Journal of Insect Physiology 84 (2016) 70–89

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Journal of Insect Physiology

journal homepage: www.elsevier .com/ locate/ j insphys

Review

Insect-induced effects on plants and possible effectors used by gallingand leaf-mining insects to manipulate their host-plant

http://dx.doi.org/10.1016/j.jinsphys.2015.12.0090022-1910/� 2015 Elsevier Ltd. All rights reserved.

⇑ Corresponding author.E-mail address: david.giron@univ-tours.fr (D. Giron).

David Giron a,⇑, Elisabeth Huguet a, Graham N. Stone b, Mélanie Body c

a Institut de Recherche sur la Biologie de l’Insecte, UMR 7261, CNRS/Université François-Rabelais de Tours, Parc Grandmont, 37200 Tours, Franceb Institute of Evolutionary Biology, University of Edinburgh, Edinburgh EH9 3JT, United KingdomcDivision of Plant Sciences, Christopher S. Bond Life Sciences Center, 1201 Rollins Street, University of Missouri, Columbia, MO 65211, United States

a r t i c l e i n f o

Article history:Received 21 October 2015Received in revised form 21 December 2015Accepted 22 December 2015Available online 23 December 2015

Keywords:Gall-inducing insectsLeaf-minersPlant manipulationEffectors

a b s t r a c t

Gall-inducing insects are iconic examples in the manipulation and reprogramming of plant development,inducing spectacular morphological and physiological changes of host-plant tissues within which theinsect feeds and grows. Despite decades of research, effectors involved in gall induction and basic mech-anisms of gall formation remain unknown. Recent research suggests that some aspects of the plantmanipulation shown by gall-inducers may be shared with other insect herbivorous life histories. Here,we illustrate similarities and contrasts by reviewing current knowledge of metabolic and morphologicaleffects induced on plants by gall-inducing and leaf-mining insects, and ask whether leaf-miners can alsobe considered to be plant reprogrammers. We review key plant functions targeted by various plant repro-grammers, including plant-manipulating insects and nematodes, and functionally characterize insectherbivore-derived effectors to provide a broader understanding of possible mechanisms used in host-plant manipulation. Consequences of plant reprogramming in terms of ecology, coevolution and diversi-fication of plant-manipulating insects are also discussed.

� 2015 Elsevier Ltd. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 712. Induced effects in host-plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74

2.1. Altering plant morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74

2.1.1. Induced cellular modifications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 742.1.2. Possible targeted plant developmental pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 762.1.3. Patterns of diversification within galling lineages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

2.2. Controlling plant nutritional quality . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

2.2.1. Improvement of nutrient concentrations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 772.2.2. Disruption of plant defense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 772.2.3. Multitrophic selection on gall induction traits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

2.3. Hijacking the plant signaling network . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

3. Effectors used by insects to manipulate plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

3.1. Effectors of gall-inducing insects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 793.2. Other insect derived effectors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

3.2.1. Glucose oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 803.2.2. ATP-hydrolyzing enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 803.2.3. Calcium-binding proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 813.2.4. Other candidates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

3.3. Insect effectors and plant growth regulators. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 813.4. Microbial partners and plant manipulation in insect–plant interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

3.4.1. Insect symbionts and suppression of plant defense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

D. Giron et al. / Journal of Insect Physiology 84 (2016) 70–89 71

3.4.2. Insect microbial symbionts and impact on plant nutritional status and morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 823.4.3. Insect microbial symbionts and effectors encoded by horizontally transferred genes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 823.4.4. Microbes and insect plant specialization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

4. Concluding thoughts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

1. Introduction

Nutrition is the cornerstone ofmost interactions between organ-isms. With more than 4 million estimated species, insects areamong the most significant evolutionary successes on Earth(Novotny et al., 2002). The origin of this success can be directlylinked to the diversity of their feeding strategies, of which herbivoryis the most common (Schoonhoven et al., 2005; Slansky andRodriguez, 1987). However, plant tissues are typically suboptimalnutritionally, due to unbalanced ratios and/or low levels of keynutrients and frequent requirement to detoxify plant defensive alle-lochemicals (Schoonhoven et al., 2005). The ability of phytophagousarthropods to exploit plant resources requires them to employ asuite of pre- and post-ingestive mechanisms to address the nutri-tional mismatch between what plants provide and what insectsrequire (Behmer, 2009; Raubenheimer et al., 2009; Schoonhoven

Fig. 1. Examples of plant manipulation by gall-inducing and leaf-mining insects. (Gracillariidae) on maple leaf. (b) ‘Blister galls’ induced by a gall midge fly (unidentified spgall midge fly Schizomyia vitiscoryloides (Diptera, Cecidomyiidae) on grape nodes. (d) ‘Tomon grape flower buds. (e) Green-island (around a serpentoid mine) induced by the leaf-mgall’ induced by the Cynipid wasp Callirhytis lanata (Hymenoptera, Cynipidae) on oak leCynipidae) on rose leaf bud. (h) ‘Tube galls’ and ‘pocket-like galls’ respectively induced baphid, Daktulosphaira vitifoliae (Hemiptera, Phylloxeridae), on grape leaf. (i) Green-(Lepidoptera, Gracillariidae) on apple-tree leaf. Credit photo: D. Giron (a) and M. Body (

et al., 2005). These strategies include associations with one or moresymbiotic partners providing new metabolic pathways (Douglas,2009, 2013; Moran et al., 2008), symbioses in which plants haveevolved food rewards specifically for insects (e.g. Heil and McKey,2003), and/or intricate interactions that involve insect reprogram-ming of host-plant development, resulting in new structures bene-fiting the parasitic herbivore at the expense of the plant (Giron et al.,2013; Price et al., 1987; Stone and Schönrogge, 2003).

Insect galls are structures composed of plant tissues thatdevelop in response to stimuli produced by the gall-inducer, andpresent inducer-specific phenotypes and patterns of tissue differ-entiation (Stone and Schönrogge, 2003). The need for inductionby another organism distinguished true galls from other structuresthat have evolved to accommodate insects but which are underplant control – such as the domatia of myrmecophytes occupiedby guarding ants (e.g. Raine et al., 2002). Galls are among the most

a) Green-island induced by the leaf-miner Phyllonorycter joannisi (Lepidoptera,ecies; Diptera, Cecidomyiidae) on honeysuckle leaf. (c) ‘Hazelnut gall’ induced by theato galls’ induced by the gall midge fly Janetiella brevicauda (Diptera, Cecidomyiidae)iner Lyonetia clerkella (Lepidoptera, Lyonetiidae) on a Prunus tree’s leaf. (f) ‘Woollyaf. (g) ‘Bedeguar gall’ induced by the Cynipid wasp Diplolepis rosae (Hymenoptera,y the gall midge fly Schizomyia viticola (Diptera, Cecidomyiidae) and the phylloxeraisland induced by the spotted tentiform leaf-miner Phyllonorycter blancardellab–i).

Table 1Plant alteration by manipulating organisms. Comparison between leaf-mining insects, gall-inducing insects and gall-inducing nematodes for plant nutrient, defense, morphology and phytohormone alterations (selected publications).Credit photo: William Wergin and Richard Sayre (nematode), Mélanie Body (leaf-miner), Mélanie Body and Ryan Richardson (galling insect).

Leaf-mining insect Galling insect Root-knot nematode phytopathogen

Nutrients Nutrition optimization Nutrition optimization Nutrition optimization� Alteration of source-sink relationship (Body et al., 2013;Giron et al., 2007)

� Control of sugar and amino acid composition (Body, 2013;Body et al., 2013)

� Alteration of source-sink relationship (Compson et al., 2011;Larson and Whitham, 1991)

� Control of sugar and amino acid composition (Dardeau et al.,2015; Liu et al., 2007; Saltzmann et al., 2008; Stuart et al.,2012)

� Alteration of source-sink relationship (Kaplan et al., 2011;Kyndt et al., 2013)

� Control of mineral, starch, sugar and amino acid composition(Nasr et al., 1980)

Selective feeding Selective feeding Selective feeding� Nutrient-rich cells (Body et al., 2015)� Nutrient-rich tissues (Body et al., 2015; Scheirs et al., 2001)

� Nutritive tissue (Harris et al., 2006; Shorthouse andRohfritsch, 1992)

� Nutritive tissue (Caillaud et al., 2008; Favery et al., 2016)

Defenses Defense inhibition Defense inhibition Defense inhibition� Phenolic compounds (Giron et al., unpublished) � Phenolic compounds (Liu et al., 2007; Stuart et al., 2012)

� Tannins (Ikai and Hijii, 2007)� Proteases (Liu et al., 2007)� Volatiles (Tooker and De Moraes, 2007)

� Phenolic compounds� Reactive Oxygen Species (ROS) (Dubreuil et al., 2007; Faveryet al., 2016)

Physical mechanisms� Vein-cutting (Whiteman et al., 2011)

Morphology ? Induction of nutritive tissues (Harris et al., 2006; Stuart et al.,2012)

Induction of giant cells at feeding site (Caillaud et al., 2008; Faveryet al., 2016)

Alteration of cell-walls at feeding site Alteration of cell-walls at feeding site Alteration of cell-walls at feeding site� Chemistry (Body, 2013) � Chemistry and morphology (Harris et al., 2006, 2010; Khajuria

et al., 2013; Liu et al., 2007; Stuart et al., 2012)� Chemistry and morphology (Favery et al., 2016; Rosso et al.,2012)

Regulation of stomata opening (Pincebourde and Casas, 2006) Induction of stomata formation (Nabity et al., 2013) ?? Formation of new vascular elements (Wool et al., 1999; Wool,

2005)?

Phytohormones Phytohormone accumulation/enhanced signaling Phytohormone accumulation/enhanced signaling Phytohormone accumulation/enhanced signaling� CKs (Body et al., 2013; Giron et al., 2007; Zhang et al., 2016)� SA (Zhang et al., 2016)

� CKs (Mapes and Davies, 2001b)� IAA (AUX) (Mapes and Davies, 2001a; Tooker and De Moraes,2011a, 2011b)

� CKs (Kyndt et al., 2013; Rosso et al., 2012; Siddique et al., 2015)� AUX (Kyndt et al., 2013; Rosso et al., 2012)� GAs (Kyndt et al., 2013)� BRs (Kyndt et al., 2013)

Phytohormone decrease/reduced signaling Phytohormone decrease/reduced signaling Phytohormone decrease/reduced signaling� ABA (Zhang et al., 2016)� JA (Zhang et al., 2016)

� ABA (Tooker and De Moraes, 2011a, 2011b)� SA (Tooker and Helms, 2014)� JA (Tooker and Helms, 2014)

� SA (Favery et al., 2016; Kyndt et al., 2013)� JA (Ji et al., 2013)� ET (Kyndt et al., 2013)

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Table 2Effectors involved in plant manipulation. Potential effectors involved in plant manipulation (selected publications). Effectors for which functional data is available are shaded in gray. HGT: horizontal gene transfer. PCWDE: plant cell-wall degrading enzymes; see text for other abbreviations. Credit photo: William Wergin and Richard Sayre (nematode), Mélanie Body (leaf-miner, caterpillar and aphid), Mélanie Body and Ryan Richardson (galling insect).

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emblematic examples of plant manipulation by insects. By ‘repro-gramming’ expression of the plant genome, the insect forces theplant to create specialized nutritional and sometimes protectiveresources that benefit the insect at the expense of the plant’sgrowth and reproduction. Gall induction involves hijacking of plantcellular machinery and development, creating completely newstructures with features and functions of a novel plant organ(Fig. 1) (Mani, 1964; Shorthouse and Rohfritsch, 1992; Shorthouseet al., 2005a; Stone and Schönrogge, 2003). Gall structures rangefrom highly complex and differentiated structures including nutri-tive inner tissues and complex protective outer tissues (e.g. in cyni-pids, aphids, thrips and gall midges) to less dramatic structurescomprising nutritive tissues only (as in the Hessian fly Mayetioladestructor, whose open galls at the base of its host-plant are pro-tected between the leaf sheaths). The Hessian fly shares with manyother gallers the ability to induce a highly differentiated nutritivetissue that provides the developing larva with a diet rich in nutri-ents (Harris et al., 2006; Saltzmann et al., 2008). Nutritive tissuesare both more nourishing and less well defended than are non-gall tissues on the same plant and are a typical feature of manygall-inducing organisms, including insects, mites, nematodes, fungi,bacteria and viruses (Bronner, 1992; Favery et al., 2016; Harriset al., 2006; Mani, 1992; Oliveira et al., 2016; Pointeau et al.,2013; Rohfritsch, 1992; Stone and Schönrogge, 2003).

In insects, mechanisms underlying gall induction and growthare largely unresolved, driving a very active research area that isthe subject of many of the papers in this Special Issue. We suggestthat looking at similarities in the responses elicited by plant repro-grammers in a diversity of systems may identify key plant func-tions that are targeted during plant manipulation. An overview ofplant functions targeted by these reprogrammers (Table 1) and ofthe insect herbivore-derived effectors that could possibly beinvolved (Table 2) may facilitate inference of the stimuli and pro-cesses involved in gall induction. We first review current knowl-edge of the effects on plants induced by leaf-mining and gall-inducing insects, highlighting ways in which leaf-miners can alsobe considered plant reprogrammers (Table 1). We then compareplant metabolic and morphological responses to plant-reprogramming insects and nematodes and identify candidateinsect effectors for plant manipulation (Table 2). We explore theconsequences of plant manipulation for the ecology, coevolutionand diversification of plant-manipulating insects (Fig. 2).

2. Induced effects in host-plants

Insects that manipulate plant development show strategies thatremodel the cell content and structure of plant tissues at their feed-ing site (Fig. 1). These changes lead to demonstrably improvednutritional value and/or nutrient access, and have been interpretedas adaptations that address themismatch between the nutrient sta-tus of unmodified host-plant tissues and the herbivore’s require-ments (Table 1) (Body, 2013; Body et al., 2013; Dardeau et al.,2014a; Harris et al., 2006; Liu et al., 2007; Moura et al., 2008;Oliveira and Isaias, 2010; Souza et al., 2000). Induction of nutritivetissues is a key feature of many gall-inducing insects (Harris et al.,2006; Rohfritsch, 1992; Stone and Schönrogge, 2003). The evolutionof nutritive tissues may be associated with the development ofadditional tissues that partially or wholly surround the gall-inducer. These additional tissues have been hypothesized to protectthe inducer from abiotic stress and/or natural enemies, though (ashighlighted for Hessian fly above), not all galls have them (Harriset al., 2006; Pincebourde and Casas, 2016; Stone and Schönrogge,2003). Similarly, the degree of nutritive tissue differentiation alsovaries among gall-inducers, being most obvious where the insectsactively consume the tissues (e.g. cynipid and chalcid wasps) ratherthan feeding on them suctorially (e.g. aphids).

2.1. Altering plant morphology

2.1.1. Induced cellular modificationsGall-inducers manipulation of host-plant development results

in complex tissue reorganization, sometimes effectively resultingin new plant organs (Mani, 1964; Harper et al., 2004; Rohfritsch,1992; Shorthouse et al., 2005a). In response to gall induction stim-uli from the ovipositing mother and/or her offspring, host tissuesusually dedifferentiate and gall development often involves a com-bination of cell division (hyperplasia) and growth (hypertrophy)(Carneiro et al., 2014, 2015; Dias et al., 2013; Oliveira and Isaias,2010; Suzuki et al., 2015). Cell divisions occur in several planes,increasing the number of cell layers and hence the thickness ofthe host tissues. In leaves, both the homogenization of the par-enchyma and cell hypertrophy are common phenomena in gall for-mation and are related to the modification of the mesophyll intonutritive and protective tissues (Mani, 1964; Moura et al., 2008;Oliveira and Isaias, 2010; Pincebourde and Casas, 2016; Priceet al., 1987; Rohfritsch, 1992; Souza et al., 2000; Stone andSchönrogge, 2003). Gall tissue differentiation can also involvechanges in cell ultrastructure rather than hypertrophy and hyper-plasia. For example, imaging studies of development of the nutri-tive tissues of Hessian fly galls show nutritive cells to exhibit anincrease in cytoplasmic staining, in numbers of cellular organelles(mitochondria, proplastids, Golgi, and rough endoplasmic reticu-lum), along with development of numerous small vacuoles andan irregularly shaped nucleus (Harris et al., 2006).

Nutritive tissue can be very localized or more extensive, cover-ing most of the inner surface in many closed galls (Shorthouse andRohfritsch, 1992). In some cases, immature stages actively feed ona lining of nutritive tissues, pupating when all has been consumed.In cynipid galls, the nutritive tissues are typically surrounded by ahard schlerophyllous shell surrounding the larval chamber, withthese and other gall tissues supplied by vascular tissues connectedto the vascular system of the host-plant (Stone et al., 2002). Newsieve tubes (phloem elements) are usually formed below the innergall surface, supplying and enveloping the nutritive tissue, such asin galls induced by Geoica wertheimae aphids (Rohfritsch, 1992;Wool et al., 1999; Wool, 2005).

Characteristics of nutritive cells vary depending on the familyand species of gall-inducer, and can show differentiation betweenthe inner ‘typical nutritive tissue’ on which the inducer feeds, andmore peripheral ‘storage nutritive tissue’ (Bronner, 1992). Typicalfeatures of nutritive cells induced by galling insects, and alsoroot-knot nematodes, include endoreduplicated or hypertrophiednuclei and nucleoli, reduced and fragmented vacuoles, increasednumber of cellular organelles (mitochondria, proplastids, Golgi,and rough endoplasmic reticulum), dense cytoplasm, and frequentcommunications between cell layers and vascular tissues throughnumerous/enlarged plasmodesmata (Bronner, 1992; Dardeauet al., 2014a; Favery et al., 2016; Harper et al., 2004; Harris et al.,2006; Isaias and Coelho De Oliveira, 2012; Jones and Payne,1978; Oliveira et al., 2016; Rodiuc et al., 2014; Schönrogge et al.,2000). Additionally, cells usually have thin walls because of theinhibition of cell-wall fortification and expansion. For example,within Hessian fly (M. destructor) feeding sites, genes involved inthe biosynthetic pathways for cell-wall components (such as cellu-lose synthase) and loosening (such as xyloglucan endotransgluco-sylase (XET), b-expansin, endo-1,4-b glucanases, pectinesterase,and polygalacturase) are downregulated (Liu et al., 2007;Khajuria et al., 2013). Variations in cell-wall composition (espe-cially polysaccharides) during gall development have just startedto be explored, highlighting the functionality of cell-walls in gallontogenetic steps and their role in insect nutrition or plant resis-tance (Carneiro et al., 2014; Khajuria et al., 2013). Gall parenchymacells generally show reduced intercellular spaces, characteristic of

Free-feeding ac�ve herbivore

Leaf-miners

Gall-inducers

Sedentary surface-feeding insects

Differen�al plant growthcauses feeding depressions

Gall-inducers

HYPOTHESIS #1 HYPOTHESIS #2

1. Enemies2. Abiotic factors

1. Nutrition2. Enemies3. Abiotic factors

1. Nutrition

1. Enemies2. Abiotic factors3. Nutrition

1. Nutrition

Free-feeding ac�ve herbivore

Gall-inducersLeaf-miners

HYPOTHESIS #1 HYPOTHESIS #2

1. Nutrition

1. Enemies2. Abiotic factors3. Nutrition

1. Abiotic factors2. Nutrition3. Enemies

Gall-inducersLeaf-miners

1. Enemies2. Abiotic factors3. Nutrition

?

HYPOTHESIS #1

Ancestral plant-manipulator

(A)

(B)

Fig. 2. Origin of the gall-inducing habit. (A) The two routes to gall formation from free-feeding active herbivores suggesting leaf-miners as an intermediate primitive form ofendophytophagy (adapted from Price et al. (1987)). (B) Possible routes for the evolution of plant-manipulating insects. Leaf-miners and gall-inducers may have evolvedindependently (dashed arrows), finding similar solutions to face analogous threats (adaptive convergence) or they may have evolved from a common plant-manipulatingancestor (solid arrows) leading to shared ancestral plant manipulation strategies (homologies). Possible selective factors are listed in possible order of importance forimmediate impact on herbivore fitness. Credit photo: M. Body.

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young and fast-growing tissues. Such lack of intercellular spacesindicates low potential for gas exchange and consequent reducedphotosynthetic capacity in some galls (Carneiro and Isaias, 2015;Moura et al., 2008).

Gall tissues act as a strong resource sink for photo-assimilates,and can be associated with massive changes in plant growth, meta-bolism, and investment (Allison and Schultz, 2005; Anderson andHarris, 2006, 2008; Bagatto et al., 1996; Fay et al., 1993; Larsonand Whitham, 1991; Raman et al., 2006; Rehill and Schultz, 2003,2012), linked directly to the feeding site and hence growth of thegall-inducer (Compson et al., 2011; Harris et al., 2006). Inducedchanges in organization of the plant vascular tissues facilitate nutri-ent translocation and insect access to host-plant nutrients (Woolet al., 1999). Nutritive cells also have an intense metabolic activitycontrolled by the larvae, resulting in the up-regulation of proteinand/or sugar synthesis directly at the insect’s feeding site(Bronner, 1992; Giron et al., 2007; Larson and Whitham, 1991;Stone and Schönrogge, 2003). Modified cell-wall morphology andcomposition may help insects to feed because plant cell contentsbreak down and move from cell to cell through compromisedcell-walls (Bronner, 1992; Harris et al., 2006; Rohfritsch, 1992). Thisprocess shows a contrast in resistant and susceptible strains ofwheat attacked by Hessian fly; cytological studies show cell-wallbreakdown in susceptible plants (Harris et al., 2006) and fortifica-tion of the cell-wall in resistant plants (Harris et al., 2010). Thebreakdown of nutritive cells in the uppermost layer of the nutritivetissue has been observed for a number of gall-inducing insectsincluding cynipid gallwasps and cecidomyiid gall midge larvae(Bronner, 1992; Rohfritsch, 1992). Cell harvest in the nutritive tis-sue by insects most likely results from a combination of increasedpermeability of plant cell-walls, enlarged plasmodesmata, mechan-ical action of larval mouthparts, high turgor pressure in the nutri-tive cells, and possibly secretion and injection of proteases intocells by gall-inducing insects (Harris et al., 2006).

As far as we know, while leaf-miners can influence the nutritivequality of host-plant tissues (see below), no morphologically dis-tinct nutritive tissue has yet been found in insect mines. There isno obvious tissue differentiation involving hyperplasia or hyper-trophy of cells around mining insect larvae. Morphological studieson mines remain scarce (e.g. Body et al., 2015; Jeanneau, 1971;Melo De Pinna et al., 2002; Scheirs et al., 2001) (Table 1) and thisremains an area for further investigation.

2.1.2. Possible targeted plant developmental pathwaysPlant galls can be viewed as novel plant organs whose develop-

ment is triggered by induction of existing plant developmentalpathways in unusual (ectopic) places and/or combinations(Harper et al., 2004). Galls often contain plant tissue types thatare found nowhere else on the plant – such as the brightly coloredtissues, spines and extrafloral nectaries induced by some cynipidgallwasps on oaks (Stone and Schönrogge, 2003). The same gall-inducers are also able to cause the development of tissues thatshow very high investment in metabolites: in oak cynipid galls,this includes heavy investment in nucleic acids and fatty acids inthe internal nutritive tissues on which the gall-inducer feeds(Harper et al., 2004; Schönrogge et al., 2000; Stone et al., 2002)and high levels of (metabolically cheap) tannins in outer gall tis-sues as putative antiherbivore defenses (Allison and Schultz,2005; Rehill and Schultz, 2012). In several oak cynipid galls, devel-opment is associated with expression of biotin carboxylase carrierprotein (BCCP), a component of the triacylglycerol lipid synthesispathway that is associated with lipid food storage in seeds of Bras-sica napus. This led to proposal that gallwasps may cause novelexpression of seed developmental pathways in gall induction(the ‘galls as seeds’ hypothesis; Harper et al., 2004; Schönroggeet al., 2000). Oak cynipid galls also show endoreduplication in

nutritive tissues, in which chromosomes are duplicated withoutcell division, hence increasing their nucleic acid content (Harperet al., 2004; Schönrogge et al., 2000). Other hypotheses under cur-rent investigation consider that development of at least some gallsinvolves plant reproductive pathways. If this were the case, thevegetative tissues must obtain reproductive competency and somefloral and/or fruit identity traits must be expressed (Schultz, per-sonal communication; Ferreira and Isaias, 2014; Von Aderkaset al., 2005). By creating appropriate signals in gall tissues, gall-inducers may take advantage of existing suites of plant develop-mental pathways. The ‘galls as seeds’ and ‘galls as ectopic fruits’hypotheses may emerge as useful hypotheses for the identificationof candidate plant existing pathways targeted by gall-inducinginsects.

2.1.3. Patterns of diversification within galling lineagesA striking feature of gall morphologies is that while some

aspects are conserved within lineages, others can be very differenteven in closely related species. In cynipid gallwasps, for example,many species share highly similar inner nutritive tissues on whichthe larvae feed. In contrast, outer gall tissues are often very differ-ent between species, as well as highly diagnostic (Bailey et al.,2009). While we do not yet know which mechanisms underliethe development of either of these tissue types, we can hypothe-size that when revealed the genes responsible will show contrast-ing patterns of selection – stabilizing in the case of nutritivetissues, and diversifying in the case of enemy-driven diversificationof defensive structures (Bailey et al., 2009). The interactionbetween gall location and gall induction has also yet to be studiedin detail for most galler radiations. It is clear that whatever mech-anisms are used by gall-inducers, the location of the gall on theplant is not a strong constraint to gall induction: both cynipid gall-wasps and cecidomyiid gall midges provide examples of speciesradiations on a single host taxon in which there have been frequentshifts between gall locations (such as leaf, bud, fruit, flower) on thesame plant host (Cook et al., 2002; Joy and Crespi, 2007). We doknow that maternal oviposition can be highly site specific – inthe case of rose gallwasps, to particular leaves in a rose bud(Shorthouse et al., 2005b). It is thus probable that for cynipids atleast, gall traits are the result of both maternal behavior (oviposi-tion site selection, number of eggs laid) and induction stimuli pro-duced by the larva (Stone and Schönrogge, 2003). We also knowthat when forced to oviposit on atypical parts of their host-plant,at least some gallwasps can still induce galls (Folliot, 1964). Instylet-feeding gall-inducers, foundress and nymph feeding behav-ior determine gall structure and development. Patterns of diversi-fication in some stylet-feeding galling lineages suggestevolutionary responses to selection for enhanced nutrition(Crespi and Worobey, 1998; Inbar et al., 2004). There is strongempirical evidence that ovipositing females can play a role ingall-induction due to specific host selection behaviors (e.g. gall-wasps: Atkinson et al., 2002; Hessian fly: Harris et al., 2003),and/or to the delivery of gall-induction stimuli during ovipositionto initiate the process of plant tissue dedifferentiation (e.g. saw-flies: Roininen et al., 2005; cynipids: Sliva and Shorthouse, 2006).Females not only decide where to lay the eggs but also how manyeggs to lay in one place (Atkinson et al., 2002, 2003; Bailey et al.,2009). Assuming that induction stimuli contributed by each egg/larva are important for determining signal strength for inductionand sink strength for the plant, maternal oviposition behaviorsare key for the gall induction and larval food provisioning pro-cesses. We propose that the observed diversity in gall structurescan be brought about by changes in inducer behavior such aschanges in the time and/or place in which otherwise similar stim-uli are applied as well as in evolution in the effectors they use tocontrol plant development.

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2.2. Controlling plant nutritional quality

2.2.1. Improvement of nutrient concentrationsThe ability to alter the physiological state of plant tissues, par-

ticularly of the cells nearest to the feeding site, has been welldescribed for root-knot nematodes and a range of gall-inducingarthropods (Abrahamson and Weis, 1987; Bronner, 1983;Dardeau et al., 2015; Favery et al., 2016; Harper et al., 2004;Hartley, 1998; Hartley and Lawton, 1992; Larson and Whitham,1991; Mani, 1964; Nabity et al., 2013; Nasr et al., 1980;Rohfritsch, 1988; Schönrogge et al., 2000; Shorthouse, 1986;Wool et al., 1999). Nutritive tissues are kept in an active metabolicstate by the galling insect and tend to contain high levels of nutri-ents including minerals, lipids, proteins, amino acids, sugars and/orstarch (Bronner, 1992; Liu et al., 2007; Nasr et al., 1980; Saltzmannet al., 2008; Stuart et al., 2012). Enhanced nutritional value of galltissues surrounding larval chambers in closed galls results fromintense proteosynthesis (mostly structural and enzymatic pro-teins) and, at the same time, proteolysis resulting in an accumula-tion of soluble amino acids and a net increase of soluble nitrogen(Koyama et al., 2004; Nabity et al., 2013; Palct and Hassler,1967). These products may also come directly from the host-plant through the vascular system (Bronner, 1983). Additionally,significant active transport of sugars toward the insect’s feedingsite is frequently observed due, at least partially, to increasedinvertase activity (Larson and Whitham, 1991; Nabity et al.,2013; Rehill and Schultz, 2003). Glucose in excess can be trans-formed into lipids and in cynipid galls numerous lipid dropletscan be observed in nutritive cells nearest to the larval chamber(Bronner, 1983). Starch is also accumulated in nutritive storage tis-sues in cynipid galls and directly in the nutritive tissue of cecido-myiid galls due to lower amylase activity (Bronner, 1992).Finally, excess of sugars may lead to a cascade of effects includingthe up-regulation of anthocyanin synthesis explaining why somegalls develop red coloration (Arnold et al., 2012; Belhadj et al.,2008; Connor et al., 2012; Inbar et al., 2010). Examples of enhancednutritional value of transformed plant tissues include galls inducedby the cynipid wasp Neuroterus quercusbaccarum on oak and theDaktulosphaira vitifoliae phylloxera on grape (Bronner, 1992;Hartley, 1998; Kovácsné-Koncz et al., 2011; Nabity et al., 2013;Warick and Hildebrandt, 1966; Witiak, 2006).

Insects inducing open galls are also able to significantlyimprove the nutritional quality of their feeding site. Infestationby the galling insect Phloeomyzus passerinii triggers the accumula-tion of both free and protein-bound amino acids at the feeding site,the strength of the mobilizing sink being more pronounced whensoil fertilization is reduced (Dardeau et al., 2015). The Hessian flyM. destructor alters the flow of nutrients in infested plants, result-ing in improved food quality (Harris et al., 2006; Liu et al., 2007;Saltzmann et al., 2008; Stuart et al., 2012). Evidence for increasednutrient flow comes from changes in plant growth (Andersonand Harris, 2006, 2008) and the strong up-regulation of genesencoding various transporter proteins, including general sugartransporters, glucose transporters, a myo-inositol transporter,ammonium transporters, and amino acid transporters (Liu et al.,2007). Further evidence for improved food quality comes fromthe strong up-regulation of genes involved in nutrient metabolism,including (i) genes involved in carbohydrate degradation such asa-glucosidase and invertase; (ii) genes in glycolysis and the citricacid cycle such as enolase, pyruvate dehydrogenase, and oxidore-ductases; and (iii) genes involved in amino acid synthesis such asaspartate aminotransferase and methionine synthase (Liu et al.,2007). These alterations indicate that excess imported carbohy-drates are degraded and converted into intermediates for aminoacid synthesis (Liu et al., 2007). This observation is consistent withthe finding that Hessian fly attack causes a dramatic shift from

carbon-containing compounds to nitrogen-containing compounds(Zhu et al., 2008), the increase in amino acids resulting in betterfood quality for Hessian fly larvae.

In most cases, gall-inducing insects are thought to have anadvantage over non-galling insects because the gall-inducer con-centrates nutrients in the gall, making it a better food source thanequivalent ungalled plant tissue (Abrahamson and Weis, 1987;Bronner, 1983; Hartley, 1998; Hartley and Lawton, 1992; Mani,1964; Price et al., 1986, 1987; Rohfritsch and Shorthouse, 1982;Shannon and Brewer, 1980; Stone and Schönrogge, 2003; Stoneet al., 2002). However, despite the fact that several studies supportPrice et al.’s (1987) original nutrition hypothesis, gall-inducinginsects do not always benefit from high nutrient concentrationsper se (Hartley, 1998; Hartley and Lawton, 1992). In its broadestform, the nutrition hypothesis for the adaptive nature of gallsstates that galling insects control nutrient levels in galls for theirown benefit (Diamond et al., 2008; Hartley, 1998; Hartley andLawton, 1992). Thus, the thistle gall-inducing fly Urophora carduiand two cynipid galling-wasps (Neuroterus quercus-baccarum andAndricus lignicola) benefit from manipulating their host-plants ina way that prevents nitrogen levels from increasing in the galls(Gange and Nice, 1997; Hartley and Lawton, 1992). In these spe-cies, nitrogen increase reduced insect survival rates.

Among insects, manipulation of the nutritional value of planttissues is not restricted to gall-inducers; the leaf-miner caterpillarPhyllonorycter blancardella modifies nutrient profiles in minedapple-tree leaf tissues (Body, 2013; Body et al., 2013;Engelbrecht et al., 1969; Giron et al., 2007; Kaiser et al., 2010). Thismanipulation allows the insect to maintain nutrient-rich green tis-sues (sugars, free and protein-bound amino acids) (‘green-island’phenotype; Fig. 1) and to create an enhanced nutritional microen-vironment in fallen leaves, which are otherwise degrading (Body,2013; Body et al., 2013; Giron et al., 2007). Further evidence ofplant manipulation by leaf-mining insects comes from the obser-vation that radioactively labeled nutrients are preferentially trans-ported and accumulate in the feeding area, as first highlighted byMothes and coworkers in the 1960s. This suggests that insect-derived effectors create a new source-sink relationship, thus caus-ing nutrient mobilization and accumulation at attack sites to feedinsect metabolic needs (Mothes and Engelbrecht, 1961).

2.2.2. Disruption of plant defenseTo maximize their fitness, herbivorous insects also need to

manipulate and reduce plant defensive responses (Hartley, 1998;Hartley and Lawton, 1992; Price et al., 1986, 1987). Such abilityto alter plant defense expression has been demonstrated in severalgall-inducing insect lineages. Phenolic compounds, for example,are substantially lower in tissues close to the insect feeding siteand accumulate at the periphery of host-plant tissues infected byP. passerinii and six Pontania sawfly species (Abrahamson andWeis, 1987; Dardeau et al., 2014b; Nyman and Julkunen-Tiitto,2000). This result has also been shown for tannins that are essen-tially absent from nutritive tissues in cynipid wasp galls (Andricusand Trigonaspis spp.) in comparison to surrounding oak leaf tissues,though present at very high levels in outer non-nutritive tissues(Allison and Schultz, 2005; Ikai and Hijii, 2007; see also Hartley,1998). The Hessian fly M. destructor has also been shown to copewith plant defenses by hijacking phenylpropanoid pathways inwheat. Indeed, the down-regulation of five out of the seven pro-tease inhibitor genes and many genes involved in the phenyl-propanoid pathway of the host-plant leads to lowerconcentrations of chalcone, flavonoids, isoflanoids and lignin atthe insect feeding site (Liu et al., 2007). These modifications arefavorable to Hessian fly development as many of thesephenylpropanoids may be toxic to insects (Stuart et al., 2012).For example, isoorientin, an abundant flavonoid in wheat, is toxic

78 D. Giron et al. / Journal of Insect Physiology 84 (2016) 70–89

to corn earworm larvae, Helicoverpa zea (Widstrom and Snook,1998). Most researches have focused on insect strategies to coun-teract plant chemicals including excretion, sequestration anddegradation of phytotoxins. However, few examples have consid-ered the direct modulation of plant metabolism by insects as away to circumvent plant defenses at source. As a result, the mech-anisms by which host defense levels are manipulated remain to bedemonstrated in any insect gall-inducer.

Modulation of plant secondary metabolism also occurs in leaf-mining insects. For example, mining of apple leaves by the leaf-miner P. blancardella appears to prevent mobilization by the treeof its main phenolic defense compounds (dihydrochalcone and fla-vonol) (Giron et al., unpublished data). While naturally senescingcontrol leaves displayed a two-fold increase of these defensivecompounds, levels in tissues of miner-inhabited leaves remainedstable and low. The dihydrochalcone phloridzin – the predominantsecondary compound of apple leaves – deters aphids (Dreyer andJones, 1981; Schoonhoven and Derksen-Koppers, 1976) and beetles(Fulcher et al., 1998) and inhibits the biosynthesis of the insectmolting hormone ecdysone (Mitchell et al., 1993). Down-regulation of this biosynthetic pathway may therefore benefit theleaf-miner. The intimate association with the host-plant and theimportant nutritional constraints associated with the endophy-tophagous lifestyle shared by most gall-inducers and all leaf-miners may have led to selection on each of these trophic groupsto evolve strategies to disrupt plant defense.

2.2.3. Multitrophic selection on gall induction traitsThe nutritive tissues discussed above are a fundamental compo-

nent of most insect galls. The non-nutritive tissues of many of themore complex galls seem to have no direct role in galler nutrition,and hypotheses for their evolution are generally of two kinds (Priceet al., 1987; Stone and Schönrogge, 2003): (i) better protectionagainst unfavorable abiotic conditions (the microenvironmenthypothesis), (ii) improved protection from attack by natural ene-mies and infestation by pathogens (the enemy hypothesis) (Fig. 2).

Gall tissues commonly harbor inhabitants in addition to thegall-inducer, including (from the gall-inducer’s perspective) harm-ful pathogens, gallivores (herbivores that specialize in eating galltissues), specific parasitoids or predators, and commensals (suchas inquilines) (Raman et al., 2005). These trophic divisions arenot absolute, because some predators and parasitoids also feedon gall tissues (Stone et al., 2002). Gall-inducing insects not onlyneed to extract nutrients from their host-plant and counteractplant defenses but they also need to escape mortality imposedby these other gall inhabitants, as well as natural enemies (suchas insectivorous birds and small mammals) living outside the gall(Stone and Schönrogge, 2003). It follows that given the ability ofgall-inducers to manipulate plant development, they should haveevolved the ability to induce gall tissue traits that confer protectionagainst these natural enemies (Stone and Schönrogge, 2003; Baileyet al., 2009).

With the general exception of galls induced by eriophyiid mites,most arthropod-induced galls are attacked by insect and/or verte-brate natural enemies. The enemy hypothesis states that much ofthe complexity and structure of gall tissues that are not directlyassociated with inducer nutrition has evolved to exclude predatorsand parasites (Price et al., 1987; Stone and Schönrogge, 2003). Inoak cynipid galls, several gall structural traits have been shownnot only to reduce enemy attack rates within species, but also tostructure the communities of natural enemies (Bailey et al.,2009). Defense can also be chemical rather than physical. Whileinterior gall tissues on which the inducer feeds are very low insecondary plant toxins (as noted above), outer gall tissues oftencontain high levels of phenolic compounds and it has beenproposed that the galling insect derives some benefit from actively

sequestering phenolics and tannins in the surrounding gall tissuesto repel generalist herbivores (Abrahamson et al., 1991; Allisonand Schultz, 2005; Carneiro et al., 2014; Hartley, 1992; Ikai andHijii, 2007; Mang et al., 2011; Rehill and Schultz, 2012). Plant phe-nolic accumulation may also be an adaptive way to sequesterexcess carbon that has accumulated in the gall (Arnold et al., 2012).

Galls are damaged by two groups of herbivores (i) those thatspecifically target gall tissues, often deliberately killing the gall-inducer (such as the tortricid moths Pammene amygdalana and P.gallicola; Schönrogge et al., 1995; see also Abe, 1995, 1997), and(ii) those, such as some leaf-feeding caterpillars, that essentiallyconsume gall tissues accidentally, while feeding on plant parts onwhich the gall develops (Ejlersen, 1978). Gall inhabitants can alsosuffer high mortality through attack by fungi (e.g. Stone et al.,1995; Taper et al., 1986). While in some gall-inducers higher tan-nin levels are correlated with higher fitness (Rehill and Schultz,2012), there is also some evidence that aphids in high tannin gallsalso have high levels of protective antioxidants to cope with a hightannin environment. Two hypotheses have been advanced for suchhigh tannin levels. One is that tannins serve as anti-feedants toreduce attack by generalist herbivores or to protect galling insectsfrom parasitoids, while a second is that tannins serve to reducemortality imposed by fungi (Carneiro et al., 2014; Cornell, 1983;Schultz, 1992; Taper and Case, 1987; Taper et al., 1986; Wilsonand Caroll, 1997). Evidence to date shows that high tannin levelsbenefit survival of cynipid gallwasps, while there is little evidencethat high tannin levels exclude specialist gallivores, which cancause high mortality even in galls with very high tannin levels(Schönrogge et al., 1995). It remains possible that high tanninlevels may confer protection against generalist herbivores, causingthem to avoid feeding on plant parts with gall-induced high tanninlevels. There is evidence that some galling aphids can induce hostvolatiles that repel large vertebrate herbivores (Rostas et al., 2013).The important point is that in either case, gall-inducers in a rangeof taxa can induce high tannin levels in external gall tissues. Whileevidence points to a defensive selective advantage for such manip-ulation in some cases, high tannin levels could indicate the role ofthe gall as a strong sink for host-plant nutrients, rather than alwaysconferring a direct advantage (Rehill and Schultz, 2012). Highlevels of phenolics may also have indirect benefits for the plant-manipulating insects by stimulating cell growth through anincrease of auxin levels thus contributing to cell redifferentiationin galls (Carneiro et al., 2014; Oliveira et al., 2016).

Just as selection may have shaped gall traits that protect theinducer, we might also expect other gall inhabitants to haveevolved some ability to modify gall tissues to their advantage.Some cynipid inquilines can indeed significantly influence gallmorphology, producing tougher galls that are likely to reduceattack rates by their own parasitoid natural enemies (Brooks andShorthouse, 1998).

2.3. Hijacking the plant signaling network

Signaling pathways involved in plant defense against insects areknown to be complex and ultimately lead to the synthesis of manydifferent secondary metabolites (Pieterse and Dicke, 2007). Theplant hormones salicylic acid (SA), jasmonic acid (JA) and ethylene(ET) have emerged as key players in the regulation of signaling net-works involved in these responses (Van Peocke and Dicke, 2004;Von Dahl and Baldwin, 2007). Other plant hormones includingabscissic acid (ABA), gibberellins (GA), auxins (AUX) and cytokinins(CK) have also been reported to play a role in plant defensiveresponses, but are reported in fewer studies. Mimicry or manipula-tion of these phytohormones is one route towards insect control ofplant development (Giron et al., 2013). Recent reviews and papersnicely address the role played by phytohormones in gall induction

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processes and their potential role in the evolution of the gall-inducing habit (Bartlett and Connor, 2014; Giron and Glevarec,2014; Tooker and Helms, 2014).

Because many galls grow via hypertrophy and/or hyperplasia,attention has focused on phytohormones that influence cell growthand division, such as AUX and CKs (Bartlett and Connor, 2014;Davies, 2004; Straka et al., 2010; Tooker and Helms, 2014;Tooker and De Moraes, 2011a; Yamaguchi et al., 2012). Severalstudies show that gall-inducing insects such as the Hessian fly M.destructor and the goldenrod elliptical-gall caterpillar Gnori-moschema gallaesolidaginis, are able to induce indole-3-acetic acid(IAA) accumulation at the initiation site (Tooker and De Moraes,2011a, 2011b; Zhu et al., 2010, 2011). IAA is the main AUX in planttissues and is well known to cause plant cells to grow and divide;therefore, increase of IAA could play a role in generating the nutri-tive cells at larval feeding sites. Studies on the pteromalid waspTrichilogaster acacialongifoliae, the psyllid Pachypsylla celtidis andthe tephritid fly Eurosta solidaginis report high concentrations ofCKs in galls (Dorchin et al., 2009; Mapes and Davies, 2001a,2011b; Straka et al., 2010) – and particularly trans-Zeatin (tZ) inthe case of the maize orange leafhopper galler Cicadulina bipunc-tata (Tokuda et al., 2013). Selective elevation of tZ concentrationmay change the AUX:CKs ratio, critically affecting cell division pat-terns and tissue differentiation of plants and potentially playing akey role in the development and maintenance of galls (Schalleret al., 2015; Yamaguchi et al., 2012). Increased CK levels can alsofavor nutrient translocation toward the insect’s feeding site anddelay senescence by increasing the activity of extracellular inver-tase (Lara et al., 2004; Walters and McRoberts, 2006; Walterset al., 2008). Indeed, several studies show that extracellular inver-tases are induced in various plants by biologically relevant concen-trations of CKs (Ehneß and Roitsch, 1997; Godt and Roitsch, 1997).The latter work also found that hexose transporters were co-induced with extracellular invertase by elevated levels of CKs.The coordinated up-regulation by CKs of the two functionallylinked key enzymes of an apoplasmic phloem unloading pathwaymay account for the transport of nutrients to CK-treated tissueand the accumulation of nutrients at attack sites first highlightedby Mothes and coworkers in the 1960s. Other benefits gained fromhigher levels of IAA or CKs are linked to their negative crosstalkwith JA, thus lowering JA-mediated induced defenses (Erb et al.,2012; Tooker and De Moraes, 2011a, 2011b; Tooker and Helms,2014; Ueda and Kato, 1982). This may also account for lower levelsof ABA and SA frequently observed in galls (Tooker and De Moraes,2009, 2011a, 2011b; Tooker et al., 2008). Finally, interfering withCK and AUX composition within host-plant tissues could alsoexplain anthocyanin accumulation associated with some gallsand cause the striking red coloration frequently associated withgall induction (Connor et al., 2012; Inbar et al., 2010; Lewis et al.,2011; Russo, 2007; Stern et al., 2010; White, 2010).

Maybe the best example of similarities between gall-inducingand leaf-mining insects comes from data on phytohormones. Sev-eral leaf-miner species, such as P. blancardella, Stigmella argyropezaand S. argentipedella, display ‘green-islands’ (photosyntheticallyactive green patches in otherwise senescent leaves; Fig. 1) aroundmining caterpillars on yellow leaves (Body et al., 2013; Engelbrechtet al., 1969; Giron et al., 2007; Gutzwiller et al., 2015; Kaiser et al.,2010; Walters et al., 2008; Zhang et al., 2016) and show the abilityto modify phytohormonal profiles in mined leaf tissues of theirhost-plant (Body et al., 2013; Engelbrecht et al., 1969; Gironet al., 2007; Kaiser et al., 2010; Zhang et al., 2016). For example,CK concentrations of mined apple-tree tissues were shown to behigher than in unmined areas (Body et al., 2013; Giron et al.,2007; Zhang et al., 2016). Similarly to gall-inducers, the manipula-tion of the plant CK content by P. blancardella allows the insect tocreate nutrient-rich tissues favorable to its development even

when senescence occurs during the autumn (Body, 2013; Bodyet al., 2013; Giron et al., 2007). More globally, this leaf-miner leadsto strong reprogramming of plant phytohormonal balance, associ-ated with increased nutrient mobilization, inhibition of leaf senes-cence and mitigation of plant direct and indirect defense (Zhanget al., 2016).

3. Effectors used by insects to manipulate plants

Although insect-induced galls are cited as iconic examples ofextended phenotypes (Dawkins, 1982), in which plant tissue devel-opment is mainly controlled by insect genes, insect effectorsinvolved in gall induction are only starting to be characterizedand basic mechanisms of gall formation are unknown. Several linesof evidence indicate that effectors secreted by the ovipositingmother or her gall-inhabiting offspring are indeed involved in gallinduction. First, in some cases the site of gall induction differs fromthe feeding sites of the gall-inducing insects suggesting a maternaleffect (Matsukura et al., 2009; Sopow et al., 2003). Secondly, appli-cation of extracts prepared from gall-inducing insects can inducephysiological or morphological changes reminiscent of gall induc-tion (Schönrogge et al., 2000; Yamaguchi et al., 2012 and refer-ences herein). Many galling insects at least have the potential todeliver effectors precisely to plant tissues from salivary glandsusing their mouthparts, or venom glands using ovipositors(Stuart et al., 2012; Vårdal, 2006).

Initial pioneering experiments suggesting that insect effectorscould be involved in plant manipulation involved applying insectoral secretions to plants or plant products and determining theirimpact on plant defense suppression (Bartlett and Connor, 2014;Kahl et al., 2000; Lawrence et al., 2007) and the subsequent conse-quences on insect performance (Consales et al., 2012). Insectextracts were also tested for their capacity to induce morphologicalchanges in plants (e.g. Yamaguchi et al., 2012) or physiological sig-natures in suspensions of host-plant cells (Schönrogge et al., 2000).Alternatively, ablation of caterpillar salivary glands was also devel-oped to determine the role of saliva in insect–plant interactions(Musser et al., 2006). Since then, several insect herbivore effectorshave been characterized (Table 2) but their modes of action in theplant are, in most cases, unknown.

Work on insect herbivore-derived effectors involved in plantmanipulation is an emerging field (e.g. Acevedo et al., 2015;Guiguet et al., 2016; Zhao et al., 2016). Here, we present function-ally characterized effectors from insects across a range of feedingmodes, as relatively few data are so far available for leaf-minersand galling insects. We predict that –omics approaches (genomics,transcriptomics, metabolomics) will lead to rapid increases in datafor both model (Hessian fly, root knot nematode) and non-modelgall-inducers and other herbivory modes. Effectors functionallycharacterized in other plant-parasitic systems (but also in animalparasites – see Guiguet et al., 2016) may be potentially of valuein deriving mechanistic hypotheses for insect reprogrammers ofplant development.

3.1. Effectors of gall-inducing insects

To date, Hessian fly is the only gall-inducing insect for whichthe genome and salivary gland transcriptome and proteome arepublished (Chen et al., 2008, 2010a; Zhao et al., 2015). The genomeprovided evidence for hundreds of transcripts encoding candidateeffectors (Zhao et al., 2015). Genome sequencing and map-basedcloning identified four candidate effectors, the Avirulence (Avr)effector genes (Aggarwal et al., 2014; Zhao et al., 2015, 2016). Foreach of these, loss-of-function mutations permit the insect toescape the effector-triggered immunity (ETI) elicited by the corre-sponding, Avr-specific, resistance (R) gene (Harris et al., 2015).

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RNA-interference-knockdown of the gene (vH13) encoding one ofthe effectors confirmed that knockdown of that gene allows larvaeto survive on H13-resistant plants (Aggarwal et al., 2014). This sug-gests that the vH13-encoded protein is an Avr-encoded effector, aprotein that elicits ETI when secreted into plant cells that harbora corresponding R gene. The specific functions of vH6, vH9, vH13and vH24 are not currently known and functional studies of thefour Avr effector candidates are now underway. The vH24-encoded effector possibly counteracts the protein kinase activitiesinvolved in the phytohormone signaling that is associated withbiotic stress (Zhao et al., 2016) while vH6 and vH9 are suspectedto contain F box domains and leucine-rich repeats that are knownto mediate protein-protein interactions involved in the regulationof hormonal signaling, plant development and plant defense(Zhao et al., 2015).

Based on transcriptomic and proteomic analyses, a small per-centage of Hessian fly putative secreted proteins showed similarityto proteases and protease inhibitors also expressed in the gut, andto lipase-like proteins (Stuart et al., 2012). One lipase-like gene wasshown to be highly expressed in salivary glands of first instar lar-vae, which is critical in modulation of plant development. Thislipase-like gene was hypothesized to either be directly involvedin the establishment of larval feeding sites and cytological changesobserved in plant tissues during larval feeding, or to catalyze theformation of a second messenger involved in signal transductionleading to reprogramming of host cell tissues (Shukle et al.,2009). However, the majority of the salivary gland transcriptsencoded small proteins (named secreted salivary gland proteins,SSGPs), which lack sequence similarity to known proteins (Chenet al., 2004), and for which little functional annotation is possible.Similar diversity of novel secreted proteins is seen during gallinduction by cynipid wasps (Hearn and Stone , unpublished data).SSGP coding genes were classified into multigenic families on thebasis of sequence similarities (Chen et al., 2008) and certain genesappeared to be under strong positive selection suggesting theyencode effectors interacting with the host-plant (Chen et al.,2010b). SSGPs undergoing similar molecular evolution were alsoidentified in the Asian rice gall midge Orseolia oryzae, further sup-porting the possibility that some of these proteins are effectors(Chen et al., 2010b).

The recent sequencing of the M. destructor genome has allowedidentification of the full putative effector repertoire, the genomicorganization and remarkable expansion of the SSGP-encoding generepertoire (Zhao et al., 2015). Interestingly, the largest gene familyto be identified is composed of 426 SSGP-71 family members thatcode for secreted proteins containing a cyclin-like F box domain inthe N-terminus and a series of leucine-rich repeats (LRRs). Inplants, F-box-LRR proteins interact with Skp1-like proteins thatare involved in targeting proteins for degradation in the protea-some, and they play essential roles in hormonal signaling, plantdevelopment and plant defense. Zhao et al. (2015) propose thatSSGP-71 proteins act as F-box-LRRmimics, in the same way as sim-ilar proteins described in bacterial plant pathogens (Angot et al.,2006; Hicks and Galan, 2010), thereby hijacking the plant protea-some leading to modified plant physiology. In accordance with thishypothesis, a direct protein–protein interaction was observedbetween an SSGP-71 protein and wheat Skp6 protein (Zhao et al.,2015).

In depth functional approaches are now required to determinethe role played by these proteins in the gall forming process, thefact that RNA interference (RNAi) has been successfully appliedto this galling insect model opens the path to unravel effectorsinvolved in gall formation in this species (Aggarwal et al., 2014).More generally, with the progress made in RNA-seq and proteomicapproaches, it is likely that more insect putative gall-inducingeffectors will be characterized in other gall-inducing and

leaf-mining species allowing comparative approaches that maylead to the identification of convergent mechanisms involved inthese interactions. Recently, a metatranscriptome of fig flowersallowed the identification of potential candidate effectors pro-duced by fig wasps. Among the highlighted candidates are proteinssuch as icarapin, serine proteases, acid phosphatase, lipase, chy-motrypsin and metalloproteinase that have already been identifiedas venom components in other hymenopteran species (Martinsonet al., 2015). Whether these candidates actually play a role in gallformation awaits functional analyses. However, transcriptomicand proteomic analyses do not always reveal the full set of effec-tors used by insects to attack their host-plants (Guiguet et al.,2016; Zhao et al., 2016) requiring complementary tools such asgenomic and classical genetic approaches to capture the entireset of effectors used in plant-insect interactions.

3.2. Other insect derived effectors

3.2.1. Glucose oxidaseIn caterpillars of the moth H. zea, one salivary component, the

enzyme glucose oxidase (GOX), has been shown to suppress theproduction of nicotine, an inducible defensive compound producedby tobacco host-plants (Musser et al., 2002). This enzyme has sincebeen identified in the caterpillars of other Lepidoptera and inphloem-feeding insects such as aphids, suggesting suppression ofplant defenses via the secretion of this enzymemight be a commonstrategy used by different herbivores (Carolan et al., 2011;Eichenseer et al., 2010; Harmel et al., 2008; Hogenhout and Bos,2011). In certain lepidopteran–plant interactions, GOX is proposedto act on defensive response suppression by eliciting a SA burst,which then negatively impacts JA and ET signaling pathways thatregulate many induced defenses targeted against chewing herbi-vores (Diezel et al., 2009; Musser et al., 2005). GOX may also playa direct role in the inhibition of plant oxidative enzymes (i.e. lipox-igenases, peroxidases) implicated in defense against insects(Eichenseer et al., 2010). Whether this enzyme is also present insecretions of galling or leaf-mining insects remains to bedetermined.

3.2.2. ATP-hydrolyzing enzymesThree ATP-hydrolyzing enzymes (apyrase, ATP synthase and

ATPase 13A1) were detected in the salivary glands H. zea caterpil-lars, and the purified recombinant proteins were shown to be ableto suppress defensive genes regulated by JA and ET pathways intomato plants (Wu et al., 2012). These hydrolyzing enzymes pre-sumably act by degrading extracellular ATP (eATP) released bythe plant during wounding and insect feeding (Guiguet et al.,2016). Indeed, when touched or wounded, plant cells release eATPwhich acts as an early signaling molecule leading to increased con-centration in intracellular calcium, production of nitric oxide andreactive oxygen species (ROS) and phosphorylation of mitogen-activated protein kinases, ultimately leading to changes in geneexpression (Cao et al., 2014; Choi et al., 2014; Clark et al., 2014;Guiguet et al., 2016). In plants, eATP signaling impacts diverse pro-cesses including growth, development and responses to stress andpathogens (Cao et al., 2014; Clark et al., 2014). Levels of eATP arecontrolled by endogenous plant ATP-hydrolyzing enzymes, in par-ticular apyrases (Clark et al., 2014). Therefore, insect-derived apyr-ase enzymes constitute excellent candidate effectors to manipulatethese different aspects of plant physiology (Guiguet et al., 2016). Inaccordance, since their characterization in H. zea, ATP-hydrolyzingenzymes have also been identified in oral secretions of other insectherbivores including Hessian fly (Chen et al., 2008; Hattori et al.,2015). It will be of great interest to investigate whether theseenzymes are secreted by other galling or leaf-mining insects giventheir potential wide range of action on plant growth and immune

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responses (Guiguet et al., 2016). It is noteworthy that these effec-tors are also used by insect parasites in their interaction with ani-mal hosts (see Guiguet et al., 2016).

3.2.3. Calcium-binding proteinsCalcium-binding proteins have been identified in the salivary

secretions of several insect herbivores including the Hessian fly(Afshar et al., 2013; Carolan et al., 2009; Chen et al., 2008;Elzinga and Jander, 2013; Hattori et al., 2012; Nicholson et al.,2012). In most cases, the mode of action of these proteins isunknown, but one hypothesis is that they could suppress plantdefense by interfering with early Ca2+ signaling triggered by insectfeeding (Guiguet et al., 2016). In the root-knot nematode, a calreti-culin plays a role in the suppression of basal plant defenses (Faveryet al., 2016; Jaouannet et al., 2012). In the case of phloem-feedinginsects, calcium-binding proteins may play a role in limitingphloem sieve-tube occlusion, which is triggered by calcium. Inthe vetch aphid, for example, calcium-binding proteins have beenproposed to interfere with specific legume proteins (forisomes)that normally occlude sieve elements in response to calcium-triggered stress (Will et al., 2007; but see Knoblauch et al., 2014for a counter-view). This molecular interaction between salivaryproteins and calcium presumably provides these phloem-feedinginsects with a continuous flow of phloem-sap and may constituteone key mechanism at the basis of their successful colonizationof plants. Calcium-binding proteins are also used by insect para-sites in their interactions with animal hosts suggesting calcium isa central target for the disruption of cellular functions and a possi-ble route of plant manipulation (see Guiguet et al., 2016).

3.2.4. Other candidatesTranscriptomic and proteomic analyses of aphid salivary glands

or aphid saliva, combined in some cases with functional genomicapproaches, have revealed the presence of other potential effectorproteins (Elzinga and Jander, 2013; Hogenhout and Bos, 2011).Some effectors have predictable functions, such as phenoloxidaseand peroxidase involved in detoxification of phenols and glucosedehydrogenase that could interfere with JA-regulated defenseresponses (Elzinga and Jander, 2013; Will et al., 2013). Functionalstudies involving gain-of-function (i.e. in planta overexpression ofeffectors) and/or loss-of-function (i.e. RNAi of effector expression)experiments have been successfully carried out for a small numberof aphid effectors. It will certainly be possible to develop similarbioinformatics and functional pipelines (i.e. see Bos et al., 2010)in order to characterize effectors produced by galling or leaf-mining insects. Overexpression of the Myzus persicae C002 effectorin tobacco or Arabidopsis led to an increase in aphid reproduction(Bos et al., 2010; Pitino and Hogenhout, 2013), while reducingC002 gene expression by RNA silencing led to modified feedingbehavior and lethality in Acyrthosiphon pisum (Mutti et al., 2006,2008) and reduced fecundity in Myzus persicae (Pitino et al.,2011). Molecular evolutionary analyses have revealed that thisaphid-specific C002 gene is fast-evolving, suggesting that it mayplay an adaptive role in aphids and could be involved in a co-evolutionary arms race with host-plants (Ollivier et al., 2010;Pitino and Hogenhout, 2013). Other effectors with a beneficialeffect on aphid reproduction and colonization have been identified,such as the M. persicae ‘‘Progeny Increase to Overexpression” pro-teins, PIntO1 and PIntO2 (Pitino and Hogenhout, 2013) and theMacrosiphon euphorbiae Me23 effector, that which increases thisaphid’s fecundity specifically on tobacco plants but not on tomato(Atamian et al., 2013). Although it is still probably too early to gen-eralize, the aphid effectors characterized to date seem to showspecificity toward the aphid species (Pitino and Hogenhout,2013) and/or host-plant (Atamian et al., 2013), suggesting someeffectors are involved in defining host range. However, the modes

of action of these different effectors are unknown, and whetherthey are involved in suppression of plant defense or in modifica-tion of the nutritional environment is open to speculation. OneM. persicaeMp10 protein has been shown to suppress the oxidativeburst response induced by a bacterial elicitor. However, in planta,Mp10 expression causes a decrease in aphid fecundity, suggestingthat Mp10 plays the role of an avirulence-like effector in theinsect–plant interaction (Bos et al., 2010). More recently, a macro-phage migration inhibitory factor (MIF) protein has been shown tobe secreted in aphid saliva and play a crucial role for aphid sur-vival, fecundity and feeding on the host-plant. The ectopic expres-sion of aphid MIF in leaf tissues inhibits major plant immuneresponses including the expression of defense-related genes, cal-lose deposition and hypersensitive cell death (Naessens et al.,2015). Here again, MIF proteins are also used by parasites in theirinteractions with vertebrates – including nematodes, ticks and pro-tozoa – suggesting common strategies among parasites of animaland plant species to disrupt the host immune response (Naessenset al., 2015).

3.3. Insect effectors and plant growth regulators

Long standing candidate insect effectors include molecules thatare likely to regulate plant growth, defense and/or nutritional statusand many organisms are now known to be able to produce phyto-hormones. This includes gall-inducing bacteria (e.g. Stes et al.,2011, 2013), nodulating bacteria (e.g. Frugier et al., 2008), plant-associated fungi and viruses (e.g. Walters et al., 2008; Baliji et al.,2010), plant-parasitic nematodes (e.g. Favery et al., 2016; Siddiqueet al., 2015) and molluscan herbivores (Kästner et al., 2014). Highlevels of phytohormones have also been detected in the body, salivaor accessory glands of galling insects suggesting their ability to pro-duce and deliver these effectors to the plant (Bartlett and Connor,2014; Giron et al., 2013; Tooker and Helms, 2014). Several studieshave identified active auxin (IAA) in gall-inducing insects (Dorchinet al., 2009; Mapes and Davies, 2001a; Tanaka et al., 2013; Tookerand DeMoraes, 2011a, 2011b; Yamaguchi et al., 2012) and in secre-tions and frass produced by the European corn borer Ostrinia nubi-lalis (Dafoe et al., 2013). ABA and CKs have also been found inseveral galling insect species (Ohkawa, 1974; Dorchin et al., 2009;Giron et al., 2013; Mapes and Davies, 2001b; Straka et al., 2010;Tanaka et al., 2013; Tooker and De Moraes, 2011a, 2011b;Yamaguchi et al., 2012). CKs have also been found in the labialglands of several leaf-miners including the birch green miner(Engelbrecht, 1971; Engelbrecht et al., 1969) and the apple-treeleaf-miner (Body et al., 2013; Engelbrecht et al., 1969).

Collectively these studies suggest that insects could be thesources of these hormones – rather than simply manipulating theplant phytohormonal balance/signaling – allowing them to hijackplant machinery for their own benefit. However, in most studies,it is not clear whether these molecules are synthesized de novoin the insect, or derived from ingested and sequestered plant tis-sue. Recently, an IAA biosynthetic pathway was identified in thesilk worm Bombyx mori and a gall-inducing sawfly, suggesting thatinsects have the capacity to synthesize IAA in general (Suzuki et al.,2014). Moreover, two gall-inducing insects, a sawfly and the Japa-nese mugwort gall-midge Rhopalomyia yomogicola, have recentlybeen shown to synthesize IAA de novo from tryptophan and arealso suspected of synthesizing CKs (Tanaka et al., 2013;Yamaguchi et al., 2012). In vitro experiments and silencing of CK-synthesizing genes (or other key regulatory genes involved in phy-tohormone synthesis) as well as the use of plant mutants shouldhelp to determine the ability of insects to produce and deliver phy-tohormones and the role played by these effectors in plant manip-ulation – as recently demonstrated for plant-parasitic nematodes(Siddique et al., 2015).

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3.4. Microbial partners and plant manipulation in insect–plantinteractions

In the challenge to determine the exact origin of effectorsinvolved in plant manipulation by insects, we should keep in mindthat microbial partners associated with insects may be involved.Many invertebrates have intimate relationships with bacterialsymbionts and it is also well known that genes involved in AUXand CK synthesis can be of bacterial origin (Giron and Glevarec,2014). Microbial partners associated with insects, or even withplants, can be key actors in insect–plant interactions (Douglas,2013; Frago et al., 2012; Sugio et al., 2015). Many phytophagousinsects are associated with microbial symbionts that provideessential nutrients or defense against parasites (Bennett andMoran, 2015; Gibson and Hunter, 2010; Oliver et al., 2010). Thereis also growing evidence that insect-associated microbes are activeplayers in plant manipulation to the benefit of the insect host(Body et al., 2013; Kaiser et al., 2010; Su et al., 2015; Sugio et al.,2015). Ongoing developments in genomic and transcriptomic tech-nology mean that it is increasingly possible to identify the sourcegenomes of expressed genes. We can also explore which non-plant gene products are present at which stage in gall induction,and so identify candidate genes for more targeted expressionanalyses.

3.4.1. Insect symbionts and suppression of plant defenseSeveral groups have reported the involvement of insect sym-

bionts in the suppression of plant defenses. A very elegant studyshowed the Colorado potato beetle (Leptinotarsa decemlineata) torelease bacteria in its oral secretions, resulting in the activationof a plant microbial defense response. This induction of theSA-signaling pathway led, in turn, by negative cross-talk, to down-regulation of the JA-responsive anti-herbivore response. Thebeetles benefited from this manipulation by improved larvalgrowth (Chung et al., 2013). These results show that the herbivoreliterally exploits symbiotic bacteria, using them as effectorsand a ‘smoke screen’, to disrupt plant perception and evadeanti-herbivore defenses. Plant defense suppression involvinginsect-associated bacteria was also suggested in the maize–cornrootworm interaction, in which Wolbachia infection was positivelycorrelated with ability of the moth larvae to inhibit defense geneexpression in the maize (Barr et al., 2010). However, further workshowed that endosymbiont-free insects do not elicit differentmaize defense responses to Wolbachia-infected insects (Robertet al., 2013) suggesting that symbiont effects can be context-dependent. In the whitefly Bemisia tabaci, saliva of whitefliesharboring the facultative symbiont Hamiltonella defensa is able tosuppressed JA-related defenses in tomato compared to saliva fromnon-infected controls (Su et al., 2015). Putative non proteinaceouseffectors were identified in the saliva, and it will be now very inter-esting to determine their origin and exactly how H. defensa medi-ates the suppression of plant defenses in this system. Hamiltonellamay serve as a nutrient provider in whiteflies (Luan et al., 2015),illustrating the multiple ways in which a symbiont can impactoverall insect fitness. Taken together, work to date shows thatinsect-associated microorganisms can influence plant defensereactions. The role of these partners in different plant–insect inter-actions may therefore have to be evaluated in more detail. Forexample, feeding by the silverleaf whitefly has been shown toinduce SA defenses and to suppress JA responses (Zarate et al.,2007). It remains to be seen whether this ability is endogenousto the insect or is symbiont-associated. Very interestingly, it wasrecently shown in an interaction involving Scaptomyza flava leaf-miner, that the larvae can vector Pseudomonas syringae bacteriato and from feeding sites and that the larvae perform better onplants infected with P. syringae. Here the suggested mechanism is

that P. syringae acts by suppressing anti-herbivore defenses medi-ated by reactive oxygen species (Groen et al., 2016). It is also pos-sible that symbionts may be involved in the observed down-regulation of plant defenses in the leaf-miner P. blancardella.

Plant-associated symbionts (bacteria and viruses) may also playa role. They are involved in shaping the behavior of insect vectorsand could therefore potentially affect the impact of insect herbi-vores on plants (Casteel and Hansen, 2014; Casteel and Jander,2013; Casteel et al., 2012; Mauck et al., 2010; Stafford et al.,2011). Several plant pathogens are vectored by insects and canmodify plant defenses (Casteel and Hansen, 2014; Casteel et al.,2012). For example, the tomato spotted wilt virus vectored bythe western flower thrips decreased JA-dependent defenses, viathe activation of the SA-signaling pathway, to the benefit of theinsect (Abe et al., 2012). Similarly, the secretion by phy-topathogenic phytoplasma of the SAP11 protein led to down regu-lation of lipoxygenase gene expression and of JA synthesis, therebyincreasing the survival and fecundity of the leafhopper vector(Sugio et al., 2011).

3.4.2. Insect microbial symbionts and impact on plant nutritionalstatus and morphology

The first survey of bacteria associated with the gut of a gallinginsect, the Hessian fly, has recently revealed a predominance ofPseudomonas species (Bansal et al., 2014), the genomes of whichwere identified in whole genome sequencing of M. destructor(Zhao et al., 2015). It remains to be seen whether these bacteriaor other microbes associated with the insect modify host-plantnutrition and development, leading to gall induction. Some gallmidges have a symbiotic association with biotrophic fungi thatare essential for invasion of plant stems and access to vascular tis-sue, for providing larvae with highly nutritious food and for galldevelopment (Rohfritsch, 2008). The molecular mechanismsunderlying such tripartite interactions involving fungi will alsobe very interesting to uncover. Curing the apple-tree leaf-minerP. blancardella of its endosymbiotic Wolbachia bacteria resulted inthe loss of the CK-induced green-island phenotype on apple-treeleaves, and in the absence of detectable CKs in larvae comparedto non-treated controls (Body et al., 2013; Kaiser et al., 2010).These results suggest that these insects have the ability to modifythe phytohormonal profile in mined leaf tissues and to deliver CKsto the plant via their association with symbiotic bacteria (Zhanget al., 2016). So far, no other studies have linked insect symbiontsto the plant metabolic status, but it will be of great interest todetermine whether similar mechanisms are used in other plant–insect interactions.

Cornell (1983) suggested that viruses or viral proteins could beinvolved in the delivery of gall-inducing stimuli in cynipids. Amodel for this hypothesis is shown in some endoparasitoid wasps,which have incorporated polydnavirus viral coat protein genes intotheir own genomes, producing virus-like particles with which theydeliver other gene products to their lepidopteran hosts, resulting inimmune system suppression (Bézier et al., 2009; Quicke, 2015).However, no evidence for this theory yet exists in gall-inducingwasps.

3.4.3. Insect microbial symbionts and effectors encoded by horizontallytransferred genes

Recent studies have shown that microbial effectors impactinginsect–plant interactions can be encoded by genes incorporatedinto insect genomes following horizontal gene transfer (HGT) frombacteria (Boto, 2014). HGT has long been recognized as an impor-tant process governing the evolution of prokaryote genomes, andincreasing genome sequencing shows the same process to bewidespread in the evolution of eukaryote genomes, leading tothe acquisition of novel traits (Syvanen, 2012). HGT in arthropods

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appears to be particularly widespread, probably because of theclose association of many insects with endosymbionts, althoughthe adaptive significance of these horizontally transferred genesis not always evident (Hotopp et al., 2007; Kondo et al., 2002;Nikoh et al., 2008; Werren et al., 2010). In some cases, HGT are pro-posed to facilitate genome reduction of endosymbionts, sometimesleading to complex nested symbioses within phytophagous insects(Husnik et al., 2013; Sloan et al., 2014).

Two of the most convincing examples of HGT concern genesderived from non-endosymbiotic bacteria, most probably involvedin the adaptation of insects to a herbivorous lifestyle. Horizontallytransferred genes coding enzymes involved in degradation ofpolysaccharides in plant cell-walls have been identified in twophytophagous beetles (Acuña et al., 2012; Pauchet and Heckel,2013). The genome of the coffee berry borer beetle includes anexpressed and functional mannanase gene that is phylogeneticallyrelated to Bacillus genes, and is presumed to facilitate the beetle’sfeeding within the coffee berry (Acuña et al., 2012). Similarly, thegenome of the mustard leaf beetle has acquired two functionalxylanases, probably through tandem gene duplication, after HGTof a gamma-proteobacteria xylanase gene (Pauchet and Heckel,2013). Interestingly, HGT of genes encoding plant cell-wall degrad-ing enzymes have also been identified in cynipid wasps (Hearn andStone, unpublished data) and in plant parasitic nematodes (Abadet al., 2008; Danchin and Rosso, 2012; Danchin et al., 2010;Favery et al., 2016; Opperman et al., 2008), suggesting acquisitionof genes by HGT in general may play an important role intransitions to plant parasitic lifestyles, or to herbivory on specifichost-plants or tissues. Thirty candidate HGT events were recentlyidentified in the Hessian fly, again with most encoding enzymesinvolved in carbohydrate metabolism, suggesting they allow theinsect to better digest plant carbohydrates (Zhao et al., 2015). HGTsinvolved in direct plant defense suppression, modification of plantmetabolism or gall induction have so far not been identified, butgrowing research into the genomics and transcriptomics of non-model plant-herbivore interactions is increasing the chance thatsuch interactions, if present, will be found.

3.4.4. Microbes and insect plant specializationSymbiotic bacteria associated with insects could play a role in

plant adaptation, specialization or expansion of host range(Henry et al., 2013; Medina et al., 2011; Oliver et al., 2010; Tojuand Fukatsu, 2011; Tsuchida et al., 2011). Chu et al. (2013) recentlyshowed that the gut microbiome of the western corn rootwormDiabrotica virgifera virgifera is an important actor in facilitatingadaptation to soybean herbivory. Interspecific transfer ofendosymbionts between stinkbug species has also been shown toimprove the recipeinet’s performance on soybean (Hosokawaet al., 2007). In Japan, host-plant specialization of pea aphid isaffected by facultative endosymbiotic bacteria and experimentaltransfer of the facultative symbiont Regiella insecticola in vetchaphids allowed some adaptation to clover plants (Tsuchida et al.,2011). However, to date the mechanisms by which these bacteriaallow adaptation to host-plants is unknown. They could involvedirect impacts on the insect such as modifications of gut enzymeactivities (Chu et al., 2013), but manipulation of plant defensesor plant nutrients could also be involved. The pea aphid, A. pisum,encompasses multiple plant-specialized biotypes, each adapted toone or a few legume species. Facultative symbiont communitiesdiffer strongly between biotypes and plant specialization is animportant structuring factor of bacterial communities associatedwith the pea aphid complex (Gauthier et al., 2015; Peccoud et al.,2015). Here again, the role played by bacterial symbionts in plantspecialization is uncertain. Whether galling or leaf-mining insectsharbor symbiotic bacteria that could also be involved in plantadaptation remains little unexplored. Given their roles in so many

aspects of insect nutritional ecology, it seems highly likely thatsymbionts will be part of the story, some of the time (Gibson andHunter, 2010).

4. Concluding thoughts

Galls are commonly distinguished from other insect-generatedshelters such as rolled leaves and leaf mines by the fact that theyinvolve active differentiation and growth of plant tissues (Crespiet al., 1997; Shorthouse and Rohfritsch, 1992; Stone andSchönrogge, 2003; Williams, 1994). It is generally assumed thatleaf-miners do not manipulate their host-plant, but constitute anintermediate evolutionary step toward more sophisticated plantmanipulation strategies (Fig. 2) (Price et al., 1987). However, theoldest leaf-miner fossil records date to the Triassic period of theMesozoic era, 248–206 million years ago (mya) (Labandeira,2002; Labandeira et al., 1994; Rozefelds, 1988; Rozefelds andSobbe, 1987), postdating the oldest gall-inducing lineages(300 mya). Rather than seeing leaf-miners as antecedents tosophisticated plant manipulators, this review presents several linesof evidence that leaf-miners are true plant reprogrammers, in someways paralleling gall-inducers in their ability to modify source-sinkrelationships, plant morphology, nutrient availability and defensesto the insect’s advantage (Table 1). The extent to which similaritiesbetween leaf-miners and gallers reflect shared ancestral traits orconvergent evolution remains to be explored.

Feeding strategies used by plant reprogrammers presumablyevolved to face similar constraints partially imposed by the endo-phytic lifestyle shared by leaf-miners and most gall-inducers(Dempewolf et al., 2005; Stone and Schönrogge, 2003). Like insectsin closed galls, leaf-miners simultaneously live in and eat theirplants with no possibility to escape in case of inadequate food sup-ply (due to plant defensive mechanisms and/or seasonal variationsof the plant quality) or attack by natural enemies (Sinclair andHughes, 2010; Stone and Schönrogge, 2003). The main hypothesesfor the adaptive significance of gall-induction (nutrition, microcli-mate, defense) can also be applied to leaf-miners (Connor andTaverner, 1997; Sinclair and Hughes, 2010). Performance of gall-inducers and leaf-miners is also highly dependent on the oviposi-tion choice of their mother, a characteristics shared with all insectsthat have relatively sessile developmental stages.

The ‘‘plant response approach” adopted here highlights key can-didate plant functions that need to be disrupted for the insect toestablish itself and feed successfully. The development of newsequencing techniques applicable to small and/or non-model spe-cies will accelerate identification of insect effectors. A fascinatingpossibility is that insects could manipulate plant developmentthrough synthesis of plant small interfering RNAs incorporatedinto their genomes. As this field develops, access to sequences forcandidate effectors will allow comparative studies across insectradiations, and comparison with other lineages, such as plant-parasitic nematodes. Merging studies on altered patterns of plantgene expression – and related plant metabolic and morphologicalresponses – and salivary effectors produced by plant-manipulating insects will undoubtedly help to identify targetedplant functions. This should contribute to identify strategies usedby insects to manipulate their host-plant paving the way towardthe discovery of mechanisms underlying gall induction andgrowth. One of the most significant bottlenecks to this end is theidentification and characterization of plant receptors that perceivethe herbivore-specific elicitors and effectors (Acevedo et al., 2015).

More well-assembled genomes for plant-manipulators and theirhosts are now required, so that we can use the techniques of classi-cal quantitative genetics and genomics (e.g. genome-wide associa-tion studies, map-based cloning, or sliding window pairwisegenome comparisons between pairs of divergent lineages inducing

84 D. Giron et al. / Journal of Insect Physiology 84 (2016) 70–89

contrasting gall phenotypes) to identify candidate genes involved ingall phenotypes or metabolism, and then link these to genomicregions (e.g. Zhao et al., 2015, 2016). A full understanding of thetranscriptomics and metabolomics of non-galled/non-mined tis-sues on the same plants is also needed before we can highlightthe true impacts of plant reprogrammers (e.g. Body, 2013; Zhanget al., 2016). Cytological and histochemical analyses are still amongthe best ways to understand altered plant development (Harriset al., 2006, 2010; Oliveira et al., 2016) and in situ hybridization(FISH) can be used for example to show spatial and temporalexpression patterns of candidate genes targeted by plant repro-grammers. Finally, model systems that have been studied in suffi-cient detail (e.g. Body, 2013; Harris et al., 2003; Giron et al., 2013;Stone and Schönrogge, 2003) and for which molecular resourcesare available (e.g. Giron and Huguet, 2011; Whiteman et al.,2011; Zhao et al., 2015) could be used to develop RNA interferenceapproaches to knock out candidate gene expression, and so betterunderstand the impacts of specific inducers and plant genes. As col-leagues in plant pathology have already experienced (Schneiderand Collmer, 2010), developing gain- and loss-of-functionapproaches is challenging but is now required to understand theproximal mechanisms at the basis of insect-induced plant manipu-lations both in leaf-mining and gall-inducing species.

Acknowledgements

This study has been supported by the ANR – France Grant No.ANR-05-JCJC-0203-01 and the Région Centre Projects – France2010-00047141 and 2014-00094521 to D. Giron. Further supportwas provided by the National Centre of Scientific Research (CNRS)– France and the University François-Rabelais de Tours – France.Graham Stone’s work was supported by UK NERC Grant NE/J010499. We also like to thank M.O. Harris, H.M. Appel, A. Dussu-tour, P. Abad, G. Dubreuil, J.C. Schultz, M. Dicke and J. Casas forfruitful discussions.

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