38, 255-266 (1990)
Aphid Carboxylesterases: Biochemical Aspects and Importance the Diagnosis of Insecticide Resistance’ YEHIA
A. I. ABDEL-AAL,*
A. WOLFF,RICHARD P. LAMPERT*
M. ROE,* AND
Plant Protection Department, College of Agriculture, Assiut University, Assiut, A.R., Egypt, and Department of Entomology, Box 7613, North Carolina State University, Raleigh, North Carolina 276957613 Received July 5, 1990; accepted August 29, 1990 The activity of general carboxylesterases from susceptible and malathion-resistant strains of the green peach aphid, Myzus persicae, and the tobacco aphid, M. nicotianae, was examined using 10 I-naphthohc esters, The activity showed a parabolic relationship with the number of carbon atoms in the substrate acyl moiety. The maximal activity was obtained with I-naphthyl propionate and I-naphthyl butyrate, respectively, for the above species. No interstrain differences in the general profde of the structure-activity relationship were observed. However, an elevated carboxylesterase activity in the resistant strains when compared to the reference-susceptible strain was observed with each of the tested substrates, and the bestowed activity correlated with the degree of malathion resistance. The zymograms and intensity of the carboxylesterase bands when resolved by wide range isoelectric focusing (PH 3.5-9.5) and stained with Fast Blue B salt using I-naphthyl acetate, 1-naphthyl propionate, and 1-naphthyl caproate as substrates correlated favorably with the general activity. From the above results a falter paper esterase test was developed and used for the diagnosis of the level of resistance in individual aphids. This test showed an excellent agreement with the susceptibility bioassay test of malathion against the green peach aphid. However, the test did not make possible an unambiguous discrimination between susceptible and resistant tobacco aphids. The possible impact of this and other more specific tests on the management of aphid populations is discussed. o IWOacademic press, IIIC.
In the past 50 years more than 500 arthropod species have become resistant to the toxicological action of insecticides (1). This number is likely to be underestimated because information on pesticide resistance in many economically important arthropods is lacking. The mechanism of resistance is complex and sometimes varies in different biotypes of the same species, even for the same insecticide. There are at least two toxicologically significant mechanisms of insecticide resistance. These are target site resistance and metabolic resistance (2). A great deal of information is available on three major enzyme systems which influ’ The test described in this manuscript is currently under negotiation between NCSU research and Agdia for any future commercial rights. 2 To whom reprint requests should be addressed.
ence the toxicity of insecticides, i.e., microsomal polysubstrate monooxygenases, glutathione transferases , and hydrolases . The latter group is the easiest group to study both under in vivo and in vitro conditions for several technical and biochemical reasons. With the overlapping substrate specificity of detoxifying hydrolases, resistance can sometimes be related to carboxylesterase activity toward surrogate, noninsecticidal substrates. Since the adaptation of Gomori’s assay technique (3) to study insect esterases (4), naphthyl acetates have been frequently used for the diagnosis of resistant biotypes (5). Both the intensive use of chemical control and the introduction of resistant plant cultivars have resulted in the development of new adapted biotypes of aphids. In this respect the genus Myzus seems to have made the broadest ecological adjustment.
ow8-3575Bo$3.00 Copyright 0 1990 by Academic Press, Inc. All rights of reproduction in any form ~served.
Resistance in one species, M. persicae, has been reported from 31 countries involving a total of at least 69 different insecticides representing organochlorines, organophosphates (OPS),~carbamates, and pyrethroids (6). In general, insecticide-resistant biotypes are not avoidable, even in species which reproduce entirely by parthenogenesis. Esterase activity appeared to be the most important mechanism of insecticide resistance in all cases studied in the green peach aphid (GPA), M. persicae, in England (7). Biochemical, serological, and nucleic acid probe assays (8) were successfully used to study the structure of the GPA populations and their dynamics in relation to the agrochemical ecosystem in Great Britain. Unfortunately, no comparable results for this and other aphid species are available in the United States. The tobacco aphid (TA) has been recently recognized as a new species, M. nicotianae Blackman (9), and is presumably isolated from the GPA by being permanently parthenogenetic (9, 10). This recognition has created several controversial issues regarding the evolutionary origin of the TA and whether or not its resistance to insecticides has resulted from an evolutionary duplication of some structural gene(s) present in its presumed taxonomic origin, the GPA. The answers to some of these questions require comparative biochemical and molecular studies. The results from this study represent some biochemical comparisons of carboxylesterases in a susceptible and two OP-resistant strains from each of the above two species found in the United States. A simple filter paper esterase test was also developed and used successfully to distinguish between individuals of susceptible and OP-resistant GPA. 3 Abbreviations used: BSA, bovine serum albumin; FB, Fast Blue B salt; FG, Fast Garnet GBC salt; GPA, green peach aphid; OPs, organophosphorus insecticides; IEF, isoelectric focusing; RAE, resistance associated esterase; R, resistant; S, susceptible; SAR, structure-activity relationship; TA, tobacco aphid.
ET AL. MATERIALS
This new ELISA reader was purchased, along with the SOFTmax software, from Molecular Devices Corp. (MDC), Palo Alto, California. Version 2.01 of SOFTmax supports the temperature regulation features of the instrument. The temperature of the microplate chamber is regulated, displayed on the screen, and updated every 6 set, thereby providing accurate kinetic measurements. Falcon Pro-Bind assay plates (96 flat-bottom we&) were purchased from Becton-Dickinson and Co., Lincoln Park, New Jersey. Chemicals
Fast Blue B salt, dodecyl sulfate sodium salt, and I-naphthol were obtained from Aldrich, Milwaukee, Wisconsin. Anhydrous mono- and di-basic sodium phosphate, Triton X-100, Fast Garnet GBC salt, and naphtholic esters, except for lnaphthyl propionate, were from Sigma, St. Louis, Missouri. I-Naphthyl propionate was synthesized previously (11). Certified A.C.S. organic solvents used in the present study were obtained from Fisher Scientific, Fair Lawn, New Jersey. The Bio-Rad dye reagent was from Bio-Rad Laboratories, Richmond, California. All the commercial reagents were of technical grade with the highest purity available and were used without any further purification. Aphids
The insects used in the present study were from a susceptible (S) and two OPresistant (R) strains each of the green peach aphid, M. persicae (GPA), and the tobacco aphid, M. nicutiunae (TA). The susceptible strain of the green peach aphid was obtained through the courtesy of Dr. Susan Halbert, College of Agriculture, Southeast Idaho Research gi Extension Center, Department of Plant, Soil, and Entomological Sciences, University of Idaho, Parma, Idaho. This strain was maintained on pep-
per plants under standard greenhouse conditions for our work. Two other strains of GPA were collected from pepper and sweet potato plants that were grown in separate greenhouses, under routine insecticidal pressure, at North Carolina State University. When tested in the laboratory, these strains were found to be resistant to malathion and will be referred to as OP-resistant strains. Contamination with other aphids was not completely excluded, especially with the greenhouse strains of the GPA. The susceptible strain and two OP-resistant strains of the TA have been maintained on tobacco plants in the laboratory of E.P.L. at North Carolina State University since 1983 and 1986, respectively. Resistance spectrum to several OPs, including malathion, in the TA strains has been recently reported (12). Except as otherwise stated, apterous adults of homogenous weights from the above strains were used in the present study. Toxicity
Malathion was tested for its toxicity against apterous adults of the GPA using a slide dip technique recommended by FAO and adapted for aphids (12). The 24-hr LCsO values were 91, 323, and 13,445 ppm, respectively for S, R,, and R2 strains. The corresponding values against the TA from a previous study (12) were 24, 84, and 92 ppm, respectively. The S and R, strains of the TA were green and R2 was red. Microplate
Assay for Esterases
Aphids were homogenized using a glass homogenizer in an ice bath at a fresh weight concentration of 1.36 mg/ml in a sodium phosphate buffer (0.2 M; pH 7.4), containing 0.15% (w/v) Triton X-100. The homogenate was then centrifuged at 10,000, for 15 min at 4°C and the supernatant was kept at -70°C until used for enzyme assay. The supernatant was always diluted in the above buffer and the dilution factor was dependent on the enzyme source, the substrate used, and the incubation time. When
prepared and stored as above, no change in activity was observed during a period of several months. Carboxylesterase activity was measured in 96 flat-bottom well microplates using lnaphtholic esters as substrates in an endpoint assay. The assay was a modified and miniaturized procedure of the original assay (3) that was previously adapted for insect esterases (4). To each of the first 27 wells, 75 ~1of sodium phosphate buffer (0.2 M; pH 7.4) was added. An extra 100 pl of buffer was added to each of the first three wells that served as controls for the standards that follow. A series of seven freshly prepared concentrations of l-naphthol was added in 100 p+lof the above buffer to each of the next 24 wells (3 wells/concentration). The final micromolar concentration of lnaphthol ranged from 18.75 to 150. The rest of the microplate (69 wells) was used for the enzymatic and nonenzymatic hydrolysis of the substrate. For the enzymatic reaction, 75 ~1 of the enzyme preparation in sodium phosphate buffer (0.2 M; pH 7.4) was added to the wells. Wells for the nonenzymatic reaction received 75 p,l buffer instead of the enzyme solution. The reaction was then started by adding 100 ~1 of 0.001 M of substrate in the working buffer. The microplate was immediately automixed and incubated in the Thermomax reader for 5-30 min at 30°C. The incubation period was chosen so that the activity was within the linear initial rate for the enzyme and substrate concentrations used. At the end of the assay period, 25 ~1 of 0.8% (w/v) Fast Blue B salt and 3.4% (w/v) dodecyl sulfate sodium salt in distilled water was added to each well, and the plate was returned to the Thermomax chamber for automixing and incubation. Ten minutes later, the absorbance was read at 595 nm. The Thermomax reader coupled with version 2.01 of SOFTmax permits the automatic calculation of the standard curve between I-naphthol concentrations and the absorbance of the diazo1-naphtholic dye and the calculation of esterase activity.
analysis was used to calculate the initial rates. The specific activity was the mean of Apterous adults from each strain were at least six replicates and two separate dehomogenized at 20 mg fresh wt/ml distilled terminations. In most of the enzyme and water containing 0.1% Triton X-100, and protein measurements, the standard deviathe homogenate was then centrifuged at tion was less than 5% of the mean. 5000, for 5 min. The supematant was applied to LKB Ampholine PAGplate (pH Filter Paper Assay for Esterases range 3.5-9.5) at 5 @/lane and the gel was This test was based on some modificafocused at 5°C and a constant power of 25 tions published procedures (14-17). W for 1.5 hr on an LKB 2117 Multiphor II Twelveof apterous adults from each of the electrophoresis unit (Bromma, Sweden). tested strains were placed in a 1Zcavity After electrofocusing, the gel was incu- porcelain plate, one aphid per cavity. Fifty bated in 0.02% (w/v) 1-naphthyl acetate in microliters of sodium phosphate buffer (0.2 sodium phosphate buffer (0.2 M; pH 7.4) M; pH 7.4) was added to each cavity and for 30 min. The gel was then gently washed the aphids were homogenized using the botin water and incubated for 20 min in 0.13% tom of a clean, 13 x lOO-mm test tube. The (w/v) Fast Blue B salt in distilled water. The homogenate from each cavity was then stained gel was subsequently washed and transferred to a marked 10 x 75-mm test stored in distilled water at 4°C. The inten- tube and the tube was then kept in an ice sity of the esterasebands was measured us- bath. After all the individuals had been hoing an LKB 2202 UltroScan laser densitom- mogenized, 5 or 10 ~1 of each of 10-36 hoeter. The results from the absorbance mea- mogenates was withdrawn and blotted onto surements at 633 nm were displayed on a a rectangular strip (17 by 5.5 cm) of Whatlinear chart recorder. The peak height for man No. 1filter paper. The filter paper was the densitometric tracing was used to quan- immersed in 0.1% (w/v) of substrate in sotify the band intensity. dium phosphate buffer containing l-2% (v/ v) ethanol for 1 or 2 min. The filter paper Microplate Assay for Protein was then briefly blotted between tissue paThe protein assay was based on the orig- pers and transferred to a 0.15% (w/v) soluinal procedure of Bradford (13). A microas- tion of FB or FG in distilled water for 4 min. say technique was developed so that either The presence of esterase was revealed by bovine serum albumin (BSA) as a standard the development of purple color which then or the unknown protein sample was added turned dark blue/grey at the site of the samin 50 pl of sodium phosphate (0.2 M; pH ple deposit. The filter paper strip was 7.4) buffer to 200 ~1 distilled water contain- briefly dipped in water, then a 10% acetic ing 25% (v/v) Bradford reagent in the wells acid solution, immediately blotted between of a microplate. BSA was used in seven tissue paper, and allowed to dry in the dark. different concentrations ranging from 12.5 The dried strip was kept between two layto 100 p&ml sodium phosphate buffer (0.2 ers of aluminum foil in a plastic bag. Dark M; pH 7.4), three replicates each. After and cold storage stabilized the color indefadding the Bradford reagent, the plate was initely. FB took less than 4 min for color introduced to the Thermomax chamber for development, while FG required longer inautomixing and absorbance measurements cubation. However, FG was less sensitive at 595 nm, 10 min after the addition of the to light and the color developed was more stable. Therefore, FG was used in our labBradford dye. A IZchannel micropipeter was used to oratory for routine filter paper esterase astransfer the substrate and the chromogenic say tests, while FB was used for the mireagents to the microplate. Least-squares croplate assay where the speed of color Electrofocusing
development was critical for accurate quantitative measurements. Except for R, (Duplin green) of the TA, individual aphids used in this test had an average weight of 228-400 kg/aphid. It has always been observed that Duplin green aphids of the same physiological age are always smaller in size as compared to R2 (Duplin red) and S (Clayton old) of the TA. In order to quantify the results obtained in the filter paper assay, 10 different concentrations of 1-naphthol, with less than 1.5-fold increment intervals, were prepared by diluting a stock naphtholacetone solution in sodium phosphate buffer. Five microliters from each concentration was then applied to a filter paper in a row of three replicates and the strip was processed in exactly the same way as for the esterase test. The final concentrations ranged from 0.07 to 1.44 t&spot. The reference and esterase tests were examined for the staining intensity of deposits using an LKB 2202 UltroScan laser densitometer as described above. RESULTS
6 8 l m g protein
100 l m g protein
FIG. 1. The activity of general carboxylesterases from the green peach aphid, Myzus persicae, and the tobacco aphid, hi. nicotianae, toward IO naptholic esters. The endpoint microassay was used to monitor the release of I-naphthol at 30°C. Fast Blue B salt and sodium dodecyl sulfate were added at the end of the reaction to couple the released naphthol and the absorbance was measured at 595 nm. The specific activity was the mean of at least six replicates and two determinations. In general, the standard deviation was less than 5% of the mean of specific activity.
The activity of carboxylesterases from the susceptible and two OP-resistant strains of the GPA and the TA toward 10 naphtholic esters is shown in the histograms of Fig. 1. For the resistant and susceptible TA, the specific activity appears to gradually increase as the number of acyl carbons (n) increases from 2 (acetate) to 4 (butyrate). The maximal activity was followed by a slight decrease when 1-naphthyl valerate (n = 5) and 1-naphthyl caproate (n = 6) were used as substrates. However, a drastic decrease (about five-fold, as compared with the caproate) in the activity of the enzymes from all three strains of the TA was observed when I-naphthyl caprylate (n = 8) was used. For substrates with a longer acyl moiety, i.e., 1-naphthyl nonanoate (n = 9), 1-naphthyl caprate (n = lo), lnaphthyl laurate (n = 12), and 1-naphthyl myristate (n = 14), there was a gradual de-
crease in the specific activity as the length of the acyl moiety increased. My&ate had no detectable activity, even at higher homogenate concentrations, and longer incubation periods. In the case of general carboxylesterases from the GPA, the highest activity was observed with 1-naphthyl propionate (n = 3) one carbon shorter than the best substrate for the enzymes from the TA. The activity in the susceptible strain of the GPA was higher even than the two resistant strains of the TA. This high activity enabled us to determine the activity for the whole series, including myristate. As shown in Fig. 1, there are no interstrain differences in the general profile of the structure-activity relationship @AR) in both aphid species. Nevertheless, resistance to malathion was always linked to high carboxylesterase activity toward all the tested substrates.
Esterase activity from homogenates of the tested strains were resolved on isoelectric focusing (IEF) gels using a wide range (pH 3.5-9.5). The resolved esterase isozymes were visualized using a naphtholic ester-fast blue staining protocol. When lnaphthyl acetate, 1-naphthyl caproate (data not shown), and I-naphthyl propionate (Fig. 2A) were used as substrates, the band intensities paralleled the SAR profile (Fig. l), with I-naphthyl propionate being the most reactive substrate to the IEF-resolved isozymes. The data presented in Fig. 2 also confirmed the information obtained from the total esterase activity (Fig. 1)in that the homogenate from resistant aphids exhibited darker bands than those of the reference susceptible strain. There are at least three esterases (see arrows in Fig. 2) that are A
associated with resistance (resistance-associated esterases, RAEs) in the GPA. The most anodic RAEs in this species appear to be a multiple of at least two isozymes. Only one isozyme is apparent in this region in resistant tobacco aphids. This anodic RAE is appreciably missing in the susceptible individuals of TA. It is rather interesting that the cathodic RAEs are only associated with one resistant strain (R2) of the TA, while the other strain (R,) shows similar band intensity of those isozymes as the susceptible strain. The latter strain (R,) showed a green body coloration while the former (R2) was red. The question as to whether the high activity of this particular esterase is associated with body coloration requires further study since in both casesthe numbers sampled were too small for a definite conclusion to be drawn. The scanned IEF gels (Fig. 2B) confirm the above observation for
FIG. 2. (A) Aphid carboxylesterase isozymes separated by wide range @H 3.5-9.5) isoelectric focusing and stained for esterase activity using I-naphthyl propionate and Fast Blue B salt. Lanes from left to right are the following. For M. persicae: susceptible (S, lane I) and resistant (R, lane 2 and R,, lane 3); for M. nicotianae: resistant (R,, lane 4 and R,, lane 5) and susceptible (S, lane 6). (B) Densitometric tracings of the IEF gel described above. Band intensity was detected in terms of the peak heightfor the densitometric scan at 633 nm. Note that the zone with the question mark represents the esterase isozyme which is apparently missing in the susceptible tobacco aphid.
RAE IN Myzus
ample of the filter paper esterase test of individuals from sample populations of susceptible and insecticide-resistant cultures of the GPA. The substrate used to generate this figure was 1-naphthyl propionate and the diazonium salt was Fast Garnet GBC. Figures 3B and 3C are a semiquantification of the actual filter paper spot test using a known concentration of 1-naphthol or the 1-naphthol released from the esterase activity of sample homogenates, respectively. A linear relationship between the amount of I-naphthol and the spot intensity, in terms Esterase Spot Test of the peak height of the absorbance meaShown in Fig. 3A is a representative ex- surements in the laser densitometric trac-
the quantitative and qualitative differences between resistant and susceptible strains of both species. The overall difference in total esterase activity between strains of the GPA and TA (Fig. 1) can be explained mainly by differences in the BAEs (see arrows in Fig. 2). As seen in the densitometric tracing, the esterase zone with the highest retention time split into a doublet in all the tested strains of the GPA. However, the peak of this zone is sharper in resistant TA as compared to that of the GPA.
40 30 20
0.36 0.27 0.18 0.14 0.09 0.07
R2 Rl Individuals FIG. 3. (A) An example of afilterpaper spot test of carboxylesterases from susceptible (S), resistant (R,), and extremely resistant (RJ strains of the green peach aphid. Each spot represents the activity of one-tenth of an individual aphid previously homogenized in SO J of phosphate buffer (0.2 M; pH 7.4). The homogenate was blotted on a strip ofjilter paper and then dipped for 2 min in a mixture of 99 ml of buffer and 1 ml of ethanol containing 0.1 g of I-naphthyl propionate. The released I-naphthol was visualized by immersing the filter paper strip in 0.15% (w/v) Fast Garnet GBC in distilled water. (B) A linear relationship between the concentration of I-naphthol and the spot intensity was obtained when known concentrations of I-naphthol were spotted on thefilterpaper, stained with’t!ze diazonium salt, and scanned for spot intensity at 633 nm. (0 Densitometric quantification (in terms of peak heights of the intensity tracing as scanned at 633 nm) of the spots from individual aphids. Values above bars are means for S, R,, or R,.
ing, up to 0.72 kg/spot, was obtained (Fig. 3B). I-Naphthol concentrations higher than 0.72 pg did not result in a linear increase in intensity. The values for the susceptible and resistant aphids (Fig. 3C) fall in the range of the linear relationship (Fig. 3B). The average peak heights of the tested individuals are depicted above the histograms of Fig. 3C. The standard deviation of these values indicates a variation of less than 25% of the mean, and this value decreases significantly among resistant populations. It is not clear whether the variability is due to differences in individual esterases or to lower sensitivity of the scanning at the lower absorbances. The average peak height (Fig. 3C) appears to be a function of the susceptibility of the tested individual and supports the visual interpretation of the actual filter paper test. There is no conflict among the filter paper test (Figs. 3A and 3C), the activity of the total (Fig. 1) and IEF-resolved (Fig. 2) esterases, and the physiological response of the tested individuals to the effect of malathion. DISCUSSION
A microassay procedure was developed to study the activity of general carboxylesterases in an aliquot of an individual aphid. The procedure was based on the original technique (3) of coupling the released naphthol with azo compounds. The activity of esteraseshydrolyzing 10carboxylic acid esters of 1-naphthol was examined in OP-susceptible and OP-resistant strains of the TA and its presumed taxonomic origin, the GPA. In each of the above species the activity to any of the tested substrates was higher in the resistant than in the susceptible aphids. The specific activity appreciably correlates with the level of resistance to malathion. Beranek and Oppenoorth (18) provided some strong evidence that in an OP-resistant strain of GPA, the enzyme hydrolyzing methyl paraoxon in vitro was identical to the enzyme responsible for the elevated I-naphthyl acetate hydrolyzing activity in this strain. If this applies to the
situation in the strains tested, one could successfully monitor resistance by assaying for the general carboxylesterase activity. The specific activity of esterases from both the GPA and TA showed a parabolic relationship in relation to the number of carbon atoms in the acyl moiety of the l-naphtholic ester. This structure-activity relationship is independent of the level of activity in the aphid strains tested. This, in addition to the finding (8) that gene amplification is the genetic mechanism of elevated carboxylesterase in GPA, may suggest a quantitative rather than a qualitative difference between our aphid strains tested. The esterases from the two species of aphids reacted the same way to all the 10 naphtholic esters tested. These esterases hydrolyzed l-naphthyl propionate and 1-naphthyl butyrate faster than all the other substrates. The activity of the enzymes decreased steadily after the butyrate and markedly after the caprylate. No reaction was detected with myristate, especially with the TA. A similar pattern was previously obtained with the GPA (19). However, this SAR cannot be extrapolated to explain any innate specificity of the aphid esterases in relation to the chemical structure of the tested substrates for at least two reasons. First, the specific activity was measured at high substrate concentrations and represents the activity of several isozymes that may be unrelated. Second, an almost identical SAR was observed for the anodic B-esterases from six species of the nematode Meloidogyne (20). Alternatively, the SAR may be a function of only pure electronic effects of the alkyl chain next to the carbonyl in the acylated enzyme. This does not exclude some hydrophobic and steric effects on the affinity of the enzymes to the tested substrates. The effect on the electrophilicity of the carbonyl carbon in the acyl enzymes may explain the indifferent SAR better since in our study and in that of the nematode esterase study (20), high substrate concentrations were used. Under these conditions (substrate saturation conditions),
different activities may only reflect effects on the catalytic rate constant of the test enzymes. The results of the general esterases (Fig. 1) indicated that the widest activity gap between resistant and susceptible aphids was found when I-naphthyl propionate and lnaphthyl butyrate were used as substrates. Therefore, these two substrates, as well as their 2-naptholic analogs, were tested in the development of the filter paper test. The 2-naphthyl propionate and butyrate did not clearly distinguish between the resistant and susceptible individuals of the two tested species. This was not surprising since in a previous study (21), an OPresistant strain of the GPA showed higher activity than a susceptible one toward lnaphthyl acetate and butyrate but not toward 2-naphthyl acetate or butyrate. On the other hand, 1-naphthyl propionate and butyrate, and to some extent 1-naphthyl acetate, discriminated between susceptible and only one resistant strain (RJ of the TA, i.e., the red form. However, no test other than the body coloration is required for determining the frequency of resistant red TA since no susceptible red TA has ever been observed under field conditions in North Carolina. The test was more appropriate in distinguishing between the susceptible and resistant individuals of GPA with the acetate, propionate, and butyrate of l-naphthyl, in close agreement with their reactivity in the microassay test. The spot intensity in the filter paper test was dependent on the substrate, the amount of homogenate used for each spot, and incubation time in the substrate solution (data not shown). I-Naphthyl propionate, one-tenth of an individual GPA, and an incubation time of 2 min were found to be the best conditions for detecting differences in esterase activity between susceptible and resistant strains (Fig. 3). Under these conditions the low activity GPA variant was not apparent when the color was developed. Therefore, the test for resistance depended on the presence or absence of an easily vi-
sualized spot. This adds to the simplicity of the test since no reference-susceptible individuals are required, and any apparent color development is a sign of higher esterase activity and an anticipation of a resistant individual. The average spot intensity (Fig. 3C) for susceptible, resistant, and extremely resistant cultures of the GPA was 3.25,8.55, and 19.00 mm, respectively. These figures are in excellent agreement with the specific activity (0.71, 2.21, and 8.19 pmol/min/mg) toward 1-naphthyl propionate with a correlation coefficient of 0.989. When increased concentrations of lnaphthol were deposited on a filter paper strip and subsequently exposed to Fast Garnet GBC, a linear relationship was observed between the concentration and the height of the peak of the staining intensity as determined by the densitometer. The fact that less than 1.5 increment intervals are detectable indicates that individual variations in esterase activity in this range can be evaluated by the filter paper esterase test. This test is convenient for rapidly testing total esterase activity and for accurately evaluating the proportion of susceptible and resistant individuals in a field population where resistance is due, at least partially, to detoxification by esterases. In comparing the stained IEF-resolved esterase activity with the intensity of the total esterase activity on filter paper, it appeared that the filter paper test correlated more favorably with the toxicity test. Therefore, the filter paper test can be used routinely with relative ease for the purpose of largescale surveys for resistance. The test can provide a warning by detecting accurately and rapidly the proportion of individuals with extremely high resistance. The biggest advantage of the assay is that because of its simplicity it can be used by greenhouse owners where electrophoresis and spectrophotometric equipment are not available and for advisory work where quantitative data are not critically needed. It does not require a highly qualified staff, it is easy, and the reagents are inexpensive. Occa-
sional confirmation of the filter paper test by bioassays of clones derived from sample populations would be essential in order to ensure that any new level or type of resistance could be detected. When the test is used in the field, samples representing the ecotone should be examined to cover the possible distribution between the extremes of clonal structure and new windbome immigrants of the aphid populations. To the best of our knowledge, this is the first use of this test in the GPA. However, it seems likely that this test is applicable to other species of aphids since in a previous report (17) a test was used with I-naphthyl acetate and the total aphid homogenate to discriminate between susceptible and OP-resistant individuals of the cabbage aphid (Brevicoryne brussicae) and the pea aphid (Acyrthosiphon pisum). In previous studies with filter paper esterase tests (1417, 22, 23), the ratiomue of using surrogate substrates of carboxylesterases, contrary to our test, was not based on any substrate structureactivity relationship. This rational optimization may be critical for obtaining the least ambiguous discrimination between individuals of different phenotypes. The isoelectric focusing studies showed that the change in total esterase activity (Figs. 1 and 3) that correlated with resistance in the GPA was mainly due to changes in similar activities in both resistant and susceptible individuals (Fig. 2). However, the difference between susceptible and resistant TA appears to be both qualitative and quantitative. In particular, one of the RAE bands was noticeably missing in the susceptible TA. Therefore, changes in total esterase activity reflected changes solely in the resistance-associated enzymes. Similar but not identical esterase patterns were obtained in susceptible and pirimicarb-resistant Aphis gossypii when resolved on IEF gels (24) and in resistant and susceptible Phorodon humuli when the activity was resolved on 7.5% polyacrylamide rod gels by electrophoresis (25). The apparent absence of one RAE-
isozyme from the susceptible TA and the clear-cut association between autosomal translocation and the activity of this particular isozyme and insecticide resistance (11) may argue against gene amplification, similar to that found in the GPA (g), as the only mechanism of elevated esterase activity in resistant phenotypes of the TA. Active and null alleles, common for the genes that code for some esterases in other insect species (26, 27), may be present in aphids with translocated and normal autosomes, respectively, and may explain the intensity variation in TA esterases.The role of translocation in the activation of this resistanceassociated esterase through frame shift mutation of some DNA-related sequences remains a possibility. Comparisons of IEF zymograms of TA differing in insecticide resistance revealed almost identical pattern, and no qualitative variation was observed when aphids were examined over a period of more than a year. It is interesting that although R, and R, of the TA were equally resistant to malathion, R, did not show a significant elevated activity over the susceptible strain in one of the resistanceassociated enzymes (Fig. 2). This may be due to differences in the overall mechanism of resistance between the two strains or to differences in alleles encoding for enzymes with the same OP-hydrolyzing activity in R, and R, but with varying hydrolytic capacity toward I-naphthyl propionate. This question is still to be solved since these two strains are not coisogenic, because in addition to the resistance genes they also have different body coloration, i.e., R1 is a green form and R2 is red. Further work is needed to study esterase polymorphism in aphids in relation to insecticide resistance, as well as to any epistatic selective fitness advantage for the gene combination of some of these isozymes. The information from such a study is critical to understanding the persistence of resistance and any expected resurgence of the normal susceptible aphid populations in the absence of insecticide selective pressure. It has been recently
RAE IN Myzus
shown, for example, that resistant phenotypes of one of the species tested possessa high reproductive potential and a shorter life cycle than the susceptible phenotypes in the absence of any intentional exposure to insecticides (28). In conclusion, insecticide resistance in both the green peach aphid and tobacco aphid from the United States is linked to high carboxylesterase activity toward lnaphtholic esters. This linkage relationship was found to be due to some polymorphic, IEF-detectable esterases. Several assays were developed to detect the frequency and level of resistance in individuals of the two species tested. The speed and simplicity of some of these assaysmake them desirable for use under both laboratory and field conditions .
ACKNOWLEDGMENTS Y.A.1.A has been awarded a Rockfeller Biotechnology Career Felloaiship to study the molecular biology of insecticide resistance in aphids. This work was partially funded by grants from RJ Reynolds Tobacco Co, Rhone-Poulenc Ag Co., Valent U.S.A. Corp., and the North Carolina Agriculture Research Service. The authors acknowledge the use of Dr. J. Baker’s greenhouse to rear one of the resistant green peach aphid strains. They are also grateful to Ms. R. Eckel for confirming the taxonomic identities of the strains tested from the two aphid species. The editorial assistance of Doug Anspaugh and David Goodlett was very helpful. This manuscript was reviewed by W. C. Dauterman, G. Kennedy, and G. Rock for the Journal Series of the North Carolina Agricultural Research Service, Raleigh, NC.
REFERENCES 1. L. B. Brattsten, Insecticide resistance: Research and management, Pestic. Sci. 2.6, 329 (1989). 2. R. L. Metcalf, Insect resistance to insecticides, Pestic. Sci. 26, 333 (1989). 3. G. Gomori, Human esterases, J. Lab. Clin. Med. 42, 445 (1953). 4. K. van Asperen, A study of housefly esterases by means of a sensitive calorimetric method, J. Insect Physiol. 8, 401 (1%2). 5. T. M. Brown and W. G. Brogdon, Improved detection of insecticide resistance through conventional and molecular techniques, Annu. Rev. Entomol. 32, 145 (1987). 6. G. P. Georghiou, Overview of insecticide resistance, in “Managing Resistance to Agrochemi-
cals from Fundamental Research to Practical Strategies (M. B. Green, H. M. LeBaron, and W. K. Moberg, Eds.), Chap. 2, pp. l&41. Amer. Chem. Sot. Washington, DC, 1990. G. D. Moores, R. H. ffrench-Constant and A. L. Devonshire, Immunoassay for detecting insecticide resistance in aphids, Pestic. Sci. 26, 324 (1989). A. L. Devonshire, Insecticide resistance in Myzus persicae: From field to gene and back again, Pestic. Sci. 26, 375 (1989). R. L. Blackman, Morphological discrimination of a tobacco-feeding form from Myzus persicue (Suizer) (Hemiptera: Aphididae), and a key to new world Myzus (Nectarosiphon) species, Bull. Entomol. Res. 77, 713 (1987). G. Boiteau and D. T. Lowery, Comparison of a yellow form of the green peach aphid, Myzus persicae (Sulzer), and a green form of the tobacco aphid, Myzus nicotianae Blackman, coexisting on greenhouse potato in New Brunswick, Canad. Entomol. 121, 1029 (1989). Y. A. I. Abdel-Aal, E. P. Lampert, R. M. Roe, and C. D. Harlow, Karyoty& variation in relation to esterase activity and insecticide resistance in Myzus nicotianae Blackman, Acta Phytopathbl. Hung., in press. C. D. Harlow and E. P. Lampert, Resistance mechanisms in two color forms of the tobacco aphid (Homoptera: Aphididae), J. Econ. Entomol., in press. M. M. Bradford, A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72, 248 (1976). K. Ozaki, The resistance to organophosphorus insecticides of the green rice leafhopper, Nephotettix cincticeps Uhler, and the smaller brown planthopper, Laodelphax striatellus Fallen, Rev. Plunt Prof. Res. 2, 1 (1%9). N. Pasteur and G. P. Georghiou, Filter paper test for rapid determination of phenotypes with hiih esterase activity in organophosphate resistant mosquitoes, Mosq. News 41, 181 (1981). N. Pasteur and G. P. Georghiou, Improved filter paper test for detecting and quantifying increased esterase activity in organophosphateresistant mosquitoes (Diptera: Culicidae) . J. Econ. Entomol. 82, 347 (1989). R. Marullo, G. L. Lovei, A. Tallarico, and E. Tremblay, Quick detection of resistant phenotypes with high esterase activity in two species of aphids (Homoptera, Aphididae), J. Appl. Entomol. 106, 212 (1988). A. P. Beranek and F. J. Oppenoorth, Evidence that the elevated carboxylesterase (esterase 2) in organophosphorus-resistant Myzus persicae (Sulz.) is identical with the organophosphate-
hydrolyzing enzyme, Pestic. Biochem. Physiol. 7, 16 (1977). A. P. Beranek, Esterase variation and organophosphate resistance in populations of Aphisfabae and Myzus persicae, Entomol. Exp. Appl. 17, 129 (1974). P. R. Esbenshade and A. C. Triantaphyllou, Partial characterization of esterases in Meloidogyne (Nematoda), Camp. Biochem. Physiol. B83, 31 (1986). K. I. Sudderuddin, An in vitro study of esterases, hydrolysing non-specific substrates, of an OPresistant strain of the green peach aphid, Myzus persicae (Sulz.), Camp. Biochem. Physiol. B 44, 1067 (1973). Z. I. Al-Khatib, Isolation of an organophosphate susceptible strain of Culex quinquefasciatus from a resistant field population by discrimination against esterase-2 phenotypes, J. Amer. Mosq. Control Assoc. 1, 105 (1985). A. T. Rees, W. N. Field, and J. M. Hitchen, A simple method of identifying organophosphate insecticide resistance in adults of the yellow fe-
ver mosquito, Aedes aegypti, J. Amer. Mosq. Control Assoc. 1, 23 (1985). C. Furk, D. F. Powell, and S. Heyd, Pirimicarb resistance in the melon and cotton aphid, Aphis gossypii Glover, Plant Puthol. 29, 191 (1980). G. A. Lewis and D. S. Madge, Esterase activity and associated insecticide resistance in the damson-hop aphid, Phorodon humuli (Schrank) (Hemiptera: Aphididae), Bull. Entomol. Res. 74, 227 (1984). R. M. Roberts and W. K. Baker, Frequency distribution and linkage disequilibrium of active and null esterase isozymes in natural populations of Drosophila montana, Amer. Nat. 107, 709 (1973). N. Pasteur, G. Pasteur, F. Bonhomme, J. Catalan, and J. B&ton-Davidian, “Practical Isozyme Genetics,” Ellis Horwood Limited, Chichester, 1988. E. P. Lampert and C. A. Dennis, Life history of two color morphs of the green peach aphid (Homoptera: Aphididae) on flue-cured tobacco, Tab. Sci. 31, 91 (1987).