Properties of copper-dependent o-diphenol oxidase activity in the potato aphid Macrosiphum euphorbiae (thomas).код для вставкиСкачать
Archives of Insect Biochemistry and Physiology 627-37 (1987) Properties of Copper-Dependent o-Diphenol Oxidase Activity in the Potato Aphid Macrosiphum euphorbiae (Thomas) P a u l J. S k i b a a n d Christopher A. Mullin Pesticide Research Laboratory and Graduate Study Center, Department of Entomology, Pennsylvania State University, University Park Potato aphid Macrosiphum euphorbiae (Thomas) was found t o contain high amounts of o-diphenol oxidase activity. Enzyme activity was largely distributed into the postmitochondrial supernatant from Brij-35 extracted aphids and occurs in a latent form that was activated up t o 45-fold by pretreatment with isopropanol. The aphid enzyme has a broad pH optimum near 6, and utilized L-dopa (K, = 1.4 mM, V, = 348 nmol/min-mg protein), dopamine, and 4 methylcatechol the best out of the twelve substrates tested. In addition, this activity is a typical copper-dependent oxidase in that it is potently inhibited by phenylthiourea (50% inhibition at 30nM) and other copper chelators, including salicylhydroxamic acid. The above properties are common t o most insect tyrosinases. However, the aphid enzyme lacked the o-hydroxylase and laccase components and the optimal activity at higher temperatures that are typical of cuticular tyrosinases of other insects. The high levels of o-diphenol oxidase in aphids compared t o other insects is surprising, since the major function associated with these enzymes, that of melanization and sclerotization of cuticle, is of much less importance t o aphids. The possibility that aphids use this enzyme t o metabolize dietary phenolics i s discussed. Key words: tyrosinase, catechol oxidase, polyphenol oxidase, Aphididae, Homoptera INTRODUCTION Insect tyrosinases (o-diphenol: oxygen oxidoreductase; EC 18.104.22.168, and monopheno1:oxygen oxidoreductase; EC 22.214.171.124) have received much at- Acknowledgments: We thank the Pennsylvania Agricultural Experiment Station (JournalSeries No. 7617) for their support. Paul Skiba is now at the Department of Chemistry, Montana State University, Bozeman, MT 59717. Received March 4,1987; accepted June 2,1987. Address reprint requests to Dr. Christopher A. Mullin, Pesticide Research Laboratory, Pennsylvania State University, University Park, PA 16802. 0 1987 Alan R. Liss, Inc. 28 Skiba and Mullin tention, because of their major roles in cuticular sclerotization [l-31 and melanization reactions associated with wound healing and immunity . Numerous insect species have been studied, although the molecular properties have been best established for dipteran and lepidopteran tyrosinases [5,6].The exceptional levels of insect o-diphenol oxidase activities as compared with other phyla, particularly within the cuticle, have allowed their detection as early as 1902 .A proenzymatic form of the enzyme commonly occurs in the hemolymph and is often processed by proteases, either in the hernolymph or the cuticle, to give the active tyrosinase. This assures the proper timing for melanization, wound healing, and sclerotization reactions, Cuticsince substrates regularly co-occur in the hemolymph and cuticle ular a-diphenol oxidases include both the tyrosinases, which may be derived from the hemolymph enzyme, and laccases (EC 126.96.36.199) that act on both oand p-diphenols [1,2]. Aphids are primary pests of many major crops including small grains, potato, sorghum, and forage legumes. Damage results from imbibing of plant sap, injection of salivary phytotoxins, andlor their vectoring of plant diseases . These phloem-sucking insects are largely soft-bodied in all stages of development, and lack extensive areas of sclerotized cuticle outside of the appendages . Pigmentation is mostly due to polycyclic phenolic quinones circulating in the hemolymph [lo], rather than the usual selective melanization of the cuticle. Well-developed phenol oxidase systems in aphids would thus be unexpected, and this may explain the general absence of study of these enzymes within aphids and other soft-bodied homopterans. Nevertheless, phenol oxidases have been detected within the saliva of aphids and other phloem-sucking insects [ll], although characterization of this enzyme among hornopterans has been limited to the work on armored scales . Recently, we found high levels of o-diphenol oxidase activity within whole body extracts of the potato aphid, Macrosiphum euphorbiue (Thomas). These levels were higher than activities within the hemolymph of representative chewing insects such as lepidopteran larvae and a coleopteran adult (Skiba and Mullin, unpublished). The general properties of this activity are presented here. [n. MATERIALS AND METHODS Aphids A pink morph of the potato aphid that had been field collected locally in the fall of 1983 was continuously reared parthenogenetically on potato (var. Norchip and Katadhin) at 22°C using a long day (1653, L:D). Apterous late nymphs and adults were used for all assays. Chemicals Substrates, detergents, inhibitors, and coenzymes used here were purchased from Sigma Chemical Co., St. Louis, MO, and Aldrich Chemical Co., Milwaukee, WI, except for 1-(4-hydroxypheny1)-2-thiourea (Fairfield Chemical Co., Blythewood, SC) and methylhydroquinine (Eastman Kodak Co., Properties of Aphid o-Diphenol Oxidase 29 Rochester, NY). All other solvents, buffers, and other chemicals were reagent grade or better. Enzyme Preparation and Assay Aphids were collected onto 243-pm controlled pore nylon mesh filters (Tetko, Elmsford, NY), using cold water aspiration followed by a wash with 0.1 M potassium phosphate (pH 6, buffer A). A 20% (WN) homogenate of the aphids in buffer A containing 0.5% Brij 35 was made in smooth glass with five passes of a motor-driven Teflon pestle. After centrifugation at 12,0009for 10 min, the supernatant was removed for enzyme assay. For some studies, this supernatant was further centrifuged at 100,OOOg for 1 h in a Beckman L5-75 ultracentrifuge. All preparations and assay procedures were at ice temperature unless noted otherwise. o-Diphenol oxidase activity was determined by following the continuous oxidation of 3-(3,4-dihydroxyphenyl)-L-alanine(L-dopa) to dopaquinone imine (dopachrome) spectrophotometrically at 475 nm using a molar absorptivity of 3,600 M-' cm-' for the latter 16,131. Enzyme, buffer A, and 15 pl of 2propanol in a total volume of 50 p1 were preincubated on ice for 5 min, and then diluted to 1ml with a warm L-dopa solution in buffer A at a 5 mM final concentration. Inhibitors, when present, were added in 10 pl of ethanol just prior to substrate. Increase in absorbance at 475 nm was followed at 30°C in a Perkin-Elmer Lambda 3B UV-visible spectrophotometer. Activities were calculated from the initial increase in absorbance after compensating for boiled enzyme and solvent controls, and are expressed in nmol of dopachrome formed per min per mg of protein. Protein was determined by the method of Lowry et a1 , using bovine serum albumin as the standard. RESULTS Enzyme Preparation Brij 35, a polyethyleneglycol-lauryl alcohol ether, gave up to a five-fold increase in activity within the 12,OOOg supernatant over that of aphid homogenates lacking this nonionic detergent; hence it was used routinely in the homogenization buffer. This detergent has negligible UV-visible absorbance above 240 nm, and does not directly interfere with 0-diphenol oxidase determinations. Further addition of the detergent to the assay buffer was unnecessary for maintenance of enzyme activity; unexpectedly, some activity was lost (Table 1).The subcellular fate of o-diphenol oxidase from the detergent extracted aphids is shown in Figure 1. Over 90% of the homogenate activity distributed into the postmitochondrial supernatant, of which 70% was soluble and 21% remained particulate-bound. Since the relative specific activity of the 100,OOOg pellet remained nearly as high as the 100,OOOg supernatant, all subsequent characterization was done with the 12,0009 supernatant fraction. Optimal Conditions for In Vitro Measurement of Aphid o-Diphenol Oxidase Most tyrosinases are present naturally in a proenzymatic form that requires some conformational change or proteolytic processing to become fully 30 Skiba and Mullin TABLE 1. Treatments to Optimize In Vitro Activity of Potato Aphid o-Diphenol Oxidase Concentration Addition Percentage of controla YO Prior to assay 2-Propanol Acetone 1-Propanol Tetrahydrofuran Ethanol 2-Methyl-1-propanol 2-Butanol 2-Pentanone Dioxane Methanol 2-Butanone Dimethylsulfoxide No solvent Trypsin Trypsin, 2-propanol During assay Boiled enzyme, 2-propanol No L-dopa Brij 35 20-30 20-30 20 20 20 30 30 20 30 20 20 20 100 f 9 87 7 84 .rt 22 82 20 34 28 f 4 25 17 4 9 + 1 7 4 f l 1f1 2 + 1 0 75 2 + + + - 3 N.F. units 3 units, 30 30 + O f 1 0 + 1 78 f 9 - 0.5 (wh) aAssay with L-Dopa conducted as in Materials and Methods; enzyme preincubations were for 5 min at 4°C with solvents or at 30°C with trypsin. Percentages are means SE for up to 13 determinations, with a 2-propanol activity of 170 nmolimin-mg protein. S 0 20 40 60 80 100 % OF TOTAL PROTEIN Fig. 1. Subcellular distribution of o-diphenol oxidase in potato aphid. Relative specific activity of PI (pellet at 12,OOOg), P2 (pellet at lOO,OOOg), and S (supernatant at 100,OOOg) is its percentage of total hornogenate activity divided by percentage of total protein; averages of two determinations, Properties of Aphid o-Diphenol Oxidase 31 active [4,6]. Various treatments were attempted to maximize the measurement of aphid o-diphenol oxidase. A 5-min treatment with mildly polar solvents such as 2-propanol and acetone greatly potentiated the activity found in detergent extracts of the aphid (Table 1).Highest activation occurred within a 20-30% range of exposure of the enzyme to solvent for both acetone (Fig. 2) and 2-propanol (data not shown), although the enzyme was more sensitive to high concentrations of the 2-propanol than to acetone, where activity with the former decreased to 40% of optimum at a 50% concentration. This up to 45-fold solvent-dependent increase in L-dopa oxidase activity could be markedly reversed by substitu~onof more, or less, polar water miscible solvents such as dimethyl sulfoxide and dioxane respectively, or by attempted proteolytic activation with trypsin (Table 1). A 5-min solvent treatment on ice was superior to that at the incubation temperature. Optimal conditions for measurement of the detergent-solubilized, 2-propanol activated enzyme were then determined. The aphid enzyme has a broad optimum of pH 5.5 to 6.5 in potassium phosphate-citrate buffers (Fig. 3). Oxidation of L-dopa at pH 6 occurs maximally between 25 to 30°C (Fig. 4). Dopachrome formation was linear with time for usually 1.5 min after a short lag period (Fig. 5A), and thus initial velocities were determined during this kinetic phase. Rates were also linear with up to 434 pg of protein per incubation, although a yet unexplained synergistic effect was sometimes observed at higher concentrations of enzyme (Fig. 58). Determination of steady-state kinetic parameters under optimal assay conditions using both a low (109 pg) and a high (434 pg) protein content gave an apparent Km of 1.4 It 0.4 mM and a V, of 348 It 10 nmollmin-mg protein (Fig. 5C); hence suitable saturation of the enzyme occurs with the routine substrate concentration of 5 mM. Substrates and Inhibitors Twelve compounds were investigated as potential substrates of aphid tyrosinase (Table 2), using both rate of oxidation to chromophores at 475 nm % ACETONE (v/v) Fig. 2. Effect of increasing acetone concentration on activation of aphid o-diphenol oxidase. Enzyme was preincubated on ice for 5 min with the indicated amount of acetone, and then assayed as in Materials and Methods. Mean & SE for up to seven determinations. 32 Skiba and Mullin L T PH Fig. 3. Effect of pH on aphid o-diphenol oxidase. Mean f SE for two to six determinations. See text. 250 T 200 - 150 - 100 i 50‘ 10 ’ ‘ ’ ’ ’ 20 ’ ‘ ’ ’ ’ ‘ 30 TEMPERATURE (“C) * ‘ ’ ’ 4h ‘ Fig, 4. Effect of temperature on aphid o-diphenol oxidase. The isopropanol-activatedenzyme was incubated at the assay temperature for 2 min prior to addition of an isothermal substrate solution. Mean & SE for two to seven determinations. and subsequent product-dependent oxidation of NADH to estimate reaction rates. Dopamine (3-hydroxytyramine), 4-methylcatechol, and L-dopa were the best substrates found. L-Dopa was oxidized at least five times faster than the D-enantiomer. All of the substrates oxidized were o-diphenols.The aphid enzyme did not catalyze the o-hydroxylation of a number of L-tyrosine compounds at detectable rates, indicating that the tyrosine hydroxylase component often associated with tyrosinases is either absent or much reduced in the aphid preparation. The lack of oxidation of the p-diphenol, methylhydroquinone, establishes the aphid enzyme as an o-diphenol oxidase and not a laccase. Over 50 compounds were screened as potential inhibitors of aphid odiphenol oxidase (Skiba and Mullin, unpublished). Determinations of 50% Properties of Aphid o-Diphenol Oxidase 800 - 100 33 - 400 - 1 0 3 2 0.0 0.4 0.2 TIME (min) 0.6 PROTEIN (mg) 20 - C 0: Krn = 1.4 rnM rnax I -1 fi 0 . . . * g * 1 * ’ = * 348 nrnol/rnin-mg protein ” * 2 * * ‘ 3 . . . . I 4 Fig. 5. Kinetics of aphid o-diphenol oxidase at 3 O O C . Linearity of activity with time (A), amount of protein (B), and the effect (C) of L-dopa concentration S on the initial velocity V in pmollmin-mg protein using the plot method of Lineweaver-Burk. See text for details. inhibitory concentration (Iso) for five of the best inhibitors are shown in Figure 6. In decreasing order of inhibitor potency, the 150 (pM) values were: phenylthiourea (0.030), 4-hydroxyphenylthiourea (0.27), tropolone (18), 5hydroxydopamine (50), and sesamol (85). At least four of the best seven inhibitors of aphid a-diphenol oxidase, including the two thioureas, salicylhydroxamic acid and 4-phenyl-3-thiosemicarbazide, are known to be strong chelators of copper. Enzyme Stability Although the detergent-solubilized preparations of aphid o-diphenol oxidase lost greater than 70% of their activity within 24 h on ice, much greater stability was obtained if the preparation was immediately stored at -80°C. Under these conditions, only 25% of the original activity was lost over a 6- 34 Skiba and Mullin TABLE 2. Substrate Utilization by Aphid o-Diphenol Oxidase Compound AAlmin-mg protein 475 nm 340 nma Dopamine 4-Methylcatechol L-Dopa D-dopa Caffeic acid Chlorogenic acid Quercetin p-Coumaric acid N-acetyl-L-tyrosine L-tyrosine ethyl ester Pyrogallol Methylhydroquinone 0.80 0.46 0.72 0.09 0.06 0.05 0 0 0 0 0 0 Percentage of L-dopab + + + -0.97 -0.76 -0.51 144 22 108 25 100 14 13 t: 1 9 + 1 7t: 1 0 0 0 0 0 __ C _. __ C - ._ - 0 ._ "Followed the decrease in 6-NADH absorbance included in reaction at 0.1 mM. A NADHenzyme incubation and a NADH-substrate incubation served as controls, both of which gave a AAImin of less than 6% of the complete incubation. bComposite value for enzyme-dependent oxidation of substrate to products absorbing at 475 nm averaged with rates of 0-NADH oxidation by enzyme-generated products; mean t: SE. All compounds at 5 mM. 'High substrate absorbance at 340 nm interfered with assay. 7[ 8 phenylt t ' 7 P 6 5 4 - LOG CONCENTRATION (M) 3 Fig. 6. Log concentration-probit plots for some inhibitors of aphid o-diphenol oxidase. week period, after which the preparation was stable to at least seven months of storage (Fig. 7). DISCUSSION Aphid L-dopa oxidase exhibits characteristics that are similar to copperdependent o-diphenol oxidases from other insect groups. Thus, pH optima of between 6 and 7 have been found for lepidopteran [13,15-181, dipteran , orthopteran , and other homopteran  phenol oxidases. Best substrates for the potato aphid enzyme, like lepidopteran [13,16], dipteran , and orthopteran 120,211 o-diphenol oxidases, include L-dopa, dopamine, and 4-methylcatechol; the apparent K, for L-dopa, as found here, is usually Properties of Aphid o-Diphenol Oxidase 35 b 8 0 5 10 15 20 25 30 35 STORAGE TIME (weeks) Fig. 7. Stability of aphid o-diphenol oxidase with storage time at - 8 O O C . Means of at least two determinations. about 1 mM [18,21], although higher values have been reported for the cuticular enzyme 1131. The aphid phenol oxidase, however, had a strong preference for L-dopa over D-dopa, which is not the case in a cockroach enzyme 120,211. Also, aphid oxidase lacked the o-hydroxylase and laccase (pdiphenol oxidase) components of the activity often present in cuticular [16,18,19]but not necessarily hemolymph activities [20,21] from other insects. The temperature optimum of the aphid preparation was exceptional in being about 20°C lower than that typical of other insect phenol oxidases [12,13,16,19], although an optimum of 25°C has been previously reported for the Spodoptevu littoralis larvae [lq. Insect tyrosinase is thought to be biosynthesized largely in the oenocytoids of the hemolymph as a proenzyme, and then released into the plasma for transport to the cuticle, where most proteolytic activation occurs 122,231. Some controversy remains over the extent to which cuticular forms of the enzyme are processed hemolymph protyrosinases. In the potato aphid, significant amounts of the enzyme remain particulate-bound in the "microsomal" pellet after solubilization with relatively high amounts of the nonionic detergent Brij 35. Although anionic detergents such as sodium dodecyl sulfate and sodium oleate are known to activate tyrosinases from grasshopper egg and frog skin [24,25], we have not directly demonstrated an activator role for Brij 35. It does, however, greatly improve the yield of o-diphenol oxidase obtained in the 12,0009 supernatant of the aphid homogenate. More study is necessary to determine the tissue localization of phenol oxidases in aphids. Aphid phenol oxidase, like most insect tyrosinases, occurs in a latent form that requires activation by solvent, detergent, or proteolytic treatments. Our preliminary studies with trypsin did not yield much success, so solvents were investigated as potential activators. Isopropanol greatly potentiated the aphid enzyme up to 45 times, and was followed in efficacy by acetone, 1propanol and tetrahydrofuran. Previous studies have demonstrated the potency of acetone and some other solvents as activators of tyrosinase in Orthoptera [20,21,25] and other Homoptera 1121, but there appears to be an 36 Skiba and Mullin absence of parallel studies in holometabolous insects such as Lepidoptera. Solvent activation has been reported more effective than proteolytic activation for hemolymph tyrosinase in a cockroach . It is well established that endogenous proteolytic activators and additions of proteases are effective in potentiating the phenol oxidases of Diptera  and Lepidoptera [13,18]. We did not observe an endogenous activator within whole body homogenates of aphids; by contrast, isopropanol was an extremely efficient activator, presumably because it elicited an expedient conformational change in the enzyme. Because of its lower volatility, isopropanol was better than acetone in kinetic studies of the phenol oxidase at elevated temperatures. Aphid o-diphenol oxidase behaves as a typical copper-dependent oxidase in being potently inhibited by phenylthiourea (150 = 30 nM), the best copper chelator and inhibitor known for insect phenol oxidases [16,19,20,27]. Other copper chelators including salicylhydroxamic acid (150 < 50 pM), also strongly inhibited the aphid enzyme in a manner similar to that observed for mushroom tyrosinase 1281, Mushroom tyrosinase inhibitors, including tropolone  and sesamol , were similarly effective on the aphid enzyme. We found that 5-hydroxydopamine is also a good inhibitor of the enzyme. Comparative studies between three species each of aphids and holometabolous insects in our laboratory indicate that apterous aphids have equal or higher o-diphenol oxidase activities than that of gypsy moth larvae, tobacco hornworm prepupae, or Western corn rootworm adults (Skiba and Mullin, unpublished). This is surprising, since the major function assigned to these enzymes, that of melanization and sclerotization of cuticle, is of much less importance to aphids than most other insects, since they lack extensive areas of hardened or melanized cuticle  and resort to phenolic glucosides for their pigmentation [lo]. Alternatively, aphids may use this enzyme to oxidize dietary phenolics and thereby metabolize these plant resistance factors, or to harden the stylet sheath. These roles have been suggested elsewhere , but they require experimental evidence. Aphids may retain high levels of phenol oxidases to retain immunity against endoparasitoids and microbial pathogens ; however, some selectivity may be necessary to preserve the microbial endosymbionts required for the aphid’s life. Determining the adaptive significance of aphid tyrosinases should prove useful in the design of chemical methods for their selective control. LITERATURE CITED 1. Andersen SO: Cuticular sclerotization. In: Cuticule Techniques in Arthropods. Miller TA, ed. Springer Verlag, New York,, pp 18.5-215 (1980). 2. Brunet PCJ: The metabolism of the aromatic amino acids concerned in the cross-linking of insect cuticle. Insect Biochem 10,467 (1980). 3. Hackman RH: Chemistry of the insect cuticle. In: The Physiology of Insecta. Rockstein M, ed. Academic Press, New York, Vol. 6, pp 21.5-270 (1974). 4. Soderhall K: Prophenoloxidase activating system and melanization-a recognition mechanism of arthropods? A review. Dev Comp Immunol6, 601 (1982). 5. Mason HS: Comparative biochemistry of the phenolase complex. Adv. Enzymol 16, 105 (1955). 6. Robb DA: Tyrosinase. In: Copper Proteins and Copper Enzymes. Lontie R, ed. CRC Press, Boca Raton, Vol. 2, pp 207-241 (1984). Properties of Aphid o-Diphenol Oxidase 37 7. Hopkins TL, Morgan TD, Kramer KJ: Catecholamines in haemolymph and cuticle during larval, pupal and adult development of Manduca sexfa (L). Insect Biochem 14, 533 (1984). 8. Blackman, RL, Eastop, VF: Aphids on the World’s Crops: An Identification and Information Guide. John Wiley & Sons, New York, 466 pp (1984). 9. Brey PT, Ohayon H, Lesourd M, Castex H, Roucache J, Latge JP: Ultrastructure and chemical composition of the outer layers of the cuticle of the pea aphid Aqrfkosiphon pisum (Harris). Comp Biochem Physiol 82A, 401 (1985). 10. Thomson RH: Naturally Occurring Quinones. Academic Press, New York, pp 576-633 (1971). 11. Miles PW: The saliva of Hemiptera. Adv Insect Physiol 9, 183 (1972). 12. Ishaaya I: Observations on the phenoloxidase system in the armored scales Aonidiella auranfii and Ckrysomphalus aonidum. Comp Biochem Physiol39B, 935 (1971). 13. Aso Y, Kramer KJ, Hopkins TL, Whetzel SZ: Properties of tyrosinase and dopa quinone imine conversion factor from pharate pupal cuticle of Manduca sexfa L. Insect Biochem 14, 463 (1984). 14. Lowry OH, Rosenbrough NJ, Farr AL, Randall RJ: Protein measurement with the folin phenol reagent. J Biol Chem 193, 265 (1951). 15. Ashida M, Ohnishi E: Activation of prephenol oxidase in hemolymph of the silkworm, Bombyx mori. Arch Biochem Biophys 122,411(1967). 16. Barrett FM: Purification of phenolic compounds and a phenoloxidase from larval cuticle of the red-humped oakworm, Symmerista cannicosfa Francl. Arch Insect Biochem Physiol I, 213 (1984). 17. Ishaaya I: Studies of the haemolymph and cuticular phenoloxidase in Spodoptera litforalis larvae. Insect Biochem 2,409 (1972). 18. Yamazaki HI: Cuticular phenoloxidase from the silkworm, Bombyx mori: Properties, solubiliation and purification. Insect Biochem 2, 431 (1972). 19. Barrett, FM, Andersen SO: Phenoloxidases in larval cuticle of the blowfly, Calliphora vicina. Insect Biochem 11, 17 (1981). 20. Fisher CW, Brady UE: Activation, properties and collection of haemolymph phenoloxidase of the American cockroach, PeripZanefa americana. Comp Biochem Physiol 75C, 111(1983). 21. Preston JW, Taylor RL: Observations on the phenoloxidase system in the haemolymph of the cockroach Leucopkaea maderae. J Insect Physiol 16, 1729 (1970). 22. Aso Y, Kramer KJ, Hopkins TL, Lookhart GL: Characterization of haemolymph protyrosinase and a cuticular activator from Manduca sexfa (L). Insect Biochem 15, 9 (1985). 23. Iwama R, Ashida M: Biosynthesis of prophenoloxidase in hemocytes of larval hemolymph of the silkworm, Bombyx mori. Insect Biochem 16, 547 (1986). 24. Wittenberg C, Triplett EL: A detergent-activated tyrosinase from Xenopus laevis. I. Purification and partial characterization. J Biol Chem 260, 12535 (1985). 25. Bodine JH, Allen TH: Enzymes in ontogenesis (Orthoptera). IV. Natural and artificial conditions governing the action of tyrosinase. J Cell Comp Physiol 12, 409 (1938). 26. Schweiger A, Karlson P: Zum tyrosinstoffwechsel der insekten. X. Die aktivierung der praphenoloxydase und das aktivator-enzym. Hoppe-Seyler’s Z Physiol Chem 329, 210 (1962). 27. Raghavan KG, Nadkarni GB: Tyrosine metabolism in carbidopa and phenylthiourea administered Corcyra cephalonica. Insect Biochem 8, 53 (1978). 28. Rich PR, Wiegand NK, Blum H, Moore AL, Bonner WD Jr: Studies on the mechanism of inhibition of redox enzymes by substituted hydroxamic acids. Biochim Biophys Acta 525, 325 (1978). 29. Kahn V, Andrawis A: Inhibition of mushroom tyrosinase by tropolone. Phytochemistry 24, 905 (1985). 30. Metcalf RL, Fukuto TR, Wilkinson CF, Fahmy MH, El-Aziz SA, Metcalf ER: Mode of action of carbamate synergists. J Agric Food Chem 14, 555 (1966). 31. Miles PW: Redox reactions of hemipterous saliva in plant tissues. Entomol Exp Appl 24, 534 (1978).