Premature production of late larval storage proteins in larvae of trichoplusia ni parasitized by euplectrus comstockii.код для вставкиСкачать
Archives of Insect Biochemistry and Physiology 26:97-109 (1 994) Premature Production of Late Larval Storage Proteins in Larvae of Trichoplusia ni Parasitized by Euplectrus comstockii Thomas A. Coudron, Davy Jones, and Grace Jones Biological Control of Insects Research Laboratoy, U S . Department of Agriculture, A R S , Columbia, Missouri (T.A.C.); Graduate Center for Toxicology (D.J.),and School of Biological Sciences (G.J.), University of Kentucky, Lexington Investigations were conducted to determine the titer of storage proteins in larvae of the cabbage looper, Trichoplusia ni (Hubner), that were parasitized by the ectoparasitoid Euplectrus comstockii Howard (Hymenoptera: Eulophidae). A gradual increase was noted in the titer of the storage proteins present in the hernolymph of parasitized third and fourth instar larvae and in the hemolymph of isolated thoracic and abdominal tissues of fourth instar larvae. The final amount present in parasitized third and fourth instar larvae was similar to thatfound in nonparasitized fifth instar larvae. The stimulation of storage proteins in envenomed larvae demonstrates the ability (competence) of early larval stages to produce a gene product that normally occurs in the last larval stadium of the lepidopteran larval host. The gene expression necessary for storage protein production in isolated tissues may be altered by mechanisms separate from inherent developmental processes and the intact endocrine system. o 1994 WiIey-Liss, Inc.* Key words: Hymenoptera, venom, host hemolymph, plasma proteins, regulation INTRODUCTION Peptides and proteins found in the hernolymph of insects are known to serve numerous functions that include intercellular communication, transport and defensive mechanisms, and energy storage 11-31. Storage proteins have been found in many species of insects and are commonly thought to supply peptides and/or amino acids required for late larval development and metamorphosis [1,4-61. Other roles attributed to these proteins include cuticular associations 171 and ligand binding and the transport of xenobiotics . Acknowledgments:A.I. Soldevila, J. Derks, and S. Brandt are recognized for their technical assistance. Received January 21, 1993; accepted April 5, 1993. Address reprint requests to Dr. T.A. Coudron, USDA ARS BCIRL, P.O. Box 7629, Columbia, M O 65205-5001. Mention of a trademark, warranty, proprietary product, or vendor does not constitute a guarantee by the U.S. Departmentof Agriculture and does not imply its approval to the exclusion of other products or vendors that may also be suitable. 0 1994 Wiley-Liss, Inc. *This article is a US Government work and, as such, is in the public domain in the United States of America. 98 Coudron et al. In their native form, the storage proteins often have a molecular weight greater than 400,000 and are composed of glycosylated subunits in hexamer or octamer aggregates. The molecular weights of the subunits range from 70,000 to 90,000 [3,9-111. The fat body of feeding larvae and nymphs has been shown to be the primary site of synthesis and secretion of the storage proteins [3,9,101. Although the fat body is accepted as the primary site of synthesis, secondary sites of synthesis have been implicated [12-151, which suggests there may be several regulatory mechanisms controlling the expression of these genes [161. Sequestration of the storage proteins by the fat body of holometabolous insects occurs just prior to the larval to pupal molt [3,17,181. Production of storage proteins is thought to be under a developmental regulation. This regulation allows for an abundant accumulation of storage proteins, to as much as 80% of the total hemolymph protein by weight in the hemolymph of the last larval instar of homometabolous insects [9,10,19,201. Trichoplusia ni (Hubner) is one of the lepidopteran insects studied in which this phenomenon has been reported .However, our understanding of the regulation, processing, transport, and biological functions of the storage proteins is limited. Parasitic Hymenoptera are known to alter the physiology, biochemistry, and behavior of the host, resulting in the enhancement of the host as a source of nutrients . One alteration is in the constituent protein profile of the host. These alterations include changes in the normal concentrations as well as alterations of the time of protein occurrence in the hemolymph of the host. Parasitism of larvae of T. ni by Hyposoter exiguae caused a decrease in the concentration of several proteins normally found in the hemolymph of the host [23,24]. Parasitism of Spodoptera frugiperda (J.E. Smith) by Cotesia mayginiventris caused an early production of several high molecular weight proteins that were not characterized further [251. A recorded decrease in the concentration of storage proteins in larvae of Pieris rapae (L.), parasitized by Cotesia glomevata , was proposed to be a result of uptake of host hemolymph proteins by the parasitoid .The reduced production of arylphorin by larvae of Manduca sexta (L.), parasitized by C. congvegata, was proposed to be a result of the inhibitory effects of parasitism on host fat body, food consumption, and growth [281. In contrast, an increase in the concentration of arylphorin was reported in larvae of T. ni parasitized by Chelonus SPP. 129-311. The eulophid, Euplectrus comstockii Howard is an ectoparasitoid of larvae of several lepidopteran insects 1321. As an ectoparasitoid, it relies entirely on a venom to induce alterations in certain physiological and biochemical events of its larval hosts. The venom, which is injected into the host during a stinging process that precedes oviposition , alters the development of the host by arresting the larval-larval ecdysal process [331. The arrestment is the result of action of the venom on the epidermal tissue and is distinct in its action from the action of other venoms of ectoparasitoids that cause paralysis of the host. In the present paper, we report the effect of the venom from E. comstockii on the expression of storage proteins in the hemolymph of envenomed larvae of T . ni. Storage Proteins in Envenomed larvae 99 MATERIALS AND METHODS Insects The E. comstockii colony was derived from a stock collected in Missouri during 1985 to 1989. Stock material was reared from field collected larvae of T. ni, Helicoverpa zed (Boddie), Plathypena scabra (Fabricius),and AZypia octomaculata (Fabricius). A continuous colony of the parasitoid was maintained in the laboratory since 1985 according to the procedure described by Coudron and Puttler .Both parasitized and nonparasitized larvae of T. ni were maintained on a semisynthetic modified wheat germ diet , and reared at 23-26"C, 40-60% RH*, and 12:12 (L:D) photoperiod 1331.Under these conditions, each of the third and fourth larval instars of T. ni lasted for 48 h, and larvae in the fifth larval instar began wandering after 54 h, spinning a cocoon within 72 h, and pupated within 90 h. Parasitism Unsexed larvae of T. ni that were in synchronous growth were used for control and treatment studies . Larvae that were ligated, parasitized, or ligated and parasitized were handled in the same manner as control (nonligated and nonparasitized) larvae. Nonligated parasitized larvae were parasitized within 4 h of ecdysis into the designated stadium. For ligated larvae, a short strand of sewing thread was used to create a ligature block between the thorax and the abdomen at 20 h post-ecdysis into the fourth stadium 1331. This process provided for the isolation of thoracic tissue (anterior to the ligature) or abdominal tissue (posterior to the ligature) from the remainder of the body prior to the endogenous release of 20-hydroxyecdysterone.Ligated larvae that were to be parasitized were exposed to the parasite until either the thoracic tissue or the abdominal tissue was parasitized. Parasitism of the isolated tissue occurred within 2 h of ligation. To remove the parasite, T. ni were anaesthetized with carbon dioxide within 24 h of oviposition, and the parasite eggs punctured thereby killing the parasite approximately 24 h before egg eclosion. No attempt was made to remove the eggs since they were firmly attached to the host cuticle, and the removal process would greatly disturb the host larvae. Protein Preparation and Determination Host hemolymph was obtained from a severed proleg according to the procedure described by Kelly and Coudron 1351. Ten microliters of hemolymph were collected at each time point without dilution, and stored at -20°C. The total number of insects bled for each time point was approximately 6 for third instar larvae, 4 for fourth instar larvae, 2 for fifth instar larvae, 8 for ligated larvae bled from the thorax, and 6 for ligated larvae bled from the abdomen. Three replicate samples were taken for gel electrophoresis analysis and two replicate samples were taken for immunoblotting analysis. *Abbreviations used: IgG = immunoglobulin with a molecular weight of 150,000; JH = juvenile hormone; L:D = 1ight:dark; RH = relative humidity; SDS-PAGE = sodium dodecyl sulfatr-polyacrylamide gel electrophoresis. 100 Coudron et al. Hemolymph proteins were separated by SDS-PAGE as described by Laemmli [361. Hemolymph samples were first centrifuged at 12,0009for 8 min (4°C) to remove hemocytes and any debris. Equal volumes containing various concentrations of the sample supernatant were applied to 20 cm x 20 cm, 10% SDS-PAGE gels and electrophoresed at 150 V for 5 h. Protein bands within the gel matrix were visualized with either a Coomassie blue stain or by a silver staining method as described by Morrissey 1371. Production of three separate polyclonal antibodies against one acidic and two basic storage proteins found in the hemolymph of T. ni has been previously described [38,39]. Each of these three antisera is specific for one of the three storage proteins. The immunoblotting techniques used, including electroblotting onto nitrocellulose and the use of 12% labeled goat anti-rabbit IgG as a secondary antibody and autoradiography to detect the presence of storage protein in the hemolymph samples, has been previously described by Soldevila and Jones . Immunoblots were probed with a mixture of the three antisera at a 1:1,000 antiserum dilution to give a strong signal. RESULTS Hemolymph Proteins in Non Ligated Larvae A comparison of the electrophoretic pattern of hemolymph proteins in control and nonligated parasitized larvae is shown in Figure 1. The growth time between ecdysis to the next larval stadium was approximately 48 h for both third and fourth instar nonparasitized larvae, and approximately 90 h in the fifth instar prior to pupation for the nonparasitized larvae. Examination of hemolymph protein samples from control fifth instar larvae revealed the presence of relatively abundant proteins, which appeared as an intense staining of one to three bands at molecular weights of approximately 70,000-80,000 in the hemolymph of late (48 h) fifth instar larvae. By comparison with other reports of hemolymph proteins, the pattern of these proteins bore similarities in time of appearance, abundance, and electrophoreticpattern with that of the storage proteins characterized from T. ni. No detectable quantities of these proteins were observed in the hemolymph from control third and fourth instar larvae. In contrast, the protein composition of the hemolymph from both third and fourth instar parasitized larvae did contain the characteristic banding pattern associated with the storage proteins. The relative abundance of these proteins, as determined by the intensity of the Coomassie blue staining of these bands, increased with the time after parasitism in both the third and fourth instars. Confirmation of the storage proteins was obtained by immunoblotting with the antibodies against three storage proteins found in T. ni (Fig.2). Control third (not shown) and fourth instar larvae did not contain detectable amounts of storage proteins. Most notably, strong immunoreactionswith antibodies for one acidic and two basic storage proteins were detected in the hemolymph of third (not shown) and fourth instar parasitized larvae within 48 h after parasitism (Fig. 2, arrows). The response signal increased in the fifth instar control larvae with an increased time in the stadium (data not shown). A similar temporal variability was observed in the response signal in the third (data not shown) 6 24 6 24 120 6 48 24 96 72 48 Control 4 t h Parasitized 3rd 6 24 48 72 96 Parasit ixed 4 t h 120 6 24 48 Control 5 t h Fig. 1. Production of storage proteins in parasitized larvae of Trichoplusia ni. Larval instar i s indicated for 3rd, 4th, and 5th instars of control and parasitized larvae with 3rd, 4th, and3!ith, respectively. The hour of sampling after the time of last larval ecdysis is indicated numerically. The positions and Mrvalues (x 10 ) for molecular weight standards are shown at the right. 48 Control 3rd Std 31 45 66.2 97.4 102 Coudron et al. Cont ro I 6 Parasitized 24 48 24 48 72 96 120 Fig. 2. lmmunoblot of one acidic and two basic storage proteins in the hemolymph from control and nonligated parasitized larvae of Trichoplusia ni. The hour of sampling after ecdysis to the 4th instar is indicated numerically. Top arrows point to the acidic protein and the bottom arrows point to the basic proteins. and fourth instar parasitized larvae, resulting in an increased signal with additional time after parasitism. Hemolymph Proteins in Ligated Larvae The ligature block between the thorax and the abdomen prevented the endogenous ecdysteroids released at approximately 28 h post-ecdysis into the fourth stadium from reaching the abdomen of the insect 1351. This was confirmed by the absence of ecdysis of the nonparasitized isolated abdomen and the complete apolysis into the next larval stadium of the isolated thorax. However, neither the isolated abdomen nor the isolated thorax showed any signs of ecdysis after parasitism. A comparison of the electrophoreticpattern of hemolymph proteins from the isolated thoraces and isolated abdomens in ligated control and ligated parasitized fourth instar larvae is shown in (Fig. 3). Ligation and parasitism studies of third instar larvae were not conducted due to the minute amount of hemolymph extracted during the bleeding process. Ligation did alter the protein composition in nonparasitized larvae. The Coomassie staining increased for several bands in the isolated thoraces and isolated abdomens of ligated control larvae (Fig. 3). However, the relative abundance and electrophoretic pattern of these Storage Proteins in Envenomed larvae Control tig. thorax 6 24 48 Pa r a sit iz ed lip. abdomen 72 6 103 24 48 72 Iig. abdomen lig. t h o r a x 24 48 72 24 48 72 96 Std 97.4 66.2 45 31 Fig. 3. Production of storage proteins in isolated abdomens of parasitized 4th instar larvae of Trichoplusia ni. The hour of sampling after the time of ligation and subsequent parasitism is indicated numerically. The positions and Mrvalues (x 10-3 for molecular weight standards are shown at the right. bands differed from that of the storage proteins observed in the nonligated control and nonligated parasitized larvae (Fig. 1). Examination of the hemolymph proteins in the isolated thoraces and isolated abdomens of parasitized fourth instar larvae revealed the presence of storage proteins, which appeared to intensify with time after parasitism but to a lesser degree than observed in nonligated parasitized third and fourth instar larvae. Confirmation of the identity of the storage proteins was again obtained by immunoblotting with the antibodies against the three forms of the storage proteins found in T. ni (Fig. 4). A positive immunoreaction for both the acidic and basic forms of the storage proteins were detected within 48 h of parasitism for the hemolymph taken from the thoraces and the abdomens of the ligated and parasitized larvae but not that from the ligated control larvae (Fig. 4, arrows). The signal intensified from 48 to 96 h post-parasitism. An interesting result observed was that the acidic protein failed to appear in the thorax section of ligated larvae post-parasitism, whereas it did appear in abdomen section of ligated larvae. 104 Coudron et al. Cont ro I lig. thorax lig. abdomen 6 2 4 4 6 7 6 6 2 4 4 8 7 Z Parasitized lig. thorax 8448 lig. abdomen =a4487296 Fig. 4. lmmunoblot of one acidic and two basic storage proteins in the hemolymph from the isolated tissues of ligated control and ligated parasitized larvae of Trichoplusia ni. The hour of sampling after the time of ligation and subsequent parasitism is indicated numerically. Top arrows point to the acidic protein and the bottom arrows point to the basic proteins. It should be noted that Coomassie staining detected an abundant protein migrating between the acidic and basic proteins at the location established for arylphorin . Also, the immunoblots showed a distinctly arched band shape for the basic proteins, which is a diagnostic feature of high abundance of arylphorin . DISCUSSION The expression of precocious or delayed events is common in lepidopteran larvae parasitized by hymenopteran wasps 1221. The precocious spinning of a cocoon reported in the penultimate instar of T. ni parasitized by Chelonus near curvimaculatus was thought to be due to a premature decline in the juvenile hormone level in the host 1411. The delay or suppression of metamorphosis and the creation of a supernumerary larval stage in the lepidopteran hosts of Cotesia and Copidosoma species was thought to be due to an abnormally high level of juvenile hormone and altered ecdysteroid levels in the parasitized host [28,421. A delay in larval-pupal metamorphosis reported in larvae of Anastrepha suspensa (Loew) parasitized by Biosteres Zongicaudattks was an apparent result of an elevated level of juvenile hormone in the parasitized larvae [431. All of these examples involve the penultimate or last larval stadium. Unique to the results reported here is the expression of a last larval stadium event, that of the production of storage proteins, in both the third and fourth larval stadium of a lepidopteran host that has five larval instars. We also report the early expression of storage proteins in isolated thoraces and isolated abdomens. These observations suggest that the venom activates Storage Proteins in Envenomed larvae 105 the expression of the storage proteins by acting directly on a particular tissue rather than indirectly via the intact neuroendocrine center. This is similar to the developmental arrest effect of the venom on the larvae which results in arrest of the larval-larval ecdysis process . In particular, these results demonstrate the ability (competence) of early larval stages and isolated tissues of penultimate instar larvae to produce gene products that normally occur in the intact last larval stadium of lepidopteran larvae. An interesting observation is that the ligated insects in these experiments become starved larvae following the placement of the ligature block. Thus, the abdominal region of a fourth instar ligated larvae is in essence an isolated and starved tissue that when parasitized demonstrated the ability to produce storage proteins. In contrast, starvation of the last instar larvae of Munduca sextu inhibited the production of storage protein mRNA 1121. Last instar larvae of GalZeriu that were fed for 24 h and then starved were reported to transcribe mRNA coding for the storage proteins. However, when the larvae of Gulleviu were starved at the onset of the last instar, there were little or no storage protein transcripts detected. Clearly, the effect of the Euplectrus venom is different from that of starving the larvae. Additionally, it was observed that the venom stimulated only the abdominal tissues to produce the acidic protein. This could indicate that the abdominal tissues were unique in their ability to produce this gene product, or that the expression of the protein was considerably delayed or impaired in the thoracic tissues. There is preliminary evidence for the independent regulation of hemolymph proteins in starved larvae of G. melZoneZZu . It is also plausible that the expression of this protein was impaired in the thoracic tissue due to a high accumulation of endogenous ecdysteroids which have been reported in parasitized larvae [351. Limited information is available on the mechanisms by which the juvenile hormones and ecdysteroids regulate gene expression of the storage proteins in insects. Production of certain storage proteins in normally developing lepidopteran larvae occurs in the last larval instar when endogenous JH titers are low or undetectable. In Bombyx mori, storage proteins were expressed and accumulated in the hemolymph of penultimate instar larvae that were deprived of JH and in the last instar larvae in which the JH titer is naturally low 145,461. Storage protein production in Spodopteru Iitura declined in larvae that were treated with JH or a JH analog, but increased in allatectomized larvae [471. The appearance of an acidic storage protein with a molecular weight of 76,000 in the last larval instar of T . ni was suppressed by treatment with JH as was the concentration of the level of its mRNA 1381. JH also suppressed mRNA abundance for the two basic proteins [391. These results show that JH suppresses the expression of storage proteins by regulating the level of mRNA of storage proteins. However, a decline in level of glycolipoprotein storage proteins in the hemolymph of Galleria larvae treated with JH was thought to be due to the induction of a supernumerary larval molt rather than the direct action of JH on the production and/or stability of the gene transcripts . The expression of storage proteins in M. sextu  and in HeZiothis virescens (Fabricius)[161appeared to be negatively regulated by 20- hydroxyecdysterone. The ecdysteroid, 20-hydroxyecdysterone,caused a decline in the level of storage 106 Coudron et al. protein mRNA transcripts in injected intact and prothorax-ligated larvae of Galleria I441 and when applied to GaZZeria fat body in vitro 1491. Similarly, mRNA transcripts of the storage proteins were not found in newly ecdysed last instar larvae of Galleria when the endogenous level of 20-hydroxyecdysterone was high 1491. Comparative results were found in in vitro studies with larval fat body preparations from Calliphora uicina where 20-hydroxyecdysterone repressed storage protein gene expression 1501. Therefore, 20-hydroxyecdysterone appears to cause a cessation of the storage protein gene expression. A previous study 1351has shown that ecdysteroids, including 20-hydroxyecdysterone, are absent in the hemolymph of larvae of T. ni parasitized by E. plathypenae prior to, and during, the accumulation of the storage protein time as reported here. Deprivation of the ecdysteroid hormone from parasitized larvae of the early instars would eliminate the suppression of storage protein production by this hormone. This may provide the first indication as to why the venom alters the ecdysteroid titer of the host when the arrestment of the larval-larval ecdysis was found to be regulated independent of 20-hydroxyecdysterone 1331. Alterations of the JH titer in parasitized larvae have not been determined. However, a decline in the JH titer in parasitized larvae is likely, given that the production of storage proteins follows parasitism. Another interesting observation is that the acidic and basic storage proteins may be independently regulated. The rate at which the basic proteins appeared in isolated and parasitized thoracic regions was faster than the rate of appearance of the acidic protein. An inverse relation existed in the parasitized abdomen region where the rate at which the acidic proteins appeared was faster than that of the basic proteins. Thus, the acidic and basic proteins appear to be independently regulated and that regulatory mechanism may differ among various tissues of the insect. The production of storage proteins appears to occur in every larval instar after parasitism by E. cornstockii, and the time of occurrence is approximate to the time of the parasitoid egg hatch and larval feeding. The protein profile of the hemolymph of the host at the time of parasitoid feeding is considerably different than the profile at the time of oviposition. A major change includes the concentration of the storage proteins as discussed here, as well as other proteins of lower molecular weights. The alteration in the protein content would assure the parasitoid of a particular protein profile in the hemolymph of the host regardless of the larval instar of the parasitized host. The timing of the storage protein production would imply a role of direct benefit to the developing parasitoid as opposed to serving some function that regulates a specific stage of host development. It is plausible that the storage proteins serve a nutritional function for the parasitoid as they do for the last larval instar of the host. LITERATURE CITED 1. Wyatt GR, Pan ML (1978): Insect plasma proteins. Annu Rev Biochem 47779. 2. Gotz P, Boman HG (1985): Insect immunity. In Kerkut GA, Gilbert LI (eds): Comprehensive Insect Physiology, Biochemistryand Pharmacology. Orlando, FL:Academic Press, pp 543-485. Storage Proteins in Envenomed larvae 107 3. Levenbook L (1985): Insect storage proteins. In Kerkut GA, Gilbert LI (eds): Comprehensive Insect Physiology, Biochemishyand Pharmacology.Orlando, FL: AcademicPress, pp 307-346. 4. Thomson JA, Fadok KA, Shaw DC, Whitten MI, Foster GG, Bert LM (1976):Genetics of lucilin, a storage protein from the sheep blowfly Lucilia cuprina (Calliphoridae).Biochem Genet 14145. 5. Tojo S, Betchaku T, Ziccardi VJ, Wyatt JR (1978):Fat body protein granules and storage proteins in the silkmoth, Hyalophora cecropia. J Cell Biol78:823. 6. Telfer WH, Keim PS, Law JH (1983): Arylphorin, a new protein from Hyalophora cecropia: Comparisons with calliphorin and manducin. Insect Biochem 13:601. 7. Scheller K, Zimmerman HP, Sekeris CE (1982): Calliphorin, a protein involved in cuticle formation of the blowfly, Calliphora vicina (L.) Insect Biochem 12:277. 8. Haunerland NH, Bowers WS (1986):Binding of insecticides to lipophorin and arylphorin, two hemolymph proteins of Heliothis zea. Arch Insect Biochem Physiol3:87. 9. Kinnear JF,Thomson JA (1975): Nature, origin and fate of the major haemolymph proteins in Callipkoru. Insect Biochem 5531. 10. Kramer SJ, Mundall EC, Law JH (1980): Purification and properties of manducin, an amino acid storage protein of the haemolymph of larval and pupal Manduca sextu. Insect Biochem 10:279. 11. Miller SG, Silhacek DL (1982): Identification and purification of storage proteins in tissues of the greater wax moth, Galleria mellonellu (L.). Insect Biochem 12:277. 12. Riddiford LM, Hice RH (1985): Developmental profiles of the mRNAs for Manduca arylphorin and two other storage proteins during the final larval instar of Manduca sexta. Insect Biochem 15:489. 13. Palli SR, Locke M (1987): The synthesis of hemolymph proteins by the larval epidermis of an insect, Culpodes ethlius (Leipdoptera: Hesperiidae). Insect Biochem 17711. 14. Palli SR, Locke M (1987): The synthesis of hemolymph proteins by the larval midgut of an insect, Calpodes ethlius (Lepidoptera: Hesperiidae). Insect Biochem 17:561. 15. Fife HU, Palli SR, Locke M (1987):A function for the pericardial cell in an insect. Insect Biochem 17829. 16. Leclerc RF, Miller SG (1990): Identification and molecular analysis of storage proteins from Heliothis virescens. Arch Insect Biochem Physioll4131. 17. Riddiford LM, Law JH (1983): Larval serum proteins of Lepidoptera. In Scheller K (ed): The Larval Serum Proteins of Insects. New York: Georg Thieme Verlag, pp 75-85. 18. Haunerland NH,Bowers WS (1987): Larval serum proteins as transport vehicles for hydrophobic compounds. Biol Chem Hoppe Seyler 368:572. 19. Sekeris CE, Scheller K (1977): Calliphorin, a major protein of the blowfly: Correlation between the amount of protein, its biosynthesis, and the titer of translatable Calliphorin-mRNA during development. Dev Biol59:12. 20. Tojo S, Nagata M, Kobayashi M (1980):Storage proteins in the silkworm, Bornbyx mori. Insect Biochem 10:289. 108 Coudron et al. 21. Jones G, Hiremath ST, Hellmann GM, Woznaiak M, Rhoads RE (1987): Inhibition and stimulatory control of developmentally regulated hemolymph proteins in Trichoplusia ni. In Law JH (ed): UCLA Symp Cell Biol New Ser vol49. New York Alan R. Liss, pp 295-304. 22. Coudron TA (1991):Host-regulating factors associated with parasitic Hymenoptera. In Hedin PA (ed): Naturally Occumng Pest Bioregulators.ACS Symp Ser No. 449, pp 41-65. 23. Smilowitz Z (1973): Electrophoreticpatterns in hemolymph protein of cabbage looper during development of the parasitoid Hyposoter exigune. Ann Entomol SOCAm 66:93. 24. Thompson SN (1982): Effects of parasitization by the insect parasite Hyposofer cxiguue on the growth, development and physiology of its host Trichoplusia ni. Parasitology 84:491. 25. Ferkovich SM, Greany PD, Dillard CR (1983): Changes in haemolymph proteins of the fall armyworm Spodopterafrugiperda associated with parasitism by the braconid parasitoid Cotesiu marginiventris. J Insect Physiol29:933. 26. Smilowitz Z, Smith CL (1977): Hemolymph proteins of developing Pieris vapue larvae parasitized by Apunteles glomerutus. Ann Entomol SOCAm 70:447. 27. Kawai T, Maeda S, On0 H, Kai H (1983):Parasitic effects of wasp, Apunteles glomcratus, on the hemolymph storage proteins in host cabbage worm, Pieris rapae. J Fac Agric Tottori Univ 18:18. 28. Beckage NE, Templeton TJ (1986):Physiological effects of parasitism by Apunteles congregatus in terminal-stage tobacco hornworm larvae. J Insect Physiol32:299. 29. Jones D (1989):Protein expression during parasite redirection of host (Trichoplusia ni) biochemistry. Insect Biochem 19:445. 30. Kunkel JG, Grossniklaus C, Karpells ST, Lanzrein C (1990): Arylphorin of Trichoplusia ni: Characterization and parasite induced precocious increase in titer. Arch Insect Biochem Physiol 13:117. 31. Jones D, Jones G, Rudnicka M, Click A (1985):Precocious expression of the final larval instar developmental program in larvae of Trichoplusia ni pseudoparasitized by Ckelonus spp. Comp Biochem Physiol83B:339. 32. Coudron TA, Puttler B (1988): Response of natural and factitious hosts to the ectoparasite Euplectvus plathypeme (Hymenoptera: Eulophidae). Arch Insect Biochem Physiol81:932. 33. Coudron TA, Kelly TJ, Puttler B (1990): Developmental responses of Trickoplusiu ni (Lepidoptera: Noctuidae) to parasitism by the ectoparasite Euplectrus plathypenae (Hymenoptera: Eulophidae). Arch Insect Biochem Physioll383. 34. Wilkinson JD, Morrison RK, Peters PK (1972): Effect of Calco oil red N-1700 dye incorporated into a semi-artificial diet of the imported cabbageworm, corn earworm, and cabbage looper. J Econ Entomol65:264. 35. Kelly TJ, Coudron TA (1990):Total and specificecdysteroids in the haemolymph of Trichoplusia ni (Lepidoptera Noduidae) and its parasite, Euplectrus plafkypenue (Hymenoptera: Eulophidae). J Insect Physiol36:463. 36. Laemmli UK (1970): Cleavage of structural proteins during assembly of the head bacteriophage T4. Nature 227680. 37. Morrissey JH (1981): Silver stain for proteins in polyacrylamide gels: A modified procedure with enhanced uniform sensitivity. Anal Biochem 117307. Storage Proteins in Envenomed Larvae 109 38. Jones G, Hiremath ST, Hellmann GM, Rhoads RE (1988): Juvenile hormone regulation of mRNA levels for a highly abundant hemolymph protein in larval Trickoplusiuni. J Biol Chem 263:1089. 39. Jones G, Manczak M, Horn M (1993): Hormonal regulation and properties of a new group of basic hemolymph proteins expressed during insect metamorphosis. J Biol Chem 268:1284. 40. Soldevila AI, Jones D (1991): Immunoanalysis of unique protein in Trickoplusiu ni larvae parasitized by the braconid wasp Chelonus near curimaculutus. Insect Biochem 21:845. 41. Jones D (1985): Endocrine interaction between host (Lepidoptera) and parasite (Cheloninae: Hymenoptera). Is the host or the parasite in control? Ann Entomol SOCAm 78(2):141. 42. Beckage NE, Riddiford LM (1982): Effects of parasitism by Apunteles congvegufus on the endocrine physiology of the tobacco hornworm Munducu sextu. Gen Comp Endocrinol47308. 43. Lawrence PO, Baker RC, Tsai LW, Miller CA, Schooley DA, Geddes LG (1990): Arch Insect Biochem Physiol 13:53. 44. Kumaran AK, Ray A, Tertakian JA, Memmel NA (1987): Effects of juvenile hormone, ecdysteroids and nutrition on larval hemolymph protein gene expression in Galleria rnellonella. Insect Biochem 171053. 45. Tojo S, Kiguchi K, Kimura S (1981): Hormonal control of storage protein synthesis and uptake by the fat body in the silkworm Bombyx mori. J Insect Physiol27491. 46. Izumi S, Tojo S, Tomino S (1980): Translation of fat body mRNA from the silkworm, Bombyx mori. Insect Biochem 10:429. 47. Tojo S, Morita M, Agui N, Hiruma K (1985): Hormonal regulation of phase polymorphism and storage-protein fluctuation in the common cutworm Spodoptera litura. J Insect Physiol 31:283. 48. Webb BA, Riddiford LM (1988):Synthesis of two storage proteins during larval development of the tobacco hornworm, Munduca sexta. Dev Biol130:671. 49. Ray A, Memmel NA, Kumaran AK (1987):Developmental regulation of the larval hemolymph protein genes in Galleriu mallonella. Wilhelm Rowc Arch Dev Biol 196:414. 50. Pau R, Levenbook L, Bauer AC (1979):Inhibitory effect of B-ecdysone on protein synthesis by blowfly fat body in vitro. Experientia 35:1449.