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Allatostatin in ovaries, oviducts, and young embryos in the cockroach Diploptera punctata
Journal of Insect Physiology 49 (2003) 1103–1114 www.elsevier.com/locate/jinsphys
Allatostatin in ovaries, oviducts, and young embryos in the cockroach Diploptera punctata A.P. Woodhead, M.E. Thompson, K.K. Chan, B. Stay ∗ Department of Biological Sciences, University of Iowa, Biology Building, Iowa City, IA 52242, USA Received 16 June 2003; received in revised form 4 August 2003; accepted 4 August 2003
Abstract The quantity and localization of -Phe-Gly-Leu-amide allatostatins (-F-G-L-amide AST) was determined by ELISA and immunohistochemistry in ovaries and oviducts and in pre-dorsal closure embryos. AST in the cytoplasm of basal oocytes gradually increased from 4 to 35 fmol/ovary pair from the start (day 2) to the completion of vitellogenesis (day 6), then rapidly increased to 121 fmol/ovary pair during choriogenesis. In oviducts, AST-immunoreactivity was found in nerves to the muscle layer and in epithelial cells. AST-immunoreactivity in oviduct epithelial cells increased during vitellogenesis. A marked increase in quantity of AST in oviduct tissue between completion of chorion formation and immediately after ovulation appears to result from AST released from oocytes as they travel down the oviducts because AST content of newly ovulated eggs was 40% lower than late stage chorionated oocytes, and these oocytes released AST when incubated in saline. AST in embryos, localized in yolk cells, decreased as embryos approached dorsal closure. That this material in ovaries and embryos is AST was confirmed by its ability to inhibit JH synthesis in vitro and identification by MALDI–TOF mass spectrometry of a peptide with a mass corresponding to that of a Diploptera punctata AST. These findings indicate likely novel functions for ASTs: facilitation of ovulation and utilization of yolk. 2003 Elsevier Ltd. All rights reserved. Keywords: Cockroach allatostatin; Ovary; Oviduct; Embryo; Diploptera punctata
1. Introduction Allatostatins (ASTs), -F-G-L-amide peptides, were first isolated from brain extract of the viviparous cockroach Diploptera punctata (Woodhead et al., 1989; Pratt et al., 1989, 1991) by their ability to inhibit juvenile hormone (JH) synthesis by corpora allata (CA). Subsequently, the gene for the ASTs was cloned and found to encode a family of 13 ASTs, all with Y/F-X-F-G-L/Iamide at the C terminus (Donly et al., 1993). The -F-GL-amides occur in other insects and in other invertebrates, and have been found to have many functions including myo- and neuromodulation and stimulation of digestive enzyme activity (reviewed in Weaver et al., 1998; Bendena et al., 1999; Stay, 2000; Ga¨de, 2002). JH synthesis by the CA is regulated not only by neuro-
Corresponding author. Tel.: +1-319-335-1096; fax: +1-319-3351069. E-mail address: [email protected] (B. Stay). ∗
0022-1910/$ - see front matter 2003 Elsevier Ltd. All rights reserved. doi:10.1016/j.jinsphys.2003.08.006
secretions such as AST but also by the ovary that requires JH for vitellogenesis. Ovariectomized female Dictyoptera and Orthoptera typically have lower rates of JH synthesis than are found during vitellogenesis (Stay et al., 1983; Wennauer et al., 1989; Couillaud and Girardie, 1984). Since the ovary is required for the cycle of JH synthesis (Stay et al., 1983), this implies that the ovary is the source of both stimulatory and inhibitory factors. Evidence for both stimulation and inhibition of JH synthesis was found by implanting ovaries at different stages of development into ovariectomized females (Rankin and Stay, 1984, 1985). Ovaries with vitellogenic basal oocytes that grew to a mean length of 1.42 mm in 48 h stimulated JH synthesis (Rankin and Stay, 1984). The inhibitory effect of a larger ovary was demonstrated by finding less stimulation of JH synthesis by a stimulatory ovary when it was implanted with an ovary in which basal oocytes grew to a mean of greater than 1.47 mm (Rankin and Stay, 1985). How the ovary of D. punctata elicits stimulation and inhibition of the CA has yet to be determined. However,
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the ovaries of several insect species contain extractable material that inhibits JH synthesis by CA in vitro. Extracts of vitellogenic ovaries inhibited JH synthesis in Locusta migratoria (Gadot et al., 1987; Applebaum et al., 1990; Ferenz and Aden, 1993) and in the cricket Gryllus bimaculatus (Hoffmann et al., 1996). Although the nature of this inhibitory material in L. migratoria was not determined, recently, -F-G-L-amide ASTs were found in ovaries of the cricket G. bimaculatus (Witek and Hoffmann, 2001). HPLC separation of vitellogenic ovarian extract yielded fractions that inhibited JH synthesis by CA in vitro, and immunoreacted with antibody to a cricket -F-G-L-amide. Immunohistochemistry with this antibody demonstrated AST in the cortical cytoplasm of oocytes at the anterior pole not only in vitellogenic ovaries from adults but also in non-vitellogenic ovaries of penultimate and last-instar larvae. Garside et al. (2002), in studies on D. punctata, have provided evidence that ASTs are synthesized not only in ovaries but also in oviducts. They measured AST mRNA expression in ovary and oviducts during the first reproductive cycle. Although the quantity of AST mRNA/total RNA was highest in lateral oviducts, less in the common oviduct, and very little in the ovary, in all of these tissues, the quantity increased during vitellogenesis with the peak toward the end of vitellogenesis. Localization of the mRNA by in situ hybridization was not determined, and immunohistochemistry revealed AST only in nerves on the oviduct. In a search for ovarian factors that regulate JH synthesis, we conditioned saline with ovaries at different developmental stages and tested the ability of this conditioned saline to affect JH synthesis by CA. Ovaries with chorionated basal oocytes released a fast-acting, reversible inhibitory factor. We report here that this factor is AST and determine its quantity and distribution in ovary and oviducts during the first vitellogenic cycle and in early embryos. Quantitative measurements were made with ELISA and localization determined by immunohistochemistry, both with monoclonal antibody to DippuAST 7. The effect of ovary-conditioned saline and ovary and embryo extract on JH synthesis in vitro and the identification of a peptide in ovary-conditioned saline with the mass of an AST by matrix assisted laser desorption/ionization–time-of-flight (MALDI–TOF) mass spectrometry confirmed the presence of ASTs in the ovary and early embryo.
2. Materials and methods 2.1. Animals Newly molted adult females were collected daily and cohorts were maintained on lab chow (Purina, St. Louis,
MO) and water at 27 °C in a 12 h/12 h light/dark cycle. At this temperature, females typically drop the spermatophore between days 5 and 6 and oviposition typically occurs by day 7 or 8. 2.2. Timing of chorion stages and oviposition A survey was made of the stages of chorion development (classification by Stay et al., 1984: A = early, B = mid, C = late, and D = complete) in basal oocytes of ovaries from 5- to 7-day females (n = 130). The proportion of females at each stage was assumed to be in proportion to the duration of each stage. These data were used to estimate the age of the female in days at the midpoint of each chorion stage and the proportion of a day during which basal oocytes were in each stage (stage: midpoint age, duration—A: 5.9, 0.9; B: 6.6, 0.6; C: 7.1, 0.3; D: 7.4, 0.3). In a subset of these day 7 females, the proportion that oviposited by day 7 was used to estimate the age of the female at oviposition (7.5 d). Duration of oviposition was determined from observations of ovipositing females (0.02 d). 2.3. Preparation of conditioned saline and tissue extract Ovaries were removed from chilled females and transferred to a dish with a small amount of Yeager’s insect saline (Buck, 1953). Adhering fat body and trachea were removed, basal oocyte length was measured with an ocular micrometer, and the stage of chorion formation was assessed. For the preparation of ovary-conditioned saline, 5–8 pairs of ovaries, rinsed in phosphate-buffered saline (PBS) (Kingan, 1989), were transferred to a 1.5 ml polypropylene tube with 0.4 ml PBS and placed on a rotary shaker for 2 h at room temperature. After centrifugation, the saline was removed from ovaries and stored at ⫺80 °C. For preparation of ovary homogenate, 3–7 pairs of ovaries, rinsed in 0.9% NaCl (saline), were homogenized in 0.4 ml saline in a mini glass tissue grinder. The homogenate and a 0.1 ml saline rinse were transferred to a 1.5 ml tube, boiled for 10 min and centrifuged for 4 min at 8000 × g. The supernatant was stored at ⫺80 °C. For preparation of oviduct homogenate, the common oviduct, along with the pair of lateral oviducts, were removed as above for ovaries and, in addition, extraneous nerves were trimmed. Five to twenty oviducts were homogenized in 0.2–0.4 ml saline. The homogenate was extracted and stored as described above. Embryo batches were expelled from females by gentle dorsal–ventral pressure on the abdomen to evert the brood sac. Three to five batches were homogenized in 0.4 ml saline. The homogenate was extracted and stored as described above.
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2.4. C18 solid-phase extraction
2.7. Immunohistochemistry
Before ovary-conditioned saline or tissue extract samples were used for ELISA, bioassay, or HPLC they were purified by solid-phase extraction to obtain AST as described previously (Woodhead et al., 1989) except that Sep-Pak light cartridges (Waters, Milford, MA) were used with 4 ml rinses and 0.8 ml eluates. Each Sep-Pak cartridge was loaded with extract of approximately 5– 10 ovary pairs, 10–40 oviducts or 5–10 embryo batches. The 17–40% acetonitrile (ACN) eluate was used for further analysis.
Females were chilled on ice before abdominal tergites and gut were removed to expose reproductive organs. Unless otherwise stated in Results, the following protocol was used. After ovaries and oviducts were carefully and quickly cleared of fat body and trachea under Yeager’s insect saline, the solution was replaced with fixative (4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.2). Following 2–5 min fixation in situ, ovaries or oviducts and ovaries were removed and fixation continued overnight at 4 °C. Fixative was removed by repeated changes of PBS (5 × 20 min) and treated for 1 h with Tris-buffered saline (0.05 M Tris, 0.35 M NaCl) with 2% Triton X-100, 2% GS and 3% ovalbumin (TBS+) before incubation with the same monoclonal antibody used for ELISA. Tissue was exposed overnight at 4 °C to antibody in cell culture supernatant diluted 1:1 with TBS+ unless otherwise stated in Results. After repeated washes in TBS with 0.2% Triton X-100, anti-AST antibody was visualized with sheep anti-mouse Cy3 (Sigma, 1:300 in TBS+ 4 h at room temperature or overnight at 4 °C). After removal of secondary antibody, nuclei were stained with SYTOX Green (Molecular Probes, Eugene, OR) 1:1000 in TBS+ for 30 min before tissue was again washed as above and placed in 0.1 M Tris. Tissues were mounted in either 90% glycerol with 0.1 M n-propyl gallate or, after dehydration in a series of increasing concentrations of ethyl alcohol, in benzyl benzoate:benzyl alcohol (2:1 v/v). For paraffin sections, oviducts were fixed in Bouin’s fixative overnight at 4 °C, washed in 70% ethanol, dehydrated in a series of increasing concentrations of ethanol, cleared in methyl benzoate, and rinsed in toluene before infiltration with Paraplast (Electron Microscopy Science, Fort Washington, PA). Ten micrometer sections were cut, mounted on slides and, after removal of embedding medium, were rehydrated and treated with antibodies as for whole mounted tissue and mounted in glycerin. Images of tissues were obtained with a confocal laser microscope (Bio-Rad MRC 1024 with krypton/argon lasers and Nikon Optophot microscope). Excitations at 488 and 568 nm were recorded separately. Merging of labels was performed using Bio-Rad or Confocal Assistant software. Controls with no primary antibody showed no staining.
2.5. ELISA The antibody, 5F10 (described previously in Stay et al., 1992), is a monoclonal antibody to Dippu-AST 7 (APSGAQRLYGFGL-NH2). Cell culture supernatant was purified using caprylic acid followed by ammonium sulfate precipitation (Harlow and Lane, 1999). The antibody was then concentrated and lyophilized. This antibody is at least 2–3 orders of magnitude more sensitive to Dippu-AST 7 compared to Dippu-ASTs 2, 5, 8 and 9 (Stay et al., 1992) and Dippu-ASTs 4 and 11 (Woodhead, unpublished). AST quantities are expressed as Dippu-AST 7 equivalents. The ELISA procedure is adapted from Kingan’s method (1989). All solutions for ELISA were made with PBS, unless stated otherwise. At room temperature, titer plates (Corning Easywash, Corning, NY) were coated with AST coupled to ovalbumin (50 µl/well) for 3 h and then rinsed once with 0.05% Tween 20, blocked for 30 min with 1% goat serum (GS) (Sigma, St. Louis, MO) and rinsed again with 0.05% Tween 20 (100 µl/well). Sample fractions from solid-phase extraction and standards (25–2500 fmol Dippu-AST 7/well) were dried and then resuspended in 1% GS to which an equal volume of antibody solution, diluted 1:1000 with 1% GS, was added. Solutions were added in duplicate to the plate (100 µl/well) and stored at 4 °C overnight. After the plate was rinsed twice with 0.05% Tween 20, goat antimouse alkaline phosphatase (Promega, Madison, WI; 1:3000 in 1% GS) was added to the wells and incubated at room temperature for 2 h. The plate was rinsed four times with 0.05% Tween 20. The alkaline phosphatase was reacted with 0.4 mg phosphatase substrate (Sigma 104)/ml 10% diethanolamine. The absorbance of the reaction product was measured at 405 nm using a Titertek Multiscan MCC/340 plate reader. 2.6. Protein measurements The quantity of protein in oviducts was measured using Bradford’s (1976) method. Two to four oviducts were homogenized in 0.2 ml 1 M NaOH and, after centrifugation, the supernatant was used for protein analysis.
2.8. Delipidation and bioassay of ovary and embryo extracts Although ELISA AST determinations were made from the 17–40% ACN fraction from solid-phase extraction, lipid interfered with the in vitro assay for JH synthesis by CA. Therefore for bioassays, extract was delipidated before the C18 solid-phase extraction step by vortexing the sample with an equal volume of ice-cold
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n-hexane that had been saturated with 0.9% NaCl. After centrifugation at 4 °C, the hexane was removed, this procedure was repeated and the aqueous moiety was retained for solid-phase extraction. The sensitivity of CA to conditioned saline and tissue extracts was tested in the sequential in vitro JH assay, described in Stay et al. (1991), except that the labeled (60 methionine was l-[methyl-14C]methionine mCi/mmol, Amersham, Piscataway, NJ). The test CA were from 2-day virgin females (n = 4–6). Briefly, right and left glands were equilibrated at room temperature for 2 h in TC199 (GIBCO, Grand Island, NY) before rates of JH synthesis in untreated radioactive medium were determined in a 2 h incubation at 27 °C. Then glands were transferred to radioactive medium with conditioned saline/tissue extract (treated) or without (controls) for another 2 h. Treated and control glands were members of a pair. For statistical analysis, JH synthesis rates of right and left glands were compared for the first 2 h (untreated, untreated) and the second 2 h (treated, untreated) for ovary and embryo extract. For ovary-conditioned saline, rates of right (treated) and left (untreated) glands in a 2 h assay were compared. Percent inhibition was determined as described previously (Stay et al., 1991).
3. Results 3.1. Quantity of AST in ovaries and young embryos The amount of AST extracted from ovary homogenate during 10 d of the first reproductive cycle is compared to the amount of AST released by ovaries in vitro for 2 h (Fig. 1B) and shown in relation to mean basal oocyte length of the ovaries from which homogenates were prepared (Fig. 1A). Basal oocytes become vitellogenic between days 2 and 3 and continue vitellogenesis until days 5–6 when the spaces between follicle cells close and follicle cells begin to produce an eggshell (chorion). The AST content of ovary homogenates gradually increased from day 2 to early (A) chorion stage, then
2.9. HPLC and mass spectrometry The 17–40% ACN eluate of delipidated embryo extract and, alternatively, ovary-conditioned saline from C18 Sep-Pak was reduced in volume and applied to a C18 column (Waters). A gradient of 10–35% ACN over 50 min was used with 0.115% TFA as solvent A and ACN with 0.1% TFA as solvent B. Half minute fractions were collected, and aliquots of fractions with elution times of synthetic ASTs were tested by ELISA and bioassay. Several fractions were combined using elution times of synthetic ASTs as a guide and were applied to a C8 column (Jones Chromatography, Lakewood, CO). A gradient of 10–40% ACN over 45 min was used with the same solvents as for C18 HPLC. Half minute fractions were collected, and aliquots of fractions with elution times of Dippu-ASTs 7–9 were submitted for MALDI– TOF mass spectrometry analysis (Molecular Analysis Facility, University of Iowa). MALDI–TOF mass spectra were obtained using a Bruker Biflex III system (Bruker Daltonics, Billerica, MA) with α-cyano-4-hydroxycinnamic acid (Sigma) as matrix and angiotensin II and ACTH as calibrants. 2.10. Statistics Student’s t-test was used to compare AST and protein content data. For JH bioassays, rates of synthesis were compared using the Mann–Whitney U-test.
Fig. 1. AST contained and released by the ovary during the first vitellogenic cycle in relation to oocyte growth. (A) The mean length of basal oocytes in ovaries used for homogenate preparation in B. Each point is the mean of 27–85 measurements, SEM is equal to or less than 0.01 mm. The second value on day 5 represents measurements taken after females dropped the spermatophore. Choriogenesis (Chor) begins on day 6 and continues through day 7. The arrow indicates oviposition (Ovip). (B) AST released into saline by ovaries (—ο—) and AST content of ovary homogenates (—쎲—). Stages of choriogenesis: A = early, B = mid, C = late and D = complete. Sample sizes for AST released are: day 3, stages A and B, n = 1; stages C and D, n = 2 and 3, respectively. For ovary homogenate, each point represents a mean value (±SEM) of 3–7 samples.
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rapidly increased between the early (A) and late (C) chorion stages. In contrast, AST released by ovaries increased only slightly from day 3 to the mid (B) chorion stage, then rapidly increased between the mid (B) and late (C, D) chorion stages. The ovaries at these late chorion stages released about half their AST. After oviposition, the AST content of ovaries, now containing small non-vitellogenic basal oocytes, returned to a low level similar to that of previtellogenic ovaries on day 2. The basal oocytes (usually 12) are fertilized just after they leave the common oviduct and are aligned in two parallel rows, partially covered by an ootheca and then retracted into the brood sac where embryogenesis takes place. Transfer of nutrient to embryo from mother does not begin until after dorsal closure, on day 12–13 (Stay and Coop, 1973). Just after ovulation, AST content of eggs was 72.6 ± 11.6 fmol / batch (Fig. 2), a loss of about 40% of their AST (Fig. 2) compared to D stage ovaries (just before ovulation, Fig. 1) in which AST occurs in basal oocytes (see below Section 3.3). One day after oviposition AST content of embryos was 122.7 ± 6.6 fmol / batch, indicating rapid synthesis of AST by embryos in the day after oviposition (Fig. 2). However, between days 4 and 8, AST content of embryos decreased to about 50 fmol/batch and remained at this level on day 12. 3.2. Quantity of AST and protein in oviducts AST content of oviducts (i.e., both common and lateral) at specific stages of the first egg development cycle was also determined (Fig. 3). On day 2 at the start of vitellogenesis the AST content of oviducts was about half that of oviducts from day 7 females with completely chorionated basal oocytes (4.9 ± 0.8 vs. 8.4 ± 1.4 fmol / female, respectively). Over a shorter time span, from pre- to immediately post-ovulation on day 7, about
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Fig. 3. AST and protein content in oviduct homogenates. Oviducts were dissected on day 7 before ovulation (day 7 pre-ovul), immediately after ovulation (day 7.5 post-ovul), and 1 d post-ovulation (day 8.5 post-ovip). AST content (black bars) of post-ovulation oviducts was significantly greater than AST content of pre-ovulation oviducts (P ⬍ 0.02) and 1 day post-oviposition (P ⬍ 0.01); each bar represents the mean (±SEM) of 3–4 samples. Protein content (gray bars) of day 7 pre-ovulation oviducts was significantly greater than day 2 oviducts (P ⬍ 0.05) and day 7.5 post-ovulation oviducts (P ⬍ 0.01); each bar represents the mean (±SEM) of three samples.
half a day, AST content of oviducts increased to 18.4 ± 2.5 fmol / female, a significantly greater AST content than that of pre-ovulation oviducts (P ⬍ 0.02). One day after the batch of eggs was deposited in the brood sac, the AST content of the oviducts had decreased to the pre-ovulation level (8.9 ± 1.2 fmol / female). It should be noted that there is far less AST in day 7 oviducts (10– 20 fmol/female) than in day 7 (late chorion stage) ovaries (120–130 fmol/pr, Fig. 1B). The oviducts visibly increase in diameter as vitellogenesis progresses (see Fig. 7A and C). To determine whether this increase represented growth, the protein content of oviducts on day 2 and day 7 pre- and postovulation was determined (Fig. 3). Indeed, oviduct protein significantly increased from day 2 to day 7 pre-ovulation (from 42.6 ± 1.9 to 73.1 ± 2.2 µg / female, respectively, P ⬍ 0.01). Immediately after ovulation (day 7.5), protein in the oviducts significantly decreased to a level similar to that of day 2 oviducts (43.9 ± 8.8 µg / female, P ⬍ 0.05) and remained low 1 d post-oviposition (34.2 ± 3.1 µg / female). 3.3. Immunohistochemical localization of AST in ovary, oviduct and embryo
Fig. 2. AST content in embryo homogenates. Each point represents the mean (±SEM) of 3–6 samples.
The paired ovaries of D. punctata usually consist of six ovarioles each held together by a thin tracheated sheath. The ovarioles contain a series of follicles, oocytes surrounded by follicle cells, and only the basal follicles undergo vitellogenesis in each reproductive cycle (Fig. 4A and C). At the completion of vitellogen-
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Fig. 4. Diagrams of ovary and oviducts. (A) Innervation by a branch of ventral nerve 7 (VN7) of the common (CO) and lateral (LO) oviduct and their muscle attachments (M). Spermatheca (S) and accessory gland (AG) adhere closely to lateral oviduct (shown only on a portion of one lateral oviduct). The muscle from the copulatory bursa (Bu) to the ventral body wall is shown cut away where it extends over the lateral oviduct. (B) Median longitudinal section of a penultimate (PO) and basal oocyte (BO) shows basal oocyte surrounded by the chorion (Ch) with anterior micropyle (arrow), under the follicle cell layer (FCL). The pedicel (Pe) is at the base of the ovariole. (C) Cross-section of an oviduct. Muscle layers (M) surround a folded epithelium (E) covered by a thin cuticle with barbs projecting into the lumen (drawn from electron micrographs of lateral oviduct of a 4-day mated female).
esis the chorion secreted by the follicle has an anterior opening, the micropyle, through which sperm enter the oocyte as it leaves the common oviduct. Examination of follicles at several stages in the first vitellogenic cycle revealed increasing amounts of AST-immunoreactivity in the basal oocytes and none between or in the follicle cells (Fig. 5A–C). In mid- and late stages of chorion formation, immunoreactivity in intact basal oocytes is almost undetectable. This late stage is when most AST is detected by ELISA and when most AST is released from ovaries incubated in vitro (Fig. 1B). In ovaries incubated in vitro before fixation, immunoreactivity was conspicuous at the micropylar end of the basal oocytes (Fig. 5D). When chorionated oocytes were cut after fixation and exposed for several days to antibody, immunoreactivity was obvious in the oocyte cytoplasm (Fig. 5E). Oocytes removed from the female as they left the genital chamber showed immunoreactivity at the micropyle and in a few places on the surface of the chorion (Fig. 5F). Occurrence of AST at the micropyle and on the surface of the chorion suggests that AST is released from the micropyle as oocytes travel down the oviducts. To investigate this possibility ovaries and oviducts of females in the process of ovulation were exposed and fixed in situ. Fig. 6A shows one oocyte leaving the follicle cell layer and three oocytes in the lateral oviducts. The oocyte partially separated from the follicle cells showed immunoreactivity not only at the micropyle but also in the tiny lumen of the follicle cell sheath (Fig.
6B). Immunoreactivity was also evident in the lumen of the lateral oviduct below the ovary in which all basal oocytes had been ovulated (Fig. 6C). The possibility that some of the immunoreactivity was also in epithelial cells of the oviduct was investigated in confocal sections of oviducts from females prior to ovulation. Oviducts from 2-day females, with oocytes at the start of vitellogenesis, showed strong immunoreactivity in nerves on the surface of the oviducts (Fig. 7A) but, at low magnification, little immunoreactivity was evident in the epithelial cells of the lateral or common oviducts, whereas some immunoreactivity was evident at higher magnification (Fig. 7B). In contrast, in oviducts from 7-day females, in addition to strong immunoreactivity in nerves (Fig. 7C), in lowmagnification confocal sections immunoreactivity was obvious at the level of epithelial cells (Fig. 7D). In crosssection, the distinction between the epithelial and muscle layers is obvious (Fig. 4C). Cross-sections of oviducts from females in late vitellogenesis confirmed that AST occurs in epithelial cells. Confocal images of such sections showed immunostaining at the apex of the epithelial cells and in the space between the cells and the cuticular lining (Fig. 7E and F). Thus the amount of AST in pre-ovulation oviducts is a combination of AST in the epithelial cells and in the nerves to the muscle cell layer of the oviduct. AST in embryos occurs in yolk cells (Fig. 7G) The nerves serving the muscle layers of the oviducts are branches of ventral nerve 7 (VN7) from the last
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Fig. 5. Fluorescent AST-immunoreactivity in basal oocytes and an ovulated egg. (A–C) show increasing reactivity in vitellogenic oocytes from 2-, 4-, and 6-day females, respectively. Inset in C shows follicle cells, with green nuclei, devoid of AST-immunoreactivity (red) which is present only in the oocyte cytoplasm. (D) Portion of an ovary with completely chorionated oocytes fixed after incubation in buffered saline for 2 h shows strong immunoreactivity only at the micropylar ends of the basal oocytes. (E) Basal follicle with chorionated oocyte that was cut open before exposure to antibody reveals immunoreactivity in oocyte cytoplasm. (F) Ovulated egg fixed as it emerged from the genital chamber shows immunoreactivity at the micropyle (top) and a small amount on the surface of the intact chorion. Scale bars = µm.
abdominal ganglion (Fig. 4A). A variable distribution of nerves occurs in oviducts even though great care is exercised in their removal from the female (e.g., Fig. 7A and B). However, remnants of nerve seen in whole mounts most frequently occur on the common oviduct and near the ovary on the lateral oviducts. In two sets of experiments, oviducts (triplicates of each of three stages) with attached nerves were exposed to a more dilute antibody (1:40) in order to distinguish potential differences in immunoreactivity that might be stage specific. Little obvious difference is seen in intensity of immunoreactivity at different stages of the first egg development cycle. Fig. 8 shows similar immunoreactivity in low-magnification confocal images of VN7 as it contacts the lateral oviducts from 2-day females, and lateral oviducts from 7-day females with chorionated oocytes, and after a batch of eggs was oviposited. At high magnification the immunoreactive axons of the nerve as it contacts the oviduct show varicosities that appear to be on the surface of the nerve where they could be acting as neurohemal release sites (Fig. 8D). 3.4. Effect of ovary-conditioned saline and ovary and embryo extracts on JH synthesis in vitro Inhibition of JH synthesis by CA in vitro confirmed the presence of allatostatic material in ovaries and
embryos. Bioassay with saline conditioned by C and D stage chorionated basal oocytes did not inhibit JH synthesis rates significantly at 1 ovary equivalent/CA (10.5 ± 8.6%) however, at 5 ovary equivalents/CA inhibition was 45.7 ± 10.0%; rates were significantly lower than rates of untreated controls (P = 0.01). Treatment with higher doses did not increase inhibition presumably because of interfering material in the extract. Therefore extracts of ovaries with C and D stage chorionated basal oocytes and 1-day embryos were further purified by delipidation and tested on CA in vitro. Based on ELISA quantification, CA were treated with the equivalent of 10⫺8 M AST (8.9 ovary pair equivalents and 7.6 embryo batch equivalents/CA). JH synthesis rates of CA treated with ovary and embryo extract were significantly inhibited compared to untreated controls (62.8 ± 3.1%, P ⬍ 0.05; 52.4 ± 3.2%, P ⬍ 0.01, respectively). For all bioassays, there was no significant difference in rates of JH synthesis between groups of untreated CA in the first 2 h incubation. Because batches of embryos retain much ovarian AST and are more easily obtained than late-stage ovaries, extract of 1-day embryos (190 batches) was purified by HPLC on a C18 column. Material with the retention time of Dippu-AST 2 (AYSYVSEYKRLPVYNFGL-NH2) and material from two other HPLC fractions not expected to contain AST were tested. Inhibition by these
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Fig. 6. Ovaries and oviducts fixed in the process of ovulation. (A) Only one basal oocyte remains partially within the follicle cell layer in the left ovary (long arrow). Box on the left is shown at higher magnification in B. Three oocytes are in the distended lateral oviducts (short arrows). Nerves on the surface of the common oviduct (CO) show strong immunoreactivity. Box on the right is shown at higher magnification in C. Note: some ovarioles were removed to facilitate the mounting of tissue. (B) Median section of anterior portion of a follicle with oocyte (O) departing the follicle cell layer (FC). Immunoreactivity (red) appears at the micropyle of the chorion (long arrow) and in the narrow lumen of the follicle cells (short arrows). (C) Median confocal section of oviduct after passage of six oocytes. AST-immunoreactivity (red) occurs in the lumen and in oviduct epithelial cells. Nuclei are green. Scale bars = µm.
other fractions was 13 ± 6% and 5 ± 7%, whereas the Dippu-AST 2 fraction inhibited JH synthesis rates 70.0 ± 6.0% when CA were treated with the equivalent of 10⫺8 M Dippu-AST 2, estimated from ELISA measurements for Dippu-AST 7. This assumes that all the ASTs are produced in approximately equal amounts (see Discussion). 3.5. MALDI–TOF mass spectrometry MALDI–TOF mass spectrometry analysis of supernatant of boiled saline extract of D. punctata brains identified the masses of eight of the 13 peptides of the Dippu-AST family (data not shown). However, no masses of any ASTs were detected in similarly prepared ovary-conditioned saline or extract of ovaries. This likely was due to the approximately 100× greater quantity of soluble proteins relative to ASTs in ovarian tissue compared to brain tissue, and also the greater quantity of lipid in the ovarian extract. Therefore, fractions from the C18 HPLC separation of conditioned saline from 400 ovary pairs expected to contain Dippu-ASTs 7–9 were
further purified on a C8 column. MALDI–TOF mass spectrometry analysis of the fractions expected to contain Dippu-ASTs 7 and 9 identified an (M+H)+ ion observed at M/Z 1335.51, which corresponds to the monoisotopic mass of Dippu-AST 7 (1334.69). Masses of Dippu-ASTs 8 and 9 were not found. This was likely due to loss of AST material during HPLC purification steps and the inability to detect AST masses when dilute solutions were concentrated for MALDI–TOF analysis. 4. Discussion This study of the quantity and localization of AST in ovaries, oviducts, and embryos in relation to the stages of vitellogenesis and choriogenesis and embryonic development in D. punctata strongly suggests new roles for this multifunctional -Phe-Gly-Leu-amide family of peptides. It is likely that AST in oocytes, epithelial cells of the oviduct, and nerves to the oviduct muscles, functions in the process of ovulation, and after ovulation, AST in the fertilized egg (embryo) functions in yolk utilization.
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Fig. 7. AST-immunoreactivity in pre-ovulation oviducts and a young embryo. (A) Common and lateral oviducts of a 2-day female with basal oocytes in early vitellogenesis. Note immunoreactivity in nerves on the surface of common oviduct (CO) and lateral oviducts near the ovary (arrows). Boxed area is shown at higher magnification in B. (B) Portion of a common oviduct in a confocal section at the level of epithelial cells shows AST-immunoreactivity (small red dots); larger spots are likely varicosities of nerves. Nuclei are pale green; barbs on cuticular lining are bright green (photographed with 60× objective). (C) Common and lateral oviducts of a 7-day female with completely chorionated basal oocytes. Note distribution of nerves as in A. Boxed area is shown at higher magnification in D. (D) AST-immunoreactivity (red) in common oviduct in a confocal section at the level of epithelial cells (photographed with 20× objective); barbs not visible at this magnification. (E) Cross-section, cut in paraffin, of common oviduct of a female at a late stage of vitellogenesis shows folded epithelium (e) and surrounding muscle cell layer (m). (F) Portion of the cross-section in E at higher magnification shows AST-immunoreactivity in apex of epithelial cells and in space between cells and cuticular lining. Barbs (short arrows) on the cuticle layer (long arrow) are bright green. (G) Part of a 9-day, pre-dorsal closure, embryo (Emb). Yolk cells (YC) are strongly AST-immunoreactive; nuclei are green. Scale bars = µm.
4.1. Function of ovarian AST during ovulation We have confirmed, for D. punctata, Witek and Hoffmann’s (2001) observation in cricket ovaries that AST occurs in oocytes and not in follicle cells. Our study shows the occurrence of AST principally in the basal oocytes and an increase in quantity closely correlated with stage of development. Although AST increases from the start of vitellogenesis, the greatest increase is during choriogenesis. At this stage, when ovaries were
incubated in saline, AST-immunoreactive material accumulated at the micropyle of the chorion and was found in the conditioned saline. That the release of AST also occurs physiologically was demonstrated by finding AST-immunoreactivity at the micropyle, between follicle cell layers and in the lumen of the oviduct in tissue fixed in a female in the process of ovulation. Further evidence to support this hypothesis is the increase in amount of AST in oviducts immediately after ovulation on day 7; and that this increase is correlated with a 40%
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Fig. 8. AST-immunoreactivity in a piece of ventral nerve 7 as it contacts the lateral oviduct in a 2-day female (A); a 7-day female with chorionated basal oocytes (B); and a 7-day female a few minutes after ovulation (C). High magnification confocal image of a portion of the nerve in A shows nuclei (green) of nerve sheath cells and an AST-immunoreactive axon (red) on the surface of the nerve. These preparations were exposed to antiDippu-AST 7 diluted 1:40 in order to detect possible differences in intensity of reactivity with stage of vitellogenesis. Scale bars = µm; A–C are the same magnification.
reduction in AST in the eggs measured immediately after ovulation. Thus the AST in oviducts could be, at least in part, that released by the oocytes from the micropyle. Indeed AST was observed at the micropyle and on the surface of newly ovulated eggs. The amount of AST in oocytes and the timing of its release indicate that oocyte AST is unlikely to function as a signal for the decline in JH synthesis during vitellogenesis, as this occurs before choriogenesis begins (Rankin and Stay, 1984; Pratt et al., 1990). Thus, if release of AST by oocytes were to have a regulatory role in JH production it would be to maintain low rates of synthesis at this time. However, this is unlikely since the AST released by oocytes during ovulation, if it reached the hemolymph, would not increase the AST concentration sufficiently to inhibit CA (Stay et al., 1994). Release of AST from nerves within the CA more effectively inhibits JH production than elevated AST in the hemolymph (Stay et al., 1994). Although the fraction of the AST in the CA that is within nerves cannot be determined by ELISA, it is interesting to note that extract of a pair of CA contains 0.3 pmol equivalents of Dippu-AST 7 (Lloyd et al., 2000) whereas the maximum quantity contained in a pair of ovaries is 0.12 pmol equivalents. 4.2. Functions of AST in oviducts Quantitative measurements of AST in extracts of oviducts cannot distinguish between material in nerves, epithelial cells or in the lumen. However, since the amount of AST-immunoreactivity seen in whole mounts of nerves to the oviduct does not appear to be different on day 2 or day 7 before or after ovulation, we presume that changes in quantity of AST in oviducts is due to peptides in the epithelial cells and/or in the lumen of the oviducts. The presence of AST-immunoreactivity in epithelial cells of pre-ovulation oviducts, as seen in
whole mount images at the level of the epithelial cells and in cross-sections cut in paraffin, clearly shows that, in addition to AST in nerves and AST that may be released by the oocytes during ovulation, AST in epithelial cells contributes to the quantity of AST extractable from oviducts both before and after ovulation. Comparison of the quantity of AST in oviducts of day 2 and day 7 pre-ovulation stages suggests that AST in oviduct cells increases during vitellogenesis as does the AST mRNA quantified by Garside et al. (2002). How the AST in the cells of the oviduct functions is yet to be determined. One hypothesis is that it facilitates the secretion of a lubricating material during ovulation. The sharp decrease in protein content of the oviducts from pre-ovulation to immediately post-ovulation on day 7 would indicate that a substance is released during ovulation. Electron micrographs of lateral oviducts reveal many mitochondria and large areas of glycogen in epithelial cells (Stay, unpublished). The glycogen could be a source of carbohydrate for a mucoprotein secretion. The abundance of mitochondria suggests that AST could be acting to regulate the ion transport necessary for the movement of sufficient water to facilitate the flow of the material out of the cells during ovulation. Further study will be required to test this hypothesis. The role of AST in the nerves to the oviduct is also yet to be determined. AST-immunoreactivity is clearly evident in nerves that innervate the oviduct muscle layer, the anchoring muscles of the common and lateral oviducts, and the muscles that pass over the lateral oviducts. Therefore, it seems likely that AST functions to regulate activity of these muscles in the complex process of movement of the oocytes along and out of the oviducts. Although no effect of AST on oviduct muscle in vitro has yet been demonstrated in D. punctata (Lange et al., 1995; Garside et al., 2002) or L. migratoria (Lange et al., 1993), AST is a strong inhibitor of proctolin-induced muscle contraction in the hindgut of D. punctata (Lange
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et al., 1993). Perhaps in the oviducts AST modulates muscle activity in conjunction with another (unidentified) modulator or neurotransmitter. It is also conceivable that the sensitivity of oviduct muscle to AST is stage specific and the sensitive stage has not been examined. The varicosities seen on the surface of the branch of VN7 leading to the oviducts suggest that these axons also release into the hemolymph and may affect muscles of the reproductive organs humorally. 4.3. Function of AST in early embryos Of the AST accumulated in the basal oocytes at the end of choriogenesis, 40% is lost during ovulation, but one day later the amount of AST in the embryos has increased to the level of pre-oviposition ovaries. This suggests that at the start of development, embryos supplied with a large store of AST by the mother are synthesizing AST well before the nervous system begins to express these peptides at about day 15, three days postdorsal closure (Stay, unpublished). In the embryo the yolk of the oocyte is incorporated into yolk cells and these are clearly AST-immunoreactive. This suggests that AST is modulating activity of nutrient digestion, as has been demonstrated in midgut of adult D. punctata (Fuse´ et al., 1999). Most of the yolk is depleted shortly after the embryo begins to drink the nutrient milk produced by the brood sac (Stay and Coop, 1973) and the amount of AST in embryos has decreased markedly by dorsal closure (day 12). 4.4. The identity of AST-immunoreactive material The quantity and localization of AST in the ovaries, oviducts and embryos is based on immunoreactivity to an antibody against Dippu-AST 7. The presence of Dippu-AST 7 was confirmed by MALDI-TOF spectrometry on ovary-conditioned saline. The ability of ovary-conditioned saline, and ovary and embryo extract to inhibit JH synthesis by CA in vitro is further evidence that the material is allatostatin. Not surprisingly, ASTimmunoreactivity occurs in vitellogenic oocytes of other cockroaches: Periplaneta americana, Leucophaea maderae and Schultesia lampyridiformis, and in chorionated blowfly oocytes (Stay, personal observation). It will be of interest to determine whether locust ovarian allatostatic activity is due to a -F-G-L-amide peptide and whether it also occurs in oviduct cells. Inhibition of JH synthesis by a fraction of embryo extract that eluted from a C18 column at the time of Dippu-AST 2 indicates that ASTs in addition to DippuAST 7 are present in extract of embryos. It is very likely that all members of the Dippu-AST family are produced, as multiple members have been isolated in D. punctata from brain (Woodhead et al., 1989), CA (Stay and Woodhead, 1993), hemolymph (Woodhead et al., 1993)
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and antennal heart muscle (Woodhead et al., 1992). Also multiple members of the P. americana AST family were identified from antennal heart muscle of that species (Predel et al., 1999). The quantitative RT–PCR analysis of mRNA for Dippu-ASTs in ovaries and oviducts during the first vitellogenic cycle in D. punctata (Garside et al., 2002) is convincing evidence that AST detected immunologically is synthesized in these tissues. The mRNA results, expressed as the amount of AST mRNA/total RNA, give the impression that the amount and increase of AST message during the vitellogenic cycle are more prominent in oviducts than in ovaries. However, if total RNA in vitellogenic ovaries is much greater than that in oviducts this would explain the apparent lower quantity of AST mRNA in ovaries. By conditioning saline with ovaries we hoped to obtain factors involved in the coordination of increase and decrease in JH production with ovarian development. We have found a factor, AST, that can modulate JH synthesis. However, because of its appearance late in the first egg development cycle, AST in the mature oocytes and oviducts most likely does not function to regulate JH synthesis, but more likely has totally different functions, such as facilitation of ovulation and utilization of yolk in the embryo. Acknowledgements The authors would like to thank John Hass for his help with tissue preparations and Shelley Plattner for his technical assistance with figures. This work is supported by NIH grant AI 15230. References Applebaum, S.W., Hirsch, J., El-Hadi, F.A., Moshitzky, P., 1990. Trophic control of juvenile hormone synthesis and release in locusts. In: Epple, A., Scanes, C.G., Stetson, M.H. (Eds.), Progress in Comparative Endocrinology. Wiley-Liss, New York, pp. 186– 192. Bendena, W.G., Donly, B.C., Tobe, S.S., 1999. Allatostatins: a growing family of neuropeptides with structural and functional diversity. Annals of the New York Academy of Sciences 897, 311–329. Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72, 248–254. Buck, J.B., 1953. Physical properties and chemical composition of insect blood. In: Roeder, K.D. (Ed.), Insect Physiology. Wiley and Sons, New York, pp. 147–190. Couillaud, F., Girardie, A., 1984. Ovariectomy and juvenile hormone JH-3 biosynthesis in African locust Locusta migratoria. Comptes Rendus des Seances de L’Academie des Sciences Serie III Sciences de la Vie 298, 157–162. Donly, B.C., Ding, Q., Tobe, S.S., Bendena, W.G., 1993. Molecular cloning of the gene for the allatostatin family of neuropeptides from the cockroach Diploptera punctata. Proceedings of the National Academy of Sciences, USA 90, 8807–8811.
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Ferenz, H.-J., Aden, E., 1993. Ovarian control of juvenile hormone biosynthesis in Locusta migratoria (Orthoptera: Acrididae). Entomologia Generalis 18, 9–17. Fuse´ , M., Zhang, J.R., Partridge, E., Nachman, R.J., Orchard, I., Bendena, W.G., Tobe, S.S., 1999. Effects of an allatostatin and a myosuppressin on midgut carbohydrate enzyme activity in the cockroach Diploptera punctata. Peptides 20, 1285–1293. Ga¨ de, G., 2002. Allatoregulatory peptides—molecules with multiple functions. Invertebrate Reproduction and Development 41, 127– 135. Gadot, M., Rafaeli, A., Applebaum, S.W., 1987. Partial purification and characterization of locust allatotropin I. Archives of Insect Biochemistry and Physiology 4, 213–223. Garside, C.S., Koladich, P.M., Bendena, W.G., Tobe, S.S., 2002. Expression of allatostatin in the oviducts of the cockroach Diploptera punctata. Insect Biochemistry and Molecular Biology 32, 1089–1099. Harlow, E., Lane, D., 1999. Using Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Hoffmann, K.H., Lorenz, M.W., Klein, P.M., 1996. Ovarian control of juvenile hormone-III biosynthesis in the cricket species Gryllus bimaculatus (Ensifera: Gryllidae). Entomologia Generalis 20, 157–167. Kingan, T.G., 1989. A competitive enzyme-linked immunosorbent assay: applications in the assay of peptides, steroids, and cyclic nucleotides. Analytical Biochemistry 183, 283–289. Lange, A.B., Chan, K.K., Stay, B., 1993. Effect of allatostatin and proctolin on antennal pulsatile organ and hindgut muscle in the cockroach, Diploptera punctata. Archives of Insect Biochemistry and Physiology 24, 79–92. Lange, A.B., Bendena, W.G., Tobe, S.S., 1995. The effect of the thirteen Dip-allatostatins on myogenic and induced contractions of the cockroach (Diploptera punctata) hindgut. Journal of Insect Physiology 41, 581–588. Lloyd, G.T., Woodhead, A.P., Stay, B., 2000. Release of neurosecretory granules within the corpus allatum in relation to the regulation of juvenile hormone synthesis in Diploptera punctata. Insect Biochemistry and Molecular Biology 30, 739–746. Pratt, G.E., Farnsworth, D.E., Siegel, N.R., Fok, K.F., Feyereisen, R., 1989. Identification of an allatostatin from adult Diploptera punctata. Biochemical and Biophysical Research Communications 163, 1243–1247. Pratt, G.E., Farnsworth, D.E., Feyereisen, R., 1990. Changes in the sensitivity of adult cockroach corpora allata to a brain allatostatin. Molecular and Cellular Endocrinology 70, 185–195. Pratt, G.E., Farnsworth, D.E., Fok, K.F., Siegel, N.R., McCormack, A.L., Shabanowitz, J., Hunt, D.F., Feyereisen, R., 1991. Identity of a second type of allatostatin from cockroach brains: an octadecapeptide amide with a tyrosine-rich address sequence. Proceedings of the National Academy of Sciences, USA 88, 2412–2416. Predel, R., Kellner, R., Rapus, J., Ga¨ de, G., 1999. Allatostatins from the retrocerebral complex and antennal pulsatile organ of the American cockroach: structural elucidation aided by matrix-assisted laser desorption/ionization-time-of-flight mass spectrometry. Regulatory Peptides 82, 81–89.
Rankin, S.M., Stay, B., 1984. The changing effect of the ovary on rates of juvenile hormone synthesis in Diploptera punctata. General and Comparative Endocrinology 54, 382–388. Rankin, S.M., Stay, B., 1985. Ovarian inhibition of juvenile hormone synthesis in the viviparous cockroach, Diploptera punctata. General and Comparative Endocrinology 59, 230–237. Stay, B., 2000. A review of the role of neurosecretion in the control of juvenile hormone synthesis: a tribute to Berta Scharrer. Insect Biochemistry and Molecular Biology 30, 653–662. Stay, B., Coop, A., 1973. Developmental stages and chemical composition in embryos of the cockroach, Diploptera punctata, with observations on the effect of diet. Journal of Insect Physiology 19, 147–171. Stay, B., Woodhead, A.P., 1993. Neuropeptide regulators of insect corpora allata. American Zoology 33, 357–364. Stay, B., Tobe, S.S., Mundall, E.C., Rankin, S., 1983. Ovarian stimulation of juvenile hormone biosynthesis in the viviparous cockroach, Diploptera punctata. General and Comparative Endocrinology 52, 341–349. Stay, B., Ostedgaard, L.S., Tobe, S.S., Strambi, A., Spaziani, E., 1984. Ovarian and haemolymph titres of ecdysteroid during the gonadotrophic cycle in Diploptera punctata. Journal of Insect Physiology 30, 643–651. Stay, B., Joshi, S., Woodhead, A.P., 1991. Sensitivity to allatostatins of corpora allata from larval and adult female Diploptera punctata. Journal of Insect Physiology 37, 63–70. Stay, B., Chan, K.K., Woodhead, A.P., 1992. Allatostatin-immunoreactive neurons projecting to the corpora allata of adult Diploptera punctata. Cell and Tissue Research 270, 15–23. Stay, B., Sereg Bachmann, J.A., Stoltzman, C.A., Fairbairn, S.E., Yu, C.G., Tobe, S.S., 1994. Factors affecting allatostatin release in a cockroach (Diploptera punctata): nerve section, juvenile hormone analog and ovary. Journal of Insect Physiology 40, 365–372. Weaver, R.J., Edwards, J.P., Bendena, W.G., Tobe, S.S., 1998. Structures, functions and occurrence of insect allatostatic peptides. In: Coast, G.M., Webster, S.G. (Eds.), Recent Advances in Arthropod Endocrinology. Cambridge University Press, Cambridge, pp. 3–32. Wennauer, R., Kassel, L., Hoffmann, K.H., 1989. The effects of juvenile hormone, 20-hydroxyecdysone, precocene II, and ovariectomy on the activity of the corpora allata (in vitro) in adult female Gryllus bimaculatus. Journal of Insect Physiology 35, 299–304. Witek, G., Hoffmann, K.H., 2001. Immunological evidence for FGLamide- and W2W9-allatostatins in the ovary of Gryllus bimaculatus (Ensifera, Gryllidae). Physiological Entomology 26, 49–57. Woodhead, A.P., Stay, B., Seidel, S.L., Khan, M.A., Tobe, S.S., 1989. Primary structure of four allatostatins: neuropeptide inhibitors of juvenile hormone synthesis. Proceedings of the National Academy of Sciences, USA 86, 5997–6001. Woodhead, A.P., Stoltzman, C.A., Stay, B., 1992. Allatostatins in the nerves of the antennal pulsatile organ muscle of the cockroach Diploptera punctata. Archives of Insect Biochemistry and Physiology 20, 253–263. Woodhead, A.P., Asano, W.Y., Stay, B., 1993. Allatostatins in the haemolymph of Diploptera punctata and their effect in vivo. Journal of Insect Physiology 39, 1001–1005.
Allatostatin in ovaries, oviducts, and young embryos in the cockroach Diploptera punctata A.P. Woodhead, M.E. Thompson, K.K. Chan, B. Stay ∗ Department of Biological Sciences, University of Iowa, Biology Building, Iowa City, IA 52242, USA Received 16 June 2003; received in revised form 4 August 2003; accepted 4 August 2003
Abstract The quantity and localization of -Phe-Gly-Leu-amide allatostatins (-F-G-L-amide AST) was determined by ELISA and immunohistochemistry in ovaries and oviducts and in pre-dorsal closure embryos. AST in the cytoplasm of basal oocytes gradually increased from 4 to 35 fmol/ovary pair from the start (day 2) to the completion of vitellogenesis (day 6), then rapidly increased to 121 fmol/ovary pair during choriogenesis. In oviducts, AST-immunoreactivity was found in nerves to the muscle layer and in epithelial cells. AST-immunoreactivity in oviduct epithelial cells increased during vitellogenesis. A marked increase in quantity of AST in oviduct tissue between completion of chorion formation and immediately after ovulation appears to result from AST released from oocytes as they travel down the oviducts because AST content of newly ovulated eggs was 40% lower than late stage chorionated oocytes, and these oocytes released AST when incubated in saline. AST in embryos, localized in yolk cells, decreased as embryos approached dorsal closure. That this material in ovaries and embryos is AST was confirmed by its ability to inhibit JH synthesis in vitro and identification by MALDI–TOF mass spectrometry of a peptide with a mass corresponding to that of a Diploptera punctata AST. These findings indicate likely novel functions for ASTs: facilitation of ovulation and utilization of yolk. 2003 Elsevier Ltd. All rights reserved. Keywords: Cockroach allatostatin; Ovary; Oviduct; Embryo; Diploptera punctata
1. Introduction Allatostatins (ASTs), -F-G-L-amide peptides, were first isolated from brain extract of the viviparous cockroach Diploptera punctata (Woodhead et al., 1989; Pratt et al., 1989, 1991) by their ability to inhibit juvenile hormone (JH) synthesis by corpora allata (CA). Subsequently, the gene for the ASTs was cloned and found to encode a family of 13 ASTs, all with Y/F-X-F-G-L/Iamide at the C terminus (Donly et al., 1993). The -F-GL-amides occur in other insects and in other invertebrates, and have been found to have many functions including myo- and neuromodulation and stimulation of digestive enzyme activity (reviewed in Weaver et al., 1998; Bendena et al., 1999; Stay, 2000; Ga¨de, 2002). JH synthesis by the CA is regulated not only by neuro-
Corresponding author. Tel.: +1-319-335-1096; fax: +1-319-3351069. E-mail address: [email protected] (B. Stay). ∗
0022-1910/$ - see front matter 2003 Elsevier Ltd. All rights reserved. doi:10.1016/j.jinsphys.2003.08.006
secretions such as AST but also by the ovary that requires JH for vitellogenesis. Ovariectomized female Dictyoptera and Orthoptera typically have lower rates of JH synthesis than are found during vitellogenesis (Stay et al., 1983; Wennauer et al., 1989; Couillaud and Girardie, 1984). Since the ovary is required for the cycle of JH synthesis (Stay et al., 1983), this implies that the ovary is the source of both stimulatory and inhibitory factors. Evidence for both stimulation and inhibition of JH synthesis was found by implanting ovaries at different stages of development into ovariectomized females (Rankin and Stay, 1984, 1985). Ovaries with vitellogenic basal oocytes that grew to a mean length of 1.42 mm in 48 h stimulated JH synthesis (Rankin and Stay, 1984). The inhibitory effect of a larger ovary was demonstrated by finding less stimulation of JH synthesis by a stimulatory ovary when it was implanted with an ovary in which basal oocytes grew to a mean of greater than 1.47 mm (Rankin and Stay, 1985). How the ovary of D. punctata elicits stimulation and inhibition of the CA has yet to be determined. However,
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the ovaries of several insect species contain extractable material that inhibits JH synthesis by CA in vitro. Extracts of vitellogenic ovaries inhibited JH synthesis in Locusta migratoria (Gadot et al., 1987; Applebaum et al., 1990; Ferenz and Aden, 1993) and in the cricket Gryllus bimaculatus (Hoffmann et al., 1996). Although the nature of this inhibitory material in L. migratoria was not determined, recently, -F-G-L-amide ASTs were found in ovaries of the cricket G. bimaculatus (Witek and Hoffmann, 2001). HPLC separation of vitellogenic ovarian extract yielded fractions that inhibited JH synthesis by CA in vitro, and immunoreacted with antibody to a cricket -F-G-L-amide. Immunohistochemistry with this antibody demonstrated AST in the cortical cytoplasm of oocytes at the anterior pole not only in vitellogenic ovaries from adults but also in non-vitellogenic ovaries of penultimate and last-instar larvae. Garside et al. (2002), in studies on D. punctata, have provided evidence that ASTs are synthesized not only in ovaries but also in oviducts. They measured AST mRNA expression in ovary and oviducts during the first reproductive cycle. Although the quantity of AST mRNA/total RNA was highest in lateral oviducts, less in the common oviduct, and very little in the ovary, in all of these tissues, the quantity increased during vitellogenesis with the peak toward the end of vitellogenesis. Localization of the mRNA by in situ hybridization was not determined, and immunohistochemistry revealed AST only in nerves on the oviduct. In a search for ovarian factors that regulate JH synthesis, we conditioned saline with ovaries at different developmental stages and tested the ability of this conditioned saline to affect JH synthesis by CA. Ovaries with chorionated basal oocytes released a fast-acting, reversible inhibitory factor. We report here that this factor is AST and determine its quantity and distribution in ovary and oviducts during the first vitellogenic cycle and in early embryos. Quantitative measurements were made with ELISA and localization determined by immunohistochemistry, both with monoclonal antibody to DippuAST 7. The effect of ovary-conditioned saline and ovary and embryo extract on JH synthesis in vitro and the identification of a peptide in ovary-conditioned saline with the mass of an AST by matrix assisted laser desorption/ionization–time-of-flight (MALDI–TOF) mass spectrometry confirmed the presence of ASTs in the ovary and early embryo.
2. Materials and methods 2.1. Animals Newly molted adult females were collected daily and cohorts were maintained on lab chow (Purina, St. Louis,
MO) and water at 27 °C in a 12 h/12 h light/dark cycle. At this temperature, females typically drop the spermatophore between days 5 and 6 and oviposition typically occurs by day 7 or 8. 2.2. Timing of chorion stages and oviposition A survey was made of the stages of chorion development (classification by Stay et al., 1984: A = early, B = mid, C = late, and D = complete) in basal oocytes of ovaries from 5- to 7-day females (n = 130). The proportion of females at each stage was assumed to be in proportion to the duration of each stage. These data were used to estimate the age of the female in days at the midpoint of each chorion stage and the proportion of a day during which basal oocytes were in each stage (stage: midpoint age, duration—A: 5.9, 0.9; B: 6.6, 0.6; C: 7.1, 0.3; D: 7.4, 0.3). In a subset of these day 7 females, the proportion that oviposited by day 7 was used to estimate the age of the female at oviposition (7.5 d). Duration of oviposition was determined from observations of ovipositing females (0.02 d). 2.3. Preparation of conditioned saline and tissue extract Ovaries were removed from chilled females and transferred to a dish with a small amount of Yeager’s insect saline (Buck, 1953). Adhering fat body and trachea were removed, basal oocyte length was measured with an ocular micrometer, and the stage of chorion formation was assessed. For the preparation of ovary-conditioned saline, 5–8 pairs of ovaries, rinsed in phosphate-buffered saline (PBS) (Kingan, 1989), were transferred to a 1.5 ml polypropylene tube with 0.4 ml PBS and placed on a rotary shaker for 2 h at room temperature. After centrifugation, the saline was removed from ovaries and stored at ⫺80 °C. For preparation of ovary homogenate, 3–7 pairs of ovaries, rinsed in 0.9% NaCl (saline), were homogenized in 0.4 ml saline in a mini glass tissue grinder. The homogenate and a 0.1 ml saline rinse were transferred to a 1.5 ml tube, boiled for 10 min and centrifuged for 4 min at 8000 × g. The supernatant was stored at ⫺80 °C. For preparation of oviduct homogenate, the common oviduct, along with the pair of lateral oviducts, were removed as above for ovaries and, in addition, extraneous nerves were trimmed. Five to twenty oviducts were homogenized in 0.2–0.4 ml saline. The homogenate was extracted and stored as described above. Embryo batches were expelled from females by gentle dorsal–ventral pressure on the abdomen to evert the brood sac. Three to five batches were homogenized in 0.4 ml saline. The homogenate was extracted and stored as described above.
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2.4. C18 solid-phase extraction
2.7. Immunohistochemistry
Before ovary-conditioned saline or tissue extract samples were used for ELISA, bioassay, or HPLC they were purified by solid-phase extraction to obtain AST as described previously (Woodhead et al., 1989) except that Sep-Pak light cartridges (Waters, Milford, MA) were used with 4 ml rinses and 0.8 ml eluates. Each Sep-Pak cartridge was loaded with extract of approximately 5– 10 ovary pairs, 10–40 oviducts or 5–10 embryo batches. The 17–40% acetonitrile (ACN) eluate was used for further analysis.
Females were chilled on ice before abdominal tergites and gut were removed to expose reproductive organs. Unless otherwise stated in Results, the following protocol was used. After ovaries and oviducts were carefully and quickly cleared of fat body and trachea under Yeager’s insect saline, the solution was replaced with fixative (4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.2). Following 2–5 min fixation in situ, ovaries or oviducts and ovaries were removed and fixation continued overnight at 4 °C. Fixative was removed by repeated changes of PBS (5 × 20 min) and treated for 1 h with Tris-buffered saline (0.05 M Tris, 0.35 M NaCl) with 2% Triton X-100, 2% GS and 3% ovalbumin (TBS+) before incubation with the same monoclonal antibody used for ELISA. Tissue was exposed overnight at 4 °C to antibody in cell culture supernatant diluted 1:1 with TBS+ unless otherwise stated in Results. After repeated washes in TBS with 0.2% Triton X-100, anti-AST antibody was visualized with sheep anti-mouse Cy3 (Sigma, 1:300 in TBS+ 4 h at room temperature or overnight at 4 °C). After removal of secondary antibody, nuclei were stained with SYTOX Green (Molecular Probes, Eugene, OR) 1:1000 in TBS+ for 30 min before tissue was again washed as above and placed in 0.1 M Tris. Tissues were mounted in either 90% glycerol with 0.1 M n-propyl gallate or, after dehydration in a series of increasing concentrations of ethyl alcohol, in benzyl benzoate:benzyl alcohol (2:1 v/v). For paraffin sections, oviducts were fixed in Bouin’s fixative overnight at 4 °C, washed in 70% ethanol, dehydrated in a series of increasing concentrations of ethanol, cleared in methyl benzoate, and rinsed in toluene before infiltration with Paraplast (Electron Microscopy Science, Fort Washington, PA). Ten micrometer sections were cut, mounted on slides and, after removal of embedding medium, were rehydrated and treated with antibodies as for whole mounted tissue and mounted in glycerin. Images of tissues were obtained with a confocal laser microscope (Bio-Rad MRC 1024 with krypton/argon lasers and Nikon Optophot microscope). Excitations at 488 and 568 nm were recorded separately. Merging of labels was performed using Bio-Rad or Confocal Assistant software. Controls with no primary antibody showed no staining.
2.5. ELISA The antibody, 5F10 (described previously in Stay et al., 1992), is a monoclonal antibody to Dippu-AST 7 (APSGAQRLYGFGL-NH2). Cell culture supernatant was purified using caprylic acid followed by ammonium sulfate precipitation (Harlow and Lane, 1999). The antibody was then concentrated and lyophilized. This antibody is at least 2–3 orders of magnitude more sensitive to Dippu-AST 7 compared to Dippu-ASTs 2, 5, 8 and 9 (Stay et al., 1992) and Dippu-ASTs 4 and 11 (Woodhead, unpublished). AST quantities are expressed as Dippu-AST 7 equivalents. The ELISA procedure is adapted from Kingan’s method (1989). All solutions for ELISA were made with PBS, unless stated otherwise. At room temperature, titer plates (Corning Easywash, Corning, NY) were coated with AST coupled to ovalbumin (50 µl/well) for 3 h and then rinsed once with 0.05% Tween 20, blocked for 30 min with 1% goat serum (GS) (Sigma, St. Louis, MO) and rinsed again with 0.05% Tween 20 (100 µl/well). Sample fractions from solid-phase extraction and standards (25–2500 fmol Dippu-AST 7/well) were dried and then resuspended in 1% GS to which an equal volume of antibody solution, diluted 1:1000 with 1% GS, was added. Solutions were added in duplicate to the plate (100 µl/well) and stored at 4 °C overnight. After the plate was rinsed twice with 0.05% Tween 20, goat antimouse alkaline phosphatase (Promega, Madison, WI; 1:3000 in 1% GS) was added to the wells and incubated at room temperature for 2 h. The plate was rinsed four times with 0.05% Tween 20. The alkaline phosphatase was reacted with 0.4 mg phosphatase substrate (Sigma 104)/ml 10% diethanolamine. The absorbance of the reaction product was measured at 405 nm using a Titertek Multiscan MCC/340 plate reader. 2.6. Protein measurements The quantity of protein in oviducts was measured using Bradford’s (1976) method. Two to four oviducts were homogenized in 0.2 ml 1 M NaOH and, after centrifugation, the supernatant was used for protein analysis.
2.8. Delipidation and bioassay of ovary and embryo extracts Although ELISA AST determinations were made from the 17–40% ACN fraction from solid-phase extraction, lipid interfered with the in vitro assay for JH synthesis by CA. Therefore for bioassays, extract was delipidated before the C18 solid-phase extraction step by vortexing the sample with an equal volume of ice-cold
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n-hexane that had been saturated with 0.9% NaCl. After centrifugation at 4 °C, the hexane was removed, this procedure was repeated and the aqueous moiety was retained for solid-phase extraction. The sensitivity of CA to conditioned saline and tissue extracts was tested in the sequential in vitro JH assay, described in Stay et al. (1991), except that the labeled (60 methionine was l-[methyl-14C]methionine mCi/mmol, Amersham, Piscataway, NJ). The test CA were from 2-day virgin females (n = 4–6). Briefly, right and left glands were equilibrated at room temperature for 2 h in TC199 (GIBCO, Grand Island, NY) before rates of JH synthesis in untreated radioactive medium were determined in a 2 h incubation at 27 °C. Then glands were transferred to radioactive medium with conditioned saline/tissue extract (treated) or without (controls) for another 2 h. Treated and control glands were members of a pair. For statistical analysis, JH synthesis rates of right and left glands were compared for the first 2 h (untreated, untreated) and the second 2 h (treated, untreated) for ovary and embryo extract. For ovary-conditioned saline, rates of right (treated) and left (untreated) glands in a 2 h assay were compared. Percent inhibition was determined as described previously (Stay et al., 1991).
3. Results 3.1. Quantity of AST in ovaries and young embryos The amount of AST extracted from ovary homogenate during 10 d of the first reproductive cycle is compared to the amount of AST released by ovaries in vitro for 2 h (Fig. 1B) and shown in relation to mean basal oocyte length of the ovaries from which homogenates were prepared (Fig. 1A). Basal oocytes become vitellogenic between days 2 and 3 and continue vitellogenesis until days 5–6 when the spaces between follicle cells close and follicle cells begin to produce an eggshell (chorion). The AST content of ovary homogenates gradually increased from day 2 to early (A) chorion stage, then
2.9. HPLC and mass spectrometry The 17–40% ACN eluate of delipidated embryo extract and, alternatively, ovary-conditioned saline from C18 Sep-Pak was reduced in volume and applied to a C18 column (Waters). A gradient of 10–35% ACN over 50 min was used with 0.115% TFA as solvent A and ACN with 0.1% TFA as solvent B. Half minute fractions were collected, and aliquots of fractions with elution times of synthetic ASTs were tested by ELISA and bioassay. Several fractions were combined using elution times of synthetic ASTs as a guide and were applied to a C8 column (Jones Chromatography, Lakewood, CO). A gradient of 10–40% ACN over 45 min was used with the same solvents as for C18 HPLC. Half minute fractions were collected, and aliquots of fractions with elution times of Dippu-ASTs 7–9 were submitted for MALDI– TOF mass spectrometry analysis (Molecular Analysis Facility, University of Iowa). MALDI–TOF mass spectra were obtained using a Bruker Biflex III system (Bruker Daltonics, Billerica, MA) with α-cyano-4-hydroxycinnamic acid (Sigma) as matrix and angiotensin II and ACTH as calibrants. 2.10. Statistics Student’s t-test was used to compare AST and protein content data. For JH bioassays, rates of synthesis were compared using the Mann–Whitney U-test.
Fig. 1. AST contained and released by the ovary during the first vitellogenic cycle in relation to oocyte growth. (A) The mean length of basal oocytes in ovaries used for homogenate preparation in B. Each point is the mean of 27–85 measurements, SEM is equal to or less than 0.01 mm. The second value on day 5 represents measurements taken after females dropped the spermatophore. Choriogenesis (Chor) begins on day 6 and continues through day 7. The arrow indicates oviposition (Ovip). (B) AST released into saline by ovaries (—ο—) and AST content of ovary homogenates (—쎲—). Stages of choriogenesis: A = early, B = mid, C = late and D = complete. Sample sizes for AST released are: day 3, stages A and B, n = 1; stages C and D, n = 2 and 3, respectively. For ovary homogenate, each point represents a mean value (±SEM) of 3–7 samples.
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rapidly increased between the early (A) and late (C) chorion stages. In contrast, AST released by ovaries increased only slightly from day 3 to the mid (B) chorion stage, then rapidly increased between the mid (B) and late (C, D) chorion stages. The ovaries at these late chorion stages released about half their AST. After oviposition, the AST content of ovaries, now containing small non-vitellogenic basal oocytes, returned to a low level similar to that of previtellogenic ovaries on day 2. The basal oocytes (usually 12) are fertilized just after they leave the common oviduct and are aligned in two parallel rows, partially covered by an ootheca and then retracted into the brood sac where embryogenesis takes place. Transfer of nutrient to embryo from mother does not begin until after dorsal closure, on day 12–13 (Stay and Coop, 1973). Just after ovulation, AST content of eggs was 72.6 ± 11.6 fmol / batch (Fig. 2), a loss of about 40% of their AST (Fig. 2) compared to D stage ovaries (just before ovulation, Fig. 1) in which AST occurs in basal oocytes (see below Section 3.3). One day after oviposition AST content of embryos was 122.7 ± 6.6 fmol / batch, indicating rapid synthesis of AST by embryos in the day after oviposition (Fig. 2). However, between days 4 and 8, AST content of embryos decreased to about 50 fmol/batch and remained at this level on day 12. 3.2. Quantity of AST and protein in oviducts AST content of oviducts (i.e., both common and lateral) at specific stages of the first egg development cycle was also determined (Fig. 3). On day 2 at the start of vitellogenesis the AST content of oviducts was about half that of oviducts from day 7 females with completely chorionated basal oocytes (4.9 ± 0.8 vs. 8.4 ± 1.4 fmol / female, respectively). Over a shorter time span, from pre- to immediately post-ovulation on day 7, about
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Fig. 3. AST and protein content in oviduct homogenates. Oviducts were dissected on day 7 before ovulation (day 7 pre-ovul), immediately after ovulation (day 7.5 post-ovul), and 1 d post-ovulation (day 8.5 post-ovip). AST content (black bars) of post-ovulation oviducts was significantly greater than AST content of pre-ovulation oviducts (P ⬍ 0.02) and 1 day post-oviposition (P ⬍ 0.01); each bar represents the mean (±SEM) of 3–4 samples. Protein content (gray bars) of day 7 pre-ovulation oviducts was significantly greater than day 2 oviducts (P ⬍ 0.05) and day 7.5 post-ovulation oviducts (P ⬍ 0.01); each bar represents the mean (±SEM) of three samples.
half a day, AST content of oviducts increased to 18.4 ± 2.5 fmol / female, a significantly greater AST content than that of pre-ovulation oviducts (P ⬍ 0.02). One day after the batch of eggs was deposited in the brood sac, the AST content of the oviducts had decreased to the pre-ovulation level (8.9 ± 1.2 fmol / female). It should be noted that there is far less AST in day 7 oviducts (10– 20 fmol/female) than in day 7 (late chorion stage) ovaries (120–130 fmol/pr, Fig. 1B). The oviducts visibly increase in diameter as vitellogenesis progresses (see Fig. 7A and C). To determine whether this increase represented growth, the protein content of oviducts on day 2 and day 7 pre- and postovulation was determined (Fig. 3). Indeed, oviduct protein significantly increased from day 2 to day 7 pre-ovulation (from 42.6 ± 1.9 to 73.1 ± 2.2 µg / female, respectively, P ⬍ 0.01). Immediately after ovulation (day 7.5), protein in the oviducts significantly decreased to a level similar to that of day 2 oviducts (43.9 ± 8.8 µg / female, P ⬍ 0.05) and remained low 1 d post-oviposition (34.2 ± 3.1 µg / female). 3.3. Immunohistochemical localization of AST in ovary, oviduct and embryo
Fig. 2. AST content in embryo homogenates. Each point represents the mean (±SEM) of 3–6 samples.
The paired ovaries of D. punctata usually consist of six ovarioles each held together by a thin tracheated sheath. The ovarioles contain a series of follicles, oocytes surrounded by follicle cells, and only the basal follicles undergo vitellogenesis in each reproductive cycle (Fig. 4A and C). At the completion of vitellogen-
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Fig. 4. Diagrams of ovary and oviducts. (A) Innervation by a branch of ventral nerve 7 (VN7) of the common (CO) and lateral (LO) oviduct and their muscle attachments (M). Spermatheca (S) and accessory gland (AG) adhere closely to lateral oviduct (shown only on a portion of one lateral oviduct). The muscle from the copulatory bursa (Bu) to the ventral body wall is shown cut away where it extends over the lateral oviduct. (B) Median longitudinal section of a penultimate (PO) and basal oocyte (BO) shows basal oocyte surrounded by the chorion (Ch) with anterior micropyle (arrow), under the follicle cell layer (FCL). The pedicel (Pe) is at the base of the ovariole. (C) Cross-section of an oviduct. Muscle layers (M) surround a folded epithelium (E) covered by a thin cuticle with barbs projecting into the lumen (drawn from electron micrographs of lateral oviduct of a 4-day mated female).
esis the chorion secreted by the follicle has an anterior opening, the micropyle, through which sperm enter the oocyte as it leaves the common oviduct. Examination of follicles at several stages in the first vitellogenic cycle revealed increasing amounts of AST-immunoreactivity in the basal oocytes and none between or in the follicle cells (Fig. 5A–C). In mid- and late stages of chorion formation, immunoreactivity in intact basal oocytes is almost undetectable. This late stage is when most AST is detected by ELISA and when most AST is released from ovaries incubated in vitro (Fig. 1B). In ovaries incubated in vitro before fixation, immunoreactivity was conspicuous at the micropylar end of the basal oocytes (Fig. 5D). When chorionated oocytes were cut after fixation and exposed for several days to antibody, immunoreactivity was obvious in the oocyte cytoplasm (Fig. 5E). Oocytes removed from the female as they left the genital chamber showed immunoreactivity at the micropyle and in a few places on the surface of the chorion (Fig. 5F). Occurrence of AST at the micropyle and on the surface of the chorion suggests that AST is released from the micropyle as oocytes travel down the oviducts. To investigate this possibility ovaries and oviducts of females in the process of ovulation were exposed and fixed in situ. Fig. 6A shows one oocyte leaving the follicle cell layer and three oocytes in the lateral oviducts. The oocyte partially separated from the follicle cells showed immunoreactivity not only at the micropyle but also in the tiny lumen of the follicle cell sheath (Fig.
6B). Immunoreactivity was also evident in the lumen of the lateral oviduct below the ovary in which all basal oocytes had been ovulated (Fig. 6C). The possibility that some of the immunoreactivity was also in epithelial cells of the oviduct was investigated in confocal sections of oviducts from females prior to ovulation. Oviducts from 2-day females, with oocytes at the start of vitellogenesis, showed strong immunoreactivity in nerves on the surface of the oviducts (Fig. 7A) but, at low magnification, little immunoreactivity was evident in the epithelial cells of the lateral or common oviducts, whereas some immunoreactivity was evident at higher magnification (Fig. 7B). In contrast, in oviducts from 7-day females, in addition to strong immunoreactivity in nerves (Fig. 7C), in lowmagnification confocal sections immunoreactivity was obvious at the level of epithelial cells (Fig. 7D). In crosssection, the distinction between the epithelial and muscle layers is obvious (Fig. 4C). Cross-sections of oviducts from females in late vitellogenesis confirmed that AST occurs in epithelial cells. Confocal images of such sections showed immunostaining at the apex of the epithelial cells and in the space between the cells and the cuticular lining (Fig. 7E and F). Thus the amount of AST in pre-ovulation oviducts is a combination of AST in the epithelial cells and in the nerves to the muscle cell layer of the oviduct. AST in embryos occurs in yolk cells (Fig. 7G) The nerves serving the muscle layers of the oviducts are branches of ventral nerve 7 (VN7) from the last
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Fig. 5. Fluorescent AST-immunoreactivity in basal oocytes and an ovulated egg. (A–C) show increasing reactivity in vitellogenic oocytes from 2-, 4-, and 6-day females, respectively. Inset in C shows follicle cells, with green nuclei, devoid of AST-immunoreactivity (red) which is present only in the oocyte cytoplasm. (D) Portion of an ovary with completely chorionated oocytes fixed after incubation in buffered saline for 2 h shows strong immunoreactivity only at the micropylar ends of the basal oocytes. (E) Basal follicle with chorionated oocyte that was cut open before exposure to antibody reveals immunoreactivity in oocyte cytoplasm. (F) Ovulated egg fixed as it emerged from the genital chamber shows immunoreactivity at the micropyle (top) and a small amount on the surface of the intact chorion. Scale bars = µm.
abdominal ganglion (Fig. 4A). A variable distribution of nerves occurs in oviducts even though great care is exercised in their removal from the female (e.g., Fig. 7A and B). However, remnants of nerve seen in whole mounts most frequently occur on the common oviduct and near the ovary on the lateral oviducts. In two sets of experiments, oviducts (triplicates of each of three stages) with attached nerves were exposed to a more dilute antibody (1:40) in order to distinguish potential differences in immunoreactivity that might be stage specific. Little obvious difference is seen in intensity of immunoreactivity at different stages of the first egg development cycle. Fig. 8 shows similar immunoreactivity in low-magnification confocal images of VN7 as it contacts the lateral oviducts from 2-day females, and lateral oviducts from 7-day females with chorionated oocytes, and after a batch of eggs was oviposited. At high magnification the immunoreactive axons of the nerve as it contacts the oviduct show varicosities that appear to be on the surface of the nerve where they could be acting as neurohemal release sites (Fig. 8D). 3.4. Effect of ovary-conditioned saline and ovary and embryo extracts on JH synthesis in vitro Inhibition of JH synthesis by CA in vitro confirmed the presence of allatostatic material in ovaries and
embryos. Bioassay with saline conditioned by C and D stage chorionated basal oocytes did not inhibit JH synthesis rates significantly at 1 ovary equivalent/CA (10.5 ± 8.6%) however, at 5 ovary equivalents/CA inhibition was 45.7 ± 10.0%; rates were significantly lower than rates of untreated controls (P = 0.01). Treatment with higher doses did not increase inhibition presumably because of interfering material in the extract. Therefore extracts of ovaries with C and D stage chorionated basal oocytes and 1-day embryos were further purified by delipidation and tested on CA in vitro. Based on ELISA quantification, CA were treated with the equivalent of 10⫺8 M AST (8.9 ovary pair equivalents and 7.6 embryo batch equivalents/CA). JH synthesis rates of CA treated with ovary and embryo extract were significantly inhibited compared to untreated controls (62.8 ± 3.1%, P ⬍ 0.05; 52.4 ± 3.2%, P ⬍ 0.01, respectively). For all bioassays, there was no significant difference in rates of JH synthesis between groups of untreated CA in the first 2 h incubation. Because batches of embryos retain much ovarian AST and are more easily obtained than late-stage ovaries, extract of 1-day embryos (190 batches) was purified by HPLC on a C18 column. Material with the retention time of Dippu-AST 2 (AYSYVSEYKRLPVYNFGL-NH2) and material from two other HPLC fractions not expected to contain AST were tested. Inhibition by these
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Fig. 6. Ovaries and oviducts fixed in the process of ovulation. (A) Only one basal oocyte remains partially within the follicle cell layer in the left ovary (long arrow). Box on the left is shown at higher magnification in B. Three oocytes are in the distended lateral oviducts (short arrows). Nerves on the surface of the common oviduct (CO) show strong immunoreactivity. Box on the right is shown at higher magnification in C. Note: some ovarioles were removed to facilitate the mounting of tissue. (B) Median section of anterior portion of a follicle with oocyte (O) departing the follicle cell layer (FC). Immunoreactivity (red) appears at the micropyle of the chorion (long arrow) and in the narrow lumen of the follicle cells (short arrows). (C) Median confocal section of oviduct after passage of six oocytes. AST-immunoreactivity (red) occurs in the lumen and in oviduct epithelial cells. Nuclei are green. Scale bars = µm.
other fractions was 13 ± 6% and 5 ± 7%, whereas the Dippu-AST 2 fraction inhibited JH synthesis rates 70.0 ± 6.0% when CA were treated with the equivalent of 10⫺8 M Dippu-AST 2, estimated from ELISA measurements for Dippu-AST 7. This assumes that all the ASTs are produced in approximately equal amounts (see Discussion). 3.5. MALDI–TOF mass spectrometry MALDI–TOF mass spectrometry analysis of supernatant of boiled saline extract of D. punctata brains identified the masses of eight of the 13 peptides of the Dippu-AST family (data not shown). However, no masses of any ASTs were detected in similarly prepared ovary-conditioned saline or extract of ovaries. This likely was due to the approximately 100× greater quantity of soluble proteins relative to ASTs in ovarian tissue compared to brain tissue, and also the greater quantity of lipid in the ovarian extract. Therefore, fractions from the C18 HPLC separation of conditioned saline from 400 ovary pairs expected to contain Dippu-ASTs 7–9 were
further purified on a C8 column. MALDI–TOF mass spectrometry analysis of the fractions expected to contain Dippu-ASTs 7 and 9 identified an (M+H)+ ion observed at M/Z 1335.51, which corresponds to the monoisotopic mass of Dippu-AST 7 (1334.69). Masses of Dippu-ASTs 8 and 9 were not found. This was likely due to loss of AST material during HPLC purification steps and the inability to detect AST masses when dilute solutions were concentrated for MALDI–TOF analysis. 4. Discussion This study of the quantity and localization of AST in ovaries, oviducts, and embryos in relation to the stages of vitellogenesis and choriogenesis and embryonic development in D. punctata strongly suggests new roles for this multifunctional -Phe-Gly-Leu-amide family of peptides. It is likely that AST in oocytes, epithelial cells of the oviduct, and nerves to the oviduct muscles, functions in the process of ovulation, and after ovulation, AST in the fertilized egg (embryo) functions in yolk utilization.
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Fig. 7. AST-immunoreactivity in pre-ovulation oviducts and a young embryo. (A) Common and lateral oviducts of a 2-day female with basal oocytes in early vitellogenesis. Note immunoreactivity in nerves on the surface of common oviduct (CO) and lateral oviducts near the ovary (arrows). Boxed area is shown at higher magnification in B. (B) Portion of a common oviduct in a confocal section at the level of epithelial cells shows AST-immunoreactivity (small red dots); larger spots are likely varicosities of nerves. Nuclei are pale green; barbs on cuticular lining are bright green (photographed with 60× objective). (C) Common and lateral oviducts of a 7-day female with completely chorionated basal oocytes. Note distribution of nerves as in A. Boxed area is shown at higher magnification in D. (D) AST-immunoreactivity (red) in common oviduct in a confocal section at the level of epithelial cells (photographed with 20× objective); barbs not visible at this magnification. (E) Cross-section, cut in paraffin, of common oviduct of a female at a late stage of vitellogenesis shows folded epithelium (e) and surrounding muscle cell layer (m). (F) Portion of the cross-section in E at higher magnification shows AST-immunoreactivity in apex of epithelial cells and in space between cells and cuticular lining. Barbs (short arrows) on the cuticle layer (long arrow) are bright green. (G) Part of a 9-day, pre-dorsal closure, embryo (Emb). Yolk cells (YC) are strongly AST-immunoreactive; nuclei are green. Scale bars = µm.
4.1. Function of ovarian AST during ovulation We have confirmed, for D. punctata, Witek and Hoffmann’s (2001) observation in cricket ovaries that AST occurs in oocytes and not in follicle cells. Our study shows the occurrence of AST principally in the basal oocytes and an increase in quantity closely correlated with stage of development. Although AST increases from the start of vitellogenesis, the greatest increase is during choriogenesis. At this stage, when ovaries were
incubated in saline, AST-immunoreactive material accumulated at the micropyle of the chorion and was found in the conditioned saline. That the release of AST also occurs physiologically was demonstrated by finding AST-immunoreactivity at the micropyle, between follicle cell layers and in the lumen of the oviduct in tissue fixed in a female in the process of ovulation. Further evidence to support this hypothesis is the increase in amount of AST in oviducts immediately after ovulation on day 7; and that this increase is correlated with a 40%
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Fig. 8. AST-immunoreactivity in a piece of ventral nerve 7 as it contacts the lateral oviduct in a 2-day female (A); a 7-day female with chorionated basal oocytes (B); and a 7-day female a few minutes after ovulation (C). High magnification confocal image of a portion of the nerve in A shows nuclei (green) of nerve sheath cells and an AST-immunoreactive axon (red) on the surface of the nerve. These preparations were exposed to antiDippu-AST 7 diluted 1:40 in order to detect possible differences in intensity of reactivity with stage of vitellogenesis. Scale bars = µm; A–C are the same magnification.
reduction in AST in the eggs measured immediately after ovulation. Thus the AST in oviducts could be, at least in part, that released by the oocytes from the micropyle. Indeed AST was observed at the micropyle and on the surface of newly ovulated eggs. The amount of AST in oocytes and the timing of its release indicate that oocyte AST is unlikely to function as a signal for the decline in JH synthesis during vitellogenesis, as this occurs before choriogenesis begins (Rankin and Stay, 1984; Pratt et al., 1990). Thus, if release of AST by oocytes were to have a regulatory role in JH production it would be to maintain low rates of synthesis at this time. However, this is unlikely since the AST released by oocytes during ovulation, if it reached the hemolymph, would not increase the AST concentration sufficiently to inhibit CA (Stay et al., 1994). Release of AST from nerves within the CA more effectively inhibits JH production than elevated AST in the hemolymph (Stay et al., 1994). Although the fraction of the AST in the CA that is within nerves cannot be determined by ELISA, it is interesting to note that extract of a pair of CA contains 0.3 pmol equivalents of Dippu-AST 7 (Lloyd et al., 2000) whereas the maximum quantity contained in a pair of ovaries is 0.12 pmol equivalents. 4.2. Functions of AST in oviducts Quantitative measurements of AST in extracts of oviducts cannot distinguish between material in nerves, epithelial cells or in the lumen. However, since the amount of AST-immunoreactivity seen in whole mounts of nerves to the oviduct does not appear to be different on day 2 or day 7 before or after ovulation, we presume that changes in quantity of AST in oviducts is due to peptides in the epithelial cells and/or in the lumen of the oviducts. The presence of AST-immunoreactivity in epithelial cells of pre-ovulation oviducts, as seen in
whole mount images at the level of the epithelial cells and in cross-sections cut in paraffin, clearly shows that, in addition to AST in nerves and AST that may be released by the oocytes during ovulation, AST in epithelial cells contributes to the quantity of AST extractable from oviducts both before and after ovulation. Comparison of the quantity of AST in oviducts of day 2 and day 7 pre-ovulation stages suggests that AST in oviduct cells increases during vitellogenesis as does the AST mRNA quantified by Garside et al. (2002). How the AST in the cells of the oviduct functions is yet to be determined. One hypothesis is that it facilitates the secretion of a lubricating material during ovulation. The sharp decrease in protein content of the oviducts from pre-ovulation to immediately post-ovulation on day 7 would indicate that a substance is released during ovulation. Electron micrographs of lateral oviducts reveal many mitochondria and large areas of glycogen in epithelial cells (Stay, unpublished). The glycogen could be a source of carbohydrate for a mucoprotein secretion. The abundance of mitochondria suggests that AST could be acting to regulate the ion transport necessary for the movement of sufficient water to facilitate the flow of the material out of the cells during ovulation. Further study will be required to test this hypothesis. The role of AST in the nerves to the oviduct is also yet to be determined. AST-immunoreactivity is clearly evident in nerves that innervate the oviduct muscle layer, the anchoring muscles of the common and lateral oviducts, and the muscles that pass over the lateral oviducts. Therefore, it seems likely that AST functions to regulate activity of these muscles in the complex process of movement of the oocytes along and out of the oviducts. Although no effect of AST on oviduct muscle in vitro has yet been demonstrated in D. punctata (Lange et al., 1995; Garside et al., 2002) or L. migratoria (Lange et al., 1993), AST is a strong inhibitor of proctolin-induced muscle contraction in the hindgut of D. punctata (Lange
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et al., 1993). Perhaps in the oviducts AST modulates muscle activity in conjunction with another (unidentified) modulator or neurotransmitter. It is also conceivable that the sensitivity of oviduct muscle to AST is stage specific and the sensitive stage has not been examined. The varicosities seen on the surface of the branch of VN7 leading to the oviducts suggest that these axons also release into the hemolymph and may affect muscles of the reproductive organs humorally. 4.3. Function of AST in early embryos Of the AST accumulated in the basal oocytes at the end of choriogenesis, 40% is lost during ovulation, but one day later the amount of AST in the embryos has increased to the level of pre-oviposition ovaries. This suggests that at the start of development, embryos supplied with a large store of AST by the mother are synthesizing AST well before the nervous system begins to express these peptides at about day 15, three days postdorsal closure (Stay, unpublished). In the embryo the yolk of the oocyte is incorporated into yolk cells and these are clearly AST-immunoreactive. This suggests that AST is modulating activity of nutrient digestion, as has been demonstrated in midgut of adult D. punctata (Fuse´ et al., 1999). Most of the yolk is depleted shortly after the embryo begins to drink the nutrient milk produced by the brood sac (Stay and Coop, 1973) and the amount of AST in embryos has decreased markedly by dorsal closure (day 12). 4.4. The identity of AST-immunoreactive material The quantity and localization of AST in the ovaries, oviducts and embryos is based on immunoreactivity to an antibody against Dippu-AST 7. The presence of Dippu-AST 7 was confirmed by MALDI-TOF spectrometry on ovary-conditioned saline. The ability of ovary-conditioned saline, and ovary and embryo extract to inhibit JH synthesis by CA in vitro is further evidence that the material is allatostatin. Not surprisingly, ASTimmunoreactivity occurs in vitellogenic oocytes of other cockroaches: Periplaneta americana, Leucophaea maderae and Schultesia lampyridiformis, and in chorionated blowfly oocytes (Stay, personal observation). It will be of interest to determine whether locust ovarian allatostatic activity is due to a -F-G-L-amide peptide and whether it also occurs in oviduct cells. Inhibition of JH synthesis by a fraction of embryo extract that eluted from a C18 column at the time of Dippu-AST 2 indicates that ASTs in addition to DippuAST 7 are present in extract of embryos. It is very likely that all members of the Dippu-AST family are produced, as multiple members have been isolated in D. punctata from brain (Woodhead et al., 1989), CA (Stay and Woodhead, 1993), hemolymph (Woodhead et al., 1993)
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and antennal heart muscle (Woodhead et al., 1992). Also multiple members of the P. americana AST family were identified from antennal heart muscle of that species (Predel et al., 1999). The quantitative RT–PCR analysis of mRNA for Dippu-ASTs in ovaries and oviducts during the first vitellogenic cycle in D. punctata (Garside et al., 2002) is convincing evidence that AST detected immunologically is synthesized in these tissues. The mRNA results, expressed as the amount of AST mRNA/total RNA, give the impression that the amount and increase of AST message during the vitellogenic cycle are more prominent in oviducts than in ovaries. However, if total RNA in vitellogenic ovaries is much greater than that in oviducts this would explain the apparent lower quantity of AST mRNA in ovaries. By conditioning saline with ovaries we hoped to obtain factors involved in the coordination of increase and decrease in JH production with ovarian development. We have found a factor, AST, that can modulate JH synthesis. However, because of its appearance late in the first egg development cycle, AST in the mature oocytes and oviducts most likely does not function to regulate JH synthesis, but more likely has totally different functions, such as facilitation of ovulation and utilization of yolk in the embryo. Acknowledgements The authors would like to thank John Hass for his help with tissue preparations and Shelley Plattner for his technical assistance with figures. This work is supported by NIH grant AI 15230. References Applebaum, S.W., Hirsch, J., El-Hadi, F.A., Moshitzky, P., 1990. Trophic control of juvenile hormone synthesis and release in locusts. In: Epple, A., Scanes, C.G., Stetson, M.H. (Eds.), Progress in Comparative Endocrinology. Wiley-Liss, New York, pp. 186– 192. Bendena, W.G., Donly, B.C., Tobe, S.S., 1999. Allatostatins: a growing family of neuropeptides with structural and functional diversity. Annals of the New York Academy of Sciences 897, 311–329. Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72, 248–254. Buck, J.B., 1953. Physical properties and chemical composition of insect blood. In: Roeder, K.D. (Ed.), Insect Physiology. Wiley and Sons, New York, pp. 147–190. Couillaud, F., Girardie, A., 1984. Ovariectomy and juvenile hormone JH-3 biosynthesis in African locust Locusta migratoria. Comptes Rendus des Seances de L’Academie des Sciences Serie III Sciences de la Vie 298, 157–162. Donly, B.C., Ding, Q., Tobe, S.S., Bendena, W.G., 1993. Molecular cloning of the gene for the allatostatin family of neuropeptides from the cockroach Diploptera punctata. Proceedings of the National Academy of Sciences, USA 90, 8807–8811.
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A.P. Woodhead et al. / Journal of Insect Physiology 49 (2003) 1103–1114
Ferenz, H.-J., Aden, E., 1993. Ovarian control of juvenile hormone biosynthesis in Locusta migratoria (Orthoptera: Acrididae). Entomologia Generalis 18, 9–17. Fuse´ , M., Zhang, J.R., Partridge, E., Nachman, R.J., Orchard, I., Bendena, W.G., Tobe, S.S., 1999. Effects of an allatostatin and a myosuppressin on midgut carbohydrate enzyme activity in the cockroach Diploptera punctata. Peptides 20, 1285–1293. Ga¨ de, G., 2002. Allatoregulatory peptides—molecules with multiple functions. Invertebrate Reproduction and Development 41, 127– 135. Gadot, M., Rafaeli, A., Applebaum, S.W., 1987. Partial purification and characterization of locust allatotropin I. Archives of Insect Biochemistry and Physiology 4, 213–223. Garside, C.S., Koladich, P.M., Bendena, W.G., Tobe, S.S., 2002. Expression of allatostatin in the oviducts of the cockroach Diploptera punctata. Insect Biochemistry and Molecular Biology 32, 1089–1099. Harlow, E., Lane, D., 1999. Using Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Hoffmann, K.H., Lorenz, M.W., Klein, P.M., 1996. Ovarian control of juvenile hormone-III biosynthesis in the cricket species Gryllus bimaculatus (Ensifera: Gryllidae). Entomologia Generalis 20, 157–167. Kingan, T.G., 1989. A competitive enzyme-linked immunosorbent assay: applications in the assay of peptides, steroids, and cyclic nucleotides. Analytical Biochemistry 183, 283–289. Lange, A.B., Chan, K.K., Stay, B., 1993. Effect of allatostatin and proctolin on antennal pulsatile organ and hindgut muscle in the cockroach, Diploptera punctata. Archives of Insect Biochemistry and Physiology 24, 79–92. Lange, A.B., Bendena, W.G., Tobe, S.S., 1995. The effect of the thirteen Dip-allatostatins on myogenic and induced contractions of the cockroach (Diploptera punctata) hindgut. Journal of Insect Physiology 41, 581–588. Lloyd, G.T., Woodhead, A.P., Stay, B., 2000. Release of neurosecretory granules within the corpus allatum in relation to the regulation of juvenile hormone synthesis in Diploptera punctata. Insect Biochemistry and Molecular Biology 30, 739–746. Pratt, G.E., Farnsworth, D.E., Siegel, N.R., Fok, K.F., Feyereisen, R., 1989. Identification of an allatostatin from adult Diploptera punctata. Biochemical and Biophysical Research Communications 163, 1243–1247. Pratt, G.E., Farnsworth, D.E., Feyereisen, R., 1990. Changes in the sensitivity of adult cockroach corpora allata to a brain allatostatin. Molecular and Cellular Endocrinology 70, 185–195. Pratt, G.E., Farnsworth, D.E., Fok, K.F., Siegel, N.R., McCormack, A.L., Shabanowitz, J., Hunt, D.F., Feyereisen, R., 1991. Identity of a second type of allatostatin from cockroach brains: an octadecapeptide amide with a tyrosine-rich address sequence. Proceedings of the National Academy of Sciences, USA 88, 2412–2416. Predel, R., Kellner, R., Rapus, J., Ga¨ de, G., 1999. Allatostatins from the retrocerebral complex and antennal pulsatile organ of the American cockroach: structural elucidation aided by matrix-assisted laser desorption/ionization-time-of-flight mass spectrometry. Regulatory Peptides 82, 81–89.
Rankin, S.M., Stay, B., 1984. The changing effect of the ovary on rates of juvenile hormone synthesis in Diploptera punctata. General and Comparative Endocrinology 54, 382–388. Rankin, S.M., Stay, B., 1985. Ovarian inhibition of juvenile hormone synthesis in the viviparous cockroach, Diploptera punctata. General and Comparative Endocrinology 59, 230–237. Stay, B., 2000. A review of the role of neurosecretion in the control of juvenile hormone synthesis: a tribute to Berta Scharrer. Insect Biochemistry and Molecular Biology 30, 653–662. Stay, B., Coop, A., 1973. Developmental stages and chemical composition in embryos of the cockroach, Diploptera punctata, with observations on the effect of diet. Journal of Insect Physiology 19, 147–171. Stay, B., Woodhead, A.P., 1993. Neuropeptide regulators of insect corpora allata. American Zoology 33, 357–364. Stay, B., Tobe, S.S., Mundall, E.C., Rankin, S., 1983. Ovarian stimulation of juvenile hormone biosynthesis in the viviparous cockroach, Diploptera punctata. General and Comparative Endocrinology 52, 341–349. Stay, B., Ostedgaard, L.S., Tobe, S.S., Strambi, A., Spaziani, E., 1984. Ovarian and haemolymph titres of ecdysteroid during the gonadotrophic cycle in Diploptera punctata. Journal of Insect Physiology 30, 643–651. Stay, B., Joshi, S., Woodhead, A.P., 1991. Sensitivity to allatostatins of corpora allata from larval and adult female Diploptera punctata. Journal of Insect Physiology 37, 63–70. Stay, B., Chan, K.K., Woodhead, A.P., 1992. Allatostatin-immunoreactive neurons projecting to the corpora allata of adult Diploptera punctata. Cell and Tissue Research 270, 15–23. Stay, B., Sereg Bachmann, J.A., Stoltzman, C.A., Fairbairn, S.E., Yu, C.G., Tobe, S.S., 1994. Factors affecting allatostatin release in a cockroach (Diploptera punctata): nerve section, juvenile hormone analog and ovary. Journal of Insect Physiology 40, 365–372. Weaver, R.J., Edwards, J.P., Bendena, W.G., Tobe, S.S., 1998. Structures, functions and occurrence of insect allatostatic peptides. In: Coast, G.M., Webster, S.G. (Eds.), Recent Advances in Arthropod Endocrinology. Cambridge University Press, Cambridge, pp. 3–32. Wennauer, R., Kassel, L., Hoffmann, K.H., 1989. The effects of juvenile hormone, 20-hydroxyecdysone, precocene II, and ovariectomy on the activity of the corpora allata (in vitro) in adult female Gryllus bimaculatus. Journal of Insect Physiology 35, 299–304. Witek, G., Hoffmann, K.H., 2001. Immunological evidence for FGLamide- and W2W9-allatostatins in the ovary of Gryllus bimaculatus (Ensifera, Gryllidae). Physiological Entomology 26, 49–57. Woodhead, A.P., Stay, B., Seidel, S.L., Khan, M.A., Tobe, S.S., 1989. Primary structure of four allatostatins: neuropeptide inhibitors of juvenile hormone synthesis. Proceedings of the National Academy of Sciences, USA 86, 5997–6001. Woodhead, A.P., Stoltzman, C.A., Stay, B., 1992. Allatostatins in the nerves of the antennal pulsatile organ muscle of the cockroach Diploptera punctata. Archives of Insect Biochemistry and Physiology 20, 253–263. Woodhead, A.P., Asano, W.Y., Stay, B., 1993. Allatostatins in the haemolymph of Diploptera punctata and their effect in vivo. Journal of Insect Physiology 39, 1001–1005.