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2002

Development of diagnostic tools for detectingexpression of resistance-associated esterases in thetobacco budworm, Heliothis virescens (F.)Huazhang HuangLouisiana State University and Agricultural and Mechanical College, hhuazha@lsu.edu

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Recommended CitationHuang, Huazhang, "Development of diagnostic tools for detecting expression of resistance-associated esterases in the tobaccobudworm, Heliothis virescens (F.)" (2002). LSU Doctoral Dissertations. 3869.https://digitalcommons.lsu.edu/gradschool_dissertations/3869

DEVELOPMENT OF DIAGNOSTIC TOOLS FOR DETECTING EXPRESSION OF RESISTANCE-ASSOCIATED ESTERASES IN THE TOBACCO BUDWORM,

HELIOTHIS VIRESCENS (F.)

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and

Agricultural and Mechanical College in partial fulfillment of the

requirements for the degree of Doctor of Philosophy

in

The Department of Entomology

By Huazhang Huang

B. Sc. (Applied Chemistry), Beijing Agricultural University, 1995 M. Sc. (Pesticide Science), China Agricultural University, 1998

December, 2002

ACKNOWLEDGMENTS I would like to express the deepest appreciation to my major professor, Dr. James A.

Ottea, for his constructive suggestions, enlightenment, encouragement and guidance. I

am grateful to the members of my Gradate Advisory Committee, Drs. Robert M.

Strongin, Lane D. Foil, Michael J. Stout and Jeffrey W. Hoy for their invaluable

suggestions and critical review of this dissertation.

I am deeply indebted to the LSU Agricultural Center and Drs. Frank Guillot, James

Fuxa for offering me a research assistantship and LSU Graduate School for providing a

Dissertation Fellowship. These financial supports made the completion of my graduate

study possible.

I would like to thank Mrs. Donald Cook for providing me susceptible insects and some

insecticides. Special thanks are expressed to Drs. Jianzhong, Huixin Fei and Lixin Mao

for their advice on statistical procedures used in this study. I also sincerely thank Dr.

Ibrahim Sanna for her help in rearing insects and Dr. Min He for helping me on NMR.

I am greatly grateful to Drs. Perich Michael, Carlton Chris, Jianzhong, Huixin Fei, and

Mrs. Donald Cook, Zou li, Ms. Moseley Victoria for their first help in my accident.

Special gratitude goes to all faculty, staff, and fellow graduate students in entomology

department, and all Chinese people for their kindness, assistance and friendship during

these years.

I also express my very special thanks to my parents, my sisters and my girlfriend for

their constant love, understanding, encouragement and support throughout this study.

Finally, I deeply thank the Almighty God to give me a second life.

ii

TABLE OF CONTENTS Page

ACKNOWLEDGMENTS…………………………………………………………….. ii

LIST OF TABLES …………………………………………………………………… v

LIST OF FIGURES…………………………………………………………………… vi

ABSTRACT…………………………………………………………………………… vii

CHAPTER 1. LITERATURE REVIEW .....................................................................................1 1.1 Introduction…………………………………………………………………… 1

1.2 History of Insecticide Resistance in H. virescens…......……………………… 1 1.3 Insecticide Resistance Mechanisms …………......…………………………… 2 1.4 Available Techniques for Detecting Esterases..………………………………. 17 1.5 Specific Objectives……………………………………………………………. 22 1.6 References..............................…………………………………………………. 23 2. DEVELOPMENT OF PYRETHROID SUBSTRATES FOR ESTERASES

ASSOCIATED WITH PYRETHROID RESISTANCE IN HELIOTHIS VIRESCENS (F.). ..................................................................................................41

2.1 Introduction…………………………………………………………………... 41 2.2 Materials and Methods………………………………………………………... 43 2.3 Results………………………………………………………………………... 52 2.4 Discussion…………………………………………………………………..... 56 2.5 References …….........................……………………………………………... 63 3. DEVELOPMENT OF METHODS TO DETECT ESTERASES ASSOCIATED

WITH PYRETHROID RESISTANCE IN HELIOTHIS VIRESCENS (F.).......... 69 3.1 Introduction…………………………………………………………………... 69 3.2 Materials and Methods………………………………....…………………….. 72 3.3 Results………………………………………………………………………... 75 3.4 Discussion……………………………………………………………………. 80 3.5 References……………………………………………………......................... 91 4. INVESTIGATING THE RELATIONSHIP BETWEEN PYRETHROID

STRUCTURE AND RESISTANCE IN THE TOBACCO BUDWORM, HELIOTHIS VIRESCENS (F.) .............................................................................95

4.1 Introduction……………………………………….………………………….. 95 4.2 Materials and Methods……………………………………………………… .98 4.3 Results…………………….…………………………………………………. .99 4.4 Discussion……………….……………………………………………………101 4.5 References ........................……………………………………………………104

iii

5. SUMMARY AND CONCLUSION .…………………………………………. 107 APPENDIX A SPECTRA OF GC-MS OF CIS-TTPA ........…………………………. 110

APPENDIX B SPECTRA OF GC-MS OF TRANS-TTPA …….……………………. 111

VITA……………………………………………………………………………………112

iv

LIST OF TABLES

page Table 2.1: Identification and analysis of products by GC-MS and 1H NMR………… 49 Table 2.2: Susceptibility of larvae to profenofos and cypermethrin………………….. 54 Table 2.3: Optimal conditions for continuous spectrophotometric assays……………. 55 Table 2.4: Esterase activities (nmol min-1 mg prot-1) toward pyrethroid and non- pyrethroid esters ………………………………………………………….. 57

Table 3.1: Effects of compounds on profenofos and cypermethrin toxicity in fifth -stadium larvae from susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens ………………………………………………… 78 Table 3.2: Susceptibility of larvae from susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens ……………………………………………. 81 Table 4.1: Susceptibility of larvae from the susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens…………………………………………… 100 Table 4.2: Structures of pyrethroids…………………………………………………… 102

v

LIST OF FIGURES page

Figure 2.1: Polyacrylamide gels of esterase activities toward 1- and 2-naphthyl acetate in

individual larvae from susceptible (LSU-S), profenofos-selected (OP-R) and cypermethrin-selected (PYR-R) strains of H. virescens. Each lane contains 75 µg of protein from whole body homogenates of individual fifth stadium (day 1) larvae……………………………………………………………………...58

Figure 3.1: Mortality measured following sequential, topical application of various doses

of synergists and 0.15 µg cypermethrin onto fifth stadium (day 1) larvae of H. virescens from the LSU-S strain……………………………………………..76

Figure 3.2: Filter paper assay with 1-NA and second stadium larvae of susceptible (LSU-

S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 1-naphthol. …………………… ...82

Figure 3.3 Filter paper assay with 1-NPA and second stadium larvae of susceptible (LSU-

S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 1-naphthol………………………..83

Figure 3.4 Filter paper assay with 2-NPA and second stadium larvae of susceptible (LSU-

S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 2-naphthol………………………..84

Figure 3.5 Filter paper assay with cis -TTPA and second stadium larvae of susceptible

(LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of thiophenol………………………..85

Figure 3.6 Filter paper assay with trans-TTPA and second stadium larvae of susceptible

(LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of thiophenol………………………..86

vi

ABSTRACT

Esterase-based metabolic resistance was studied using biochemical and biological

assays with organophosphate (OP)- and pyrethroid (PYR)- resistant tobacco budworms,

Heliothis virescens. In biochemical assays, results suggest that: (1) esterase activities

toward all substrates used were enhanced in both resistant strains compared with the

susceptible strain, suggesting that esterases were involved in resistance; (2) esterase

profiles differed depending on the strain and substrate used, and these differences were

visualized by using native polyacrylamide gel electrophoresis; 3) esterase activities

toward some pyrethroid substrates were significantly higher (P ≤ 0.01) in the PYR-R

strain than those in the OP-R strain. These results suggest that pyrethroid substrates may

be useful indicators for detecting esterases associated with pyrethroid resistance. Finally,

biochemical assays were modified for use on solid materials, and esterase substrates were

tested in filter paper assays. Whereas some differences in color intensity were detected

between susceptible and resistant strains, these differences were not dramatic. Thus,

utility of these substrates in such assays appears limited at this time, but further research

is warranted.

In biological assays, two approaches were taken to improve the precision with which

esterases associated with pyrethroid resistance were detected. First, bioassays were used

to test effects of pyrethroid substrates and traditional synergists (e.g., piperonyl butoxide)

on insecticide toxicity. Non-toxic pyrethroid esters enhanced pyrethroid toxicity to a

greater extent than DEF, a compound widely used as an esterase inhibitor. In addition,

synergism of profenofos and cypermethrin toxicity in both resistant strains by an oxidase

vii

inhibitor, 1, 2, 4-trichloro-3 (2-propynyloxy) benzene, suggests that P450

monooxygenases were also involved in resistance. The second method tested was to

utilize bioactivated insecticides to detect esterases. Absence of negative cross-resistance

to insecticides (i.e., acephate and indoxacarb) that are activated by esterases suggests that

detoxication of these compounds in resistant insects proceeds more rapidly than their

activation by esterases. Finally, levels of cross-resistance to tefluthrin and trans-

fenfluthrin were lower than to permethrin and cypermethrin in both resistant strains,

suggesting that resistance to insecticides in which sites for detoxifying enzymes (e.g.,

oxidases) are blocked develops more slowly than resistance to those in which metabolic

sites are present.

viii

1

CHAPTER 1

LITERATURE REVIEW

1.1 Introduction

The tobacco budworm, Heliothis virescens (F.), is a devastating cotton pest. Larvae of

this polyphagous insect damage a wide range of field, horticultural and ornamental crops

(Fitt, 1988). In addition, populations of H. virescens and its close relative, Helicoverpa

zea (Boddie), can increase rapidly to very large size and were reported to cause almost

one-third of all insect damage to American cotton from 1981 to 1998 (Suguiyama and

Osteen, 1988; Head, 1992, 1993; Williams, 1994-99). During this period, the economic

status of this complex consistently ranked as the number one pest in the southern cotton

ecosystem. Wide-spread plantings of transgenic cotton have effectively managed H.

virescens throughout much of midsouth. However, due to relatively low toxicity of

currently expressed Bt toxins to H. zea, the H. virescens / H. zea complex remains the

third most damaging of all cotton pests in this country (Williams, 2000, 2001).

1.2 History of Insecticide Resistance in H. virescens

Resistance is an inevitable consequence of the repeated applications of any new class

of insecticides, and it is one of the major reasons that H. virescens is so destructive

(Sparks, 1981; Sparks et al., 1993). Resistance has been described as “the ability of an

insect population to survive a dose of poison that is lethal to the majority of individuals in

a normal population of the same species” (Anonymous, 1957). Over the past 40 years,

field populations of H. virescens have developed resistance to almost all classes of

insecticides. During the 1960’s, organochlorine insecticides (e.g., DDT) could not

adequately control the tobacco budworm (Graves, 1967; Lingren and Bryan 1965;

2

Lowry, 1966), which led to increased reliance on organophosphates (OPs) and

carbamates. Shortly thereafter, resistance to many OP insecticides was reported (Graves

and Clower, 1971; Harris, 1972), and the pyrethroids became the cornerstone of chemical

management strategies in the cotton ecosystem during the late 1970’s (Wolfenbarger et

al., 1981). However, reduced efficacy of these highly effective and environmentally-

compatible insecticides to manage H. virescens spread rapidly throughout the mid-south

because of their heavy use and pre-existing cross-resistance to DDT (Plapp and Hoyer,

1968; Nicholson and Miller, 1985; Sparks et al., 1988). At present, populations of this

pest are being managed effectively by transgenic cotton that expresses a crystalline toxin

(Cry1Ac) from Bacillus thuringiensis (Bt). However, three Bt-resistant strains of H.

virescens have been developed by laboratory selection using Cry1Ac-impregnated

artificial diet (Stone, 1989; Gould, 1992, 1995). In addition, Elzen (1997) collected a

field strain of H. virescens with a significant resistance to a foliar formulation of Bt that

was applied in Mississippi from 1993 through 1995.

1.3 Insecticide Resistance Mechanisms

A comprehensive knowledge of mechanisms by which insects develop insecticide

resistance is a prerequisite for developing resistance management strategies and

diagnostic techniques for detecting and monitoring the expression of resistance

mechanisms in field populations of insects (Hammock and Soderlund, 1986). Insects

develop resistance to insecticides primarily though three mechanisms: decreased

penetration, reduced target site sensitivity and enhanced metabolism (Plapp, 1976;

Oppenoorth, 1985). Expression of all three mechanisms has been reported in H.

virescens (Sparks et al, 1993). Moreover, development of resistance to a particular

3

insecticide may be accompanied by the development of high levels of resistance or

hypersensitivity to other insecticides, due to similar (i.e., cross-resistance) or dissimilar

(i.e., multiple cross-resistance) resistance mechanisms (Adkisson and Nemec, 1967;

Leonard et al., 1988; Pittendrigh, 2000). Cross-resistance can increase or decrease

insecticide toxicity. For example, the toxicities of a number of structurally divergent Bt

toxins decreased in H. virescens following selection with a single Bt toxin (CryIAc) in

the laboratory (positive cross-resistance; Gould, 1992). In contrast, the susceptibility of

pyrethroid-resistant H. virescens to chlorfenapyr (a non-pyrethroid) increased with the

development of resistance to pyrethroids (negative cross-resistance; Pimprale et al.,

1997).

1.3.1 Penetration Resistance

Reduced rate of cuticular penetration of an insecticide delays, but does not prevent

the attainment of an effective amount of an insecticide that can kill the insect. Thus,

reduced cuticular penetration as a single resistance mechanism confers only low levels of

resistance and cross-resistance to an insecticide. However, when combined with other

mechanisms, reduced cuticular penetration may enhance resistance in more than an

additive fashion (Scott, 1990). In addition, reduced cuticular penetration is thought to be

expressed in many cases of insecticide resistance (Sawicki and Lord, 1970; Oppenoorth,

1985) and has been detected in many resistant insects, such as the house fly, Musca

domestica (Plapp and Hoyer, 1968), American cockroach, Periplaneta americana

(Soderlund et al., 1983), diamondback moth, Pultella xylostella (Noppun et al., 1989),

and the cotton bollworm, H. armigera (Ahmad and McCaffery, 1988). Despite its

prevalence and importance, this mechanism is poorly understood.

4

Reduced cuticlar penetration as a resistance mechanism is also expressed in

insecticide-resistant H. virescens. For example, the cuticular penetration of endrin into

resistant larvae of H. virescens was six-fold slower than that into susceptible larvae

(Pollens and Vinson, 1972). However, resistance due to a six-fold decrease in the

amount of absorbed endrin was amplified by co-expressed resistance mechanisms (e.g.,

enhanced metabolism, target-site insensitivity) to produce extremely high levels of

resistance to endrin in H. virescens. In contrast, reduced penetration was a minor

resistance factor in profenofos-resistant H. virescens (Kanga and Plapp, 1995). Finally,

reduced penetration was detected in both larvae and adults of cypermethrin-resistant H.

virescens, but difference between resistant and susceptible strains were not dramatic

(Ottea et al., 2000).

1.3.2 Target-Site Resistance

Reduced sensitivity of insecticide target sites is a second major insecticide resistance

mechanism. This mechanism results from functional modifications of target sites in

resistant insects compared with susceptible ones. Decreased sensitivity of all major

insecticide target sites (e.g., acetylcholinesterases (AChEs), the voltage-sensitive sodium

channel, and gama-aminobutyric acid (GABA)-gated chloride channel) has been reported

in a number of insects. To date, reported modifications have been exclusively point

mutations of structural genes encoding the target site protein (Feyereisen, 1995; ffrench-

Constant et al., 2000).

Target-site resistance to an insecticide usually confers cross-resistance to other

compounds that share the same target site. For example, decreased sensitivity of sodium

channel to DDT confers cross-resistance to all pyrethroids, which also act at this target

5

(Narahashi, 1983). Thus, it is believed that reduced susceptibility of the tobacco

budworm to the pyrethroids existed before their widespread use in the field because of

previous, extensive applications of DDT in the field (Davis et al., 1975; Brown et al.,

1982; Leonard et al., 1988).

1.3.2.1 Reduced Sensitivity of AChEs

The major target site for both OPs and carbamates (O’Brien, 1960; Eldefrawi, 1985) is

AChE, which acts by hydrolyzing excess neurotransmitter (acetylcholine) in some

synapses of the nervous system. Reduced sensitivity of AChE to these insecticides is

well-studied and has been expressed in a number of insects (Oppenoorth, 1985), such as

M. domestica (Walsh et al., 2001), the aphid, Nasonovia ribisnigri (Rufingier et al.,

1999), the Colorado potato beetle, Leptinotarsa decemlineata (Zhu et al., 1996), and the

fruit fly, Drosophila melanogaster (Mutero et al., 1992). Reduced sensitivity of AChEs

to OPs and carbamates is also expressed in H. virescens as a major resistance mechanism.

For example, compared with a susceptible strain, AChE activity from a methyl parathion-

resistant strain of H. virescens was 22-fold less sensitive to inhibition by methyl paraoxon

in both larvae and adults. This resistant AChE was also less sensitive to structurally-

analogous OPs, and to the N-methyl carbamates (Brown and Bryson, 1992). Similar

results were demonstrated in thiodicarb-selected larvae (Zhao et al, 1996) and OP- and

carbamate-resistant adults of H. virescens (Kanga et al., 1995).

Reduced sensitivity of AChE to OPs and carbamates is conferred by point mutations.

Although genetic and biochemical bases of reduced sensitivity of AChE have been

examined in a number of insects, they have been most well-studied in D. melanogaster,

M. domestica and L. decemlineata. In the nucleotide sequence of AChE in D.

6

melanogaster, Tyr109 was first reported as a potential site of a point mutation conferring

resistance to insecticides (Mutero et al., 1992). Subsequently, amino acid substitutions at

four different sites (Phe115 to Ser, Ileu199 to Val, Gly303 to Ala, and Phe368 to Tyr) were

found in OP-resistant D. melanogaster (Fournier et al., 1993) and at five different sites

(Val180 to Leu, Gly262 to Ala or Val, Phe327 to Tyr and Gly365 to Ala) in M. domestrica

(Walsh et al., 2001). The expression of varied combinations of mutations in a single

AChE gene resulted in different patterns of resistance among OPs, and levels of

resistance increased in an additive way. To date, more mutations in AChEs of D.

melanogaster have been reported as being possibly involved in OP and carbamate

resistance, but their toxicological significance has not been demonstrated (Villate et al.,

2000). Interestingly, reduced sensitivity of AChEs to OPs and carbamates also occurs by

replacement of a different amino acid (Ser238 to Gly) in an azinphosmethyl-resistant strain

of the Colorado potato beetle, L. decemlineata (Zhu et al., 1996).

1.3.2.2 Reduced Sensitivity of GABA-gated Chloride Channels

The GABA-gated chloride channel is the major target site for a number of

insecticides including cyclodienes (e.g., α-endosulfan), lindane, fipronil (Casida, 1993;

Bloomquist, 2001), the spinosyns (Watson, 2001), and possibly a secondary target site of

type II pyrethroids (Gammon and Casida, 1983) and the avermectins (Bloomquist, 1994).

To date, reduced sensitivity of GABA-gated chloride channels as a resistance mechanism

has been widely studied only in cyclodiene-resistant insects (Bloomquist et al., 1992;

ffrench-Constant, 1994). In addition, replacement of Ala302 in the GABA receptor

subunit gene (termed resistance to dieldrin or Rdl) from three different orders is the only

mutation that is known to confer reduced target site sensitivity at this site (ffrench-

7

Constant et al., 1995). There have been no direct studies associating reduced sensitivity

of GABA-gated chloride channel to resistance in H. virescens (McCaffery, 1998).

1.3.2.3 Reduced Sensitivity of Voltage-gated Sodium Channels

Three classes of insecticides (pyrethroids/DDT analogs, N-alkylamides, and

dihydropyrazoles) can modify sodium channel function, although modifications result

from binding to three distinct target sites on this protein (Soderlund and Knipple, 1995).

Thus, a domain-specific modification of sodium channels as a resistance mechanism

usually does not confer cross-resistance among these three classes, although negative

cross-resistance between pyrethroids and N-alkylamides (Elliott et al., 1987) or

dihydropyrazoles (Khambay et al., 2001) has been reported. Moreover, reduced

sensitivity of the alpha subunit of voltage-gated sodium channel has been widely studied

in DDT and pyrethroid resistance and has been identified in a range of insect species,

such as D. melanogaster (Vais et al., 2001) and M. domestica (Lee et al., 1999). Reduced

sensitivity of sodium channels to pyrethroids is expressed in pyrethroid-resistant H.

virescens (McCaffery, 1991; Ottea et al., 1995; Park and Taylor, 1997).

Substitutions of two different amino acids though point mutations are associated with

pyrethroid/DDT-resistant phenotypes: kdr (knock-down resistance) and super-kdr. First,

substitution of Leu1014 with Phe results in the kdr phenotype in D. melanogaster (Vais et

al., 2001) and M. domestica (Lee et al., 1999), which shifts the voltage dependence of

both activation and steady-state inactivation by approximately 5mV towards more

positive potentials and facilitates Na channel inactivation (Vais et al., 2001). In addition,

incorporation of a second mutation (Met918 to Thr) led to expression of the super-kdr

phenotype in both D. melanogaster (Vais et al., 2001) and M. domestica (Lee et al.,

8

1999), which also increased sodium channel inactivation and caused a more than 100-

fold reduction in deltamethrin sensitivity in D. melanogaster (Vais et al., 2001).

Similarly, replacement of Thr929 with Ile in D. melanogaster confers super-kdr-like

resistance (Vais, 2001).

1.3.3 Metabolic Resistance

Enhanced metabolism of insecticides decreases the attainment of the effective amount

of insecticides that can kill insects. Thus, metabolic resistance may significantly decrease

the susceptibility of insects to insecticides. Three major detoxifying enzymes are

associated with insecticide resistance: cytochrome P450 monooxygenases (CYPs),

glutathione S-transferases (GSTs) and esterases (ESTs) (Bull, 1981; Oppenoorth, 1985).

Most insecticides can be detoxified by at least one of these enzymes in insects and

enhanced detoxication is often involved in resistance. Moreover, the broad substrate

spectrum of each of these enzymes and the multiplicity of individual enzymes (ffrench-

Constant et al., 1999) is believed to cause cross-resistance among different classes of

insecticides, regardless of their target sites.

1.3.3.1 Resistance Associated with CYPs

Detoxifying CYPs are a superfamily of important enzymes that catalyze multiple

oxidative reactions and are capable of metabolizing a variety of endogenous and

exogenous substrates (Guengerich, 1996). Metabolism by CYPs generally leads to

detoxication of substrates, although activation is also possible. For example, the efficacy

of phosphorothioate insecticides is based on the activation of P = S to P = O by CYPs,

which results in more toxic insecticides than parent compounds (O’Brien, 1960;

Oppenoorth, 1972; Prestwich, 1990).

9

Elevated CYP activities have been detected in a number of resistant insects (Agosin,

1985; Feyereisen, 1999). Increased epoxidation by CYPs was expressed in resistant fall

armyworms, Spodoptera frugiperda (Yu, 1991). Moreover, enhanced demethylation by

CYPs was present in insecticide-resistant diamondback moth larvae (Yu and Nguyen,

1992) and permethrin-resistant soybean loopers, Pseudoplusia includens (Thomas et al.,

1996). Similarly, elevated deethylation by CYPS was detected in D. melanogaster

(DeSousa et al., 1995) and heightened levels of hydroxylation by CYPs were measured in

M. domestica (Li et al., 1989). Finally, enhanced CYP activities including dealkylation

(Rose et al., 1995; Zhao et al., 1996) and hydroxylation (Shan and Ottea, 1997) were also

expressed in insecticide-resistant H. virescens.

Elevated CYP activities are generally thought to occur by over-production of CYP

proteins. Rose et al. (1995) demonstrated that elevated CYP activities toward three

substrates (p-nitroanisole, benzopyrene and benzphetamine) in multi-resistant H.

virescens varied from 3- to 33-fold and were associated with enhanced cytochrome P450

content in microsomes from the gut (3.7-fold), fat body (4.4-fold) and carcass (4-fold) of

resistant insects. In addition, over-expression of CYP6 and CYP9 genes conferred

pyrethroid resistance in H. virescens (Pimprale, 1998). Similarly, over-expression of a

number of CYPs that were associated with insecticide resistance was present in CYP6A2,

CYP4E2, CYP6A9 genes of D. melanogaster (Amichot et al., 1994; Brun et al., 1996;

Maitra et al., 1996), CYP6A1, CYP6D1 of M. domestica (Cariño et al., 1992; Liu and

Scott, 1998), and CYP6B2 of H. armigera (Wang et al., 1995).

Enhanced transcriptional expression is possibly the underlying mechanism of CYP

over-expression that confers insecticide resistance. Pimprale (1998) compared the CYP6

10

and CYP9 genes from pyrethroid-resistant and -susceptible H. virescens and showed that

structural gene polymorphism and gene amplification were not likely to be responsible

for the pyrethroid resistance measured. However, altered transcriptional expression of

CYP genes was demonstrated and is believed to underlie resistance in this pest. In

studies with M. domestica, an elevated rate of transcription of CYP6D1 from pyrethroid-

resistant adults is an underlying cause of insecticide resistance (Liu and Scott, 1998), and

enhanced levels of transcription were mapped to factors on autosomes 1 and 2 (Scott et

al., 1998).

1.3.3.2 Resistance Associated with GSTs

Detoxifying GSTs are a family of enzymes that catalyze the conjugation of

glutathione with electrophilic substrates including insecticides (Soderlund, 1997). The

GSTs are involved in O-dealkylation or dearylation of OPs (Hayes and Wolf, 1988).

High frequencies of profenofos resistance were moderately correlated with GST activity

toward 1-chloro-2, 4-dinitrobenzene in larvae of H. virescens that were collected in

Louisiana cotton fields during the 1995 cotton growing season (Harold and Ottea, 1997).

Moreover, GSTs are involved in the dehydrochlorination of DDT in M. domestica (Clark

and Shamaan, 1984) and are the primary metabolic mechanism of resistance to this

insecticide. Finally, a recent study suggests that GSTs act as antioxidant-defense agents

and confer pyrethroid resistance in Nilaparvata lugens and possibly in other insects

(Vontas et al., 2001).

Enhanced activities of GSTs that confer insecticide resistance result from both

quantitative and qualitative alterations in gene expression. First, there is evidence for

over-expression of one or more GST isoforms in resistant insects. For example, the high

11

activity found in an insecticide-resistant strain of M. domestica is correlated with high

level of GST1 transcript (Fournier et al., 1992). Similar phenomena were also found in

insecticide-resistant Aedes aegypti (Grant et al., 1991), Anopheles gambiae

(Prapanthadara et al., 1993; Ranson et al., 2001), and P. xyostella (Ku et al., 1994).

Moreover, qualitative differences of GSTs were also present between susceptible and

resistant insects. For example, most subcellular fractions from susceptible M. domestica

had higher conjugation activities toward 1-chloro-2, 4-dinitrobenzene than the fractions

from the Cornell-R strain, but all fractions from the susceptible strain had lower

conjugation activities toward 1, 2-dichloro-4-nitrobenzene than fractions from the

Cornell-R strain (Chien et al., 1995). In addition, quantitative and qualitative alterations

can be co-expressed and confer resistance in one resistant strain. For example, GSTs

from a DDT-resistant strain of A. gambiae had an altered GST profile and one of GSTs

was increased 8-fold in a resistant strain compared with the susceptible strain

(Prapanthadara et al., 1993).

1.3.3.3 Resistance Associated with ESTs

Esterases are a large group of proteins (Benning, 1994; Cousin, 1997; Kim, 1997) that

share a highly-conserved catalytic domain characterized by an α / β hydrolase fold (Ollis,

1992) in their three-dimensional structures. They use water to hydrolyze endogenous and

exogenous esters and produce alcohols and acids as products.

A systematic nomenclature for classifying esterases remains to be established, and

multiple classifications are used. For example, based on the nature of inhibition by OPs,

esterases can be classified into three types (A, B, C; Aldridge, 1953). Moreover, based

on the identity of the functional amino acid (either serine or cysteine) within the catalytic

12

triad, esterases can be categorized into serine- or cysteine-type esterases (Yan et al.,

1994). Finally, esterases can be named according to the substrates they hydrolyze, such

as phosphorotriester hydrolase (PTEH, EC 3.1.1.2), AChEs, carboxylesterases (CarbEs;

EC 3.1.1.1), or amidases. Although there are multiple classifications, none of them is

completely satisfactory for naming individual esterases (La Du, 1992).

1.3.3.3.1 Esterase-based Resistance in Insects

Esterase-mediated metabolic resistance is widespread and has been detected in almost

all pests and against all classes of insecticides containing an ester moiety. First, enhanced

metabolism by esterases is a major mechanism in OP resistance, and has been detected in

a number of dipterans (Holwerda and Morton, 1983; Newcomb et al., 1997; Hemingway,

1981; Whyard et al., 1995), homopterans (Abdel-Aal et al., 1992; Owusu et al., 1996;

Chen and Sun, 1994; Chiang and Sun, 1996), coleopterans (Conyers et al., 1998;

Mathews, 1980; Argentine et al., 1989), lepidopterans (Beeman and Schmidt, 1982), and

hymenopterans (Baker et al., 1998). Similarly, elevated esterase activities are associated

with pyrethroid resistance in the sweet potato whitefly, Bemisia tabaci (Gennadius)

(Prabhaker et al., 1988), Leptinotarsa decemlineata (Lee, 1996), and the rice green

leafhopper, Nephotettix cincticeps (Ulher) (Chiang, 1996). Moreover, esterase-associated

resistance to carbamates has also been demonstrated in methomyl-resistant tentiform leaf

miners, Phyllonorycter blancard (F.) (Pree, 1989), and carbaryl-resistant S. frugiperda

(Yu, 1992). Finally, an esterase-based resistance mechanism has also been detected in

abamectin-resistant tomato leafminers, Tuta absoluta (Meyrick) (Siqueira et al., 2001)

and L. decemlineata (Say) (Argentine, 1991).

13

Enhanced esterase activities are also expressed in insecticide-resistant H. virescens. A

field-derived strain of H. virescens expressed an enhanced phosphorotriester hydrolase

activity as one of three mechanisms conferring resistance to methyl parathion (Konno et

al., 1990). More recent studies have shown that increased esterase activities toward

model (i.e., non-insecticide) substrates are associated with resistance to carbamate, OP,

and pyrethroid insecticides (Goh, 1995; Zhao, 1996), and are correlated with frequencies

of profenofos resistance in field-collected and laboratory-selected strains of H. virescens

(Harold and Ottea, 1997).

1.3.3.3.2 Esterases Associated with Insecticide Resistance

There are a large number of cases in which esterases are involved in insecticide

resistance. However, there are two relatively specific esterases (PTEH and CarbEs) that

are most often associated with metabolic resistance.

Levels of PTEHs (EC 3.1.1.2) are relatively low in arthropods compared with

mammals (Dauterman, 1982). However, in resistant insects, these enzymes effectively

cleave phosphorus esters. For example, an increase in PTEH activity was observed in all

of the malathion-resistant strains of M. domestica examined by Kao (1984). Moreover,

PTEH activity toward diazoxon was detected in a resistant strain of M. domestica and

was present in all subcellular fractions examined (Oi et al., 1990). In addition, a field-

derived resistant strain of H. virescens displayed increased PTEH activities as one of

three mechanisms contributing to methyl parathion resistance (Konno et al., 1989, 1990).

The CarbEs can hydrolyze both aromatic and aliphatic carboxylesters and have also

been shown repeatedly to be associated with insecticide resistance (Soderlund, 1997). As

was seen in GSTs, resistance arises from both quantitative and/or qualitative changes of

14

CarbE expression. Quantitative increases of CarbEs that confer resistance have been

shown to occur by over-expression of the corresponding genes. It is interesting to note

that, in many cases, over-expression of CarbEs does not result in a significant change in

overall rates of insecticide hydrolysis. Rather, the insecticide is sequestered from its

target site, and actual hydrolysis occurs very slowly (Devonshire, 1977; Herath et al.,

1987; Karunaratne et al., 1995). This type of resistance has been well studied in the

green peach aphid, Myzus persicae. A single isozyme (termed E4) from both susceptible

and OP-resistant aphid strains exhibited identical catalytic properties toward α-naphthyl

acetate (1-NA) and paraoxon, but massive amounts of E4 (approximately 3% of the

insect’s total proteins) are expressed in resistant aphids (Devonshire, 1977, 1979; 1982).

Moreover, further studies showed that E4 is very inefficient in catalyzing the hydrolysis

of insecticidal substrates, such as OPs, carbamates and pyrethroids (Devonshire, 1982).

These results suggest that the observed resistance to OPs, carbamates and pyrethroids

resulted primarily from sequestration of these insecticidal substrates by over-expressed

CarbEs rather than from increased efficiency of detoxication of these substrates.

Similarly, CarbEs from permethrin-resistant L. decemlineata possibly sequester

insecticides in a non-specific way, and sequestration is associated with cross-resistance to

other hydrophobic insecticides, such as other pyrethroids, DDT, and abamectin (Lee et

al., 1998). In addition, resistance conferred by sequestration by over-expressed CarbEs

also occurred in culicine mosquitoes and the brown planthopper, Nilaparvata lugens

(Mouches et al., 1986; Field and Devonshire, 1998; Karunaratne et al., 1998; Small and

Hemingway, 2000; Hemingway, 2000). Finally, abundance of an over-expressed CarbE

15

was quantitatively correlated with pyrethroid resistance in several field-collected strains

of H. armigera (Gunning et al., 1996).

Overproduction of sequestering esterases may result from either gene amplification

(e.g., multiple copies) or gene expression (e.g., methylation; Field et al., 1988; Vaughan

and Hemingway, 1995; Field et al., 1996; Rooker et al; 1996). Field et al. (1996)

demonstrated that approximately 12 copies of a 24 kb amplicon containing the E4 gene

were responsible for insecticide resistance in the peach-potato aphid M. persicae (Sulzer).

Similarly, high levels of OP resistance (approximately 800-fold) in C. pipiens resulted

from over-expressed aliesterase (Ali-E) B1 by B1 gene amplification, which contained at

least 250 copies of a 30 kb amplicon, including a highly conserved "core" of 25 kb

(Mouches et al., 1990). However, gene amplification cannot fully account for the levels

of resistance expressed. For example, the level of resistance (60-fold) measured in M.

persicae is substantially higher than the extent of gene amplification (12 copies),

suggesting that enhanced E4 gene expression or additional resistance mechanisms is also

responsible for producing resistance measured in these aphids (Field et al., 1996). A

similar conclusion may be possible to explain high levels of OP resistance in C. pipiens,

because the copy number of esterase gene (at least 250) is far below the level of

resistance (800-fold; Rooker et al., 1996). In addition, reversion in M. persicae from

resistant to susceptible phenotypes (Field et al., 1996; Hick et al., 1996; Field et al., 1999)

did not arise from the loss of amplified esterase DNA sequence, but rather from

decreased gene expression due to methylation at a certain site (CCGG in M. persicae) in

the amplified DNA sequence, which silences expression of the amplified E4 gene. A

similar phenomenon has been observed for the greenbug aphid, Schizaphis graminum

16

(Ono, 1999). These results suggest that both gene amplification and transcriptional

control affect esterase expression in resistant insects.

Qualitative changes in resistance-associated CarbEs are thought to occur as a result

of point mutations in structural genes for these proteins. Qualitative changes in CarbEs

are nicely illustrated by a phenomenon known as the “mutant aliesterase hypothesis.”

This theory was proposed after the observation that increased degradation of OPs is

associated with decreased CarbE activities toward “general” substrates, e.g., 1-NA

(Oppenoorth and van Asperen, 1961; Lewis and Sawicki, 1971; Hughes and Raftos,

1985). Recent research demonstrates that OP-hydrolase activities (with a diethyl

preference) of a mutated Ali-E increases in parathion-/ diazinon-resistant L. cuprina at

the expense of activities toward diverse carboxylesters including Ali-E substrates, 1-NA

and the carboxylester moiety of malathion. In contrast, OP-hydrolase activities (with a

dimethyl, rather than diethyl preference) of a mutated Ali-E increased in malathion-

resistant L. cuprina at the expense of activities toward Ali-E substrates and 1-NA, but not

the carboxylester moiety of malathion (Campbell et al., 1997). These and other studies

(Bell and Busvine, 1967; Townsend and Busvine, 1969; Huges and Raftos, 1985) led to

the conclusion that esterase-associated metabolic resistance to OPs in M. domestica, C.

putoria, and L. cuprina can be categorized into two related but distinct phenotypes,

resulting from different changes in the same major esterase genes (Campbell et al., 1997).

Present evidence from nucleotide sequence of structural genes suggests that point

mutations in CarbE genes are responsible for substrate specificity and insecticide

resistance. For example, a single substitution (Trp251-Leu) within the E3 CarbE of L.

cuprina results in resistance to malathion (Campbell et al., 1998). Conversely, a second

17

substitution (Gly137-Asp) within the same gene confers broad cross-resistance to a range

of OPs, but not malathion (Campbell et al., 1998). This same substitution within the E7

CarbE gene confers OP resistance in both L. cuprina and M. domestica (Claudianos et

al., 1999).

As noted earlier, quantitative and qualitative changes of CarbEs can co-occur in

resistant insects. Owusu et al. (1996) demonstrated that both changes of CarbEs are

responsible for dichlorvos resistance in the cotton aphid, Aphis gossypii. Similarly, both

kinds of changes of CarbEs also lead to the high levels of malathion resistance in C.

tarsalis (Ziegler et al., 1987). In addition, the relative resistance of several field-collected

strains of H. armigera was quantitatively correlated with the abundance and activities of

an over-expressed CarbE, suggesting that quantitative and qualitative changes have

occurred in expression of this CarbE (Gunning et al., 1996).

1.4 Available Techniques for Detecting Esterases The ability to detect esterases associated with resistance in a field situation is an

important component of insecticide resistance management strategies, because incisive

decisions regarding effective implementation of these strategies are primarily based on

precise detection of insecticide resistance mechanisms in early stages in the development

of resistance. Technology is available to develop a number of different types of

diagnostic assays (Brown and Brogdon, 1987) and each has advantages and

disadvantages. In general, these methods can be evaluated based on their expense,

specificity, the time and equipment required for development/implementation, and

ultimately, their field utility.

18

1.4.1 Chromatographic Techniques

Chromatographic techniques directly detect enhanced metabolism of an insecticide

by measuring either the disappearance of insecticide substrates or appearance of products.

These techniques include thin-layer chromatography (TLC), gas chromatography (GC)

(Abernathy and Casida, 1973) and high-pressure liquid chromatography (HPLC) (Yu,

1988, 1990). The cheapest and most facile one of these techniques is TLC, which can be

used to measure qualitative differences in metabolism, but, unless radioactive substrates

are used, the quantitative results are not possible. Whereas both GC and HPLC are

powerful analytic tools, they are highly dependent on instrumentation, fairly labor-

intensive and thus, difficult to use in a field situation.

1.4.2 Immunological Techniques

Immunoassays are analytical methods based on the specific immuno-reaction between

an antibody and antigen, and may be used for the quantitation of either product or

reactant in a solution. Enzyme Linked Immunosorbant Assays (ELISAs) have been used

successfully to detect esterases associated with insecticide resistance (Zhu et al., 1994;

Karunaratne et al., 1995, 1998; Siegfried et al., 1997), and are able to detect both low

abundance and over–expressed esterases (Devonshire et al., 1986). However, generation

of a specific antibody requires identification and purification of the enzyme of interest.

Moreover, the specificity of the generated antibody may be suspect (Karunaratne et al.,

1995). On the one hand, they may be so specific that their utility is limited to a single

esterase that may be unique to a particular insect. On the other hand, resistance-

associated esterases that result from a single point mutation may not be readily

distinguished from wild type enzymes (Siegfried, 1997).

19

1.4.3 Molecular Techniques

Monitoring techniques based on molecular biology are used for detecting

polymorphisms at the nucleotide level between insect strains. These techniques include

the polymerase chain reaction (PCR) and restriction endonucleases, single-stranded

conformational polymorphism, and PCR amplification of specific alleles (ffrench-

Constant et al., 1995; Zhu et al., 1999). The advantages of these techniques are that a

change in one nucleotide may be detected, making them the least ambiguous of presently

used diagnostic assays. In addition, this is the only type of method that allows absolute

detection of heterozygous individuals (ffrench-Constant et al., 1995). However, the

limitation of these techniques may be directly attributed to their unparalleled specificity:

the genetic change responsible for resistance must be identified prior to establishing this

method (ffrench-Constant, 1994), and this is both time-consuming and expensive.

Further, these methods are highly dependent upon instrumentation. Thus, their field

utility is limited. Thus, molecular monitoring techniques will probably never replace the

use of insecticide bioassays in the initial diagnosis, or even necessarily in the routine

monitoring of resistance (ffrench-Constant et al., 1995).

1.4.4 Bioassays with Synergists

Traditional bioassays with synergists are extremely fast and easy to use, but their

specificity is often questionable. Synergist assays rely on specific inhibition of

resistance-associated enzymes; however, synergists may reduce (or increase) the

penetration of insecticides and, thus, decrease (or increase) the accessibility of

insecticides to their targets (Martin et al., 1997; Sanchez et al., 2001). Moreover,

multiple forms of insecticide detoxifying enzymes may be expressed concurrently, and

20

the specificity of these enzymes to inhibition by any individual synergist is variable (Jao

and Casida, 1976; Khayrandish et al., 1993; Brown et al., 1996; Shan and Ottea, 1997).

As a result, the absence of synergism might indicate that: (1) the targeted enzyme is not

responsible for resistance, (2) a resistant-associated enzyme is present but not inhibited

by the synergist (ffrench-Constant et al., 1990; Brown et al., 1996) or (3) the synergist

affects the accessibility of insecticides to their target sites.

1.4.5 Biochemical Assays

Biochemical assays may be based on the hydrolysis of an actual insecticide or

surrogate substrate, such as 1-naphthyl acetate (1-NA) by esterases. Most often,

hydrolysis results in a color change that can be read by spectrophotometry, and the

intensity of this change is directly proportional to the activity of the enzyme of interest.

Based on the substrate and modes of measurement, three assays are possible.

1.4.5.1 pH Indicator Assays

A pH indicator assay is a colorimetric, pH responsive method for screening hydrolase

libraries, which is based on a pH change that occurs as the reaction proceeds (i.e., as

carboxylic acid are generated) (Moris-Varas, 1999). The advantage of this method is that

a target insecticide containing an ester bond is hydrolyzed to generate an acid. Thus, it is

straightforward to detect whether insecticide hydrolysis is a major mechanism of

resistance, and to detect the frequency of resistant individuals in a population. Moreover,

it is cheaper and safer compared with using a radiolabeled insecticide. However, these

assays are time-intensive compared with other biochemical assays because the change of

pH is usually very slow. In addition, it is difficult to use this method to visualize

esterases in electrophoretic gels because the color generated is most often transient.

21

1.4.5.2 Fluorescent Indicator Assays

Fluorescent indicator assays are based on the appearance of a fluorescent product

after hydrolysis of a surrogate substrate by an esterase, which can be detected by

fluorescent spectrophotometer. They are robust assays and have been used in a number

of cases to detect esterases (Homolya et al., 1993; Kitson et al., 2000; Shan et al, 2001);

however, the fluorescent product is invisible to the naked eye, and cannot be seen without

special lighting, thus their utility is limited for field situations and electrophoresis.

1.4.5.3 Colorimetric Assays

Colorimetric assays increase the appearance of visibly colored metabolites and/or its

conjugated complexes after hydrolysis of a surrogate substrate by an esterase. Many of

the surrogate substrates for these assays may be used to visualize esterases following

separation by polyacrylamide gel electrophoresis (PAGE; Matsumura and Brown, 1961;

Hemingway, 1981, 1989; Gopalan et al., 1997; Baker et al., 1998; Harold and Ottea,

2000). In addition, minor adaptations of these techniques for use on solid materials (e.g.,

filter paper “squash” assays) increase their potential use in field situations (Harold and

Ottea, 2000).

Colorimetric assays are widely utilized because they are economical, easy and fast

tools for detecting expression of enzymes associated with resistance. Furthermore,

compared with other methods, they have potential to be used extensively in the field

environment, although, to date, their use has been limited. However, the disadvantages

of these assays are similar to those of synergist bioassays: surrogate substrates may or

may not be suitable substrates for resistance-associated esterases, or the isozymes that

metabolize model substrates may not necessarily be those that detoxify insecticides

22

(McCaffery, 1998). For example, 1-NA is used extensively as a model substrate for

esterases, but is a poor indicator of resistance-associated esterases from pyrethroid-

resistant H. virescens (Dowd, 1987; Ottea et al., 2000). In addition, purified GSTs from

rat liver and M. domestica also have activity toward 1-NA, suggesting that some of

“esterase activity” measured with this substrate may result from the activity of GSTs

(Lalah et al., 1995).

1.5 Specific Objectives

I) Synthesize and evaluate substrates for resistance-associated ESTs from

the tobacco budworm, Heliothis virescens (F.).

* Synthesize and analyze pyrethroid substrates

* Optimize conditions for biochemical assays with substrates

* Evaluate substrates as indicators of resistance-associated ESTs from H.

virescens

II) Investigate the use of existing and novel compounds as tools for detecting

esterases associated with resistance.

* Evaluate whether synthesized pyrethroid substrates can be synergists of

cypermethrin toxicity in profenofos- and cypermethrin-resistant strains of H.

virescens

* Evaluate whether acephate, indoxacarb, tefluthrin and trans-fenfluthrin are

diagnostic compounds of esterases associated with insecticide resistance

* Evaluate whether pyrethroid substrates can be used as indicators of

esterases associated with resistance in a rapid “ squash” assay to detect

resistant individuals of H. virescens

23

III) Determine the degree to which resistance to profenofos or cypermethrin

confers cross-resistance to other organophosphates or pyrethroids in H.

virescens.

* Investigate the relationship between pyrethroid structure and cross-

resistance

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Dowd, P. F., Sparks, T. C., 1987. Inhibition of trans-permethrin hydrolysis in Pseudoplusia inculdens (Walker) and use of inhibitors as pyrethroid synergists. Pestic. Biochem. Physiol. 27 (2): 237-245.

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41

CHAPTER 2

DEVELOPMENT OF PYRETHROID SUBSTRATES FOR ESTERASES ASSOCIATED WITH PYRETHROID RESISTANCE IN HELIOTHIS

VIRESCENS (F.)

2.1 Introduction

Esterases are a large group of proteins (Benning, 1994; Cousin, 1996; Kim, 1997) that

share a highly-conserved catalytic domain characterized by an α / β hydrolase fold (Ollis,

1992) in their three dimensional structures. Esterases play critical roles in insect

physiology and detoxify a broad range of xenobiotics including insecticides (Dauterman,

1985). Enhanced esterase activity is a major mechanism of insecticide resistance

(Oppenoorth, 1984) and has been detected in a number of insects (Devonshire, 1991;

Hemingway, 2000). The role of esterases in insecticide resistance has been actively

studied in lepidopteran pests (McCaffery, 1991), where they have been shown to be

associated with resistance to organophosphorus (OP; Konno et al., 1990; Harold and

Ottea, 1997), carbamate (Goh et al., 1995; Zhao et al., 1996) and pyrethroid insecticides

(Konno et al., 1990; Goh et al., 1995; Zhao et al., 1996; Gunning, 1996, 1997).

The ability to detect esterases associated with resistance in a field situation is an

important component of insecticide resistance management strategies. Technology is

available to develop a number of different types of diagnostic assays (Brown and

Brogdon, 1987) and each has advantages and disadvantages. In general, these methods

can be evaluated based on their expense, specificity, the time and equipment required for

development, and ultimately, their field utility. Molecular techniques detect mutations in

specific genes that are associated with resistance. A change in one nucleotide may be

detected by this technique, making it the least ambiguous of presently used diagnostic

assays (ffrench-Constant et al., 1995). In addition, this is the only method that allows

42

absolute detection of heterozygous individuals. However, the limitation of this method

may be directly attributed to its unparalleled specificity: the genetic changes responsible

for resistance must be identified prior to establishing this method, and this is both time-

consuming and expensive. Further, this method is highly dependent upon

instrumentation; thus, its field utility is limited. Similar constraints limit the field utility

of immunoassays, which use antibodies directed against resistance-associated proteins,

and chromatographic techniques, which measure directly the enhanced metabolism of an

insecticide.

In contrast, insecticide bioassays with synergists and biochemical assays for

measuring enzyme activities are extremely fast and easy to use, and have potential field

utility. However, the specificity of these techniques is often questionable. These assays

rely on specific detection or inhibition of resistance-associated enzymes; however,

multiple forms of detoxifying enzymes may be expressed concurrently. For example, the

specificity of these enzymes to inhibition by any individual synergist is variable (Jao and

Casida, 1976; Brown et al., 1996; Shan and Ottea, 1997), and lack of synergism of

insecticide toxicity might indicate either that the targeted enzyme is not responsible for

resistance or a resistance-associated enzyme is present but not inhibited by the synergist

used (ffrench-Constant and Roush, 1990; Brown et al., 1996). Similarly, biochemical

assays have been used to detect changes in esterases expressed in insecticide-resistant

insects. In addition, many of the substrates for these assays may be used to visualize

esterases following separation by polyacrylamide gel electrophoresis (PAGE; Matsumura

& Brown, 1961; Hemingway, 1981, 1989; Gopalan et al., 1997; Baker, 1998; Harold and

Ottea, 2000). The disadvantages of these assays are similar to those of synergist

bioassays: model (i.e., non-insecticide) substrates for esterases may not be selectively

43

hydrolyzed by resistance-associated esterases. For example, 1-naphthyl acetate (1-NA),

which has been used extensively as a model substrate, is a poor indicator of resistance-

associated esterases from pyrethroid-resistant Heliothis virescens (Dowd et al., 1987).

Developing specific substrates is vital for enhancing the precision of quantitative

biochemical assays for detecting resistance. The hypothesis tested here is that the

specificity of detection of resistance-associated esterases can be enhanced by using

substrates that are structurally-similar to the insecticide that is resisted. The approach

used is to modify the structure of a commonly used pyrethroid insecticide such that the

product of hydrolysis is detectable by spectrophotometry. A similar approach has been

used to study esterases associated with pyrethroid hydrolysis in the cattle tick Boophilus

microplus (Riddles, 1983) and in human serum (Butte, 1999).

The major objectives of this Chapter are to: (1) identify whether non-pyrethroid

substrates are accurate indicators of esterases associated with pyrethroid resistance in H.

viresecens, and (2) if not, develop and test pyrethroid substrates that are structurally-

similar to conventional pyrethroids. Results in this study suggest that esterases

associated with pyrethroid resistance in H. virescens were more easily detected by

pyrethroid substrates than non-pyrethroid substrates.

2.2 Materials and Methods

2.2.1 Chemicals

Fast blue B salt (90% purity), 1-NA (98% purity), benzenethiol (97% purity), S-

methyl thiobutanoate (98% purity), 1-naphthol (99% purity), 2-naphthol (99% purity),

acetyl chloride (97% purity), thionyl chloride (99% purity), phosphorus pentoxide (98%

purity), 5,5’-dithio (2-nitrobenzoic acid) (DTNB, 99% purity), 4-nitrophenol (99%

purity) and benzoyl chloride (99% purity) were obtained from Aldrich Chemical Co.

44

(Milwaukee, WI). 2-Naphthyl acetate (2-NA) and Silica gel (28-200 mesh, average pore

diameter 22 Å) were purchased from Sigma Chemical Co., Inc. (St. Louis, MO). The

methyl ester of 3 – (2, 2 –dichlorovinyl) – 2, 2 – dimethylcyclopropane carboxylate (PA;

cis/trans = 40/60) was purchased from Fisher Scientific (Pittsburgh, PA). Technical

grade profenofos (O-(4-bromo-2-chlorophenyl)–O-ethyl–S-propyl phosphorothioate:

89%) was purchased from NOVARTIS (Greensboro, NC) and cypermethrin (a racemic

mixture of trans/cis, 1R/S and α R/S isomers: 96.2%) was from FMC Corporation

(Princeton, NJ). All other chemicals were analytical quality and purchased from

commercial suppliers.

2.2.2 Insects

A susceptible strain of H. virescens (LSU-S) was established in 1977 (Leonard et al.,

1988) and has been reared in the laboratory since that time without exposure to

insecticides. All larvae for resistance selection were fifth instars. Larvae from the LSU-

S strain were selected for five consecutive generations with topical applications of

profenofos at doses corresponding to the LD60 for each generation (3.5, 4.6, 8.5, 13.6 and

23.8 µg/individual). After one year without selection, individual larvae were treated with

25µg profenofos (an approximate LD20) and the next generation (OP-R strain) was used

for biochemical and toxicological assays.

Similarly, larvae from the LSU-S strain were selected for four consecutive generations

with topical applications of cypermethrin at doses corresponding to the LD60 (0.22, 0.4,

0.5 and 0.6 µg/individual). After one generation without selection, larvae were treated

with 1 µg of cypermethrin (6% mortality at 72 hours), and the next generation (PYR-R)

was used for biochemical and biological assays.

45

2.2.3 Bioassays

Insecticide susceptibility was measured and compared among strains using a topical

bioassay. Fifth instars (day 1), weighing 180±20 mg, were treated on the mid-thoracic

dorsum with 1 µl aliquots of profenofos or cypermethrin (in acetone) or with acetone

alone (control). The dose-mortality relationship for each compound was assessed from at

least five doses with 30 insects treated per dose. After treatment, larvae were maintained

at 27°C, and mortality was recorded after 72 hours. The criterion for mortality was the

lack of coordinated movement within 30 seconds after being prodded. Results were

corrected for control mortality with Abbott’s formula (Abbott, 1925), then analyzed by

Finney’s method (Finney, 1971). Resistance ratios (RR) were calculated as the LD50 of

resistant strain / LD50 of susceptible strain.

2.2.4 Synthesis

Preparation of PA followed the procedure of Shan et al. (1997). A mixture of the

methyl ester of PA (22.3 g, 0.1 mol), sodium hydroxide (12 g, 0.3 mol) and ethanol-water

(1:1, 250 ml) was refluxed for 12 hrs, concentrated under reduced pressure, diluted with

150 ml water, and extracted with ethyl ether (3 x 30 ml). The aqueous phase was

acidified with concentrated HCl (37.4%, 30 ml), and the oil precipitate was extracted into

ether (3 x 30 ml), washed with water (2 x 25 ml), and then dried (anhydrous Na2SO4)

overnight. The solution was filtered and ethyl ether was evaporated under reduced

pressure to obtain a solid product, which was re-crystallized from hexane / ether (1:1, v /

v). Yield 18.0 g (86%).

The method of Foggassy et al. (1986) was modified and used for separating trans and

cis isomers of PA. A mixture of PA (20.9 g, 0.1 mol) and benzene (100 ml) was stirred

at 27°C for 5 hrs. The suspension was filtered and the filter residue was recrystallized

46

from benzene to give 5.0 g of pure cis – PA (60% recovery). After removal of benzene

from the filtrate, the solid product was stirred 5 hrs at 27°C in 100 ml petroleum ether,

and then filtered to give 2.5 g solid product, which was recrystallized from hexane to give

1.8 g pure trans –PA (14% recovery).

Preparation of substrates (Scheme 2.1-2.2)

47

Two methods were adopted to prepare esters. First, thionyl chloride (0.1 mole, 8.7

ml) was added dropwise into a mixture of PA (0.025 mole, 5.25 g) and 100 ml benzene

(60% in hexane) and then refluxed for 3 hrs at 50-60 °C. The solvent and excess thionyl

chloride were removed from the reaction mixture and an oil product (PA-chloride) was

obtained (Riddles, 1983). A mixture containing 50 ml benzene (60% in hexane) and p-

nitrophenol (3.5 g, 0.025 mol) was prepared and refluxed for 4 hrs at 50-60°C. The

reaction mixture was washed with saturated NaHCO3 (20 ml x 3) and 50% ethanol-water

(50 ml x 2), and then purified by column chromatography with hexane: ethyl acetate

(70:30) as the developing phase to obtain 5.2 g p-NPPA (62.5% yield). A similar method

was used to prepare thiophenyl acetate (TPA) and 1- or 2-naphthyl PA ester (1-NPA & 2-

NPA). The yields of TPA, 1-NPA and 2-NPA were 72%, 79.7% and 80.5%,

respectively.

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The second method used was direct esterification between an acid and an alcohol

under dehydrating condition (such as polyphosphate ester; Adams, 1979; Imamoto,

1982). A reaction mixture containing cis/trans-PA (0.01 mol), thiophenol (0.011 mol),

polyphosphate ester (20 ml), chloroform (80 ml) and pyridine (10 drops) was stirred 12

hrs at room temperature. The final mixture was washed with saturated NaHCO3 (2 X 50

ml), then extracted with chloroform (2 X 40 ml). The combined organic phase was dried

(anhydrous Na2SO4) overnight. After removal of solvent, the solid product was purified

by column chromatography with a mixture of hexane: ethyl acetate (70: 30) as the

developing phase. The yield of cis-TTPA was 7.30 g (96.5 %).

Identity of products was determined by NMR and GC-MS (Table 2.1). 1H NMR

spectra were obtained on a Brucker Ac 200 spectrometer using tetramethylsilane as the

internal standard and CDCl3 as solvent. Gas chromatography-mass spectrometry (GC-

MS) was performed on a Finnigan GCQ (Trace GC 2000 coupled with Polaris MSD). A

silica capillary MS column, DB-5MS (30 m × 0.25 mm × 0.25 µm; J &W Scientific,

Folsom, CA), was operated at 600C for 1 min, increased to 1500C (at 5 0C/min

increments), where it was held for 15 min at this temperature, then increased to 3000C at

50C/min, where it was finally held for 10 min. The injector port was operated in splitless

mode at 2000C, and helium was used as carrier gas at 1.2 ml/min. The mass spectral

detector (MSD) was set on full scan model (m/z 41-400). Purity was calculated by

relative peak area.

2.2.5 Enzyme Preparation

Homogenates from individual larvae were used as enzyme source for measuring

esterase activities towards all substrates. Individual larvae were dissected and the

digestive system was evacuated. The remaining carcass was homogenized using an all-

49

Table 2.1. Identification and analysis of products by GC-MS and 1H NMR

Substrates purity GC (Retention MS m/z 1H NMR time, min) (Relative Intensity) (CDCl3) cis-TPPA 98.60% (90.54 18.19 127.03(100),191.07 (88.71), δ7.39-7.45 S cis-TTPA, 91.05(84.92), 63.02(65.93), 5H(aromatic 8.06% trans- 193.05(65.28), 165.01(40.4), protons); TTPA) 129.04(38.63), 241(10.93), δ6.18-6.27 q 77(13.3). 1H (C=C-H); δ 2.21-2.29 q 2H(2 cyclopropane protons);1.24- 1.31 d 6H(2- CH3) trans-TTPA 94% (no 18.36 127.04(100), 163.01(74.12), d7.39-7.45 S cis-TTPA) 91.06 (74), 191.06(71.85), 5H(aromatic 165.06(48.37),193.03(45.06), protons);d5.63 129.11 (32.5), 77(11.8) -5.77 d 1H (C=C-H); d 2.44-2.49 q 1H (C=C-C-H); d 2.02-2.04 d 1H (H-C-C=O); d1.24-1.27 d 6H (2 -CH3) 1-NPA 98.67% (cis 21.34 (cis) cis: 191.01(100), Butte (1999) 51.89%, 21.52 (trans) 127.04 (88.16), 193.01(68.57), trans 46.78%) 91.06(53.04),115.05(41.69), 129.04(29.44), 225 (26.2) trans: 191.02(100), 127.05(89.12), 193.01(69.32), 91.06(58.66), 225(40.4), 115.09(31.06), 129(29.52) 2-NPA 92.21% (cis 21.85(cis) cis: 127.05(100), 191.02 (87.25), ND 40.29%, trans 22.03 (trans) 91.05(68.68), 225.02(59.72), 51.92%) 193.03 (58.66), 144.1(57.3), 209 (47.1), 129.11(31.57) trans: 127.06(100), 191.03(89.41), 224.98(87.51), 193.08(77.05), 91.06(65.42), 209.01(65.26), 144.11(60.56), 129.05 (31.83)

Table continued

50

p-NPPA 96.43% (cis 10.07(cis) cis: 187.08(100), 127.11 (53.75), ND 6.08%, trans 10.19(trans) 91.05(49.75), 225.03(39.87), 90.35%) 226.99(35.09), 189.06(33.44) trans: 187.07(100), 127.05(58.76), 91.06(40.5), 189.04(34.7), 163.02(25.94) TPA 99.6% 12.89 110.02(100), 43.01(32.66), ND 109.07(13.5), 66.02(12.37), 65.02(11.05) ND: not determined.

51

glass homogenizer with 500 µl of ice-cold 0.1M sodium phosphate buffer (pH 8.0).

Homogenates were centrifuged at 12,600 g for 15 min at 40C. The resulting supernatants

were then held in ice and used in enzyme assays within 20 minutes of preparation.

2.2.6 Optimization of Biochemical Assays

Assays with all substrates were optimized (e.g., tissue, pH, temperature, substrate

concentrations, and protein dependence) with cytosol from an existing, pyrethroid-

resistant strain (CSA) of H. virescens (Shan et al., 1997). Individual larvae from the CSA

strain were dissected and the digestive system was evacuated. A whole body was rinsed

with 1.15% KCl and 0.1 M, pH 7.6 sodium phosphate buffer, then homogenized using a

all-glass homogenizer with 0.1M pH 7.6 sodium phosphate buffer containing 1mM

dithiothreitol, 1mM EDTA, 1mM phenylthiourea, and 4mM phenylmethylsufonyl

fluoride (Rose, 1995). The homogenate was centrifuged at 15,000g for 15 min. The

resulting supernatant was filtered through glass wool and further centrifuged at 115,000g

for 1 hour. The supernatant was used as the enzyme resource for optimizing the

conditions of biochemical assays. Activities of esterases toward all substrates (1-NA, 1-

NPA, 2-NPA, cis-TTPA, trans-TTPA, SMTB, TPA and p-NPPA) were measured using

the methods of Gomori (1953) and Ellman (1961) with modifications. The total volumes

of each individual well of a microtiter plate were 250 µl for 1-NA, 1-NPA, 2-NPA and

300 µl for SMTB, TPA, cis-TTPA, trans-TTPA and p-NPPA. The rate of change in

absorbency during the initial 10 min (or 1 min for TPA) was measured using a

Thermomax Microtiter Reader (Molecular Devices, Palo Alto, CA). Data were corrected

for non-enzymatic activity using incubations without protein as the control. Changes in

optical density were converted to nmol/min-1.mg prot-1 using the following extinction

52

coefficients: 9.25 mM-1250 µl-1 (Grant, 1989) for 1-NA, 1-NPA, 2-NPA; 8.34 mM-1300

µl-1 for cis-TTPA, trans-TTPA, TPA and SMTB; and 11.4 mM-1300 µl-1 for p-NPPA.

Native polyacrylamide gel electrophoresis (PAGE) was used to visualize esterases

from three individuals for each strain using a vertical eleptrophoretic unit (Hoefer

Scientific Instruments, San Francisco, CA) and 5% polyacrylamide (Sambrook et al.,

1989). Protein concentrations in homogenates were adjusted to contain 75 µg protein in

30 µl of pH 7.0 sodium phosphate buffer containing 0.1% Triton X-100, which was

mixed with 10 µl of a 6X tracking dye containg 0.25% bromophenol blue and 40%

sucrose in double-distilled water. Electrophoresis occurred in an electrode buffer

consisting of 100 mM Tris, 2.4 mM EDTA, and 100 mM boric acid (pH 8.0) at constant

voltage (150 V). When the dye marker migrated to within 1 cm of the gel base, each gel

was removed and stained in darkness at 250C with 100 ml of 0.1 M, pH 7.0 sodium

phosphate buffer containing 0.5 mM 1-NA and 2-NA, and 0.2% fast blue B for 30 min at

room temperature. Gels were de-stained in water and fixed in 5% acetic acid if

necessary. Protein concentrations were measured from diluted homogenates by the

method of Bradford (1976) using bovine serum albumin as the standard.

2.2.7 Statistical Analysis

Data from biochemical assays were analyzed using ANOVA – Tukey’s HSD test (P

≤0.01).

2.3 Results

2.3.1 Insecticide Susceptibility

Moderate levels of resistance to profenofos and cypermethrin were expressed in

fifth instars from the OP-R and PYR-R strains (Table 2.2). Resistance was greatest

against the insecticides to which larvae were selected: RRs of 18.1 to profenofos and 19.6

53

to cypermethrin were measured in OP-R and PYR-R strains, respectively. Moreover,

profenofos selection conferred cross-resistance (12.9-fold) to cypermethrin in the OP-R

strain. In contrast, cypermethrin selection conferred relatively low levels (4.53-fold) of

cross-resistance to profenofos in the PYR-R strain.

2.3.2 Optimization of Assay Conditions

Optimal conditions for enzymatic assays differed depending on the substrate used.

First, differences were measured in optimal pH among the substrates (Table 2.3).

Activities with all substrates except p-NPPA were greater in phosphate than in Tris/HCl

buffers, and optimal pHs differed. In contrast, highest activities with p-NPPA were

measured with Tris/HCl buffer at pH 8.6. Moreover, optimal temperatures for activities

also varied with substrates, and optimal amounts of proteins for assays (expressed as the

maximal value that was within linear range of activities for reactions) also varied among

substrates (Table 2.3).

2.3.3 Esterase Activities toward Non-pyrethroid Substrates

Esterase activities toward non-pyrethroid substrates were generally similar between

insecticide-resistant OP-R and PYR-R strains, but were significantly greater (Tukey’s

HSD; P ≤ 0.01) than that in insecticide-susceptible LSU-S larvae (Table 2.4).

Differences were not dramatic, and range from 1.1- and 1.2- fold (for 1-NA) to 1.9- and

2.2- fold (for SMTB) in the OP-R and PYR-R strains, respectively.

Both quantitative and qualitative differences in banding patterns among LSU-S, PYR-

R and OP-R larvae were observed in polyacrylamide gels stained with 1- and 2-NA (Fig.

2.1). The staining of some bands (e.g., bands 1 and 6, Rm: 0.25 and 0.83, respectively)

in both resistant strains was more intense than those in the susceptible strain. Moreover,

54

Table 2.2 Susceptibility of larvae to profenofos and cypermethrin Strain a Insecticide LD50 (95% FL) Slope ( ± SD) Chi-square RRb

LSU-S Profenofos 2.90 (2.40-3.56) 2.63 (0.40) 1.38 1.00 Cypermethrin 0.19 (0.16-0.24) 2.81 (0.39) 2.90 1.00 OP-R Profenofos 52.3 (46.0-59.2) 3.88 (0.57) 1.76 18.1 Cypermethrin 2.46(1.86-3.43) 6.49 (0.88) 1.06 12.9 PYR-R Profenofos 13.2 (10.4-16.2) 2.49 (0.39) 2.17 4.53 Cypermethrin 3.75 (3.16-4.33) 3.34 (0.48) 1.56 19.6

aStrains--LSU-S: insecticide-susceptible; OP-R: profenofos–selected; PYR-R: cypermethrin-selected. bRR= resistance ratio= LD50 (resistant strain/susceptible strain)

55

Table 2.3 Optimal conditions for continuous spectrophotometric assays

Substrate Final conc. Protein Buffer /pH Temp. (°C) Wavelength (nm)

(mM) (µg/assay) Peak Assay

1-NA 2.15 60 Phosphate / 7.0 25 450 450 1-NPA & 2-NPA 0.67 60 Phosphate / 7.0 25 435 450 SMTB 2.00 30 Phosphate / 7.6 29 435 450 TPA 1.33 30 Phosphate / 8.0 28 414 405

p-NPPA 0.17 360 Tris-HCl / 8.6 27 412 405 cis /trans-TPPA 1.33 120 Phosphate /7.6 31 412 405

56

some bands (e.g., bands 2 and 3, Rm: 0.37 and 0.50, respectively) were present in the

OP-R strain, but not in the LSU-S or PYR-R strains. In contrast, band 4 (Rm: 0.68) was

present in both LSU-S and PYR-R strains, but was not apparent in OP-R larvae. Finally,

band 5 (Rm: 0.73) was visible in LSU-S and OP-R strains, but not in larvae from the

PYR-R strain.

2.3.4 Esterase Activities toward Pyrethroid Substrates

As with non-pyrethroid substrates, esterase activities toward pyrethroid esters were

significantly greater (Tukey’s HSD; P ≤ 0.01) in insecticide-resistant strains (OP-R and

PYR-R) than those in insecticide-susceptible strain (LSU-S) (Table 2.4). Differences

between resistant and susceptible strains were greater than those measured with non-

pyrethroid esters, and ranged from 3.1- and 2.8- fold (for 2-NPA) to 8.4- and 2.7- fold

(for cis-TTPA) in the PYR-R and OP-R strains, respectively. Moreover, esterase

activities toward some pyrethroid substrates (e.g., 1-NPA, cis-TTPA and trans-TTPA)

were significantly greater in PYR-R strain than those in OP-R strain (Table 2.4), and

differences ranged from 1.3- fold for 1-NPA to 3.4- fold for cis-TTPA in PYR-R

compared with OP-R larvae. In contrast, esterase activities toward other pyrethroid

substrates (e.g., 2-NPA, p-NPPA) were similar between PYR-R and OP-R strains (Table

2.4).

2.4 Discussion

Enhanced levels of esterase activities are associated with insecticide resistance in

the tobacco budworm, H. virescens (McCaffery, 1998). In previous studies with non-

pyrethroid substrates (especially 1-NA), increased esterase activities were measured in

larvae that are resistant to OPs such as methyl parathion, profenofos and azinphosmethyl

57

Table 2.4 Esterase activities (nmol min-1 mg prot-1) toward pyrethroid and non- pyrethroid esters a

Esters Strainb Non-pyrethroid LSU-S OP-R PYR-R 1-NA 73.5(9.51)B 91.5(12.9)A 85.1(15.9)A TPA 345(79.5)B 582(126)A 621(158)A SMTB 98.5(15.5)C 214(33.9)A 188(40.8)B Pyrethroid 1-NPA 16.6(2.19)C 39.6(5.55)B 50.9(3.47)A 2-NPA 12.6(3.24)B 35.2(7.09)A 38.6(3.28)A cis-TTPA 0.82(1.62)C 2.02(0.69)B 6.88(0.87)A trans-TTPA 1.43(1.64)C 6.57(1.10)B 10.6(1.37)A

p-NPPA 0.42(0.36)B 1.21(0.65)A 0.96(0.72)A

a Activities (nmol min-1 mg prot-1) are expressed as means (± SD) based on assays with 30 larvae from each strain. In tests with LSU-S larvae, no activities were detectable in 5, 4, and 1 individual with cis-TTPA, trans-TTPA and p-NPPA, respectively. Values within rows followed by different letters are significantly different (ANOVA – Tukey’s HSD test, P ≤0.01) bStrain: LSU-S: insecticide susceptible; OP-R: profenofos–selected; PYR-R: cypermethrin-selected.

58

Figure 2.1 Polyacrylamide gels of esterase activities toward 1- and 2-naphthyl acetate in individual larvae from susceptible (LSU-S), profenofos-selected (OP-R) and cypermethrin-selected (PYR-R) strains of H. virescens. Each lane contains 75 µg of protein from whole body homogenates of individual fifth stadium (day 1) larvae.

59

(Goh et al., 1995; Zhao et al., 1996; Harold and Ottea, 2000). Moreover, Goh et al.

(1995) concluded that substantially increased esterase activities toward 1-NA may be

involved in resistance in a thiodicarb-resistant strain. Enhanced esterase activities also

confer pyrethroid resistance to H. virescens (Dowd et al., 1987; Graves et al., 1991; Goh

et al., 1995; Shan and Ottea, 1997). In the present study, further evidence is presented

that esterases are associated, at least in part, with resistance and cross-resistance in both

OP-R and PYR-R strains.

Detecting and monitoring insecticide resistance and associated mechanisms in insect

populations is possible using biochemical assays with model substrates (Brown and

Brogdon, 1987; Devonshire, 1987). Model substrates have been widely used in studies to

evaluate whether enhanced esterase activities play a critical role on insecticide resistance.

In previous studies, esterase activities toward 1-NA have been associated with insecticide

resistance in several insects (Abdel et al., 1993). High frequencies of profenofos

resistance were detected in larvae of H. virescens from 11 field strains collected in

Louisiana during 1995, and there was a strong correlation between profenofos resistance

ratios and esterase activities toward 1-NA (Harold and Ottea, 1997). Similarly, activity

toward S-methyl thiobutanoate (SMTB) is a good indicator of esterases associated with

malathion resistance (Kao et al., 1985), and TPA has been shown to have a similar

specificity for esterases as 1-NA (Chambers, 1973).

Staining electrophoretic gels with model substrates (e.g., 1- & 2- NA) can be used to

visualize quantitative and qualitative differences in esterases associated with resistance.

In a previous study, Harold and Ottea (1997) demonstrated that one esterase was

consistently over-expressed and a second band under-expressed in OP-resistant H.

virescens from both laboratory-selected and field-collected strains. In the present study

60

using a different OP-resistant strain, under-expression of a band migrating to the same

position (Rm= 0.68, Fig. 2.1) was detected and provides further evidence that resistance

to profenofos was accompanied by under-expression of this esterase. However, in the

present study, two bands (Rm = 0.37 and 0.5, Fig., 2.1) were also over-expressed in the

OP-resistant strain. This difference between these two studies may result from the

involvement of different esterases in resistance, or from different selection regimes (e.g.,

insecticide used and selection pressure) used to generate resistant strains.

The pattern of esterase banding was simpler in pyrethroid-resistant (PYR-R) than OP-

resistant (OP-R) larvae. Because both strains originated from the same parent colony

(i.e., LSU-S), it is likely that these differences in banding patterns are associated with

structural differences between profenofos and cypermethrin. Whereas both under-

expression (band 5, Rm = 0.73, Fig. 2.1) and over-expression (band 1, Rm = 0.25, Fig.

2.1) of esterases were observed in the PYR-R strain compared with the LSU-S strain,

differences between these strains in intensity of staining were not dramatic.

Results from biochemical assays using model substrates to determine causal

relationship between enzyme activities and insecticide resistance may be equivocal and

must be interpreted carefully. Isozymes that metabolize model substrates may not

necessarily be those that detoxify insecticides (McCaffery, 1998). The “mutant ali-

esterase” hypothesis originated from the observation that esterase activities toward 1-NA

actually decreased (rather than increased) with malathion resistance in some resistant

strains of M. domestica and other insects (Oppenoorth and van Asperen, 1960; Townsend

et al., 1969; Beeman et al., 1982; Hughes et al., 1985; Campbell et al., 1998). Similarly,

the model substrate, 1-NA, is a poor indicator of resistance-associated esterases from

pyrethroid-resistant H. virescens (Dowd et al., 1987; Ottea et al., 2000).

61

In the present study, esterase activities toward non-pyrethroid substrates including 1-

NA, SMTB and TPA were variable. Activities toward non-pyrethroid substrates did not

appear to be associated with either OP or pyrethroid resistance because activities were

not dramatically different (ranging from 1.1- and 1.2-fold for 1-NA to 1.9- and 2.2-fold

for SMTB, Table 2.4) in resistant PYR-R and OP-R strains compared with susceptible

LSU-S larvae. Moreover, banding patterns of esterases stained with 1- and 2-NA were

very similar between the PYR-R and LSU-S strains. In addition, the trend in esterase

activities toward non-pyrethroid substrates was dissimilar to that measured in bioassays

with cypermethrin. In contrast, activities toward pyrethroid substrates appeared to be

associated with pyrethroid (but not OP) resistance because activities were dramatically

different (ranging from 3.1-fold for 2-NPA to 8.4-fold for cis-TTPA, Table 2.4) in the

PYR-R strains compared with LSU-S strain. Moreover, the pattern of esterase activities

toward these substrates (except for 2-NPA and p-NPPA) was very similar to that

measured in bioassays with cypermethrin, but not with profenofos. Thus, esterases

associated with pyrethroid resistance may be more easily detected by pyrethroid

substrates than non-pyrethroid substrates. However, whether esterases hydrolyzing

pyrethroid substrates are those hydrolyzing cypermethrin is still questionable. In

previous reports, carboxylesterases from both mammals and insects have been reported to

prefer a pyrethroid with a primary alcohol ester (e.g., permethrin) to a pyrethroid with a

secondary alcohol ester (e.g., cypermethrin) (Abernathy et al., 1973; Holden, 1979).

However, Jersey et al. (1985) found that one esterase in the cattle tick, Boophilus

microplus, hydrolyzed α-cyano-substituted pyrethroid, trans-cypermethrin, but not trans-

permethrin.

62

Hydrolysis of geometric isomers of pyrethroid-similar esters strongly depends on the

sources or three-dimensional structures of esterases. More rapid hydrolysis of trans-

isomers compared with cis-isomers may be due to less steric hindrance by the

dichlorovinyl group of attack on the carbonyl carbon atom (Riddles, 1983). In previous

studies, pyrethroid esterases from pyrethroid-resistant Amblyseius fallacis (Chang et al.,

1986), Trichoplusia ni (Ishaaya and Casida, 1980) and Spodoptera littoralis (Ishaaya et

al., 1983) hydrolyzed trans-permethrin faster than cis-permethrin. In the present study,

trans-TTPA also can be more easily hydrolyzed by esterases than cis-TTPA. However,

hydrolysis of pyrethroid substrates cannot be explained only by chemical stability. For

example, the ratio of hydrolysis of trans-TTPA over cis-TTPA in the PYR-R strain was

1.54 (trans-TTPA and cis-TTPA: 10.59 and 6.88 nmol.min-1.mg prot-1, respectively,

Table 2.4). However, the ratio in the OP-R strain was 3.25 (trans-TTPA and cis-TTPA:

6.57 and 2.02 nmol.min-1.mg prot-1, respectively, Table 2.4). Similarly, isomer

specificity of pyrethroid esterases for trans- and cis-permethrin in Amblyseius fallacies

was observed in pyrethroid-resistant strains, but not in susceptible insects (Chang et al.,

1986). Conversely, esterases, from resistant rice green leafhoppers, Nephotettix

cincticeps (Chang et al., 1996), green lacewings, Chrysopa carnea (Ishaaya and Casida,

1981; Ishaaya, 1993) and the rice brown planthopper, Nilaparvata lugens (Chen and Sun,

1994), can degrade cis-isomers of permethrin and/or cypermethrin, faster (from 2- to 5-

fold) than the corresponding trans-isomers. These studies may imply that the sources of

esterases or three-dimensional structures of esterases play a primary role in the hydrolysis

of geometric isomers of pyrethroid esters, and the chemical stability of pyrethroid esters

may not be the major factor affecting enzymatic hydrolysis.

63

In summary, enhanced levels of esterase activities may play an important role in both

profenofos- and cypermethrin-resistant strains. Moreover, pyrethroid substrates appear to

be better indicators of esterases associated with pyrethroid resistance or cross-resistance

to pyrethroid insecticides than non-pyrethroid esters. However, whether esterases

detoxifying cypermethrin are those hydrolyzing pyrethroid substrates is still not clear.

2.5 References

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Adams, J. H., Lewis, J. R., Paul, J. G., 1979. Preparation of aryl esters using

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Butte, W., Kemper, K., 1999. A spectrophotometric assay for pyrethroid-cleaving

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amino acid substitutions in the ali-esterase, E3, confer alternative types of organophosphorus insecticide resistance in the sheep blowfly, Lucilia cuprina. Insect Biochem. Mol. Biol. 28 (3): 139-150.

Chambers, H., 1973. Thioesters as substrates for insect carboxylesterases. Ann. Entomol.

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Chen, W. L., Sun, C. N., 1994. Purification and characterization of carboxylesterases of a

rice brown planthopper, Nilaparvata lugens Stal. Insect Biochem. Mol. Biol. 24(4): 347-355.

Chiang, S. W., Sun, C. N., 1996. Purification and characterization of carboxylesterases

of a rice green leafhopper Nephotettix cincticeps Uhler. Pestic. Biochem. Physiol. 54 (3): 181-189.

Cousin, X., Hotelier, T., Giles, K., Lievin, P., Toutant, J. P., Chatonnet, A., 1997.

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Physiology, Biochemistry and Pharmacology. Volume 12. Insect control. Pergamon Press, Oxford, New York. pp 713-730.

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Ellman, G. L., Courtney, K. D., Anders, V., Featherstone, R. M., 1961. A new and rapid colormetric determination of acetylcholinesterase activity. Biochem. Pharmacol. 7: 88-95.

ffrench-Constant, R. H., Roush, R. T., Mortlock, D., Dively, G. P., 1990 Resistance

detection and documentation: The relative roles of pesticide and biochemical assays. In Pesticide Resistance in Arthropods (Eds., Roush R. T., Tabashnik B. E.). Chapman and Hall, NY, 1990, p4.

Finney, D. J., 1971. Probit analysis. 3rd Ed. Cambridge [Eng.] University Press, pp.333. Foggassy, E., F. Faigl, R. Soos and J. Rakoczi, 1996. Process for the separation of

isomeric cyclopropane carboxylic acids. US. Patent, 1996, 4,599,444. Goh, D. K. S., Anspaugh, D. D., Motoyama, N., Rock, G. C., Roe, R. M., 1995. Isolation

and characterization of an insecticide-resistance associated esterase in the tobacco budworm, Heliothis virescens (F.). Pestic. Biochem. Physiol. 51 (3): 192-204.

Gopalan, N., Bhattacharya, B. K., Prakash, S., Rao, K. M., 1997. Characterization of

carboxylesterases from malathion-resistant Culex quinquefasciatius Say (Diptera: Culicidae) mosquitoes. Pestic. Biochem. Physiol. 57(2): 99-108.

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glutathione S-transferase and general esterase in individual mosquitoes using an EIA reader. Insect Biocehm. 19 (8): 741-751.

Graves, J. B., Leonard, B. R., Micinski, S., Martin, S. H., 1991. Situation on tobacco

budworm resistance to pyrethroids in Louisiana during 1991. Proc. Beltwide Cotton Prod. Res. Conf. Memphis, Tenn.: National Cotton Council of America. 2: 743-746.

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Gunning, R. V., Moores, G. D., Devonshire, A. L., 1997. Esterases and fenvalerate

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Harold, J. A., Ottea, J. A., 1997. Toxicological significance of enzyme activities in

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Harold, J. A, Ottea, J. A., 2000. Characterization of esterases associated with profenofos resistance in the tobacco budworm, Heliothis virescens (F.). Arch. Insect Biochem. Physiol. 45 (2): 47-59.

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69

CHAPTER 3

DEVELOPMENT OF METHODS TO DETECT ESTERASES ASSOCIATED WITH PYRETHROID RESISTANCE IN HELIOTHIS VIRESCENS (F.)

3.1 Introduction

Insecticide resistance is a major impediment to the effective management of field

populations of the tobacco budworm, Heliothis virescens (Sparks, 1981; Sparks et al.,

1993). Resistance may result from reduced sensitivity of an insecticide’s target site to the

action of an insecticide, enhanced enzymatic detoxication of the insecticide or reduced

rates of penetration (Plapp, 1976; Oppenoorth, 1985). Expression of all three of these

mechanisms has been reported in H. virescens (Sparks et al, 1993).

The ability to discriminate among these three mechanisms is essential for

development and effective implementation of insecticide resistance management

strategies. Such knowledge would allow growers to make timely and rational decisions

about what to do once resistance has developed in their fields. For example, when

resistance is due to reduced sensitivity of a target site, classes of insecticides that act at

different target sites must be used as a countermeasure. Further, if resistance is due to

enhanced metabolism (and the enzymes responsible have been identified), synergists may

be used. At present, there are a limited number of techniques available to discriminate

insecticide resistance mechanisms. However, most methodologies are designed for use in

the laboratory and their stringent experimental conditions and need for instrumentation

make them difficult to utilize in field situations.

Biological assays have potential to be used as diagnostic assays in field situations.

They are cheap and there are no special requirements for equipment. Thus, they are

suitable for use by growers and consultants. Bioassays with esterase inhibitors have been

70

widely used in laboratory settings to detect whether enhanced esterase activities are

associated with insecticide resistance. However, the results of such assays are often

equivocal. The synergist, S, S, S-tributyl phosphorotrithioate (DEF), is extensively used

as an inhibitor of esterases to detect involvement of these enzymes in resistance, but DEF

also inhibits both glutathione S-transferases (Armes et al., 1997) and NADPH-dependent

monooxygenases (Sanchez et al., 2001). Thus, the presence of synergism (i.e.,

significantly increased toxicity) with esterase inhibitors in bioassays may not be an

accurate indication that esterases are associated with insecticide resistance. In addition, a

single inhibitor/synergist may not block the activity of all toxicologically-relevant (i.e.,

resistance-associated) enzymes; thus, the absence of synergism in assays with a single

compound might not eliminate involvement of esterases in resistance, because a resistant-

associated esterase may be present but not be inhibited by the synergist used (ffrench-

Constant and Roush, 1990; Brown et al., 1996). Finally, non-metabolic effects (e.g.,

reduced penetration) of synergists on insecticide toxicity may be present (Bull and

Patterson, 1993; Sanchez et al., 2001).

Improving the precision with which an esterase-based resistance mechanism is

detected with bioassays is important and necessary. Bioassays with an insecticide that is

bioactivated (i.e., toxicity increased) have potential to detect esterases associated with

insecticide resistance and offer advantages over bioassays using an insecticide plus a

synergist. For example, the esterase-based metabolites of some insecticides (e.g.,

acephate and indoxacarb) are more toxic than the parent compounds, and the rate of

insecticide bioactivation by esterases may dramatically influence toxicity (Mahajna et al.,

1997; Wing et al., 2000). A testable hypothesis is that if these insecticides were activated

71

faster than they are inactivated, negative cross-resistance to these insecticides would be

evident in insects expressing resistance-associated esterases. A second class of chemicals

with potential use as diagnostic compounds is insecticides in which metabolically-labile

sites for non-hydrolytic enzymes (e.g., oxidases) are blocked. For example, both trans-

and cis-isomers of 1 (R)- fenfluthrin suppressed resistance to cypermethrin in a

cypermethrin-resistant strain of H. virescens because the major target sites for

metabolism by cytochrome P450 monooxygenases (i.e., ring hydroxylation) in fenfluthrin

were blocked with halogens (Shan et al., 1997). A second, structurally-similar pyrethroid

is tefluthrin, which has a phenyl ring substituted with four fluorines and a methyl group.

Finally, bioassays with non-toxic pyrethroids acting as inhibitors or competitive

substrates of esterases associated with pyrethroid resistance have potential use in

diagnostic assays.

In addition to diagnostic bioassays, biochemical assays on solid materials may be used

in field situations to detect esterase-mediated resistance in early stages of development of

H. virescens. Simple colorimetric enzyme assays blotted onto a paper matrix (e.g., filter

paper or “squash” assays) have been developed to detect enhanced esterase activities

toward 1-naphthyl acetate (1-NA; Ozaki, 1969; Pasteur and Georghiou, 1980, 1981,

1989), and can be used in cases where resistance is positively (or negatively) correlated

with esterase activities toward a non-insecticide substrate. Thus, these methods have

potential to provide an efficient and rapid means to determine the frequency of resistant

individuals in field populations of insects. Using such a filter paper assay, differences in

esterase activities toward 1-NA between susceptible and resistant H. virescens were

observed (Harold and Ottea, 2000). In addition, such assays have been used in other

72

insects, including the leafhoppers, Nephotettix cincticeps and Laodelphas striatellus

(Ozaki, 1969) and a number of mosquitoes including Culex pipiens (Pasteur and

Georghiou, 1980, 1981), C. quinquefasciatus (Alkhatib, 1985; Pasteur and Georghiou,

1989), Aedes aegypti (Trees et al., 1985).

It is essential to improve the precision and expand the applications of these simple and

rapid field assays. To date, these assays have been only used with organophosphate-

resistant insects and the surrogate (i.e., non-insecticide) substrate used has been limited to

1-NA. However, 1-NA is a poor indicator of resistance-associated esterases from

pyrethroid-resistant H. virescens (Dowd et al., 1987; Ottea et al., 2000).

The present study has shown that, compared with non-pyrethroid substrates,

pyrethroid esters are better indicators of esterases associated with pyrethroid-resistance in

H. virescens. The major goals of Chapter 3 are to: 1) develop a filter paper assay

(“squash assay”) with pyrethroid substrates to detect esterases associated with pyrethroid

resistance; and 2) determine whether bioactivated insecticides (acephate and indoxacarb)

and novel pyrethroid esters can be used as diagnostic compounds for esterases associated

with insecticide resistance in H. virescens.

3.2 Materials and Methods

3.2.1 Chemicals

Fast blue B salt (90% purity), 1-naphthyl acetate (98% purity), benzenethiol (97%

purity), 1-naphthol (99% purity), 2-naphthol (99% purity), 5,5’-dithio(2-nitrobenzoic

acid) (DTNB, 99% purity) and benzenethiol (97% purity) were obtained from Aldrich

Chemical Co. (Milwaukee, WI).

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Pyrethroid esters tested as synergists included: [1/2-naphthyl, 3–(2, 2–dichlorovinyl)–

2, 2 – dimethylcyclopropane carboxylate (1/2-NPA), p-nitrophenyl, 3–(2, 2–

dichlorovinyl)– 2, 2 – dimethylcyclopropane carboxylate (p-NPPA), cis/trans -

thiophenyl, 3–(2, 2–dichlorovinyl)– 2, 2 –dimethylcyclopropane carboxylate (cis/trans-

TPPA)], and were synthesized as described in Chapter 2. In addition, S, S, S-tributyl

phosphorotrithioate (DEF; 98.7% from Bayer Corporation, Kansas City, MO), piperonyl

butoxide (PBO; 90-95% from Fluka Chemical Corporation, Milwaukee, WI) and 2, 3, 6-

trichloro-3 (2-propynyloxy) benzene (TCPB; > 95% purity (synthesized by Shan, 1997)

were also tested as synergists.

3.2.2 Insects

A susceptible strain of H. virescens (LSU-S) was established in 1977 (Leonard et al.,

1988) and has been reared in the laboratory since that time without exposure to

insecticides. All larvae for resistance selection were fifth instars. Larvae from the LSU-S

strain were selected for five consecutive generations with topical applications of

profenofos at doses corresponding to the LD60 for each generation (3.5, 4.6, 8.5, 13.6 and

23.8 µg/individual). After one year without selection, individual larvae were treated with

25 µg profenofos (an approximate LD20), and the survivors (OP-R strain) were used for

biological assays.

Similarly, larvae from the LSU-S strain were selected for four consecutive generations

with topical applications of cypermethrin at doses corresponding to the LD60 (0.22, 0.4,

0.5 and 0.6 µg/individual). After one generation without selection, larvae were treated

with 1 µg of cypermethrin (6% mortality at 72 hours), and the next generation (PYR-R)

was used for biochemical and biological assays.

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3.2.3 Bioassays

Fifth instars (day 1), weighing 180±20 mg, were treated on the mid-thoracic dorsum

with 1 µl aliquots of either profenofos or cypermethrin (in acetone) or with acetone alone

(control). The dose-mortality relationship for each compound was assessed from at least

five doses with 30 insects treated per dose. After treatment, larvae were maintained at

27°C, and mortality was recorded after 72 hours. The criterion for mortality was the lack

of coordinated movement within 30 seconds after being prodded. Results were corrected

for control mortality with Abbott’s formula (Abbott, 1925), then analyzed by Finney’s

method (Finney, 1971). Resistance ratios (RR) were calculated as LD50 of resistant strain

/ LD50 of susceptible strain.

Synergism of cypermethrin or profenofos toxicity was evaluated with LSU-S, OP-R

and PYR-R larvae. Compounds were applied to the mid-abdominal dorsum 30 min prior

to application of insecticide. Larvae were treated with 1µl acetone or synergist alone as

control. Doses of pyrethroid esters (1/2-NPA, cis/trans-TTPA, and p-NPPA) were

optimized for maximal mortality with fifth-stadium LSU-S larvae by topical application

of different concentrations (0-70 µg/larva) of synergists followed by cypermethrin (0.15

µg/larva). All pyrethroid esters administered were non-toxic to LSU-S larvae at 70

µg/larva. The dose used for tests with PBO, DEF and TCPB was 50 µg/larva (Shan et

al., 1997). Synergism ratio (SR) was calculated as: LD50 (without a synergist)/LD50 (with

a synergist). Synergism was indicated when SR > 1 and antagonism when SR < 1.

3.2.4 Filter Paper Assays

A “squash assay” was performed on filter paper based on earlier methods (Pasteur and

Georgiou, 1981; Harold and Ottea, 2000) with modifications. Second stadium larvae

75

(approximately 10-25 mg) were individually squashed through a micro-centrifuge tube

(cut open at the distal end and washed with double-distilled water) using a spatula tip

capped with a 0.5 ml micro-centrifuge tube on a Whatman #3 filter paper moistened with

phosphate buffer. The pH values of phosphate buffer used were 7.0 for 1-NA and 1- or

2-NPA, or 7.6 for cis or trans-TTPA. The filter paper containing a homogenized larva

was incubated atop a second filter paper that was soaked for 1 minute in a substrate

solution that was identical to that used for biochemical assays (see Chapter 2).

Incubation time for filters varied: 1 min for 1-NA and cis-TTPA; 30 sec for trans-TTPA

and 2 min for 1-/ 2-NPPA. Esterase activities were visualized as color production on

the undersurface of the filter paper, which were compared with a color scale prepared

with 1- or 2- naphthol (0-0.08 µmoles), or thiophenol (0-0.9 µmoles). Photographs were

taken immediately following reactions.

3.3 Results

3.3.1 Dose-response Relationship of Pyrethroid Esters

Generally, synergism of cypermethrin toxicity by pyrethroid esters was quantitatively

similar: toxicity increased, then decreased with increasing dose of synergists (Figure 3.1).

Moreover, maximal levels of mortality measured (50%-63%) were also similar among

compounds. In addition, responses measured between geometric isomers (cis- and trans-

TTPA) or stereoisomers (1- and 2-NPA) were also similar. However, optimal doses of

synergists were different and ranged from 20 µg/larva (for 1- and 2-NPA) to 50 µg/larva

(for p-NPPA).

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Figure 3.1 Mortality measured after sequential topical application of various doses of synergists followed by 0.15 µg cypermethrin onto fifth stadium (day 1) larvae of H. virescens from the LSU-S strain.

77

3.3.2 Synergism by Non-pyrethroid Compounds Effects of non-pyrethroid compounds on insecticide toxicity varied depending on the

insect strain and insecticide used for assays (Table 3.1). First, DEF antagonized

profenofos toxicity in the LSU-S strain, but had no significant effect on profenofos

toxicity in the OP-R strain, or cypermethrin toxicity in the LSU-S or PYR-R strain.

Moreover, in tests with LSU-S larvae, PBO synergized cypermethrin toxicity, but was an

antagonist of profenofos toxicity. No effect of PBO was observed in either the OP-R or

PYR-R strains. In contrast, TCPB was the most potent synergist of profenofos toxicity in

the LSU-S and OP-R strains, and significantly increased cypermethrin toxicity in the

PYR-R larvae, but no significant effect was observed on cypermethrin toxicity in the

LSU-S strain.

3.3.3 Synergism by Pyrethroid Compounds

All pyrethroids esters significantly increased cypermethrin toxicity in both OP-R and

PYR-R strains (Table 3.1). In the OP-R strain, SRs were similar among the five

pyrethroid esters, and ranged from 2.93 (for 1-NPA) to 3.67 (for cis-TTPA). Similarly, in

tests with PYR-R larvae, SRs for the five pyrethroid esters ranged from 1.80 (for 2-NPA)

to 3.34 (for p-NPPA).

3.3.4 Insecticides as Diagnostic Compounds

Susceptibility to indoxacarb, acephate, tefluthrin and trans-fenfluthrin was

significantly lower in larvae from the OP-R strain than that in the LSU-S larvae (Table

3.2). Resistant was highest to indoxacarb (15.8-fold) but relatively low levels were also

expressed toward acephate (2.76-fold), tefluthrin (3.30-fold) and trans-fenfluthrin (2.59-

fold). Similarly, susceptibility to these compounds was also lower in the PYR-R larvae

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Table 3.1 Effects of compounds on profenofos and cypermethrin toxicity in fifth-stadium larvae from susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens. Straina Treatmentb(µg) LD50 (95% fiducial limit)c SRd

LSU-S cypermethrin 0.19 (0.16-0.24) ---- cypermethrin + PBO (50) 0.09 (0.08-0.10)* 2.05 cypermethrin + DEF (50) 0.22 (0.16-0.38) 0.85 cypermethrin + TCPB (50) 0.12 (0.08-0.18) 1.66 profenofos 2.90 (2.40-3.60) ---- profenofos + PBO (50) 7.24 (5.71-10.4)‡ 0.40 profenofos + DEF (50) 8.02 (6.52-11.2)‡ 0.36 profenofos + TCPB (50) 1.63 (1.23-1.97)* 1.78

OP-R cypermethrin 2.46 (1.86-3.43) ---- cypermethrin + 1-NPA(20) 0.84 (0.55-1.26)* 2.93 cypermethrin + 2-NPA(20) 0.68 (0.56-0.81)* 3.62 cypermethrin + cis-TTPA(30) 0.67 (0.56-0.78)* 3.67 cypermethrin + trans-TTPA (40) 0.70 (0.42-1.05)* 3.51 cypermethrin + p-NPPA(50) 0.79 (0.53-1.14)* 3.11 profenofos 52.3 (46.0-59.2) ----- profenofos + PBO (50) 67.7 (58.8-79.8) 0.77 profenofos + DEF (50) 59.5(42.5-91.2) 0.88 profenofos + TCPB (50) 8.35 (6.29-10.3)* 6.26 PYR-R cypermethrin 3.75 (3.16-4.33) ----- cypermethrin + 1-NPA(20) 1.84 (1.27-3.00)* 2.04 cypermethrin + 2-NPA (20) 2.08 (1.80-2.48)* 1.80 cypermethrin + cis-TTPA(30) 1.75 (1.32-2.69)* 2.14 cypermethrin + trans-TTPA(40) 1.42 (1.17-1.76)* 2.64 cypermethrin + p-NPPA(50) 1.11 (0.91-1.36)* 3.34 cypermethrin + PBO (50) 3.28 (2.55-4.20) 1.14 cypermethrin + DEF (50) 4.81 (3.84-6.07) 0.78 cypermethrin + TCPB (50) 1.74 (1.43-2.13)* 2.16 aStrain-----LSU-S: insecticide-susceptible; OP-R: profenofos–selected; PYR-R: cypermethrin-selected btreatment: candidate synergists: 1-NPA: 1-naphthyl, 3–(2, 2–dichlorovinyl)–2, 2–dimethylcyclo- propane carboxylate; 2-NPA: 1-naphthyl, 3–(2,2–dichlorovinyl)–2, 2–di- methylcyclopropane carboxylate; p-NPPA: p-nitrophenyl, 3–(2, 2–dichloro- vinyl)–2, 2– dimethylcyclopropane carboxylate; cis/trans-TTPA: cis/trans-thiophenyl, 3–(2, 2–

table continued

79

dichlorovinyl)–2, 2–dimethylcyclo-propane carboxylate; DEF: S, S, S-tributyl phosphorotrithioate; PBO: piperonyl butoxide; TCPB: 2, 3, 6-trichloro-3 (2-propynyloxy) benzene. c(95% fiducial limit): ‡ antagonism, * synergism dSR =LD50 (without a synergist)/LD50 (with a synergist)

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and resistance was greatest toward indoxacarb (5.57-fold), but was also expressed toward

tefluthrin (2.18-fold) and trans-fenfluthrin (2.91-fold). However, the difference in LD50s

for acephate between LSU-S and PYR-R larvae was not statistically significant.

3.3.5 Filter Paper Assays Differences in color intensity for each product used in filter paper assays were

observed within and between the susceptible (LSU-S) and resistant (PYR-R and OP-R)

strains, but differences were not dramatic (Figures 3.2-3.7). For all substrates, color

production was more intense in tests with OP-R larvae than with LSU-S larvae.

However, no qualitative differences could be observed between PYR-R and LSU-S

larvae.

3.4 Discussion

Simple and rapid field assays to monitor and detect resistance and resistance

mechanisms in early stages of development of H. virescens are essential for providing

timely and accurate information for effective implementation of resistance

countermeasures. Insecticide bioassays have been successfully developed and used for

monitoring insecticide resistance in populations of H. virescens in cotton (McCutchen et

al., 1989; Ernst and Dittrich, 1992; Kanga and Plapp, 1995). For example, a glass vial

technique has been developed to monitor pyrethroid resistance in larval stages of H.

virescens (McCutchen et al., 1989), and this technique was further developed to monitor

resistance to biodegradable insecticides (e.g., organophosphates and carbamates) (Daly

and Fitt, 1990; Kanga and Plapp, 1995; Kanga et al., 1995). This technique not only

provides timely and accurate data for the stage when H. virescens is an actual pest, but

also provides consultants with a predictive bioassay that is useful in detecting resistance

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Table 3.2 Susceptibility of larvae from the susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens Straina Insecticide LD50 (95% FLb) Slope (SDc) Chi-square RRd LSU-S indoxacarb 0.27 (0.22-0.33) 4.46 (0.64) 2.05 1 acephate 25.8 (22.3-29.4) 4.26 (0.61) 1.65 1 trans-fenfuthrin 1.64 (1.22-2.12) 2.99 (0.40) 3.73 1 tefluthrin 0.51 (0.42-0.69) 2.65 (0.52) 0.05 1 OP-R indoxacarb 4.29 (3.18-5.87)* 1.62 (0.23) 1.11 15.8 acephate 69.1 (55.0-86.8)* 3.64 (0.53) 4.76 2.76 trans-fenfluthrin 4.30 (3.70-4.88)* 4.94 (0.64) 0.16 2.59 tefluthrin 1.68 (1.38-2.01)* 3.00 (0.419) 2.38 3.30 PYR-R indoxacarb 1.51 (1.02-2.06)* 1.68 (0.29) 0.44 5.57 acephate 39.4 (24.8-62.5) 5.09 (0.61) 9.48 1.53 trans-fenfluthrin 4.83 (3.91-6.25)* 4.49 (0.65) 3.18 2.91 tefluthrin 1.11 (0.96-1.31)* 1.77 (0.28) 6.75 2.18

aStrain--LSU-S: insecticide-susceptible; OP-R: profenofos–selected; PYR-R: cypermethrin-selected

bFL= fiducial limits. *: Significantly different from the corresponding value for the LSU-S strain. cSD=standard deviation. dRR=resistance ratio, defined as LD50 (resistant strain) / LD50 (susceptible strain).

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Figure 3.2 Filter paper assay with 1-NA and second stadium larvae of susceptible (LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 1-naphthol.

83

Figure 3.3 Filter paper assay with 1-NPA and second stadium larvae of susceptible (LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 1-naphthol.

84

Figure 3.4 Filter paper assay with 2-NPA and second stadium larvae of susceptible (LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of 2-naphthol.

85

Figure 3.5 Filter paper assay with cis-TTPA and second stadium larvae of susceptible (LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of thiophenol.

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Figure 3.6 Filter paper assay with trans-TTPA and second stadium larvae of susceptible (LSU-S) and resistant (PYR-R and OP-R) strains of H. virescens. Color scale developed with known concentrations of thiophenol.

87

before making a decision regarding subsequent management strategies. In addition, other

bioassays (e.g., topical bioassays, plant residue bioassays) also are used to monitor

insecticide resistance of field-collected populations of this pest (Elzen, 1991; Ernst and

Dittrich, 1992; Harold and Ottea, 1997).

Currently available bioassays provide no information on the biochemical mechanism

underlying resistance. A synergist may be incorporated into these assays to identify the

metabolic enzyme associated with resistance; however, synergists may not be specific for

the detoxifying enzymes associated with resistance (Armes et al., 1997; Sanchez et al.,

2001) or they may affect the accessibility of insecticides to their target sites (Bull and

Patterson, 1993; Sanchez et al., 2001). In the present study, three approaches were taken

in an attempt to improve the precision with which esterase-based resistance is detected.

The first method was to evaluate and compare effects of novel pyrethroid esters and

traditional synergists (PBO and DEF) on the toxicity of profenofos and cypermethrin in

resistant strains of H. virescens. All pyrethroid esters significantly increased the toxicity

of cypermethrin in the susceptible (LSU-S) and resistant (OP-R and PYR-R) strains,

further suggesting that esterases were associated with pyrethroid resistance in these

strains. However, SRs of each pyrethroid ester did not follow in the same order (i.e.,

LSU-S < OP-R < PYR-R) of esterase activities toward these substrates. Two

explanations are possible for this difference: 1) the optimized dose of each pyrethroid

ester with susceptible larvae may not be optimal for synergism in resistant strains because

of differential hydrolytic activities of esterases in these three strains; or 2) other, non-

hydrolytic, detoxifying enzymes, possibly cytochrome P450 monooxygenases (CYPs)

88

and glutathione S-transferases, may contribute to resistance (Brattsten et al., 1977; Vontas

et al., 2001).

The most widely used esterase synergist (DEF) was not a good indicator of esterases

associated with insecticide resistance in H. virescens. Antagonism of profenofos toxicity

was measured following pretreatment with DEF in both LSU-S and OP-R strains, but the

decreased LD50 was only statistically significant (P ≤ 0.05) in the LSU-S strain.

Antagonism by DEF of cypermethrin toxicity also was observed in both susceptible and

OP-resistant strains, but was not statistically significant (P ≤ 0.05). A similar result was

observed in field populations of cypermethrin-resistant Helicoverpa armigera, where

esterases were the predominant resistance mechanism and pyrethroid toxicity was

antagonized by DEF (Tikar et al., 2001). These studies suggest that DEF may not be a

specific inhibitor of esterases associated with cypermethrin resistance in these insects.

Significant synergism by TCPB of profenofos toxicity in the OP-R strain (SR of 6.26)

and of cypermethrin toxicity in PYR-R larvae (SR of 2.16) indicated that CYPs are

associated with resistance in these strains. In addition, there were only low levels of

cross-resistance (less than 3.5-fold) in both resistant strains to fenfluthrin and tefluthrin,

which have fluorine atoms or methyl groups blocking sites of attack for oxidases, in both

resistant strains.

Effects of the oxidase synergists, PBO and TCPB, varied with the strains tested. For

example, in the LSU-S strain, PBO acted as a synergist of cypermethrin toxicity and an

antagonist of profenofos toxicity, but did not significantly affect either profenofos

toxicity in the OP-R strain, or cypermethrin toxicity in the PYR-R strain. In contrast,

TCPB was a synergist of both profenofos toxicity in the OP-R strain and cypermethrin

89

toxicity in the PYR-R strain. These differences may result from multiplicity of CYPs

associated with insecticide resistance. Profenofos susceptibility is a function of both

bioactivation (Wing et al., 1983) and detoxication by CYPs. In the present study, CYPs

associated with profenofos activation may have been inhibited by PBO, which resulted in

antagonism of profenofos toxicity. In contrast, CYPs associated with detoxication of

profenofos and cypermethrin were inhibited by TCPB (Brown et al., 1996), resulting in

synergism of profenofos and cypermethrin toxicity. Moreover, differences in polarity of

these two compounds and/or variation of cuticular structure of larvae from these strains

may have produced differential effects on accessibility of these insecticides on their

targets.

Insecticides that can be activated or inactivated by esterases have potential to detect

esterase-mediated resistance. The hypothesis tested here was that negative cross-

resistance to acephate or indoxacarb (insecticides that are activated by esterases), or

positive cross-resistance to tefluthrin and fenfluthrin (that are insecticides inactivated by

esterases) would be present when esterase activity was enhanced with resistance. That is,

if resistance-associated esterases also hydrolyze acephate and indoxacarb, these

insecticides should be more toxic to resistant than to susceptible insects. However, low

levels of resistance to both compounds were measured in resistant strains. These results

suggest that either the bioactivation of these insecticides by esterases was much slower

than their detoxication, or other mechanisms (e.g., CYPs) were involved in resistance in

both resistant strains, which agrees with bioassay data with TCPB. In either case, the

utility of acephate and indoxacarb as indicators of esterase-mediated resistance in these

strains appears limited. In contrast, only low levels of cross-resistance to tefluthrin and

90

fenfluthrin (two pyrethroids with metabolically-blocked alcohol moiety) were expressed

in the resistant strains. This finding suggests that these compounds are not metabolized

by either esterases or oxidases in these strains, and may have potential use in diagnostic

bioassays to detect metabolism resistance.

Simple filter paper assays, based on biochemical assays, provide an efficient and

rapid means to distinguish resistant phenotypes in field populations of H. virescens, and

this technique has potential for utilization in field situations. In the present study,

differences in color intensity for each substrate used were observed within each strain,

suggesting that expression of esterase activities toward these substrates was

heterogeneous in these strains. However, differences in color developed using 1-NA, or

cis and trans-TTPA between OP-R and PYR-R strains were not obvious, although color

developed using 1 and 2-NPA in the OP-R strain was more intense than that in the PYR-

R strain. Patterns of color intensity among strains did not parallel (LSU-S < OP-R <

PYR-R) that measured for esterase activities in biochemical assays (Chapter 2). In

addition, the intensity of yellow color developed with cis and trans-TTPA is difficult to

discriminate with the naked eye. Thus, potential application of these substrates in filter

paper assays needs further evaluation in studies with a greater number of individuals.

In summary, novel pyrethroid substrates for esterases may be used as synergists of

pyrethroid toxicity in strains expressing esterase-mediated resistance. In some cases,

pyrethroid substrates may be used in filter paper assays in field situations to distinguish

resistant phenotypes in populations of H. virescens. In addition, the potential utility of

bioactivated insecticides (e.g., acephate and indoxacarb) and novel pyrethroid

insecticides (e.g., tefluthrin and fenfluthrin) in diagnostic bioassays for esterases

91

associated with pyrethroid resistance is limited, but they remain candidates for such

assays in other resistant insects.

3.5 References

Abbott, W.S., 1925. A method of computing the effectiveness of an insecticide. J.

Econ. Entomol. 18: 265. Al-khatib, Z. I., 1985. Isolation of an organophosphorate-susceptible strain of Culex

quinquefasciatus from a resistant field population by discrimination against esterase-2 phenotype. J. Am. Mosq. Control Assoc. 1: 105-107.

Armes, N. J., Wightman, J. A., Jadhav, D. R., Rao, G. V. R., 1997. Status of insecticide

resistance in Spodoptera litura in Andhra Pradesh, India. Pestic. Sci. 50 (3): 240-248.

Brattsten, L. B., Wilkinson, C. F., Eisner, P., 1977. Herbivore-plant interactions: mixed

function oxidases and secondary plant substances. Science. 196: 1349-1352. Brown, T.M., Bryson, P.K., Payne, G.T., 1996. Synergism by propynyl aryl esters in

permethrin-resistant tobacco budworm larvae, Heliothis virescens. Pestic. Sci. 46 (4), 323-331.

Bull, D. L., Patterson, R. S., 1993. Characterization of pyrethroid resistance in a strain of

the German cockroach, Blattella germanica (Dictyoptera: Blattellidae). J. Econ. Entomol. 86 (1): 20-25.

Daly, J. C., Fitt, G. P., 1990. Monitoring for pyrethroid resistance in relation to body

weight in adult Helicoverpa armigera (Hubner) (Lepidoptera: Noctuidae). J. Econ. Entomol. 83 (3): 705-709.

Dowd, P. F., Sparks, T. C., 1987. Inhibition of trans-permethrin hydrolysis in

Pseudoplusia inculdens (Walker) and use of inhibitors as pyrethroid synergists. Pestic. Biochem. Physiol. 27 (2): 237-245.

Elzen, G. W., 1991. Pyrethroid resistance and carbamate tolerance in a field population of

the tobacco budworm in the Mississippi delta. Southwestern Entomologist Supplement. 15: 27-32.

Ernst, G. H., Dittrich, V., 1992. Comparative measurements of resistance to insecticides

in three closely-related Old and New World bollworm species. Pestic Sci. 34(2): 147-152.

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ffrench-Constant, R.H., Roush, R.T., Mortlock, D., Dively, G.P., 1990. Resistance detection and documentation: the relative roles of pesticide and biochemical assays. In: Roush R. T., Tabashnik B.E., (eds) Pesticide Resistance in Arthropods. Chapman and Hall, NY p 4.

ffrench-Constant, R., Aronstein, K., Anthony, N., Coustau, C., 1995. Polymerase chain

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Harold, J. A., Ottea, J. A., 2000. Characterization of esterases associated with profenofos

resistance in the tobacco budworm, Heliothis virescens (F.). Arch. Insect Biochem. Physiol. 45 (2): 47-59.

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resistance to organophosphorus, carbamate, and cyclodiene insecticides in tobacco budworm adults. J. Econ. Entomol. 88 (5): 1150-1157.

Kanga, L. H. B., Plapp, F. W. Jr., 1995. Development of a technique to monitor

resistance to biodegradable insecticides in field populations of tobacco budworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 88 (3): 487-494.

Kanga, L. H. B., Plapp, F. W. Jr., Elzen, G. W., Wall, M. L., Lopez, J. D. Jr., 1995.

Mornitoring for resistance to organophosphorus, carbamate, and cyclodiene insecticides in tobacco budworm adults (Lepidoptera: Noctuidae). J. Econ. Entomol. 88 (5): 1144-1149.

Leonard, B. R., Graves, J. B., Sparks, T. C., 1988. Variation in resistance of field

populations of tobacco budworm and bollworm (Lepidoptera: Noctuidae) to selected insecticides. J. Econ. Entomol. 81(6): 1521-1528.

Mahajna, M., Quistad, G. B., Casida, J. E., 1997. Acephate insecticide toxicity: safety

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McCutchen, B. F., Plapp, F. W. Jr., Nemec, S. J., Campanhola, C., 1989. Development of diagnostic monitoring techniques for larval pyrethroid resistance in Heliothis spp (Lepidoptera Noctuidae) in cotton. J. Econ. Entomol. 82(6): 1502-1507.

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Pasteur, N. and Georghiou, G. P., 1981. Filter paper test for rapid determination of

phenotypes with highest esterase activity in organophosphate resistant mosquitoes. Mosq. News. 41: 181-183.

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quantifying increased esterase activity in organophosphate resistant mosquitoes (Diptera: Culicidae). J. Econ. Entomol. 347-353.

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resistance mechanism in tobacco budworm, Heliothis virescens, Abstract of Ph.D. Dissertation, Louisiana State University

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Sparks, T.C., 1981. Development of insecticide resistance in Heliothis zea and Heliothis virescens in North America cotton and crop pests. Bull. Entomol. Soc. Am. 27 (3): 186-192.

Sparks, T. C., Graves, J. B., Leonard, B. R., 1993. Insecticide resistance and the tobacco

budworm: past, present and future. Rev. Pestic. Toxicol. 2: 149-184. Tikar, S. N., Rao, N. G. V., Arya, D. R., Moharil, M. P., 2001. Pyrethroid resistance

monitoring and mechanisms in Helicoverpa armigera (Hubner) in central India. Crop Research Hisar. 22 (3): 474-478.

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Vontas, J. G., Small, G. J. H., Hemingway, J., 2001. Glutathione S-transferases as

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CHAPTER 4

INVESTIGATING THE RELATIONSHIP BETWEEN PYRETHROID STRUCTURE AND RESISTANCE IN THE TOBACCO BUDWORM,

HELIOTHIS VIRESCENS (F.)

4.1 Introduction Resistance is defined as “the ability of an insect population to survive a dose of poison

that is lethal to the majority of individuals in a normal population of the same species”

(Anonymous, 1957). Resistance of insects to insecticides continues to be a serious

barrier to successful management of insect pests (Metcalf, 1994). Field populations of H.

virescens have developed resistance to almost every class of insecticide over the past 40

years (McCaffery, 1998). Insects can develop resistance through reduced target site

sensitivity, enhanced metabolism and decreased penetration (Plapp, 1976; Oppenoorth,

1985). All these mechanisms are expressed in H. virescens (Sparks et al., 1993;

McCaffery, 1998).

Development of metabolic or target resistance to a particular insecticide may result in

hypersensitivity (i.e., negative cross-resistance) to other insecticides that have not been

previously applied. Mechanistically, negative cross-resistance may arise in a number of

ways. First, enhanced activities of detoxifying enzymes accompanied with resistance to

one insecticide may increase the bioactivation of another. For example, increased

activities of cytochrome P450 monooxygenases were possibly responsible for observed

negative cross-resistance to cypermethrin in a chlorpyrifos-selected strain of the German

cockroach, Blattella germanica (Scharf et al., 1997), to diazinon in the pyrethroid

resistant horn fly, Haematobia irritans (Cilek et al., 1995), to chlorpyrifos in dicofol-

selected two-spotted spider mites, Tetranychus urticae (Hatano et al., 1992) and to

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chlorfenapyr in pyrethroid-resistant H. virescens (Pimprale et al., 1997) and Haematobia

irritans (Sheppard and Joyce, 1998). Similarly, enhanced activities of esterases were

associated with observed negative cross-resistance to proinsecticides in insecticide-

resistant peach-potato aphid, Myzus persicae (Hedley et al., 1998). In addition to

increased metabolic activation, structural changes in insecticide target sites accompanied

with resistance to an insecticide may become more sensitive to other insecticides that

share the same target site. For example, the alteration in acetylcholinesterase resulting

from selection with dimethoate conferred negative cross-resistance to propoxur in the

olive fruit fly, Bactrocera oleae (Vontas et al., 2001). Finally, allosteric effects at target

sites may result in negative cross-resistance between two types of insecticides that share

the same target, but different sites. For example, due to allosteric effects of sodium

channels, negative cross-resistance to N-alkylamides (Elliott et al., 1986) or

dihydropyrazoles (Khambay et al., 2001) is expressed in pyrethroid resistant Musca

domestica.

In contrast, development of metabolic or target resistance to a particular insecticide

may often result in high levels of resistance (i.e., cross-resistance) to other insecticides

that have not been previously applied. For example, DDT and pyrethroids share the same

target site on the voltage-sensitive sodium channels. Thus, the wide variation in

susceptibility of H. virescens to the pyrethroids before their widespread field applications

is thought to result from cross-resistance associated with reduced sensitivity of this target

site arising from the previous and extensive use of DDT (Davis et al., 1975; Brown et al.,

1982; Leonard et al., 1988). Similarly, resistance due to reduced sensitivity of

acetylcholinesterases in H. virescens conferred cross-resistance among many (but not all)

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OPs and carbamates (Brown and Bryson, 1992; Heckel et al., 1998). Similar metabolic

mechanisms underlying resistance to OPs, carbamates and pyrethroids in H. virescens

conferred cross-resistance among these three classes of insecticides (Campanhola and

Plapp, 1987; Leonard et al., 1988; Zhao et al., 1996). Finally, cross-resistance to

structurally divergent Bt toxins in H. virescens was exhibited following selection with a

Bt toxin (Cry1Ac) in the laboratory (Gould, 1992) and is not accompanied by significant

alterations in toxin binding. Generally, metabolic resistance is believed to confer a

broader cross-resistance than target site insensitivity (Casida and Quistad, 1998).

The spectrum of cross-resistance not only depends on the target sites and metabolic

pathways of an insecticide, but also on the insect being selected and the chemical

structure of the insecticide used for selection. Both natural pyrethrins and synthesized

pyrethroids share the same target site (i.e., voltage sensitive sodium channels), but

pyrethroid-resistant M. domestica had very high levels of resistance to synthesized

pyrethroids (e.g., permethrin: 204-fold; phenothrin: 283-fold), but had only 7-fold

resistance to natural pyrethrins (Katsuda, 1999). Similarly, permethrin-selected resistant

strains of H. virescens expressed varying degrees of cross-resistance to other pyrethoids

(Brown and Bryson, 1996). Moreover, the pyrethroid, fenfluthrin, was not resisted by a

strain of H. virescens that was selected for high levels (i.e., 58-fold) of resistance to

cypermethrin (Shan et al., 1997). These studies suggest that the chemical structure of an

insecticide is an important determinant of metabolism and cross-resistance.

In the present study, the degree to which cross-resistance is conferred between OPs

and pyrethroids by selection for enhanced esterase activities was examined. The

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objective of this chapter was to investigate the relationship between pyrethroid structure

and cross-resistance in H. virescens.

4.2 Materials and Methods

4.2.1 Chemicals

Permethrin (94.6%), bifluthrin (96%), tefluthrin (97%), cypermethrin (96%) and

acephate (97.6%) were kindly provided by FMC Corporation (Princeton, New Jersey).

Indoxacarb (100%) and spinosyn A (100%) were kindly offered by Du Pont DE Nemours

(Wilmington, DE) and DowElanco Corporation (Indianapolis, IN), respectively. trans-

Fenfluthrin was originally synthesized by Shan et al. (1997) and re-crystallized before

use.

4.2.2 Insects

A susceptible strain of H. virescens (LSU-S) was established in 1977 (Leonard et al.,

1988) and has been reared in the laboratory since that time without exposure to

insecticides. All larvae for resistance selection were fifth instars. Larvae from the LSU-S

strain were selected for five consecutive generations with topical applications of

profenofos at doses corresponding to the LD60 for each generation (3.5, 4.6, 8.5, 13.6 and

23.8 µg/individual). After one year without selection, individual larvae were treated with

25 µg profenofos (an approximate LD20), and the survivors (OP-R strain) were used for

biochemical and toxicological assays.

Similarly, larvae from the LSU-S strain were selected for four consecutive generations

with topical applications of cypermethrin at doses corresponding to the LD60 (0.22, 0.4,

0.5 and 0.6 µg/individual). After one generation without selection, larvae were treated

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with 1 µg of cypermethrin (6% mortality at 72 hours), and the next generation (PYR-R)

was used for biochemical and biological assays.

4.2.3 Bioassays

Fifth instars (day 1), weighing 180±20 mg, were treated on the mid-thoracic dorsum

with 1 µl aliquots of either profenofos or cypermethrin (in acetone) or with acetone alone

(control). The dose-mortality relationship for each compound was assessed from at least

five doses with 30 insects treated per dose. After treatment, larvae were maintained at

27°C, and mortality was recorded after 72 hours. The criterion for mortality was the lack

of coordinated movement within 30 seconds after being prodded. Results were corrected

for control mortality with Abbott’s formula (Abbott, 1925), then analyzed by Finney’s

method (Finney, 1971). Resistance ratios (RR) were calculated as LD50 of resistant strain

/ LD50 of susceptible strain. Values were considered statistically different if their 95%

fiducial limits (FL) did not overlap.

4.3 Results

Resistance to all insecticides tested (except for acephate) was present in both OP-R

and PYR-R larvae (Table 4.1). Resistance to different insecticides varied depending on

the strain used for assays. Resistance was highest to the insecticide used for selection

(i.e., to profenofos in the OP-R strain and cypermethrin in the PYR-R strain). In

addition, pyrethroid resistance was higher to insecticides with 3-phenoxybenzyl

alcohol/aldehyde (e.g., cypermethrin and permethrin) than those with 3-phenyl 2-

methylbenzyl alcohol (e.g., bifenthrin), and with fenfluorobenzyl (e.g., fenfluthrin) or 4-

methyl tetrafluorobenzyl alcohol (e.g., tefluthrin). Moreover, negative cross-resistance

was not measured for either acephate or indoxacarb. In contrast, high levels of cross-

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Table 4.1 Susceptibility of larvae from the susceptible (LSU-S) and resistant (OP-R and PYR-R) strains of H. virescens Straina Pesticide LD50 (95% FLb) Slope (SDc) Chi-square RRd LSU-S profenofos 2.90 (2.40-3.56) 2.63 (0.40) 1.38 cypermethrin 0.19 (0.16-0.24) 2.81 (0.39) 2.90 permethrin 0.14 (0.11-0.19) 1.91 (0.25) 0.93 tefluthrin 0.51 (0.42-0.69) 2.65 (0.52) 0.05 bifenthrin 0.05 (0.04-0.06) 3.12 (0.42) 3.08 trans-fenfuthrin 1.64 (1.22-2.12) 2.99 (0.40) 3.73 spinosyn A 1.84 (1.35-2.66) 1.66 (0.29) 0.49 indoxacarb 0.27 (0.22-0.33) 4.46 (0.64) 2.05 acephate 25.8 (22.3-29.4) 4.26 (0.61) 1.65 OP-R profenofos 52.3 (46.0-59.0)* 3.88 (0.57) 1.76 18.1 indoxacarb 4.29 (3.18-5.87)* 1.62 (0.23) 1.11 15.8 cypermethrin 2.46 (1.86-3.43)* 6.49 (0.88) 1.06 12.9 permethrin 1.03 (0.54-1.75)* 2.05 (0.31) 3.99 7.20 bifenthrin 0.17 (0.13-0.24)* 1.77 (0.34) 2.03 3.47 tefluthrin 1.68 (1.38-2.01)* 3.00 (0.42) 2.38 3.30 trans-fenfluthrin 4.30 (3.70-4.88)* 4.94 (0.64) 0.16 2.59 acephate 69.1 (55.0-86.8)* 3.64 (0.53) 4.76 2.76 spinosyn A 4.03 (2.85-5.49)* 1.52 (0.22) 0.64 2.19 PYR-R cypermethrin 3.75 (3.16-4.33)* 3.34 (0.48) 1.56 19.6 permethrin 5.59 (4.27-7.91)* 1.77 (0.28) 0.57 12.4 bifenthrin 0.26 (0.18-0.41)* 2.97 (0.38) 8.40 5.10 trans-fenfluthrin 4.83 (3.91-6.25)* 4.49 (0.65) 3.18 2.91 tefluthrin 1.11 (0.96-1.31)* 1.77 (0.28) 6.75 2.18 profenofos 13.2 (10.4-16.2)* 2.49 (0.39) 2.17 4.53 spinosyn A 5.49 (3.79-7.93)* 1.30 (0.21) 0.19 2.99 indoxacarb 1.51 (1.02-2.06)* 1.68 (0.29) 0.44 5.57 acephate 39.4 (24.8-62.5) 5.09 (0.61) 9.48 1.53 aStrain--LSU-S: insecticide-susceptible; OP-R: profenofos–selected; PYR-R: cypermethrin-selected

bFL= fiducial limits. *: Significantly different from the corresponding datum in the LSU-S strain. cSD=standard deviation. dSR=synergism ratio, defined as LD50 (without a synergist) / LD50 (with a synergist).

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resistance to indoxacarb existed in both OP-R (RR: 15.8) and PYR-R (RR: 5.57) strains.

In addition, a low level of resistance to spinosyn A was measured in OP-R (RR: 2.19) and

PYR-R (RR: 2.99) larvae.

4.4 Discussion

Chemical structure, as it relates to metabolic stability, is a major determinant of cross-

resistance; however, the influence of structure on cross-resistance spectra is poorly

understood. In OP-resistant Lucilia cuprina, esterases are primarily responsible for

resistance (Campbell et al., 1998). Insects resistant to diazinon (a diethoxy ester)

displayed 2-fold more resistance toward other diethoxy OPs than their dimethoxy

analogs. Similarly, these strains did not show cross-resistance to either diisopropyl or

OPs with chiral phosphorus atoms. In contrast, strains selected with malathion (a

dimethoxy ester) displayed 2-5 times more resistance toward the dimethoxy OPs than

their diethoxy analogs. Moreover, these strains showed only slight resistance (<3-fold) to

either the diisopropyl or optically active OPs, including the diisopropyl analog of

malathion.

Structure activity relationships, as they relate to cross-resistance to pyrethroids, have

received limited attention. In the present study, a relationship between pyrethroid

structure and resistance (Table 4.2) has been examined with profenofos- and

cypermethrin-resistant H. virescens in which esterases and oxidases were primarily

responsible for resistance. Four type-I (non-cyano) pyrethroids (permethrin, bifenthrin,

tefluthrin and trans-fenfluthrin) showed much lower resistance ratios than a type-II

pyrethroid (cypermethrin) in both OP-R and PYR-R strains. Similar results were

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obtained previously with a different, cypermethrin-selected strain of H. virescens (Shan

et al., 1997). In previous studies, permethrin-selected H. virescens expressed greater

resistance to type-I (permethrin) than type-II (cypermethrin) pyrethroids (Jensen et al.,

1984; Brown et al., 1996). These results suggest that the insecticide used for resistance

selection has a major effect on the degree of cross-resistance to other insecticides. In

addition, the variability seen in resistance among the pyrethroids tested in this study

suggests that target site resistance is not a major mechanism of resistance in the PYR-R

strain.

Differences in resistance among four different type-I pyrethroids were present in both

OP-R and PYR-R strains. The phenoxybenzyl moiety of cypermethrin and permethrin is

a major site for enzymatic detoxication (Little et al., 1989; Lee et al., 1989). Tefluthrin

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and trans-fenfluthrin are structurally similar: both have fluorinated phenyl rings, thus, a

major site for detoxication by oxidases is no longer present. Resistance to these two

insecticides was very similar (from 2.5- to 3.3-fold), but much lower than that to

permethrin in both OP-R and PYR-R strains, suggesting that: 1) oxidases were involved

in resistance in both resistant strains, which agreed with results from bioassays with

TCPB (Chapter 3); 2) a substitution of one chlorine atom in trans-fenfluthrin with a

methyl group in tefluthrin did not change the degree of resistance in both OP-R and PYR-

R strains; 3) these two insecticides cannot be used as diagnostic compounds of esterases

associated with pyrethroid resistance (Chapter 3), but are useful for insects with oxidase-

mediated metabolic resistance. Moreover, the alcohol moiety of bifenthrin (i.e., 3-

phenyl 2-methyl benzyl alcohol) probably subjects to some metabolisms. This was

supported by higher resistance ratios of bifenthrin than those of tefluthrin and trans-

fenfluthrin, in the PYR-R strain.

The low level of resistance to non-phenoxybenzyl pyrethroids probably reflects the

contribution of a penetration mechanism. Similar levels of resistance to spinosyn A in

both OP-R and PYR-R strains (less than 3-fold) further suggest that a penetration

resistance may be present in both resistant strains.

In summary, the spectrum of cross-resistance to pyrethroids in both profenofos- and

cypermethrin-resistant strains was somewhat specific and was primarily dependent on the

insecticide used for resistance selection. However, resistance to tefluthrin and trans-

fenfluthrin, in which some metabolic sites for detoxifying enzymes (e.g., oxidases) are

blocked, may develop more slowly than those (e.g., permethrin, bifenthrin) in which sites

for detoxication are present. These results suggest that simple bioassays with such

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compounds may be used to detect metabolic resistance in insects. In addition, similar

modification in alcohol moiety of existing pyrethroids may result in insecticides that will

be active against tobacco budworms expressing metabolic resistance.

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Pimprale, S. S., Besco, C. L., Bryson, P. K., Brown, T. M., 1997. Increased susceptibility

of pyrethroid-resistant tobacco budworm (Lepidoptera: Noctuidae) to chlorfenapyr. J. Econ. Entomol. 90 (1): 49-54.

Plapp, F. W., 1976. Biochemical genetics of insecticide resistance. Annu. Rev. Entomol.

21: 179-197. Scharf, M. E., Neal, J. J., Bennett, G. W., 1997. Changes of insecticide resistance levels

and detoxication enzymes following insecticide selection in the German cockroach, Blattella germanica (L.). Pestic. Biochem. Physiol. 59 (2): 69-79.

Shan, G., Hammer, R. P., Ottea, J. A., 1997. Biological activity of pyrethroid analogs in

pyrethroid-susceptible and -resistant tobacco budworms, Heliothis virescens (F.). J. Agric. Food Chem. 45 (11): 4466-4473.

Sheppard, D. C., Joyce, J. A., 1998. Increased susceptibility of pyrethroid resistant horn

flies (Diptera: Muscidae) to chlorfenapyr. J. Econ. Entomol. 91 (2): 398-400. Sparks, T. C., Graves, J. B., Leonard, B. R., 1993. Insecticide resistance and the tobacco

budworm: Past, present and future. Rev. Pestic. Toxicol. 2: 149-184. Vontas, J. G., Cosmidis, N., Loukas, M., Tsakas, S., Hejazi, M. J., Ayoutanti, A., 2001.

Altered acetylcholinesterase confers organophosphate resistance in the olive fruit fly Bactrocera oleae. Pestic. Biochem. Physiol. 71 (2): 124-132.

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CHAPTER 5

SUMMARY AND CONCLUSIONS

Precise detection of esterase-mediated resistance and cross-resistance is a prerequisite

of effective implementation of insecticide resistance management strategies. In this

study, non-pyrethroid and novel pyrethroid substrates were evaluated as biochemical

indicators of esterases associated with pyrethroid-resistance in laboratory populations of

H. virescens. Moreover, novel pyrethroid substrates were evaluated as synergists of

pyrethroid toxicity in H. virescens, and results were compared with traditional synergists

of esterases. In addition, insecticides that are either bioactivated or detoxified by

esterases were assessed as indicators of esterases associated with pyrethroid resistance in

H. virescens. Finally, a relationship between pyrethroid structure and the development of

cross-resistance was studied with both OP- and pyrethroid-resistant H. virescens.

Esterase activities toward non-insecticide substrates were measured in the susceptible

(LSU-S) strain, and strains selected with profenofos (OP-R) or cypermethrin (PYR-R).

Esterase activities toward non-pyrethroid substrates (e.g., 1-NA, SMTP, TPA) and novel

pyrethroid esters (e.g., 1 and 2-NPA, cis and trans-TTPA and p-NPPA) were

significantly (P ≤ 0.01) enhanced in both resistant strains compared with those in the

susceptible LSU-S strain, suggesting that esterases were involved in resistance in these

resistant strains. However, significant differences (P ≤ 0.01) in esterase activities

between OP-R and PYR-R strains were detected only with SMTB, 1-NPA, cis and trans-

TTPA. In addition, patterns of esterase activities toward pyrethroid esters among three

strains matched those measured in that of resistance from bioassays with cypermethrin

(i.e., LSU-S < OP-R < PYR-R), but not profenofos. Thus, novel pyrethroid substrates

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appear to be better indicators of esterases associated with pyrethroid resistance or cross-

resistance to pyrethroid insecticides than non-pyrethroid esters.

Traditional synergists and novel pyrethroid substrates for esterases were evaluated as

markers of metabolic resistance in three strains. Novel pyrethroid substrates for esterases

significantly increased cypermethrin toxicity in both susceptible and resistant strains,

whereas the widely used esterase inhibitor, DEF, antagonized both profenofos and

cypermethrin toxicity in these strains. These results suggest that novel pyrethroid esters

were better inhibitors of esterases associated with pyrethroid-resistance in H. virescens

than DEF. Moreover, the oxidase inhibitor, TCPB, synergized both profenofos and

cypermethrin toxicity in both resistant strains, suggesting that cytochrome P450

monooxygenases were also involved in resistance in these strains.

Attempts were made to develop simple and rapid filter paper assay with non-

pyrethroid (e.g., 1-NA) and novel pyrethroid substrates (e.g., 1 and 2-NPA, cis and trans-

TTPA) using larvae from LSU-S, OP-R and PYR-R strains. Differences in color

intensity for each product were observed within and between the susceptible (LSU-S) and

resistant (OP-R and PYR-R) strains, but differences were not dramatic. For all substrates,

color production was more intense in the tests with OP-R larvae than with LSU-S larvae.

However, no qualitative differences could be observed between PYR-R and LSU-S

larvae.

Insecticides that are either bioactivated (e.g., acephate and indoxcarb) or detoxified

(e.g., tefluthrin and fenfluthrin) by esterases were investigated as indicators of esterases

associated with pyrethroid resistance in H. virescens. Strong positive cross-resistance to

indoxacarb was present in both OP-R and PYR-R strains, no or a low level of cross-

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resistance to acephate was present in both OP-R and PYR-R strains. These results

suggest that acephate and indoxacarb may not be useful as indicators of esterases in H.

virescens. In addition, low levels of resistance to tefluthrin and trans-fenfluthrin in both

OP-R and PYR-R strains were present, which suggests that these insecticides cannot be

used as indicators of esterases in H. virescens.

The spectrum of cross-resistance to pyrethroids in both profenofos- and

cypermethrin-resistant strains was somewhat specific, and was primarily dependent on

the insecticide used for resistance selection. Resistance to a type-II pyrethroid (e.g.,

cypermethrin) developed faster than that to type-I (e.g., permethrin, fenfluthrin, tefluthrin

and bifenthrin). However, resistance to the insecticides (e.g., tefluthrin, trans-fenfluthrin)

in which metabolic sites for detoxifying enzymes (e.g., oxidases) are blocked appears to

develop more slowly than to those (e.g., permethrin, bifenthrin) in which sites of

metabolism are present. These results suggest that simple bioassays with such

compounds may be used to detect metabolic resistance in insects. In addition, similar

modification in alcohol moiety of existing pyrethroids may result in insecticides that will

be active against tobacco budworms expressing metabolic resistance.

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APPENDIX A SPECTRA OF GC-MS OF CIS -TTPA

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APPENDIX B SPECTRA OF GC-MS OF TRANS-TTPA

112

VITA

Huazhang Huang, the second son of Zhenchu Huang and Qiaoyun Ge, was born on

August 27, 1971, in Qiuxi, Sichuan Province, China. He graduated from Qiuxi Sanzhong

High School, Sichuan Province, China, in 1991. He received his bachelor of science

degree in applied chemistry in 1995 from Beijing Agricultural University, Beijing, China.

He obtained his master of science in pesticide science under the guidance of Drs.

Xuefeng Li and Wenji Zhang in 1997 from China Agricultural University, Beijing,

China. Huazhang is presently a candidate for the degree of Doctor of Philosophy in

entomology at Louisiana State University.