Genotype to phenotype, the molecular and physiological dimensions of resistance in arthropods

Accepted Manuscript Title: Genotype to phenotype, the molecular and physiological dimensions of resistance in arthropods Author: René Feyereisen, Wannes Dermauw, Thomas Van Leeuwen PII: DOI: Reference:

S0048-3575(15)00005-X http://dx.doi.org/doi: 10.1016/j.pestbp.2015.01.004 YPEST 3777

To appear in:

Pesticide Biochemistry and Physiology

Received date: Accepted date:

5-12-2014 7-1-2015

Please cite this article as: René Feyereisen, Wannes Dermauw, Thomas Van Leeuwen, Genotype to phenotype, the molecular and physiological dimensions of resistance in arthropods, Pesticide Biochemistry and Physiology (2015), http://dx.doi.org/doi: 10.1016/j.pestbp.2015.01.004. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Genotype to phenotype, the molecular and physiological dimensions of resistance in

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arthropods

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René Feyereisena, Wannes Dermauwb, Thomas Van Leeuwenc

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a

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b

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Belgium

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c

INRA, Institut Sophia Agrobiotech, Sophia Antipolis, France Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, Ghent,

Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Amsterdam,

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Netherlands

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Corresponding author:

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INRA

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Sophia Antipolis

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France

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Phone: + 492386450

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E-mail: [email protected]

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Highlights:

Dr. René Feyereisen

 The different classes of mutations associated with resistance are discussed  A survey of mutations in each class is provided  Point mutations in target sites are still abundantly described  Gene duplications, amplifications and gene disruptions are increasingly important  Mutations affecting gene regulation are frequent but difficult to describe with precision Graphical Abstract

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ABSTRACT 1 Page 1 of 46

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The recent accumulation of molecular studies on mutations in insects, ticks and mites conferring

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resistance to insecticides, acaricides and biopesticides is reviewed. Resistance is traditionally

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classified by physiological and biochemical criteria, such as target-site insensitivity and metabolic

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resistance. However, mutations are discrete molecular changes that differ in their intrinsic

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frequency, effects on gene dosage and fitness consequences. These attributes in turn impact the

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population genetics of resistance and resistance management strategies, thus calling for a molecular

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genetic classification. Mutations in structural genes remain the most abundantly described, mostly

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in genes coding for target proteins. These provide the most compelling examples of parallel

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mutations in response to selection. Mutations causing upregulation and downregulation of genes,

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both in cis (in the gene itself) and in trans (in regulatory processes) remain difficult to characterize

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precisely. Gene duplications and gene disruption are increasingly reported. Gene disruption appears

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prevalent in the case of multiple, hetero-oligomeric or redundant targets.

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Keywords: selection, point mutation, gene duplication, gene amplification, gene disruption, transposable element

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1. Introduction

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Theodosius Dobzhansky is well known for his landmark phrase "nothing makes sense in

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biology except in the light of evolution, sub specie evolutionis" [1]. Less well known is the fact that

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in one of the pillars of the modern evolutionary synthesis, Genetics and the Origin of Species,

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Dobzhanksy pointed out that “[insecticide resistance is] probably the best proof of the effectiveness

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of natural selection yet obtained” ([2]; see also [3]). This bold statement is remarkable for at least

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two reasons. One, that it was made even before the introduction of DDT for vector and pest control,

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when resistance was not the problem that it is today. Two, that insecticide resistance did not

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become a favorite research topic of evolutionary biologists, with the notable exception of James

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Crow ([4]; see also [5]). It seems that by 1951, when DDT resistance had emerged as a major

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problem in entomology, Dobzhansky did not belabor the point. In one of the first conferences on

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insecticide resistance, he delivered (in absentia) a 16 line summary of the idea that resistance "can

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be understood only as a special case of the far more general phenomenon of adaptability of

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populations of sexual species to environmental change" [6] . James Crow at the same conference

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presented his first results on laboratory selection of a DDT-resistant strain of Drosophila, but

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modestly wondered "whether any of these interests are such as to lead to much improvement in

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methods of control" [7]. Undeterred by the fact that "the phenomenon of resistance has received

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surprisingly little attention from evolutionary biologists", Georghiou and Taylor [8-10] and others

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[11, 12] proceeded to lay the foundations of modern resistance management, by analyzing the

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genetic, biological and operational parameters which control the dynamics of resistance. These

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factors were then extensively modeled by population geneticists (a field already reviewed by [13]).

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It is undeniable that the success in resistance management for Bt transgenic crops [14, 15] owes

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much to the theoretical work on the design of resistance management strategies to classical,

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chemical pesticides.

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Resistance is agreed to be an evolutionary phenomenon, with the same factors driving the

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dynamics of resistance to classical chemical pesticides, to biopesticides and to transgenic crops. It is

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somewhat odd then that resistance to Bt toxins does not squarely fit in the usual classification of

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resistance (target site, metabolism...) and is not usually discussed in that framework. The traditional

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classification of resistance distinguishes toxicodynamic and toxicokinetic changes in the physiology

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and biochemistry of resistant strains, and often also includes behavioral changes. Alteration of the

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target site is a toxicodynamic change, whereas increased metabolism, decreased penetration,

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sequestration or increased excretion are toxicokinetic changes. These broad classifications are

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useful because they are phenotypic descriptions of underlying genetic changes. Also, for classical

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pesticides they allow some degree of intervention and management. For instance, increased

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metabolism and detoxification can be answered by the use of synergists. An altered target site

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would be better dealt with by a pesticide with a different mode of action. Such knowledge can help

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design resistance management strategies [16]. A comprehensive documentation of the various

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modes of action is therefore a key task of IRAC [17].The difficulty of classifying resistance to

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toxins from entomopathogens, most notably Bacillus thuringiensis toxins is perhaps due to the

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complex mode of action - what constitutes metabolism, what precisely can be considered a target

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site when Bt toxins can bind to multiple gut proteins?

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The population genetic approach that gives the theoretical framework for resistance

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management follows the effects of operational factors (selection dose, schedule and alternance,

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observance of refugia) [9] and biological factors (life history traits) [8] on genotype frequencies. So

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perhaps a classification of resistance based on the genotype, more precisely the type of mutation,

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rather than its ultimate toxicokinetic or toxicodynamic phenotypic outcome would be helpful. This

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additional dimension of resistance is always implicit, but has rarely been formulated. It was first 3 Page 3 of 46

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introduced by Cariño and Feyereisen [18], adapted by Mullin and Scott [19] and extensively

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developed by Taylor and Feyereisen [20]. The advances in the molecular analysis of resistance call

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for an update of the latter study. In this paper, we present a tabular summary of this additional

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molecular genetic dimension of resistance to insecticides and acaricides. We show that it can

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integrate our knowledge on mutations causing Bt resistance in the same framework as mutations

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causing resistance to classical chemical pesticides. Furthermore, we hope that this "state of the art",

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describing how often different classes of mutations can lead to resistance in the field but also in

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laboratory selection, will be useful in the continued quest for resistance management strategies.

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2. Different classes of mutations

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Different classes of mutations affecting a gene implicated in resistance can be

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distinguished, and fall broadly in three categories. The first are mutations affecting the coding

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sequence of the gene and thereby structurally alter the gene product. The second are mutations

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causing an increase in gene dosage or expression. The third are mutations causing a decrease in

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gene dosage or expression. The latter two classes can be further distinguished in mutations affecting

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the whole gene (such as duplication and amplification, or disruption and loss) or just the trans

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regulation or the cis regulatory elements of the gene. By using the classes of mutations as a basis for

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classification, the nature of the selecting agent, a synthetic pesticide, a biopesticide, a transgenic

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crop making its own biopesticide, is not a factor. It is then possible to describe resistance in two

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dimensions, a molecular genetic dimension, and a biochemical/physiological dimension (the

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classical way). To avoid hair-splitting discussions on whether proteolytic processing of a Bt toxin

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can be classified as metabolism, or whether mutations in aminopeptidase or cadherin genes are truly

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target site mutations, we can then simply add gut toxin resistance as a separate class. This two-

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dimensional classification is described schematically in Figure 1. Where do we find the well

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documented cases of resistance to insecticides and acaricides in this two-dimensional grid ?

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3. Mutations affecting the coding sequence of a gene

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These include non-synonymous nucleotide changes causing point mutations in the protein

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sequence. A very extensive literature covers such point mutations causing a decreased sensitivity of

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the main target sites while there are far fewer point mutations reported in toxicokinetic resistance

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(see 3.5 below) (Table 1, [21-48]). Point mutations, as well as small insertions or deletions (indels)

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or even transposable element insertions can introduce a premature stop codon. The gene is then

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disrupted and the protein, if expressed, is mostly non-functional.

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3.1. Mutations in the voltage sensitive sodium channel (VSSC) gene

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Mutations in this target of DDT and pyrethroid insecticides have been recently analyzed in

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the excellent reviews from Ke Dong’s laboratory [49, 50], and we include in our analysis a few

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recent additional papers [51-54]. Clearly this is a snapshot of our knowledge base at this time, but it

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already gives an impressive account of the parallel evolution that has taken place as a result of DDT

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and pyrethroid selection. Sixty one different mutation positions or combinations of up to five

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individual mutations have been reported in 51 different species. Not all have been conclusively

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shown to contribute to resistance. Two mutations form the "core" of the VSSC structural changes,

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the kdr mutation at Leu1014 and the super-kdr mutation at Met918, with 81 reported cases of one or

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both mutations (Figure 2).

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3.2. Mutations in the acetylcholinesterase genes

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The biochemical characterization of acetylcholinesterase (AchE) activity with decreased

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sensitivity to inhibition by organophosphate or carbamate insecticides is relatively straightforward,

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but the molecular basis of this insensitivity is made somewhat complicated because most insects

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have two AchE genes (Ace 1 and 2), whereas higher Diptera have only one (now designated as Ace

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2). Within the Acari, a single gene has been reported for the spider mite T. urticae [55], while up to

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three genes have been reported for ticks [56]. Outside the Cyclorrhapha, AchE1 appears to be the

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main catalytic enzyme of insects, although this may not always be the case [57]. The work on

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Drosophila and house fly Ace is therefore not immediately translatable to other insects or Acari,

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and resistance mutations as well as mutation combinations have now been documented for both

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genes. Over a dozen sites can be found mutated in insensitive AchE from various organisms, and

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other polymorphisms in resistant strains have been compiled in the ESTHER database [58]. Only

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four mutations are found at orthologous positions in the alignments between AchE1 and AchE2

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(Table 2, [59-91]). This indicates that despite the important phylogenetic distance between the two

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Ace genes [92] there is a high degree of constraint in the way both enzymes can retain activity while

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discriminating against the OP and carbamate substrates. Multiple alleles with one or more

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mutations can be found in field collected strains that indicate multiple origins of resistance. In the

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house fly, at least three origins of the Phe290Tyr;Gly227Ala/Val alleles were shown, each then

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complemented with the Val150Leu or Ala201Ser mutations [93]. In Drosophila, the four most 5 Page 5 of 46

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frequent mutations, Ile129Val, Gly227Ala, Phe290Tyr and Gly328Ala can be found independently

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or in various combinations [59, 60]. In another study, several origins of the I120V mutation were

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then seen complemented with the Gly227Ala and Phe290Tyr mutations [94].

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Heterologous production of each of the main AchE2 mutants of Drosophila alone or in

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combination (15 allelic variants) allowed testing of the decreased sensitivity towards 17 OPs and

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carbamates [59]. Although the combination of mutations generally increases the insensitivity of the

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enzyme [59, 60], there is an important contribution of the nature of the pesticide, and some

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compensation of multiple mutations for fitness cost of the individual mutations [95]. The respective

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roles of high recombination rates within the Ace locus [96] or of recurrent point mutations in

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achieving the mutation combinations that yield high resistance and low fitness costs are still unclear

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[97].

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In addition to the functional constraint, there is also a molecular constraint, as shown in

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mosquitoes. Here the key carbamate resistance mutation Gly119Ser has until now been found only

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when Gly is encoded by GG(CT) in the susceptible populations of the species. The Gly GG(CT)

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codons can be mutated to Ser by a G to A transition. In contrast, the GG(AG) codons would need

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two adjacent mutations to obtain Ser, a very rare event. This explains why Anopheles gambiae can

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become resistant through insensitive AchE while Aedes aegypti so far has not [98]. Such a

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constraint helps to predict the resistance risk in closely related mosquito species as indeed the Ace 1

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sequence including the Gly119 codon is phylogenetically constrained.

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3.3. Mutations in the GABA receptor gene

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Mutations in the GABA receptor gene (Rdl) are less diverse than in the VSSC and Ace

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genes. A mutation at Ala301 was found in 27 species from 6 orders of arthropods. This mutation in

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the second membrane spanning domain, lining the ion channel pore is most commonly Ala3010Ser,

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with some reports of Ala301Gly or Asn (Table 3, [37-39, 99-119]). Mutation at this site ( also

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designated as position TM2 A2', review in [120]) confers high resistance to cyclodiene insecticides

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and the initial work on Drosophila led to the first cloning of an insect GABA receptor and the

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precise identification of the dieldrin target site [121]. Cyclodiene resistance is very widespread

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[122]. It is likely therefore that the point mutation at Ala301 is one of the most common examples

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of parallel evolution at the molecular level. The Rdl gene is duplicated in cyclodiene resistant

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Myzus persicae, with one locus carrying Ala301Ser alleles (apparently not related to resistance) and

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the other carrying either the wild type Ala301 or the endosulfan resistance-conferring Ala301Gly

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allele [109]. A two nucleotide mutation Thr305Leu was reported in dieldrin-resistant southern cattle

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tick [119] and this position (known as position T6') is important for the interaction with cyclodienes

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[123]. Resistance to fipronil has been associated with mutations at Ala301 in combination with a

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second mutation, Thr350Met or Arg357Gln [37-39]. The Ala301Gly mutation of Drosophila

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simulans [37] appears to be optimal for fipronil resistance when tested in transgenic D.

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melanogaster, with the associated Thr350Met mutation in the third transmembrane domain possibly

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contributing to the reduction of fitness cost [124]. At least one of the Heliothis virescens Rdl genes

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is Ser301 in the wild type state, and this confers decreased sensitivity of the homo-oligomeric

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receptor to fipronil [125]. There are reports of other mutations found in combination with the

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Ala301 mutation (Val332Ile in Anopheles funestus, [101] and Ile281Thr in Bemisia tabaci, [113])

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but their relevance to the resistance phenotype has not been clearly demonstrated. The GABA

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receptor case illustrates the danger of the snowball effect, when several early studies report just one

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mutation, later studies in other organisms often merely check for that mutation on a partial PCR

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product of the gene and do not check the full sequence for possible additional mutations that might

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prove to be important.

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3.4 Point mutations in the genes for other targets

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Point mutations in the genes for most other known targets have now been documented and

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these are listed in Table 1. In most cases, these are single reports of single mutations, and strong

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evidence that the mutation is responsible for the resistance phenotype is not always available.

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However, these are also often among the first reports of the (presumed) molecular basis of

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resistance to some of the new classes of pesticides, so that their value for early monitoring is very

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important. As with Rdl, a resistance-associated point mutation can be instrumental in the

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identification of a target, for instance chitin synthase and cytochrome b as target of etoxazole and

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bifenazate in spidermites [29, 40]. In the latter case, similar mutations have been found in the Qo

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pocket of cytochrome b in several spider mite species and strains [126, 127]. As noted previously

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[20], point mutations in known target sites are easiest to document precisely, so there may be a bias

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in the relative importance of such mutations in the resistance literature. In addition, experimental

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mutagenesis followed by selection can be used to "predict" the likely sites of resistance point

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mutations [128, 129]. In some cases documented in spider mites using large initial sample sizes of

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high standing genetic variation, target-site resistance can be selected in the laboratory without the

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use of mutagens and using strains that were never exposed to the selecting compound [40, 130].

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An additional complexity is that the copy number of target-sites is not the same for all species. For

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example, while GABA- and glutamate-gated chloride channels consist of proteins encoded by

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single copy genes (Rdl and GluCl, respectively) in insect of which genomes have been sequenced 7 Page 7 of 46

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so far (except Lepidoptera), the spider mite T. urticae has three Rdl orthologues, and six GluCl

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orthologues [41]. A fine modulation of function is apparently achieved in insects by alternative

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splicing, and by multiple gene copies in mites. In the genome sequence of the London strain (an

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acaricide susceptible strain,[73]), TuRdl2 and TuRdl3 carry the Ala301Ser and Thr305Leu

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substitutions, while TuRdl3 has a Ala301His and Thr305Ile substitution. As the tick R. microplus

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has Ala301 in the wild type Rdl, this is not a general feature of Acari, and these mutations

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potentially reflect the massive selection with organochlorines in the past. It also suggests that an

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accumulation of mutations in the different receptor copies is needed to develop resistance. A similar

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accumulation of mutations was also reported for GluCls and abamectin resistance, as resistance

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strains carry the multiple substitutions Gly323Asp in TuGluCl1 and Gly323Glu in TuGluCl3 [41].

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Whether the copy number of the target-site significantly influences resistance development is at

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present not known, and multiple copies might also confer an advantage, as they might partially

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compensate for fitness-costs (see section 4 and 5 on gene copy number and redundancy)

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3.5. Point mutations and toxicokinetic resistance

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Two mutually exclusive mutations in the αE7 esterase gene of some higher Diptera are

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linked to broad OP resistance distinguishable by their selectivity towards to diazinon resistance

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(Gly137Asp) and to malathion resistance (Trp251Leu/Ser). The Gly137Asp mutation is located in

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the oxyanion hole of the enzyme and the Trp251Leu in the acyl binding pocket ([32, 33]; review in

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[131]). The αE7 is a carboxylesterase (also known as ali-esterase or B-esterase) of unknown

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endogenous substrate, although its X-ray structure suggests activity on fatty acid methyl esters

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[132]. The mutations, in particular Gly137Asp abolish the ali-esterase activity and confer a

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comparatively small, but toxicologically significant OP hydrolase activity. This can be confirmed

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by the resistance obtained by transgenic expression of the Lucilia cuprina Asp137 enzyme in

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Drosophila [133]. In contrast the Asp137 variant of the B1 esterase of C. pipiens does not confer

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resistance in transgenic flies [134].

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A single point mutation Leu119Phe in the GSTe2 gene in Anopheles funestus confers

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DDT and pyrethroid resistance [21], and the effect of this mutation on resistance is enhanced by

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overexpression of the gene. X-ray crystallography of the enzyme shows that the mutation opens the

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active site relative to the wild type enzyme and to the A. gambiae GSTe2 [135] and favors binding

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of DDT.

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More complex than a single non-silent nucleotide change, a recent example of gene

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conversion involving two adjacent, recently duplicated P450 genes was reported in the cotton

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bollworm Helicoverpa armigera. Here, the hybrid gene CYP337B3 arose (probably) from a gene

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conversion event resulting from unequal crossing over at the locus encompassing CYP337B1 and

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CYP337B2 [47, 48]. This has happened independently at least twice, and in each case the new gene

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encodes a P450 enzyme uniquely capable of metabolizing fenvalerate. However, CYP337B3 may

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not be the only gene involved in fenvalerate resistance in China [136].

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3.6 Point mutations causing ectopic expression or disrupting expression

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In some cases a stop codon can cause ectopic production of the protein: A stop codon at

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Leu569 of the maltase gene Cpm1 of Culex pipiens preserves the enzymatic function of the N-

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terminal portion of the protein, but prevents its C-terminal GPI anchoring to the midgut membrane.

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This causes resistance to Bin toxins of Bacillus sphaericus which bind to the now soluble Cpm1

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protein in the midgut lumen without toxic consequences [46]. A similar result is achieved in two

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other resistant alleles. In Cpm1BP a stop codon at Gln396 leads to a nonfunctional protein while in

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Cpm1BPdel a transposon insertion in exon 2 unmasks a cryptic intron splice site, leading to the

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production of a modified protein that can no longer bind the Bin toxin [137]. A single nucleotide

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deletion in the orthologous Cqm1 maltase gene of Culex quinquefasciatus also introduces a

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premature stop codon and lack of expression [138].

282

Small indels causing a change in subcellular location have been reported for

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acetylcholinesterase. In the olive fruit fly, a 3 Gln deletion at the C-terminal portion of the protein

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increases the efficiency of GPI anchoring of the enzyme in the synaptic cleft, thereby effectively

285

decreasing sensitivity to OPs [44, 45].

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The effects of nonsense mutations, indels and transposon insertions can lead to different

288

results: expression of a modified protein, lack of expression of a functional protein or increased

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expression, and we have therefore classified such cases as structural mutation, up or downregulation

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accordingly. These mutational events are sometimes more difficult to identify precisely than simple

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point mutations, but future work may show that a finer grain of the molecular classification of

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resistance can provide more insights.

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4. Gene upregulation and resistance

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Upregulation of genes involved in pesticide detoxification is very common, but the precise

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molecular mechanism has in most cases remained obscure (Table 4, [88, 91, 109, 137-201]). There

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are therefore very many reports of resistance associated with elevated esterases, glutathione S-

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transferase and cytochromes P450s [202-204]. There are far fewer reports where the mutation

300

causing this upregulation has been described. Upregulation is however not restricted to

301

toxicokinetic resistance, and an increase in the amount of target is one way to decrease the effect of

302

its inhibition. For instance, amplification of the Ace gene in spider mites has been reported (see

303

below) [88, 147]. In some cases, rather than causing increased metabolism, increased GST levels

304

can attenuate oxidative stress caused by the pesticide [205], or serve to sequester rather than

305

metabolize the pesticide [206].

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4.1 Gene duplication and gene amplification

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Gene duplication is a simple way to increase gene product, and recent work has shown

310

that the frequency of gene duplication can be much higher that the frequency of mutations at a

311

single nucleotide. In Drosophila melanogaster a new whole gene duplication rate of 1.75 x 10-7 per

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gene per generation was estimated, versus a single nucleotide mutation rate of 5.5 x 10 -9 per site per

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generation [207]. The rate of partial gene duplication or deletion is even higher. As shown in Table

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4, gene duplications have been reported for both toxicokinetic and toxicodynamic resistance, but

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not in any case of gut toxin resistance. It is likely that many more cases of gene duplication linked

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to resistance will be found, as the techniques to detect them are becoming more rigorous.

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Gene amplification has long been known to occur with esterases linked to OP resistance in

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mosquitoes and aphids (reviews in [158, 159, 208]). The precise mechanism, such as onion-skin

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multiple replication, perhaps followed by translocations or reintegration of extrachromosomal

320

elements with or without the aid of transposable elements, multiple sequential tandem duplications,

321

etc. is still more obscure. Chromosomal regions with amplified genes are difficult to sequence and

322

assemble, and this has not facilitated the precise description of the amplification events. Because

323

such processes leading to gene amplification are more complex than gene duplication, we have

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therefore kept amplification (often with only an estimate of the number of gene copies) separate

325

from "simple" duplication in this survey. In the Est-3-Est-2 locus of Culex mosquitoes encoding the

326

A and B esterases, various resistant alleles are found throughout the world, following amplification

327

of either one gene, or both tightly linked genes. In China, the A11-B11 allele results from a

328

duplication of the Est-3 (A11) gene followed by an amplification of the pair of genes (A11-B11

329

allele) [156]. The phenotypic consequence of esterase amplification in mosquitoes and aphids is a

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combination of two toxicokinetic mechanisms, increase in sequestration and pesticide hydrolysis.

331

The ratio of the two is dependent on the pesticide and enzyme [157, 209]. Moderate levels of

332

amplification of the Ace gene have been observed in monocrotophos resistant T. urticae. Here,

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multiple copies of the gene for an insensitive AChE can restore normal level of catalytic activity in

334

the monocrotophos resistant mites ([147, 148], also reviewed in this issue by Lee et al. [210]).

335 336

Amplification is not restricted to esterases, but has now also been reported for GST genes

337

[91, 211] and P450 genes [161, 164]. In the case of the CYP6CY3 gene in Myzus persicae

338

(nicotianae), the relative roles amplification and overexpression are difficult to distinguish, and

339

neonicotinoid resistance may in fact be a (cross-) tolerance due to an earlier host shift on tobacco

340

[212]. The CYP9J26 gene and the ABC half transporter gene ABC B4 are both amplified about 6-7

341

times in a pyrethroid resistant strain of Aedes aegypti from the Caribbean [164]. Gene amplification

342

is usually associated with a proportional increase in gene expression, but the two are not precisely

343

matched. In the coamplified A2-B2 esterases of Culex quinquefasciatus, the B2 esterase is

344

expressed ten times more than A2 [213]. In the case of amplified esterases in aphids, expression can

345

be modulated by methylation, so that the expression can be silenced reversibly [214]. The size of

346

the amplicon can be large enough to encompass other genes, as seen with the coamplification of the

347

A2-B2 esterases with an aldehyde oxidase in C. quinquefasciatus [215]

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4.2 Gene upregulation

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Mutations that positively affect gene expression without change in gene copy number can

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be separated between cis and trans upregulation. Cis regulation implies a mutation in the regulatory

355

(mostly in the 5' UTR) sequences of the gene, and at first sight it might seem difficult to achieve up

356

(or down) regulation instead of dysregulation (causing expression at the wrong time or place). Yet

357

examples of transposable element insertion in 5'-UTR regions are now well established, notably in

358

the case of the Cyp6g1 gene of Drosophila [144, 165]. The CYP6G1 enzyme metabolizes both

359

DDT and neonicotinoids and this causes cross resistance to both classes of compounds. The Accord

360

insertion in the Cyp6g1 gene carries its own specific transcriptional enhancers [216]. In Culex

361

pipiens, CuRE1 insertion in the CYP9M10 gene is similarly associated with an upregulation of a

362

pyrethroid metabolizing P450 ([146, 217] see however [167]). The CYP6D1 gene of the house fly is

363

overexpressed in pyrethroid-resistant strains such as LPR. It has been mapped to chromosome 1,

364

and overexpression is controlled by factors on chromosome 1 and 2, so at least one factor is acting

365

in trans [175]. A cis regulation is also suggested however: the 5' UTR region of resistant strains is

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characterized by a 15-bp deletion that causes a decreased binding of the Gfi-1 zinc finger

367

transcription repressor [218]. Imidacloprid resistance in the Q biotype of Bemisia tabaci is

368

associated with overexpression of CYP6CM1, and a mutation in cis is suggested by the fact that

369

three SNPs in the intron of the gene are strongly associated with resistance [221]. The R and S

370

alleles are therefore different in some (still unknown) respect. A 48-fold AC tandem repeat

371

microsatellite in the promoter of the CYP6CY3 gene of Myzus persicae nicotianae was shown to

372

cause overexpression of this nicotine and neonicotinoid-metabolizing P450 [161]. Cis upregulation

373

of CYP6ER1 [222] or of CYP6AY1 [168] in imidacloprid- and buprofezin-resistant strains of N.

374

lugens is strongly suggested by the close association of promoter variants with resistance. In the

375

latter case, analysis of promoter activity in vitro confirmed the higher activity of the resistant

376

variant [168]. Another example of upregulation in cis was provided by the study of Berrada and

377

Fournier [223] who transformed Drosophila with an extra copy of the Ace gene, and favored its

378

movement in the genome by crosses with a fly strain carrying an active source of transposase. After

379

34 generations of exposure to parathion, these authors obtained a strain with resistance caused by

380

the overexpression of the Ace transgene. Although this is a fully experimental situation, it clearly

381

demonstrated that resistance can be achieved by overexpression of a (sensitive) target, and that this

382

overexpression is dependent on the genomic insertion site of the transgene (i.e. a cis regulatory

383

effect).

Page 12 of 46

384

Trans upregulation is perhaps the most difficult to describe in molecular detail, as the

385

target gene, as well as the actual mutation in trans need to be identified. Trans upregulation

386

typically targets more than one gene (pleiotropic effect) and the mutation can be anything that

387

affects the level (synthesis, metabolism) or nature (structure or post-translational modification) of

388

the trans acting factor (a transcription factor, or a ligand thereof). Genetic evidence therefore

389

usually precedes molecular evidence. In a DDT-resistant strain of Aedes aegypti, overexpression of

390

GST-2 involves a trans acting locus [176]. Overexpression of CYP6A1 and CYP6D1 in the house

391

fly are each regulated in trans [172, 175]. The overproduction of the diazinon-metabolizing

392

CYP6A1 is genetically linked to the αE7 esterase locus [173]. It was suggested that the absence of

393

the wild-type Gly137 allele of the αE7 gene releases the transcriptional repression of CYP6A1. This

394

did not resolve the question of the nature of the repressor. While Sabourault et al. proposed a role

395

for an endogenous product of the αE7 esterase enzyme, it is also possible that a functional

396

regulatory gene is located close to the αE7 gene in wild type flies. In any case, pleiotropic

397

upregulation of CYP6A1, along with other genes (CYP12A1, GST1) confers a higher level of

398

resistance to diazinon in house flies than the mutant ali-esterase alone [204]. Upregulation of some

399

GPCR genes in permethrin-resistant C. quinquefasciatus strains appears causally related to the

400

upregulation of key P450 genes and resistance, as shown by RNAi experiments, but how these

401

GPCRs regulate P450 expression is unclear [224]. The upregulation of CYP6A2 in some

402

Drosophila strains is correlated with DDT resistance, and caused by both cis and trans factors [219,

403

220]. The CncC/Keap1 pathway is constitutively active in those strains and both CncC disruption or

404

Keap1 overexpression can block overexpression. A chromosome 3 factor from the resistant strains

405

[220] causes the activation of the CncC/Keap1 pathway [225], but although both CncC and Keap1

406

map to the appropriate region on chromosome 3 [220], their coding region shows no mutation in the

407

resistant strains [225]. Furthermore, overexpression of CYP6A2 alone in transgenic flies does not

408

confer DDT resistance [226].

409

These examples show the molecular complexity of trans upregulation where mutations in

410

genes with regulatory functions affect multiple target genes, and where some but not all target

411

genes contribute to resistance. These pleiotropic effects predictably cause fitness deficits [20].

412

Increasing evidence for pleiotropic up regulation of multiple genes related to detoxification in

413

resistant strains of mosquitoes and Lepidoptera (reviewed in [227]) suggests that resistance

414

mutations with trans effects are widespread. However, their precise identification remains a

415

challenge.

416 417

5. Gene downregulation and resistance 13 Page 13 of 46

418 419

The pioneering work of Wilson on resistance to juvenile hormone analogs in Drosophila

420

led to the discovery of Met, a transcription factor of the bHLH-PAS family that serves as a juvenile

421

hormone receptor. Several alleles of the Met (Methoprene tolerant) gene were found, and the

422

common feature of these alleles is disruption (e.g. nonsense mutations, transposon insertions) of the

423

gene leading to reduced levels or no transcript being produced. Wilson [228] predicted the general

424

importance of transposons as mutational agents for resistance, as well as the importance of

425

functional redundancy [180]. Indeed, gene disruption in the case of Met is made possible by a

426

degree of functional redundancy with a close paralog, Gce [229]. The importance of redundancy

427

was also noted by Heckel [230]. All the currently known examples of gene disruption linked to

428

resistance are examples where some redundancy is available, for toxicodynamic resistance as well

429

as for gut toxin resistance (Table 4).

430

Disruption of several types of nicotinic acetylcholine receptor subunits, either

431

experimentally (by EMS mutagenesis, [184, 192] or naturally [186, 188]) has been linked to

432

spinosad or neonicotinoid resistance. Cyromazine resistance obtained experimentally in Drosophila

433

by disruption of a phosphatidylinositol kinase-like kinase (PIKK) gene [195] is perhaps also made

434

possible by some degree of redundancy.

435

RNAi approaches, as in the cyromazine resistance study, are now widely available to test

436

the risk of resistance by gene disruption. For instance RNAi of the ryanodine receptor in

437

Leptinotarsa decemlineata and Sogatella furcifera can confer chloranthranilipole resistance [231].

438

Decreased expression and 42 bp deletion in resistant Plutella xylostella RyR has been reported

439

[197] confirming that gene disruption of the RyR can be found in natural settings, or that the

440

Gly4946Glu point mutation is already an adaptive solution with minimal fitness cost. Experimental

441

RNAi works well in Coleoptera but does not routinely work in all arthropods. Other ways to disrupt

442

or modify genes, for instance the CRISPR/Cas or TALEN techniques work in Lepidoptera where

443

RNAi is less dependable [232].

444

The greatest diversity of disruptive mutations is found in gut toxin resistance. The genes

445

encoding gut proteins that bind toxins are all members of multigene families (cadherins,

446

aminopeptidases, alkaline phosphatases, ABC transporters for Bt toxins, maltases for Bacillus

447

sphaericus toxins). Disruptive mutations (review in [233], see also [191]) include transposon

448

insertion, indels of various sizes causing truncations of the proteins in intracellular or extracellular

449

domains, exon skipping, and single amino acid substitutions.

450

Page 14 of 46

451

There are not many examples of cis and trans downregulation of genes linked to

452

resistance. Perhaps this is due to the fact that gene disruption is a more effective way to remove a

453

target with non-vital function. Down regulation of alkaline phosphatases appears widespread in

454

Lepidoptera [177]. Closer to the actual mutation(s), two examples of trans downregulation of

455

aminopeptidase genes linked to Cry1Ab and Cry1Ac resistance have been reported [174, 201]. In

456

the latter case, down regulation of APN1 seems associated with upregulation of APN6, thus

457

supporting the idea that downregulation of a resistance gene is possible when a redundant gene can

458

compensate for the loss of function. Decreased activation of protoxins [178, 179, 233, 234] as well

459

as decreased activation of proinsecticides is well known, but to date there is no study providing the

460

molecular genetic details on the mutations underlying such lack of activation as resistance

461

mechanism.

462 463

6. Selection, mutation, recombination and drift.

464 465

Mutation and selection, along with recombination and genetic drift are generally

466

considered the four fundamental forces of evolution [235]. While selection is the main

467

"operational" factor in the dynamics of resistance, the other three forces are stochastic events with

468

non-random outcomes. We have described above the variety of mutational events that generate the

469

variability upon which selection can occur, leading to resistance. Below, we will briefly mention

470

some aspects of selection, recombination and drift as they relate to resistance.

471 472

6.1 Selection

473

Selection is far from uniform in space and time. In the case of Bt crops, the operational

474

control of selection is the key aspect of resistance management, with a high degree of selection on

475

the protected crop maintaining resistance functionally recessive, and the absence of selection in the

476

refuge maintaining the pool of susceptibility. This operational control is working despite the

477

sometimes surprisingly high frequencies of resistance alleles in untreated populations [15, 236], and

478

despite the ease with which disruptive mutations at various steps of the mode of action of Bt toxins

479

can lead to resistance.

15 Page 15 of 46

480

More troublesome is the collateral damage of pesticide use leading to highly variable

481

degrees of selection on non-target organisms. The use of pesticides in agriculture can cause the

482

selection of mosquitoes (e.g. [237-239]), and this can negatively impact vector control. Similarly,

483

runoff of pesticides can be toxic to aquatic copepods, resulting in the selection of the same

484

resistance mutations as seen in insect pests [240]. This selection beyond the intended target is

485

perhaps best documented in Drosophila where field-collected strains can easily be shown to be

486

resistant to a number of pesticides in a variety of ways, even though the fruit fly is only rarely

487

considered a target pest, except in some wine-growing areas [241].

488

Selection is therefore necessary for resistance to develop, but it is not as simple a factor as

489

may appear. This has led to debates about the value of laboratory selection as model for field-

490

evolved resistance and about the intensity of selection in determining the type and number of

491

resistance genes being selected [5, 242, 243]. The population size and its standing genetic variation

492

is of paramount importance. Perhaps the type of selecting agent (does it have one target or more ?)

493

and the ecological history/trajectory of the resistant species should be integrated into such

494

discussions as well. Does feeding on alkaloid-containing plants such as nicotine predispose to

495

neonicotinoid resistance [161]? Do frequent host plant changes and polyphagy predispose to

496

metabolic resistance [244]? Did long term evolution along entomopathogenic bacteria contribute to

497

the redundancy of the gut proteins that toxins use to kill insects?

498 499

6.2 The importance of recombination

500

The importance of recombination in driving the evolution of resistance is increasingly

501

well recognized. As a source of unequal crossing over, recombination is a powerful generator of

502

copy number variation, and the multiple cases of resistance linked to gene duplications have been

503

discussed above. When recombination occurs in genomic regions carrying clusters of genes

504

encoding detoxification enzymes, it can cause gene conversion, which is a way of generating new

505

sequences by molecular lego. Pyrethroid resistance in H. armigera discussed above (section 3.5) is

506

a remarkable example, and the rapid progress in genome sequencing may uncover other cases that

507

were difficult to detect with older gene by gene cloning techniques. Depending on the divergence

508

between the two original sequences, gene conversion can amount to the simultaneous mutation at

509

many sites of an enzyme. The probability of a "gain of function" as in fenvalerate metabolism is

510

low, but gene conversion can be seen as a jump forward through multiple point mutational changes.

511

(i.e. crossing a fitness valley). Recombination between close, sibling taxa (introgression) can carry

512

adaptive mutations as shown for the kdr and Ace mutations in Anopheles gambiae [245, 246].

513

Page 16 of 46

514

Recombination can result in the accumulation of different types of mutations at the same

515

locus, although recurrent mutations can also be invoked. It was predicted by functional expression

516

of the mutant channels in Xenopus oocytes that a single crossing over event between a Ser989Pro +

517

Val1016Gly carrying VSSC haplotype and a Phe1534Cys carrying haplotype would synergistically

518

increase resistance to pyrethroids in Aedes aegypti to very high levels [247]. Such a recombination

519

event may have already occurred in Burma [248]. When recombination generates tandem duplicates

520

of the resistance gene, one R copy and one S copy, the dissociation of alleles at meiosis is

521

effectively prevented. Thus R;S duplicates represent the best of both worlds, or “segregation

522

avoidance” because the R and S copies are inherited as a unit. Heterozygote advantage is

523

maintained without the risk of producing deleterious homozygous individuals at every generation,

524

i.e. S/S when the pesticide is present would be lethal, R/R when the pesticide is absent would have

525

deleterious fitness costs. The generation of such R;S duplicates is complex, and it is not always

526

obvious whether the R;S haplotype is most favored as homozygote or heterozygote, because the

527

imbalance in gene dosage has different effects depending on the remaining endogenous function of

528

the R gene. Such cases have been documented, for instance for insensitive AchE in Culex pipiens

529

where insecticide selection actually favors the retention of the duplicated gene [139, 249]. Copy

530

number variation of the VSSC gene in Aedes aegypti has been reported in Brazil, with multiple

531

alleles carrying different point mutations [145]. An R;S duplication of Rdl has recently been

532

documented in Drosophila [130]. In the latter case, the duplication was related to ectopic

533

recombination between transposable elements.

534 535 536 537

6.3 Genetic drift

538

factor contributing to the genetic variation in populations. In the context of arthropod resistance,

539

genetic drift might affect mutation frequencies especially in small populations, such as genetically

540

isolated greenhouse populations, or at the time when ‘exotic, alien pests’ colonize a new region. In

541

such cases, drift might lead to fixation or loss of resistance mutations without selection pressure.

Genetic drift, the change in allele frequencies in populations due to random sampling, is a

17 Page 17 of 46

542

In contrast, genetic drift has been shown to be the major contributing factor in the evolution of

543

mitochondrial genes. A number of compounds target the mitochondrial electron transfer process,

544

and in some cases, mitochondrial genes have been implicated in resistance. This is the case for

545

strobilurin fungicides [250] as well as the antimalarial drug atovaquone [251]. Within arthropods,

546

mutations in the mitochondrially encoded cytochrome b cause target-site resistance to bifenazate,

547

and the inheritance of mitochondrial mutations was investigated in heteroplasmic mites [40]. For

548

mitochondrial genes, there is no recombination during meiosis and transmission to the offspring is

549

usually uniparental. While a diploid cell contains two copies of a nuclear gene, mitochondrial genes

550

are present in somatic cells in the range of thousands of copies. The emergence of any new

551

mitochondrial haplotype must be caused by a single mutation in a single copy of the mitochondrial

552

DNA of a (precursor of) female germline cell, resulting in a heteroplasmy (the existence of two

553

copies within one individual) in the next generation. As such, mitochondrial target-sites could be

554

considered relatively safe in the light of resistance development, as a single mutated mitochondrial

555

DNA copy is diluted by thousands of wild type copies. However, the frequency of mitochondrial

556

mutations has been shown to rapidly change in the offspring of individuals exhibiting heteroplasmy,

557

due to the rapid sorting and unequal segregation of mitochondria, also referred to as ‘genetic

558

bottlenecking’. Mutation frequencies in cytochrome b in the offspring of a number of individuals

559

with different starting frequencies were followed, and it was shown that this non-Mendelian

560

inheritance is subjected to rapid fixation by drift [40].

561 562

The dynamics of resistance is particular in organisms with haplodiploid reproduction

563

(arrhenotoky or pseudo-arrhenotoky), such as tetranychid mites, thrips, and whiteflies. Although

564

most studies are based on complex theoretical modeling and only limited experimental data is

565

available [252-254], recessive resistance mutations might be fixed more rapidly in populations by

566

fast selection on haploid males. However, the strength of this effect depends on many factors, of

567

which fitness differences between R males and RR females (gene dosage) are probably most

568

important [254]. Other factors influencing resistance dynamics are sexual dimorphism (size), type

569

of mutation (gain of function, loss of function) and selection pressure in the field. Similar dynamics

570

might occur in species with functional haplo-diploidy, where males fail to express and transmit

571

paternal resistance genes [255]. In addition, organisms with rare or seasonal sexual stages such as

572

aphids are limited in recombination to generate novel gene-combinations. However, a mutation or

573

gene combination, once acquired, can spread rapidly in populations under selection pressure [256].

574 575 576

7. Conclusions

Page 18 of 46

577 578

Our survey of the classes of mutations involved in different types of resistance reveals a

579

broader range of mutations than surveys of phenotypic evolution in eukaryotes in general, where cis

580

regulation has an important share [257, 258]. In particular, we find relatively more cases of gene

581

duplication or amplification, and of gene disruption. The latter were not present in an earlier survey

582

of resistance [20], but are predominant in resistance to gut toxins. The relationships between

583

different classes of mutations and their dominance or fitness costs can be analyzed anew. We hope

584

that the data and patterns described here can now be exploited in the continued quest for rational

585

resistance management strategies.

586 587

Acknowledgements

588 589

WD is a postdoctoral fellow of the Fund for Scientific Research Flanders (FWO)

590 591

19 Page 19 of 46

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Figure Legends

Figure 1 - Genotype and phenotype of resistance in arthropods. Resistance classified along molecular and biochemical/physiological dimensions reveals the broad specrum of mutational changes described in the literature.

Figure 2 - Summary of mutation diversity in the voltage sensitive sodium channel (VSSC) gene or athropods. A: VSSC diversity amoung different orders; B: Relative importance of the kdr and super kdr mutations. The data were compiled from [49] and updated. See text for details. Table 1 - Mutation(s) affecting the coding sequence of a gene, resulting in either toxicodynamic, toxicokinetic or gut toxin resistance mutations affecting coding sequence

toxicodyn amic resistanc e

point mutation:

A301S/G in GABA Rdl

one position

species *

ref.

toxicoki netic resistan ce

cyclodienes

see Table 3

Tab le 3

L119F in GSTe2

DDT

A. funestu s

[2 1]

R81T in nAchR beta 1 G275E in nAchR alpha 6

neonicotinoi ds

M. persicae

[22]

spinosad

F. occident alis, T. palmi

[23, 24]

G4946E in RyR

diamides

[2527]

L64I in βAOR

amitraz

I1017F in CHS1

etoxazole clofentezine hexythiazox spiromesife n

P. xylostell a B. microplu s T. urticae T. vaporaro rium

[31]

E7 esterase

OPs

L. cuprina, L. sericata M. domesti ca, C. hominiv orax A. calandr ae

[3 236 ]

E645K in ACCase point mutation:

pesticide

species

re f.

gut toxin resista nce

pestici de

spec ies

re f.

[28] [29, 30]

Ace 1

OPs, carbamates

see Table 2

Tab le 2

Ace 2

OPs, carbamates fipronil

see Table 2 D. simulans , S. furcifera

Tab le 2 [3739]

> one position

GABA Rdl

pestic ide

Page 41 of 46

indels and nonsense mutations

VSSC1

DDT, pyrethroids

see Figure 2

cyt b Q(o) site GluCl

bifenazate

RyR

chlorantrani liprole

T. urticae T. urticae P. xylostell a

Ace 2 Delta3Q1

OPs

abamectin

B. oleae

Fig ure 2 [40] [41, 42] [43]

[44, 45]

Cpm1 maltas e L569St op2

B. sphaeri cus

[4 6]

C. pipie ns

bin toxin

recombin CYP337 fenvale H. [4 ation: B33 rate armiger 7, gene a 48 conversio ] n *species abbreviations: Frankliniella(F.) occidentalis, Thrips(T.) palmi, Trialeurodes(T.) vaporarorium, Lucilia(L.) cuprina, Lucilia(L.) sericata, Cochliomyia(C.) hominivorax, Anisopteromalus(A.) calandrae, Bactrocera(B.) oleae, 1 small indel may favor GPI anchoring; 2 ectopic expression; 3 from CYP337B1 and B2

Table 2 – Overview of mutations in arthropod Ace1 and Ace2 position in mature Torpedo AchE*

mutation in Ace 1

species**

ref.

E73 F78 E81K G119

D128

G119S

[62]

G119S

A. gambiae, C. pipiens A. albimanus

S119G

T. urticae

[64]

G119A

N. lugens

[65]

D128E

T. urticae

[64]

V150

G227

A201S

G227A

A. gossypii

[61, 69, 70]

P. xylostella

[71]

C. suppressalis

[72]

T. urticae

[73]

P. xylostella

[71]

V238 N280

T280A

T. urticae

[76]

F290

F290V

C. pomonella

[77]

C. pipiens

[79]

G328

G328A

T. urticae

species**

ref.

E73G F78S F78L E81K

D. melanogaster D. melanogaster A. gossypii D. melanogaster

[59] [60] [61] [59]

I129V/T

D. melanogaster

[60]

I129V

M. domestica

[66]

I129V

B. oleae

[67]

V150L

M. domestica

[66, 68]

G227A

D. melanogaster

[60]

G227A/V

M. domestica

[66, 68]

G227A

H. irritans

[74]

S238G

L. decemlineata

[75]

F290Y

D. melanogaster

[78]

M. domestica

[66, 68]

[63]

V129

A201

mutation in Ace 2

[76]

C. hominivorax

[80]

F290L

R. padi

[81]

G328A

D. melanogaster

[59]

M. domestica

F330

F330S

N. lugens

[65]

F331

S331F

M. persicae

[83]

F331W

C. capitata

[82]

C. tritaeniorhynchus

[84]

Page 42 of 46

L336

S331F

A. gossypii

[69, 70, 85]

F331W

B. tabaci

[86]

F331W

T. urticae

[76]

F331W

T. kanzawai

[87]

F331C

T. urticae

[64]

F331Y/W

T. evansi

[88]

F331H

N. lugens

[65]

F331H

L. striatellus

[89]

L336S

S. avenae

[90]

G396

G396S

B. oleae

[91]

W435

W435R

S. avenae

[90]

A441G

A441

P. xylostella

[71]

*mutations at positions with bold font have been experimentally validated ** species abbreviations: Anopheles(A.) albimanus, Aphis(A.) gossypii, Chilo(C.) supressalis, Haematobia(H.) irritans, Cydia(C.) pomonella, Rhopalosiphum(R.) padi, Ceratitis(C.) capitata, Tetranychus(T.) evansi, Tetranychus(T.) kanzawai, Laodelphax(L.) striatellus, Sitobion(S.) avenae

Table 3 – Overview of mutations in the arthropod Rdl gene position in Rdl of D. melanogaster* A301 TM2 A2’ position

mutation

species**

pesticide

ref.

A301S

D. melanogaster

dieldrin

[99]

A301S/G

D. simulans

cyclodienes

[100]

A301G

D. simulans

fipronil, dieldrin

[37]

A301S

A. funestus

dieldrin

[101]

A301G

A. gambiae

dieldrin

[102]

A301S

dieldrin

[102]

A301S/G

A. arabiensis Anopheles sp. (6 species from Indonesia)

cyclodienes

[103]

A301S

H. irritans

cyclodienes

[104]

A301S

C. quinquefasciatus

cyclodienes

[105]

A301S

A. albopictus

cyclodienes

[105]

A301S

M. domestica

cyclodienes

[100]

A301S

C. felis

fipronil (?)

[106, 107]

A301N

L. striatellus

fipronil

[38]

A301S

T. castaneum

dieldrin

[100, 108]

A301G

M. persicae

[109]

A301S

P. xylostella

endosulfan fipronil, dieldrin, endosulfan

A301N

S. furcifera

fipronil

[39, 111]

A301S

O. oryzae

fipronil

[112]

A301S

B. tabaci

cyclodienes

[113]

A301S

H. hampei

endosulfan

[114, 115]

A301S

B. germanica

cyclodienes

[116, 117]

A301S

P. americana

cyclodienes

[100]

[110]

A301S

D. vergifera

aldrin

[118]

I281

I281T (with A301S)

B. tabaci

cyclodienes

[113]

T305

T305L

B. microplus

dieldrin

[119]

V332

V332I (with A301S)

A. funestus

dieldrin

[101]

T350

T350M (with A301G)

D. simulans

fipronil/dieldrin

[37]

Page 43 of 46

R357

R357Q (with A301N)

fipronil

S. furcifera

[38, 39]

* mutations at positions with bold font have been experimentally validated ** species abbreviations: Anopheles(A.) arabiensis, Aedes(A.) albopictus, Ctenocephalides(C.) felis, Tribolium(T. castaneum, Oulema(O.) oryzae, Hypothenemus(H.) hampei, Blatella(B.) germanica, Periplaneta(P) americana, Diabrotica(D.) vergifera

Table 4 - Mutations affecting gene expression levels resulting in either toxicodynamic, toxicokinetic or gut toxin resistance mutations affecting gene expression levels

toxicodynam ic resista nce

pesticide

specie s*

ref .

UPREGULATI ON duplication

cis UP

pesticide

specie s*

ref .

gut toxin resista nce

toxi n

species *

ref .

many reports1 Ace 1

OPs

C. pipiens P.xylos tella

amplification

toxicokinetic resistan ce

GABA/ Rdl

dieldrin

VSSC1

deltamethri n monocroto phos chlorpyriph os

Ace 1

A. gambia e M. persica e Drosop hila A. aegypti T. urticae T. evansi

[13 914 1]

E7 aliesterase

OPs

L. cuprin a

[14 2]

[10 9, 14 3]

Cyp6g1

DDT neonicotin oids

Droso phila

[14 4]

[14 5] [88 , 14 7, 14 8]

CYP9M 10 A2-B2, A4-B4, A5-B5, A8-B8, A9-B9 A1, A3B1, B1 esterase s A11-B11 esterase s E4 and FE4 esterase s

permethrin

C. qqf

OPs

C. pipiens C. qqf

[14 6] [14 915 5]

GST amplific ation2 GST amplific ation3 CYP6C Y3 (+ cis) CYP9M 6

pyrethroid s

CCEae3 a

temephos

ABC B4

pyrethroid s

Cyp6g1

DDT neonicotin oids

3

C. ttr 3

OPs

C. pipiens

[15 6]

OPs carbamate s pyrethroid s

M. persic ae S. gramin um N. lugens N. lugens

[15 715 9]

M. domes tica M. nicotia nae A. aegypt i A. aegypt i A. aegypt i Droso phila

[16 0]

terachlorvi nphos nicotine/oi ds permethrin

[91 ]

[16 1] [16 2] [16 3] [16 4] [14 4, 16 5]

Page 44 of 46

trans UP

mutations affecting gene expression levels

toxicodynam ic resista nce

pesticide

specie s*

ref .

Cyp12a 4 CYP9M 102 CYP9M 10 CYP6C Y3

lufenuron

CYP6A Y1 CYP6P9 a and b

imidaclopri d pyrethroid s

CYP6P4

pyrethroid s

CYP6A1

diazinon

CYP6D1

pyrethroid s

GST-212

DDT

M. domes tica A. aegypt i

toxicokinetic resistan ce

pesticide

specie s*

permethrin

Droso phila C. qqf

permethrin

C. qqf

nicotine/oi ds

M. nicotia nae N. lugens A. funest us

3 3

A. arabie nsis M. domes tica

[16 6] [14 6] [16 7] [16 1] [16 8] [16 9, 17 0] [17 1] [17 2, 17 3] [17 5]

APN1

Cry1 Ac

T. ni

[17 4]

toxi n

species *

ref .

ALP4

Cry1 Ac Cry1 Fa

[17 7]

protoxi n activati on4 protoxi n activati on4 ABC C2

Cry1 Ab

H. virescen s H. armigera S. frugiperd a M. unipunct a

Cry1 Ab

P. interpun ctella

[17 9]

Cry1 Ac

H. virescen s

[18 2]

ABC C2

Cry1 Ac

H. armigera

[18 5]

ABC C214

Cry1 Ac

P. xylostell a T. ni B. mori

[18 7]

S. frugiperd a P. gossypie lla H. armigera H. virescen s

[19 1]

[17 6]

ref .

gut toxin resista nce

DOWNREGU LATION no mechanism

disruption

Met5

methopren e

Drosop hila

nAchR a66

spinosyns

Drosop hila

nAchR a67

spinosad

P. xylostel la

nAchR a68

spinosad

[18 8]

ABC C215

Cry1 Ab

nAchR a69

spinosad

[19 0]

ABC C216

Cry1 Ac

nAchR a110

nitenpyram

B. dorsali s P. xylostel la Drosop hila

[19 2]

cadheri n17

Cry1 Ac

nAchR b211 PIKK CG327 4312

nitenpyram

Drosop hila Drosop hila

[19 2] [19 5]

cadheri n18 cadheri n19

Cry1 Ac Cry1 Ac

cyromazin e

[18 0, 18 1] [18 3, 18 4] [18 6]

[17 8]

[18 9]

[19 3] [19 4] [19 6]

Page 45 of 46

RyR13

cis DOWN

chlorantran ilipole

P. xylostel la

[19 7]

cadheri n20

Cry1 Ac

P. gossypie lla H. armigera

[19 8]

cadheri n21

Cry1 Ac

Cqm1 maltas e22 Cpm1 maltas e23 APN1

bin toxin

C. qqf3

[13 8]

bin toxin

C. pipiens

[13 7]

[19 9]

Cry1 H. [20 Ac armigera 0] trans DOWN APN Cry1 T. ni [17 Ac 4] APN Cry1 O. [20 Ab nubilalis 1] * species abbreviations: Culex(C.) quinquefascatus, Culex(C.) tritaeniorhynchus, Schizaphis(S.) graminum, Spodoptera(S.) frugiperda, Mythimna(M.) unipuncta, Plodia(P.) interpunctella, Trichoplusia(T.) ni, Bactrocera(B.) dorsalis, Bombyx(B.) mori, Ostrinia(O.) nubilalis, Pectinophora(P.) gossypialis 1 many reports of P450, CCE and GST overexpression without explicit molecular mechanism [202-204]; 2 5’ CuRE insertion cis acting, also duplication; 3 qqf, quinquefasciatus; ttr, tritaeniorhynchus; 4 no mechanism; 5 many types of disruptive mutations; 6 multiple types of gene disruption, EMS mutants; 7 several mutants - stop codons; 8 several types of indels - stop codon; 9 multiple types of splice variants truncated transcripts; 10 EMS 11 bp deletion; 11 EMS 53 bp deletion; 12 PIKK, phosphoinositide 3-kinase-related kinases, EMS nonsense mutation; 13 14 aa deletion; 14 30 bp deletion in P. xylostella; only mapped in T.ni; 15 single Tyr codon insertion; 16 deletion in ATP binding domain; 17 three deletions alleles; 18 stop codon; 19 disruption by retrotransposon insertion; 20 alternative splicing - many forms; 21 indel exon skipping; 22 1 nt deletion - stop codon, no expression; 23 transposon insertion - cryptic splice site - truncated receptor: other allele: indel; both disrupt

Page 46 of 46