Critical Reviews in Plant Sciences, 32:458–482, 2013Copyright C© Taylor & Francis Group, LLCISSN: 0735-2689 print / 1549-7836 onlineDOI: 10.1080/07352689.2013.809293

Resistance Mechanisms Against Arthropod Herbivoresin Cotton and Their Interactions with Natural Enemies

S. Hagenbucher,1 D. M. Olson,2 J. R. Ruberson,3 F. L. Wackers,4 and J. Romeis1

1Agroscope Reckenholz-Tanikon Research Station ART, Reckenholzstr. 191, 8046 Zurich, Switzerland2Crop Protection and Management Research Unit, USDA-ARS, Tifton, Georgia, USA3Department of Entomology, University of Georgia, Tifton, Georgia, USA4Lancaster Environment Centre, Lancaster University, LA1 4YQ Lancaster, United Kingdom

Table of Contents

I. INTRODUCTION ..................................................................................................................................................................................................459

II. DIRECT RESISTANCE MECHANISMS ..................................................................................................................................................461A. Morphological Traits ........................................................................................................................................................................................461

1. Impact of trichomes on herbivores .......................................................................................................................................................4622. Impact of trichomes on natural enemies .............................................................................................................................................462

B. Plant Secondary Metabolites .........................................................................................................................................................................4631. Terpenoids .....................................................................................................................................................................................................463

1.1. Terpenoid distribution .....................................................................................................................................................................4631.2. Impact of terpenoids on herbivores ............................................................................................................................................4641.3. Induction of glands and terpenoids ............................................................................................................................................466

2. Impact of other plant metabolites on herbivores ..............................................................................................................................4683. Impact of secondary plant metabolites on natural enemies .........................................................................................................469

III. INDIRECT RESISTANCE MECHANISMS .............................................................................................................................................469A. Volatiles ................................................................................................................................................................................................................469

1. Release of volatile compounds ..............................................................................................................................................................4692. Arthropod response to cotton volatiles ...............................................................................................................................................470

B. Extrafloral Nectaries ........................................................................................................................................................................................471

IV. INSECT-RESISTANT TRANSGENIC COTTON ..................................................................................................................................472

V. INFLUENCE OF ENVIRONMENTAL CONDITIONS ON COTTON ARTHROPOD RESISTANCE .....................473

VI. CONCLUSIONS .....................................................................................................................................................................................................474

ACKNOWLEDGMENTS ..............................................................................................................................................................................................475

REFERENCES ...................................................................................................................................................................................................................475

Address correspondence to J. Romeis, Agroscope ART, Reckenholzstrasse 59, 8046 Zurich, Switzerland. E-mail: joerg.romeis@agroscope.admin.ch

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ARTHROPOD HERBIVORES IN COTTON 459

Cotton plants (genus Gossypium) are grown on more than 30million hectares worldwide and are a major source of fiber. Theplants possess a wide range of direct and indirect resistance mech-anisms against herbivorous arthropods. Direct resistance mecha-nisms include morphological traits such as trichomes and a rangeof secondary metabolites. The best known insecticidal compoundsare the terpenoid gossypol and its precursors and related com-pounds. Indirect resistance mechanisms include herbivore-inducedvolatiles and extrafloral nectaries that allow plants to attract andsustain natural enemy populations. We discuss these resistancetraits of cotton, their induction by herbivores, and their impact onherbivores and natural enemies. In addition, we discuss the use ofgenetically engineered cotton plants to control pest Lepidopteraand the influence of environmental factors on the resistance traits.

Keywords Bt-cotton, Gossypium, gossypol, host plant resistance, op-timal defense theory, plant defense

I. INTRODUCTIONCotton (Gossypium spp.) is one of the most important natu-

ral sources of textile fiber, accounting for around 50% of allfibers used by humans (Matthews and Tunstall, 1994). Thegenus Gossypium belongs to the tribe Gossypiae, together withthe genera Cephalohibiscus, Cienfuegosia, Gossypioides, Ham-pea, Kokia, Lebronnecia and Thespesia (Fryxell, 1979). Amongother traits, this tribe is characterized by small lysigenous glands(glands formed through cell lysis) on the plants’ surface. Thegenus Gossypium contains around 50 species (Wendell et al.,2010) that have evolved in tropical and subtropical parts ofSouth and Central America, the Caribbean, Australasia, Africaand Oceania (Fryxell, 1979). A distinct group of six Gossyp-ium species (G. hirsutum, G. barbadense, G. tomentosum, G.lancelotum, G. darwinii and, G. mustelinum) in South and Cen-tral America are tetraploid, whereas all other cotton species arediploid (Fryxell, 1979).

Four species of Gossypium are of economic importance:G. hirsutum, G. barbadense, G. arboreum and G. herbaceum.They were most likely domesticated independently of one an-other (Fryxell, 1979). Whereas the dominant cultivated cottonspecies is G. hirsutum, the so-called upland cotton, the otherthree Gossypium species are grown only on comparably smallareas. Gossypium hirsutum accounts for at least 90% of all cottonproduced (PROTA4U 2013). In the United States, for example,it is predicted for 2013 that more than 12 million acres of G.hirsutum and 238,000 acres, or 1.9%, of G. barbadense (Pimacotton) will be planted (http://www.cotton.org). More than 33million hectares of cotton were grown in 2010, producing ca.23 million metric tons of lint and ca. 43 million metric tons ofseed (http://faostat.fao.org; James, 2011). The top three cottonproducers are China (5 million hectares, 6 million metric tonsof lint), India (11.1 million hectares, 5.7 million metric tons)and the United States (4.3 million hectares, 3.9 million metrictons) (http://faostat.fao.org; James, 2011).

Cotton harbors a rich arthropod biodiversity. Hargreaves(1948) lists more than 1300 herbivore species in cotton, in-

cluding specialists like the boll weevil, Anthonomus grandis(Coleoptera: Curculionidae) and polyphagous insects like He-liothis/Helicoverpa spp. (Lepidoptera: Noctuidae) (Matthewsand Tunstall, 1994). Additionally, cotton supports a high diver-sity of entomophagous arthropods: over 600 species of preda-tors, including dragonflies, beetles, and spiders were reportedfrom cotton fields in Arkansas (U.S.) alone (Whitcomb and Bell,1964).

Despite intensive pest management, the many arthropod her-bivores of cotton cause considerable damage (Matthews andTunstall, 1994; King et al., 1996). For example, in the UnitedStates, yield losses due to arthropod pests averaged 3.1% be-tween 2006 and 2012 (Figure 1). It needs to be taken intoaccount that even a small percentage of yield loss translatesinto a high economic loss. In 2012 total yield reduction fromarthropod pests in the United States was 2.04%, represent-ing a loss of >700,000 bales of cotton valued at >381 mil-lion US$. If management costs are included, the total eco-nomic damage was >1 billion US$ (http://www.biochemistry.msstate.edu/resources/cottoncrop.asp).

Many key pests of cotton are in the order Lepidoptera, anumber of which damage the reproductive organs, and therebycausing severe losses to fiber production. Some of the mostnotorious pest species belong to the family Noctuidae, namelythe polyphagous New World species Heliothis virescens andHelicoverpa zea and the Old World and Australasia speciesHelicoverpa armigera and Helicoverpa punctigera (the latteroccurring only in Australia). The globally distributed pink boll-worm Pectinophora gossypiella (Gelechiidae) is a highly spe-cialized cotton herbivore and a serious pest. In the United States,however, this species has been nearly eradicated (section IV).Other important pests include Earias species, such as E. vitella,E. fabia, E. insulana, E. biplaga, and E. cupreoviridis (Noc-tuidae), which also act as stem borers, and Diparopsis species(Noctuidae), which are major cotton pests in Africa. Defoliat-ing caterpillars can also contribute to yield losses, although ahealthy cotton plant can tolerate losses of up to 20% of its totalleaf area with little or no yield reduction (Sadras and Felton,2010). Foliage feeders that can be of considerable importanceinclude Spodoptera spp. (Noctuidae) and Alabama argillacea(Noctuidae).

There are few coleopteran pest species of cotton, mostlyfrom the family Curculionidae (Matthews and Tunstall, 1994;Naranjo et al., 2008), of which A. grandis is devastating to thecotton industry in the Americas. The weevil’s original distribu-tion was probably in Central America. Since the immigrationof the weevil into the United States from Mexico (ca. 1890s),accumulated costs of control and yield loss reached 15 billionUS$ (http://www.cotton.org), justifying the initiation of the bollweevil eradication program in 1978. This program succeededin eliminating the weevil as a serious pest from all previouslyinfested states of the United States except Texas, where the pro-gram is still in active phases (Hardee and Henneberry, 2004;Allen, 2008). Elsewhere in the Americas the weevil is still a

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FIG. 1. Summary of major arthropod pests of cotton in the USA. Numbers indicate the average percent yield loss for the period 2006 to 2012 across theentire United States. Bold letters denote the major production regions where the pest group primarily impacts cotton production: W, West; SW, Southwest; MS,Midsouth; SE, Southeast. Insects attacking cotton bolls (fruit) generally attack squares (flower buds) as well. Lines pointing to lint indicate that whiteflies andaphids can affect yield quality through the deposition of honeydew. Shaded boxes indicate pests, which are affected significantly by Bt-transgenic cotton. Figureredrawn from Naranjo and Luttrell (2009), which provide comparable yield loss data for the periods 1986–1995 and 1996–2005. Loss figures summarized fromthe Mississippi State University archive of Beltwide Cotton Crop Loss data: http://www.biochemistry.msstate.edu/resources/cottoncrop.asp (color figure availableonline).

serious pest. In Brazil it is estimated to cause 51 to 74 millionUS$ economic losses/year (Oliveira et al., 2013).

The order Hemiptera contains a number of important cot-ton pest species, including the two aphid species Acyrthosiphongossypii and Aphis gossypii (Aphididae). Damage is mainlycaused by the transmission of viral diseases and by contamina-tion of cotton fibers with honeydew (causing so-called stickycotton). In Africa, A. gossypii is responsible for losses in cot-ton of up to 6.5% (Matthews and Tunstall, 1994). Whiteflies(Aleyrodidae), especially Bemisia tabaci, cause similar prob-lems. The cotton stainers, Dysdercus spp. (Pyrrhocoridae), at-tack cotton and cause regional yield losses in the tropics. Severalspecies of leafhoppers (Cicadellidae), such as Amrasca devas-tans in India and Pakistan, can seriously damage cotton byinjecting toxins that interfere with photosynthesis, thus causingsymptoms known as hopperburn. In Pakistan, yield losses ofnearly 25% due to A. devastans were reported (Ahmad et al.,

2005). Plant bugs (Miridae), such as Lygus spp., and stink bugs(Pentatomidae), such as Nezara viridula, attack squares, devel-oping seeds in bolls, and the meristematic tissue. Other pestsof cotton include Orthoptera, Thysanoptera and spider mites(Acari: Tetranychidae) (Matthews and Tunstall, 1994).

Historically, cotton has been among the crops most heavilytreated with insecticides, with up to 22.5% of all global in-secticides being applied to cotton, of which about half used tomanage pest Lepidoptera (Naranjo et al., 2008; Fitt, 2008). Thispattern has changed, however, with the introduction of new tech-nologies (primarily, genetically engineered insect-resistant cul-tivars) and improved Integrated Pest Management (IPM) strate-gies that reduced insecticide use significantly (Naranjo et al.,2008; Naranjo, 2011).

Cotton plants have evolved a range of direct and indirectmechanisms that contribute to their resistance against arthropodherbivores in natural ecosystems (Figure 2). The complex of

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ARTHROPOD HERBIVORES IN COTTON 461

FIG. 2. Arthropod resistance mechanisms of cotton.

resistance mechanisms makes cotton plants an ideal model-system to study the evolution and functioning of multiple resis-tances in plant-herbivore-carnivore interactions. In this reviewwe describe the various resistance mechanisms of cotton andtheir impact on pest species and their antagonists.

II. DIRECT RESISTANCE MECHANISMSPlant arthropod resistance can be divided in two basic cate-

gories, direct and indirect resistance (Price et al., 1980; Sabeliset al., 1999). Direct resistance mechanisms can be chemical orphysical and are defined by having a direct impact on the her-bivore by negatively affecting important life-history parameterssuch as survival, development time, size, longevity, or fecundity(Zangerl and Berenbaum, 1993; Li et al., 2000; Awmack andLeather, 2002; Gols et al., 2008). Direct resistance mechanismsare often not lethal, but they frequently help protect the plant bytriggering avoidance behavior in the mobile stages of the herbi-vore. The adult herbivore may reject the plant as an ovipositionsite, while larval stages can respond by moving to tissues, whichfeature lower levels of resistance, or migrate to more suscepti-ble neighboring plants (Anderson et al., 2011). The plant maythereby deflect herbivores onto neighboring plants with which

it competes for light and nutrients (van Dam et al., 2000, 2001;Wackers et al., 2007; Anderson et al., 2011).

A. Morphological TraitsSeveral morphological features of cotton provide some de-

gree of resistance to arthropod pests. These include frego bracts,okra-shaped leaves, and trichomes. There is evidence that theopen twisted bracts of cotton with the frego bract genotype areless preferred for oviposition by A. grandis (Jenkins and Parrott,1971; Mitchell et al., 1973). Compared to normal leaf plants,cotton cultivars with spiny, okra-shaped leaves (i.e., a leaf shapewith lengthened lobes and decreased lamina expansion) tendto suffer less damage from different herbivores, including thecotton specialist P. gossypiella (Wilson et al., 1986; Naranjoand Martin, 1993). The mechanisms underlying this effect arelittle understood. Wilson et al. (1986) observed a 13% reductionin the number of P. gossypiella larvae penetrating the boll wallof okra-leaf cotton, and Naranjo and Martin (1993) found thatthe mechanism was a combination of reduced oviposition onbolls coupled with fewer eggs being deposited once a boll isselected for oviposition. In support of this, choice-experimentswith Creontiades signatus (Hemiptera: Miridae) revealed that

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females laid an average of three times more eggs on the normal-leaf than on the okra-leaf cotton (Armstrong et al., 2009). How-ever, the authors could not rule out the possibility that highertrichome density on the okra-shape leaves played a role. Com-paring different okra- and normal-leaf cotton cultivars in a fieldstudy in Arizona, Chu et al. (2002) recorded 30–40% fewerB. tabaci eggs, nymphs and adults on the okra-leaf cultivars.The authors suggested that the reduction in whitefly coloniza-tion was partly caused by less favorable micro-environmentalconditions on the okra-shaped leaves. An unfavorable micro-environment was also implicated in the increased resistance ofokra-leaf cotton to the spider mite Tetranychus urticae (Koch)(Acari: Tetranychidae) (Wilson, 1994).

1. Impact of trichomes on herbivoresTrichomes are hair-like structures on the plant surface that

can protect the plant by forming a physical barrier, or, in thecase of glandular trichomes, by producing chemical repellents,toxins or sticky substrates. Trichomes provide antixenotic andantibiotic plant resistance in numerous plant species (Peteret al., 1995; Smith, 2005) and can affect plant surface-dwellingarthropods by hampering their movement. As a consequence,arthropods need more time to move between feeding sites andincreases their exposure to natural enemies and unfavorableenvironmental conditions (e.g., Smith et al., 1975; Ramalhoet al., 1984).

The density and types of trichomes vary among Gossypiumspecies (Kosmidou-Dimitropoulou et al., 1980; Desai et al.,2008) and cultivars (Kosmidou-Dimitropoulou et al., 1980;Bryson et al., 1983; Navasero and Ramaswamy, 1991). The dif-ferent trichome types are best described for G. hirsutum, whichpossesses unicellular needle-like, single or multiple branchedtrichomes, stellate (star-like) trichomes and globose (spherical)glanded trichomes (Navasero and Ramaswamy, 1991; Bondadaand Oosterhuis, 2000). Three principal trichome phenotypesare defined: (i) glabrous = smooth leaf without trichomes; (ii)hirsute = medium length trichomes of normal density; and(iii) pilose = high density of short trichomes. In general, tri-chome density is higher on the lower leaf surface compared tothe upper surface (Navasero and Ramaswamy, 1991).

The interactions between herbivores and trichomes are com-plex and not all species are equally affected by trichomes.Several studies reported decreased feeding damage by pestspecies and/or increased yields in cotton cultivars with highpubescence (hairiness). The species affected include P. gossyp-iella (Wilson and George, 1986), the boll weevil A. gran-dis (Wannamaker, 1957; Wessling et al., 1958), Lygus line-olaris (Hemiptera: Miridae) (Meredith and Schuster, 1979),and Pseudatomoscelis seriatus (Miridae) (Walker et al., 1974).Other studies on pubescent cotton genotypes reported a reducedabundance of pest species, including A. gossypii (Kamel andElkassaby, 1965), Empoasca lybica (Hemiptera: Cicadellidae)(Butler et al., 1991), and A. devastans (Sikka et al., 1966;Batra and Gupta, 1970). In the field Spodoptera litura (Lep-

idoptera: Noctuidae) laid fewer eggs and had 60% increasedmortality on pubescent cotton cultivars compared to glabrouscultivars (Kamel, 1965). Female L. hesperus preferred piloseplants for oviposition (about 30% higher) over hirsute andglabrous leaf plants in the greenhouse, although the nymphsperformed poorer on the pilose plants (nymphal weights 37%lower) than on glabrous plants (Benedict et al., 1983). Un-der field conditions, this adverse effect resulted in significantlylower damage caused by Lygus spp. to hirsute plants whencompared to glabrous plants (Wilson and George, 1986). Sim-ilarly, increased larval mortality on highly pubescent cottonhas been observed for P. gossypiella and H. virescens, possi-bly because of impaired mobility (Smith et al., 1975; Ramalhoet al., 1984). However, these effects on herbivores are not nec-essarily caused exclusively by trichomes. For example, Navonet al. (1991) reported that H. armigera larval weight gain wasreduced by about 90% on the highly pubescent cotton culti-var Texas 172 when compared to the less pubescent cultivarAcala SJ-2. Since the trichome density was four times higheron Texas 172 compared to Acala SJ-2 trichomes appeared tobe responsible for the reduced weight gain. However, the effecton H. armigera larval weight was still present after shaving thepubescent cultivar, which points to another factor(s) playing arole.

In other instances, trichomes may also benefit herbivores.Several studies report that plants with higher trichome densitiesare more attractive as oviposition sites. This positive ovipo-sition response to trichomes has been reported for E. fabia(Mehta and Saxena, 1970), H. virescens and H. zea (Luke-fahr et al., 1971), L. lineolaris and L. hesperus (Benedict et al.,1983), and B. tabaci (Wilson and George, 1986; Gruenhagen andPerring, 2001; Ashfaq et al., 2011). Oviposition by H. virescensand H. zea was by a factor of 1.6 to 5.2 higher on hirsute cot-ton types compared to glabrous plants (Lukefahr et al., 1971).In addition, some herbivores also perform better on pubescentplants than on glabrous cotton. For example, feeding damageby Bucculatrix thurberiella (Lepidoptera: Bucculatricidae) wasincreased by 76% on hirsute cotton compared to glabrous cot-ton (Wilson and George, 1986). Similarly, the abundance ofA. gossypii nearly doubled in pubescent cotton when comparedto glabrous cotton (Weathersbee and Hardee, 1994). Tetranychusurticae developed faster on hairy than on glabrous cotton, butthis did not result in yield differences (Reddall et al., 2011). Inthe field, B. tabaci abundance increased linearly with increasingtrichome density up to about 70 trichomes per 13.7 mm2 whereabundance declined thereafter (Butler et al., 1991). These ef-fects may partly be due to reduced attack by natural enemies ashas been suggested by Gruenhagen and Perring (2001).

2. Impact of trichomes on natural enemiesAttributes that restrict the movement of an herbivore can

have the same effect on natural enemies. Trichomes canreduce the efficacy of parasitoids and predators and influ-ence the structure of predator communities (Schuster and

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ARTHROPOD HERBIVORES IN COTTON 463

Calderon, 1986; Obrycki, 1986; Hare, 2002). For example, sev-eral studies have reported a lower efficacy of Trichogrammaspp. (Hymenoptera: Trichogrammatidae) egg parasitoids onpubescent versus glabrous cotton cultivars (Romeis et al., 2005).Thus, cotton trichomes can contribute to an enemy-free spaceand thereby increase herbivore survival on pubescent plants(Gruenhagen and Perring, 2001).

Overall, trichomes appear to be an important resistance fac-tor in cotton. Their net effect, however, is often difficult to assesssince they can reduce damage by some herbivores but promoteothers and impair the efficacy of some natural enemies. Indi-vidual studies are furthermore difficult to compare due to theuse of different plant types, environmental conditions, measure-ment end-points (e.g., yield vs. herbivore abundance) and oftenundefined trichome densities.

B. Plant Secondary MetabolitesGossypium plants produce a range of compounds with insec-

ticidal properties. This includes terpenoids, flavonoids, tannins,and anthocyanins. Of those the cotton terpenoids received thegreatest attention, and are the best studied resistance mechanismof cotton.

1. TerpenoidsCotton plants produce a set of closely related terpenoids. The

best known are gossypol; hemigossypol; hemigossypolone; andheliocides H1, H2, H3, and H4 (Stipanovic et al., 1988; Altmannet al., 1990; Benson et al., 2001; Bezemer et al., 2004; Meyeret al., 2004). All of these are closely related, with H1 and H4, aswell as H2 and H3, being isomers of each other. Several othercompounds from the “family” of gossypol-related terpenoids,e.g., hemigossypol or desoxyhemigossypol, are known, but theirfunction is not well understood (Howell et al., 2000; Stipanovicet al., 1977b). The terpenoids are contained in small subepider-mal and intracellular pigment glands that are characteristic forall Gossypium species (Fryxell, 1979; Gershenzon and Croteau,1992), and in cytoplasmic granules of the root epidermis (Maceet al., 1974). The pigment glands can be found in all exter-nal tissues, including the seed, but are mainly concentrated onleaves and squares. The final root tips, however, do not containpigment glands. Similar glands are also present in other generaof the tribe Gossypiae, such as Kokia (Fryxell, 1979).

The best studied cotton terpenoid is gossypol, which is an op-tically active substance that occurs as an enantiomeric mixture of(+)-gossypol and (-)-gossypol (Jaroszewski et al., 1992a). Theratio of the enantiomers varies among plant tissues (Jaroszewskiet al., 1992b). Since gossypol is toxic to humans and non-ruminant animals (Risco and Chase, 1997), its presence in cot-ton seeds limits their use as food and feed, despite the factthat cottonseed is very rich in oil and high-quality protein (Caiet al., 2009). Consequently, one of the breeding goals in cottonhas been to select cultivars with low gossypol content resultingin cultivars that do not possess the gossypol-producing glands(McMichael, 1960; Cai et al., 2009). Seeds of these gland-

less plants contain less than 0.001% gossypol and are therebyacceptable for human and animal consumption (Fisher et al.,1988). However, as an unintended consequence, the glandlessplants became highly susceptible to arthropod pests (Bottgeret al., 1964; Jenkins et al., 1966). This provided the first indi-cation that gossypol is an important arthropod resistance com-pound. Subsequent spray experiments with purified gossypolconfirmed its insecticidal activity (Bottger et al., 1964). In re-cent years, RNAi-knockdown of δ-cadinene synthase gene(s)has been used to engineer plants that produce ultra-low gossy-pol cottonseed without affecting terpenoid production in the restof the plant (Sunilkumar et al., 2006; Palle et al., 2013; Rathoreet al., 2012). The seeds of these plants contain 0.5 μg gossy-pol/mg seed, which is less than 10% of the concentration in theparental line, Coker 312 (6 μg/mg) (Sunilkumar et al., 2006).The trait was stable under field conditions suggesting that theRNAi-based product has the potential to be commercially viable(Palle et al., 2013).

After examining wild strains of G. hirsutum that were partic-ularly resistant to H. virescens, it became clear that cotton mustpossess more insecticidal compounds than gossypol (Shaverand Lukefahr, 1971). The increased resistance found in theseplants could not be explained with gossypol alone (Shaver andLukefahr, 1971), giving the first indication that cotton producesseveral antibiotic compounds. A few years later those were iden-tified and described as hemigossypolone and the heliocides H1,H2, H3 and H4 (Gray et al., 1976; Stipanovic et al., 1977a,1978a, b). In addition, cotton plants produce a range of other ter-penoids, like caryophyllene, raimondal, ocimene, myrcene andlimonene, which also appear to be involved in plant resistance(Stipanovic et al., 1980; Elzen et al., 1985; Smith et al., 1992).

1.1. Terpenoid distribution. In general, terpenoid concen-tration, composition and distribution differ among Gossypiumspecies (Altmann et al., 1990; Stipanovic et al., 2005) and culti-vars (Stipanovic et al., 1988; Altmann et al., 1990; Hedin et al.,1991, 1992a; Wu et al., 2010) and can be dependent on en-vironmental conditions (Section V). Also within the plants aconsiderable variation in concentration and composition of ter-penoids exists (Table 1). As predicted by the optimal defensetheory (ODT) (Figure 3), terpenoid concentrations are highestin tissues that are most valuable to the plant. For example, youngleaves contain 2.5 to 5 times more terpenoids compared to ma-ture leaves (McAuslane et al., 1997; Bezemer et al., 2004; Ha-genbucher et al., 2013). Another example is the way terpenoidsare allocated to the individual parts of the squares (flower bud),where stigma and anthers contain about 10 times more gossypolthan the bracts (Hedin et al., 1992a).

Gossypol is the only terpenoid found in seeds (Stipanovicet al., 1988; Sunilkumar et al., 2006). It is the predominantterpenoid in squares and, together with hemigossypol, in roots(Hedin et al., 1984; Stipanovic et al., 1988; Khoshkoo et al.,1993; Benson et al., 2001; Bezemer et al., 2004). In contrast, themixture of terpenoids in leaves is dominated by the heliocides(esp. H1 and H2) and hemigossypolone (Stipanovic et al., 1988),

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TABLE 1Terpenoid Content (μg/g Dry Weight) in Different Cotton Plant Parts (Selected Studies). G – Gossypol,

HGQ – Hemigossypolone, HG – Hemigossypol, H1-H4 – Helicocides 1–4

Gossypium species (cultivar)Plantpart G HGQ HG H1 H2 H3 H4 Reference

G. hirsutum (Acala1517-70) Leaf 387 NA NA NA NA NA NA Meyer et al., 2004Stem 143 NA NA NA NA NA NASeed 6780 NA NA NA NA NA NA

G. hirsutum (OR19) Leaf 798 NA NA NA NA NA NA Meyer et al., 2004Stem 275 NA NA NA NA NA NASeed 10980 NA NA NA NA NA NA

G. hirsutum (Sikora 1-4/649) Leaf 929 3010 ND 2170 1090 390 1450 Benson et al., 2001Square 3620 585 ND 1850 989 351 1210Bollcoat 701 331 ND 5210 952 396 3450

G. hirsutum (Stoneville 213) Square 1400 400 NA 600 700 NA NA Hedin et al., 1992G. hirsutum (DH118) Square 1400 1500 NA 1900 1100 NA NA Hedin et al., 1992G. hirsutum (DPL 147RF) Leaf

(young)2014 6764 NA 284∗ 642∗ 642∗ 284∗ Hagenbucher et al.,

2013G. hirsutum (Stoneville 213) Leaf

(young)191 3062 NA 321 780 317 94 McAuslane and

Alborn, 1998G. herbaceum Leaf 23 825 ND 163∗ 389∗ 389∗ 163∗ Bezemer et al., 2004

Roots 10717 ND 1060 ND ND ND ND

NA = not analyzed; ND = not detected; ∗H1+H4 and H2+H3 were not analyzed separately.

while gossypol is a minor compound and hemigossypol appearsto be completely absent (Table 1). For a semiquantative overviewof the terpenoid composition in different cotton tissues see alsoStipanovic et al. (1977b).

FIG. 3. The Optimal Defense Theory.

1.2. Impact of terpenoids on herbivores. Cotton ter-penoids have direct toxic effects on a range of cotton herbi-vores. Most research has been conducted with gossypol andlepidopteran larvae. Feeding a gossypol-rich artificial diet tocaterpillars increased mortality and reduced growth rates in anumber of species, including H. virescens (Bottger and Patana,1966; Lukefahr et al., 1977), H. zea (Bottger and Patana,1966), H. armigera (Mao et al., 2007), P. gossypiella (Lukefahret al., 1977), Trichoplusia ni (Lepidoptera: Noctuidae) (Bottgerand Patana, 1966), Estigmene acrea (Lepidoptera: Arctiidae)(Bottger and Patana, 1966), E. insulana (Meisner et al., 1977c),and E. vitella (Dongre and Rahalkar, 1980). Studies with both H.zea and H. virescens did not detect any difference in the activityof the two gossypol enantiomers (Stipanovic et al., 2006, 2008).

The sensitivity to gossypol differs among herbivore species.For example, the percentage of gossypol in artificial diets thatcaused 50% mortality (LD50) varied from 0.04% (w/w) forE. acrea to 0.2% for H. zea (Bottger and Patana, 1966). Forcomparison, the gossypol content in cotton tissue is typicallyaround 0.1% of the dry mass of leaves and even higher in squaresand seeds (Table 1). Thus, gossypol and related compounds canbe a tremendous obstacle for herbivores to overcome. At lowconcentrations, however, gossypol can have a hormetic effect asdemonstrated for H. virescens and H. armigera where larvae ondiets with a low gossypol concentration (ca. around 0.0125%,w/v) performed better than those on pure diet (Stipanovic et al.,1986; Celorio-Mancera et al., 2011). Susceptibility to cottonterpenoids also tends to vary among larval instars, with olderlarvae being less sensitive (Lukefahr and Houghtaling, 1969;

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Shaver and Parrott, 1970; Belcher et al., 1983; Hedin et al.,1992a).

There is little information available on the comparativetoxicity of the different cotton terpenoids. Studies with H.virescens revealed the following order of toxicity (from mostto least toxic): gossypol > H1 > H3 > hemigossypolone > H2(Lukefahr et al., 1977; Stipanovic et al., 1977b). For P.gossypiella, however, toxicity was equal for gossypol,hemigossypolone, H1 and H2 (Lukefahr et al., 1977;Stipanovic et al., 1977b).

Effects of cotton terpenoids on caterpillars were also ob-served in planta. Larvae of H. virescens maturing on glandlesscotton had a significantly higher growth rate than those matur-ing on glanded cotton (Lukefahr et al., 1966). In contrast, thespecialist cotton-feeder A. argillacea appears to be unaffectedby terpenoids as it survived equally well on leaves of a glandedand glandless cotton cultivar and even gained more weight onthe glanded plants (Montadon et al., 1986, 1987). In general,it is difficult to relate the sensitivity of caterpillars measuredin artificial-diet studies to their performance on cotton plantsbecause larvae on plants can avoid feeding on glands or prefer-entially feed on tissue with lower terpenoid content (Lukefahrand Houghtaling, 1969; Belcher et al., 1983; Parrott et al., 1983,1990; Hedin et al., 1992a; Anderson et al., 2001).

Terpenoids also affect larval feeding behavior. Gossypol forexample is a strong feeding deterrent, as shown for Spodopteralittoralis (Lepidoptera: Noctuidae) (Meisner et al., 1977a) andneonate H. virescens that avoid consuming glands (Belcheret al., 1983; Parrott et al., 1983; Parrott, 1990; Hedin et al.,1992a). Female H. virescens typically oviposit on the terminalsof cotton plants. After hatching the larvae consume some leafmaterial and quickly move to small squares where they prefer-entially feed on the margin of the calyx, which contains about10% of the gossypol present in the anthers and petals (Hedinet al., 1992a). When larvae are older (late 2nd or 3rd instar) andless sensitive to gossypol, they burrow through the calyx, and thepetals into the anthers (Belcher et al., 1983; Parrott, 1990; Hedinet al., 1992a). Larvae of H. zea feed less, increase searching be-havior on glanded compared to glandless plants, and also preferolder leaves on glanded plants for feeding (Schmidt et al., 1988).Additionally, the rate at which larvae dropped off the plant wasbetween 4 and 5.5 times higher on glanded plants (Schmidtet al., 1988). A similar avoidance strategy has been reported forSpodoptera exigua (Lepidoptera: Noctuidae) and S. littoralislarvae that avoid gossypol-rich young leaves by migrating toolder leaves (McAuslane and Alborn, 2000; Anderson et al.,2001; Bezemer et al., 2003).

Less is known about the effect of cotton terpenoids on non-lepidopteran herbivores. Spray applications of gossypol to hostplants exerted adverse effects on L. hesperus, A. gossypii and A.grandis (Bottger et al., 1964). Compared to most lepidopteranspecies, the coleopteran A. grandis appears to be less sensitiveto gossypol (Stipanovic et al., 1977b; Moore, 1983), possiblybecause it is a specialist herbivore of Gossypium. Stipanovicet al. (1977b) reported that the weevil is insensitive to gossy-

pol and individual heliocides up to a concentration of 0.2%,which corresponds to the LD50 of the relatively insensitiveH. zea (Bottger and Pattana, 1966). In contrast, Moore et al.(1983) observed a decrease in fecundity and reduced resistanceto abiotic stresses in the weevil when this particular concentra-tion was mixed into artificial diets. Other experiments indicatedthat a broad range of arthropods from mirids (L. hesperus) toleafhoppers (Spanogonicus albofasciatus; Hemiptera: Miridae)and pill bugs (Porcellio spp.; Isopoda: Porcellionidae), sufferedless mortality and thus caused more damage on glandless com-pared to glanded cotton (Bottger et al., 1964). Du et al. (2004)found that the life span and fecundity of A. gossypii was reducedby half after feeding on a high gossypol cultivar (gossypol con-centration of 1.12%) compared to a cultivar with a gossypolcontent that was 20 times lower. In support of this, Hagenbucheret al. (2013) showed that aphid abundance was significantly re-duced on caterpillar-induced plants that had higher terpenoidlevels than undamaged plants. Studies with glanded and gland-less cotton suggest that cotton terpenoids do not affect spidermites (Agrawal and Karban, 2000). It is not clear, however, ifthe mites are unaffected by the terpenoids or simply avoid theglands due to their unique feeding patterns (Brody and Karban,1989; Agrawal and Karban, 2000).

Relatively few data exists about the mode of action and fateof gossypol after ingestion by herbivores. Binding to proteinscould explain the toxicity of gossypol to insects, since it is analkylating agent that can react with the amino group of aminoacids (Felton, 1996). For example, Meisner et al. (1977b) re-ported that gossypol inhibits amylase and protease activity inS. littoralis. When larvae had fed on cotyledons of glanded cot-ton for three days their protease activity was reduced by 73.8%and amylase activity by 79% when compared to larvae feedingon glandless cotton. A study with A. grandis showed a reversalof toxic effects of gossypol when weevils were maintained on ahigh-protein diet. While a 0.2% (w/w) gossypol concentrationin the diet reduced the fecundity of females from 9.8 eggs/dayto 4.8 eggs/day compared to a control diet (containing 0.0055%gossypol), the fecundity increased from 4.8 to 11.7 eggs/daywhen the protein concentration in the gossypol-containing dietwas raised from 3.25% to 6.5% (Moore, 1983). Some herbi-vores appear to adapt to cotton terpenoids, as has been shownfor S. exigua for which larval survival and adult fecundity onhigh terpenoid cotton cultivars increased with generations (Wuet al., 2010).

Studies with the generalist H. virescens and the specialist A.argillacea revealed that the larvae are very efficient in excretinggossypol (Montadon et al., 1987). During the last two instars,A. argillacea excreted 71% and 83% of the ingested gossypol,while the excretion levels were 45% and 68% for H. virescens(Montadon et al., 1987). Studying the metabolic fate of 14C-labeled gossypol in H. virescens, Rojas et al. (1992) observedthat most (52%) of the ingested gossypol was eliminated withthe frass, nearly 10% was metabolized to CO2, while only 24%of the gossypol or its products remained in the insect. Within thecaterpillars, radioactivity accumulated mainly in the fat body,

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466 S. HAGENBUCHER ET AL.

cuticle, and gut tissues, and was transferred to a limited extent(2.4%) to the adult stage (Rojas et al., 1992). Gossypol canthus be used as a marker to identify moths that have devel-oped on cotton (Orth et al., 2007). For the aphid A. gossypii,hemigossypolone and gossypol were found to pass through thegut and were detectable in the honeydew (Hagenbucher, 2012).A recent study with H. armigera larvae demonstrated a key roleof the P450 mono-oxygenase CYP6AE14 in the detoxificationof gossypol (Mao et al., 2007). The same enzyme appears toparticipate in the metabolism of some insecticides (Tao et al.,2012).

Cotton terpenoids are also reported to be toxic to the root-knotnematode Meloidogyne incognita (Tylenchidae: Heteroderi-dae) (Veech, 1979; Hedin et al., 1984) and to play a role inresistance against different pathogens, including Verticilliumdahlia (Hypocreales: Insertae sedis) (Mace et al., 1990), Fusar-ium oxysoporum f.sp. vasinfectum (Hypocreales: Nectriaceae)(Zhang et al., 1993), and Rhizoctonia solani (Cantharellales:Ceratobasidiaceae) (Puckhaber et al., 2002).

Several minor terpenoids with insecticidal properties arefound in cotton plants. Caryophyllene and caryophyllene ox-ide, two terpenes found together with gossypol in the glandsof G. hirsutum and other cotton species, are not lethal to H.virescens, but they cause sublethal effects on larval weight, andact synergistically with gossypol. Adding caryophyllene ox-ide to a gossypol rich diet reduced the larval weight by about30% compared to larvae exposed only to the same amount ofgossypol (Gunasena et al., 1988). In G. raimondii, the predom-inant terpenoid is a unique sesquiterpenoid called raimondal.This compound is also concentrated in the glands and is sev-eral times more toxic to H. virescens than gossypol (Stipanovicet al., 1980, 1990; Smith et al., 1992).

1.3. Induction of glands and terpenoids. For a plant, in-duction of resistance in response to pest attack is a way tooptimize the allocation of resistance, increase its efficacy andminimize associated costs (Karban and Myers, 1989). Induc-tion can be herbivore-specific, is triggered by a range of elicitorsand regulated by complex biochemical pathways. Key signals inthese processes are jasmonic acid-, salicylic acid-, and ethylene-dependent pathways, which interact with each other (Beckersand Spoel, 2006).

The cotton plant’s response to an attack by herbivores typi-cally consists of an increase in gland density as well as in theirterpenoid content in young or developing leaves (McAuslaneet al., 1997; Agrawal and Karban, 2000; Opitz et al., 2008).This induction can be elicited by a range of different above- andbelow-ground herbivores, including caterpillars (Alborn et al.,1996; McAuslane et al., 1997; Agrawal and Karban, 2000; Ha-genbucher et al., 2013), spider mites (Karban and Carey, 1984),thrips (Spence et al., 2007), wireworms (Coleoptera: Elateri-dae) (Bezemer et al., 2003, 2004), and root-knot nematodes(Veech, 1979; Khoshkhoo et al., 1993). In addition, terpenoidinduction can be elicited by infection with bacterial and fun-gal pathogens (Mace et al., 1990; Zhang et al., 1993; Abrahamet al., 1999; Howell et al., 2000; Puckhaber et al., 2002). Plants

also respond to the application of jasmonic acid (Omer et al.,2001; Opitz et al., 2008; Meszaros et al., 2011) or growth reg-ulators (Hedin and McCarty, 1991; Khoshkhoo et al., 1993).Mechanical damage can also induce terpenoid production, butthe response tends to be weaker than following herbivory orjasmonic acid treatment (Karban, 1985; Opitz et al., 2008).

Some herbivores can suppress the induction of terpenoid pro-duction. The best documented case is H. zea (Bi et al., 1997;Olson et al., 2008), where suppression of resistance inductionis most likely caused by the caterpillar’s saliva. Studies with to-bacco revealed that glucose oxidase, a protein from the saliva ofH. zea, is responsible for this suppression (Musser et al., 2005).Damage caused by H. zea caterpillars appears to induce a dis-tinct set of non-terpenoid resistance compounds in cotton, suchas several oxidases, including peroxidases and lipoxygenase, re-active oxygen species and phenolic compounds (Bi et al., 1997).In addition, feeding by H. zea causes an overall reduction in thenutritional quality of cotton plant tissue and leads to increasedlignification and strengthening of cell walls (Bi et al., 1997).Also the mealybug Phenacoccus solenopsis (Hemiptera: Pseu-dococcidae) appears to actively suppress terpenoid induction.While mealybugs perform better on cotton plants that were al-ready damaged by P. solenopsis, they were adversely affectedwhen the plants were induced with jasmonic acid (Zhang et al.,2011).

The induction of secondary metabolites in cotton is systemic,but depends on the age of the plant and the site of damage(McAuslane et al., 1997; Anderson et al., 2001; Anderson andAgrell, 2005). When the plant is attacked above-ground, noor only weak responses are induced in mature leaves, whilenewly developing leaves exhibit a strong increase in terpenoidsregardless of where the attack occurs on the plant (McAuslaneet al., 1997; Bezemer et al., 2004; Anderson and Agrell, 2005)(Figure 4). These findings fit well with the ODT, as matureleaves are less valuable to the plant compared to young leaves(Anderson and Agrell, 2005). As a consequence, caterpillarsmay migrate to the less defended, less valuable mature leavesor they may leave the induced plant altogether (McAuslaneand Alborn, 2000; Bezemer et al., 2003; Anderson and Agrell,2005). In choice experiments (McAuslane and Alborn, 1998), S.exigua larvae preferred leaves from glanded undamaged plants33 times more than leaves from glanded damaged plants. In thecase of glandless plants, undamaged leaves were only preferred2.6-fold over damaged leaves.

The systemic induction of terpenoids can mediate indirectcompetition between different species of herbivores. Recently,Hagenbucher et al. (2013) reported a 40% reduction in A.gossypii abundance on caterpillar-induced cotton plants thatcontained 2.5 times more terpenoids compared to undamagedplants. That the induction of plant resistance can mediate inter-actions between below- and above-ground herbivores was firstdescribed in cotton. The growth rate of S. exigua larvae wasreduced on G. herbaceum plants that suffered below-grounddamage by A. lineatus (Bezemer et al., 2003, 2004). Feeding byA. lineatus triggered below-ground terpenoid production as well

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FIG. 4. Patterns of systemic resistance induction in cotton plants after foliar herbivory by caterpillars (damaged leaf indicated by the arrow) or below-groundherbivory on the roots. The darker the grey, the greater the level of induction of terpenoids, volatiles and extrafloral nectaries (EFN). Question mark indicates thatno information is available about this plant part.

as above-ground. Terpenoid concentrations in immature leavesincreased by a factor of 2.3 as a response to root herbivory,while above-ground feeding by S. littoralis increased the ter-penoid concentration by a factor of 13.5 (Bezemer et al., 2004).Even though below-ground damage also caused an increase interpenoid levels of mature leaves (by a factor of 4.2), the absoluteterpenoid content of mature leaves was still about 50% lowerthan in immature leaves. In contrast, leaf damage did not triggera higher rate of terpenoid synthesis in the roots (Bezemer et al.,2003, 2004) (Figure 4). When the cotton plants were exposedto above- and below-ground herbivores at the same time, root

and foliar terpenoid content did increase but to a lesser extentthan above- or below-ground herbivory alone (Bezemer et al.,2004). Below-ground herbivory by A. lineatus also has a strongeffect on the behavior of above-ground herbivores. When givena choice between plants damaged by A. lineatus and undamagedplants, S. littoralis females deposited 81% of their eggs on theundamaged plants; the difference in the distribution of eggs wasmanifested both in the number of egg masses and in the meannumber of eggs per mass deposited (Anderson et al., 2011).

Terpenoid induction in cotton can also be triggered by differ-ent plant pathogens including F. oxysporum f.sp. vasinfectum

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468 S. HAGENBUCHER ET AL.

(Zhang et al., 1993), Rhizobium rhizogenes (Rhizobiales:Rhizobiacae) (Triplett et al., 2008), V. dahlia (Bianchini et al.,1999), and Xanthomonas spp. (Xanthomonadales: Xanthomon-adaceae) (Abraham et al., 1999). The biocontrol agent Tricho-derma virens (Hypocreales: Hypocreaceae) mediates resistanceagainst R. solani via an increase of hemigossypol and the closelyrelated desoxyhemigossypol (Howell et al., 2000; Puckhaberet al., 2002). It is not clear if nematodes can induce resistance incotton. While Olson et al. (2008) found that M. incognita doesnot induce terpenoid production, Khoshkhoo et al. (1993) foundthat damage from the same species in combination with plantgrowth regulators did induce resistance. The induction foundby Khoshkhoo et al. (1993) may have been facilitated by thegrowth regulators.

Little is known about the lag time between the onset ofdamage and the expression of the terpenoid-based resistance.Terpenoid-induction by the pathogen R. solani in the roots ofcotton seedlings was detectable within 8 h after inoculation(Rathore et al., 2012). McAuslane et al. (1997) reported thatyoung leaves expressed a high feeding deterrence for S. exigualarvae within one day after being damaged by conspecifics. El-evated terpenoid levels were still present after seven days, andthe amount of terpenoids was even higher than that recorded oneday after the initial damage. Studies with S. littoralis indicatethat terpenoid levels were still elevated 14 days after caterpillarsceased feeding (Anderson et al., 2001), while terpenoid levelswere back to constitutive levels four weeks after feeding by H.virescens larvae had stopped (Hagenbucher, 2012).

Changes in terpenoid concentration in G. hirsutum af-ter arthropod damage differ among compounds and cultivars(Table 2). Seven days after caterpillar damage, terpenoid lev-els in G. hirsutum was increased by 47–149% for hemigossy-polone, 112–124% for gossypol, 339–772% for H1, 149–883%for H4, 26–79% for H2, and 25–135% for H3 (McAuslaneet al., 1997; McAuslane and Alborn, 1998; Anderson and Agrell,2005; Opitz et al., 2008). The total terpenoid amount increasedby 89–144% (McAuslane et al., 1997; McAuslane and Alborn,1998; Anderson and Agrell, 2005; Opitz et al., 2008). The ratioof H1 + H4 to H2 + H3 changed from 0.38–0.69 to 1.28–1.48.

The ratio of hemigossypolone to all heliocides changed overtime; from 10.6: 1 at the first day to 1.3: 1 seven days afterinduction (McAuslane et al., 1997). Such a shift in terpenoidratios could be explained by a photochemical reaction betweenhemigossypolone and the volatile terpenoid ocimene, with theend product being H1 (McAuslane et al., 1997). Other factorsthat affect the induction of terpenoids include plant age and size(McAuslane et al., 1997; Agrawal and Karban, 2000; Andersonet al., 2001; Anderson and Agrell, 2005) and the site where thedamage occurs (section II B 1.3).

There is evidence that factors other than terpenoid glandsalso contribute to the increased resistance of induced plants. Forexample, Agrawal and Karban (2000) reported an equal reduc-tion in spider mite population growth on damaged glanded andglandless cotton plants, implying that terpenoids are not a majorfactor in spider mite population growth. Further evidence comesfrom studies with the glandless line WbMgl, in which the pro-duction of terpenoids like hemigossypol and desoxyhemigossy-pol is inducible by the pathogen Xanthomonas campestris pv.malvacearum (Xcm) (Davis and Essenberg 1995; Davis et al.,1996; Abraham et al., 1999).

2. Impact of other plant metabolites on herbivoresCotton produces other compounds that might contribute to

the plant’s resistance against herbivores. These include differentflavonoids (Hedin et al., 1968, 1988), some of which have beenshown to inhibit growth of lepidopteran larvae when added toartificial diet (Shaver and Lukefahr, 1969; Chan et al., 1978a;b) or in planta (Hedin et al., 1992b). Flavonoids appear to bea particularly important resistance factor in the Asian cottonspecies G. arboreum, which contains three to four times lessgossypol (Altman et al., 1990; Hedin et al., 1992b) than G.hirsutum, but is as resistant to H. virescens (Hedin et al., 1992b).There is evidence that two flavonoids absent in G. hirsutum areresponsible for this resistance: gossypetin 8-0-glucoside andgossypetin 8-0-rhamnoside (Hedin et al., 1992b).

Artificial diet studies indicate that cotton tannins can alsoadversely affect several insect species, including H. virescens

TABLE 2Relative Increase of Terpenoid Concentration (%) in Terminal Cotton Leaves, Seven Days after Spodoptera spp. Damage,

Compared to Undamaged Control Plants (Selected Studies). TT – Total Terpenoid; G – Gossypol, HGQ – Hemigossypolone,HG – Hemigossypol, H1-H4 – Helicocides 1–4

Gossypium species (cultivar) TT G HGQ H1 H2 H3 H4 Reference

G. hirsutum (Deltapine 90) 144 112 118 772 68 61 883 Anderson and Agrell, 2005G. hirsutum (Deltapine 90) 89 141 47 339 79 135 179 McAuslane et al., 1997G. hirsutum (Stoneville 213) 118 124 149 487 42 45 351 McAuslane and Alborn, 1998G. hirsutum (Deltapine

acala-90)115 NA 97 386 26 25 149 Opitz et al., 2008

G. herbaceum 103 113 114∗ 126∗ 69∗ 69∗ 126∗ Bezemer et al., 2004

∗H1+H4 and H2+H3 were not analyzed separately.

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and P. gossypiella (Waiss et al., 1977; Chan et al., 1978a, b).While there is evidence from field experiments that tanninsprovide some level of resistance to pests such as A. gossypii, B.tabaci and T. urticae (Lane and Schuster, 1980; Mansour et al.,1997) the impact on caterpillars is inconclusive (Zummo et al.,1983, 1984; Smith et al., 1992). Anthocyanins, like cyanidin-3-β-glucoside, can also contribute to insect resistance (Hedinet al., 1983).

3. Impact of secondary plant metabolites on natural enemiesThe effects of plant arthropod resistance can cascade into

higher trophic levels and have consequences for the naturalenemies of herbivores (Turlings and Benrey, 1998; Ode et al.,2006). Such effects can be negative (Turlings and Benrey, 1998)or positive; for example, when the immune system of the hostis weakened due to the ingestion of toxic compounds (Rhoades,1983; Benrey and Denno, 1997) or when the development timeof hosts is prolonged, providing predators or parasitoids with alarger window of attack (Clancy and Price, 1987; Benrey andDenno, 1997).

Little is known about the impact of cotton terpenoids on nat-ural enemies. Adverse effects on Campoletis sonorensis (Hy-menoptera: Ichneumonidae), an endoparasitoid of H. virescens,were reported when the host consumed a diet containing gossy-pol at a concentration of more than 0.1% (w/w): the parasitoidsdeveloped slower, had a lower emergence rate, and adults weresmaller (Gunasena et al., 1989). Low concentrations of gossy-pol (0.013%, w/w) in the host’s diet, however, had a positiveimpact on the weight of the adult parasitoid. In a study withPropylaea japonica (Coleoptera: Coccinellidae), the ladybirdswere fed aphids reared on one of three different cotton cultivarswith gossypol concentrations of 0.06%, 0.44% and 1.12% (Duet al., 2004). While the life-span of adult aphids was roughlyhalved on the cultivar with the highest gossypol concentrationcompared to the other cultivars, no negative impact on P. japon-ica survival and fecundity was detected. In contrast, predatorsfeeding on aphids from the high-gossypol cultivar did showa 10% shorter development time and a ca. 30% higher adultweight compared to the low gossypol cultivar. This might beexplained by an increased fatty acid content in aphids reared onthe high gossypol cultivars (Du et al., 2004). For the aphid par-asitoid Lysiphlebus japonica (Hymenoptera: Braconidae), Sunet al. (2011) did not observe a difference in parasitism whenparasitoids were provided A. gossypii reared on the same culti-vars as used by Du et al. (2004). Similar results were reportedfor Lysiphlebus testaceipes (Hymenoptera: Braconidae) whencomparing the parasitism rate for aphids feeding on caterpillar-induced or uninduced cotton despite the fact that the aphidscontained terpenoids at a concentration of 270–800 ng/mg dryweight (Hagenbucher, 2012).

These tri-trophic studies provide only limited insights intothe terpenoid sensitivity of natural enemies as the observed ef-fects could also be a consequence of other factors, e.g., an alteredquality of the herbivore as a prey or host. Direct feeding studies

to assess the impact of cotton terpenoids on natural enemies arerare and have only been conducted with gossypol. When thepredatory bug Podius nigrispinus (Hemiptera: Pentatomidae)was presented with gossypol solutions of various concentra-tions together with Tenebrio molitor (Coleoptera: Tenebrion-idae) pupae as prey throughout its entire life, adverse effectson development time, egg production and hatching rate wereobserved at certain gossypol concentrations (Evangelista Junioret al., 2011). Bugs feeding on a high gossypol diet (0.2%; w/v)lived longer (43.6 days) than those feeding on a diet containing0.1% (w/v) gossypol (39.2 days), while those on a gossypol freediet lived 48.5 days (Evangelista Junior et al., 2011). A possibleexplanation is that the binding of gossypol forces the femalesto reabsorb eggs, thus prolonging their longevity. Comparableresults were found for the two parasitoids L. testaceipes andEretmocerus eremicus (Hymenoptera: Aphelinidae) when fedsugar solutions containing different concentrations of gossypol.Female longevity was reduced by gossypol only at intermedi-ate concentrations (0.000001 to 0.0001%, w/v) while a higherconcentration (0.001%) had no effect (Hagenbucher, 2012).

Another mechanism by which induced resistance in cottoncan influence arthropod food-webs is indicated by a study withthe omnivorous thrips Frankliniella occidentalis (Thysanoptera:Thripidae). This species feeds directly on cotton tissue as wellas on other arthropods living on the plant. On plants previouslydamaged by spider mites, F. occidentalis adults reduced theiruptake of plant material by 50% and doubled the consumption ofspider mites compared to thrips on undamaged plants (Agrawalet al., 1999). This might be a way of avoiding the increasedterpenoid levels in the plant tissue resulting from mite herbivory.

III. INDIRECT RESISTANCE MECHANISMSInstead of acting on herbivores directly, indirect resistance

mechanisms act by attracting or enhancing the effectivenessof natural enemies (Price et al., 1980; Sabelis et al., 1999;Turlings and Wackers, 2004). Examples of such indirect resis-tance mechanisms include the emission of signals that are usedas host-/prey-finding cues by natural enemies, the provisioningof food, and the provisioning of shelter or oviposition substrates.

A. VolatilesPlants emit a blend of volatile compounds that typically

change in quantity and in composition after herbivore damage.As such, they can be used by herbivores and their natural ene-mies as host location cues. A number of studies have assessedthe impact of cotton volatiles for plant-arthropod interactions.

1. Release of volatile compoundsCotton produces a large number of volatiles, mainly terpenes

and lipoxygenase-derived compounds (Minyard et al., 1965,1966; Elzen et al., 1985; Loughrin et al., 1994). Similar to theterpenoids, the volatile blends differ among cotton species andcultivars (Elzen et al., 1985). Glands may play a role in stor-age and production of volatiles since glandless cotton cultivars

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470 S. HAGENBUCHER ET AL.

release fewer volatile compounds at lower concentrations whencompared to glanded cultivars (Elzen et al., 1985).

Undamaged cotton plants emit a limited range of volatilesat relatively small amounts. These mainly comprise green-leaf volatiles and terpenoid compounds (McCall et al., 1994;Loughrin et al., 1994, 1995; Rose et al., 1996, 1998). Theamount and range of emitted volatiles is typically increased inresponse to damage by lepidopteran herbivores (McCall, 1994;Rose et al., 1996). Similar to the induction of terpenoids, me-chanical damage is not as effective in inducing the release ofvolatiles as actual herbivory. This is attributed to the presenceof certain elicitors in the saliva of herbivores, like volicitin frombeet armyworm, S. exigua (Alborn et al., 1997; Rose and Tum-linson, 2004). Like other plants such as tobacco and maize, thevolatile blend emitted by cotton plants varies when damagedby different lepidopteran species (DeMoraes et al., 1998; Roseet al., 1998).

The process of volatile release by cotton following dam-age by caterpillars is not static and the blend changes overtime (Loughrin et al., 1994; Rose et al., 1996; Pare and Tum-linson, 1997a; Turlings and Wackers, 2004). A mixture oflipoxygenase-derived volatiles (green-leaf volatiles, e.g., (Z)-3-hexenal, (Z)-3-henxenyl acetate) and terpenes (e.g., α-pineneand myrcene) dominate the blend in the first hours after thedamaging event. These volatiles are either stored in the glands,synthesized by stored intermediates, or created quickly fromfatty acids by hydroperoxidation (Loughrin et al., 1994; Pareand Tumlinson, 1997a, b). The release of these volatiles is alsotriggered by mechanical damage (Rose et al., 1996). After in-duction, the early or constitutive volatiles are complemented bya broad range of acyclic terpenes that are produced de novoas a response to damage (e.g., (E)-β-ocimene, linalool, (E)-α-farnesene, (E)-β-farnesene) (Loughrin et al., 1994; McCallet al., 1994; Pare and Tumlinson, 1997a, b). The volatiles areinduced systemically (Figure 4), and the elevated release canbe detected for at least 48 h (Rose et al., 1996; Rose and Tum-linson, 2004). The volatile blend emitted from leaves does notdiffer when damage occurs on leaves or squares (Rose andTumlinson, 2004). However, the volatile blend released fromdamaged squares differs from that of damaged leaves since thelatter contains more green-leaf volatiles (Rose and Tumlinson,2004). Loughrin et al. (1995) reported that during the first 24 hafter caterpillar damage, naturalized cotton (cultivated cottonthat spread into the wild) released up to 10 times the amountof volatiles measured in commercial cultivars, indicating thatthere could be some scope for preservation and/or enhancementof innate arthropod resistance properties in breeding programs.

Studies on the plant bug L. hesperus, revealed that piercing-sucking herbivores can also systemically induce volatiles incotton (Rodriguez-Saona et al., 2002; Williams et al., 2005).Extracts from the salivary glands of L. hesperus were capable ofinducing the emission of the same volatile blend as measured forplants damaged by caterpillars (Rodriguez-Saona et al., 2002).Thus, the volatile emission appears to be triggered by an uniden-

tified elicitor present in the salivary glands that activates thesame biosynthetic pathway as caterpillar damage. OvipositingL. hesperus females also caused the emission of constitutivevolatiles, most likely as a result of mechanical damage to theglands on the plant surface when the bug punctured the leaf sur-face to insert eggs into the plant tissue (Rodriguez-Saona et al.,2002).

The influence of phloem-feeding herbivores with piercing-sucking mouthparts on the induction and release of volatilesis inconclusive. While B. tabaci could not induce the releaseof volatiles (Rodriguez-Saona et al., 2003), the cotton aphidA. gossypii could (Hegde et al., 2011). Among the compoundswere green-leaf volatiles, methyl salicylate and homoisoprenoidcompounds, but no terpenoids (Hegde et al., 2011). This meansthat the aphid volatile blend is distinctly different from thecaterpillar-induced volatiles. The mealybug P. solenopsis alsotriggers volatile release in cotton, but primarily causes the emis-sion of 3-hexen-1-ol acetate, cyclohexane, and β-caryophyllene,resulting in a volatile blend different from that released as a re-sponse to A. gossypii or caterpillar infestation (Zhang et al.,2011).

The impact of above- versus below-ground damage for cot-ton volatile induction has not been as well-studied as it hasfor other resistance mechanisms. However, Olson et al. (2008)found that below-ground damage by the nematode M. incog-nita did not affect volatile emissions above-ground, whereas thecombination of leaf–feeding by H. zea and root herbivory byM. incognita increased the concentrations of volatiles comparedto plants with leaf damage only.

Like most resistance mechanisms, volatile release in cottonis under the control of the jasmonic acid pathway and emissionof de novo produced volatiles can be induced by application ofmethyl-jasmonate. The constitutive volatiles are not affected bythis, as they are mainly released by physical damage to the tissue(Rodriguez-Sanoa et al., 2001; Zhang et al., 2011). Treatmentwith salicylic acid also triggers the release of volatiles witha blend similar to that released after P. solenopsis infestation(Zhang et al., 2011).

Under field conditions, plants are often attacked concurrentlyby a range of arthropod species. As plants react differently todifferent herbivores, such concurrent infestations can lead tosurprising results. For example, the total amount of emittedvolatiles from cotton plants infested with both B. tabaci andS. exigua larvae were decreased by about 60% compared toplants that were attacked by caterpillars only (Rodriguez-Saonaet al., 2003). A possible explanation for this effect is cross-talk between the jasmonic acid pathway (induced by caterpillarfeeding) and the salicylic acid pathway (induced by whiteflyfeeding) (Zarata et al., 2007). Another explanation could be thesuppression of induced plant resistance by B. tabaci.

2. Arthropod response to cotton volatilesMany cotton volatiles are attractive to parasitic wasps in

wind tunnel and olfactometer studies (e.g., Elzen et al., 1983;

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McCall, 1993; Rose et al., 1998), or induce a positive response inelectroantennogram recordings (e.g., Li et al., 1992; Hou et al.,1997; Gouinguene et al., 2005; Williams et al., 2008; Ngumbiet al., 2009). Thus, the volatiles that are released in response toherbivore attack might benefit the plant by attracting parasitoids.This includes parasitoids of caterpillars (e.g., McCall et al.,1993; Cortesero et al., 1997; DeMoraes et al., 1998; Rose et al.,1998; Olson et al., 2008, 2009), lepidopteran eggs (Moraeset al., 2011), aphids (Hou et al., 1997), and eggs of Lygus spp.(Manrique et al., 2005; Williams et al., 2008). Loughrin et al.(1994) reported that volatile emissions by cotton vary over thecourse of a day, but the peak coincides with the main activitywindow of caterpillar parasitoids like Cotesia marginiventris(Hymenoptera: Braconidae). In addition to the volatiles that areemitted by the cotton plant, some parasitoids also respond tovolatiles released from the feces of caterpillar hosts (Rose et al.,1997). These volatiles act as specific host-finding cues for thewasps and include unprocessed cotton volatiles and compoundsmodified by caterpillar metabolism (Alborn et al., 1996; Roseet al., 1997).

The reaction of parasitoids to cotton volatiles may vary ac-cording to their level of specialization. While both the generalistC. marginiventris and the specialist Microplitis croceipes (Hy-menoptera: Braconidae) were attracted by herbivore-damagedcotton plants, only the generalist showed a preference for freshlyand artificially (mechanical) damaged plants over undamagedplants (Rose et al., 1998). Apparently, the largely unspecificgreen leafy volatiles and constitutive compounds released fromfreshly damaged plants serve as host-finding cue for the gen-eralist (Rose et al., 1998). The Helicoverpa specialist M. cro-ceipes can differentiate between volatiles emitted as a responseto damage caused by its host H. zea and damage caused by thenon-host species S. exigua when both feed on cotton, but notwhen they feed on cowpea (McCall et al., 1993). Similarly theH. virescens parasitoid Toxoneuron nigriceps (Hymenoptera:Braconidae) can distinguish cotton volatiles induced by its hostfrom those released in response to damage by the non-hostspecies H. zea (De Moraes et al., 1998).

Few studies dealt with the response of predators to cottonvolatiles. Hegde et al. (2011) reported that adult Chrysoperla lu-casina (Neuroptera: Chrysopidae) showed antennal responses toodors released by aphid-infested cotton plants. Green lacewings,Chrysoperla carnea (Neuroptera: Chrysopidae) and the preda-tory beetle Collops vittatus (Coleoptera: Melyridae) were at-tracted to insect traps distributed in cotton fields that containedcaryophyllene and its derivates, compounds emitted in the bou-quet of undamaged and damaged cotton plants (Flint et al., 1979,1981).

Herbivores also detect and respond to cotton volatiles.Olfactometer studies revealed that A. gossypii prefers thevolatile blend released from uninfested cotton plants over thatfrom aphid-infested plants (Hegde et al., 2011). Similarly,H. armigera is capable of detecting several typical volatile com-pounds of cotton that are induced after herbivore damage; e.g.,

the terpene α-pinene (Yan et al., 2004). Spodoptera littoralisfemales were reported to preferentially lay eggs on undamagedcotton plants, as compared to plants damaged by conspecificlarvae (Anderson and Alborn, 1999) or wireworms (Andersonet al., 2011). This oviposition preference for undamaged plantswas only visible in older plants (6 leaf stage) and was even re-versed in young plants (3-4 leaf stage) (Anderson and Alborn,1999). The role of plant volatile cues in those experiments wasdemonstrated recently (Zakir et al., 2013). The recognition andavoidance of damaged plants allows herbivores to avoid infe-rior host plants and competition (Anderson and Alborn, 1999;Anderson et al., 2011). The plant benefits by receiving fewerand smaller herbivore egg batches and by deflecting the herbi-vore to nearby plants competing for light, water and nutrients(Anderson et al., 2011).

B. Extrafloral NectariesCotton provides sugars to attract and sustain natural enemies

in the form of extrafloral nectaries (EFN). Such nectaries arefound in all Gossypium species, except G. tomentosum and G.gossypioides (Fryxell, 1979). Nectaries are located on the lowersurface of the leaves, on one or more of the principal veins, andalso on the bracts (Fryxell, 1979). The most common predatorsthat attend extrafloral nectar are ants (Wackers and Bonifay,2004). Indirect evidence that EFNs have evolved to attract antscomes from the two species lacking extrafloral nectaries (Wack-ers and Bonifay, 2004). Gossypium tomentosum is endemic toHawaii (Fryxell, 1979), an area of the world devoid of ants untilrecently (Wilson, 1996), while G. gossypioides is atypical inthat it grows at high altitudes where ant activity is suppressed(Wackers and Bonifay, 2004). Rudgers (2002) and Rudgerset al. (2003) describe a mutualistic association between the wildG. thurberi and the ant Forelius pruinosus (Hymenoptera:Formicidae). The ants reduce herbivory by preying on cater-pillars or by disturbing their feeding activities and consequentlyenhance seed production. There is some evidence that cottonplants with EFNs support a larger beneficial arthropod com-munity (Schuster et al., 1976; Hennenberry et al., 1977; Adjei-Maafo and Wilson, 1983) and can have positive effects on differ-ent life-history parameters of nectar feeding predators and par-asitoids (Lindgren and Lukefahr, 1977; Schuster and Calderon,1986). Under agricultural conditions, the direct correlation be-tween the availability of EFN and the abundance of predators,however, may be masked by confounding factors like abundanceof prey, weed coverage, and plot size. Most importantly, agro-nomic practices, such as tillage destroy ant nests and the use ofpesticides eliminates many (predatory) arthropods (Naranjo andGibson, 1996). When predator numbers are restricted, it is likelythat the value of EFNs as a resistance factor is also limited.

The production of extrafloral nectar appears to follow theODT. Constitutive foliar nectar production by undamagedG. hirsutum plants ranged between 79 and 204 μg per plantper day, while nectaries on the bracts exceed foliar nectar secre-tion by a factor of between 80 and 110 (Wackers and Bonifay,

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472 S. HAGENBUCHER ET AL.

2004). The production of bracteal nectar shows a distinct peakon the day of anthesis (where an average 11.7 mg of nectar wasproduced per day), followed by a prolonged secretion duringfruit maturation (Wackers and Bonifay, 2004). Valuable repro-ductive tissue is thus considerably better resourced for defendersthan the less valuable vegetative tissue. Similar to terpenoids andvolatiles, foliar nectar production can be induced by above- andbelow-ground herbivore damage (Wackers and Bezemer, 2003;Wackers and Bonifay, 2004). After induction by S. littoralisfeeding, foliar nectar production was enhanced by a factor of12, primarily at the damaged leaf (Wackers et al., 2001). Thisallows the plant to “guide” ants and other predators to the site ofattack. In addition, there was a limited systemic response: nec-tar production also showed some increase on adjacent youngerleaves (Wackers et al., 2001) (Figure 4). Four days after removalof the herbivore, nectar production decreased to pre-treatmentlevels (Wackers et al., 2001). Damage of roots by wireworms(A. lineatus) also increased foliar nectar production. However,in this instance the induced nectar is evenly distributed over theplant (Wackers and Bezemer, 2003) (Figure 4).

In contrast to the foliar nectar, the bracteal nectar shows highlevels of constitutive production, and is not further induced byherbivory. This indicates that the cotton plant has evolved twoseparate strategies for nectar-mediated resistance in its foliageand its reproductive organs. The high level of bracteal nectarsecreted irrespective of herbivore damage represents a preven-tative strategy, recruiting ants as a standing army that will helpto prevent damage to valuable flowers and squares. In contrast,the low levels of constitutive foliar nectar production, combinedwith the high level of nectar induction, represents a curativestrategy, recruiting ants once herbivores initiate feeding (Wack-ers and Bonifay, 2004).

Similar to natural enemies, herbivores may also utilize cottonextrafloral nectar as a food source. Several studies comparingherbivore pressure between nectariless cotton cultivars and cul-tivars producing extrafloral nectar have shown larger herbivorepopulations on the latter (Lukefahr and Rhyne, 1960; Lukefahret al., 1965; Schuster et al., 1976; Henneberry et al., 1977;Adjei–Maafo and Wilson, 1983; Flint et al., 1988; Scott et al.,1988). However, these studies were conducted in agriculturalfields that are characterized by impoverished ant populations.Studies done in natural habitats typically show the reverse pat-tern (Rudgers 2002, 2003).

IV. INSECT-RESISTANT TRANSGENIC COTTONCotton cultivars with improved arthropod resistance are con-

tinuously developed through methods of conventional breed-ing and genetic engineering (Hardee and Henneberry, 2004;Naranjo et al., 2008; Naranjo, 2011). After the success of the bollweevil eradication program in the USA (USDA-Aphis, 2006),caterpillars, especially the so called budworm-bollworm com-plex of Helicoverpa spp., H. virescens and P. gossypiella, be-came the most serious cotton pests. This pest complex has been

largely controlled by resistant cotton cultivars derived throughgenetic engineering (Naranjo and Luttrell, 2009; Naranjo 2011).Plants that express Cry toxins derived from the bacterium Bacil-lus thuringiensis Berliner (Bt) with high specificity to lepi-dopteran species, were produced (Schnepf et al., 1998; Bravoand Soberon, 2008). The first generation of Bt cotton expressedthe single protein Cry1Ac. Second generation Bt cotton plantsproduce a combination of two insecticidal proteins, which in-creases their efficacy against the lepidopteran pest complex andhelps to delay development of resistance in the target pests. Themost common dual-gene Bt cotton plants grown today are Boll-gard II R© plants, which produce Cry1Ac and Cry2Ab2 (Naranjoet al., 2008).

In 2012, Bt-transgenic cotton cultivars were grown on a totalof 22.5 million hectares worldwide (James, 2012). The high-est adoption rate was reached in Australia (95% of the cottonarea), followed by India (93%), Pakistan (82%), China (80%),the United States (88%), and Burkina Faso (58%) (James, 2012;USDA Economic Research Service, http://www.ers.usda.gov/).Using comparative farm-level data in adopting countries,Brookes and Barfoot (2011) estimated that the use of Bt cottonled to a global reduction in the volume of insecticidal activeingredients applied by 153 million kilograms globally between1996 and 2009, a 22% change over the 14-year period. In someworld areas, however, savings in insecticide use were signifi-cantly higher (Fitt, 2008; Qaim et al., 2009; Krishna and Qaim,2012).

The high efficacy of Bt cotton and high adoption levels havecaused area-wide suppression in the populations of some impor-tant cotton pests. For example, Bt cotton has been an importantfactor in the successful eradication program for P. gossypiellain the southwestern United States (Carriere et al., 2003; An-tilla and Liesner, 2008; Naranjo and Ellsworth, 2010). In sixprovinces of China, Wu et al. (2008) observed a linear declinein populations of H. armigera on cotton associated with increas-ing yields since the adoption of Bt cotton in 1997. This patternof decline was also observed in many other crops in the regionaffected by this polyphagous pest. In addition, the high speci-ficity of the expressed Cry proteins together with the decline ininsecticide use had significant positive effects on the arthropodfauna in cotton fields (Romeis et al., 2006; Marvier et al., 2007;Wolfenbarger et al., 2008; Naranjo et al., 2008; Naranjo, 2011;Lu et al., 2012). This can benefit biological control by creatingan environment, in which natural enemies can flourish (Romeiset al. 2008; Naranjo 2011; Lu et al., 2012). Lu et al. (2012)demonstrated a marked increase in the abundance of three typesof generalist predators (ladybirds, lacewings and spiders) anddecreased aphid abundance in six provinces in China. This wascaused by the adoption of Bt cotton and the reduced insecticidesprays for the 1990 to 2010 period. The reduction in insecticideuse, however, can also lead to the development of non-targetpests. An example is the increased crop damage attributed toplant and stink bugs in Bt cotton (Naranjo et al., 2008; Luet al., 2008, 2010). Additional factors that may contribute to the

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development of non-target pests in Bt cotton include the releasefrom direct resource competition by caterpillars (Whitehouseet al., 2007; Zeilinger et al., 2011) or from indirect competition,as Bt cotton plants produce less terpenoids due to lower feedingdamage from caterpillars (Hagenbucher et al., 2013).

New arthropod-resistant cotton cultivars are continuously de-veloped using tools of genetic engineering. Cotton plants ex-pressing other Cry proteins or vegetative insecticidal proteins(VIP) from B. thuringiensis have already been commercializedor will reach the market soon (Whitehouse et al., 2007; Naranjoet al., 2008; CERA, 2012). A recent study has also shown thepotential of controlling emerging hemipteran pests with a newlyidentified B. thuringiensis crystal protein (TIC807) in the future(Baum et al., 2012).

A novel strategy to control cotton pests adapts RNA inter-ference (RNAi) techniques (Price and Gatehouse, 2008). Thiscomprises the use of genetically engineered plants that ex-press double-stranded RNA (dsRNA) of the cytochrome P450CYP6AE14 of H. armigera. This enzyme plays an importantrole in the detoxification of gossypol (Luo et al., 2001). Stud-ies by Mao et al. (2007) showed that caterpillars feeding onthese plants are more susceptible to gossypol, due to silencingof the cytochrome caused by the ingested dsRNA. Thereby thecaterpillars lose an important part of their detoxification system.

The novel insecticidal traits added to cotton will interactwith the plant’s natural arthropod resistance mechanisms. Un-derstanding these interactions is important because it likely af-fects the sustainability of genetic engineering as a pest controltool. For example, the arthropod resistance compounds in cot-ton could help to increase the durability of Bt cotton. A numberof studies have shown that the efficacy of the Cry proteins isenhanced by cotton insecticidal compounds. For example, theresistance of Bt (Cry1Ab) cotton to H. virescens was increasedwhen crossed with a high-terpenoid cultivar (Sachs et al., 1996).Furthermore, the effectiveness of Bt cotton against H. armigerawas increased by a factor of 4 to 15 when plants had previ-ously been induced by caterpillar damage (Olsen et al., 2005a).A similar effect was reported by Meszaros et al. (2011) forSpodoptera frugiperda (Lepidoptera: Noctuidae) when Bt cot-ton plants were induced by the application of jasmonic acid.In contrast are the results of Olson and Daly (2000), were ex-periments revealed that the age of plants could influence thetoxicity of Cry1Ac. When leaves of fruiting cotton plants andpresquare cotton were incorporated in various diets containingCry1Ac protein it was found that leaves from the older, fruit-ing cotton plants reduced the toxicity of the toxin by 14 to726-fold (Olsen and Daly, 2000). The authors suggested thattannins were a main reason for this effect by either limitingthe availability of the Cry protein or by reducing its toxicity,a fact suggested earlier for example by Navon et al. (1993).Cotton terpenoids may also play a role in delaying the evolu-tion of resistance against certain Cry proteins in the target pests.Resistance is often linked to fitness costs, which can result inincreased susceptibility against cotton terpenoids. This is one

possible reason for the observation that Cry1Ac-resistant Lep-idoptera strains suffer higher mortality on Cry1Ac-expressingcotton compared to Cry1Ac-containing artificial diets (Tabash-nik et al., 2003; Bird and Akhurst, 2004; Anilkumar et al.,2009; Gassmann et al., 2009). For example, Cry1Ac-resistant P.gossypiella larvae were found to be more vulnerable to gossypolthan Cry1Ac-susceptible larvae (Carriere et al., 2004; Williamset al., 2011). The fitness cost related to the Cry1Ac resistancewas displayed in reduced larval growth on an artificial diet. Theresistant larvae gained less weight on the control diet than thesusceptible larvae (5.8% weight reduction), suggesting a resis-tance costs even in the absence of gossypol. This resistance costwas magnified in the presence of gossypol leading to an evenlower weight gain (12.9% weight reduction) than on the controldiet (Williams et al., 2011). In contrast, susceptibility to gossy-pol did not differ between Cry1Ac-resistant and susceptiblelarvae of H. zea (Anilkumar et al., 2009). It has been hypothe-sized that the difference in gossypol sensitivity between the twolepidopteran species is due to differences in the mechanisms ofCry1Ac-resistance (Gassmann et al., 2009). In P. gossypiella,resistance to Cry1Ac is caused by mutations in genes encodingcadherin proteins that bind the Bt toxin (Morin et al., 2003;Fabrick et al., 2011). It has been postulated that those mutationsmay disturb the integrity of the midgut membrane and increaseits permeability for gossypol, thereby amplifying its toxic effect(Carriere et al., 2004, 2006). Indeed, P. gossypiella strains car-rying these cadherin alleles contain 4.4 (heterozygote strain) or13.6 (homozygote strain) times more gossypol after feeding ongossypol-rich artificial diet than Cry1Ac-susceptible larvae witha normal cadherin genotype (Williams et al., 2011). In contrast,Cry1Ac resistance in H. zea appears to be due to altered pro-teolysis, and thus gossypol uptake is not affected (Anilkumaret al., 2008, 2009).

V. INFLUENCE OF ENVIRONMENTAL CONDITIONSON COTTON ARTHROPOD RESISTANCE

Numerous abiotic factors influence arthropod resistance incotton. For example, Stipanovic et al. (1988) found that ter-penoid concentrations in leaves of 14 cotton cultivars differedamong plants grown in five different locations. The impact ofenvironmental factors on cotton arthropod resistance, however,has mainly been investigated for Bt-transgenic cotton cultivars.Temperature appears to be particularly important. For example,Chen et al. (2005a) provide some evidence for reduced Cry pro-tein levels in cotton grown at high temperatures. Olsen et al.(2005a) reported that Bt cotton exposed to cool temperaturesin the early part of the growing season suffered increased H.armigera damage. Since the expression of the cry1Ac transgeneremained unaffected by temperature in that study, the factorsresponsible for the observed effects were likely due to someother plant compounds and/or their interactions.

Herbivore-induced cotton plants grown under conditionswith limited available water (125 ml/day) were preferred

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feeding sites for S. exigua when compared to “normal-water”plants (500 ml/day); in a choice experiment larvae consumedabout 9 times more leaf tissue from water-stressed plants, in-dicating reduced terpenoid production in these plants (Olsonet al., 2009). The parasitoid M. croceipes did not prefer water-stressed plants: in a wind-tunnel assay only about 10% of allparasitoids landed on the water-stressed plants while the rest fa-vored the unstressed plants (Olson et al., 2009). This preferenceof parasitoids for unstressed plants was explained by a reduc-tion of emitted volatiles in water-stressed plants. A sufficientsupply of water seems to be crucial for a high gossypol contentin cotton plants as irrigation was found to increase the gossypolcontent in cotton seeds by 21% (Pettigrew and Dowd, 2011).Even water-logging (an overabundance of water) had a positiveimpact on terpenoids as it increased the foliage terpenoid con-centration by as much as 100% (Luo et al., 2008). Bt toxinsare also influenced by water availability. Water deficiency wasreported to decrease Cry protein levels in Bt cotton (Benedictet al., 1996). These findings fit with an earlier report that underdrought and heat conditions, cotton plants produce heat shockproteins at the expense of normal protein synthesis (Burke et al.,1985). However, water-logging reduced Cry1Ac toxin levels byas much as much 38–50%, depending on cultivar and durationof exposure (Luo et al., 2008).

Another environmental factor that could influence cotton her-bivore resistance is the salt content of the soil. While a highsalinity (up to 0.05% NaCl) increased the foliar terpenoid con-centration by 20–59%, it had the opposite effect on Cry1Aclevels, which were reduced by 11.3–22.4%. If salt-stress wascombined with waterlogging, Cry1Ac concentrations were re-duced by as much as 72% (Luo et al., 2008). The exact effects ofthis stress, however, depend on the cultivar used and the durationof exposure.

The plants’ C:N ratio can be affected by the extensive useof nitrogen fertilizers. On the one hand, high nitrogen levelsincrease the total protein in plants (Baker et al., 1980), whichcan also increase the Bt protein levels (Pettigrew and Adamczyk,2006). On the other hand, application of nitrogen fertilizers hasbeen found to reduce the induction of terpenoids, jasmonic acidand volatiles, in response to herbivory damage (Chen et al.,2008; Olson et al., 2009). Consequently, both under- and over-fertilized plants were preferred feeding sites for S. exigua larvaeand less attractive to the parasitoid M. croceipes (Olson et al.,2009).

The rising CO2 levels in our atmosphere will influence thedefensive chemistry of Gossypium species. Elevated CO2 levelsincrease the photosynthetic rate in plants, thereby increasinggrowth rate and the carbon to nitrogen (C:N) ratio (Chen et al.,2010). According to the carbon–nutrient balance hypothesis(Bryant et al., 1983), the allocation of resistance resources willlikewise shift to substances that do not contain nitrogen. Thishas been reported from studies with Bt cotton where elevatedCO2 levels resulted in a significant decrease in the Cry protein,which contains nitrogen. In contrast, carbon-based compounds

such as gossypol and condensed tannins increased, resulting in astronger induced response after herbivory (Coviella et al., 2002;Chen et al., 2005b; Wu et al. 2007, 2011; but see Sun et al.,2011). In spite of this, H. armigera larvae consumed more Btor non-Bt cotton material under elevated CO2 conditions, mostlikely to compensate for the reduced N content (Chen et al.,2005b). Thus, an increase in CO2 levels might lead to higherplant damage by herbivores. It is, however, unclear if cottonwill adapt to elevated CO2 levels as has been reported for otherplants (Lee et al., 2001; Chen et al., 2010).

VI. CONCLUSIONSCotton possesses a broad range of arthropod resistance traits.

Most of the studies on cotton-herbivore interactions have beenconducted with lepidopteran species since they represent mostof the major cotton pests. The impact of the resistance traits onother cotton pests and natural enemies has received relativelylittle attention.

Direct resistance mechanisms include morphological traitslike trichomes. The most important mechanism of direct re-sistance in cotton plants, however, is a group of closely re-lated terpenoids: gossypol, hemigossypolone and the heliocides1–4. These compounds affect herbivores and provide resistanceagainst pathogens. Terpenoid production is inducible and theirdistribution follows the optimal-defense theory. Several minorsecondary metabolites, like tannins and flavonoids also con-tribute to direct resistance. Furthermore, cotton plants possessinducible indirect resistance traits, like the emission of volatilesand the production of extrafloral nectar that attract and sustainpopulations of beneficial insects.

Despite the number of resistance mechanisms, cotton is at-tacked by a complex of arthropod pests and still receives alot of insecticide treatments. Therefore the wealth of resistancemechanisms provided by the plant should be explored by se-lecting for cultivars and production methods that enhance theplants’ direct and/or indirect resistance traits. This needs to besupplemented with production methods that conserve predatorsand parasitoids; e.g., reduced insecticide use, use of selectiveinsecticides, reduced tillage to increase ant densities or habitatmanagement to provide beneficials with alternative food sourcesor overwintering sites.

Since 1996, genetically engineered cotton cultivars that pro-duce insecticidal Cry proteins derived from Bacillus thuringien-sis have become an important component of IPM in the majorcotton producing regions in the world. The deployment of theseBt cotton cultivars has had a major impact on the control ofkey lepidopteran pests and has caused a significant decline inthe use of chemical insecticides, with benefits for biologicalpest control and biodiversity in general. Bt cotton technology,however, targets only lepidopteran pests, which while very im-portant, represent only a fraction of the arthropod complex thataffects cotton production globally. Consequently, conventionalhost plant resistance and other components of IPM are still

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essential to control pests and diseases that are not currently tar-geted by transgenic crop technology. This includes the plant andstink bugs that have benefited from the reduction in insecticideuse in Bt cotton. In addition, these is evidence that secondaryplant compounds in cotton, terpenoids in particular, can enhancethe efficacy of Cry toxins and also increase the fitness costs as-sociated with the resistance against the Cry toxins in the targetpests. Consequently, traits introduced by means of genetic en-gineering will be most sustainable for arthropod pest controlin cotton in combination with conventional approaches to hostplant resistance and other IPM tactics.

ACKNOWLEDGMENTSWe thank Ursus Kaufmann (Agroscope) for the drawing

of Figures 2 and 4 and Steven E. Naranjo (USDA-ARS) andMichael Meissle and Jana Collatz (Agroscope) for their thor-ough review of an earlier draft of the manuscript.

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