Diversity of detoxification pathways of ingested ecdysteroids among phytophagous insects.код для вставкиСкачать
Archives of Insect Biochemistry and Physiology 65:65–73 (2007) Diversity of Detoxification Pathways of Ingested Ecdysteroids Among Phytophagous Insects Kacem Rharrabe,1,2 Salah Alla,3 Annick Maria,2 Fouad Sayah,1 and René Lafont2* The metabolic pathways of ingested ecdysteroids have been investigated in three insect species, the aphid Myzus persicae and two Lepidoptera, Plodia interpunctella and Ostrinia nubilalis. M. persicae produces mainly a 22-glucoside conjugate, whereas P. interpunctella eliminates a mixture of 20E and its 3-oxo and 3-epi derivatives, both in free form and as conjugates with various fatty acids. O. nubilalis only produces fatty acyl ester conjugates. These data point out the great diversity of detoxification mechanisms used by phytophagous insects in order to overcome the potential harmful effects of ecdysteroids present in their food. Arch. Insect Biochem. Physiol. 65:65–73, 2007. © 2007 Wiley-Liss, Inc. KEYWORDS: ecdysteroid metabolism; ecdysone; 20-hydroxyecdysone; detoxification; insect INTRODUCTION Analogues of insect moulting hormones (ecdysteroids) are widely distributed in plants where they can reach concentrations up to 3% of their dry weight (Lafont, 1997; Dinan, 2001). The major phytoecdysteroid is 20-hydroxyecdysone, which is the same molecule as the active hormone of insects. Phytoecdysteroids are generally considered as secondary metabolites that protect plants against phytophagous insects either by feeding deterrency or toxicity (Lafont, 1997, Dinan, 2001). Indeed, ingested ecdysteroids are highly toxic to insects, e.g., Pectinophora gossypiella and Spodoptera fugiperda (Kubo et al., 1981, 1983), Bombyx mori (Tanaka and Naya, 1995); S. frugiperda, Inachi io, and Aglais urticae (Blackford and Dinan, 1997a,b), Lobesia botrana (Mondy et al., 1997), and Bradysia impatiens (Schmelz et al., 2002) by endocrine disrupting leading to death. Ecdysteroids also protect plants against soil nematodes (Soriano et al., 2004). However, several insect species remain unaffected by ecdysteroids present in their food even at high concentrations. This is, for instance, the case of Heliothis virescens (Kubo et al., 1987), Heliothis armigera (Robinson et al., 1987), Spodoptera littoralis (Blackford et al., 1996), and Lacanobia oleraceae (Blackford and Dinan, 1997a). On the other hand, low doses of ecdysteroids may be beneficial, as reported in B. mori on which 20E synchronized larval development and silk production (Ninagi and Maruyama, 1996), and in Myzus persicae where the hormone increased the number of offspring (Malausa et al., 2006), and in bees where it improved fat body development (Moskalenko et al., 1992). Resistant insects have developed effective detoxification/inactivation mechanisms mainly though oxidation at C-26, oxidation/epimerization at C3, or conjugation reactions concerning secondary alcohols (at C-2, C-3, and C-22), which lead to 1 Université Abdelmalek Essaadi, Faculté des Sciences et Techniques, CEEM-Laboratoire de Biologie Appliquée et Sciences de l’Environnement, Tangier, Morocco 2 Université Pierre et Marie Curie, Laboratoire Protéines, Biochimie Structurale et Fonctionnelle, CNRS FRE 2852, Paris, France 3 INRA, Laboratoire Physiologie de l’Insecte, Signalisation et Communication, UMR-A 1272, Versailles, France Contract grant sponsor: “Pôle d’Excellence Régional: CEEM-AUF project F Sayah.” *Correspondence to: Rene Lafont, Université Pierre et Marie Curie, Laboratoire Protéines, Biochimie Structurale et Fonctionnelle, CNRS FRE 2852, Paris, France. E-mail: email@example.com © 2007 Wiley-Liss, Inc. DOI: 10.1002/arch.20191 Published online in Wiley InterScience (www.interscience.wiley.com) 66 Rharrabe et al. the formation of various polar or apolar conjugates (Lafont and Connat, 1989; Rees, 1995; Lafont et al., 2005). The present work has investigated the metabolic fate of ingested ecdysteroids in three insect species, the aphid M. persicae and the lepidoptera Plodia interpunctella and Ostrinia nubilalis. The aim was essentially qualitative and does not allow us to draw quantitative conclusions about the relative importance of the different pathways, as there were only 2–3 repeats in each case. MATERIALS AND METHODS Insects P. interpunctella larvae were taken from Errachidia province in South-East region of Morocco and reared at 28 ± 2°C under a 18L:6D photoperiod and 70 ± 5% relative humidity. Larvae were fed date fruits. Fourth instar larvae were used in this study. O. nubilalis was provided by S. Moyal (INRAVersailles) and reared on an artificial medium (Moyal, personal communication) at 25°C under a L12:D12 photoperiod and 50% relative humidity. Fifth instar larvae were used for metabolic studies. M. persicae was provided by R. Delorme (INRA, Versailles). The mass rearing was conducted on broad bean (Vicia fabae) seedlings from the Aguadulce cultivar at 21°C under a L16:D8 photoperiod and 80% relative humidity. For metabolic studies, last instar nymphs/wingless adult aphids were fed an artificial diet containing a mixture of amino acids, vitamins, mineral salts, and sucrose as described by Febvay et al. (1988). Chemicals 23,23,24,24-[3H4]-Ecdysone, specific activity Ci/ mmol (NEN, 52 Ci/mmol), and 1α,2α-[3H2]-20hydroxyecdysone (20E, 40 Ci/mmol, synthesis to be described elsewhere) were used for metabolic studies. Reference ecdysteroids (20E, 3-epi-20E, 3dehydro-20E and 22-fatty acyl esters of 20E) were synthesized or isolated in previous studies. Syn- thetic 20E glucosides (Pís et al., 1994) were a generous gift from Dr. Juraj Harmatha (Prague, Czech Republic), and 20E galactosides from the late Pr. Ziyadilla Saatov (Tashkent, Uzbekistan). Feeding Experiments Tritiated 20E dissolved in 10 µl of ethanol was incorporated into the diet of P. interpunctella larvae that were previously starved during 24 h in order to induce a high feeding rate. They were left 3 h on treated diet, then transferred in other Petridishes containing unlabeled diet, and their faeces were collected after 24 h. In the case of O. nubilalis, tritiated ecdysone (1 µCi) was evaporated on a small piece (25 mg) of artificial medium and given to three larvae. Faeces were collected after 4 and 8 h. Control experiments were performed by injecting labeled ecdysone to O. nubilalis larvae, and the faeces were collected over the next 8 h. For the aphids M. persicae, tritiated 20E was dissolved in the artificial medium (80 µl) and put between 2 parafilm layers at the top of a polypropylene cylinder (30-mm high, 4-cm diameter); groups of 30 animals were allowed to feed on it for 24 h. Extractions of Excreta Honeydew (aphids) or faeces (lepidoptera) were extracted with methanol using a sonication bath during 30 min, then centrifuged. The solvent was dried under nitrogen at 25°C. Dried samples were redissolved in 1 ml of methanol. Aliquotes (10 µl) were then assayed for radioactivity by using a Kontron Beta IV liquid scintillation counter. Then, samples were analyzed by radio-HPLC with different systems. HPLC Analyses HPLC equipment from Thermo Separation was used for the separation and identification of the various labelled metabolites present in the samples, based on their comigration with reference ecdysteroids using different chromatographic systems. Flow-rate was 1 ml/min in every case. Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. Pathways of Ingested Ecdysteroids RP-HPLC was performed by using an ACE 5C18-HL column (150 × 4.6 mm i.d.) eluted with a 17% acetonitrile-isopropanol (5:2) in 20 mM Tris-HClO4, pH 7.5, during 20 min, then 100% acetonitrile-isopropanol (5:2) for 15 min (solvent system 1). An isocratic system was also used with 17% acetonitrile-isopropanol (5:2) in 0.1% trifluoroacetic acid (TFA) (solvent system 2). RPHPLC used, alternatively, an Ultrasphere-5ODS2 column (250 mm, 4.6 mm i.d.) eluted with a gradient of acetonitrile-isopropanol (5:2, v/v) in 20 mM Tris/HClO4, pH 7.5 (8 to 40% in 40 min, then 40 to 100% in 20 min, then 100% isocratic for 20 min) (solvent system 3), The same solvent system was also used with a Spherisorb ODS2 column (250 mm, 4.6 mm i.d., solvent system 4). NP-HPLC was performed by using a Kromasil column (250 mm, 4.6 mm i.d., particle size 3.5 µm, from A.I.T.) eluted with a flow-rate of 1 ml/ min with dichloromethane-propan-2-ol-water (100:40:2.5, v/v/v) (system 5). Enzymatic Hydrolyses Polar metabolites:aliquotes of each extract were evaporated and dissolved in 1 ml of 50 mM sodium acetate buffer, pH 5.4, and incubated overnight with 1 mg β-glucuronidase from Helix pomatia (H1 type, Sigma) at 37°C. Apolar metabolites:samples were evaporated and dissolved in 1 ml 20 mM borate buffer, pH 8.5, and incubated overnight with 1 mg porcine liver esterase (Sigma) at 37°C. In both cases, ecdysteroids were then adsorbed on a C18 Sep-Pak cartridge (Millipore) and eluted with 5 ml of absolute methanol. RESULTS AND DISCUSSION Metabolism of Ingested 20-Hydroxyecdysone in P. interpunctella Larvae Ingested radiolabelled 20E was transformed into three sets of compounds of decreasing polarity (Fig. 1): the first group was polar, the second exhibited a polarity close to that of 20E, and the third group Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. 67 Fig. 1. RP-HPLC analysis of radioactive metabolites in faeces of P. interpunctella larvae following ingestion of 20E (chromatographic system 1). was apolar metabolites. The first two groups were separated from the third one using a C18 Sep-Pak cartridge (Millipore), which was eluted with 5 ml of 60% methanol (SP60) and then with 5 ml of 100% methanol (SP100). The SP60 and the SP100 (the latter after esterase hydrolysis) were analyzed by isocratic RP-HPLC using solvent system 2 (Fig. 2). Such conditions appeared very resolutive and allowed to fully characterize the different peaks. Metabolites of polarity close to 20E coeluted with reference 3-epi-20-hydroxyecdysone (3-epi-20E) and 3-dehydro-20-hydroxyecdysone (3D20E), respectively (3D20E gives two peaks under these conditions). The two polar peaks were ionizable, their retention time varied with solvent pH (it increased at acidic pH), and they remained unchanged after enzymatic hydrolyses (data not shown). So these metabolites are not conjugates, and their behaviour indicates that the more polar is probably 20hydroxyecdysonoic acid (20Eoic) and the second either 3-dehydro-20-hydroxyecdysonoic acid or 3epi-20-hydroxyecdysonoic acid. The SP100 fraction after hydrolysis gave 20E, 3-epi-20E, and 3D20E (Fig. 2B). So the apolar metabolites correspond to conjugates of 20E (and to a lower extent of 3-epi20E and 3D20E) with various fatty acids (probably 22-acyl esters as in other insect species). There were no apolar esters of ecdysonoic acids. 68 Rharrabe et al. Fig. 2. RP-HPLC analysis of radioactive metabolites in faeces of P. interpunctella larvae following ingestion of 20E (chromatographic system 2). A: Fraction SP60; B: fraction SP100 after esterase treatment. Fig. 3. HPLC analysis of radioactive metabolites in honeydew of M. persicae adults following ingestion of 20E. A: RP-HPLC (chromatographic system 4); B: NP-HPLC (chromatographic system 5). Metabolism of Ingested 20-Hydroxyecdysone in M. persicae Adults The analysis of chromatograms (Fig. 3A,B) showed that ingested 20E was excreted mostly unchanged, or transformed into a single more polar (non-ionic and hydrolysable) conjugate of 20E, which coeluted with 20E 22-glucoside in both RP-HPLC and NPHPLC systems. These conditions were able to resolve all available 20E glucosides and glucuronides (Maria et al., 2005). Thus, although the present data do not fully establish the structure of the conjugate, it appears that M. persicae would use an original detoxification mechanism, maybe connected with the presence of high glucose levels in its diet. Metabolism of Ingested and Injected Ecdysone In O. nubilalis Larvae Ingested ecdysone was rapidly excreted mainly in the form of apolar conjugates that released only ecdysone upon esterase treatment (Fig. 4A,B). In contrast, injected ecdysone was converted to 20E, 20,26-dihydroxyecdysone (20,26E) and 20-hydroxyecdysonoic acid (20Eoic) together with apolar conjugates (Fig. 4C,D). The situation thus differs from that in P. interpunctella with (1) the absence of the oxidase/epimerase system in O. nubilalis and (2) the lack of 20Eoic formation. The situation in O. Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. Pathways of Ingested Ecdysteroids 69 Fig. 4. RP-HPLC analysis of radioactive metabolites in faeces of O. nubilalis larvae following ingestion (A,B) or injection (C,D) of ecdysone (chromatographic system 3). A and C: Before esterase treatment; B and D: after esterase treatment. nubilalis is similar to that in H. armigera (Robinson et al., 1987). produced by baculoviruses was described (O’Reilly et al., 1991). The present data in M. persicae thus extend the formation of 22-glucosides as an inactivation mechanism for dietary ecdysteroids. The formation of apolar acyl esters has been widely documented among insects (Table 1). In the case of O. nubilalis, it appears that the gut barrier is particularly efficient, as none of the ingested ecdysone enters the insect body (otherwise it would have undergone 20- and/or 26-hydroxylation). A similar situation was previously encountered for instance in H. armigera (Robinson et al., 1987). In the case of P. interpunctella, the detoxification mechanisms are more complex, as they involve Diversity of Detoxification Pathways Against Ingested Ecdysteroids The formation of glucoside conjugates was proposed a long time ago to take place in Calliphora erythrocephala (Heinrich and Hoffmeister, 1970), but the structure of the conjugates was not fully established. Later on, the formation of 26-hydroxyecdysone 22-glucoside was observed in Manduca sexta embryos (Thompson et al., 1987). The formation of ecdysone 22-glucoside by an enzyme Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. 70 Rharrabe et al. TABLE 1. Metabolites of Ingested Ecdysteroids in Various Insect Species* Type of metabolite Species References 2-phosphate Locusta migratoria Manduca sexta Spodoptera littoralis Drosophila melanogaster Plodia interpunctella Locusta migratoria Manduca sexta Pieris brassicae Spodoptera littoralis Pieris brassicae Gryllus bimaculatus Heliothis virescens Heliothis armigera Phormia sp. Gryllus bimaculatus Spodoptera littoralis Ostrinia nubilalis Plodia interpunctella Myzus persicae Manduca sexta Spodoptera littoralis Bombyx mori Feyereisen et al. (1976); Modde et al. (1984); Lafont (1994) Weirich et al. (1986)b Webb et al. (1995, 1996)b Sommé-Martin et al. (1988) This study Modde et al. (1984) Weirich et al. (1986, 1991) Beydon et al. (1987) Webb et al. (1995, 1996)b Beydon et al. (1987) Hoffmann et al. (1990) Kubo et al. (1987); Zhang and Kubo (1993) Robinson et al. (1987) Sutter (1986) Thiry and Hoffmann (1992) Blackford et al. (1997) This study This study This study Weirich et al. (1986)b Webb et al. (1995, 1996)b Hikino et al. (1971) 3-dehydro3-acetate 3α-epimer 3α-phosphate 14-deoxy-a 22-acyl ester 22-glucoside 22-phosphate Side-chain cleavage to poststerone *Two reactions can be combined, e.g., the double-conjugates in Locusta (3-acetate + 2-phosphate), the 3-epimer + 3-phosphate (Pieris), and the 3-epimer or 3-dehydro + 22-acyl esters (Plodia). a 14-Dehydroxylation takes place after feeding ecdysteroids, but this reaction is most probably due to gut bacteria and not to insect tissues. b In vitro enzymatic studies with midgut cytosol. both a dehydrogenase/epimerase reaction and 22acylation. The situation is similar in S. littoralis (Webb et al., 1995, 1996; Blackford et al., 1997). The lack of 3-epimerization in O. nubilalis was an unexpected finding, as such a reaction is widespread in lepidopteran larvae (Weirich and Bell, 1997; Lafont et al., 2005). Our result is, however, consistent with the absence of 3-epimers detection in metabolic studies performed with various organs of O. nubilalis larvae (Gelman et al., 1991). Thus, there is a great diversity of detoxification pathways of ingested ecdysteroids in insects, which concern essentially the secondary hydroxyl groups (Fig. 5). From the available data, it seems that the nature of detoxification mechanisms does not correlate with the systematic position of insects, and that Lepidoptera, the most investigated insect group in this respect, show large species differences. The diversity of detoxification mechanisms in insects is probably even greater. For instance, formation of sulfate esters of ecdysteroids is highly probable in some species, although up to now it was demonstrated to occur in vitro only (Yang and Wilkinson, 1972; Shampengton and Wong, 1989; Matsumoto et al., 2003). In the most recent study, the sulfotransferase was isolated from the fat body of the fleshfly Sarcophaga peregrina, and whether it is also present in the midgut was not investigated (Matsumoto et al., 2003). Finally, the availability of 20E labeled on the nucleus will allow us to further explore the sidechain cleavage reaction, which was early described in B. mori (Hikino et al., 1971) but never observed since. When looking at ecdysteroid metabolites in insect faeces, we should keep in mind that some reactions may take place within the faeces themselves, due to microbial metabolism. For instance, this was shown for indomethacin, a prostaglandin synthesis inhibitor, when administered to Manduca sexta larvae (Miller and Stanley-Samuelson, 1996). In the case of ecdysteroids, this may apply to the formation of 14-deoxy metabolites (Hoffmann et al., 1990), and to side-chain cleavage products (see, e.g., Tom et al., 1975), but the other described reactions are probably due to insect enzymes (Lafont et al., 2005). Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. Pathways of Ingested Ecdysteroids 71 Fig. 5. The various inactivation reactions on the 20hydroxyecdysone molecule in insects. ACKNOWLEDGMENTS Dinan L. 2001. Phytoecdysteroids: biological aspects. Phytochemistry 57:325–339. The authors thank Simon Moyal for his contribution to O. nubilalis experiments, and Dr. Juraj Harmatha for providing reference 20E glucosides. Febvay G, Delobel B, Rahbe Y. 1988. Influence of the amino acid balance on the improvement of an artificial diet for a biotype of Acyrthosiphon pisum (Homoptera:Aphididae). Can J Zool 66: 2449–2453. LITERATURE CITED Beydon P, Girault JP, Lafont R. 1987. Ecdysone metabolism in Pieris brassicae during the feeding last larval instar. Arch Insect Biochem Physiol 4:139–149. Blackford M, Dinan L. 1997a. The effects of ingested ecdysteroid agonists (20-hydroxyecdysone, RH5849 and RH5992) and an ecdysteroid antagonist (cucurbitacin B) on larval development of two polyphagous lepidopterans Acherontia atropus and Lacanobia oleracea. Ent Exp Appl 83:263–276. Blackford MJP, Dinan L. 1997b. The effects of ingested 20hydroxyecdysone on the larvae of Aglais urticae, Inachis io, Cynthia cardui (Lepidoptera: Nymphalidae) and Tyria jacobaeae (Lepidoptera: Arctiidae). J Insect Physiol 43:315–327. Blackford M, Clarke B, Dinan L. 1996. Tolerance of the Egyptian cotton leafworm Spodoptera littoralis (Lepidoptera: Noctuidae) to ingested phytoecdysteroids. J Insect Physiol 42:931–936. Blackford MJP, Clarke BS, Dinan L. 1997. Distribution and metabolism of exogenous ecdysteroids in the Egyptian Cotton leafworm Spodoptera littoralis (Lepidoptera: Nocturidae). Arch Insect Biochem Physiol 34:329–346. Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. Feyereisen R , Lagueux M, Hoffmann JA. 1976. Dynamics of ecdysone metabolism after ingestion and injection in Locusta migratoria. Gen Comp Endocrinol 29:319–327. Gelman D, DeMilo AB, Thyagaraja BS, Kelly TJ, Masler EP, Bell RA, Borkovec AB. 1991. 3-Oxoecdysteroid 3b-reductase in various organs of the European corn borer, Ostrinia nubilalis (Hubner). Arch Insect Biochem Physiol 17:93–106. Heinrich G, Hoffmeister H. 1970. Bildung von Hormonglykosiden als Inaktivierungsmechanismus bei Calliphora erythrocephala. Z Naturforsch B 25:358–361. Hikino H, Ohizumi Y, Takemoto T. 1971. Catabolism of ponasterone A to ecdysterone, inokosterone and poststerone in Bombyx mori. Chem Commun 1036–1037. Hoffmann KH, Thiry E, Lafont, R. 1990. 14-Deoxyecdysteroids in an insect (Gryllus bimaculatus). Z Naturforsch 45c:703– 708. Kubo I, Klocke JA, Asano S. 1981. Insect ecdysis inhibitors from the East African medicinal plant Ajuga remota (Labiatae). Agric Biol Chem 45:1925–1927. Kubo I, Klocke JA, Asano S. 1983. Effects of ingested 72 Rharrabe et al. phytoecdysteroids on the growth and development of two lepidopterous larvae. J Insect Physiol 29:307–316. Moskalenko PG, Piletskaya IV, Kholodova YuD. 1992. Effect of ecdysterone on bees and varroa mites. Veterinariya 1:42–43. Kubo I, Komatsu S, Asaka Y, De Boer G. 1987. Isolation and identification of apolar metabolises of ingested 20hydroxyecdysone in frass of Heliothis virescens larvae. J Chem Ecol 13:785–794. Ninagi O, Maruyama M. 1996. Utilization of 20-hydroxyecdysone extracted from a plant in sericulture. Japan Agric Res Q 30:123–128. Lafont R. 1994. Phytoecdysteroids and moulting control. In: Rembold H, Benson JA, Franzen H, Weickel B, Schulf AF, editors. New strategies for locust control. ATSAF. p 39–44. Lafont R. 1997. Ecdysteroids and related molecules in animals and plants. Arch Insect Biochem Physiol 35:3–20. Lafont R, Connat J-L. 1989. Pathways of ecdysone metabolism. In: Koolman J, editor. Ecdysone, from chemistry to mode of action. Stuttgart: Georg Thieme Verlag. p 167–180. Lafont R, Dauphin-Villemant C, Warren J, Rees HH. 2005. Ecdysteroid chemistry and biochemistry. In: Gilbert LI, Iatrou K, Gill S, editors. Comprehensive molecular insect science, vol. 3. New York: Elsevier. p 125–196. Malausa T, Salles M, Marquet V, Guillemaud T, Alla S, MarionPoll F, Lapchin L. 2006. Within-species variability of the response to 20-hydroxyecdysone in peach-potato aphid (Myzus percicea Sulzer). J Insect Physiol 52:480–486. Maria A, Girault J-P, Saatov Z, Harmatha J, Dinan LN, Lafont O’Reilly DR, Howarth OW, Rees HH, Miller LK. 1991. Structure of the ecdysone glucoside formed by a baculovirus ecdysteroid UDP-glucosyltransferase. Insect Biochem 21:795–801. Pís J, Hykl, J, Budésínsky M, Harmatha, J. 1994. Regioselective synthesis of 20-hydroxy-ecdysone glycosides. Tetrahedron 50:9679–9690. Rees HH. 1995. Ecdysteroids biosynthesis and inactivation in relation to function. Eur J Entomol 92:9–39. Robinson PD, Morgan ED, Wilson ID, Lafont R. 1987. The metabolism of ingested and injected [3H]ecdysone by final instar larvae of Heliothis armigera. Physiol Entomol 12:321–330. Schmelz EA, Grebenok RJ, Ohnmeiss TE, Bowers WS. 2002. Interactions between Spinacia oleracea and Bradysia impatiens: a role for phytoecdysteroids. Arch Insect Biochem Physiol 51:204–221. R. 2005. Ecdysteroid glycosides : chromatographic properties and biological activity. J Chrom Sci 43:149–157. Shampengton L, Wong KP. 1989. An in vitro assay of 20hydroxyedysone sulfotransferase in the mosquito, Aedes togoi. Insect Biochem 19:191–196. Matsumoto E, Matsui M, Tamura H. 2003 Identification and purification of sulfotransferases for 20-hydroxyecdysteroid from the larval fat body of a fleshfly, Sarcophaga peregrina. Biosci Biotechnol Biochem 67:1780–1785. Sommé-Martin G, Colardeau J, Lafont R, 1988. Conversion of ecdysone and 20-hydroxyecdysone into 3-dehydroecdysteroids is a major pathway in third instar Drosophila melanogaster larvae. Insect Biochem 18:729–734. Miller JS, Stanley-Samuelson DW. 1996. The pharmacology of indomethacin, a prostaglandin biosynthesis inhibitor, in larvae of the tobacco hornworm, Manduca sexta. J Insect Physiol 42:893–901. Soriano I, Riley I, Potter M, Bowers W. 2004. Phytoecdysteroids : a novel defense against plant-parasitic nematodes. J Chem Ecol 30:1885–1899. Modde J-F, Lafont R, Hoffmann JA. 1984. Ecdysone metabolism in Locusta migratoria larvae and adults. Int. J Invert Reprod Dev 7:161–183. Mondy N, Caïssa C, Pitoizet N, Delbecque JP, Corio-Costet MF. 1997. Effect of the ingestion of Serratula tinctoria extracts, a plant containing phytoecdysteroids, on the development of the vineyard pest Lobesia botrana (Lepidoptera: Tortricidae). Arch Insect Biochem Physiol 35:227–235. Sutter J. 1986. Etude compareée du métabolisme de l’ecdysone chez trois Diptères adultes (Phormia species, Lucilia cuprina et L. sericata) après injection ou ingestion de [3H]-ecdysone. Travail de Certificat, Université de Neuchâtel, Suisse. [Comparative study of ecdysone metabolism in three adult Diptera (Phormia sp., Lucilia cuprine and L. sericata) after injection or ingestion of [3H]-ecdysone. Certificate Memoir. University of Neuchatel, Switzerland.] Tanaka Y, Naya S-I. 1995. Dietary effect of ecdysone and 20Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch. Pathways of Ingested Ecdysteroids 73 hydroxyecdysone on larval development of two lepidopteran species. Appl Ent Zool 30:285–294. midgut cytosol of the cotton leafworm, Spodoptera littoralis. Insect Biochem Molec Biol 26:809–816. Thiry Y, Hoffmann KH. 1992. Dynamics of ecdysone and 20hydroxyecdysone metabolism after injection and ingestion in Gryllus bimaculatus. Zool Jb Physiol 96:17–38. Weirich GF, Bell RA. 1997. Ecdysone 20-hydroxylation and 3-epimerization in larvae of the gipsy moth, Lymantria dispar L.: tissue distribution and developmental changes. J Insect Physiol 43:643–649. Thompson MJ, Feldlaufer MF, Lozano R, Rees HH, Lusby WR, Svoboda JA, Wilzer KR.1987. Metabolism of 26-[14C]hydroxyecdysone 26-phosphate in the tobacco hornworm, Manduca sexta (L.), to a new ecdysteroid conjugate: 26-[14C]-hydroxyecdysone 22-glucoside. Arch Insect Biochem Physiol 4:1–15. Weirich GF, Thompson MJ, Svoboda JA. 1986. In vitro ecdysteroid conjugation by enzymes of Manduca sexta midgut cytosol. Arch Insect Biochem Physiol 3:109–126. Tom WM, Abul-Hajj YJ, Koreeda M. 1975. Microbial oxidation of ecdysones. A convenient preparation of rubrosterone. J Chem Soc Chem Commun 1975:24–25. Weirich GF, Thompson MJ, Svoboda JA. 1991. Enzymes of ecdysteroid 3-epimerization in midgut cytosol of Manduca sexta: pH optima, consubstrate kinetics, and sodium chloride effect. Insect Biochem 21:65–71. Webb TJ, Powls R, Rees HH, 1995. Enzymes of ecdysteroid transformation and inactivation in the midgut of the cotton leafworm, Spodoptera littoralis: properties and developmental profiles. Biochem J 312:561–568. Yang RSH, Wilkinson CF. 1972. Enzymic sulphation of pnitrophenol and steroids by larval gut tissues of the southern armyworm (Prodenia eridania Cramer). Biochem J 130:487–493. Webb TJ, Powls R, Rees HH, 1996. Characterization, fractionation and kinetic properties of the enzymes of ecdysteroid 3-epimerization and phosphorylation isolated from the Zhang M, Kubo I. 1993. Metabolic fate of ecdysteroids in larval Bombyx mori and Heliothis virescens. Insect Biochem Mol Biol 23:831–843. Archives of Insect Biochemistry and Physiology June 2007 doi: 10.1002/arch.