized egg of G. bimaculatus stopped at the pseudosyncytial stage and in the condition that the anucleate pseudoenergids could be stained with dextran tetramethylrhodamine. The development of the activated unfertilized egg was therefore possibly stopped before cellularization occurred. To determine this conclusively, further examination using electron microscopy is needed.
The pseudoenergids in activated unfertilized eggs were similar to the energids in fertilized eggs in shape (Fig. 7A,B) and distribution pattern (Fig. 5), but different in forming pattern (Fig. 7E–J), and lack of nuclei (Fig. 3) and actin caps (Fig. 7D). This phenomenon of pseudoenergids coming into existence and increasing in number in activated unfertilized eggs was probably ‘pseudocleavage’, which has been reported in some insects (Counce 1973). But ‘pseudocleavage’ differed from our observation in that it occurred in a small proportion of unfertilized eggs without activation treatment.
To interpret reasonably the phenomena occurring in activated unfertilized eggs, we intended to dissociate the parts controlled by the nucleus and cytoplasm in these processes. We used aphidicolin, a specific inhibitor of DNA polymerase-α, which inhibits DNA synthesis and, consequently, nuclear replication (Raff & Glover 1988; Raff & Glover 1989). Meiosis can be completed without DNA synthesis, but DNA synthesis is indispensable to mitosis. Therefore, it was anticipated that meiosis could be completed but subsequent mitosis could not progress in aphidicolin-treated eggs, and the phenomena occurred after the completion of meiosis could be interpreted not to be involved with nuclei. We injected aphidicolin into fertilized eggs 1 h after oviposition, and confirmed at 15 h after oviposition that nuclei did not proliferate at all, while anucleate energid-like structures increased in number, and were similar to those in activated unfertilized eggs in terms of shape, distribution pattern and the lack of actin caps. In the energid-like structures in most aphidicolin-treated fertilized eggs, the forming pattern of cytoplasmic aggregation was also the same as that in activated unfertilized eggs. Therefore, it is likely that the energid-like structures of aphidicolin-treated fertilized eggs corresponded to the pseudoenergids of activated unfertilized eggs. It was curious that the energid-like structures in some aphidicolin-treated eggs proliferated by division. A possible cause of this phenomenon was that, in some aphidicolin-treated eggs, the concentration of aphidicolin was not high enough and the nuclei became abnormal, and although were not stained with DAPI, did not lose the ability to proliferate by division. At least in many of the aphidicolin-treated eggs, we succeeded in making fertilized eggs mimic activated unfertilized eggs, and confirmed that anucleate pseudoenergids could increase independently of the nucleus. These results suggest that the cytoplasm rather than the nucleus plays the primary role in early development, at least to the syncytial stage, in G. bimaculatus. The materials in the cytoplasm which controlled forming and increasing of pseudoenergids are possibly maternal messenger RNA and proteins. Identification of the materials and elucidation of the mechanisms in these events awaits further investigation.
To understand the nature of activation should provide the basis for a systematic approach for in vitro fertilization of crickets. This should be useful to analyze mutant eggs which cannot be fertilized in vivo. The information obtained in this paper can be considered a prerequisite for such a study.
We thank Kugao Oishi for critical reading of the manuscript. We thank Noriyuki Shimizu and Yuko Sakaue for cricket rearing.
This work was supported by a grant from the Ministry of Education, Culture, Sports, Science and Technology of Japan to I. S., T. M., H. O. and S. N.