Metabolic fate of carbohydrates and lipids during moulting cycle of Philosamia ricini (Lepidoptera: Saturniidae)

Insect Biochem., Vol. 9, pp. 577 to 582. © Pergamon Press Ltd. 1979 Printed in Great Britain

0020-1700/79/1201-0577 $02.00/0

METABOLIC FATE OF CARBOHYDRATES AND LIPIDS DURING MOULTING CYCLE OF P H I L O S A M I A RICIN1 (LEPIDOPTERA: SATURNIIDAE) RADHA PANT a n d SUMAN KUMAR Department of Biochemistry, The University, Allahabad-211 002, India (Received 25 January 1979; revised 17 April 1979)

Abstract--During larval-larval moults, total soluble carbohydrates and glycogen decline 50-60~o and 80-90~o respectively and total lipids increase 20~o. This indicates carbohydrate to be the predominant carbon source of chitin, a participant in energy metabolism as well as a substrate for lipid synthesis. During larval growth between moults, total soluble carbohydrates and glycogen accumulate with concurrent depletion in neutral lipids. This suggests that during this period the latter functions as an energy source for day-to-day activities while the glycogen serves as a reserve fuel. Along with glycogen, trehalose also participates as a substrate for chitin synthesis--interestingly, only during the second moult. In larval-pupal moult increasing quantities of soluble carbohydrates, free sugars and reducing substances including monosaccharides are formed. During larval-larval moult the reserves are mobilized as shown by their depletion. These changes indicate that the accumulated metabolites derived from the larval cuticular chitin are reabsorbed for new larval cuticle formation during the larval-larval moults. The higher citrate concentration observed in the larva (both in feeding and moulting) compared with the pupa suggests that citrate brings about a metabolic block in glucose metabolism possibly by inhibition of phosphofructokinase. Key Word Index: Metabolic fate of carbohydrates, metabolic fate of lipids, moulting cycle, Philosamia ricini

INTRODUCTION CONSIDERABLE work o n the m e t a b o l i s m of iipids a n d c a r b o h y d r a t e s a n d their quantitative changes accentuates their role d u r i n g o n t o g e n y a n d m e t a m o r p h o s i s o f insects (WYATT, 1967; AGRELL a n d LANDQUIST, 1973; PANT et al., 1979; PANT a n d KUMAR, ( u n p u b l i s h e d observation). However, except for a few reports (SRIDHARA a n d BHAT, 1963; ZALUSKA, 1959; BADE a n d WYATT, 1962; PANT et aL, 1978) little i n f o r m a t i o n is available o n the metabolism o f these nutrients d u r i n g moulting. T o o b t a i n some additional i n f o r m a t i o n on the role o f these metabolites in energy metabolism, as substrates for cuticle f o r m a t i o n a n d in h o r m o n a l control the present study was u n d e r t a k e n with the lepidopteran p h y t o p h a g o u s insect Philosamia ricini. MATERIALS AND METHODS Insects. P. ricini larvae were reared in the laboratory as described earlier (PANT and AGRAWAL, 1965). Experimental procedure. Larvae starved for 6 hr and prepupae were picked at random from insect colonies of known age and stage of development. They were weighed, chilled and homogenized with ice-cold, glass-distilled water in a precooled Potter-Elvehjem type of homogenizer at 10~o (w/v) tissue concentration and employed for the various metabolite and enzyme assays. Soluble acid and alkaline phosphatase (EC 3.1, 3.2 and 3.1.3.1) activity was determined according to PANT and LACY (1969). Active phosphorylase (EC 2.4.1.1) activity was assayed as described earlier (PANT and NAUT1YAL, 1974a).

577

The liberated phosphate was estimated by the method of FISKE and SUBBAROW(1925). Soluble trehalase (EC 3.2.1.28) activity was assayed according to DERR and RANDALL (1966) by measuring the rate of trehalose hydrolysis. The glucose released was estimated by the method of NELSON (1944). For the estimation of glycogen and sugars the homogenate was prepared within 2-3 min in cold glass-distilled water and inactivated by the addition of 90~o alcohol to bring the concentration of Et0H to 70?/0within 60 sec of the completion of the homogenization to prevent glycogenolysis and loss of trehalose. Total soluble carbohydrates were determined by the method of TREVELYANand HARRISON(1952). Total free sugars (glucose, mannose, galactose, amino sugars, pentoses and trehalose) and reducing substances including monosaccharides were assayed as described by CARROLet al. (1956) and NELSON(1944). Glycogen was isolated (WIENSand GILBERT, 1967a) and assayed according to CARROL et al. (1956). Trehalose was estimated by the method of WYATT and KALE(1957) taking advantage of the exceptional stability of this disaccharide both in acid and alkali. Citrate was estimated as described by PANT and AGRAWAL (1965) and inorganic phosphate by the method of FISKE and SUaBAROW (1925). Total lipids were extracted from the whole insect by the method of FOLCH et al. (1957). Concentrated extracts of the lipids were redissolved in a known volume of redistilled chloroform and employed for the determination of total lipids, free fatty acids, neutral lipids and phospholipids. Suitable aliquots (0.5-1.0 ml) of the chloroform solution were measured into microbeakers, heated to 40°C in an oven to constant weight giving an estimate of the total lipid. Phospholipid phosphorus was estimated by digesting the lipid samples in perchloric acid (6 M) and the liberated phosphate determined (ALLEN, 1940). An approximate figure

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Fig. I. Changes in total soluble carbohydrates during moulting cycle of Philosamia ricini. Each point represents the mean of three observations in duplicate with upper and lower limits. LM = larval-larval moult; S = spinning (0-36 hr); L-PM = larval-pupal moult.

Fig. 2. Changes in total lipid (0) and neutral lipids (0) during moulting cycle of Philosamia ricini. Each point represents the

for the weight of total phospholipids was calculated by multiplying the lipid phosphorus by 25 (lecithin contains approximately 4~o phosphorus, OSER, 1976). Free fatty acids were estimated colorimetrically by the method of NOVAK (1965). Nelatral lipids were calculated by subtracting the sum of phospholipids and free fatty acids from the total lipids.

with a simultaneous decrease in total lipid (Figs. 1, 2 and 3). This suggests the utilization of lipids for locomotor activities and suggests the conversion of the ingested food to glycogen which is then stored as a reserve nutrient. The high enzymic activity of tri- and di-glyceride lipases (PANT and NAUTIYAL, 1974b; PANT et al., 1978) during feeding period of this insect explains the depletion of the neutral lipids. According to STEPHEN and GILBERT (1970) lipid metabolism in insects is regulated by the juvenile hormone (JH), low concentrations stimulating lipid synthesis in fat body and high concentrations inhibiting, giving rise to an adipokinetic effect (LuSCHER and WYss-HUBER, 1964). Thus it is possible that during the feeding period of the larva the gradual

mean of three observations in duplicate with upper and lower limits. L-M = larval-larval moult: S = spinning (0-36 hr): L-PM = larval-pupal moult.

R E S U L T S AND D I S C U S S I O N During development, total soluble carbohydrates decline just prior to each larval moult of P. ricini by 50-60~o and total lipids increase by 20~o (Figs. 1 and 2). However, during the feeding period of larval development through instars 2--4, total soluble carbohydrate and glycogen both increase significantly Instar I1Tr Inslar 1l"~r Instar I

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Moulting changes in P. ricini

Glycoproteins are considered to be internal vehicles for such transfer (LIPKE et al., 1965). The change in chitinolytic activity supports this view. For instance in B. mori, total chitinolytic activity in the epidermal extracts was low in the intermoult stages but registered sharp peaks during larval and pupal moults (JEUNiAUX, 1961; KIMMURA, 1973). Similar findings have also been reported for Drosophila melanogaster by WINICtJR and MITCHELL(1974). Glycogen (the chief tissue carbohydrate) gradually increases during larval development through second to fifth instar with a significant depletion at successive larval moults as shown by the reciprocal relationship to active phosphorylase (Fig. 5). Total free sugars but not reducing substances including monosaccharides also fluctuate in a similar manner (Fig. 4). Accumulation of both glycogen and free sugars during feeding period of P. ricini could be attributed to the increase in amylase activity and gluconeogenesis (PANT and MORRIS, 1969a, b). In P. ricini the glycogen content falls rapidly during the first day of spinning but remains practically constant during subsequent larval-pupal transformation. Total soluble carbohydrates, total free sugars and reducing substances are enhanced significantly along with a simultaneous increase in total lipids from 3 days after the commencement of spinning (Figs. 2 and 4). BADE and WYATT (1962) observed that fully formed pupal cuticle contains about 50~o chitin originating from larval cuticle, no attempt being made to account for the remaining 50%. The increase in free sugars, trehalose and reducing substances during larval-pupal transformation observed in the present investigation suggests that they may originate from this remaining 50% of the larval cuticle. Trehalose (Fig. 5) although present in considerable amount just prior to the second moult, falls to a low concentration which is maintained through the subsequent instars. The fall is associated with a high trehalase activity (Fig. 5). From the above observations, it is clear that glycogen and trehalose contribute to the carbon chain of the chitin. However, the requirement for nitrogen may be fulfilled by glycogenic amino acids as suggested by a significant depletion in total free amino acids observed during moults of this insect (PANT and KUMAR, unpublished data).

depletion in lipids was due to a corresponding increase in the JH. The synthesis of lipids at ecdysis may be associated with the low concentration of JH that occurs at this time. Total soluble carbohydrates and lipids (Figs. 1 and 2) in the fifth instar larva both increase, an effect also shown by glycogen and neutral lipids which may act as potential reserve sources for pupal-adult development (Figs. 2 and 3). On commencement of spinning, both lipids and carbohydrates are utilized for energy and for silk spinning as suggested by their depletion during this period. This supports the observations of BADE and WYATT (1962). However, during the subsequent larval-pupal transformation (days 4 and 5) both these metabolites again increase (Figs. 1 and 2), possibly due to hydrolysis of insoluble larval cuticular chitin, the only precursor available in quantity. This speculation is based on the findings of several investigators where chitin functions as a reserve during moulting (CROMPTON and" BIRT, 1967; ZALUSKA, 1959). In Hyalophora cecropia some 80-85~o of the chitin is digested and resorbed during the larval-pupal stage as well as in the pupal-adult moults (PASSONEAUand WILLIAMS, 1953). Further, the role of chitin as a reserve nutrient is emphasized by LOCKE(1964) who noted that resorption of endocuticle occurs during prolonged starvation and its deposition is resumed on repletion. In Periplaneta americana, synthesis and degradation of cuticular polysaccharides occurred continuously during moulting thus emphasizing the metabolic mobility of the cuticle (LIPKr et al., 1965). Of the total carbohydrates, about 209/0 are utilized for lipid synthesis during larval ecdysis while the remainder fulfils energy needs and serves as a substrate for chitin synthesis. If however, there is an additional requirement for the de novo synthesis of chitin, it has to be met by some other potential sources of carbon and nitrogen. BADE and WYATT (1962) injected [14C]glucose into the late fourth instar H. cecropia and detected the radioactivity largely in the cuticle of the freshly ecdysed fifth instar larva. Again, when fifth instar larvae are transformed into pupae, the radioactivity was once again observed to be transferred and retained by the pupal cuticle. Such specific transfer might perhaps, involve the degradation of the old cuticle and subsequent resorption for incorporation into the new one.

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Fig. 5. C h a n g e s in trehalase activity (O) and trehalose c o n c e n t r a t i o n (O) d u r i n g m o u l t i n g cycle of Philosamia ricini. Each point represents the m e a n of three o b s e r v a t i o n s in d u p l i c a t e with u p p e r and lower limits. L M = l a r v a l - l a r v a l moult; S = spinning (0-36 hr); L - P M = l a r v a l - p u p a l moult.

Accumulation of citrate from endogenous sources may regulate glycogen metabolism through its inhibitory effect on the hepatic phosphofructokinase (UNDERWOOD and NEWSHOLME, 1965). Throughout larval development changes in the citrate level parallel those of glycogen (Figs. 3 and 6). Curiously, the higher citrate concentration observed in the larva over pupa suggests that citrate brings about a metabolic block in glucose metabolism via inhibition of phosphofructokinase. Alternatively, citrate may direct increased quantities of glucose-6-phosphate for incorporation into chitin whilst glycogen undergoes degradation during successive moults. The depletion in citrate during both spinning and larval-pupal transformation (Fig. 6) could be due to enhanced catabolism as reported earlier from this laboratory (PANT and AGRAWAL, 1965; PANT and SHARMA, 1967; PANT, 1973). During larval development alkaline phosphatase activity predominates over that of acid phosphatase (Fig. 7). However, at the commencement of spinning and at larval-pupal transformation, the position is reversed. Ample r

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evidence for the localization of alkaline phosphatase in cells where extensive synthesis of fibrous protein occurs, has been presented (DAY, 1949; BRADFIELD, 1951; DENUCE, 1952). In P. ricini alkaline phosphatase activity increases and reaches a peak during each successive larval moult when histolysis of old cuticular tissues and histogenesis of the new one proceed concurrently at a high rate (Fig. 7). The observed high activity of acid phosphatase during larval-larval and larval-pupal ecdysis (Fig. 7) suggests its involvement in active glycogen degradation as reported by several investigators [WIENS and GILBERT 1967b; GOLDSWORTHY, 1970; PANT and MORRIS 1972; PANT and KUMAR (unpublished observation)]. Inorganic phosphorus declines during each successive larval moult (Fig. 6) while phospholipids increase (Fig. 8). This indicates the incorporation of inorganic phosphates into 3 72 - I2 I m" Instar I T~7 Jnst e~ 5 4 1

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Moulting changes in P. ricini

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and biochemical changes during insect development. In Physiology o f lnsecta (Ed. by ROCKSTEIN M.) Vol. 1, 159-233. Academic Press, New York. ALLEN R. J. L. (1940) The estimation of phosphorus. Biochem. J. 34, 858-865. BADEM. L. and WYATTG. R. (1962) Metabolic conversions during pupation of the Cecropia silkworm. 1. Deposition and utilization of nutrient reserves. Biochem. J. 83, 470--477. ~o8 BRADFIELDJ. R. J. (1951) Phosphatases and nucleic acids in silk glands. Cytochemical aspects of fibrillar protein secretion. Q. Jl. microsc. Sci. 92, 87-112. CARROL N. V., LONGLEYR. W. and ROE J. H. (1956) The determination of glycogen in liver and muscle by the use of anthrone reagent. J. biol. Chem. 220, 586-593. CROMPTON M. and BIRT L. M. (1967) Changes in the amounts of carbohydrates, phosphagen and related compounds during metamorphosis of blowfly Lucilia O2 I cuprina. J. Insect Physiol. 13, 1575-1592. DAY M. F. (1949) The distribution of alkaline phosphatase in I~ Jl i illl i III I I insects. Aust. scient. Res. 2, 31-41. LM 2 I 3 LM 3 I 3 LM 4 2 4 6 0 I 5 5 DENUCE J. M. (1952) Recherches sur le systeme A g e in d o y s phosphatasique des glandes sericigenes chez le ver a soie Fig. 8. Changes in free fatty acids (O) and phospholipids (O) (B. mori L.) Experientia 8, 64-65. during moulting cycle of Philosamia ricini. Each point DERR R. F. and RANDALLD. D. (1966) Trehalase of the represents the mean of three observations in duplicate with differential grasshopper Melanoplus differentialis. J. Insect upper and lower limits. LM = larval-larval moult; Physiol. 12, 1105-1114. S = spinning (0-36 hr): L-PM = larval-pupal moult. FISKE C. H. and SUBBAROWY. (1925) The colorimetric determination of phosphorus. J. biol. Chem. 66, 375-400. phospholipids synthesized for the cellular and FOLCH J., LEES M. and SLOANESXANLEYG. H. (1957) A subcellular structures of the growing larva (SRIDHARA simple method for the isolation and purification of total and BHAT, 1965; GREEN and TZAGOLOLOGG, 1966). lipids from animal tissues. J. biol. Chem. 226, 497-509. During the larval-pupal moult inorganic phosphate G]LBYA. R. and Cox M. E. (1963) The cuticular lipids of the increases probably resulting from histolysis of larval cockroach Periplaneta americana (L.). J. Insect Physiol. 9, muscles (CROMPTON and BIRT, 1967; PANT and 671-681. GOLDWORTHYG. T. (1970) The action of hyperglycaemic MORRIS, 1972). factors from the corpus cardiacum of Locusta migratoria Neutral lipids in general, constitute 80-90% of the of glycogen phosphorylase. Gen. comp. Endocr. 14, 78-85. t o t a l lipids over the entire development (Fig. 2). GREEN D. E. and TZAGOLOLOGGA. (1966) Role of lipids in The increase in total lipids which is seen in all the the structure and function of biological membranes. J. other lipid fractions examined, combined with the Lipid Res. 7, 587-602. simultaneous depletion of carbohydrates during JEUNIAXC. (1961) Biochimie de la mue chez les arthropodes successive larval moult suggests that lipogenesis is Bull. Soc. zool. Fr. 86, 590. occurring at the expense of nonlipoidal material as KIMMURA S. (1973) Chitinolytic enzymes in the larval demonstrated by WALKER and BAILEY (1970). The development of the silkworm, B. mori L. (Lepidoptera Bombycidae). Appl. Ent. Zool. 8, 234-236. synthesized lipids may participate in the formation of L1PKEH., GRAVESB. and LETOS. (1965) Polysaccharide and new cuticular layers since presence of appreciable glycoprotein formation in the cockroach. II. quantities of lipids has been detected in the cast off Incorporation of D-glucose-C 14 into bound carbohydrate. cuticle of this insect (PANT and 1974) as in J. biol. Chem. 240, 601-608. several other insects (GILBY and Cox, 1963). Free fatty LOCKE M. (1964) The structure and formation of the acids constituting 5-10% of the total lipids, vary in a integument in insects. In Physiology o f Insects (Ed. by V-shaped manner at every larval moult (Fig. 8). ROCKSTEINM.), Vol. 3, Ch. 7, Academic Press, New York. During fifth instar development the total lipids LUSCHER M. and WYss-HUBER M. (1964) Die adenosin increase gradually and attain the maximum nukleotide im Fettkorper des adulten weibchens von Leucophaea maderae im loufe des sexualzyklus. Revue concentration prior to spinning (Fig. 2). On suisse Zool. 71, 183-194. commencement of spinning until day 3, a significant fall in neutral lipids (Fig. 2) and free fatty acids (Fig. 8) NELSONN. (1944) A photometric adaptation of the Somogyi inethod for the determination of glucose. J. biol. Chem. is observed, suggesting that lipids may be utilized as a 153, 375-380. source of energy for spinning. This observation agrees NOVAK M. (1965) Colorimetric ultra-micro method for the with that of BADE and WYATT (1962) for Hyalophora determination of free fatty acids. J. Lipid Res. 6, 431. cecropia. Phospholipids on the other hand, do not OSER B. L. (1976) Hawk's Physiological Chemistry, 14th show any change during the above period (Fig. 8). edition p. 1061, Tata McGraw-Hill, New Delhi, India. PANT R. (1973) Studies on citrate accumulation in insects. Curr. Sci. 42, 592-595. Acknowledgement--This research has been financed in part by a grant made by the US Department of Agriculture, PANT R. and AGRAWALH. C. (1965) Some quantitative changes observed in Philosamia ricini pupal haemolymph Agricultural Research Service under PL 480. during metamorphosis. Biochem. J. 96, 824-828. PANT R. and LACY P. S. (1969) Phosphatase activity in Philosamia ricini during development. Indian J. exp. Biol. REFERENCES 6, 154-156. AGRELLI. P. S. and LANDQU]STA. M. (1973) Physiological PANTR. and MORRISI. D. 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activity in Philosamia ricini (Eri silkworm) during development. Indian J. Biochem. 6, 156-157. PANT R. and MORRIS !. D. (1969b) Changes in active phosphorylase activity and glycogen content during larval and pupal development of Philosamia ricini. J. Biochem. (Tokyo) 66, 29-31. PANT R. and MORRlSI. D. (1972) Variation in glycogen, total free sugars, protein, alkaline and acid phosphatases, citrate and inorganic phosphorus level in fat body of Philosamia ricini (Eri silkworm) during development. J. Biochem. (Tokyo) 71, 1-8. PANT R. and NAUTIYALG. C. (1974a) Changes in protein, glycogen, free sugar content and active phosphorylase activity during embryogenesis of Philosamia ricini. Proc. Ind. Acad. Sci. 79, 121-126. PANT R. and NAUTIYALG. C. (1974b) Variation in lipase activity in Philosamia ricini during embryogenesis and larval-pupal-adult development. Proc. Indian Acad. Sci. 80, 230-235. PANT R. and SHARMAS. C. (1967) Changes in citrate level in tissues of Philosamia ricini during development with special reference to the pupal stage (Lepidoptera: Saturniidae) Annls ent. Soc. Am. 60, 170-172. PANT R. and SHARMAK. K. (1974) Amino acid composition of the cuticles of Philosamia ricini during various stages of larval development. Indian J. exp. Biol. 12, 192-194. PANT R., GUPTA D. K. and SHARMAB. (1978) Studies on triand di-acylglycerol hydrolases and some esterases (total-, cholesteryl- and cholin-) on larva Of Philosamia ricini during development. Indian J. exp. Biol. 16, 706-708. PANT R., KUMAR S. and S1NGH S. D. (1979) Changes in carbohydrates and lipids during embryonic development of Antheraea mylitta (L.) J. Biosei. 1, 27-33. PASSONEAUJ. V. and WmLtAMSC. M. (1953) Moulting fluid of the ¢k,eropia silkworm. J. exp. Biol. 30, 545-560. SRIDHARA S. and BHAT J. V. (1963) Alkaline and acid phosphatases of the silkworm Bombyx mori. J. Insect Physiol. 9, 693-701.

SR[DHARAS. and BHAl J. V. (1965l Lipid composition of the silkworm Bombyx mori. J. Insect Physiol. 11,449-462. STEPHEN W. f. and GILBERT L. 1. (1970) Alterations in fatty acid composition during metamorphosis of Hyalophora cecropia correlation with juvenile hormone titre. J. Insect Physiol. 16, 851. TREVELYANW. E. and HARRISONJ. S. (1952) Studies of yeast metabolism. 1. Fractionation and micro-determination of cell carbohydrates. Biochem. J. 50, 298-303. UNDERWOODA. H. and NEWSHOLMEE. A. (1965) Properties of phosphofructokinase from rat liver and their relation to the control of glycolysis and gluconeogenesis. Biochem. J. 95, 686-875. WALKER P. R. and BAILEY E. (1970) Changes in enzyme associated with lipogenesis during development of the adult male desert locust. J. Insect. Physiol. 16, 679-690. WIENS A. W. and GmBERT L. I. (1967a) Variations in glycogen content of fat body, ovary and embryo during the reproductive cycle of Leucophaea maderae. J. Insect Physiol. 13, 587-594. WIENS A. W. and GILBERT L. I. (1967b) Regulation of carbohydrate metabolism and utilization in Leueophac~ maderae. J. Insect Physiol. 13, 779-794. WINICUR S. and MITCHELL H. K. (1974) Chitinase activity during Drosophila development. J. Insect Physiol. 20, 1795-1805. WYATT G. R. (1967) The biochemistry of sugars and polysaccharides in insects. Advances in Insect Physiology' (Ed. by BEAMENT J. W. L., TREHERNE J. E. and WIGGLESWORTH V. B.), Vol. 4, pp. 28%347, Academic Press, New York. WYATT G. R. and KALFG. F. (1957) The chemistry of insect haemolymph. If. Trehalose and other carbohydrates. J. gen. Physiol. 40, 833-847. ZALUSKAH. (1959) Glycogen and chitin metabolism during development of the silkworm (Bombyx mori L.) Acta biol. exp. Lodz. 19, 339-351.