How insects overcome two-component plant chemical defence: plant β ...

Biol. Rev. (2013), pp. 000–000. 1doi: 10.1111/brv.12066

How insects overcome two-component plantchemical defence: plant β-glucosidases as themain target for herbivore adaptation

Stefan Pentzold, Mika Zagrobelny, Fred Rook and Søren Bak∗

Plant Biochemistry Laboratory, Department of Plant and Environmental Sciences, University of Copenhagen, Thorvaldsensvej 40, Copenhagen

Dk-1871, Denmark

ABSTRACT

Insect herbivory is often restricted by glucosylated plant chemical defence compounds that are activated by plantβ-glucosidases to release toxic aglucones upon plant tissue damage. Such two-component plant defences are widespreadin the plant kingdom and examples of these classes of compounds are alkaloid, benzoxazinoid, cyanogenic and iridoidglucosides as well as glucosinolates and salicinoids. Conversely, many insects have evolved a diversity of counter-adaptations to overcome this type of constitutive chemical defence. Here we discuss that such counter-adaptations occurat different time points, before and during feeding as well as during digestion, and at several levels such as the insects’feeding behaviour, physiology and metabolism. Insect adaptations frequently circumvent or counteract the activity ofthe plant β-glucosidases, bioactivating enzymes that are a key element in the plant’s two-component chemical defence.These adaptations include host plant choice, non-disruptive feeding guilds and various physiological adaptations aswell as metabolic enzymatic strategies of the insect’s digestive system. Furthermore, insect adaptations often act incombination, may exist in both generalists and specialists, and can act on different classes of defence compounds. Wediscuss how generalist and specialist insects appear to differ in their ability to use these different types of adaptations:in generalists, adaptations are often inducible, whereas in specialists they are often constitutive. Future studies aresuggested to investigate in detail how insect adaptations act in combination to overcome plant chemical defences andto allow ecologically relevant conclusions.

Key words: insect herbivore-plant interactions, two-component plant chemical defence, β-glucosidases, β-glucosides,insect adaptations, feeding guild, gut pH, sequestration, generalists and specialists.

CONTENTS

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2(1) Overview: insect herbivores and two-component plant chemical defence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2(2) Specialist versus generalist insect herbivores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2(3) The role of plant β-glucosidases and β-glucosides in two-component plant chemical defence . . . . . . . . . 4

II. How non-adapted insect herbivores are affected by two-component plant chemical defence . . . . . . . . . . . . . . . 6(1) Alkaloid glucosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6(2) Benzoxazinoid glucosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6(3) Cyanogenic glucosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8(4) Glucosinolates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8(5) Iridoid glucosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8(6) Salicinoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

III. From feeding to digestion: targets for insect herbivore adaptations to two-component plant chemcial defencein a temporal context . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9(1) Before feeding: recognition, switching and selection of host plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9(2) During feeding: impact of the feeding guild . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

* Author for correspondence (Tel: +0045 3533 3346; E-mail: [email protected]).

Biological Reviews (2013) 000–000 © 2013 The Authors. Biological Reviews © 2013 Cambridge Philosophical Society

2 Stefan Pentzold and others

(a) Piercing-sucking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10(b) Leaf-snipping versus leaf-chewing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10(c) Leaf-mining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

(3) During digestion: physiological and metabolic adaptations to counteract two-component defence . . . . 12(a) An alkaline gut pH inhibits the plant β-glucosidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12(b) Reduction of endogenous insect β-glucosidase activity in the gut . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13(c) Specialized enzyme activity of insects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

( i ) Before hydrolysis by plant β-glucosidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14( ii ) After hydrolysis by plant β-glucosidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

(d ) Sequestration: spatial separation of plant β-glucosidase and β-glucoside in the insect . . . . . . . . . . . . . . 15(e) Single amino acids counteract plant β-glucosidase activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

(4) Do generalists and specialists have different types of adaptations? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17IV. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17V. Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

VI. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

I. INTRODUCTION

(1) Overview: insect herbivores and two-componentplant chemical defence

Insect herbivores account for more than one quarter ofall living species on Earth (Scudder, 2009), and the co-evolution of phytophagous insects and their food plants hascontinued for more than 350 million years (Chaloner et al.,1991; Sinclair & Hughes, 2010). Although insect herbivorespotentially have an abundance of plant species availablefor feeding, herbivory is often restricted by the physical andchemical defence mechanisms plants have evolved to fend offinsect attacks. Whereas mechanical structures like cuticularwaxes, prickles and thorns provide plants with a physicaldefence, toxic chemical compounds provide an additionaleffective defensive barrier (Chen, 2008; Mithofer & Boland,2012). To fend off insect herbivores, more than 200,000specialized metabolites, with toxic, growth-reducing or anti-nutritive effects, are known to be produced by plant species(Zhu-Salzman, Luthe & Felton, 2008; Mithofer & Boland,2012). Chemical defence compounds can be constitutivelypresent in the plant, i.e. pre-exist in anticipation of aninsect attack (phytoanticipins) or their biosynthesis may beinducible (phytoalexins) (VanEtten et al., 1994; Chen, 2008;Mithofer & Boland, 2012).

Constitutive plant defence compounds are often stored inthe form of non-active and non-toxic glucosides in the plantand are spatially separated from bioactivating β-glucosidases.Well-known classes of these compounds are alkaloid,benzoxazinoid, cyanogenic and iridoid glucosides as well asglucosinolates and salicinoids (Halkier & Gershenzon, 2006;Morant et al., 2008; Dobler, Petschenka & Pankoke, 2011;Winde & Wittstock, 2011). Upon insect herbivore attackand tissue damage these glucosylated defence compoundscome into contact with plant β-glucosidases, resulting in therelease of toxic aglucones (Fig. 1A). Such a binary systemof components that are chemically inert when separatedis referred to as two-component plant chemical defence(Wittstock et al., 2004; Bak et al., 2006; Morant et al.,2008). In response, many insect herbivores have evolved

counter-adaptations to overcome this conditional toxicity(Fig. 1B, Table 1). The permanent presence of a specific classof β-glucosidase-activated defence compound in a particularplant species is a predictable characteristic that may havefacilitated herbivorous insects to evolve adaptations. Thisraises the questions: (i) have insect adaptations evolvedfor each specific class of defence compound, or are theregeneral applicable mechanisms that allow insect herbivoresto adapt to all classes of two-component defence, (ii) dogeneralist and specialist herbivores differ in this respect, (iii)do adaptations of insect herbivores mainly target activity ofthe key enzyme, the plant β-glucosidase, to avoid generationof toxic aglucones or are there also other ‘targets’, and (iv) doinsect herbivores combine several adaptations to overcometwo-component plant chemical defence?

Several excellent reviews have described how insectherbivores adapt to toxic chemicals in general, includingbehavioural, physiological and metabolic adaptations tomany different classes of natural plant chemical defencecompounds as well as synthetic insecticides (Brattsten, 1988;Hoy, Head & Hall, 1998; Despres, David & Gallet, 2007;Schowalter, 2011). Here we focus more specifically on two-component chemical defence, as accumulating evidencesuggests that insect herbivores are able to interfere witheither one or both components (Boeckler, Gershenzon &Unsicker, 2011; Dobler et al., 2011; Winde & Wittstock, 2011;Zagrobelny & Møller, 2011). In particular species from theLepidoptera (butterflies and moths), Coleoptera (beetles),Hemiptera (e.g. aphids), Hymenoptera (e.g. sawflies),Orthoptera (e.g. locusts and grasshoppers) and Diptera (trueflies) have evolved a remarkable diversity of adaptations.

(2) Specialist versus generalist insect herbivores

The range of host plant species an insect feeds on defines itsdegree of dietary specialization. When feeding is restrictedto a few, often closely related plant species, herbivores areconsidered specialists (Ali & Agrawal, 2012). A narrow rangeof host plant species is thought to enable high optimalperformance on the host plant and reduce interspecificcompetition, but dietary specialization may also increase


Insect adaptations to plant chemical defence 3

(A) Two-component plant chemical defence and activation by β-glucosidases after herbivory

BGD

DC-glc

Undamaged plant:Spatial separation of both components

in the plant cell or tissue

BGD

2. Non-disruptivefeeding guild

digestive tract

7. Single amino acids

1. Host-plant switching and selection

3. Alkaline gut lumen

4. Reduction of insectβ-glucosidase activity

haemolymph

5. Insect enzyme activity before (a) and after (b)

hydrolysis by plant BGDs

6. Sequestration in special tissues

DCDC-glc

Herbivory causes tissue damage:Mixing of both components

and release of toxic aglucones

Feeding

(B) Adaptations of insect herbivores to overcome two-component plant chemical defence

1. Spilosoma virginica 3. Hyphantria cunea

Digestion

5a. Athalia spp. 6. Chrysomela populi

2. Myzus persicae 4. Epilachna varivestis 5b. Zygaena filipendulae 7. Brahmaea wallichii

Fig. 1. Two-component plant chemical defence, its β-glucosidase-mediated activation into toxic aglucones after herbivory andthe diverse counter-adaptations of insect herbivores. (A) Spatial separation of both components in undamaged plants ensures thatthe defence compound stays glucosylated (DC-glc) and hence non-toxic, since the corresponding plant β-glucosidase (BGD) hasno access to its substrate. After insect herbivory that causes tissue damage, both components come into contact, and the BGDhydrolyses the glucosylated defence compound into a toxic aglucone (DC). (B) Insect herbivores have evolved different adaptationsto counteract and overcome two-component plant chemical defence. These adaptations occur at different stages of herbivoryin a temporal context: before feeding (1), during feeding (2), during digestion (3–7). Examples of insect species that have atleast one of these adaptations are pictured below and numbered according to their adaptation. For more details see Table 1.Photo credits: 1. Whitney Cranshaw, Colorado State University, Bugwood.org (S. virginica); 2. David Cappaert, Michigan StateUniversity, Bugwood.org (M. persicae); 3. Gyorgy Csoka, Hungary Forest Research Institute, Bugwood.org (H. cunea); 4. WhitneyCranshaw, Colorado State University, Bugwood.org (E. varivestis); 5a. Merle Shepard, Gerald R. Carner, and P.A.C Ooi, Insectsand their Natural Enemies Associated with Vegetables and Soybean in Southeast Asia, Bugwood.org (Athalia spp.); 5b. StefanPentzold, University of Copenhagen (Z. filipendulae); 6. Gyorgy Csoka, Hungary Forest Research Institute, Bugwood.org (C. populi);7. Alessandra & Rocco Marciano, Acremar Photos (B. wallichii).

intraspecific competition and reduce the capacity to exploitnew host plants (Schowalter, 2011; Barrett & Heil, 2012).Since specialists encounter a limited number of toxic plantchemicals, specialized enzymes for detoxification may beone strategy. It should be noted that dietary specialization is

considered a continuum and includes intermediates such asoligophagous species that feed on several plant species withinone family (Ali & Agrawal, 2012). Generalists are insects thatfeed on a broad range of plant species, often from morethan one plant family. Greater resource availability and a



higher capacity to exploit new hosts are the advantages, butgeneralists usually have a lower optimal performance on anyparticular plant species than specialists (Mody, Unsicker &Linsenmair, 2007; Barrett & Heil, 2012). The possibility ofhost-plant switching may improve insect development bydiluting the exposure to any single toxic plant chemical andbalancing nutrient intake (Bernays & Minkenberg, 1997;Behmer, 2009). Since generalists encounter a variety of toxicplant chemicals, detoxification enzymes may be less efficient(Barrett & Heil, 2012).

Both generalist and specialist insects are able to suppressthe toxicity of their food plants. Whereas generalists benefitfrom suppressing any level of toxicity from plant defencecompounds, enabling at least short-term feeding, specialistsoften suppress only high levels of toxicity and benefitfrom the presence of low to intermediate levels of plantdefence compounds (Ali & Agrawal, 2012). Sequesteringspecialists, i.e. those that selectively take up and accumulateplant chemicals in their own body, benefit from any levelof defensive chemicals, since the sequestered compoundsprotect the insect against predators (Nishida, 2002; Ali &Agrawal, 2012). Some insect herbivores can also inhibit theproduction of plant chemicals for example by modulating theplant’s salicylic acid and jasmonic acid signalling pathway(Musser et al., 2005; Zarate, Kempema & Walling, 2007;Sarmento et al., 2011). However, for insect herbivoresconfronted with two-component chemical defence, theinhibition of one component, for example the β-glucosidaseor the modification of the free aglucones once released, willbe more effective strategies.

(3) The role of plant β-glucosidases andβ-glucosides in two-component plant chemicaldefence

To be most effective against a wide range of attackingorganisms, defence compounds affect basic metabolicprocesses or evolutionarily conserved cellular mechanisms.Thus, constitutive plant defence compounds may also betoxic to the plant itself. One strategy that limits self-toxicity is to store constitutive defence compounds in aninactive form as β-d-glucopyranosides and to separate themfrom their activating β-glucosidases. Glucosylation of theplant defence compound and its storage separate from β-glucosidases, renders it chemically inert, which preventscellular damage, increases stability and solubility, facilitatesstorage and enables selective intra- and intercellular transportof the compound (Bak et al., 2006; Discher et al., 2009;Nour-Eldin et al., 2012). After tissue damage by herbivory,mixing of glucosylated plant defence compounds and plantβ-glucosidases results in hydrolysis of the defence compoundsand a fast release of toxic aglucones that serve to fend offnon-adapted herbivores (Fig. 1A) (Wittstock & Gershenzon,2002; Morant et al., 2008; Frey et al., 2009; Hopkins, van Dam& van Loon, 2009; Boeckler et al., 2011; Dobler et al., 2011;Pankoke, Bowers & Dobler, 2012). This strategy applies tomany classes of plant defence compounds such as alkaloid,benzoxazinoid, cyanogenic and iridoid glucosides as well as

glucosinolates and salicinoids (Fig. 2). Spatial separation canoccur at different levels, and varies among defence pathwaysor plant species. On a cellular level, β-glucosidases thatactivate cyanogenic glucosides are generally localized in thechloroplast in monocots, or in the apoplast and chloroplastin dicots, while their substrates are stored in the vacuoleor cytosol (Morant et al., 2008; Sanchez-Perez et al., 2009).On a tissue level, activating β-glucosidases may be stored inspecial cell types such as myrosin-cells as in the glucosinolate-myrosinase defence system, while glucosinolates are mainlystored in sulphur-rich S-cells nearby (Koroleva et al., 2000).

β-glucosidases are generally very stable enzymes due totheir compactly folded core structure, and have pH optima inslightly acidic conditions in plants as well as insects (Ruuhola,Julkunen-Tiitto & Vainiotalo, 2003; Byeon et al., 2005;Ketudat, Cairns & Esen, 2010; Pankoke et al., 2012; Terra &Ferreira, 2012). Most plant and insect β-glucosidases belongto the glycoside hydrolase (GH) family 1. GHs comprisemore than 100 described glycosyl hydrolase families, andthese families are mainly classified based on structure andamino acid sequence similarities (Davies & Sinnott, 2008).GH1 β-glucosidases catalyze the hydrolysis of a glucosidicbond between two carbohydrates or between a carbohydratemoiety and an aryl or alkyl aglucone moiety (Cantarelet al., 2009). They are retaining enzymes, i.e. the anomericconfiguration of the glucose is retained during hydrolysis(Vocadlo & Davies, 2008; Davies, Planas & Rovira, 2011).

From an evolutionary point of view, GH1 β-glucosidasesconstitute a monophyletic multigene family. In plants,there are 48 putative β-glucosidase genes in the Arabidopsis

thaliana (Brassicaceae) genome (Xu et al., 2004), and 40in the rice (Oryza sativa, Poaceae) genome (Opassiri et al.,2006). Substrate specificity of plant β-glucosidases maydiffer from a broad range as seen for the Zea mays

(Poaceae) β-glucosidase ZmGlu1 (Brzobohaty et al., 1993)to the narrow specificity reported for the Sorghum bicolor

(Poaceae) β-glucosidase SbDhr1, which hydrolyses onlydhurrin (Verdoucq et al., 2004). Insect genomes also containGH1s, and as in plants their substrate specificity range alsodiffers, but insect genomes contain far fewer β-glucosidases.For example, five putative GH1 β-glucosidases are foundin the transcriptome of the termite Coptotermes formosanus

(Isoptera, Rhinotermitidae) (Zhang et al., 2012). A blastsearch against the fully sequenced insect genomes Anopheles

gambiae, Drosophila melanogaster (both Diptera) and Apis mellifera

(Hymenoptera) showed that their genomes each harbourbetween 5 and 15 β-glucosidases. The differences in numberof GH1 genes between plant and insect genomes may bedue to the fact that plant β-glucosidases are involved inmany physiological processes such as cell-wall lignificationand degradation, phytohormone activation and activationof chemical defence compounds (Morant et al., 2008), whileinsect β-glucosidases mainly have a digestive role (Ferreira,Torres & Terra, 1998; Terra & Ferreira, 2012). The evolutionof insect β-glucosidases might have been driven by the needto hydrolyse nutritive plant β-glucosides encountered duringfeeding. Insect β-glucosidases with broad substrate specificity



Table 1. Overview of the various adaptations of insect herbivores to β-glucosidase-activated plant defence compounds. Entriesare arranged in alphabetical order of defence compound class. Examples where adaptations occur at different levels, including thebehavioural, physiological and metabolic level are indicated by *. For comparison of insect adaptations to further defence compoundclasses see Table 1 in Despres et al. (2007)

Insect herbivorespecies

Dietaryspecialisation

Defencecompound

class Adaptations of insect herbivores References

Callosobruchusmaculatus*

Generalist AG Decreased expression of a gene encoding avicine BGD results in reduced activity ofendogenous BGDs in the midgut

Desroches et al. (1995, 1997)

Lack of endogenous insect BGD activity in thehaemolymph

Sequestration of intact AGsMythimna separata Generalist BX UGT activity in the midgut and excretion of

non-toxic glucosidesSasai et al. (2009)

Ostrinia furnacalis Generalist BX UGT activity in the midgut and excretion ofnon-toxic glucosides

Kojima et al. (2010)

Spodoptera frugiperda*,S. littoralis*

Generalists BX An alkaline midgut reduces plant BGD activityHigh UGT activity in the gutExcretion of BXs

Berenbaum (1980),Dutartre et al. (2011),Glauser et al. (2011) and

CNG S. frugiperda: endogenous gut BGDs lack activitytowards CNGs

Marana et al. (2000)

Agromyzidae species Specialists CNG Leaf-miningSelection of T. ulmifolia populations with low

CNG content

Schappert & Shore (1999)

Cyrtomenus bergi Specialist CNG Piercing-sucking of mature bugs beyond theouter layer of cassava roots lacking CNGs

McMahon et al. (1995)

Diatraea saccharalis Generalist CNG Selective and inducible reduction of the activityof the endogenous insect BGD βGly2

Azevedo et al. (2003) andFerreira et al. (1997)

Epilachna varivestis Specialist CNG Reduction of endogenous insect BGD activityin the gut

Ballhorn et al. (2010)

Heliconius sara* Specialist CNG Metabolic replacement of the nitrile group by athiol

Engler et al. (2000)

SequestrationHyphantria cunea Generalist CNG Highly alkaline midgut Fitzgerald (2008)Schistocerca gregaria* Generalist CNG Leaf-snipping Ballhorn et al. (2010)Zygaena filipendulae* Specialist CNG Detoxification by β-cyanoalanine synthase Zagrobelny & Møller (2011)

SequestrationAthalia rosae* Specialist GSL Sulfatases and sulfotransferases in the gut and

haemolymphOpitz et al. (2011)

SequestrationBrevicoryne brassicae*,

Lipaphis erysimi*Specialists GSL Piercing-sucking

Sequestration of intact GSLsSpatial separation of sequestered GSLs and

endogenous insect BGDs in the body

Barth & Jander (2006),Bridges et al. (2002) andKazana et al.(2007)

Myzus persicae* Generalist GSL Piercing-suckingExcretion of intact GSLsInducible glutathione S-transferase activity

Barth & Jander (2006),Francis et al. (2005) and

IG Piercing-sucking Gange & West (1994)Pieris rapae Specialist GSL Detoxification by nitrile specifier protein and

β-cyanoalanine synthaseWittstock et al. (2004) and

Stauber et al. (2012)Plutella xylostella Specialist GSL Constitutive sulfatase activity in the larval gut Ratzka et al. (2002)S. gregaria* Generalist GSL Host-plant switching

Inducible sulfatases in the gutInducible reduction of endogenous insect BGD

activity in the midgut

Bernays et al. (1994), Falk &Gershenzon (2007) andMainguet et al. (2000)

S. exigua, S. littoralis,T. ni, M. brassicae,H. armigera

Generalists GSL Glutathione S-transferase activity, probably inthe gut

Schramm et al. (2012)

Spilosoma virginica*,Grammia incorrupta*

Generalists IG Inducible reduction of endogenous insect BGDactivity in the midgut

Pankoke et al. (2010, 2012)

Host-plant switching



Table 1. (cont.)

Insect herbivorespecies

Dietaryspecialisation

Defencecompound

class Adaptations of insect herbivores References

Amphipyra monolitha Generalist IG Inducible secretion of high levels of β-alanineinto the midgut

Konno et al. (2010)

Artopoetes pryeri Specialist IG Secretion of high levels of GABA into themidgut

Konno et al. (2010)

Brahmaea wallichii,Dolbina tancrei

Specialists IG Secretion of high levels of free glycine into theanterior part of the midgut

Konno et al. (1997)

Chrysomela populi *,Phratora vitellinae*

Specialists SA Lack of endogenous insect BGD activity in thehaemolymph

Discher et al. (2009) andKuhn et al. (2004, 2007)

Specific transportersSequestration of intact SAs

Lymantria dispar*,Malacosoma disstria*

Generalists SA Inducible reduction of endogenous insectBGD activity in the midgut

Hemming & Lindroth(2000)

Inducible glutathione-S-transferase in themidgut

Operophtera brumata Generalist SA Highly alkaline gut Ruuhola et al. (2003)Papilio glaucus

canadensisSpecialist SA Inducible reduction of endogenous insect

BGD activity in the midgutLindroth (1988)

Phyllonoryctersalicifoliella

Specialist SA Leaf-miningSelective feeding on poplar species with low

levels of SAs

Auerbach & Alberts (1992)

AG, alkaloid glucoside; BGD, β-glucosidase; BX, benzoxazinoid glucoside; CNG, cyanogenic glucoside; GABA, γ -aminobutyric acid;GSL, glucosinolate; IG, iridoid glucoside; SA, salicinoid; UGT, UDP-glucosyltransferase.

may also activate glucosylated plant defence compoundsduring digestion. However, this process is likely to be lessefficient in deterring insect feeding than the immediaterelease of high amounts of toxic aglucones by specific andhighly active plant β-glucosidases upon tissue disruptionby herbivores. Moreover, insects may also reduce theirendogenous β-glucosidase activity to limit the release oftoxic aglucones during digestion (see Section III.3b).

II. HOW NON-ADAPTED INSECT HERBIVORESARE AFFECTED BY TWO-COMPONENT PLANTCHEMICAL DEFENCE

The chemical compounds from the various classes of two-component defence differ in their chemical structure (Fig. 2),effect on non-adapted insect herbivores and distributionwithin land plant species. Whereas some classes are limitedto specific orders and families, others are found in mostgroups of land plants (Bak et al., 2006; Halkier & Gershenzon,2006). Many of the aglucones inhibit enzymes and proteinsunspecifically, but others may be targeted to specific enzymes.In the following sections, compounds from several classes oftwo-component plant chemical defence are described as wellas their effects on non-adapted insect herbivores.

(1) Alkaloid glucosides

Alkaloid glucosides are characterised by nitrogen-containingheterocyclic aglucones, such as pyrimidine for example. In

the case of vicine and convicine, the main alkaloid glucosidedefence compounds in the cotyledons of the broad bean Vicia

faba (Fabaceae), pyrimidine is linked to a β-d-glucopyranose(Desroches et al., 1995, 1997). When bruchid beetle larvaeCallosobruchus maculatus (Coleoptera, Chrysomelidae) feed onV. faba, the ingested vicine and convicine are activatedby endogenous C. maculatus β-glucosidases into the highlyreactive and free-radical-generating aglucones divicine orisouramil (Fig. 2A) (Hegazy & Marquardt, 1984; Desrocheset al., 1997; Brimer, 2011). These aglucones inhibit theactivity of glucose-6-phosphate dehydrogenases causingadverse metabolic effects such as lipid peroxidation, whichresults in high mortality rates of C. maculatus larvae feeding onV. faba (Desroches et al., 1995; McMillan, Bolchoz & Jollow,2001).

(2) Benzoxazinoid glucosides

Benzoxazinoid glucosides are found in numerous species ofthe monocot family Poaceae, including major crops such asmaize Z. mays, wheat Triticum aestivum and rye Secale cereale

(Frey et al., 2009). In addition, a few species of the eudicotorders Ranunculales and Lamiales produce benzoxazinoidglucosides following convergent evolution of both thebiosynthetic pathway and the activating β-glucosidase (Dicket al., 2012). The biosynthetic pathway of benzoxazinoidglucosides in maize is fully characterized. Indole-3-glycerolphosphate is converted mainly by cytochrome P450-dependent monooxygenases into cyclic hydroxamic acid,which is glucosylated with a β-d-glucopyranose before



GlcS

R

N O

SO3-

unstableaglucone

R N C S

Isothiocyanate

R C N

Nitrile

(D) Glucosinolates

-S

R

N O

SO3-

NSP

Sulfatase

GlcS

R

N OH

desulfo-glucosinolate

Glc O

CN

(C) Cyanogenic glucosides

HO

CN

unstableaglucone

(F) Salicinoids

O

GlcO Esterase

or alkaline pH

OH

GlcO

SalicinSalicortin Saligenin

GSTGSH-ITC

(E) Iridoid glucosides

O

OH

HOGlcO

Aucubin

O

OH

HOOH

reactive aglucone

b-CASβ-cyanoalanine

(B) Benzoxazinoid glucosides

N

O

O

OH

OOGlc

DIMBOA-glc

N

O

O

OH

OHO

UGT

BGD

DIMBOA

Linamarin

R

unknown

substitution by -SH group

excretionR = aliphatic

R = aromatic

HCN

Glc = β-D-glucopyranose(A) Alkaloid glucosides

BGD*N

NH

Vicine Divicine

OGlc

O NH2

NH2

N

NH

HO

O NH2

NH2

O

H

HO

H

HO

H

HOHH

OH

OH

BGD

BGD*

BGD

BGD*

BGD

BGD*

BGD

BGD*

OH

HO

Sulfotransferase

desulfo-glucosinolate-3-sulfate

HCN

HCN

HCN

b-CASβ-cyanoalanine

Insect enzymesPlant enzymes

Fig. 2. Chemical structure of different classes of glucosylated plant defence compounds and their activation by β-glucosidases(BGDs) into toxic aglucones and further products. Activation can be due to activity of BGDs from the plant (green) or due toinsect BGDs (orange). Importantly, adapted insects reduce the activity of their endogenous BGDs to reduce the formation ofaglucones (marked with *). Activity of further more specialized enzymes from insects convert plant defence compounds: eitherbefore their hydrolysis by plant BGDs, so that the product can no longer be activated by plant BGDs into toxic aglucones,or after their hydrolysis by plant BGDs via conjugating or detoxifying insect enzymes. Toxicity of a compound is indicated bya skull. Abbreviations: b-CAS, β-cyanoalanine synthase; DIMBOA, 2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-one; GSH-ITC,glutathione-isothiocyanate; GST, glutathione-S-transferase; NSP, nitrile specifier protein; UGT, UDP-glucosyltransferase.



final addition of a methoxy group into 2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-glucoside (DIMBOA-glucoside),a major benzoxazinoid glucoside in Poaceae (Fig. 2B)(Jonczyk et al., 2008; Frey et al., 2009; Dick et al., 2012;Dutartre, Hilliou & Feyereisen, 2012). Once activated byplant β-glucosidases, the toxic aglucone DIMBOA depletescellular glutathione levels and this leads to irreversibleinactivation of enzymes with cysteine residues in their activesite (Dixon et al., 2012). For example, when DIMBOA isadded to artificial diets, survival, mass gain and reproductionof several aphid species is strongly reduced (Escobar,Sicker & Niemeyer, 1999). Similarly, when larvae of thesilkworm Bombyx mori (Lepidoptera, Bombycidae) werereared on a DIMBOA-containing diet, the larvae died within3 days (Sasai et al., 2009). Another benzoxazinoid agluconefrom maize, 2-hydroxy-4, 7-dimethoxy-1, 4-benzoxazin-3-one (HDMBOA), was shown strongly to deter generalistherbivores such as the Egyptian armyworm Spodoptera littoralis

and the fall armyworm Spodoptera frugiperda (both Lepidoptera,Noctuidae) (Glauser et al., 2011).

(3) Cyanogenic glucosides

Cyanogenic glucosides are synthesized from amino acidsand consist of an α-hydroxy nitrile linked to a β-d-glucopyranose (Bak et al., 2006; Møller, 2010). They arepresent in more than 2650 plant species distributed among130 families within ferns, gymnosperms and angiosperms,and have independently evolved in different plant lineagesby recruiting biosynthetic genes from similar gene families(Takos et al., 2011). Despite their wide distribution, onlyabout 50 different structures of cyanogenic glucosidesare known (Bak et al., 2006; Bjarnholt et al., 2008).Cyanogenic glucosides are also found within a few arthropodspecies (Zagrobelny, Bak & Møller, 2008). For example,Zygaena filipendulae six-spot burnet moth larvae (Lepidoptera,Zygaenidae) are able to carry out both de novo biosynthesisand sequestration of cyanogenic glucosides for use in defence,and also as a nuptial gift during mating (Zagrobelny et al.,2007). The biosynthetic pathways have been elucidatedin both plants and insects, and this has revealed thatthe ability to synthesize cyanogenic glucosides has evolvedconvergently in the two kingdoms (Jensen et al., 2011). Duringherbivory, the plant tissue is damaged, and the cyanogenicglucosides come into contact with plant β-glucosidases.This releases an unstable aglucone that dissociates eitherspontaneously or enzymatically, liberating highly toxichydrogen cyanide in a process called cyanogenesis (Fig.2C) (Zagrobelny et al., 2008). Hydrogen cyanide specificallyinhibits cytochrome c oxidase, a key enzyme in themitochondrial respiratory pathway (Blom et al., 2011), whichcauses cell and tissue death within a short time, and stronglydeters various non-adapted herbivores (Møller, 2010). Forexample, fall armyworm larvae quickly die when fed onan artificial diet containing cyanide (Hay-Roe, Meagher &Nagoshi, 2011).

(4) Glucosinolates

Similar to cyanogenic glucosides, glucosinolates aresynthesized from amino acids, but consist of a β-d-thioglucopyranoside, a sulfonated oxime and a variableside chain (Halkier & Gershenzon, 2006; Hopkins et al.,2009). More than 140 different structures of glucosinolatesare known, but they are almost exclusively restricted to theorder Brassicales which includes many crop species such asoilseed rape (Brassica napus), broccoli (B. oleraceae), mustard(Sinapsis alba) and the model plant A. thaliana (Halkier &Gershenzon, 2006). Activation of glucosinolates is carried outby β-thioglucosidases, so-called myrosinases. Myrosinasesare closely related to β-glucosidases and are known fromplants and insects (Husebye et al., 2005). Myrosinaseactivity releases an unstable aglucone, which dissociatesmainly into toxic isothiocyanates, but also into nitrilesand related compounds (Fig. 2D) (Halkier & Gershenzon,2006). The aglucone produced depends on the presenceof epithiospecifier proteins, the structure of the side chaingroup and factors such as pH and the presence of ferrousions (Halkier & Gershenzon, 2006). Isothiocyanates forexample react with amino and sulfhydryl groups of proteins(Halkier & Gershenzon, 2006), which makes them toxic toe.g. the springtail Folsomia fimetaria (Collembola, Isotomidae)even at low concentrations (Jensen et al., 2010). Similarly,generalist herbivores like the cabbage looper Trichoplusia ni(Lepidoptera, Noctuidae) or the tobacco hornworm Manducasexta (Lepidoptera, Sphingidae) significantly decrease in bodymass when feeding on A. thaliana containing glucosinolatesand myrosinases, in comparison to feeding on A. thalianaknock-out mutants that lack myrosinases (Barth & Jander,2006). Adverse effects of the glucosinolate-myrosinasetwo-component defence are also known for the beetarmyworm Spodoptera exigua (Lepidoptera, Noctuidae) (Mulleret al., 2010).

(5) Iridoid glucosides

Iridoid glucosides are widespread plant defence compoundsthat can be found in more than 50 different plant familiesof the Asteridae (Dobler et al., 2011). They are derived frommonoterpenes and form a large group of several hundreddifferent compounds (Fig. 2E) (Dobler et al., 2011). All iridoidglucosides have a similar skeleton with a cyclopentane ringconnected to an oxygenated heterocyclohexane, where a β-d-glucopyranose is attached at the C1 atom. Various epoxygroups and sugars determine their structural diversity. Ininsects, iridoid glucosides are mainly activated in the gut,either by co-ingested plant β-glucosidases or by endogenousinsect β-glucosidases (Pankoke et al., 2012). The releasedaglucones are highly unstable and reactive: the pyran ringof the aglucone opens and forms irreversible bonds withproteins, which causes unspecific crosslinks in proteins,inhibits enzymes and deters non-adapted insects (Kim et al.,2000; Dobler et al., 2011). The aglucone of the seco-iridoid glucoside oleuropein from the privet tree (Ligustrumobtusifolium, Oleaceae) covalently binds to lysine residues in



proteins due to its electrophilic glutaraldehyde-like structure(Konno, Hirayama & Shinbo, 1997; Konno et al., 1999).This leads to denaturing and crosslinking of proteins, whichdecreases the nutritive value of dietary proteins and detersinsect herbivores.

(6) Salicinoids

Salicinoids are phenolic glucosides consisting of a salicylalcohol linked to a β-d-glucopyranose (Boeckler et al., 2011).Salicin, the simplest phenolic glucoside, can be modified withdifferent organic acids resulting in at least 20 more complexstructures (Fig. 2F) (Boeckler et al., 2011). Salicinoids areparticularly abundant in species of the Salicaceae (e.g. poplarsPopulus spp. or willows Salix spp.) where they can reachconcentrations of up to 30% of plant dry mass (Donaldsonet al., 2006), but are also known from the pine family(Pinaceae) and oaks (Quercus spp., Fagaceae) (Delvas et al.,2011). As a two-component defence, toxicity of salicinoidsrequires activation by β-glucosidases to form aglucones,which may be oxidized in the gut lumen of non-adaptedherbivores into tissue-damaging and protein-crosslinkingreactive oxygen species such as quinones (Ruuhola et al.,2003; Boeckler et al., 2011; Delvas et al., 2011). For example,in white spruce needles (Picea glauca, Pinaceae) the salicinoidspicein and pungenin are activated by β-glucosidases intopiceol and pungenol (MacKay, 2012). These aglucones harmthe spruce budworm Choristoneura fumiferana (Lepidoptera,Tortricidae) by increasing its larval mortality, retarding itsdevelopment and reducing its pupal mass (Delvas et al.,2011). Needles that are susceptible to budworms do notexpress β-glucosidase genes (MacKay, 2012). Thus, non-resistant needles contain mainly intact salicinoids and lackthe ability to release toxic aglucones (Delvas et al., 2011).

III. FROM FEEDING TO DIGESTION: TARGETSFOR INSECT HERBIVORE ADAPTATIONS TOTWO-COMPONENT PLANT CHEMCIALDEFENCE IN A TEMPORAL CONTEXT

As discussed above, non-adapted insects will be stronglyaffected in different ways by toxic aglucones released fromglucosylated defence compounds by plant β-glucosidaseactivity. However, other insect herbivores, includinggeneralists and specialists, have evolved a range of diverseadaptations to overcome the deleterious effects of two-component plant chemical defence, and this enables them tofeed on these plants. These adaptive processes occur beforeand during feeding as well as mainly during digestion ofthe plant material. Behavioural, physiological and metabolicadaptations may be combined, often target the plant β-glucosidase activity and apply to all classes of two-componentplant chemical defence discussed here (Table 1, Figs 1 and2). Finally, we discuss that adaptations in generalists areoften inducible, whereas those in specialists seem to beconstitutive.

(1) Before feeding: recognition, switching andselection of host plants

Insect herbivores encounter both nutrients and plant defencecompounds in their food, often resulting in a trade-off between nutrient intake and ingestion of harmfulsubstances (Singer, Bernays & Carriere, 2002; Behmer,2009). Importantly, plant defence compounds often havea pronounced spatial and temporal variation at all levels,i.e. varying concentrations in different organs and withinand among populations as well as throughout developmentalstages (Hoy et al., 1998; Gebrehiwot & Beuselinck, 2001).This heterogeneity in the plant puts a selection pressure onherbivores to evolve sensory systems for the recognition andavoidance of plant defence compounds to avoid ingestionof lethal doses. Underlying mechanisms are geneticallydetermined and can be inherited, but may also be learned(Chapman, 2003; Despres et al., 2007; Schowalter, 2011).Recognition of potentially toxic substances allows insectherbivores to generate a beneficial behavioural response.Whereas non-adapted herbivores benefit from avoiding theingestion of toxins by feeding on toxin-free organs or atdevelopmental stages where toxins are absent (Hoy et al.,1998; Despres et al., 2007), adapted herbivores may responddifferently (Hoy et al., 1998).

Insect herbivores that are adapted to low to medium levelsof defence compounds can regularly switch to other hostplants to avoid ingestion of lethal doses by dietary mixing -a feeding behaviour mainly exhibited by generalists. Host-plant switching may dilute excessive detrimental effects ofany single plant defence compound. For example, larvaeof the lepidopteran generalists yellow woolly bear Spilosomavirginica and tiger moth Grammia incorrupta (both Lepidoptera,Arctiidae) can feed on high iridoid-glucoside-containingplants like Plantago lanceolata (Plantaginaceae) (Pankoke,Bowers & Dobler, 2010; Pankoke et al., 2012). Both specieswere shown to compensate fitness costs from feeding ontoxic P. lanceolata by regular host-plant switching underfield conditions (Singer et al., 2002). Several experimentswith generalist grasshopper species have shown that host-plant switching and dietary mixing results in better growth,survival and fecundity than when feeding is limited to a singleplant species (summarized in Bernays & Minkenberg, 1997).Grasshoppers also seem behaviourally to select for dietarymixing and nutritional balance if given the opportunity(Bernays & Minkenberg, 1997). For example, the growth rateof the desert locust Schistocerca gregaria (Orthoptera, Acrididae)was significantly higher on a mixed diet containing kale (B.oleraceae, containing glucosinolates), cotton (Gossypium hirsutum,Malvaceae) and basil (Ocimum basilicum, Lamiaceae) than onany one of the single plant species (Bernays et al., 1994).

Specialists that are adapted to high levels of a certainplant defence compound and even require them for theirdevelopment, benefit from identifying plants with highconcentrations. In feeding-choice experiments, larvae ofthe six-spot burnet moth were shown to identify andto prefer Lotus corniculatus (Fabaceae) food plants withhigh contents of cyanogenic glucosides over plants with



low contents (Zagrobelny et al., 2007). Larvae reared onplants with low cyanogenic glucoside content showed adecelerated development since they need to expend energyin synthesizing cyanogenic glucosides. By contrast, larvaereared on plants with high levels of cyanogenic glucosideshad normal development since they are able to accumulatecyanogenic glucosides in their body by sequestration, whichis less costly than biosynthesis (Zagrobelny et al., 2007).

Behavioural adaptation through host-plant switching ingeneralists and selection of toxic plants in specialists alsoinvolves trade-offs and fitness costs. Insect herbivores needto invest time and energy to search for a suitable host(Despres et al., 2007). Investment costs differ in generalistsand specialists and mainly seem to depend on the level ofglucosylated plant defence compound, but also to a highdegree on the level of activating plant β-glucosidases. Themore of both components are present in the plant, the moreoften generalists need to switch host plants, and the longerthey need to search for suitable host plants, which increasescosts. By contrast, specialists need to invest less time, energyand thus costs in this case.

(2) During feeding: impact of the feeding guild

Herbivory inevitably results in tissue damage. The more planttissue is damaged, the more toxic aglucones are releaseddue to mixing of plant β-glucosidases and glucosylatedplant defence compound (Bernays & Janzen, 1988; Barth &Jander, 2006). The extent of damage, however, differs amongherbivore species and mainly depends on their feeding guild(Simberloff & Dayan, 1991; Gleadow & Woodrow, 2002;Textor & Gershenzon, 2009). A feeding guild is a groupof species that use the same environmental resources ina similar way, i.e. the technique insects use to feed onplants. In general there are different guilds of feeding suchas piercing-sucking, leaf-snipping, leaf-chewing and leaf-mining (Bernays & Janzen, 1988; Sinclair & Hughes, 2010;Ali & Agrawal, 2012). Importantly, these insect feedingguilds impact on the effectiveness of two-component plantchemical defence, because the amount of tissue damage andhence the release of aglucones will vary depending on thefeeding strategy. Whereas the non-disruptive feeding guildsof piercing-sucking and leaf-snipping efficiently preventtissue damage (Fig. 3A, B), leaf-mining is potentially moredisruptive (Fig. 3C) and leaf-chewing extensively damagesplant tissue (Fig. 3D).

(a) Piercing-sucking

Insects belonging to the order Hemiptera such asaphids (Aphidae), whiteflies (Aleyrodidae) and leafhoppers(Cicadellidae) use their stylets, highly modified mouthparts,to pierce the cuticle, epidermis and mesophyll and suck fromsingle phloem sieve elements, a rich source of high levelsof sucrose and other nutrients (Douglas, 2006; Kehr, 2006;Walling, 2008). Aphids inject watery saliva containing Ca2+-binding proteins that counteract the influx of Ca2+ andprevent closure of sieve elements by callose formation (Miles,

1999; Walling, 2008). Piercing-sucking minimizes tissuedisruption, and thereby minimizes the plant β-glucosidase-mediated activation of glucosylated defence compounds(Fig. 3A).

In the case of the glucosinolate-myrosinase defence systemin Arabidopsis plants, piercing-sucking aphids were shown toprevent toxic hydrolysis of glucosinolates. This allows thespecialist cabbage aphid Brevicoryne brassicae (Aphididae) toaccumulate intact glucosinolates in its body, often for use inits own defence (Bridges et al., 2002). Likewise, the generalistgreen peach aphid Myzus persicae (Aphididae) is able to ingestintact glucosinolates, and excretes them in their non-toxicform via the honeydew (Barth & Jander, 2006). Green peachaphids are also able to feed and develop on P. lanceolata plantswith a high content of iridoid glucosides (Gange & West,1994). Moreover, different aphid species are the dominantinsect taxa found in plant populations of Turnera ulmifolia(Turneraceae) that exhibit high concentrations of cyanogenicglucosides of up to 640 μg HCN per g dry mass (Schappert& Shore, 1999). This shows that the non-disruptive feedingguild of piercing-sucking efficiently minimizes mixing ofplant β-glucosidase and glucosylated defence compoundand hence the formation of toxic aglucones among differentclasses of two-component defences.

Even the length of the stylus of piercing-sucking insectsseems important in the avoidance of activation of two-component plant chemical defence. For example, immaturecassava bugs Cyrtomenus bergi (Hemiptera, Cydnidae) have ashort stylus and are deterred from feeding on the outer layerof cyanogenic-glucoside-containing roots of cassava Manihotesculenta (Euphorbiaceae) (McMahon, White & Sayre, 1995).By contrast, fully mature cassava bugs have a longer styletand are able to feed on cassava roots by penetrating beyondthe outer layer.

(b) Leaf-snipping versus leaf-chewing

Insect herbivores may snip leaves into quite large piecesas shown for some lepidopteran caterpillars biting off leafdiscs up to 0.6 mm2 (Bernays & Janzen, 1988). Such leaf-snipping reduces potential tissue damage and the formationof toxic aglucones markedly (Bernays & Janzen, 1988;Barbehenn, 1992). Lepidopteran species that have shortand simple mandibles and lack molars, and locusts that haverelatively large mandibles can all exhibit leaf-snipping (Fig.3B) (Bernays & Janzen, 1988; Barbehenn, 1992; Ballhorn,Kautz & Lieberei, 2010).

The specialist Mexican bean beetle Epilachna varivestis(Coleoptera, Coccinellidae) and the generalist desert locustboth feed on cyanogenic-glucoside-containing lima beans(Phaseolus lunatus, Fabaceae). Although the Mexican beanbeetles have a reduced activity of endogenous β-glucosidasesin comparison to locusts (see Section III.3b), more cyanogenicglucosides are hydrolysed during feeding by the beetles thanby the locusts (Ballhorn et al., 2010). This difference is due totheir different feeding guilds: beetles with their rather smallmandibles chew leaves and tend to crush the plant tissue(leaf-chewing). By contrast, locusts with their quite large



DC

BGD

(A)

(B)

Piercing-sucking

BGD

Leaf-snipping

Leaf-chewing

(D)

Tissue damage, aglucone formation

and toxicity

DC-glc

DC-glc

Inhibition of aglucone formation

and toxicity

DC

Insect feeding guild

Plant tissue level

Plant cell level

Molecularlevel

BGD

DC-glc

No aglucones

Completemixing

Highagluconeformation

Minimalagluconeformation

Lowmixing

No mixing 100 %

0 %100 %

0 %

(C) Leaf-mining

DC

Intermediateagluconeformation

BGD

Mediummixing

DC-glc

Fig. 3. Insect feeding guilds and their impact on the activation of glucosylated plant defence compounds by β-glucosidases. Feedingguild, potential damage to the plant on the tissue level, potential enzyme and substrate mixing upon tissue damage on the cellularlevel, and formation of toxic aglucones on the molecular level are shown. Feeding by (A) piercing-sucking insects like aphids and (B)leaf-snipping caterpillars or locusts causes minimal tissue damage, resulting in low mixing of glucosylated plant defence compound(DC-glc) and β-glucosidases (BGDs) in the cell, and thus minimal formation of toxic aglucones (DC). Leaf-mining (C) by dipteranlarvae, for example, results in medium levels of tissue damage and intermediate levels of aglucone formation, whereas leaf-chewingby beetles (D) extensively damages plant tissue. This results in complete mixing of glucosylated plant defence compound andβ-glucosidase and consequently high levels of aglucones released. Drawings of insects with permission and copyright by D. G.Mackean.

mandibles snip leaves into larger pieces (leaf-snipping), andconsequently ingest a high percentage of intact plant tissue,thereby limiting hydrolysis of cyanogenic glucosides (Fig. 3B)(Ballhorn et al., 2010). During digestion, plant tissue and cellsmay be disrupted in the gut, but physiological conditionsmight favour stabilization or detoxification of plant defencecompounds as discussed later.

Bernays & Janzen (1988) compared mandible morphologyof caterpillars between two lepidopteran families andcorrelated morphological differences to their different feedingguilds. Mandibles of Saturniidae, a leaf-snipping family, wereshort and simple. This morphology minimizes tissue damage,enables ingestion of large plant pieces and minimizes theamount of activated plant defence compound. By contrast,the mandibles of Sphingidae, a leaf-chewing family, werelong, toothed and ridged. This morphology enables crushingof plant material, ingestion of small pieces and absorptionof maximal amounts of nutrients (Bernays & Janzen, 1988).Thus, leaf-snipping enables insect herbivores to feed onpotentially toxic plants, but may be less efficient in termsof nutrient extraction. Conversely, if more plant materialis crushed, more nutrients become available, but also moreplant defence compounds are activated, which potentiallylimits leaf-chewers’ feeding on toxic plants. It should be

noted that leaf-snipping saturniid caterpillars are mainlygeneralists that feed on a variety of plants, including speciesnot protected by two-component chemical defence (Janzen,1984). The evolution of leaf-snipping and piercing-suckingmorphologies and feeding guilds thus is probably not aspecific adaptation to two-component defence; insects witha minimally disruptive feeding morphology may simplyhave been pre-adapted to feed on plants protected by two-component chemical defence.

(c) Leaf-mining

Leaf-mining insects feed inside the leaf lamina, mostly onparenchymous or epidermal tissues, causing channels, minesor blotches (Sinclair & Hughes, 2010). These herbivores areprotected from external factors such as ultraviolet radiationor externally applied insecticides, although they are oftenhighly susceptible to parasitoids (Connor & Taverner, 1997;Loch, Matthiessen & Floyd, 2004). Most leaf-mining speciesare found within the Coleoptera, Diptera, Hymenopteraand Lepidoptera (Connor & Taverner, 1997). Because theirfeeding is restricted to specific leaf tissues, the total tissuedamage by leaf-mining is lower than the damage resultingfrom leaf-chewing, but potentially higher than from piercing-sucking or leaf-snipping (Fig. 3C) (Schappert & Shore,



1999). Several studies indicate that intermediate amounts ofglucosylated defence compounds are activated during leaf-mining and that, according to the selective feeding hypothesis(Scheirs, De Bruyn & Verhagen, 2001), it is adaptive forleaf-miners behaviourally to select and feed on plants ortissues with low levels of chemical defence.

Dipteran (Agromyzidae) and lepidopteran (Gelechiidae)leaf-mining species were only found in T. ulmifolia populationswith low concentrations of cyanogenic glucosides and wereabsent from populations with high concentrations (Schappert& Shore, 1999). The aspen blotch miner Phyllonoryctersalicifoliella (Lepidoptera, Gracillariidae) avoids exposure tohigh levels of salicinoids by selectively feeding on poplarspecies with low levels of salicinoids such as Populus tremuloides(Auerbach & Alberts, 1992). Recent studies show that theextent of damage by aspen blotch miners is indeed negativelycorrelated to the total salicinoid concentration of P. tremuloidesleaves (Young et al., 2010a, b). Leaf-mining Scaptomyza flava(Diptera, Drosophilidae) larvae exhibit significantly reducedgrowth rates on glucosinolate-containing wild-type A. thalianarelative to larvae reared on plants with low levels ofglucosinolates (Whiteman et al., 2012). The two dipterangrass miners Chromatomyia milii and C. nigra prefer to feed onthe leaf mesophyll tissue and never feed on the epidermaltissue of Holcus lunatus (Poaceae) (Scheirs et al., 2001). Inanother grass species such as barley Hordeum vulgare (Poaceae)the cyanogenic glucoside epiheterodendrin accumulates inthe epidermal cells (Nielsen et al., 2002; Li et al., 2011). Itcan be imagined that leaf-mining insects that do not feed onepidermal cells would avoid exposure to such epidermallylocalized chemical defence compounds.

(3) During digestion: physiological and metabolicadaptations to counteract two-component defence

(a) An alkaline gut pH inhibits the plant β-glucosidase

Differences in pH may affect enzymatic reactions bychanging the charge of certain amino acids, affecting proteinconformation and their catalysis (Johnson & Felton, 1996;Harrison, 2001). Whereas the haemolymph of insects ismostly neutral with pH ranging from 6.4 to 7.5, thepH of the midgut lumen varies from a strongly acidicpH 3.1 to extremely alkaline pH 12.4 among differentinsect orders (Berenbaum, 1980; Dow, 1984; Schultz &Lechowicz, 1986; Johnson & Felton, 1996; Appel & Joern,1998; Harrison, 2001; Cristofoletti et al., 2003; Fitzgerald,2008; Terra & Ferreira, 2012). Regulation of midgut pHmainly involves H+ V-ATPases that are located in theapical membrane of goblet cells in the midgut (Wieczoreket al., 2003). Many larvae of Lepidoptera, Diptera andscarab beetles (Coleoptera) have highly alkaline midguts,whereas larval Orthoptera, Hemiptera and most coleopteranfamilies usually have slightly acidic to neutral midguts.Consequently, lepidopteran digestive enzymes like amylasesare evolutionarily adapted to function in an alkaline gut(Pytelkova et al., 2009). However, unfavourable highlyalkaline pH conditions in the midgut lumen may inhibit plant

β-glucosidases and prevent activation of ingested defencecompounds.

Although caterpillars of the generalist fall webwormHyphantria cunea (Lepidoptera, Arctiidae) are vulnerable tohydrogen cyanide, they can feed on leaves of black cherry(Prunus serotina, Rosaceae) containing cyanogenic glucosideswithout any adverse effects (Fitzgerald, 2008). A highalkaline midgut lumen at pH 11 was shown to preventthe hydrolysis of cyanogenic glucosides into toxic hydrogencyanide during passage of the plant parts through the gut(Fitzgerald, 2008), most likely due to inhibition of plant β-glucosidases. A direct link between an alkaline midgut andreduced plant β-glucosidase activity towards benzoxazinoidglucosides was shown in the generalist fall armyworm feedingon maize containing DIMBOA-glucosides (Dutartre et al.,2011). The larval midgut lumen with a pH of 10 wasshown to reduce plant β-glucosidase activity by more than80%, which strongly reduced the release of toxic aglucones.Caterpillars of the generalist winter moth Operophtera brumata

(Lepidoptera, Geometridae) feed on Salix species that containthe salicinoid salicortin. In the midgut lumen, plant esteraseactivity or an alkaline pH of 9.5 convert salicortin intosalicin (Fig. 2F) (Berenbaum, 1980; Ruuhola, Tikkanen &Tahvanainen, 2001). However, hydrolysis of salicin by plantβ-glucosidases to toxic saligenin is markedly reduced at analkaline pH, as Salix β-glucosidases function optimally ataround pH 5 (Ruuhola et al., 2003). The alkaline midgutprevents formation of toxic saligenin and enables larvae toingest salicortin and to excrete non-toxic salicin in their frass(Fig. 2F) (Ruuhola et al., 2001).

It is important to note that numerous insect herbivoreswith an alkaline midgut are known to feed on plants thatare not protected by two-component chemical defences, butby diverse other toxic plant chemicals (Berenbaum, 1980).Thus, a highly alkaline midgut probably did not arise asan evolutionary response to two-component plant chemicaldefences. Insect herbivores with an alkaline midgut simplymay have been pre-adapted to feed on plants protected bytwo-component chemical defences.

In a classical study that employed tannins, polyphenolglucosides that do not require β-glucosidases to exerttoxicity, it was shown that generalist caterpillars of thegypsy moth Lymantria dispar (Lepidoptera, Erebidae) adjustedthe pH of their midgut according to the amount of ingestedtannin from slightly to highly alkaline within a few hours(Schultz & Lechowicz, 1986). This alkalinization leads to re-dissociation of harmful tannin–protein complexes and makestannins non-toxic (Schultz & Lechowicz, 1986). It would beinteresting to know if such an inducible alkalinization of themidgut is also found in insects that feed on plants defendedby two-component chemical defences or if these herbivoreshave a constitutive alkaline gut.

Evolutionary costs associated with having an alkaline gutmay be related to entomopathogenic bacteria such as Bacillus

thuringiensis that produce insecticidal δ-endotoxins forminglytic pores in the cell membrane of the midgut epithelium (deMaagd, Bravo & Crickmore, 2001; Bravo, Gill & Soberon,



2007). Solubilization and activation of δ-endotoxin protoxinsrequires mainly alkaline conditions in addition to specificproteolytic enzymes (de Maagd et al., 2001; Bravo et al., 2007).Consequently, lepidopteran and dipteran larvae with theiralkaline midguts are often more vulnerable to B. thuringiensisδ-endotoxins than coleopteran larvae with a neutral gutenvironment that keeps protoxins insoluble and preventstheir activation (Lambert et al., 1992; Bradley et al., 1995; deMaagd et al., 2001).

(b) Reduction of endogenous insect β-glucosidase activity in the gut

Since endogenous insect β-glucosidases are mainly digestiveenzymes, they may also activate two-component plantchemical defence after ingestion of plant material, whichrenders them a target for insect adaptation. Indeed, severalspecialists and generalists were shown to reduce activityof their endogenous β-glucosidases, efficiently reducingformation of toxic aglucones during digestion.

Bruchid beetles C. maculatus are generalists that develop onseeds of the cow pea Vigna unguiculata (Fabaceae) (Desrocheset al., 1997). Seeds of the broad pea Vicia faba, however, exertstrong toxicity to C. maculatus larvae caused by activationof the alkaloid glucoside vicine by larval endogenous β-glucosidases (Fig. 2A) (Desroches et al., 1995). Interestingly,some larvae of C. maculatus are able to develop on vicine-containing V. faba seeds (Desroches et al., 1995). Thistolerance is due to decreased expression of a gene encodinga β-glucosidase that is active towards vicine (Desrocheset al., 1997). The decreased expression of this β-glucosidaseresults in lowered β-glucosidase activity in the midgut of thelarvae and strongly reduces formation of toxic aglucones,even when feeding on seeds containing vicine at 1% drymass. Only concentrations higher than 1.5% vicine in theexperimental diet caused lethal effects on feeding larvae, butsuch concentrations are not found naturally in V. faba seeds(Desroches et al., 1997).

The Mexican bean beetle is a specialist that feeds onlima beans containing cyanogenic glucosides without anyadverse effects (Ballhorn et al., 2010). It was found thatbean beetles strongly decrease the activity of their ownendogenous β-glucosidases in the gut relative to gut β-glucosidase activity from desert locusts feeding on the sameplant species (Ballhorn et al., 2010). After ingestion thecyanogenic glucosides are not hydrolysed in the bean beetleand can be excreted intact with the frass.

The generalist caterpillars yellow woolly bear and tigermoth feed on iridoid-glucoside-containing plants such as P.lanceolata. Both species reduce the activity of endogenousβ-glucosidases in their midgut in response to increasinglevels of the iridoid glucoside aucubin (Fig. 2E) (Pankokeet al., 2010, 2012). A decrease of endogenous midgut β-glucosidase activity enables both herbivore species to feedon toxic aucubin-containing plants at least for short periods(Pankoke et al., 2010, 2012). However, larvae feeding onplants with high concentrations of aucubin weighed less anddeveloped more slowly compared to larvae feeding on plantswith low aucubin concentrations, probably since reduction

of insect β-glucosidase activity is not complete and someaucubin is still hydrolyzed. Further adaptations for long-termfeeding may include dietary mixing by host-plant switching(see Section III.1) (Pankoke et al., 2012)

When feeding on a diet with an increasing concentrationof the salicinoid salicortin, the generalists gypsy mothand forest tent caterpillar Malacosoma disstria (Lepidoptera,Lasiocampidae) adaptively reduced the activity of theirendogenous β-glucosidases in the midgut (Hemming &Lindroth, 2000). This inducible reduction in β-glucosidaseactivity prevents formation of toxic aglucones and enablesfeeding, but it may also reduce the efficiency of digestion,which could explain why both species show slightly decreasedgrowth rates on this diet (Hemming & Lindroth, 2000).

Even at an intraspecific level, activity of endogenous insectβ-glucosidases may differ. A subspecies of the lepidopteraneastern tiger swallowtail Papilio glaucus canadensis (Lepidoptera,Papilionidae) is a specialist adapted to salicinoids and feedsextensively on poplars and willows (Lindroth, 1988). Bycontrast, the subspecies P. g. glaucus is susceptible to salicinoidsand avoids feeding on those plants. Dietary exposure of bothsubspecies to salicin and salicortin reduced the activity of aspecific β-glucosidase in P. g. canadensis, while the activity wasincreased in P. g. glaucus (Lindroth, 1988). The lower activityof endogenous midgut β-glucosidases in P. g. canadensis

explains their adaptation to dietary intake of plant salicinoids.An inducible reduction of endogenous midgut β-

glucosidase activity was shown for desert locusts whenfeeding on Schouwia purpurea (Brassicaceae), a plant whichcontains glucosinolate concentrations tenfold higher than inmost other crucifers, in comparison to feeding on non-toxicplants (Mainguet et al., 2000). Such a reduction of midgutβ-glucosidase activity enabled S. gregaria to feed without anyadverse effects for at least 1 week. Feeding for more than3 weeks on S. purpurea resulted in slower mass gain (Mainguetet al., 2000), which could reflect lower digestion rates.

Inducible and selective reduction of the endogenous insectβ-glucosidase most active on glucosylated plant defencecompounds avoids intoxication while maintaining efficientdigestion of ingested plant material. Generalist caterpillarsof the sugar cane borer Diatraea saccharalis (Lepidoptera,Crambidae) have three β-glucosidases, βGly1–3, in theirmidgut (Ferreira, Parra & Terra, 1997). Dietary exposureof the sugar cane borer to the cyanogenic di-glucosideamygdalin results in degradation to the cyanogenic mono-glucoside prunasin due to βGly1 and βGly3 activity.However, activity of βGly2, which hydrolyses prunasinfurther into toxic aglucones, was strongly reduced (Azevedo,Terra & Ferreira, 2003). Moreover, general digestion isnot affected since the remaining two β-glucosidases, whichhydrolyse plant oligosaccharides, are still active (Azevedoet al., 2003).

Adaptation may also be due to a lack of endogenousinsect β-glucosidases able to hydrolyse glucosylated plantdefence compounds. This was shown for fall armywormlarvae that can be reared on an artificial diet containingamygdalin without any adverse effects or any reduction in



insect β-glucosidase activity (Ferreira et al., 1997). Amygdalinmay be hydrolysed to the mono-glucosylated prunasin, butthe two digestive β-glucosidases present in the midgut of thefall armyworm are unable to further hydrolyse prunasin intotoxic aglucones (Marana, Terra & Ferreira, 2000). Generaldigestive processes are not affected since both β-glucosidaseshydrolyse cellulose from the ingested plant material.

Adverse costs at a trophic level for insect endogenousβ-glucosidase activity were shown for the cabbage whitebutterfly Pieris rapae (Lepidoptera, Pieridae) feeding oncabbage plants containing glucosinolates (Mattiacci, Dicke& Posthumus, 1995). Regurgitate of P. rapae contains β-glucosidase activity that elicits a blend of volatiles from theplant that was highly attractive to parasitic wasps—naturalenemies of P. rapae. By contrast, the plant β-glucosidase fromthe cabbage leaf extract was not as efficient as the insectβ-glucosidase in releasing the volatiles.

In many cases, except in that of an alkaline gut (seeSection III.3a), it is unclear how the activity of endogenousβ-glucosidases is reduced. It may be that transcription ofthe gene or translation of its mRNA is decreased, that theenzyme is inhibited by other factors than pH or that itssubstrate specificity is altered. It would be interesting toanalyse how reduction in activity of endogenous insect β-glucosidases may affect digestive processes in the gut, how thisinteracts with nutrient uptake and how insect developmentmay be influenced.

(c) Specialized enzyme activity of insects

Apart from β-glucosidases, insects possess a variety of otherand more specialized enzymes involved in counteractingplant chemical defence. Cytochrome P450 monooxygenasesare important enzymes in insect metabolism and resistanceto many kinds of toxins such as insecticides, drugs andspecialized plant metabolites (Scott, 1999; Schuler, 2011;Feyereisen, 2012). However, cytochrome P450s seem tobe of lesser importance in insect herbivores that areadapted to two-component plant chemical defence, becausedetoxification via cytochrome P450s has not been reportedon these compound classes (Schuler, 2011; Feyereisen, 2012;Tao et al., 2012). Instead, insect enzymes convert defencecompounds either (i) before their hydrolysis by plant β-glucosidases, i.e. the modified product can no longer beactivated by plant β-glucosidases into toxic aglucones, or (ii)after their hydrolysis by plant β-glucosidases, via conjugatingor detoxifying enzymes that render aglucones easier toexcrete or less toxic (Fig. 2).

( i ) Before hydrolysis by plant β-glucosidases. Insectenzymatic conversion of glucosylated defence compoundsbefore their hydrolysis by plant β-glucosidases has beenshown in specialist caterpillars of the neotropical Saralongwing Heliconius sara (Lepidoptera, Nymphalidae) thatfeed on cyanogenic-glucoside-containing Passiflora auriculata(Passifloraceae) plants. This insect species prevents plant β-glucosidase activity on cyanogenic glucosides by metabolicreplacement of the nitrile group with a thiol group inthe cyanogenic glucoside by an as yet unknown enzyme

(Fig. 2C) (Engler, Spencer & Gilbert, 2000). Moreover,after catabolism of the cyanogenic glucoside epivolkenin, thereleased nitrogen is used in the insect’s primary metabolism.

Diamondback moth Plutella xylostella (Lepidoptera,Plutellidae) caterpillars are specialized feeders on cruciferplants, which are primarily defended by the glucosinolate-myrosinase two-component system. To prevent activationof glucosinolates, larvae possess a sulfatase in their gut thatconverts all major classes of glucosinolates into desulfo-glucosinolates, which can no longer be activated bymyrosinases (Fig. 2D) (Ratzka et al., 2002). The insectsulfatase directly competes with the plant myrosinase forglucosinolates, and furthermore the released sulfate inhibitsmyrosinases (Ratzka et al., 2002). Sulfatase expression isunder tight developmental and tissue-specific control, sincetranscripts are constitutively present in the larval gut, theonly stage and organ in the diamondback moth life cyclethat is exposed to glucosinolates, but sulfatase transcriptsare absent in other tissues and developmental stages (Ratzkaet al., 2002). Desert locusts that are generalists also possessa sulfatase with broad substrate specificity in their gut.Sulfatase activity enables feeding even on S. purpurea, aplant with very high concentrations of glucosinolates (Falk &Gershenzon, 2007). Following feeding on a glucosinolate-freediet, sulfatase activity increased tenfold when locusts were fedS. purpurea, and decreased when glucosinolates were removedfrom the diet. Thus, sulfatase activity in generalist desertlocusts is highly inducible, whereas it seems constitutive inthe specialist diamondback moth larva.

The interplay between specialized insect enzymes thatare active before plant β-glucosidases and sequestrationas a further adaptation (see Section III.3d ) was shownin the sawfly Athalia rosae (Hymenoptera, Tenthredinidae).Sawfly larvae take up glucosinolates from the gut intotheir haemolymph where they are degraded to desulfo-glucosinolates by sulfatases, and subsequently sulfated at theglucose moiety by sulfotransferases (Fig. 2D) (Opitz et al.,2011). Since excess glucosinolates are transported back intothe gut and excreted via the frass, prior conversion in thehaemolymph is highly adaptive, since modified glucosinolatescan no longer be activated by the remaining plant β-thioglucosidases in the gut (Muller, 2009; Opitz et al., 2011).

( ii ) After hydrolysis by plant β-glucosidases. Even whenplant β-glucosidases activate the corresponding glucosylateddefence compounds into toxic aglucones, some insects haveevolved enzymes for counteraction. Conjugating enzymesadd a glucose or glutathione moiety to the aglucone toincrease its water solubility and excretion efficiency. Others,more specialized detoxification enzymes modify specifictarget substrates and render them non-toxic.

For conjugation, glucose is activated by linkage to uridinediphosphate via a phosphoester bond. Activated glucoseis then used to form O-glucosides, an enzymatic reactioncatalysed by family 1 UDP-glucosyltransferases (UGTs). Re-glucosylation of toxic aglucones into non-toxic glucosidesdirectly in the gut is an efficient adaptation that allowsrapid and non-toxic excretion of the glucoside. For example,



the generalist larvae of the Egyptian armyworm and fallarmyworm re-glucosylate toxic aglucones such as DIMBOAinto non-toxic DIMBOA-glucoside via UGTs in the gut andexcrete them (Fig. 2B) (Glauser et al., 2011). Similarly, UGTactivity in the rice armyworm Mythimna separata (Lepidoptera,Noctuidae) midgut leads to formation and excretion of non-toxic DIMBOA-glucosides, which enables these generalists togrow on a DIMBOA-containing diet (Sasai et al., 2009). UGTactivity, and thus resistance to plant defence compounds,can also differ between closely related species. Whereas theAsian corn borer Ostrinia furnacalis (Lepidoptera, Crambidae)is able to feed on an DIMBOA artificial diet without adverseeffects, survival of the congener adzuki bean borer Ostrinia

scapulalis is strongly affected (Kojima et al., 2010). Differencesin resistance were linked to a higher activity of midgut UGTsin O. furnacalis than in O. scapulalis.

Conjugation of reduced glutathione to aglucones isanother adaptation for some insects and is carried out byglutathione S-transferases (GSTs). GSTs have substrates withan electrophilic centre, such as α/β-unsaturated carbonylcompounds, that may be electrochemically mimickedby isothiocyanates derived from glucosinolate-myrosinaseactivation (Brattsten, 1988). Hence, the generalists cabbagemoth Mamestra brassicae, cotton bollworm Helicoverpa

armigera (both Lepidoptera, Noctuidae), fall armyworm,Egyptian armyworm and the cabbage looper, whichfeed on glucosinolate-containing plants, conjugate toxicisothiocyanates with glutathione by GST activity in theirgut and subsequently excrete non-toxic products with thefrass (Fig. 2D) (Schramm et al., 2012). Induction of GSTactivity in response to increasing glucosinolate concentrationswas shown in green peach aphids (Francis, Vanhaelen& Haubruge, 2005) as well as in the midgut of gypsymoth larvae and forest tent caterpillars when feeding onplants containing salicinoids (Hemming & Lindroth, 2000).These results suggest that detoxification by GST activitymay be widespread in generalist lepidopterans and efficienton different classes of two-component chemical defence(Schramm et al., 2012).

Detoxification enzymes are for example reportedfrom the cabbage white butterfly, a specialized feederon glucosinolate-containing crucifers. Larvae possess anendogenous nitrile specifier protein (NSP) in their midgutthat redirects glucosinolate hydrolysis towards the less toxicnitriles instead of the more toxic isothiocyanates (Fig. 2D)(Wittstock et al., 2004). NSP may act as a cofactor bindingto the plant myrosinase to modify glucosinolate hydrolysis,or it may use the aglucone directly as a substrate to generatenitriles (Wittstock et al., 2004). Insect NSP activity seems tohave evolved shortly after their host plants evolved, and tohave enabled adaptive radiation of the Pierinae subfamilywith significantly elevated species numbers in comparisonto related clades (Wheat et al., 2007). Recent studies onthe detoxification efficiency of the NSP, however, haveshown conflicting results. Whereas P. rapae larvae feeding onplants with increased glucosinolate levels showed high NSPefficiency and no adverse effects in some studies (Gols et al.,

2008; Muller et al., 2010), another study showed negativeeffects of P. rapae larvae feeding on plants with increasedglucosinolate levels, and a less efficient NSP (Kos et al.,2012). It might be that nitriles, generated by NSP activity,are still toxic to P. rapae, but to a lesser extent (Burow &Wittstock, 2009), or that higher glucosinolate concentrationsimpose higher expression of NSP, which is metabolicallymore costly (Kos et al., 2012). It would be interestingto see if NSP expression level increases with increasingglucosinolate concentrations, and whether such an inductioninvolves higher metabolic costs. Similarly, it has beenshown that young larvae grow more slowly with increasingglucosinolate concentration on Brassica napus plants (Rotem,Agrawal & Kott, 2003), whereas older larvae were muchless affected by increasing glucosinolate concentrations.After NSP activity, nitriles from aliphatic glucosinolates areexcreted unmodified with the frass, whereas nitriles fromaromatic glucosinolates are α-hydroxylated into an unstableintermediate that decomposes spontaneously to an aldehydeand cyanide (Fig. 2D) (Vergara et al., 2006; Agerbirk et al.,2010; Stauber et al., 2012). Cyanide in P. rapae is detoxifiedby β-cyanoalanine synthase and rhodanese activity into non-toxic β-cyanoalanine and rhodanide (Stauber et al., 2012).These detoxification enzymes might have enabled pieridspecies to shift from the cyanogenic-glucoside-containingFabales to the aromatic-glucosinolate-containing Brassicalesaround 75 million years ago (Stauber et al., 2012).

Larvae of the six-spot burnet moth emit toxichydrogen cyanide due to endogenous β-glucosidase activityon sequestered or biosynthesized cyanogenic glucosides(Zagrobelny et al., 2008; Zagrobelny & Møller, 2011).Detoxification enzymes such as β-cyanoalanine synthaseenable burnets to endure the permanent exposure tohydrogen cyanide (Zagrobelny et al., 2008; Zagrobelny &Møller, 2011). β-cyanoalanine synthase activity convertscyanide with the amino acid cysteine into β-cyanoalanineand subsequently into non-toxic asparagine, another aminoacid (Fig. 2C) (Zagrobelny et al., 2004; Zagrobelny & Møller,2011). In agreement with the target site of hydrogen cyanidetoxicity, β-cyanoalanine synthase activity is found mainly inmitochondria (Watanabe et al., 2008).

(d ) Sequestration: spatial separation of plant β-glucosidase andβ-glucoside in the insect

Some specialized insects are able selectively to take upand accumulate plant defence compounds in their bodytissues, e.g. the haemolymph or defence glands (Nishida,2002; Opitz & Muller, 2009). This so-called sequestrationis an efficient adaptation, since glucosylated plant defencecompounds are no longer accessible to enzymatic hydrolysisby ingested plant β-glucosidases that remain in the gutlumen. Furthermore, sequestering insects become toxic topotential predators (Kazana et al., 2007; Zagrobelny et al.,2008; Discher et al., 2009). To date, more than 250 insectspecies from different orders have been shown to sequesterglucosylated and non-glucosylated plant defence compoundsfrom over 40 plant families, but little is known about potential



membrane carriers and their transport mechanism (Opitz &Muller, 2009).

Alkaloid, benzoxazinoid, cyanogenic and iridoid gluco-sides as well as glucosinolates and salicinoids are polarcompounds due to their β-d-glucopyranose moiety. Polar-ity prevents diffusion through the gut membrane into thehaemolymph, where plant defence compounds are no longeraccessible to plant β-glucosidases that remain in the gut(Muller, 2009; Opitz & Muller, 2009). Thus, transport ofglucosylated compounds into the haemolymph has to bespecific and efficient since plant β-glucosidases immediatelyhydrolyse them after tissue damage and this hydrolysis maycontinue in the gut (Desroches et al., 1997; Pankoke et al.,2012). Ingested plant β-glucosidases thus may have driventhe evolution of efficient transporters in insects (Muller &Wittstock, 2005; Muller, 2009). As the midgut is the mostpermeable part of the digestive tract (Dow, 1986), gen-eral transporters may enable most glucosides to enter thehaemolymph, and more specialized transporters may chan-nel only specific substrates into the final tissue (Kunert et al.,2008; Burse et al., 2009).

Larvae of the leaf beetles Chrysomela populi and Phratora

vitellinae (both Coleoptera, Chrysomelidae) sequester thesalicinoid salicin from their Salix spp. food plants andtransport it intact from the gut into the haemolymphand further on into their defence glands (Kuhn et al.,2004; Burse et al., 2009). Membrane carriers with moderateselectivity transport salicin through the gut membrane intothe haemolymph (Discher et al., 2009; Opitz & Muller, 2009),whereas highly specific carriers that match the orientation ofthe functional group of the glucopyranose ensure that onlysalicin is transported into the defence glands (Kuhn et al.,2007; Discher et al., 2009). Activity of insect β-glucosidasesin the defence glands hydrolyses salicin to the deterringcompound saligenin (Fig. 2F), which may be finally oxidizedto salicylaldehyde (Kuhn et al., 2004; Opitz & Muller, 2009).Highly evolved chrysomelina leaf beetles such as Chrysomela

lapponica are able to sequester a wide variety of glucosylatedleaf alcohols into their defence glands (Hilker & Schulz, 1994;Schulz, Gross & Hilker, 1997; Burse et al., 2009). After insectβ-glucosidase activity, aglucones are esterified with butyricacid derived from the insect’s own amino acid pool finallyto produce a diversity of deterring esters (Hilker & Schulz,1994; Schulz et al., 1997).

Absence of β-glucosidase activity in the insecthaemolymph seems to enable bruchid beetles C. maculatus

to sequester intact plant defence compounds (Desrocheset al., 1997). The alkaloid glucoside vicine was shown to betransferred intact from the midgut into the haemolymph,where it is stored without chemical transformation until theadult instar stage of larval development (Desroches et al.,1997).

Sequestering insects that produce β-glucosidases that canactivate sequestered defence compounds might benefit fromseparating both components spatially in their body. As inplants, compartmentalization ensures that insect predatorsare exposed to toxic aglucones only upon tissue damage

and mixing of both components. The Brassica specialists,the cabbage aphid, and the mustard aphid Lipaphis erysimi(Hemiptera, Aphididae), sequester intact glucosinolates fromtheir host plants and possess an endogenous myrosinase.These insects prevent hydrolysis of glucosinolates by storingtheir own myrosinases in crystalline microbodies in non-flightmuscles, whereas sequestered plant glucosinolates are storedseparately in the haemolymph (Bridges et al., 2002; Kazanaet al., 2007). Six-spot burnet moth larvae sequester thecyanogenic glucosides linamarin and lotaustralin from theirL. corniculatus food plants and store them in all tissues includingthe haemolymph, where endogenous β-glucosidases arealso present (Zagrobelny et al., 2008). Non-separate storageof both components leads to a continuous hydrolysis ofcyanogenic glucosides and emission of hydrogen cyanidefor defence. Larvae of the Sara longwing sequester thecyanogenic glucoside epivolkenin from their P. auriculata hostplants (Engler et al., 2000). Sequestration is highly selectivesince only epivolkenin is taken up and accumulated, whereasthe other plant cyanogenic glucosides are not sequestered.

(e) Single amino acids counteract plant β-glucosidase activity

In addition to the various adaptations that often targetplant β-glucosidase activity, single amino acids wereshown to counteract activity of plant β-glucosidasesmainly by reducing the toxicity of released aglucones.Several lepidopteran and hymenopteran larvae secreteextraordinarily high amounts of glycine, β-alanine or γ -aminobutyric acid (GABA) into their gut lumen when feedingon plants containing the iridoid glucoside oleuropein (Konnoet al., 1997, 1998, 2010; Konno, Okada & Hirayama,2001). These single amino acids protect insect digestiveenzymes and dietary protein by competing with the aminoresidue in the lysine side chains of proteins for the protein-crosslinking activity of aglucones (Konno et al., 2010). Leavesof the privet tree are defended by oleuropein, but larvaeof the lepidopteran Brahmaea wallichii (Brahmaeidae) andDolbina tancrei (Sphingidae) or the hymenopteran Macrophyatimida (Tenthredinidae) can feed on this plant by secretingextremely high concentrations of free glycine (up to 164 mM)into their midgut (Konno et al., 1997, 2010). This is morethan 20-fold higher than seen in non-privet feeders (Konnoet al., 2010). Specific glycine-transporters mediate secretionin the anterior part of the midgut (Konno et al., 1998,2001). Secretion in the anterior midgut is adaptive, sincedenaturation and lysine decrease begin immediately afterleaf damage and it is beneficial to stop this reaction as soonas possible after ingestion (Konno et al., 2001). Interestingly,when non-adapted privet feeders, such as caterpillars ofthe Eri silkworm Samia ricini (Lepidoptera, Saturniidae) arefed privet leaves together with high levels of free glycine,the toxic effects of plant β-glucosidase-activated oleuropeinare completely prevented and the caterpillars grow withoutapparent detrimental effects (Konno et al., 2009). Otherlepidoptera such as Artopoetes pryeri (Lycaenidae) secrete highconcentrations of GABA (up to 60 mM) into their midgut,or high levels of β-alanine (up to 35 mM), as shown in



caterpillars of Amphipyra monolitha (Noctuidae) (Konno et al.,2010). In this generalist, the secretion is induced only whenfeeding on privet trees, while no secretion occurs whenfeeding on a plant species devoid of oleuropein (Konno et al.,2010). Since aglucones of other two-component defenceclasses also have protein-denaturing activities, secretion ofhigh levels of certain amino acids into the midgut may be apotential adaptation in other insect herbivores.

(4) Do generalists and specialists have differenttypes of adaptations?

Do generalists and specialists have different types ofadaptations? This question is relevant since generalistsand specialists differ in their host range and hencethe β-glucosidase-activated plant defence compounds theyencounter. Could the exposure to several defence compoundsas in generalists result in adaptations that are inducible byplant defence compounds or have broad effects on differentclasses of compounds? Conversely, for specialists, couldexposure to single defence compounds lead to constitutiveadaptations or adaptations that are highly effective, but onlyon a small number of similar compounds?

The adaptation of host-plant switching seems to supportthese notions, since this behaviour is mainly shown bygeneralists. The frequency of host-plant switching dependson the toxicity of the plant defence compound (Singer et al.,2002) and probably its concentration in the insect body.Thus, the decision to switch host plants in generalists maybe inducible by both factors. Feeding guilds such as non-disruptive piercing-sucking and leaf-snipping are efficientadaptations found in both generalists and specialists (Bernays& Janzen, 1988; McMahon et al., 1995; Bridges et al., 2002;Barth & Jander, 2006; Ballhorn et al., 2010). However,from two lepidopteran families there are morphologicalindications that generalists and specialists do differ. Mandiblestructure of generalist species within each group was similar,whereas specialists had very characteristic mandibles, eachof unique design and related to the nature of its host leaf(Bernays & Janzen, 1988). Adaptations that occur duringdigestion of plant material such as an alkaline pH in the gutlumen were mainly shown in generalists and could have abroader adaptive effect (Ruuhola et al., 2003; Fitzgerald,2008; Dutartre et al., 2011). The question of whetheralkalinity is inducible has not been answered clearly yet,because only a few studies have measured the pH of emptyas well as filled guts. In addition, studies were only carriedout on generalists showing that midgut lumen alkalinityis constitutive in fall webworm larvae (Fitzgerald, 2008),whereas it is inducible in gypsy moth larvae (Schultz &Lechowicz, 1986). A reduction of endogenous midgut β-glucosidase activity that is sensitive to and inducible by thelevel of two-component defence chemical is mainly foundin generalists (Hemming & Lindroth, 2000; Mainguet et al.,2000; Azevedo et al., 2003; Pankoke et al., 2012), and onlyin a few specialists (Lindroth, 1988; Ballhorn et al., 2010).When comparing further enzymatic adaptations, we findboth similarities and differences between generalists and

specialists. For instance, both groups possess a sulfatase witha broad specificity in their gut (Ratzka et al., 2002; Falk& Gershenzon, 2007). However, specialized enzymes fordetoxification such as NSP and β-cyanoalanine synthase aremost often found in specialists (Barrett & Heil, 2012), becausesuch enzymes act on very specific substrates (Wittstock et al.,2004; Zagrobelny & Møller, 2011). In turn, broad-substrate-specificity conjugating enzymes such as UGTs and GSTs aremainly found in generalists (Kojima et al., 2010; Schrammet al., 2012), and induction of enzyme activity such as GSTsand sulfatases has also only been found in generalists so far(Francis et al., 2005; Falk & Gershenzon, 2007). Enzymeinduction that is sensitive to the level of plant defencecompound reduces metabolic costs of toxin processing andis especially adaptive for generalists that encounter a varietyof plant defence compounds (Falk & Gershenzon, 2007).Sequestration of plant defence compounds activated by plantβ-glucosidases is mainly found in specialists (Bridges et al.,2002; Discher et al., 2009; Opitz, Jensen & Muller, 2010;Zagrobelny & Møller, 2011), and only in one generalistspecies so far (Desroches et al., 1997). If given the opportunity,most such specialized insects sequester constitutively, but theconcentration of sequestered plant defence compounds inthe insect varies during larval development, and there is abalance between uptake and turnover (Muller & Wittstock,2005). In all cases, sequestration of intact glucosylated plantdefence compounds seems to require further adaptationssuch as non-disruptive feeding, an alkaline gut lumen,reduced endogenous insect β-glucosidase activity or thepresence of other insect enzymes that counteract plant β-glucosidase activity as discussed in Section III.3c. Secretionof high levels of single amino acids has mainly been found inspecialists feeding on iridoid-glucoside-containing plants, andonly in a few generalists. Species from different lepidopteranand hymenopteran families have evolved this ability (Konnoet al., 1997, 2010), and it seems to be a constitutive adaptationin these specialists. In generalists, secretion of amino acidsmay be inducible by plant iridoid glucosides as shown in thecase of the caterpillar A. monolitha (Konno et al., 2010).

Whereas some types of adaptation are found in both gen-eralists and specialists, others appear to be limited to eithergeneralists or specialists. Most striking is the observationthat adaptations are often inducible in generalists, whereasadaptations in specialists seem to be constitutive.

IV. CONCLUSIONS

(1) The various adaptations of insect herbivores towidespread two-component plant chemical defences, i.e.β-glucosidase-activated defence compounds, occur beforeand during feeding as well as largely during digestion, andinclude the insect’s behaviour, physiology and metabolism.Moreover, adaptations from these different time points andlevels are often combined in both generalists and specialistsas well as for the different classes of two-component chemicaldefence. Adaptations between generalists and specialists may



differ. In generalists, adaptations are often inducible and mayhave broader effects, whereas they seem to be constitutiveand provide more specific adaptations in specialists.

(2) Insect adaptations often target the activity of theenzymatic key component of the plant’s two-componentdefence, the β-glucosidase, at different time points of feedingas well as at different levels. Host-plant switching may reduceexposure to and adverse effects of plant defence compoundsby preventing the accumulation of toxic levels of aglucones.Feeding guilds that are minimally disruptive, especiallypiercing-sucking and leaf-snipping, prevent mixing of plantβ-glucosidase and glucosylated defence compound. A highlyalkaline midgut directly inhibits plant β-glucosidase activity.The reduction of endogenous insect β-glucosidase activity inthe gut lumen reduces the release of toxic aglucones. Diversefurther insect enzymes modify defence compounds eitherbefore their hydrolysis by plant β-glucosidases to preventactivation, or after their hydrolysis by plant β-glucosidasesto render aglucones less toxic. Sequestration of plant defencecompounds serves to remove them from the gut beforethey are activated by ingested plant β-glucosidases. Highlevels of single amino acids that are secreted into the gutlumen counteract plant β-glucosidase activity by reducingthe toxicity of released aglucones.

(3) The midgut pH differs widely among insect orders. Asthe midgut is the target site of plant defence compoundtoxicity, but also of sequestration, it is important tomeasure the exact pH on intact tissue and to use thispH for enzyme kinetic studies. The midgut pH mayalso be a physiological key parameter when investigatingmechanisms of sequestration. On the one hand a highlyalkaline midgut pH that inhibits plant β-glucosidase activitymay facilitate sequestration of intact glucosylated plantdefence compounds, but on the other hand would requiretransporters that function efficiently under such extremephysiological conditions.

(4) Further studies are encouraged to investigate howadaptations are combined and may interact in an ecologicallyrelevant context. For example, how does the reducedactivity of endogenous insect β-glucosidases interfere withoverall digestion of plant material and how does this affectdevelopmental processes? As this strategy is more commonin generalists, does this reflect a trade-off between foodavailability and efficiency? The use of mutant or geneticallymodified plants, which differ in single defence traits such asdefence compound synthesis or activation, will allow morecontrolled and detailed studies on mechanisms, costs andbenefits of sequestration and expression patterns of insectenzymes that counteract plant chemical defence.

V. ACKNOWLEDGEMENTS

S.P. acknowledges a LIFE Ph.D. scholarship from theUniversity of Copenhagen, Denmark. We acknowledgesupport by the Villum Foundation to the research centre‘Pro-Active Plants’ and grants from the Danish Council

for Independent Research, Technology and ProductionSciences. We thank two anonymous reviewers for theirhelpful suggestions.

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(Received 23 November 2012; revised 3 September 2013; accepted 5 September 2013 )


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