- Email: [email protected]
Metabolism of juvenile hormone I in Drosophila melanogaster
Car,,,, B,,,~,w,,,Phr.,I,11. 197X.Co/ WA. pp X5,,>X9 Pwqamo~,
Pwr\
Pmrd
,o (;rwr
Hriru,,,
METABOLISM OF JUVENILE HORMONE DROSOPHILA MELANOGASTER*
I IN
T. G. Wn_soNt and L. I. G~LHERT Department
of Biological
Sciences,
Northwestern
University,
Evanston.
IL 60201. U.S.A.
(Receiord 24 June 1977) C3H]-Cls juvenile hormone (JH) was topically applied to larvae and adult Drosophila melanogaster and was later recovered as conjugated hormone and acid-dial. diol. and acid metabolites. 2. Haemolymph was relatively inactive toward JH metabolism. 3. The lack of striking differences in rates of JH metabolism among late third instar larvae and male and female adults suggests that metabolism may not be a principle mode of JH titer regulation in Drosophila. Abstract-l.
MATERIALS
INTRODUCTION
AND
METHODS
Hormone. [7-Ethyl-1,2-3H (N)]-C,, juvenile hormone (JH I), 11.17 Ci/mmole, was obtained from New England Nuclear Corp. Unlabelled JH I (mixed isomers) was obtained from Ayerst Laboratories. Drosophila sfocks. The wild-type Oregon-R strain (see Lindsley & Grell. 1968) was used throughout this study and has been maintained continuously as an inbred line for the past 20 years by Dr. R. C. King at Northwestern University. Newly-eclosed adults were weighed. and males were found to weigh 213 of females. Flies were raised on
Insect development depends to a great extent on the exposure of target tissues to critical titers of moulting hormone and juvenile hormone (JH) at specific stages. The JH titer depends in part on the rate of JH degradation, particularly by haemolymph esterases (Weirich et al., 1973; Sanburg et al., 1975; Kramer & DeKort, 1976; Nowock & Gilbert, 1976; Vince & Gilbert, 1977; see Gilbert et al., 1976; Nowock et al., 1976). Indeed, Whitmore et al. (1972) found that JH injected into Hyalophora gloueri pupae induced the biosynthesis of JH-specific esterases. The primary pathways of in oioo metabolism of radiolabelled JH are ester hydrolysis, epoxide hydration, and conjugate formation (Slade & Zibbitt, 1972; Ajami & Riddiford, 1973; Hammock et al., 1975). The rates of JH degradation vary in different orders of insects, being relatively rapid in the Lepidoptera and slow in the Diptera (Slade & Zibitt, 1972). Recent studies of Drosophila melanogaster have demonstrated that vitellogenesis is JH-dependent (Postlethwait & Weiser, 1973) and that JH is released about 12 hr after adult eclosion (Postlethwait et al., 1976). Vitellogenesis begins soon thereafter, and vitellogenic oocytes are noted 15-18 hr after eclosion. Vitellogenesis, presumably under continued support by JH, continues unabated for several weeks in wellfed females. We wished to determine if JH metabolism varied in different developmental stages of Drosophila; i.e. stages requiring JH (vitellogenic females), stages in which JH might disrupt normal development (late third instar larvae), or stages in which JH has no proven function (adult males). Clearcut differences in the rates of JH degradation among insects at these stages could indicate a role for degradative enzymes in the regulation of JH titers in Drosophila.
a standard yeast-cornmeal-agar diet. The term “late third instar larva” as used in this report refers to larvae crawling on the walls of the culture bottle. a behavioral pattern displayed several hr prior to pupariation. Hormone applicarion and extraction. [3H]-JH I (0.1 pmolel dissolved in 0.5 ~1 of nanograde acetone was applied topically with a I-PI Drummond microcap to the ventral side of the abdomen of each cold-anesthetized individual. The insects recovered after 1-2 min and were placed in a holding vial. After incubation for various lengths of time at room temperature. the treated individuals were dipped twice in separate acetone baths. which removed >99”, of the unpenetrated hormone. Penetrated hormone refers to label which is not removed from [3H]-JH treated flies after four dips in acetone. Groups of 10 individuals were homogenized in 1 ml of 2:l ethyl ether-methanol with a glass homogenizer held in an ice bath. The homogenizer was rinsed 3 x with 1 ml of homogenizing solvenr. and I ml of I”,, (NH.,)ZSO, was then added. After vigorously vortexing the mixture. the ether layer was removed. and the aqueous layer re-extracted 3 x with ether. Further ether extractions removed only an additional 5”,, of labelled material. The ether extracts were combined. partially dried over Na,SO,. and evaporated to dryness under NZ. The container vial was rinsed 2 x with ether and 2 x with acetone. and each extract was spotted together with cold standards on a plastic backed silica gel IB-F TLC sheet (Baker-Flex). Following development in 4: 1 benzeneethyl acetate (twice. IOO”, irrigation) and 9: 1 benzene-npropanol (once. 6@-70”” irrigation). the strip was visualized under U.Y. to locate the standards and then cut into 1 cm strips and counted in liquid scmtillation fluid. Recovery of isotope from the extraction procedure was 75-85”,,. Radiolabelled JH carried through this extraction procedure showed no significant breakdown. Except for the initial homogenization step at 0 C. all procedures were carried out at room temperature (22-24 C). Haemolymph merabolism srudies. Haemolymph was collected by capillary action in a I-PI microcap from an inci-
*Supported by grants AM 02818 from the National Institutes of Health and PCM 76-03620 from The National Science Foundation. t Post-doctoral fellow of the National Science Foundation (74-19929) and National Institutes of Health (ES05050-01). Present address: Center for Pathobiology, University of California, Irvine, CA 92717, U.S.A. 85
Sh Table
I.
regions
Radioactivity immediately
extracted from after C3H]-JH abdomen
Body region
~)~~~s(~~~II~~I hod) II/
I application Counts,min
---~Legs Head and thorax Abdomen
sion beneath the wing base in adults or near the midline in larvae. Care was taken to produce a shallow incision to prevent puncturing the gut. A volume of 0.50~1 of haemolymph was collected from 5 to 20 individuals. depending on the ease of collection. Haemolymph was transferred directly to another 1 ~1 microcap containing [‘HI-JH deposited on the inside wall following acetone evaporation. By gently forcing the haemolymph back and forth in the microcap with a rubber bulb for 60~90sec. >95”, of the 13H]-JH “dissolved” in the haemolymph. This haemolymph was finally transferred to a third l-111 microcap and incubated in a humid environment at room temperature. The haemolymph was then spotted onto a TLC sheet together with 4 acetone rinses of the microcap. The TLC sheet was developed and counted as described in the previous section. Mrtaholire idmtijiccltion. JH acid was synthesized by base hydrolysis of JH I (Popjak & Cornforth. 1960). JH diol by acid hydrolysis tSIade & Zibitt. 1972). and JH aciddial by base hydrolysis of JH dial. Each metabohte was purified on preparative silica gel TLC plates (Analtech) and stored in hexane at -20 C. JH actd and JH acid-diol were confirmed by methylation of an aliquot with diazomethane and comparison of the products with JH and JH diol after 2-dimensional TLC. JH diol was reacted with
0
I
I
Time of
Incubation
(mid
20
I40 SYXO
n-butyl boronic acid. which spectfically reacts with cis-hydroxyl groups (Hammock et ul.. 1974). and chromatographicdily compared with unreacted JH dial. Neither JH nor JH acid exhibited an increased R, value upon reaction with n-butyl boronic acid as did JH dioi [R, 0.08 increased to 0.59 when devjeloped in a solvent system of bemene propanol (9: 1)I. The usual positions of the metabolttes and standards after TLC were: fraction 2 tortgin). conJugated metabolites. fracions 4 and 5. acid dial; fractions 7 and R. dial: fractions 10 and II. acid: and fractions 14 and 15. JH I. Fraction numbers refer to l-cm strips of the 20-cm TLC sheet. We failed at several attempts to recover sufficient quanttties of JH metabotites for derivatization from TLC plates following topical application of radiolabelled JH I to Drosophila Therefore. we relied on cochromatography of labelled Drosophila metabolites with standards after 2-dimensional TLC (Hammock ef ui.. 1975) to establish identity of the metabolic products.
RESL:LTS L~cu~i~~f~o~ C$ u~piie~ h~r~~~e. Hormone applied to adults in acetone solution tends to spread over the entire abdomen as judged by its wet appearance. but not across the thorax. When flies are divided into three body regions and analyzed for radioactivity immediately after hormone application. 97”, of the applied hormone is confined to the abdomen (Table 1). Penetrurion of JH. The penetration of [‘HI-JH topically applied to adult females (as a function of time) is shown in Fig. l(a). Penetration appears to be discontinuous. being relatively rapid during the first 15 min after application and then tapering off thereafter. If the rate of penetration is examined as a function of adult age. it can be seen that penetration is most rapid in newly eclosed females (Fig. 1b). Pre-
Adult Age (hr)
0 Fig. t. (a) Penetration of r3H]-JH topically applied to &2-hr-old adult female Drosophila. (b) Rate of penetration of [‘HI-JH topically applied to adult Drosophila of various ages. Time of incubation was 30 min. Each value represents the average of two determinations derived from separate cultures of Oregon-R flies. 0, female; A. male.
24
48
72
Adult Age (hr) Fig. 2. Percentage of label recovered in the extraction aqueous phase after topical application of [)H]-JH to adults of various ages. Figure legends same as for Fig.
I Ibl.
Juvenile
hormone
metabolism
in Drosophila
87
300 E u” 200
0 loo
24 Adult
5
L
,,.
IO Fraction Number
oa
15
@
obtained (6-9%) during periodic examination by TLC of the C3H]-JH used in this study. Therefore, if excreted JH in Drosophila is conjugated or metabolized as in Locusta, little or no hormone is excreted within 1 hr of topical application. Fate of penetrated JH. Radioactivity which penetrated during the incubation period partitioned into either an aqueous or organic phase during extraction, depending on its solubility in ethyl ether. TLC of an aliquot of the aqueous phase resulted in ~95% of the label remaining at the origin on the chromatogram. This radioactivity probably reflects JH conjugates similar to those found in other insects (Slade & Zibitt, 1972; White, 1972). The distribution of label into aqueous and organic phases was measured as a function of adult age (Fig. 2). Newly eclosed adults show a much higher percentage of label incorporated into the aqueous phase. Male and female adults have similar percentages of aqueous label despite their weight differences. For late third instar larvae 34% of the penetrated label was recovered in the aqueous phase. Analysis of the organic phase by TLC revealed several peaks of radioactivity (Fig. 3). In addition to label which remained at the origin of the chromatogram, 3 peaks were seen which cochromatographed with cold acid-diol, diol and JH standards. A distinct peak cochromatographing with acid was usually not
sumably, penetration is facilitated at this stage of development by a soft, newly tanned cuticle. The reason for the difference in penetration between newly eclosed male and female flies is not readily apparent, but may be a reflection of different rates of cuticle maturation. A value of 24% penetration for C3H]-JH applied to late third instar larvae was measured for a 30-min incubation period. Excretion ofapplied hormone. Generally, l&20% of the applied label was found in the holding vial after a 1-hr incubation period. This could have been a result of the larvae contacting the vial as they crawled around and the adults contacting the vial during grooming. Alternatively, the label could represent excretory products. Since excretory products in Locusta have been shown to consist mainly of metabolites and conjugates (Erley et al., 1975) TLC of the Drosophila holding vial label was carried out to determine the chromatographic identity. Eight per cent of the label from vials holding larvae and 5% from vials holding adults did not chromatograph with the JH standard. These values are within the range
2. Metabolites
extracted
from C3H]-JH Metabolite
Stage of development
Acid-diol
Third instar larva O-2 hr adult female 24 hr adult female
0.019 (6) 0.038 (5) 0.044 (15)
Figures
in parentheses
72
Fig. 4. Percentage of the extracted organic phase which chromatographed as JH after topical application of C3H]-JH to adults of various ages. Figure legends same as for Fig. l(b).
20
Fig. 3. TLC of radioactivity in the extracted organic phase. [-‘HI-JH was topically applied to 10 O-2-hr-old female adults; extraction followed a 30-min incubation period. The chromatographic positions of the cold standards are indicated by the circles: acid-dial, 3-4; dial. 7; acid, 8-9; and JH, 13. Fraction 2 is the origin.
Table
46 Age (hr)
treated
larvae
and adult
females
quantity per 10 individuals (pmole) Diol Acid JH
0.005 (l)* 0.030 (4) 0.041 (14)
0.008 (2)* 0.010 (l)* 0.027 (9)
0.092 (41) 0.037 (5) 0.035 (12)
refer to: counts/min
in metabolite
total counts/min
extracted
fraction -from carcass
X 100.
Time of incubation was 30 min; each value represents the average of two determinations on ten individuals derived from separate cultures, *Indicates no discernible peak of activity, but fractions were higher than background and presumably reflect low levels of these metabolites.
T. G. WILSON AND L. I. CrlLaI,Hr
RX
Table 3. Metabolism
Developmental
of [3H]-JH by Drosophilrr ~luccr haemolymph
stage
and iLl~r~-
“Metabolitea”
Third instar larka 0 2 hr adult
(“,,i
16
13(.). 14(T) 7(. 1. 4(j)
?4 hr adult
43
Mumlucu \<‘.xtu (Sth instar. 4th day)
Time of incubation was 2 hr. except that incubation with Munducu haemolymph was 1 hr. I’” “metabolites” of a zerotime haemolymph incubation was subtracted from the Drosophil~ values in each case. 83”” of the label recovered from the Mtrnducu incubation was JH acid.
discerned, although this area of the chromatogram had higher than background radioactivity. The amount of unmetabolized JH in the organic phase was measured as a function of adult age and was found to be almost constant (Fig. 4). Quantities of metabolities in the organic phase were measured in larvae and adult females (Table 2). All three of the common JH metabolites were present in the stages of development examined, although the quantities of each varied. Third instar larvae appear to have the lowest rate of JH metabolism, judging from the high percentage of unchanged JH that was recovered. Newly eclosed females have a lower percentage of metabolites than 24 hr old females, possibly due to a higher rate of conjugation (Fig. 2). However. the absolute amounts of acid-diol. dial. and JH are about the same for the different aged adults. Although not shown in Table 2. the quantity and distribution of metabolites in males were similar to those in females. Metabolism by harmlymph. When [“HI-JH was incubated with haemolymph from Drosophila at various stages of development. little metabolism was noted. Distinct metabolite peaks were absent after TLC, and the only indication of metabolism was a greater “tailing” of JH. suggesting the formation of small amounts of polar derivatives of JH. The amount of tailing radioactivity in the fractions from the TLC of haemolymph incubations was accumulated and compared among flies in different stages of development (Table 3). As can be seen. metabolism of JH by Drosophila haemolymph is very slight compared to that in Mnrlduca haemolymph.
differences in JH metabolic rates among these stages of development may not be physiologically significant, suggesting that Drosophila may not depend primarily on metabolism to regulate its titer of JH. Using a different experimental approach Erley et al. (1975) concluded that the rates of JH excretion from injected Loc~tsta were similar for insects of different ages and sex. The ether soluble metabolites appear to be the same as reported for other insects (Ajami & Riddiford, 1973). Conjugate formation is appreciable. as Slade & Zibitt (1972) found for Sarcophnga and Ajami & Riddiford (1973) noted for other dipteran insects. Our finding that Drosophila haemolymph does not actively metabolize JH relative to Manduca is similar to observations on Sarcophaga (Weirich & Wren. 1976). Certainly, the JH esterases implicated in regulating JH titers in the haemolymph of Manduca and Leptinotarsa (Nowock & Gilbert. 1976: Kramer & DeKort, 1976) do not seem to play as dominant a role in the Diptera examined thus far. Although the rate of conjugation of the hormone is greater in newly enclosed adults than in older flies (Fig. 2). the quantities of ether-soluble metabolites, including unchanged JH. are similar for the two age groups (Table 2). This suggests the existence of steady-state levels of these metabolites prior to conjugation plus some unknown control mechanism. A steady-state Row of metabolites would explain the perplexing observation that newly eclosed adult males and females exhibit different rates of JH penetration but similar conjugation rates and levels of JH and metabolites. REFERENCES
DISCXSSION
Tnis study was initiated with the intent of comparing JH metabolism rates in Drosophila rn~/anogastrr during different stages of development. Several estimates of JH metabolism rate were used: conjugate formation, quantity of extracted unchanged JH, and “metabolite” formation by haemolymph. If the rate of metabolism is based on conjugate formation and “metabolite” formation by haemolymph, then it appears that third instar larvae and newly eclosed adults have slightly higher metabolic rates than older adults (Fig. 2 and Table 3). If quantities of unchanged JH are compared. however. the adults have a higher rate than third instar larvae (Fig. 4). The observed
AJAMI A. M. & RIDDIFOKI) L. M. (1973) Comparative metabolism of the Cecropia juvenile hormone. ./ Inst~r P/IVsiol. 19, 635-645. ERLEY D.. SOUTHARD S. & EMMERICH H. (1975) Excretion of juvenile hormone and its metabolites in the locust. Locusto miyratoria. J. Inswt Phpiol. 21. 61 70. GILBERT L. I., GOODMAN W. & NOWO~K J. (1976) The possible roles of binding proteins in Juvenile hormone metabolism and action. In il~tuulites wr /es Hormonrs d’lnrutuhrrs. pp. 413-434. Colloques mternationaux C.N.R.S. No. 251. Paris. HAMMOCK B. D.. GILL S. S. & CA~~VAJ. E. (1974) Synthesis
and morphogenetic activity of derivatives and analogs of aryl geranyl ether juvenoids. J. agrrc’. Fd Chrm 22, 379-385.
Juvenile
hormone
metabolism
HAMMOCK B., NOWOCK J., GOODMAN W.. STAMOUDIS V. & GILBERT L. I. (1975) The influence of hemolymphbinding protein on juvenile hormone stability and distribution in Manduca sexta fat body and imaginal discs in vitro. Mol. cell. Endocrin. 3, 167-184. KRAMER S. J. & DEKORT C. A. D. (1976) Age-dependent changes in juvenile hormone esterase and general carboxyesterase activity in the hemolymph of the Colorado potato beetle. Leptinorarsa decemlineara. Mol. cell. Endocrin.
4, 43-53.
LINDSLEY D. L. & GRELL E. H. (1968) Genetic variations of Drosophila melanogaster. Carnegie Inst. of Wash. Publ. No. 627. Washington. D.C. NOWOCK J. & GILBERT L. I. (1976) In cirro analysis of factors regulating the juvenile hormone titer of insects. In Insertehrate Tissue Culrure: Applications in Medicine. Biology. and Agriculture (Edited by KURSTAK. E. & MARAMOROSCH K.). pp. 203-212. Academic Press. New York. NOWOCK J.. HAMMOCK B. D. & GILBERT L. 1. (1976) The binding protein as a modulator of juvenile hormone stability and uptake. In The Juoenile Hormones (Edited by GILBERT L. I.), pp. 354373. Plenum Press, New York. POPIAK G. & CORNFORTH R. H. (1960) Gas-liquid chromatography of allylic alcohols and related branched-chain acids. J. Chromat. 4, 214-221. POSTLETHWAITE J. H.. HANDLER A. M. & GRAY P. W. (1976) A genetic approach to the study of juvenile hormone control of vitellogenesis in Drosophila melanogaster. In The Jucenile Hormones (Edited by GILBERT L. I.). pp. 449469. Plenum Press. New York.
in Drosophrlu
x9
POSTLETHWAITE J. H. & WEISER K. (1973) Vitellogenesis induced by juvenile hormone in the female sterile mutant apterous-four in Drosophila melanogaster. Nature NYM Biol. 244, 284-285.
SANBURG L. L.. KRAMER K. J., KEZDY F. J.. LAW J. H. & OBERLANDER H. (1975). Role of juvenile hormone esterases and carrier proteins in insect development. Nature 253, 266-267. SLADE M. & ZI~ITT C. H. (1972) Metabolism of Cecropia juvenile hormone in insects and in mammals. In Insecr JurL,ni/c, Hormones (Edited by MENN J. J. & B~KOZA M.). pp. 155-I 77. Academic Press. New York. VINCE R. K. & GILBERT L. I. (1977) Juvenile hormone esterase activity in precisely timed last instar larvae and pharate pupae of Manduca srxra. Insect Biochem. 7, 115-120. WEIRICH G. & WREN J. (1976) Juvenile hormone esterase in insect development. A comparative study. Physiol. zoo/.
49, 341-350.
WEIRICH G.. WKEN J. & SIDDALL J. B. (1973) Developmental changes of the juvenile hormone esterase activity in haemolymph of the tobacco hornworm. Manducu sextu. Insect
Biochem.
3, 397-407.
WHITF A. F. (1972) Metabolism of the Juvenile hormone analogue methyl farnesoate 10.1 I-epoxide in two insect species. L* Sci. II, 201.-210. WHITMORE D.. WHITMORE E & GILBERT L. I. (1972) Juvenile hormone induction of esterases: a mechanism for the regulation of juvenile hormone titer. Proc. ncltn. Acad. Sri. L.S.A. 69, 1592-1595.
Pwr\
Pmrd
,o (;rwr
Hriru,,,
METABOLISM OF JUVENILE HORMONE DROSOPHILA MELANOGASTER*
I IN
T. G. Wn_soNt and L. I. G~LHERT Department
of Biological
Sciences,
Northwestern
University,
Evanston.
IL 60201. U.S.A.
(Receiord 24 June 1977) C3H]-Cls juvenile hormone (JH) was topically applied to larvae and adult Drosophila melanogaster and was later recovered as conjugated hormone and acid-dial. diol. and acid metabolites. 2. Haemolymph was relatively inactive toward JH metabolism. 3. The lack of striking differences in rates of JH metabolism among late third instar larvae and male and female adults suggests that metabolism may not be a principle mode of JH titer regulation in Drosophila. Abstract-l.
MATERIALS
INTRODUCTION
AND
METHODS
Hormone. [7-Ethyl-1,2-3H (N)]-C,, juvenile hormone (JH I), 11.17 Ci/mmole, was obtained from New England Nuclear Corp. Unlabelled JH I (mixed isomers) was obtained from Ayerst Laboratories. Drosophila sfocks. The wild-type Oregon-R strain (see Lindsley & Grell. 1968) was used throughout this study and has been maintained continuously as an inbred line for the past 20 years by Dr. R. C. King at Northwestern University. Newly-eclosed adults were weighed. and males were found to weigh 213 of females. Flies were raised on
Insect development depends to a great extent on the exposure of target tissues to critical titers of moulting hormone and juvenile hormone (JH) at specific stages. The JH titer depends in part on the rate of JH degradation, particularly by haemolymph esterases (Weirich et al., 1973; Sanburg et al., 1975; Kramer & DeKort, 1976; Nowock & Gilbert, 1976; Vince & Gilbert, 1977; see Gilbert et al., 1976; Nowock et al., 1976). Indeed, Whitmore et al. (1972) found that JH injected into Hyalophora gloueri pupae induced the biosynthesis of JH-specific esterases. The primary pathways of in oioo metabolism of radiolabelled JH are ester hydrolysis, epoxide hydration, and conjugate formation (Slade & Zibbitt, 1972; Ajami & Riddiford, 1973; Hammock et al., 1975). The rates of JH degradation vary in different orders of insects, being relatively rapid in the Lepidoptera and slow in the Diptera (Slade & Zibitt, 1972). Recent studies of Drosophila melanogaster have demonstrated that vitellogenesis is JH-dependent (Postlethwait & Weiser, 1973) and that JH is released about 12 hr after adult eclosion (Postlethwait et al., 1976). Vitellogenesis begins soon thereafter, and vitellogenic oocytes are noted 15-18 hr after eclosion. Vitellogenesis, presumably under continued support by JH, continues unabated for several weeks in wellfed females. We wished to determine if JH metabolism varied in different developmental stages of Drosophila; i.e. stages requiring JH (vitellogenic females), stages in which JH might disrupt normal development (late third instar larvae), or stages in which JH has no proven function (adult males). Clearcut differences in the rates of JH degradation among insects at these stages could indicate a role for degradative enzymes in the regulation of JH titers in Drosophila.
a standard yeast-cornmeal-agar diet. The term “late third instar larva” as used in this report refers to larvae crawling on the walls of the culture bottle. a behavioral pattern displayed several hr prior to pupariation. Hormone applicarion and extraction. [3H]-JH I (0.1 pmolel dissolved in 0.5 ~1 of nanograde acetone was applied topically with a I-PI Drummond microcap to the ventral side of the abdomen of each cold-anesthetized individual. The insects recovered after 1-2 min and were placed in a holding vial. After incubation for various lengths of time at room temperature. the treated individuals were dipped twice in separate acetone baths. which removed >99”, of the unpenetrated hormone. Penetrated hormone refers to label which is not removed from [3H]-JH treated flies after four dips in acetone. Groups of 10 individuals were homogenized in 1 ml of 2:l ethyl ether-methanol with a glass homogenizer held in an ice bath. The homogenizer was rinsed 3 x with 1 ml of homogenizing solvenr. and I ml of I”,, (NH.,)ZSO, was then added. After vigorously vortexing the mixture. the ether layer was removed. and the aqueous layer re-extracted 3 x with ether. Further ether extractions removed only an additional 5”,, of labelled material. The ether extracts were combined. partially dried over Na,SO,. and evaporated to dryness under NZ. The container vial was rinsed 2 x with ether and 2 x with acetone. and each extract was spotted together with cold standards on a plastic backed silica gel IB-F TLC sheet (Baker-Flex). Following development in 4: 1 benzeneethyl acetate (twice. IOO”, irrigation) and 9: 1 benzene-npropanol (once. 6@-70”” irrigation). the strip was visualized under U.Y. to locate the standards and then cut into 1 cm strips and counted in liquid scmtillation fluid. Recovery of isotope from the extraction procedure was 75-85”,,. Radiolabelled JH carried through this extraction procedure showed no significant breakdown. Except for the initial homogenization step at 0 C. all procedures were carried out at room temperature (22-24 C). Haemolymph merabolism srudies. Haemolymph was collected by capillary action in a I-PI microcap from an inci-
*Supported by grants AM 02818 from the National Institutes of Health and PCM 76-03620 from The National Science Foundation. t Post-doctoral fellow of the National Science Foundation (74-19929) and National Institutes of Health (ES05050-01). Present address: Center for Pathobiology, University of California, Irvine, CA 92717, U.S.A. 85
Sh Table
I.
regions
Radioactivity immediately
extracted from after C3H]-JH abdomen
Body region
~)~~~s(~~~II~~I hod) II/
I application Counts,min
---~Legs Head and thorax Abdomen
sion beneath the wing base in adults or near the midline in larvae. Care was taken to produce a shallow incision to prevent puncturing the gut. A volume of 0.50~1 of haemolymph was collected from 5 to 20 individuals. depending on the ease of collection. Haemolymph was transferred directly to another 1 ~1 microcap containing [‘HI-JH deposited on the inside wall following acetone evaporation. By gently forcing the haemolymph back and forth in the microcap with a rubber bulb for 60~90sec. >95”, of the 13H]-JH “dissolved” in the haemolymph. This haemolymph was finally transferred to a third l-111 microcap and incubated in a humid environment at room temperature. The haemolymph was then spotted onto a TLC sheet together with 4 acetone rinses of the microcap. The TLC sheet was developed and counted as described in the previous section. Mrtaholire idmtijiccltion. JH acid was synthesized by base hydrolysis of JH I (Popjak & Cornforth. 1960). JH diol by acid hydrolysis tSIade & Zibitt. 1972). and JH aciddial by base hydrolysis of JH dial. Each metabohte was purified on preparative silica gel TLC plates (Analtech) and stored in hexane at -20 C. JH actd and JH acid-diol were confirmed by methylation of an aliquot with diazomethane and comparison of the products with JH and JH diol after 2-dimensional TLC. JH diol was reacted with
0
I
I
Time of
Incubation
(mid
20
I40 SYXO
n-butyl boronic acid. which spectfically reacts with cis-hydroxyl groups (Hammock et ul.. 1974). and chromatographicdily compared with unreacted JH dial. Neither JH nor JH acid exhibited an increased R, value upon reaction with n-butyl boronic acid as did JH dioi [R, 0.08 increased to 0.59 when devjeloped in a solvent system of bemene propanol (9: 1)I. The usual positions of the metabolttes and standards after TLC were: fraction 2 tortgin). conJugated metabolites. fracions 4 and 5. acid dial; fractions 7 and R. dial: fractions 10 and II. acid: and fractions 14 and 15. JH I. Fraction numbers refer to l-cm strips of the 20-cm TLC sheet. We failed at several attempts to recover sufficient quanttties of JH metabotites for derivatization from TLC plates following topical application of radiolabelled JH I to Drosophila Therefore. we relied on cochromatography of labelled Drosophila metabolites with standards after 2-dimensional TLC (Hammock ef ui.. 1975) to establish identity of the metabolic products.
RESL:LTS L~cu~i~~f~o~ C$ u~piie~ h~r~~~e. Hormone applied to adults in acetone solution tends to spread over the entire abdomen as judged by its wet appearance. but not across the thorax. When flies are divided into three body regions and analyzed for radioactivity immediately after hormone application. 97”, of the applied hormone is confined to the abdomen (Table 1). Penetrurion of JH. The penetration of [‘HI-JH topically applied to adult females (as a function of time) is shown in Fig. l(a). Penetration appears to be discontinuous. being relatively rapid during the first 15 min after application and then tapering off thereafter. If the rate of penetration is examined as a function of adult age. it can be seen that penetration is most rapid in newly eclosed females (Fig. 1b). Pre-
Adult Age (hr)
0 Fig. t. (a) Penetration of r3H]-JH topically applied to &2-hr-old adult female Drosophila. (b) Rate of penetration of [‘HI-JH topically applied to adult Drosophila of various ages. Time of incubation was 30 min. Each value represents the average of two determinations derived from separate cultures of Oregon-R flies. 0, female; A. male.
24
48
72
Adult Age (hr) Fig. 2. Percentage of label recovered in the extraction aqueous phase after topical application of [)H]-JH to adults of various ages. Figure legends same as for Fig.
I Ibl.
Juvenile
hormone
metabolism
in Drosophila
87
300 E u” 200
0 loo
24 Adult
5
L
,,.
IO Fraction Number
oa
15
@
obtained (6-9%) during periodic examination by TLC of the C3H]-JH used in this study. Therefore, if excreted JH in Drosophila is conjugated or metabolized as in Locusta, little or no hormone is excreted within 1 hr of topical application. Fate of penetrated JH. Radioactivity which penetrated during the incubation period partitioned into either an aqueous or organic phase during extraction, depending on its solubility in ethyl ether. TLC of an aliquot of the aqueous phase resulted in ~95% of the label remaining at the origin on the chromatogram. This radioactivity probably reflects JH conjugates similar to those found in other insects (Slade & Zibitt, 1972; White, 1972). The distribution of label into aqueous and organic phases was measured as a function of adult age (Fig. 2). Newly eclosed adults show a much higher percentage of label incorporated into the aqueous phase. Male and female adults have similar percentages of aqueous label despite their weight differences. For late third instar larvae 34% of the penetrated label was recovered in the aqueous phase. Analysis of the organic phase by TLC revealed several peaks of radioactivity (Fig. 3). In addition to label which remained at the origin of the chromatogram, 3 peaks were seen which cochromatographed with cold acid-diol, diol and JH standards. A distinct peak cochromatographing with acid was usually not
sumably, penetration is facilitated at this stage of development by a soft, newly tanned cuticle. The reason for the difference in penetration between newly eclosed male and female flies is not readily apparent, but may be a reflection of different rates of cuticle maturation. A value of 24% penetration for C3H]-JH applied to late third instar larvae was measured for a 30-min incubation period. Excretion ofapplied hormone. Generally, l&20% of the applied label was found in the holding vial after a 1-hr incubation period. This could have been a result of the larvae contacting the vial as they crawled around and the adults contacting the vial during grooming. Alternatively, the label could represent excretory products. Since excretory products in Locusta have been shown to consist mainly of metabolites and conjugates (Erley et al., 1975) TLC of the Drosophila holding vial label was carried out to determine the chromatographic identity. Eight per cent of the label from vials holding larvae and 5% from vials holding adults did not chromatograph with the JH standard. These values are within the range
2. Metabolites
extracted
from C3H]-JH Metabolite
Stage of development
Acid-diol
Third instar larva O-2 hr adult female 24 hr adult female
0.019 (6) 0.038 (5) 0.044 (15)
Figures
in parentheses
72
Fig. 4. Percentage of the extracted organic phase which chromatographed as JH after topical application of C3H]-JH to adults of various ages. Figure legends same as for Fig. l(b).
20
Fig. 3. TLC of radioactivity in the extracted organic phase. [-‘HI-JH was topically applied to 10 O-2-hr-old female adults; extraction followed a 30-min incubation period. The chromatographic positions of the cold standards are indicated by the circles: acid-dial, 3-4; dial. 7; acid, 8-9; and JH, 13. Fraction 2 is the origin.
Table
46 Age (hr)
treated
larvae
and adult
females
quantity per 10 individuals (pmole) Diol Acid JH
0.005 (l)* 0.030 (4) 0.041 (14)
0.008 (2)* 0.010 (l)* 0.027 (9)
0.092 (41) 0.037 (5) 0.035 (12)
refer to: counts/min
in metabolite
total counts/min
extracted
fraction -from carcass
X 100.
Time of incubation was 30 min; each value represents the average of two determinations on ten individuals derived from separate cultures, *Indicates no discernible peak of activity, but fractions were higher than background and presumably reflect low levels of these metabolites.
T. G. WILSON AND L. I. CrlLaI,Hr
RX
Table 3. Metabolism
Developmental
of [3H]-JH by Drosophilrr ~luccr haemolymph
stage
and iLl~r~-
“Metabolitea”
Third instar larka 0 2 hr adult
(“,,i
16
13(.). 14(T) 7(. 1. 4(j)
?4 hr adult
43
Mumlucu \<‘.xtu (Sth instar. 4th day)
Time of incubation was 2 hr. except that incubation with Munducu haemolymph was 1 hr. I’” “metabolites” of a zerotime haemolymph incubation was subtracted from the Drosophil~ values in each case. 83”” of the label recovered from the Mtrnducu incubation was JH acid.
discerned, although this area of the chromatogram had higher than background radioactivity. The amount of unmetabolized JH in the organic phase was measured as a function of adult age and was found to be almost constant (Fig. 4). Quantities of metabolities in the organic phase were measured in larvae and adult females (Table 2). All three of the common JH metabolites were present in the stages of development examined, although the quantities of each varied. Third instar larvae appear to have the lowest rate of JH metabolism, judging from the high percentage of unchanged JH that was recovered. Newly eclosed females have a lower percentage of metabolites than 24 hr old females, possibly due to a higher rate of conjugation (Fig. 2). However. the absolute amounts of acid-diol. dial. and JH are about the same for the different aged adults. Although not shown in Table 2. the quantity and distribution of metabolites in males were similar to those in females. Metabolism by harmlymph. When [“HI-JH was incubated with haemolymph from Drosophila at various stages of development. little metabolism was noted. Distinct metabolite peaks were absent after TLC, and the only indication of metabolism was a greater “tailing” of JH. suggesting the formation of small amounts of polar derivatives of JH. The amount of tailing radioactivity in the fractions from the TLC of haemolymph incubations was accumulated and compared among flies in different stages of development (Table 3). As can be seen. metabolism of JH by Drosophila haemolymph is very slight compared to that in Mnrlduca haemolymph.
differences in JH metabolic rates among these stages of development may not be physiologically significant, suggesting that Drosophila may not depend primarily on metabolism to regulate its titer of JH. Using a different experimental approach Erley et al. (1975) concluded that the rates of JH excretion from injected Loc~tsta were similar for insects of different ages and sex. The ether soluble metabolites appear to be the same as reported for other insects (Ajami & Riddiford, 1973). Conjugate formation is appreciable. as Slade & Zibitt (1972) found for Sarcophnga and Ajami & Riddiford (1973) noted for other dipteran insects. Our finding that Drosophila haemolymph does not actively metabolize JH relative to Manduca is similar to observations on Sarcophaga (Weirich & Wren. 1976). Certainly, the JH esterases implicated in regulating JH titers in the haemolymph of Manduca and Leptinotarsa (Nowock & Gilbert. 1976: Kramer & DeKort, 1976) do not seem to play as dominant a role in the Diptera examined thus far. Although the rate of conjugation of the hormone is greater in newly enclosed adults than in older flies (Fig. 2). the quantities of ether-soluble metabolites, including unchanged JH. are similar for the two age groups (Table 2). This suggests the existence of steady-state levels of these metabolites prior to conjugation plus some unknown control mechanism. A steady-state Row of metabolites would explain the perplexing observation that newly eclosed adult males and females exhibit different rates of JH penetration but similar conjugation rates and levels of JH and metabolites. REFERENCES
DISCXSSION
Tnis study was initiated with the intent of comparing JH metabolism rates in Drosophila rn~/anogastrr during different stages of development. Several estimates of JH metabolism rate were used: conjugate formation, quantity of extracted unchanged JH, and “metabolite” formation by haemolymph. If the rate of metabolism is based on conjugate formation and “metabolite” formation by haemolymph, then it appears that third instar larvae and newly eclosed adults have slightly higher metabolic rates than older adults (Fig. 2 and Table 3). If quantities of unchanged JH are compared. however. the adults have a higher rate than third instar larvae (Fig. 4). The observed
AJAMI A. M. & RIDDIFOKI) L. M. (1973) Comparative metabolism of the Cecropia juvenile hormone. ./ Inst~r P/IVsiol. 19, 635-645. ERLEY D.. SOUTHARD S. & EMMERICH H. (1975) Excretion of juvenile hormone and its metabolites in the locust. Locusto miyratoria. J. Inswt Phpiol. 21. 61 70. GILBERT L. I., GOODMAN W. & NOWO~K J. (1976) The possible roles of binding proteins in Juvenile hormone metabolism and action. In il~tuulites wr /es Hormonrs d’lnrutuhrrs. pp. 413-434. Colloques mternationaux C.N.R.S. No. 251. Paris. HAMMOCK B. D.. GILL S. S. & CA~~VAJ. E. (1974) Synthesis
and morphogenetic activity of derivatives and analogs of aryl geranyl ether juvenoids. J. agrrc’. Fd Chrm 22, 379-385.
Juvenile
hormone
metabolism
HAMMOCK B., NOWOCK J., GOODMAN W.. STAMOUDIS V. & GILBERT L. I. (1975) The influence of hemolymphbinding protein on juvenile hormone stability and distribution in Manduca sexta fat body and imaginal discs in vitro. Mol. cell. Endocrin. 3, 167-184. KRAMER S. J. & DEKORT C. A. D. (1976) Age-dependent changes in juvenile hormone esterase and general carboxyesterase activity in the hemolymph of the Colorado potato beetle. Leptinorarsa decemlineara. Mol. cell. Endocrin.
4, 43-53.
LINDSLEY D. L. & GRELL E. H. (1968) Genetic variations of Drosophila melanogaster. Carnegie Inst. of Wash. Publ. No. 627. Washington. D.C. NOWOCK J. & GILBERT L. I. (1976) In cirro analysis of factors regulating the juvenile hormone titer of insects. In Insertehrate Tissue Culrure: Applications in Medicine. Biology. and Agriculture (Edited by KURSTAK. E. & MARAMOROSCH K.). pp. 203-212. Academic Press. New York. NOWOCK J.. HAMMOCK B. D. & GILBERT L. 1. (1976) The binding protein as a modulator of juvenile hormone stability and uptake. In The Juoenile Hormones (Edited by GILBERT L. I.), pp. 354373. Plenum Press, New York. POPIAK G. & CORNFORTH R. H. (1960) Gas-liquid chromatography of allylic alcohols and related branched-chain acids. J. Chromat. 4, 214-221. POSTLETHWAITE J. H.. HANDLER A. M. & GRAY P. W. (1976) A genetic approach to the study of juvenile hormone control of vitellogenesis in Drosophila melanogaster. In The Jucenile Hormones (Edited by GILBERT L. I.). pp. 449469. Plenum Press. New York.
in Drosophrlu
x9
POSTLETHWAITE J. H. & WEISER K. (1973) Vitellogenesis induced by juvenile hormone in the female sterile mutant apterous-four in Drosophila melanogaster. Nature NYM Biol. 244, 284-285.
SANBURG L. L.. KRAMER K. J., KEZDY F. J.. LAW J. H. & OBERLANDER H. (1975). Role of juvenile hormone esterases and carrier proteins in insect development. Nature 253, 266-267. SLADE M. & ZI~ITT C. H. (1972) Metabolism of Cecropia juvenile hormone in insects and in mammals. In Insecr JurL,ni/c, Hormones (Edited by MENN J. J. & B~KOZA M.). pp. 155-I 77. Academic Press. New York. VINCE R. K. & GILBERT L. I. (1977) Juvenile hormone esterase activity in precisely timed last instar larvae and pharate pupae of Manduca srxra. Insect Biochem. 7, 115-120. WEIRICH G. & WREN J. (1976) Juvenile hormone esterase in insect development. A comparative study. Physiol. zoo/.
49, 341-350.
WEIRICH G.. WKEN J. & SIDDALL J. B. (1973) Developmental changes of the juvenile hormone esterase activity in haemolymph of the tobacco hornworm. Manducu sextu. Insect
Biochem.
3, 397-407.
WHITF A. F. (1972) Metabolism of the Juvenile hormone analogue methyl farnesoate 10.1 I-epoxide in two insect species. L* Sci. II, 201.-210. WHITMORE D.. WHITMORE E & GILBERT L. I. (1972) Juvenile hormone induction of esterases: a mechanism for the regulation of juvenile hormone titer. Proc. ncltn. Acad. Sri. L.S.A. 69, 1592-1595.