to imago, emergence (imago flies emerging within 3â4 h were collected). The method of Hammock and Sparks. [13] was used to estimate the degradation of ...
Doklady Biological Sciences, Vol. 376, 2001, pp. 72–74. Translated from Doklady Akademii Nauk, Vol. 376, No. 3, 2001, pp. 427–429. Original Russian Text Copyright © 2001 by Gruntenko, Monastirioti, Rauschenbach.
GENERAL BIOLOGY
Juvenile Hormone Metabolism in Drosophila melanogaster Imago is Controlled by Biogenic Amines N. E. Gruntenko, M. Monastirioti, and I. Yu. Rauschenbach Presented by Academician V.K. Shumnyi April 12, 2000 Received April 12, 2000
It is well known that energy metabolism in insects is controlled by biogenic amines, which also serve as releasing factors for secretion of other hormones (for review, see [1, 2]). For example, it was shown that octopamine and dopamine regulate biosynthesis and secretion of juvenile hormone in representatives of the orders Ortoptera and Lepidoptera [3–5]. However, little is known about the mechanisms of interaction between these hormones in higher Diptera. It was shown in our studies on stress response in Drosophila that exposure of fruit fly imagoes to stress caused parallel changes in the system of biogenic amines (an increase in the level of octopamine and dopamine [6, 7]) and in the juvenile hormone system (inhibition of juvenile hormone degradation [8]). These findings suggest that metabolism of juvenile hormone in Diptera is also controlled by biogenic amines. To test this suggestion, we studied juvenile hormone degradation in the D. melanogaster mutants with modified levels of biogenic amines. The D. melanogaster strains without mutations of the system of biogenic amines (wild-type and laboratory strains) were used as control groups. The goal of this work was to study the efficiency of juvenile hormone degradation under normal and stress conditions in female wild-type D. melanogaster and in mutants with double amount of dopamine or an inhibited biosynthesis of octopamine. Six strains of D. melanogaster were used: (1) TβhnM18, which is zero-mutant for the tyramine-βhydroxylase gene. Because tyramine-β-hydroxylase catalyzes octopamine biosynthesis from tyramine, the mutant flies are completely deficient in octopamine [9]; (2) Ste, which is mutant for the ebony gene of N-β-alanyldopamine synthetase. This mutation causes doubling of dopamine content [10, 11]; (3) the Canton S wild-type strain; (4) the 921500 wild-type strain isolated from a Gorno-Altaisk natural population; (5) the vermilion (v) laboratory strain; and (6) the First Multiple Seven (FM7) laboratory strain [12]. Drosophila cul-
Institute of Cytology and Genetics, Siberian Division, Russian Academy of Sciences, pr. Lavrent’eva 10, Novosibirsk, 630090 Russia
tures were grown at 25°C using the standard nutrient medium. The cultures were synchronized with respect to imago, emergence (imago flies emerging within 3–4 h were collected). The method of Hammock and Sparks [13] was used to estimate the degradation of radioactive juvenile hormone. Statistical significance of experimental results was estimated using Student’s t-test. The degrees of degradation of juvenile hormone in 24-h-old female TβhnM18 mutants, Ste mutants, and Canton S wild-type flies (control) measured under normal conditions (25°ë) are shown in Fig. 1. It follows from Fig. 1 that the rate of the juvenile hormone hydrolysis in Canton S females is significantly lower than in TβhnM18 and higher than in Ste females (p < 0.001 in both cases). However, the difference between TβhnM18, Ste, and Canton S females could also be interpreted in terms of Rate of juvenile hormone hydrolysis, pmol/min per specimen 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0 Ste Canton S M18 Fig. 1. Hydrolysis of 3H-labeled juvenile hormone in 24-h-old females of the Canton S, TβhnM18, and Ste strains of D. melanogaster under normal conditions. M18 designates TβhnM18.
0012-4966/01/0102-0072$25.00 © 2001 MAIK “Nauka /Interperiodica”
JUVENILE HORMONE METABOLISM IN DROSOPHILA MELANOGASTER IMAGO Rate of juvenile hormone hydrolysi, pmol/min per specimen 4.0
Stress reactivity, % 45
3.5
35
3.0
30
40
25
2.5
20
2.0
15
1.5
10 5
1.0
0
0.5 0
73
921500
Canton S
FM7
v
M18
Ste
v
Fig. 2. Hydrolysis of 3H-labeled juvenile hormone in 24-h-old females of the 921500, Canton S, v, and FM7 strains of D. melanogaster under normal conditions.
interstrain polymorphism. To exclude (or confirm) this suggestion, we compared the efficiency of juvenile hormone hydrolysis in Canton S females and in females of three other strains of independent origin: the wild-type strain 921500 and two laboratory strain (v and FM7), neither of which had mutations of the system of biogenic amines. The results of these experiments are shown in Fig. 2. It is seen from Fig. 2 that there was no statistically significant difference between the rates of juvenile hormone hydrolysis in the four strains of D. melanogaster studied. Therefore, the high rate of juvenile hormone degradation in TβhnM18 females and the low rate of degradation in Ste females might be caused by the corresponding mutations. The question arises as to whether these changes in the levels of biogenic amines affect the stress-induced response of the system of degradation of juvenile hormone? To answer this question, we measured the rate of juvenile hormone hydrolysis in females of the six strains exposed to heat stress (38°ë, 3 h). The stress reactivity of the system was determined as the percentage of the activity decrease in each animal exposed to stress relative to the mean value of this parameter in the sample of the given strain under normal conditions (Fig. 3). It follows from Fig. 3 that both the laboratory strains without mutations of the system of biogenic amines (FM7 and v) and wild-type strains (921500 and Canton S) are characterized by similar levels of stress reactivity (the difference between the strains was statistically nonsignificant). On the other hand, the levels of stress reactivity in strains TβhnM18 and Ste were significantly lower than in Canton S (p < 0.001 in both cases). It was shown by Thompson et al. [5] that octopamine inhibits biosynthesis of juvenile hormone in the female Diploptera punctata cockroaches. If this also were true for D. melanogaster, the absence of octopamDOKLADY BIOLOGICAL SCIENCES
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Fig. 3. Stress reactivity of the system of juvenile hormone degradation in 24-h-old females of the 921500, Canton S, v, FM7, TβhnM18, and Ste strains of D. melanogaster. M18 designates TβhnM18.
ine in the fruit flies would cause an increase in the rate of production of juvenile hormone. To maintain the titer of the hormone at the level sufficient to support reproduction, the increase in the rate of production of juvenile hormone should be accompanied by a corresponding increase in the rate of juvenile hormone hydrolysis [14]. The results of our experiments show that the rate of juvenile hormone degradation in octopamine-deficient females (strain TβhnM18) is almost twice as high as in wild-type females (Fig. 1). The results obtained with Ste females also indicate that biogenic amines regulate juvenile hormone metabolism in Drosophila. Indeed, as noted above, the dopamine content in ebony mutants is two times higher than in wild-type fruit flies [11]. If dopamine, like octopamine, were an inhibitor of juvenile hormone production in Drosophila, the rate of juvenile hormone degradation in dopamine-deficient Ste females would be decreased to compensate the juvenile hormone production inhibition. This was indeed observed in our experiments: the rate of juvenile hormone hydrolysis in Ste females was almost half as high as in Canton S females (Fig. 1). Studies of stress reactivity of the system of juvenile hormone degradation provided another evidence for interaction between biogenic amines and juvenile hormone. The stress reactivity of the system of juvenile hormone degradation in mutant fruit flies with altered levels of biogenic amines is significantly lower than in flies without these mutations (Fig. 3). ACKNOWLEDGMENTS This study was supported by the Russian Foundation for Basic Research and Foundation for Young
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Researchers, Siberian Division, Russian Academy of Sciences.
7. Rauschenbach, I.Yu., Serova, L.I., Timochina, I.S., et al., J. Insect Physiol., 1993, vol. 39, pp. 761–767.
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1. Evans, P.D., in Comprehensive Insect Physiology, Biochemistry and Pharmacology, New York: Pergamon, 1985, pp. 499–530. 2. Brown, C.S. and Nestler, C., in Comprehensive Insect Physiology, Biochemistry and Pharmacology, New York: Pergamon, 1985, pp. 435–484. 3. Piulachs, M.D. and Belles, H., Comp. Biochem. Physiol., 1989, vol. 94A, pp. 795–798. 4. Granger, N.A., Sturgis, S.L., Ebersohl, R., et al., Arch. Insect Biochem. Physiol., 1996, vol. 32, pp. 449–466. 5. Thompson, C.S., Yagi, K.J., Chen, Z.F., and Tobe, S.S., J. Comp. Physiol., 1990, vol. 160, pp. 241–249. 6. Hirashima, A., Sukhanova, M.Jh., Kuwano, E., and Rauschenbach, I.Yu., Dros. Inf. Serv., 1999, vol. 82, pp. 30–31.
9. Monastirioty, M., Linn, C.E., Jr., and White, K., J. Neurosci., 1996, vol. 16, pp. 3900–3911. 10. Hodgetts, R.B. and Konopka, R.J., J. Insect Physiol., 1973, vol. 19, pp. 1211–1220. 11. Ramadan, H., Alawi, A.A., and Alawi, M.A., Cell Biol. Int., 1993, vol. 17, pp. 765–771. 12. Turner, C. and Wilson, T.G., Arch. Insect. Biochem. Physiol., 1995, vol. 30, pp. 133–147. 13. Hammock, B.D. and Sparks, T.C., Anal. Biochem., 1977, vol. 82, pp. 573–579. 14. Soller, M., Bownes, M., and Kubli, E., Dev. Biol., 1999, vol. 208, pp. 337–351.
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