Oocyte growth, follicle cell differentiation and vitellin processing in the stick insect, Carausius morosus br. (Phasmatodea)

l~lt. J. Insect Morphol. ~ Embryol., Vol. 22. Nos 2-4, pp. 271 293, 1993 Printed in Great Britain

0020-7322/93 $6.00+ .00 © 1993 Pergamon Press Ltd

OOCYTE GROWTH, FOLLICLE CELL DIFFERENTIATION A N D V I T E L L I N P R O C E S S I N G IN T H E S T I C K I N S E C T , CARAUSIUS MOROSUS BR. ( P H A S M A T O D E A )

FRANCO GIORGI,*~" ANTONELLA CECCHETTINI,* PAOLO LUCCHESI* a n d MASSIMO MAZZINI~ *Department of Biomedicine, University of Pisa; SDepartment of Environmental Sciences, Tuscia University, Viterbo, Italy

Abstract -- Ovarian growth in stick insects (Phasmatodea) was examined ultrastructurally and cytochemically with a view to studying: (1) the kinetics of oocyte growth and the staging characteristics of ovarian follicles undergoing vitellogenesis; (2) the endocytic capability of the growing oocyte, including the post-endocytic fate of the vitellins sequestered by the oocyte during vitellogenesis; (3) the differentiation of the follicular epithelium in relation to the appearance of intercellular spaces and the extracellular release of a follicle cell product. These structural observations were interpreted in relation to the nature and kinetics of the vitellin processing in follicles undergoing vitellogenesis. Index descriptors (in addition to those in title): Ovarian development, vitellogenesis, electron microscopy.

INTRODUCTION OVER THE years, the study of ovarian growth in insects has provided biologists with a number of reliable model systems to understand reproduction and cell differentiation. Recall here that clathrin-coated vesicles were first visualized in mosquito oocytes by electron microscopy and recognized as the cell organelles for ligand internalization by endocytosis (Roth and Porter, 1964). The immunological relatedness of egg yolk proteins with several antigens of the female haemolymph, and in particular, the extraovarian origin of the major yolk precursor, now referred to as vitellogenin, were recognized as early as 1954 through the pioneering work of Telfer (1954). Since then, vitellogenin has been recognized as the major, though by no means the only (Telfer and Pan, 1988), haemolymph ligand to be internalized into the oocyte through a process of receptor-mediated endocytosis (Roth and Woods, 1982). Hitherto, all major aspects of insect ovarian growth, from synthesis of vitellogenin in the female fat body (Wyatt, 1980; Locke, 1982; Raikhel and Lea, 1983; Giorgi et al., 1989) to the receptor-mediated endocytosis of vitellogenin into the growing oocyte, have been investigated both ultrastructurally (Raikhel and Lea, 1986; Giorgi and Mazzini, 1987; Raikhel, 1987) and biochemically (Koenig et al., 1988; Koller et al., 1989; Ferenz, 1990; Dhadialla and Raikhel, 1991) in numerous insect species. Whenever possible,

-tAuthor to whom correspondence should be addressed. 271

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molecular biologists have also provided additional clues to the regulation of gene expression leading to the sex-specific and hormone-regulated synthesis of vitellogenin in the adult female fat body (Belote et a l . , 1985; Bownes, 1989; Logan and Wensink, 1990). Yolk in insects comprises of a family of evolutionary related proteins coded for by highly conserved genes. In insects, these genes are currently interpreted as having evolved from a c o m m o n ancestral gene by either gene duplication or deletion (Harnish and White, 1982). The evolutionary relatedness of vitellogenin genes in insects points to the existence of essential functions for these proteins, due perhaps to the domains being recognized as binding sites by specific receptors for either cell uptake or sorting (Yan and Postlethwait, 1990). These domains had to be maintained during evolution if vitellins were to play their role in embryogenesis. For instance, certain vitellin domains may be needed to sustain both secretion from the fat body and uptake into the oocyte by receptor-mediated endocytosis (Gochoco et a l . , 1988). Once internalized into the oocyte, insect vitellins also need to be properly delivered to the oocyte's storing c o m p a r t m e n t to avoid degradation by lysosomal intervention (Wall and Patel, 1987a, b). Additional roles of specific vitellin domains may be played during embryonic development (Kunkel and Nordin, 1985). Recent evidence from sea urchin embryos suggests that during embryonic development, yolk may provide some intermediate vitellin products capable of affecting embryo morphogenesis (Scott and Lennarz, 1989). These so-called toposomes were shown to originate by partial proteolysis of yolk polypeptides and to become part of the plasma m e m b r a n e of early blastomeres (Noll et a l . , 1985). The study of ovarian growth and differentiation in insects may provide new insights as to how vitellin gene products are first stored in the oocyte and then utilized by the growing embryo. Sorting of different gene products within the growing oocyte should be looked upon as a possible way to trigger differentiation by either activating and/or releasing vitellin products in the egg at the most appropriate time in development. Since release of yolk intermediates during embryonic development appears to be a temporally differentiated p h e n o m e n o n (Masetti and Giorgi, 1989), differential yolk partitioning into the oocyte could provide a structural substratum for such a process to occur. This p a p e r deals with ultrastructural and cytochemical analyses of oocyte growth in stick insects. It is our intention to examine how vitellins are processed following uptake into the oocyte, and sequestered into the endosomal compartment. Particular emphasis is focused on the role the follicular epithelium plays in synthesizing and secreting follicle specific products during vitellogenesis.

MATERIALS

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METHODS

General Carausius morosus (Br.) and Bacillus rossius (Rossi) (Insecta, Phasmatodea) were maintained in a rearing

room at 20-24°C with a natural photoperiod, and fed weekly with either fresh ivy or bramble leaves, respectively. Ovaries from both species were collected through dissection of adult egg-layingfemales. Ovarian follicles at different developmental stages were fixed for 2 hr in 5% glutaraldehyde-4% formaldehyde in 0.1 M cacodylate buffer at pH 7.2 maintained at 4°C (Karnovsky, 1965). More developmentally advanced follicles were punctured through the chorion and left overnight in the fixative. While small ovarian follicles were left intact, larger follicles were first fixed and then cut in half and each half processed individually for either TEM or SEM. Ovarian follicles were then washed overnight in the same buffer and post-fixed for an additional 2 hr in 1% osmium tetroxide in 0.1 M cacodylate buffer at pH 7.2. They were then dehydrated in a graded series of alcohols and, those destined for TEM analysis, were embedded in an Epon-Araldite mixture and left to

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polymerize at 60°C for 2 days. Thin sections were cut with an LKB Nova ultramicrotome, stained in uranyl acetate and lead citrate and examined in a 100SX Jeol electron microscope. Ovarian follicles to be analysed by SEM were critical-point dried in a Bomar apparatus equipped with liquid CO2 and then gold-coated using a sputtering unit working with an Argon atmosphere. Critical-point dried ovarian follicles were then glued onto metal stubs and eventually observed in a Philips scanning electron microscope.

Cytochemistry Egg-laying females of both C. morosus and B. rossius were injected with 10 txl of a Ringer solution containing horseradish peroxidase (Sigma, St. Louis, MO) at a concentration of 0.5 mg/ml and dissected after 4 or 16 hr. Ovarian follicles collected this way were fixed for 2 hr in 2.5% glutaraldehyde in 0.1 M cacodylate buffer at pH 7.2 and then thoroughly rinsed in the buffer. Prior to post-fixation in 1% osmium tetroxide, peroxidase activity was revealed cytochemically by treatment with diamino benzidine (DAB) in the presence of 1% H202 according to Graham and Karnovsky (1966). A few vitellogenic ovarian follicles were also fixed with osmium zinc iodide (OZI) for 8 hr at room temperature according to the procedure of Maillet (1963) as modified by Niebauer et al. (1969) and embedded in epoxy resins as specified above. Ruthenium red and lanthanum nitrate were administered to fixed ovarian follicles as previously reported (Giorgi et al., 1991). In vivo incorporatton studies Adult egg-laying females of C. morosus were injected with either 20 p,Ci of [35S]-methionine (specific activity 1000 Ci/mlvl), sodium [3SS]-sulphate (specific activity 40 Ci/mg), [3H]-leucine (specific activity 144 Ci/ mM), or [3H]-acetyl-glucosamine (specific activity 5.79 Ci/mM) (Amersham International, Amersham, Bucks) and maintained thereafter in small glass cages containing fresh ivy leaves. At appropriate intervals after injection, ovarian follicles of different stages of development were collected as described above.

Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) Ovarian follicles of different developmental stages were dissected from injected females in ice-cold phosphate buffered saline and homogenized in 50 mM Tris-HCl buffer at pH 6.8 containing 0.1% benzamidine, 0.1% PMFS and 10% glucose. The homogenates were then spun at 10,000 rpm for 10 min in an Eppendorf bench centrifuge. Aliquots of the supernatants were diluted with equal volumes of 5% [3mercaptoethanol and 5% SDS in 50 mM Tris-HC1 buffer at pH 6.8 and eventually boiled for 3 min. Samples were resolved by 5-15% polyacrylamide gel gradients using a 3-channel peristaltic pump connected to 2 mixing syringes. Electroplhoresis was carried out overnight at room temperature with 15 m A × 80 V. Acrylamide solutions were prepared as specified by Laemmli (1970) with 0.1% SDS added to the upper electrode buffer reservoir. After e]ectrophoresis, gels were stained for 1 hr in 0.1% Coomassie R in 30% acetic acid-10% methanol and eventually stored in 7% acetic acid. Destained gels were dried in a gel dryer connected to an Edwards vacuum pump through a Savant cooling trap. Western blotting After electrophoresis, gels were placed onto nitrocellulose sheets (Bio-Rad) and sandwiched between porous sponges. Electrotransfer was achieved using a Bio-Rad electrotransfer apparatus cooled at 16°C (Towbin et al., 1979). The nitrocellulose sheet was then blocked with a solution containing 5% dry milk and 0.05% Tween-20 in Dulbecco's phosphate-buffered saline (PBS). The nitrocellulose sheet was incubated for 2 hr in blocking buffer containing a mono-specific monoclonal antibody raised against the follicle cell product of C. morosus diluted 1 : 100. Nitrocellulose sheets were repeatedly rinsed with buffer (0.05% Tween-20 in PBS) and then incubated for 1 hr at room temperature with goat anti-mouse serum, alkaline phosphatase conjugated (BioRad). The nitrocellulose strips were then rinsed in wash buffer and incubated in the Bio-Rad colour developer for 10 min. The staining reaction was blocked by addition of 0.1 M HC1 to the incubation medium. Strips were air dried and photographed as soon as the staining reaction was completed, before the coloured bands faded. Fluorography Destained polyacrylamide slab gels were soaked in Amplify (Amersham International, Amersham, Bucks) for 30 min and then vacuum-dried as described above. Subsequently, gels were overlaid with Dupont X-ray films and placed in light-proof boxes at - 8 0 ° C for periods ranging from 1 to 4 weeks. The film sheets were then developed and fixed using commercially available procedures. Light microscope autoradiography of developing ovarian follicles Ovarian follicles were dissected from egg-laying females following in vivo exposure to [3H]-acetylglucosamine, [3H]-leucine or sodium-[3SS] sulphate (Amersham International, Amersham, Bucks), for time intervals ranging fi~om 4 to 16 hr. Soon after dissection they were fixed for 2 hr in 4% formaldehyde in 0.1 M phosphate buffer at pH 7.2 and thoroughly washed in the buffer. They were then dehydrated in a graded series of ethanol solutions, embedded in LKB embedding medium. Two-micrometre-thick sections were prepared

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using a Reichert microtomeand mounted on gelatin-coatedglass slides. LM-autoradiographywas carried out by dipping these slides into a melted K2 Ilford photographic emulsion and letting them dry on an ice-coldplate. Slideswere exposedfor 4 weeksin lightproofboxes maintainedat 4°C. They were then developedin a D 19 Kodak developer for 3 min, rinsed in running tap water and fixed in 24% sodium thiosulphate for 10 min. Once air-dried, they were stained with i% methylene blue-l% toluidine blue on a hot plate.

RESULTS

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DISCUSSION

Overview o f ovarian growth

Ovarian growth in stick insects is regulated so that ovarioles within the same ovary are out of phase with each other (Huebner, 1983). Usually up to 10 follicles may be lodged in each ovariole. They follow one another according to a developmental sequence in which the follicles more posteriorly positioned along the ovariole are also more advanced in development. Within the ovariole, only the terminal follicle is capable of attaining full growth, the preceding ones being blocked in pre-vitellogenesis or early vitellogenesis (Bradley et al., 1987). It has been demonstrated that ovulation of the terminal follicle in the ovariole triggers the subterminal follicle to enter full vitellogenic growth (Mesnier, 1984). Although the matter has not been further investigated, it is not unlikely that the interfollicular region, i.e. the follicle cells between succeeding follicles in the ovariole, might play a role in the process, perhaps by regulating the extent of intercellular communication through gap junctions. It has been determined that an ovarian follicle increases over 100 times in volume during the time interval between entrance into the terminal position of the ovariole and ovulation (Mesnier, 1980). Ovarian follicles in stick insects are of the panoistic type, that is, they include only the oocyte and a surrounding follicular epithelium. During ovarian development, oocyte enlargement is primarily due to engulfment and storage of extensive amounts of yolk. As the oocyte enlarges, the follicular epithelium differentiates, acquiring different structural characteristics that vary from stage to stage. These aspects will be dealt with separately in the following sections. Based on these structural characteristics, oogenesis in stick insects can be thought of as including a pre-vitellogenic period in subterminal follicles, and vitellogenic growth phase in terminal follicles. This latter occurs mainly by endocytic uptake of a yolk precursor or vitellogenin as long as haemolymph gains access to the oocyte surface through the intercellular spaces of the follicular epithelium (Teller, 1961). Closure of follicle cell patency and deposition of various egg coverings will eventually seal the oocyte from the haemolymph (Rubenstein, 1979). By then, the oocyte will be incapable of continuing endocytic uptake because of inaccessibility to vitellogenin. The period between cessation of the vitellogenic growth phase in terminal follicles and ovulation was estimated to last about 8-9 days in C. morosus (Giorgi et al., 1990). This period also includes the time spent for ovulation and oviposition. The minimum time interval an average ovarian follicle would require to complete vitellogenesis was thought to be about 30 days. This was determined by measuring the time required by vitellin, in C. morosus, to be processed in ovarian extracts exposed in vivo to [aac]-amino acids (unpublished observations). A question still to be answered experimentally in stick insects is whether inception of chorionogenesis depends upon completion of vitellogenesis. The observation that newly laid eggs are significantly smaller in allatectomized females, or even in senescent

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females, than in younger females, suggests that chorionogenesis is a temporally controlled phenomenon, and does not depend upon the ultimate volume attained by the ovarian follicle or on the amount of yolk stored in the ooeyte (Bradley et al., 1988). Ovarian development in stick insects is apparently a hormone-independent phenomenon, :for allatectomy does not prevent eggs from being laid (Pflugfelder, 1937). Under these experimental conditions, vitellogenin is still being synthesized and secreted by the female fat body. However, since allatectomized females exhibit a vitellogenin titre higher than control females, allatectomy would seem to primarily affect the endocytic apparatus of tlhe oocyte itself, rather than the fat body's capability to synthesize and secrete vitellogenin (Bradley et al., 1988), as appears to be the case in other insect species. Thus, even though vitellogenesis in C. morosus was reported to occur independently of any juvenile hormone supply (Pflugfelder, 1937), it may well be that other proteins in the ovarian follicle, contributing perhaps to the endocytic transfer of vitellogenin, are either quantitatively reduced or their synthesis suppressed in allatectomized females. Oocyte differentiation Ultrastructural features of the undifferentiated oocyte. The ultrastructure of developing oocytes along the ovariole has been extensively studied in stick insects (Mazzini and Giorgi, 1984, 1985). Vitellogenesis appears to begin in follicles occupying the subterminal position in the ovariole. Prior to this stage, all preceding follicles are pre-vitellogenic, that is, void of any yolk inclusion in both the cortical and central ooplasm (Fig. 1). Nevertheless, these oocytes are characterized by an extensive development of the microvilli along the oolemma (Fig. 5) and by numerous endocytic vesicles in the cortical ooplasm (Fig. 6). Along the regions of the plasma membranes where the oocyte microvilli interdigitate with those emerging from the follicle cells, there are frequent junctional contacts (Fig. 7). In vivo incorporation studies with horseradish peroxidase, showed that newly formed endocytic vesicles in pre-vitellogenic oocytes never convey this extracellular tracer to the yolk spheres (Figs 8, 9), but to multivesicular bodies for intracellular degradation (Mazzini and Giorgi, 1986). One of the most recurrent features of the pre-vitellogenic ooplasm in stick insects is the numerous multivesicular bodies containing an electron-dense matrix in various phases of condensation (Fig. 10). These should not be confused with forming yolk spheres of endogenous origin on the basis of their electron density, because no antivitellin binding sites could be detected immunochemically in the ooplasm at these developmental stages. Mazzini et al. (1986) interpreted these inclusions as cell sites used by the oocyte for processing aged ooplasmic organelles. Based on the kinetics of vitellin labelling in developing follicles, they concluded that most, if not all, yolk stored in vitellogenic oocytes has an heterologous origin, i.e. transferred to the oocyte following the endocytic pathway from the haemolymph (Mazzini et al., 1986). This conclusion excluded any possibility that these early ooplasmic organelles contribute to yolk deposition in stick insects. However, fusion between early formed yolk spheres and condensed multivesicular bodies were frequently seen in oocytes about to begin vitellogenesis (Fig. 13), suggesting that yolk spheres in vitellogenic follicles may be part of the oocyte endosomal compartment. Rather than contributing to the overall vitellin content of the yolk spheres, multivesicular bodies might allow membranes to be

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FIG. 1. A light microscope picture of a pre-vitellogenic ovarian follicle showing follicular epithelium (fc) with tightly adjoined cells and no intercellular spaces. Cortical ooplasm in oocyte (oo) is devoid of forming yolk spheres. × 550. Fro. 2. A light microscope picture showing a cross-section of follicular epithelium from a previtellogenic ovarian follicle. Note few follicle cells undergoing mitosis. × 550. FIG. 3. Scanning electron micrograph of follicular epithelium from a pre-vitellogenic ovarian follicle. fc = follicle cells; oo = oocyte. × 2200. Fro. 4. A low power electron micrograph of follicular epithelium from a pre-vitellogenic ovarian follicle. Some follicle ceils are undergoing mitosis (arrows). n = nuclei, x 1850.

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conveyed to the: yolk spheres and to b e c o m e associated with the nearby Golgi apparatus (Figs 11, 12), perhaps for m e m b r a n e recycling and/or intracellular digestion. Y o l k deposition in vitellogenic follicles. Vitellogenesis in stick insects starts in follicles occupying the terminal position in the ovariole. However, for reasons not yet fully understood, subterminal follicles only grow in volume to a limited extent (Giorgi and Mazzini, 1984). It is with the attainment of the terminal position in the ovariole that the oocyte undergoes major structural changes, leading to a fully grown oocyte. The beginning of vitellogenesis in terminal follicles is m a r k e d by the appearance of large coated pits and vesicles in the cortical ooplasm and a progressive extension of the o o l e m m a microvilli (Fig. 14). The latter interdigitate with similar projections emerging from the apica]: end of the overlying follicle cells. In vitellogenic follicles of Bacillus rossius, some of these follicle cell projections were seen to protrude deeply into the cortical ooplasm, giving rise to specific junctional contacts at the very end of the invagination (IVlazzini and Giorgi, 1985; Giorgi and Mazzini, 1985). Because no similar studies have been carried out with other species, it is difficult to envisage a functional significance for the structural relationship between the follicle cells and the oocyte upon onset of vitellogenesis. It is possible that they might provide a way for the oocyte and the follicle cells to coordinate cell differentiation. O o l e m m a microvilli are somehow e m b e d d e d in a densely fibrous matrix that permeates all intercellular spaces facing the oocyte (Fig. 14). Such a matrix can be clearly shown to be highly glycosylated, as demonstrated by specific staining with ruthenium red (Fig. 15). U n d e r these conditions, the glycosylated matrix bound to the o o l e m m a microvilli can be seen to be made of regularly spaced subunits. The use of this test does not determine whether the fibrous matrix is either a constitutive c o m p o n e n t of the o o l e m m a glycocalyx or the vitellogenin itself is bound to the oocyte surface. The current consensus is that coated pits will transform into coated vesicles, by detachment from the o o l e m m a (Roth and Porter, 1964; Pastan and Willingham, 1983), and later on into uncoated vesicles (Rhaikhel and Lea, 1986; Tsuruhara et al., 1990; Richter and Bauer, 1990). The use of various extracellular tracers in a n u m b e r of insect species, including stick insects (Mazzini and Giorgi, 1986), has clearly shown that the endocytic pathway, leading to the forming yolk spheres, entails labelling of both coated and uncoated vesicles (Figs 16, 17). This suggests that most of the newly formed endocytic vesicles lose their clathrate coat prior to conveying their highly dense contents to the yolk spheres. It has been observed repeatedly that newly formed endocytic vesicles in the cortical

FIG. 5. Scanning electron microscope view of follicle cell/oocyte interface from a pre-vitellogenic ovarian follicle of Bacillus rossius. Note presence of numerous interdigitating microvilli, fc = follicle cells; oo = oocyte x 11,100. FIG. 6. A high power micrograph of follicle cell/oocyte interface showing numerous microvilli and coated pits (cp) and vesicles (cv) in cortical ooplasm, x 42,000. FIG. 7. Follicle cell/oocyte interface from a pre-vitellogenic follicle of Bacillus rossius impregnated with lanthanum nitrate showing interdigitating cell microvilli. Several junctional complexes (arrows) are visualized in the region of contact, x 81,400. FIGS 8, 9. Cortical ooplasm of a pre-vitellogenicovarian follicle from Bacillus rossius exposed in vivo to horseradish peroxidase. Label has gained access to small coated pits (cp) and vesicles (cv). mv = microvilli, x 33,700.

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FIG. 10. A low power electron micrograph of central ooplasm of a pre-vitellogenic ovarian follicle from Bacillus rossius showing a n u m b e r of multivesicular bodies (mvb) in various phases of condensation. × 15,600. F1G. 11. Electron micrograph of central cytoplasm of a pre-vitellogenic ovarian follicle from Bacillus rossius showing a Golgi apparatus with several dilated cisternae and n u m e r o u s vesicles nearby. × 29,900. FlG. 12. Freeze-fracture replica of a similar cytoplasm region in pre-vitellogenic ovarian follicle. × 32,850. F1G. 13. A low power micrograph of central ooplasm of an early vitellogenic ovarian follicle from Bacillus rossius showing multivesicular bodies (mvb) and forming yolk spheres (y). × 29,200. FI~. 14. O o l e m m a microvilli of a vitellogenic ovarian follicle. Note presence of a fibrous coat extending from plasma m e m b r a n e , x 42,300. FIG. 15. O o l e m m a microvilli of a vitellogenic ovarian follicle as visualized after glutaraldehyde fixation in presence of ruthenium red. Arrows point to a regular spacing along cell coat. x 36,500.

Ovarian Growth in Stick Insects

F~c. 16. Low power micrograph of cortical ooplasm from a vitellogenic ovarian follicle of Bacillus rossius showing several coated pits (cp) and vesicles (cv) along with other endocytic vesicles. x 19,200. FIG. 17. Enlargement of cortical ooplasm depicted in previous picture to show membrane-bound (arrows) and core-filled endocytic vesicles (double arrows). × 48,100. F~G. 18. A vitellogenic ovarian follicle showing extent of vesicular labelling in cortical ooplasm after 8 hr fixation with osmium zinc iodide, mv = microvilli; y = yolk spheres. × 14,800. Flo. 19. Follicle cells from a vitellogenic ovarian follicle showing extent of Golgi apparatus (G) labelling after 8 hr fixation with osmium zinc iodide, x 16,650. FIGs 20, 21. Forming yolk spheres from cortical ooplasm (Fig. 20) and from deeper regions of central ooplasm (Fig. 21). Yolk spheres (Y) and Golgi apparatus (G) are closely associated. × 19,200 (Fig. 20); x 17,200 (Fig. 21).

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ooplasm are either membrane-bound or core-filled vesicles (Bassemir, 1977). While some electron-dense material adheres to the limiting membrane in the former, it is densely packed within the vesicle lumen in the latter (Fig. 17). Interestingly, the transformation of one type of vesicle into the other seems to be an essential prerequisite for the membrane-bound vitellin to be displaced to the forming yolk spheres. Not all vesicles lying close to, or surrounding, the newly formed yolk spheres in the cortical ooplasm are endocytic in origin. Ovarian follicles fixed with OZI have most of the vesicles in the cortical ooplasm, including a few scattered Golgi, labelled with electron-dense OZI deposits (Fig. 18). By comparison, the cytoplasm of the overlying follicular epithelium is void of any OZI deposit, notwithstanding the Golgi apparatus itself (Fig. 19). In early vitellogenic oocytes, the forming yolk spheres of the cortical ooplasm bear a close structural relationship with the Golgi apparatus (Fig. 20). A few OZI-positive deposits are also included within the superficial layer enclosing the forming yolk spheres. As yolk deposition proceeds, many more OZI-positive vesicles appear associated with the forming yolk spheres (Fig. 21), the rest of the cortical ooplasm also remaining heavily labelled with endogenously derived vesicles. These observations are consistent with earlier reports of oocytes of other species (Giorgi et al., 1976) showing that both the Golgi apparatus and the nearby vesicles can be labelled specifically in the cortical ooplasm of vitellogenic follicles fixed with OZI. Because of the known specificity of the OZI fixative (Niebauer et al., 1969), all vesicles lying close to the forming yolk spheres can be reliably interpreted as being contributed to by the Golgi apparatus and on their way to fusing with the superficial layer of the yolk spheres. One of the peculiarities of vitellogenic oocytes in stick insects is that yolk spheres in the cortical ooplasm undergo an apparent repetitive fusion until a common yolk compartment is formed in the central ooplasm. At present, it is not known why yolk spheres stop fusing upon reaching a certain size in some insect species, while in others, as in stick insects, fusion results in the actual disappearance of isolated yolk spheres. Whatever the mechanisms(s) controlling such a process, there are various consequences that are worth mentioning here. Our ultrastructural observations show that vitellin storage in stick insects does not entail any form of crystallization, as it has been previously reported for such species as A e d e s aegypti in insects (Roth and Porter, 1964), or X e n o p u s laevis in amphibians (Ward, 1978). This may result from the fact that yolk spheres in stick insects are not dehydrated as in other species, the yolk being stored in the ooplasm as a fluid. Perhaps the oocyte volume in stick insects does not constitute a limiting factor for yolk storage and the surface/volume ratio of the yolk compartment need not to be as high as that in oocytes with isolated yolk spheres to sustain water exchange. Both these conditions may perhaps be functionally related to the high solubility of vitellins in these species. Another noteworthy consequence of the repetitive fusion of yolk spheres in stick insects is that the entire ooplasm in fully grown oocytes will eventually become enclosed by 2 membranes lying very close to one another. One is the oolemma itself, and the other the limiting membrane contributed to by the fusing yolk spheres. Only a narrow fringe of cytoplasm intervenes between the 2 cell membranes. Yolk deposition, through repetitive fusion of newly formed yolk spheres, can be clearly followed by light microscope autoradiography following in vivo exposure to [3H]glucosamine (Figs 22-24). In early ovarian follicles exposed to [3H]-glucosamine for 4 hr, label appeared unevenly distributed in the yolk spheres of the cortical ooplasm (Figs 25, 26). In more developmentally advanced follicles, the common yolk fluid ooplasm has

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F~Gs 22-24. Light microscope autoradiographs showing vitellogenic ovarian follicles at different development :stages exposed to [3H]-glucosamine for 4, 12 and 16 hr, respectively, fc = follicle cells; oo = oocyte, x 150. FiGs 25-27. Light microscope autoradiographs showing follicle cell/oocyte interface of vitellogenic ovarian follicles exposed to [~H]-glucosamine for 4, 12 and 16 hr, respectively. Note that labelled yolk spheres in cortical ooplasm become progressively larger with development and fuse into a common yolk ooplasm, fc = follicle cells; oo = oocyte, x 600 (Figs 25, 26); x 900 (Fig. 27). FIG. 28. Light microscope autoradiograph showing a chorionogenic ovarian follicle labelled in vivo with [3H]-glucosamine for 4 hr. At this developmental stage, cortical yolk granules are labelled, while cortical ooplasm is unlabelled, fc = follicle cells; oo = oocyte, x 1000. FIGS 29, 30. Light microscope autoradiographs showing follicle cell/oocyte interface from an ovarian follicle exposed to [3H]-glucosamine for 4 hr. Note that label is restricted to cortical yolk granules (arrows) underneath endochorion and in cortical ooplasm (arrows). × 1000. FIG. 31. Low power electron micrograph of cortical ooplasm from a chorionogenic ovarian follicle showing a newly formed cortical yolk granule. Presence of translucent and electron-dense vesicles makes this granule look like a multivesicular body. Ve = vitelline envelope; y = cortical ooplasm, x 2600. FIG. 32. A scanning electron micrograph of a chorionogenic ovarian follicle showing inner endochorion (ien) with some cortical yolk granules still attached. Note presence of numerous holes along granule surface (arrows). x 1800.

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eventually extended to include all residual yolk spheres of the cortical ooplasm. When these follicles were exposed to [3H]-glucosamine for up to 16 hr, label appeared to be uniformly dispersed within the common yolk compartment (Fig. 27). Thus, it seems that fusion of yolk spheres is a continuous process that ends only with termination of vitellogenesis and transition to the chorionogenic phase that follows. These studies, along with earlier findings by other investigators, have confirmed the generally accepted view that insect oocytes undergo yolk deposition by receptor-mediated endocytosis of a yolk precursor -- or vitellogenin -- from the haemolymph. This rules out any possibility that yolk vitellins in stick insects are synthesized endogenously by the oocyte itself. This conclusion does not necessarily imply that all proteins to be stored in the yolk compartments of the oocyte are contributed, through endocytic uptake, from the haemolymph. As will be discussed later, follicle cells in stick insects, as in many other insect species, are endowed with the capability of synthesizing and releasing a major follicle cell product that is eventually transferred endocytically to the growing oocyte during vitellogenesis. Multivesicular bodies in the cortical ooplasm o f chorionogenic follicles. Yolk deposition in the cortical ooplasm of stick insects does not end with the onset of chorion formation. By the time the inner endochorion is being deposited along the oocyte/follicle cell interface, yolk spheres in the cortical ooplasm have already fused together to form a common fluid compartment. However, there is as yet another late burst of yolk deposition in the cortical ooplasm. This takes place in follicles in which the vitelline envelope has already been completed, and the inner endochorion is actually being deposited (Fig. 28). Following in vivo exposure to [3H]-glucosamine for 4 hr, a number of yolk granules in the cortical ooplasm become labelled, even though the rest of the ooplasm is completely fused and the intercellular spaces of the overlying follicular epithelium have already collapsed. Figures 29 and 30 show a chorionogenic ovarian follicle in which the follicular epithelium and the underlying cortical ooplasm have been widely separated by fixation. Under these conditions, [3H]-glucosamine-labelled cortical yolk granules can be seen associated with both the inner endochorion (Fig. 29) and the ooplasm (Fig. 30). These cortical yolk granules have been formed following the completion of vitellogenin uptake from the haemolymph and fusion of all newly formed yolk spheres. To find out how these cortical granules come to be formed in late oocytes, choriogenic follicles were examined ultrastructurally. By transmission electron microscopy, the granules appear to consist of multivesicular bodies enclosing a number of vesicles, some of which have an electron-dense content (Fig. 31). Interestingly, the granules lay in the region between the fringe of cytoplasm underneath the oolemma and the limiting membrane of the common yolk compartment. This location suggests that the cortical granules may be formed by a process not involving endocytic uptake through small vesicles, but rather bulk engulfment through a process resembling phagocytosis. Using the scanning electron microscope, the cortical yolk granules can be seen as roundish bodies lying close to the inner endochorion and superficially pierced by many holes (Fig. 32). Examination of newly laid eggs by both scanning and transmission electron microscope revealed that these granules persist in the cortical ooplasm throughout oogenesis and the initial stages of embryonic development (unpublished observations). It is not unlikely that the cortical granules might contain products other than the vitellin

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polypeptides sequestered by the oocyte at earlier stages. As will be clarified later in this paper, the folliicle cells may be the most likely candidate to synthesize material eventually sequestered in these cortical granules.

Follicle cell differentiation Uhrastructural features of undifferentiated follicle cells. Pre-vitellogenic oocytes are enclosed by a multilayered follicular epithelium characterized by tightly adjoined cells (Figs 2-4). Follicle cells are columnar cells with an apical end protruding towards the oocyte surface through long microvilli, and a basal end closely apposed to the basal lamina. At these early developmental stages, follicle cells are typically undifferentiated cells, most of the cell volume being occupied by a huge elongated nucleus, containing numerous clumps of heterochromatin material and 2 or more nucleoli (Fig. 4). The cytoplasm forms a narrow fringe around the nucleus with very few cell organelles. Adjacent cells are joined to one another through gap junctions of macular type, which are apparently randomly dispersed over the cell surface (Mazzini and Giorgi, 1985). Follicle cell differentiation during the vitellogenic growth phase. As ovarian follicles enter the terminal position in the ovariole, the follicular epithelium undergoes major structural modifications which invest both the overall cell shape and the organelle content of the cytoplasm (Figs 33-36). With the onset of rapid vitellogenic growth in terminal ovarian follicles, follicles cells become separated by wide intercellular spaces (Fig. 37). When examined by scanning electron microscopy, these intercellular spaces appear filled with fibrous material that extends from the apical to the basal end of the follicular epithelium. The origin of this material can be traced back to the secretory activity of the follicle cells in ovarian follicles that are about to undergo rapid vitellogenic growth. Initially, such material prevails towards the apical end of the follicle cells (Fig. 38). Cells not yet fully covered by this fibrous material are characterized by a number of holes along their basal end (Fig. 39). As vitellogenesis proceeds to completion, the material becomes widespread all over the follicle cell surface (Figs 40, 43), so as to hide any structural detail of the follicle cell surface. Follicle cells are polarized cells with a branched bas~tl end apposed onto the basal lamina (Fig. 41) and a roundish apical end deeply protruding towards the cortical ooplasm (Fig. 42). An extensive structural analysis of vitellogenic ovarian follicles has shown that the follicle cell cytoplasm is characterized by the presence of membrane-bounded granules. These appear to enclose material structurally similar to the one present in the intercellular spaces of the follicular epithelium (Figs 44, 45). Instances of actual continuities between the membrane enclosing these granules and the follicle cell plasma membrane, have also been encountered with a certain frequency. In addition, Golgi apparatus lying close to electron-translucent granules have also been recorded. These data suggest that, at least ultrastructurally, follicle cells in C. morosus possess all cell organelles apt to sustain a secretory activity during the vitellogenic growth phase. The identity of the material released by the follicle cells in C. morosus is discussed below. In spite of the formation of wide intercellular spaces, adjacent follicle cells remain joined to one another through extensive gap junctions. Both lanthanum impregnation (Fig. 46) and freeze-fracture analysis (Fig. 47) of vitellogenic ovarian follicles show that follicle cells are extensively joined through junctional complexes.

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36

FIGS 33-36. Light microscope pictures showing follicular epithelium of ovarian follicles staged as early vitellogenic (Fig. 33), mid-vitellogenic (Fig. 34), late vitellogenic (Fig. 35), and chorionogenic (Fig. 36) follicles. × 600. FIG. 37. Light microscope picture of follicular epithelium from a vitellogenic ovarian follicle. Note presence of wide intercellular spaces between adjacent follicle cells. × 600. F1G. 38. Scanning electron microscope view of follicular epithelium from an early vitellogenic ovarian follicle, × 3300.

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Perhaps their function is to coordinate follicle cell differentiation and to allow the follicular epitlaelium to enlarge to keep pace with the increasing oocyte volume. As long as intercellular spaces in the follicular epithelium are open, gap junctions are restricted to the follicle cell projections that extend through the intercellular spaces. However, when yolk deposition ceases, adjacent follicle cells once again become joined through extensive gap and septate junctions (Figs 48-50). Septate junctions are characterized by undulated strings of particles tightly packed with one another. Rows of particles can also be clearly visualized at the border between 3 closely apposed cells (Fig. 49). In between adjacent septate junctions, are still clusters of intramembrane particles forming gap junctions of rnacular type (Fig. 50). By the time intercellular spaces in the follicular epithelium are no longer open, the vitelline envelope and the inner endochorin are also being deposited onto the oocyte surface. The temporal coincidence of these 2 events makes it difficult to establish whether cessation of vitellogenin uptake is primarily caused by closure of the intercellular spaces in the follicular epithelium, as suggested by Rubenstein (1979), or by the formation of the egg coverings.

The secretory activity of the follicle cells The onset of vitellogenesis in terminal follicles coincides with the appearance of some fibrous material in the intercellular spaces of the follicular epithelium. This could either be haemolymph proteins trapped in the intercellular spaces on their way to the oocyte surface, or follicle cell-produced proteins released extracellularly, or both. However, material struc~turally similar to the one occupying the intercellular spaces of the follicular epithelium can also be detected within vesicles of the follicle cell cytoplasm (Fig. 45). Horseradish peroxidase administered in vivo to egg-laying females, does not gain access to these vesicles, suggesting that these are exocytic rather than endocytic vesicles (data not shown). These observations, along with the above structural analysis, suggest that the follicle cells are endowed with the capability of synthesizing and secreting a fibrous material into the intercellular spaces. To explore this possibility, ovarian follicles at different developmental stages were exposed, either in vivo or in vitro, to various radioactive precursors and the radioactivity incorporated under these conditions was revealed both fluorographically on polyacrylamide gels and autoradiographically on semi-thick sections.

FiG. 39. Enlargement of previous figure to show a n u m b e r of holes (arrows) along basal end of follicle cell surface. × 11,100. FIG. 40. Scanning electron microscope view of follicular epithelium from a mid-vitellogenic ovarian follicle, bl = basal lamina, x 2700. FIGS 41, 42. Scanning electron microscope view of basal (Fig. 41) and apical (Fig. 42) ends of follicular epithelium in a mid-vitellogenic ovarian follicle, x 4700. FlG. 43. Scanning electron micrograph of follicular epithelium from a mid-vitellogenic ovarian follicle showing a fibrous material in follicle cell interspaces, bl = basal lamina, z 6600. FXG. 44. Low power electron micrograph showing follicle cells (fc) from a vitellogenic ovarian follicle. Note that most of cell volume is m a d e up of nucleus (n), cytoplasm forming a narrow peripheral fringe, oo = oocyte, x 1800. FIr. 45. Enlargement of follicle cell surface from a vitellogenic ovarian follicle to show exocytotic vesicles close to apical plasma m e m b r a n e . They contain fibrous material similar in electron density to that present extracellularly. × 29,600.

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£"

.

FIG. 46. High resolution micrograph of a lanthanum impregnated pre-vitellogenic ovarian follicle from Bacillus rossius showing an extended portion of follicle cell plasma membrane joined by gap junctions, x 133,200. FIG. 47. A platinum replica of a freeze-fractured ovarian follicle from Bacillus rossius showing several gap junctions along plasma membranes joining adjacent follicle cells, x 53,500. FIG. 48. High resolution micrograph of a lanthanum impregnated chorionogenic ovarian follicle from Bacillus rossius showing a portion of follicle cell plasma membrane joined by septate junctions, x 114,000• FIGS 49, 50. Replicas of a freeze-fractured ovarian follicle showing cell plasma membranes joined by septate junctions. Gap junctions (gj) of macular types are also visible. × 27,500 (Fig. 49). x 31,000 (Fig. 50).

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[ 35S]-methionine

287

[ 35S]-sulphate

a

hemolymph/follicles

ovarian follicles

1.1 2.2 3.3 4.4

6 12 24 72

aemolymph/follicles He 0.8 1.5

2.0 b

B1

A1

..,~ 116 ~97

A2 ~ ....

A3

,

,~

-

=

=Wm

~

66

--,---45

51 FIG. 51. Fluorograph of haemolymph and several vitellogenic ovarian follicles exposed in vivo to [35S]-methionine for time intervals ranging from 1 to 4 hr (left panel) and for 6, 12, 24 and 72 hr (central pane]) as resolved by SDS-PAGE on 5-15% gradients. Right panel depicts a sequence of developing ovarian follicles exposed in vivo to sodium [35S]-sulphate for 24 hr. a = hr; b = ram.

In vivo incorporation studies. Vitellins were originally identified in stick insects by electrophoretic analysis of developing ovarian follicles (Giorgi and Macchi, 1980). Following short time exposures to [35S]-methionine, these female-specific polypeptides were shown to become, first labelled in the haemolymph, and later on in the ooplasm. Since then vitellins in stick insects have been purified (Giorgi et al., 1982), and extensively characterized during synthesis in the fat body (Giorgi et al., 1989) and limited proteolysis in the developing embryo (Masetti and Giorgi, 1989). Data available to date indicate that stick insects are characterized by 2 multimeric, glycosylated vitellins, referred to as vitellins A and B, both of which are synthesized by the female fat body in a hormone-independent manner (Bradley et al., 1988). In order to identify the cell products released by the follicular epithelium, developing ovarian follicles and haemolymph were pulse-chased in vivo with [35S]-methionine for time intervals ranging from i to 72 hr. Under these conditions, haemolymph was labelled within an hour after injection, whereas ovarian follicles were labelled with a minimum time interval o1! about 4 hr (Fig. 53). The protein fraction most heavily labelled in the ovarian follicle was the one indicated as B1, polypeptides A1 and A 2 requiring longer exposure times to become labelled. This labelling pattern in stick insect follicles is due to a differential processing of the vitellin polypeptides during endocytic transfer into the oocyte and subsequent sequestration into the endosomal compartment. It should be noted, however, that a 75 kDa follicle polypeptide, with no counterpart in the

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800 u

600

"~,.~400

[]

6h

[]

12h 24h

[]

u 200

0 0.8

1.5

2.0

2.5 *

follicle size (ram) (*) chorionated follicles FIG. 52. Histogram showing. 3~extent of radioactivity, incorporated in vitellogemc ovarian follicles after in vivo exposure to sodium [ -S]-sulphate. ( ) , chorionated follicles exposed to tracer for 6 and 24 hr.

5:3

54 F]Gs 53, 54. Light microscope autoradiographs showing follicular epithelium of vitellogenic ovarian follicles of different developmental stages exposed in vivo to sodium [35S]-sulphate for 24 hr. fc = follicle cells; oo = oocyte, x 1780.

haemolymph, becomes labelled after a minimum time exposure of 4 hr to [35S]methionine (Fig. 51). Prolonging the in vivo exposure from 6 to 72 hr caused the 75 kDa polypeptide fraction to become progressively more labelled along with the other vitellin polypeptides (Fig. 51). Because this polypeptide is synthesized in an ovarian autonomous manner, it is likely to represent and endogenous contribution by the ovarian follicle and perhaps be restricted to the follicle cells. To pursue this hypothesis further we studied labelling of the 75 kDa polypeptide using various radioactive tracers and attempted to establish its location within the follicle

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5

|

o o

2

4

6

8

10

time (h)

12

14

F~G. 55. Extent of radioactivity incorporated into vitellogenic ovarian follicles exposed in vitro to [35S]-methionine for time intervals ranging from 1 to 12 hr.

14

8 o

6

4

0

1

2

follicle length (ram)

3

FIG. 56. Extent of radioactivity incorporated into vitellogenic ovarian follicles of different developmental stages exposed in vitro to [35S]-methionine for 12 hr.

cell epithelium. Ovarian follicles exposed in vivo to sodium [35S]-sulphate became labelled with kinetics that depended primarily on the stage of development rather than on the exposure time (Fig. 52). In sodium [35S]-sulphate labelled follicles examined by SDS-PAGE, radioactivity was, however, exclusively associated with the 75 kDa polypeptide (Fig. 51). Light microscope autoradiography of ovarian follicles exposed in vivo to sodium [35S]-sulphate showed that label appeared confined to the intercellular spaces of the follicular epithelium (Figs 53, 54). In vivo exposure to [3H]-glucosamine resulted in a more complex labelling pattern, since vitellin polypeptides in stick insects are also glycosylated.

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STAINED PROTEINS

[35S]-METHIONINE

ovarian follicles 1.6 (ram)

ovarian follicles 2

4

6

8

12 a

IMMUNOBLOT

ovarian follicles 1.2 1.4 1.6 1.8 b

B~

i ¸ :i!! ~ ~116

A1

~-- 97

-&2

M

B2

iil ;iii¸

A3

.,,.__ 6 6 -,,-- 4 5

57 Fro. 57. Vitellogenic ovarian follicles exposed in vitro to [35S]-methionine for time intervals ranging from 1 to 12 hr and resolved by SDS-PAGE on 5-15% gradients. Right panel shows a Western blot of ovarian follicles reacting with a monoclonal antibody (Mab 4G12) specific for 75 kDa polypeptide.

In vitro incorporation studies. In in vitro cultured ovarian follicles, the incorporation of [35S]-methionine increases linearly for over a period of 12 hr (Fig. 55). The rate of incorporation relates exponentially to the follicle size, suggesting that protein synthesis gradually declines in follicles that are about to complete chorionogenesis (Fig. 56). Although the polypeptides labelled under these conditions are released into the culture medium, none corresponded in molecular weight to the 75 kDa polypeptide identified in vivo (Fig. 57). A plausible explanation to interpret this observation would be to assume that synthesis of this follicle cell-produced polypeptide is a hormone dependent process. Alternatively the newly synthesized follicle product could be highly susceptible to proteolytic degradation in in vitro cultured conditions. As a final approach to establish the identity of the follicle cell product, a monoclonai antibody was raised against the 75 kDa glycoprotein. The blot shown in Fig. 57 demonstrates the specificity of the Mab 4G12 produced against this polypeptide. Interestingly, the polypeptide is present as a 75 kDa doublet at all follicle developmental stages including the beginning of vitellogenesis. Mab 4G12 is also detecting a protein fraction of higher molecular weight, but its origin and role remain to be established (Fig. 57). The 4G12 antibody was not reactive with the follicular epithelium when tested by immunostaining on cryo-embedded sections of vitellogenic ovarian follicles. By contrast, antibodies against either vitellin A or B polypeptides used as controls, were shown to stain preferentially newly formed yolk spheres in the cortical ooplasm. The reason for this apparent failure is unclear, but could be caused by the denatured condition of the 75 kDa antigen used to raise the Mab in mice.

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CONCLUSION The observations reported in this p a p e r are consistent with the view that different gene products are eventually stored in insect oocytes. While vitellogenins are taken up from the h a e m o l y m p h by a process of receptor-mediated endocytosis throughout vitellogenesis, follicle-specific proteins are secreted by the follicular epithelium and taken up by the oocyte, following a process as yet to be defined. Follicle-specific proteins are preferentially transferred to the ooplasm just prior to the beginning of chorionogenesis, where they are eventually stored as distinct cortical organelles. In this study, we provided morphological and biochemical evidence showing that the follicular epithelium in stick insects has the ability to synthesize and secrete an ovarian autonomous 75 k D a polypeptide. A monospecific monoclonal antibody allowed us to detect it in growing ovarian follicles but we were unable to reveal it immunocytochemically on thin sections. H o w e v e r , by both autoradiography on thick sections and fluorography on polyacrylamide gels, we proved that the 75 k D a polypeptide can be specifically labelled with sodium [35S]-sulphate and [3H]-glucosamine. Both these characteristics m a k e this follicle-specific product look like a glucosaminoglycan, such as that already identified in H y a l o p h o r a c e c r o p i a (Telfer, 1979). Ovarian follicles from other insect species have also been studied in their ability to elaborate egg-specific proteins and found to m a k e a major contribution to the yolk content of newly laid eggs (Yamashita and Indrasith, 1988). Because of the ovarian origin of the 75 k D a polypeptide in stick insects, in v i t r o cultured follicles would be expected to release it into the culture medium. The failure to detect it under our culturing conditions suggest that further experiments will be required to verify the proteolytic susceptibility of this protein (Tsuchida et a l . , 1992) and/or the hormonal dependence of its synthesis. Although follicle-specific proteins could be mainly nutritional, by providing amino acids for early embryogenesis, other possibilities should also be considered. Folliclespecific proteins could provide specific binding sites for vitellogenins or for other haemolymph-derived proteins to be retained in the intercellular spaces of the follicular epithelium (Telfer et al., 1982) as a prelude to endocytosis into the oocyte. It is also possible that follicle-specific proteins could be sorted in yolk granules differing from the ones containing vitellin. This could provide a mechanism for their differential utilization during embryogenesis. These possibilities are yet to be explored and are presently being investigated in our laboratory.

REFERENCES BASSEMIR, U. 1977. Ultrastructural differentiations in the developing follicle cortex of Locustra migratoria, with special reference to vitelline membrane formation. Cell Tissue Res. 185: 247~2. BELOTE, J. M., A. M. HANDLER, M. F. WOLFNER, K. J. LIVAKand B. S. BAKER. 1985. Sex specific regulation of yolk protein gene expression in Drosophila. Cell 40: 339-48. BOWNES, M. 1989. The roles of juvenile hormone, ecdysone and the ovary in the control of Drosophila vitellogenesis. J. Insect Physiol. 35: 409-13. BRADLEY, J. T., F. GIORGIand M. MASETrI. 1988. The effects induced by allatectomy in the stick insect Carausius morosus. Boll. Zool. 55: 11. BRADLEY, J. Y., ~V[.MAZZINIand F. GIORGI.1987. Vitellogenesis in the stick insect Bacillus rossius (Rossi) (Phasmatodea, Bacillidae). 6. Kinetics of vitellogenic development. Monit. Zool. ltal. 21: 99-115. DHADIALLA, T. S. and A. S. RAIKHEL.1991. Binding of vitellogenin to membranes isolated from mosquito ovaries. Arch. Insect Biochem. Physiol. 18: 55-70. FERENZ, H. J. 1990. The locust oocyte vitellogenin receptor. Function and characteristics. Adv. Invertebr. Reprod. ~',:1034)8.

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GIORGI,F., S. BUCCI-INNOCENTIand M. RAGGHIANTI. 1976. Osmium zinc iodide staining of Golgi elements in oocytes of Triturus cristatus. Cell Tissue Res. 172: 121-31. GIORGI, F. and F. MACCH1. 1980. Vitellogenesis in the stick insect Carausius morosus. I. Specific protein synthesis during ovarian development. J. Cell Sci. 46: 1-16. GIORGI,F., G. BALDINI,A. L. SIMONINIand M. MENGHERI. 1982. Vitellogenesis in the stick insect Carausius morosus. II. Purification and biochemical characterization of two vitellins from eggs. Insect Biochem. 12: 553-62. GIORGI, F. and M. MAZZINI. 1984. Vitellogenesis in the stick insect Bacillus rossius (Rossi) (Insecta, Phasmatodea Bacillidae). 2. Ultrastructural observations on developing oocytes. Monit. Zool. ltal. 18: 259-73. GIORG~, F. and M. MAZZ1N1. 1985. Cell to cell interactions in ovarian follicles of insects with different types of oogenesis. Redia 68: 439~66. GIORGI, F. and M. MAZZIN1. 1987. Secretory and endocytic pathways of vitellogenin in the stick insects. Adv. lnvertebr. Reprod. 4: 85-92. GIORGI, F., J. T. BRADLEY, R. VIGNALI and M. MAZZINI. 1989. An autoradiographic analysis of vitellogenin synthesis and secretion in the fat body of the stick insect Bacillus rossius. Tissue Cell 21: 543-58. GIORGI, F., A. CECCHER~F1NI and M. MASETTI. 1990. Changes in the patterns of proteins stored and synthesized by developing embryos of the stick insect Carausius morosus Br. Comp. Biochem. Physiol. 95: 107-13. GIORGI, F., C.-M. YIN and J. G. STOFFOLANO. 1991. Permeability barriers and anionic sites of the ovarian basal laminae in the black blowfly Phormia regina (Meigen) (Diptera, Calliphoridae). Int. J. Invertebr. Reprod. 19: 37--44. GocHoco, C. H., J. G. KUNKEL and J. H. NORDIN. 1988. Experimental modifications of an insect vitellin affect its structure and its uptake by oocytes. Arch. Insect Biochem. Physiol. 9: 179-99. GRAHAM, R. C. and M. J. KARNOVSKY. 1966. The early stages of absorption of injected peroxidase in the proximal tubules of mouse kidney: ultrastructural cytochemistry by a new technique. J. Histochem. Cytochem. 14: 291-302. HARNISH, D. G. and B. N. WHITE. 1982. An evolutionary model for insect vitellins. J. Mol. Evol. 28: 405-13. HUEBNER, E. 1983. Oostatic hormone antigonadotropin and reproduction, pp. 319-329. In R. G. H. DOWER and H. LAUFER(eds) Endocrinology o f Insects. Alan R. Liss, New York. KARNOVSKY, M. J. 1965. A formaldehyde-glutaraldehyde fixative of high osmolality for use in electron microscopy. J. Cell Biol. 276: 137A-38A. KOENIG, R., J. H. NORDIN, C. H. GOCHOCO and J. G. KUNKEL. 1988. Studies on ligand recognition by vitellogenin receptors in follicle membrane preparations of the German cockroach Blattella germanica. Arch. Insect Biochem. Physiol. 18: 395404. KOLLER, C. N., T. S. DHADIALLAand A. S. RAIKHEL. 1989. Selective endocytosis of vitellogenin by oocytes of the mosquito, Aedes aeypti: an in vitro study. Insect Biochem 19: 693-702. KUNKEL, J. G. and NORDIN, J. H. 1985. Yolk proteins, pp. 83-111. In G. A. KERKUTand L. I. GILBERT (eds) Comprehensive Insect Physiology, Biochemistry and Pharmacology, Vol. 1. Pergamon Press, Oxford. LAEMML1,U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (Lond. ) 227: 680-82. LOCKE, M. 1982. The cell biology of fat body development, pp. 227-252. In M. LOCKE and D. S. SMITH (eds) Insect Biology in the Future. Academic Press, New York. LOGAN, S. K. and P. C. WEr~SlNK. 1990. Ovarian follicle cell enhancers from the Drosophila yolk protein genes: different segments of one enhancer have different cell-type specificities that interact to give normal expression. Gen. Dev. 4: 613-23. MAILLET, M. 1963. Le r6active au t6traoxyde d'osmium-iodure du zinc. Z. Zellforsch. 70: 397-406. MASETTI, M. and F. GIORGI. 1989. Vitellin degradation in developing embryos of the stick insect Carausius morosus. J. Insect Physiol. 35: 689-97. MAZZ1N1, M. and F. GIORGI. 1984. Vitellogenesis in the stick insect Bacillus rossius (Rossi) (Insecta, Phasmatodea Bacillidae). 1. Ultrastructural observations on ovarian follicle cells. Monit. Zool. ltal. 18: 239-57. MAZZINI, M. and F. GIORG1. 1985. The follicle cell/oocyte interaction in ovarian follicles of the stick insect Bacillus rossius (Rossi). J. Morphol. 185: 37-49. MAZZINI, M. and F. GIORGI. 1986. Endocytic pathways in ovarian follicles of stick insect Bacillus rossius (Rossi) (Phasmatodea, Bacillidae). J. Submicrosc. Cytol. 18: 577-86. MAZZINI, M., M. MASETrl and F. GIORGI. 1986. Auto- and heterosynthetic processes in developing ovarian follicles of Bacillus rossius (Rossi) (Phasmatodea, Bacillidae). Int. J. Insect Morphol. Embryol. 15: 49-64. MESN1ER, M. 1980. Study of oocyte growth within the ovariole in a stick insect Clitumnus extradentatus. J. Insect Physiol. 26: 59-65.

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