F-actin domains in the syncytial blastoderm of the dipteran Ceratitis capitata

Early embryonic development in dipteran insects requires several nuclear divisions, which lead to the formation of a large organized syncytial cell that later cellularizes. When the first somatic nuclei reach the periplasm the embryo surface bulges out and the nuclei are enclosed in cytoplasmic protuberances, also called caps. These structures, already described by scanning electron microscopy of Drosophila (Turner and Mahowald, 1976), Calliphora (Lundquist and Löwkvist, 1984) and Ceratitis (Callaini and Fanciulli, 1987), change in morphology during the mitotic cycle. Several studies on Drosophila revealed the presence in these structures of an extensive cytoskeleton of microtubules, microfilaments, spectrin, myosin, actin-binding proteins and intermediate filaments (Warn et al., 1980, 1984, 1987; Walter and Alberts, 1984; Karr and Alberts, 1986; Warn, 1986; Warn and Warn, 1986; Lutz and Kiehart, 1987; Kellog et al., 1988; Miller et al., 1989; Pesacreta et al., 1989). These cytoskeletal elements, which rearrange during the syncytial mitoses, mediate the high level of organization of the embryo cortex in which microfilaments and microtubules appear to play many important roles (Edgar et al., 1987). Actin organization has been examined by immunofluorescence techniques during the syncytial mitoses of Drosophila embryos (Warn and Magrath, 1983; Karr and Alberts, 1986; Kellogg et al., 1988; Miller et al., 1989) but, due to the small dimensions of this embryo, detailed analysis of microfilament distribution is difficult. Moreover, the structural basis for the mechanism by which nuclei are held in the proper position and neighbouring spindles separate during the syncytial mitoses has not been clarified. Therefore we chose a dipteran insect with large eggs and very similar development to Drosophila. In Cer atitis the somatic nuclei reach the embryo surface after nine intravitelline mitoses and pole cells bulge out at the posterior end of the egg during nuclear cycle 10. The somatic nuclei divide four times in the periplasm and give rise to a syncytial blastoderm that later cellularizes (Callaini and Fanciulli, 1987). In this case the actin caps above each nucleus are widely spaced and their morphological changes during the mitotic cycle are easily visible. The laser scanning confocal microscope, which provides serial optical section of thick specimens, allowed us to appreciate changes in F-actin organization in the cortical region.

Embryos of the Mediterranean fruit fly Ceratitis capitata Wied were collected from a laboratory culture, dechorionated in a 50% commercial bleach solution and washed with distilled water. Excess liquid was removed by blotting on tissue paper for different preparations.

For immunofluorescence observations the ages of the embryos were determined by direct observation with interference contrast or by counting the somatic nuclei, stained with the DNA specific dye Hoechst 33258.

After removal of excess liquid, embryos at appropriate stages were collected on a plastic film (Parafilm), allowed to develop on melting ice for 1 h and recovered at 24°C for 15 min.

The dechorionated embryos were transferred to a solution containing 2.5 ml of 4% paraformaldehyde, 5 ml of heptane and 0.5 μM taxol and gently agitated for 30 min. The embryos were then rinsed in PBS and their vitelline membrane was removed with fine needles.

For actin staining the embryos were incubated for 30 min in 1.5 μg/ml phalloidin labelled with rhodamine (Molecular Probes, Eugene, OR).

For microtubule staining the embryos were incubated 1 h in PBS containing 0.1% bovine serum albumin and reacted for 30 min with a monoclonal antibody against α-tubulin (Amersham, Bukinghamshire, UK; 1:200 dilution in PBS). After several rinses in PBS, the embryos were incubated with a rabbit anti-mouse fluorescein-conjugated IgG (Cappel, Durham, NC).

Nuclei were visualized after incubation for 3 min in 1 μg/ml of the DNA-specific dye Hoechst 33258.

Finally the embryos were washed in PBS and mounted in 90% glycerol containing 2.5% n-propyl gallate to reduce photobleaching (Giloh and Sedat, 1982).

Fluorescence observations were carried out with a Leitz Aristoplan microscope equipped with rhodamine (Rh), fluorescein and UV filters. Photomicrographs were taken with Kodak Tri-X pan film and developed in Kodak HC 110 developer for 7 min at 20°C.

The distribution of microfilaments was also described using a MRC-500 Laser Scanning Confocal apparatus (Bio-Rad Microscience, Cambridge, MA). It was mounted on a Nikon Optiphot microscope equipped with a 63× objective. This technique provides optical thin sections through the sample.

Embryos were fixed in the trialdehyde solution of Kalt and Tandler (1971) for 2 h. After rinsing in cacodylate buffer, pH 7.2, the embryos were postfixed in 1% osmium tetroxide for 2 h. After washing with distilled water, the samples were dehydrated through an ethanol series. After treatment with propylene oxide the embryos were embedded in an Epon-Araldite mixture and polymerized at 50°C for 48 h. (1 μm) Sections were obtained with an LKB ultratome and stained with 1% toluidine blue in 1% sodium borate.

Embryos were fixed in 1% paraformaldehyde in PBS for 30 min and dehydrated through an ethanol series at −20°C. Samples were infiltrated with Lowicryl resin, embedded overnight at −20°C using UV light, thin sectioned with a diamond knife and collected on nickel grids. Sections were hydrated with PBS for 10 min, blocked in 0.1% bovine serum albumin for 30 min, incubated in primary monoclonal actin antibody (Amersham, 1:200 dilution in PBS) for 1 h, washed with several changes of PBS and reacted with secondary goat anti-mouse IgG-conjugated 5 nm colloidal gold (Biocell, Cardiff, UK; dilution 1:20 in PBS) for 1 h. Sections were washed in PBS, postfixed in aqueous 1% glutaraldehyde, washed with distilled water, poststained at room temperature with 2% aqueous uranyl acetate for 5 min, and treated with lead citrate for 1 min. Sections were observed and photographed on a Philips CM10 electron microscope.

Cortical microfilaments formed a surface layer a few micrometres deep in the cortex of the pre-blastoderm embryo. By nuclear cycles 9-10, after the somatic nuclei reached the periplasm, the F-actin was concentrated in cortical caps between the nucleus and the plasma membrane. These cortical actin caps underwent cycle-specific reorganization during each mitosis of the syncytial blastoderm, very similar to that described for the early Drosophila embryo (Warn et al., 1984; Karr and Alberts, 1986). Fig. 1A, B shows a surface view of whole mounts of Ceratitis embryo during the late metaphase of nuclear cycle 11, stained with Rh-phalloidin to reveal F-actin. The conventional fluorescence microscope revealed the presence of two morphologically distinct F-actin domains. One domain is constituted of the microfilaments underlying the plasma membrane. Actin was concentrated above the somatic nuclei, where the plasmalemma was more convoluted and folded, and was reduced between the caps. In this region some microfilaments were associated with the short microprojections covering the whole embryo surface. At this time, the cortical caps appeared as fluorescent elliptical structures crossed by a narrow area apparently devoid of Factin staining. This central area increased in size throughout the mitotic cycle and the forming poles showed strong phalloidin staining (not shown). The edges of the caps were also devoid of fluorescent structures. A second F-actin domain was observed when the same preparations were examined at a focal plane just below the embryo surface. This Rh-phalloidin staining consisted of small fluorescent aggregates which localized around the syncytial nuclei throughout the mitotic cycle. The DNA staining pattern (not shown) revealed a metaphase nucleus located at the centre of each cluster of actin aggregates and below each actin cap. At this level of focus only small isolated aggregates and a diffuse fluorescence were observed.

These observations were confirmed when metaphase preparations similar to those in Fig. 1A, B were examined with a laser scanning confocal microscope in different focal planes. Top views showed that F-actin was concentrated in closely packed fluorescent folds, rather variable in size and shape, presumably corresponding to plasma membrane infoldings, and in small thin rings, presumably corresponding to submembranous actin filaments lining the microvillar-like projections (Fig. 1C). As observed by conventional immunofluorescence, the staining in the central area and at the edges of the caps is much reduced. At a focal plane just below the cortical actin layer, two additional F-actin distributions were identified. One pattern of distribution consisted of irregular aggregates also seen by fluorescence microscope; these aggregates were dispersed in the periplasm immediately beneath the bright fluorescent surface folds and concentrated in irregular clusters near the nuclear region (Fig. 1D). A second staining pattern, consisting of thin fibrillar material, probably actin bundles, crossed the regions between the clusters of aggregates. These filament bundles were clearly visible at a slightly lower focal level and formed an extensive interconnected network delimiting oval areas lacking any detectable fluorescent labelling (Fig. 1E, F). Double staining with Rh-phalloidin and anti-tubulin antibody showed that the dark areas corresponded to the spindle regions (Fig. 2A, B). Actin filaments formed a cage around the spindles, linking them together (Fig. 1E). Some actin aggregates were interspersed among the filament bundles, but most of them were concentrated around the spindles, forming a shell-like envelope which underwent morphological changes during the mitotic cycle. Actin aggregates were dispersed around the nuclei at interphase (Fig. 3A), but the distribution was no longer uniform after the nuclei entered mitosis. The fluorescent aggregates concentrated at the extremities of the mitotic apparatus at metaphase (Fig. 3B). This close disposition of the aggregates at the poles of the mitotic apparatus was more evident during spindle elongation at anaphase (Fig. 3C). Serial optical section and double labelling with anti-tubulin antibody showed that the aggregates were disposed in a cap-like envelope which surrounded the poles of the spindles. Often, very conspicuous rings can be observed (Fig. 1E, F). At higher magnification they seem to be constituted of several small F-actin aggregates disposed in circular figures (Fig. 2A, B). Consecutive optical sections suggest that these aggregates are arranged in ovoid clusters (not shown).

A further actin domain, recognized as a bright spot during metaphase, was detected inside the polar cap-like envelopes (Fig. 3B). This fluorescence expanded during anaphase (Fig. 3C) and was diffused at telophase (Fig. 3D). At interphase this pattern of actin staining was not apparent. No aggregates were observed inside the polar caps, but thin fluorescent threads, apparently connecting the bright spots with the periplasmic actin network, were seen (Fig. 3C). Double labelling with anti-tubulin antibody showed that the bright spots co-localized with the poles of the mitotic apparatus, suggesting a close relation with the centrosomal region (Fig. 2A).

To find cytoplasmic structures corresponding to the actin aggregates, we examined semithin sections of syncytial blastoderm embryos. Many dark particles, corresponding in size and numbers with the actin aggregates observed by immunofluorescence, were visible in the cortical region (Fig. 4A, B). These cytoplasmic particles were localized around the spindle region and during the mitotic cycle underwent changes in distribution that recall the behaviour of actin aggregates. Ultrastructural investigations using monoclonal anti-actin and colloidal gold-conjugated secondary antibodies revealed that actin is particularly concentrated in the cytoplasmic granules that surround the mitotic spindle (Fig. 4C).

When the embryos are exposed for 1 h at 0°C and briefly recovered at 24°C before fixation the microtubule organizing centers dissociate from the spindle regions. Short microtubule regrowth from these foci and miniasters can be observed in the cytoplasm among the spindles (Fig. 5A). Rh-phalloidin staining shows that, when the position of the microtubule organizing centres is modified by cold treatment, the actin aggregates behave differently from those in untreated embryos and, are disposed in irregular clusters in the proximity of the spindle regions (compare Fig. 5B with Fig. 1B).

Although fluorescein or rhodamine-labelled phalloidin staining (Warn et al., 1984; Warn, 1986) and anti-actin antibodies (Karr and Alberts, 1986) were found to localize in the cortex of the Drosophila embryo, microfilaments have not been detected by transmission electron microscopy of sections of fixed eggs at the syncytial blastoderm stage (Stafstrom and Staehelin, 1984). In agreement with previous reports on the Drosophila embryo, the present findings on the Ceratitis capitata embryo show that filamentous actin is associated with the plasma membrane within the microprojections and surface infoldings. Moreover, the diffuse fluorescence seen in the periplasm of the same stage of the Drosophila embryo with Rh-phalloidin and anti-actin antibodies may correspond to the network of actin filaments observed by confocal microscopy in the periplasm of the early Ceratitis embryo.

Previous studies failed to reveal significant bands of microfilaments associated with the cleavage zone of the mitotic spindles of Drosophila syncytial embryo (Warn et al., 1984; Karr and Alberts, 1986). Moreover, myosin also failed to accumulate in the region between the two separating caps (Young et al., 1991), as would be expected for a contractile ring that will be located in the equatorial plane of division between newly forming caps. This is a surprising feature because the actomyosin contractile ring is a principal and ubiquitous structural component of animal cells during mitosis (see Satterwhite and Pollard, 1992). Because contractile ring structures were observed during the cellularization of the blastoderm with Rh-phalloidin (Warn and Magrath, 1983; Warn and Robert-Nicoud, 1990), it is likely that such bands of aligned microfilaments do not exist during the syncytial mitoses. If contractile ring-like structures are absent in the syncytial blastoderm we can assume that different mechanisms are responsible for surface cap cleavage and daughter nuclei separation during the syncytial mitoses. As already observed (Warn et al., 1984), the cortical actin filaments lining the plasma membrane infoldings and the surface microprojections are the main candidates for the surface cap modifications. The complex network of filament bundles filling the whole periplasm, under the plasma membrane, together with myosin(Young et al., 1991) and actin-associated proteins (Pesacreta et al., 1989; Miller et al., 1989), may be the structural basis of the tension leading to daughter nuclei segregation at late telophase.

Our observations of a dense network of actin filaments enveloping the syncytial nuclei is consistent with the finding that the lateral proliferation and regular pattern of the nuclei in the syncytial blastodermel periplasm are sensitive to cytochalasin (Zalokar and Erk, 1976; Foe and Alberts, 1983; Edgar et al., 1987). Taken together the confocal data from Ceratitis and the results from drug-treated Drosophila embryos suggest that microfilaments may be involved in the maintenance of normal cortical organization and nuclear positioning. This agrees with the report that in Drosophila mutants in which the pattern of microfilament distribution is affected, the distance between neighbouring nuclei is shorter than that in the wild-type embryo (Hatanaka and Okada, 1991). As recently suggested, the subcortical actinbased cytoskeleton may be a likely candidate for localizing cytoplasmic information, which determines the future embryonic axes (Bearer, 1991).

Sparse actin aggregates were also found in the cortex of the precellular Drosophila embryo, lining the edges of the caps, in the region between them (Warn and Magrath, 1983; Warn, 1986; Karr and Alberts, 1986), and during the process of cellularization (Warn and Robert-Nicoud, 1990). Since during mitosis the Ceratitis embryo appeared to segregate the cytoplasmic aggregates into clusters close to the spindle regions, it is possible that this segregation is mediated by microtubules. Whether a functional relationship exists between aster microtubules and F-actin aggregate dynamics remains to be established. It is interesting to note that double labelling for actin and tubulin suggests the spatial proximity of aster microtubules and actin aggregates. Moreover, actin aggregates form shell-like clusters around the spindle regions. Because low temperature has been reported, in Drosophila embryos (Callaini and Marchini, 1989), to cause the depolymerization of aster microtubules, which regrow from sparse foci after a short recovery at 24°C, an altered distribution of the actin aggregates was to be expected in cold-treated Ceratitis embryos. Actually, actin aggregates form irregular clusters in this conditions, supporting the hypothesis that the fate of F-actin aggregates is correlated to the behaviour of the aster microtubules. Additional data suggested that the centrosome plays a role in the organization of the actin cytoskeleton in the cortex of the Drosophila embryo. The actin cap above each nucleus was densest over the centrosome (Karr and Alberts, 1986), and centrosomes were able to direct the formation of pole cells in the absence of nuclei (Raff and Glover, 1989). Double-staining experiments suggest that the position of myosin caps is also determined by centrosomes (Young et al., 1991). The spatial relationship between microtubules and microfilaments suggests that the ability of the centrosome to organize actin is mediated by the microtubules that it nucleates. When Drosophila embryos were microinjected with anti-tubulin antibodies, the cortical microtubules were locally inactivated and the formation and maintenance of actin caps was affected (Warn et al., 1987).

The nature and function of these aggregates present an intriguing question. Phalloidin has a much higher affinity for F-actin than for G-actin (Wieland, 1977), so the aggregates may be mostly composed of microfilaments. Structures corresponding to aggregates of actin filaments have not been detected by transmission electron microscopy. This may be due to the difficulty of specimen fixation or, alternatively, the actin may be present in short polymers (Wehland and Weber, 1981), which are not detected by conventional observation. Immunogold labelling showed a discrete population of actin in the cytoplasmic granules which surround the mitotic spindles. Their positions correspond to those of cortical granules observed by light microscopy in semithin sections stained with toluidine blue. It is possible that the formation of these structures is dependent on the presence of actin-binding proteins. Because the accumulation of these aggregates during the early stages of embryogenesis seems to follow distinct patterns coupled to the nuclear cycle, these structures presumably play a role during mitotic division. Some observations in the early Drosophila embryo indicate that the sparse cortical aggregates may be interpreted as storage forms of actin, which is gradually recruited during the mitotic cycle (Warn and Robert-Nicoud, 1990). Moreover, the finding that the aggregates were depleted when F-actin was polymerized in Rh-phalloidin-microinjected Drosophila embryos confirms this storage role (Planques et al., 1991).

The co-localization of a discrete population of actin in the centrosomal region was of further interest. This is not surprising because actin is consistently present in centrosome preparations in significant amounts (Bornens et al., 1987; Komesli et al., 1989). The present study, however, provides direct evidence for such a pattern of localization. The functional importance of this F-actin domain is an intriguing question. Microfilament inhibitors have been demonstrated to prevent centriole splitting in TPA-treated polymorphonuclear leukocytes (Euteneuer and Schliwa, 1985) and centrosome separation in sea-urchin eggs (Schatten et al., 1988), suggesting that actin plays a role in centrosome dynamics. Moreover, the observation that fluorescence denoting F-actin was concentrated at the poles of the spindles during metaphase and anaphase, and diffused at telophase when the centrosomal material expanded, suggests a close relationship between F-actin and centrosome behaviour. Another possibility is that actin plays a role in anchoring centrosomes to nuclei (Goldestein and Vale, 1992). The localization of actin in the region occupied by the pericentriolar process complex in the sperm of Hydractinia has been demonstrated by immunofluorescence (Kleve and Clark, 1980) and, recently, an actin-like protein, named centractin, has been discovered in association with the centrosomes of vertebrate cells (Clark and Meyer, 1992). Whether a direct functional relationship exists between cortical F-actin and centrosome dynamics remains to be established. The finding of a discrete amount of actin localized at the poles of the mitotic apparatus in close continuity with the peripheral network of microfilaments may constitute the morphological basis of this interaction.

We are grateful to Dr. M. Suffness for providing taxol. We thank Prof. G. Gabbiani for comments on the manuscript. This work was supported by a grant from the MURST to R. Dallai.