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Properties of copper-dependent o-diphenol oxidase activity in the potato aphid Macrosiphum euphorbiae (thomas).

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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 1.10.3.1, and
monopheno1:oxygen oxidoreductase; EC 1.14.18.1) 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 [4].
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 [5].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 1.10.3.2) 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
[8]. These phloem-sucking insects are largely soft-bodied in all stages of
development, and lack extensive areas of sclerotized cuticle outside of the
appendages [9]. 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 [12].
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 [14], 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
[19], orthopteran [20], and other homopteran [12] phenol oxidases. Best
substrates for the potato aphid enzyme, like lepidopteran [13,16], dipteran
[19], 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 [21]. It is well established that
endogenous proteolytic activators and additions of proteases are effective in
potentiating the phenol oxidases of Diptera [26] 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
[29] and sesamol [30], 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 [9] 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 [31],
but they require experimental evidence. Aphids may retain high levels of
phenol oxidases to retain immunity against endoparasitoids and microbial
pathogens [4]; 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.
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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).
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