Phytochemistry 102 (2014) 97–105

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Phytochemistry

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3-b-D-Glucopyranosyl-6-methoxy-2-benzoxazolinone (MBOA-N-Glc)is an insect detoxification product of maize 1,4-benzoxazin-3-ones

http://dx.doi.org/10.1016/j.phytochem.2014.03.0180031-9422/� 2014 Elsevier Ltd. All rights reserved.

⇑ Corresponding author at: Neuchâtel Platform of Analytical Chemistry, Univer-sity of Neuchâtel, 2000 Neuchâtel, Switzerland. Tel.: +41 32 718 25 34.

E-mail addresses: daniel.maag@unine.ch (D. Maag), claudio.dalvit@unine.ch(C. Dalvit), damien.thevenet@unine.ch (D. Thevenet), angela.koehler@unine.ch(A. Köhler), fwouters@ice.mpg.de (F.C. Wouters), vassao@ice.mpg.de (D.G. Vassão),gershenzon@ice.mpg.de (J. Gershenzon), jean-luc.wolfender@unige.ch(J.-L. Wolfender), ted.turlings@unine.ch (T.C.J. Turlings), merb@ice.mpg.de(M. Erb), gaetan.glauser@unine.ch (G. Glauser).

Daniel Maag a,e, Claudio Dalvit b, Damien Thevenet c, Angela Köhler a, Felipe C. Wouters d,Daniel G. Vassão d, Jonathan Gershenzon d, Jean-Luc Wolfender e, Ted C.J. Turlings a, Matthias Erb d,Gaetan Glauser a,b,⇑a Laboratory of Fundamental and Applied Research in Chemical Ecology, University of Neuchâtel, 2000 Neuchâtel, Switzerlandb Neuchâtel Platform of Analytical Chemistry, University of Neuchâtel, 2000 Neuchâtel, Switzerlandc Laboratory of Organic Chemistry, University of Neuchâtel, 2000 Neuchâtel, Switzerlandd Max Planck Institute for Chemical Ecology, 07745 Jena, Germanye Laboratory of Phytochemistry and Bioactive Natural Products, University of Geneva, University of Lausanne, 1211 Geneva, Switzerland

a r t i c l e i n f o

Article history:Received 13 January 2014Received in revised form 10 March 2014Available online 5 April 2014

Keywords:Zea maysPoaceaeSpodopteraOstrinia nubilalisPlant defenceSecondary metabolites1,4-Benzoxazin-3-onesMBOAMBOA-N-GlcDetoxification

a b s t r a c t

In order to defend themselves against arthropod herbivores, maize plants produce 1,4-benzoxazin-3-ones(BXs), which are stored as weakly active glucosides in the vacuole. Upon tissue disruption, BXs come intocontact with b-glucosidases, resulting in the release of active aglycones and their breakdown products.While some aglycones can be reglucosylated by specialist herbivores, little is known about how theydetoxify BX breakdown products. Here we report on the structure of an N-glucoside, 3-b-D-glucopyran-osyl-6-methoxy-2-benzoxazolinone (MBOA-N-Glc), purified from Spodoptera frugiperda faeces. In vitroassays showed that MBOA-N-Glc is formed enzymatically in the insect gut using the BX breakdownproduct 6-methoxy-2-benzoxazolinone (MBOA) as precursor. While Spodoptera littoralis and S. frugiperdacaterpillars readily glucosylated MBOA, larvae of the European corn borer Ostrinia nubilalis were hardlyable to process the molecule. Accordingly, Spodoptera caterpillar growth was unaffected by the presenceof MBOA, while O. nubilalis growth was reduced. We conclude that glucosylation of MBOA is an importantdetoxification mechanism that helps insects tolerate maize BXs.

� 2014 Elsevier Ltd. All rights reserved.

Introduction

Plants produce a variety of chemicals to defend themselvesagainst arthropod herbivores. These metabolites can function asfeeding deterrents, repellents, or toxins (Gatehouse, 2002; Howeand Jander, 2008). One,4-benzoxazin-3-ones (BXs) are nitrogen-containing metabolites derived from indole-3-glycerol phosphatethat are found in many grasses including maize, wheat and rye.In most BX producing species, they represent the predominantclass of secondary metabolites (Macias et al., 2009, 2004;Niemeyer, 2009). Following their synthesis, BXs are stored as

inactive 2-O-b-D-glucosides in the vacuole. Upon tissue disruptionduring herbivory, BXs come into contact with b-glucosidases fromthe plastids, which hydrolyse the glucosidic bond and release theactive aglycones (Czjzek et al., 2000; Morant et al., 2008).

The most abundant BXs in maize are the constitutively present2-b-D-glucopyranosyloxy-4-hydroxy-7-methoxy-1,4-benzoxazin-3-one (DIMBOA-Glc) and the highly inducible 2-b-D-glucopyrano-syloxy-4,7-dimethoxy-1,4-benzoxazin-3-one (HDMBOA-Glc)(Glauser et al., 2011). Their respective aglycones, DIMBOA andHDMBOA, degrade spontaneously to 6-methoxy-2-benzoxazoli-none (MBOA) (Hashimoto and Shudo, 1996; Macias et al., 2004;Maresh et al., 2006). Such degradation is favoured at high pHvalues, which are found in the gut of many lepidopteran insects(Raun et al., 1966; Skibbe et al., 1996). During the degradationprocess, highly reactive a-oxo-aldehydes are formed, which reactwith nucleophilic moieties such as proteinogenic amino acids(Niemeyer, 2009).

Possibly due to their unspecific mode of action, BXs are toxic toa wide range of insect herbivores (Niemeyer, 2009). Resistance of

98 D. Maag et al. / Phytochemistry 102 (2014) 97–105

different maize genotypes to the European corn borer Ostrinianubilalis, for instance, is positively correlated with their DIMBOAcontents (Barry et al., 1994). Likewise, a positive correlation be-tween the mortality of larvae of the stalk corn borer Sesamia non-agrioides on different maize lines and their BX concentrations hasbeen observed (Niemeyer, 2009). Furthermore, DIMBOA exerts atoxic effect on S. nonagrioides caterpillars when administered inartificial diet (Ortego et al., 1998). Apart from BX aglycones, theirbreakdown product MBOA has also been reported to be toxic toherbivores. O. nubilalis larvae for instance suffer from higher mor-tality and increased developmental time when MBOA is added toartificial diet (Campos et al., 1988; Houseman et al., 1992). Thisis accompanied by a decrease of protease activity in the midgutsof these larvae (Houseman et al., 1992). High concentrations ofMBOA also lower the growth of another maize pest, Diatraea gran-diosella, the southwestern corn borer (Hedin et al., 1993).

OHO

HOO

OH

2(Hofmann e

1

OHO

HO

OH

OH

NO

1'2'3'

4'5'

6'

23

4

5

O

6OCH3

1

7

7a4a

Fig. 1. Chemical structures of 3-b-D-glucopyranosyl-6-methoxy-2-benzoxazolinone (1),tert-butyl-(2-hydroxy-4-methoxyphenyl) carbamate (3).

m/z100 200 300 400 500 600

%

100166.0505

350.0853

208.0611

328.1034166.0505208.0611

(b)

(a) Spodoptera frugiperda frass

1

DIMBOA-Glc

DIMBOA

MBOA

PLANT

DIET WITH MB

Feeding subs

Pseu

min

%

%

Fig. 2. Chromatogram and mass spectra of (1) obtained by UHPLC-QTOF-MS. (a) Charactafter feeding on MBOA-containing artificial diet or maize plants. Shown are the extractedmass spectrum of (1) in positive ionisation mode with proposed fragments. (c and d)ionisation mode. Following in-source fragmentation, the fragment at m/z 166.05, corfragmented.

However, some insect herbivores can transform BXs to lesstoxic forms. Larvae of the rice armyworm (Mythimna separata)for instance are able to enzymatically convert DIMBOA to DIM-BOA-Glc, which allows them to tolerate high concentrations of thistoxin in their diet (Sasai et al., 2009). Glucosylation of DIMBOA alsooccurs in Spodoptera littoralis and Spodoptera frugiperda (Glauseret al., 2011). Contrary to DIMBOA, little is known about the poten-tial detoxification of MBOA by insect herbivores. Accumulation of aputative glucoside of MBOA in the frass of two Spodoptera spp.after feeding on maize leaves has been observed, but its structureremained elusive (Glauser et al., 2011).

In the present study, we report on the structural elucidation ofthis novel benzoxazinoid derivative, 3-b-D-glucopyranosyl-6-methoxy-2-benzoxazolinone (MBOA-N-Glc, compound (1);Fig. 1), using a variety of spectral techniques including ultrahighpressure liquid chromatography coupled to quadrupole

O

N

HO

OCH3

t al. 2006)3

OCH3

HN

HO

O

O

1-(2-hydroxy-4-methoxyphenylamino)-1-deoxy-b-glucoside 1,2-carbamate (2) and

m/z50 100 150 180

%

10095.0447

67.0440

65.0392

166.0504110.0603

107.0365151.0267

122.0605

138.0555

(c)

OA

trate

m/z50 100 150 180

%

10095.0454

67.043565.0398

166.0507

110.0606

107.0372

151.0270

122.0606138.0549

(d)

do-MS3

%

%

eristic chromatograms of a frass extract from fourth instar S. frugiperda caterpillarsion chromatograms for m/z 166.05 in positive ionisation mode. (b) High resolution

MS2 experiment on MBOA (c) and pseudo-MS3 experiment on (1) (d) in positiveresponding to [MBOA+H]+, was selected in the quadrupole analyzer and further

F1

F2

2’-H/2’-OH

3’-H/3’-OH

4’-H/4’-OH6’-H/6’-OH

6’-H/6’-OH

(6’/4’)-OH

(6’/3’)-OH

(6’/2’)-OH

(a) (b)

Fig. 3. Expanded region of the 2D 1H DQF COSY (a) and ROESY (b) spectra of (1) in DMSO-d6. The COSY cross peaks for the four exchangeable protons are labelled. In theROESY spectrum the negative cross peaks (red colour) originate from ROE effects whereas the positive cross peaks (black colour) originate from exchange effects. Exchangecross peaks for the OH protons with themselves are indicated in the spectrum. In addition the four black cross peaks observed in the upper left of the 2D ROESY spectrumoriginate from the exchange of the four OH with the residual water signal at 3.32 ppm. (For interpretation of the references to colour in this figure legend, the reader isreferred to the web version of this article.)

carbamate (3)

MBOA

MBOA-N-Glc (1)

Fig. 4. UV–vis absorption spectra of MBOA (solid), (1) (dashed) and tert-butyl-(2-hydroxy-4-methoxyphenyl) carbamate (3) (dotted) in aqueous solution (pH 4).

D. Maag et al. / Phytochemistry 102 (2014) 97–105 99

time-of-flight mass spectrometry (UHPLC-QTOFMS) and nuclearmagnetic resonance (NMR) spectroscopy. We hypothesised thatthe enzymatic conversion of MBOA to MBOA-N-Glc in the insectgut provides an effective mechanism to alleviate its toxicity andsimultaneously facilitates its excretion from the insect body. In aseries of experiments we assessed the toxicity of MBOA to differentinsect herbivores that are commonly found on maize plants(Dutton et al., 2005) and possess different capacities to glucosylateMBOA.

Results

3-b-D-Glucopyranosyl-6-methoxy-2-benzoxazolinone (1) is excretedin the faeces of S. frugiperda

We previously reported the presence of a putative glucoside ofMBOA in the frass of S. frugiperda after feeding on maize leaves(Glauser et al., 2011). To confirm that S. frugiperda indeed producesthis glucoside, we devised an experiment in which S. frugiperda lar-vae were allowed to feed either on maize leaves or on an artificialdiet containing synthetic MBOA. After 24 h of feeding, frass wascollected and analysed by UHPLC-QTOFMS. In both cases (leavesand artificial diet), a minor peak of MBOA and a predominant peakof the putative glucoside (1) were observed (Fig. 2a). The high res-olution (HR) mass spectrum of (1) in positive ionisation modeshowed an [M+H]+ ion at m/z 328.1034 (C14H18NO8) together witha sodium adduct at m/z 350.0852 (Fig. 2b). A major fragment atm/z 166.0506 (C8H8NO3) corresponding to a neutral loss of162.0528 Da, typical of a dehydrated glucose moiety, was ob-served. In addition, a less abundant fragment at m/z 208.0610(C10H10NO4; [M+H-120]+) was present. To further elucidate thenature of the m/z 166.0506 fragment, a pseudo-MS3 experimentwas performed on (1) and compared with the MS2 fragmentationof MBOA (Fig. 2c and d). The HR mass spectra obtained by fragmen-tation of the m/z 166 ion selected in the first quadrupole were vir-tually identical for both compounds demonstrating that the m/z166.0506 fragment of (1) most likely corresponded to MBOA. This

MBOA-N-Glc

Inte

nsity

(cps

)

5.0e5

0

5.0e5

0Inte

nsity

(cps

)

(a) Gut suspension + MBOA + UDP-glucoseMBOA

MBOA(b) Boiled gut suspension + MBOA + UDP-glucose

(c) pH dependency of in vitro MBOA-N-Glc forma�on

min

min

0

20000

40000

60000

80000

100000

4.0 5.4 7.0 8.4 10.0

MBO

A-N-

Glc (

rela

�ve)

pH

Fig. 5. MBOA is glucosylated enzymatically in the gut of S. frugiperda. (a,b) HPLCchromatograms of in vitro assays containing MBOA and UDP-glucose after incuba-tion with gut homogenate from S. frugiperda (a) and after incubation with boiled guthomogenate from S. frugiperda (b). (c) pH optimum of in vitro MBOA-N-Glcformation. MBOA-N-Glc was quantified relatively and measured peak areas aregiven.

100 D. Maag et al. / Phytochemistry 102 (2014) 97–105

strongly suggested that the purified compound was indeed a glu-coside of MBOA.

However, as a related isomeric carbamate structure (2) (Fig. 1)had been reported in plants and was proposed to derive from (1)via spontaneous intramolecular rearrangement (Hofmann et al.,2006; Sicker et al., 2001), additional proof was sought to unambig-uously elucidate the structure of the putative molecule (1). There-fore, (1) was purified from S. frugiperda frass by semi-preparativeHPLC and a sample of the compound was dissolved in DMSO-d6for NMR analyses, keeping the presence of water to a minimum.This was necessary for the detection of the fast exchanging OH res-onances, i.e. the key signals for the unequivocal structure elucida-tion of the compound. The 1H and 13C NMR data were consistentwith the presence of the MBOA moiety in the molecule. The 1HNMR spectrum showed typical signals for a trisubstituted aromaticring and signals in the sugar region (Fig. S1). The proton spin-sys-tem of the aromatic ring was assigned in the 2D DQF-COSY spec-trum and the through space connectivities between the protonsof the aromatic ring and the protons of the glucoside were assignedin the 2D ROESY spectrum (Fig. S2). Fig. 3 depicts an expanded re-gion of the 2D DQF-COSY (Fig. 3a) and 2D ROESY (Fig. 3b) spectra,each containing the signals of the glucoside moiety of the mole-cule. The four OH exchangeable proton resonances were easilyrecognised in the 2D ROESY spectrum due to the presence of ex-change cross peaks between themselves and with the residualwater signal at 3.32 ppm. All four OH resonances displayed COSYcross peaks with the vicinal protons of the glucoside moiety. Forthe OH at 4.60 ppm two COSY cross peaks were observed with

the two vicinal protons of the CH2 group in position 60 (see ‘Exper-imental’ section for the assignments of all the proton and carbonresonances obtained by the analysis of the 2D DQF COSY, 2DROESY, 2D HSQC and 2D HMBC). The N-linkage between the aro-matic and the glucoside moieties was established with the obser-vation in the HMBC of the cross peaks between the anomericproton H-10 and the carbon C-4a and the carbonyl of the NCOOgroup (C-2). In summary, the NMR data were consistent withstructure (1), 3-b-D-glucopyranosyl-6-methoxy-2-benzoxazoli-none (MBOA-N-Glc), which was also in good agreement with theMS data. The alternative structure (2) proposed in the literature(Hofmann et al., 2006) was not compatible with the NMR dataobtained.

To provide a third line of evidence for the structure of (1), atert-butyl-(2-hydroxy-4-methoxyphenyl) carbamate (3) (Fig. 1)analogous to (2) was synthesised and the UV spectra of MBOA,MBOA-N-Glc (1) and the synthesised carbamate (3) were com-pared (Fig. 4). MBOA and (1) showed a maximum in absorptionat 286 nm. The presence of a carbamate in the chromophore of(3) led to a 4 nm shift of the absorption maximum (282 nm). More-over, the 2D ROESY NMR spectrum of (3) in DMSO-d6 clearlyshowed an exchangeable proton at 9.61 ppm originating from theOH on the aromatic ring, which was missing in compound (1) (datanot shown). To our knowledge, this is the first time that 3-b-D-glucopyranosyl-6-methoxy-2-benzoxazolinone (1) is reported.

MBOA is glucosylated enzymatically in the gut of S. frugiperda

In order to determine whether the glucosylation of MBOAoccurs enzymatically in the insect gut, in vitro assays with gut sus-pensions were conducted. Incubation of MBOA with a homoge-nised gut suspension of S. frugiperda and UDP-glucose led to theformation of the glucoside (Fig. 5a). Boiling of the gut homogenateprior to incubation with MBOA to denature any proteins resulted ina total loss of glucosylation activity, as no MBOA-N-Glc could bedetected (Fig. 5b). Furthermore, the pH optimum of in vitroMBOA-N-Glc formation was determined (Fig. 5c). S. frugiperdagut homogenate showed highest glucosylation activity at pH 7and minor activity at pH 8.4. At acidic or highly alkaline pH valuesalmost no activity was observed.

Resistance towards MBOA is associated with MBOA glucosylationcapacity

Previous studies reported toxic effects of MBOA on O. nubilalis(Campos et al., 1988; Houseman et al., 1992). We thereforehypothesised that this insect may not be able to glucosylate MBOA.To determine the MBOA glucosylation efficiency of O. nubilalis andcompare it with S. frugiperda and S. littoralis, caterpillars of thethree species were allowed to feed on MBOA-containing artificialdiet, and the accumulation of MBOA and MBOA-N-Glc in the frasswas monitored. In the frass of S. littoralis and S. frugiperda, almostno MBOA, but high concentrations of MBOA-N-Glc were present(Fig. 6a and b). In contrast, O. nubilalis predominantly accumulatedMBOA, but very little MBOA-N-Glc in the frass (Fig. 6c). Quantita-tive analyses (Table 1) revealed that the MBOA-N-Glc/MBOA ratiosin the frass of S. littoralis and S. frugiperda were about two orders ofmagnitude higher than in O. nubilalis (ANOVA, F2,10 = 275.99,P < 0.001). To determine whether MBOA glucosylation is associatedwith MBOA tolerance, the effect of MBOA on the performance ofthe different herbivore species was assayed in a no-choice feedingexperiment. The concentration of 330 lg MBOA/g fresh weight(FW) diet that was used in this assay is within the physiologicalrange of BX production in maize and potential conversion duringdigestion by Spodoptera (Glauser et al., 2011). MBOA had a negativeeffect on the growth of O. nubilalis (Fig. 6c): growth was reduced by

ns

MBOA

Gro

wth

redu

ctio

n re

lativ

e to

die

t with

out

MBO

A (%

)

ns

MBOA

**

MBOA

8.19x104

8.99x104

4.19x104

Fig. 6. MBOA resistance in Spodoptera spp. is associated with glucosylation efficiencies. (a–c) Characteristic UHPLC-QTOF-MS chromatograms of frass extracts from S. littoralis(a), S. frugiperda (b) and O. nubilalis (c) after feeding on MBOA-containing artificial diet. Shown are the extracted ion chromatograms for m/z 166.05 in positive ionisationmode. Feeding on MBOA-containing artificial diet for 7 days reduced the growth of O. nubilalis but not of S. littoralis or S. frugiperda as determined by no-choice feeding assayscontaining 330 lg MBOA/g FW diet. Differences in growth were calculated for each species relative to a control and expressed as percentage of growth reduction(mean ± standard error; N = 27–30). Asterisks indicate significant differences between treatments (⁄⁄P < 0.01).

Table 1Accumulation of MBOA-N-Glc and MBOA in the frass of different insect species.*

Species MBOA-N-Glc (lg/g DW) MBOA (lg/g DW) Ratio (MG/M)

S. littoralis 2261.2 ± 115.6 107.4 ± 25.6 18.60 ± 3.86aS. frugiperda 2736.2 ± 52.8 148.0 ± 18.1 19.78 ± 2.19aO. nubilalis 242.5 ± 22.8 873.8 ± 88.2 0.28 ± 0.03b

* Concentrations are given as means ± SE. MBOA-N-Glc/MBOA ratios were calcu-lated for each insect species based on the individual concentrations (mean ± SE).Different letters indicate statistically significant differences at P < 0.001. DW, dryweight, MG, MBOA-N-Glc, M, MBOA. (N = 3 for S. littoralis, N = 6 for S. frugiperda,N = 4 for O. nubilalis).

D. Maag et al. / Phytochemistry 102 (2014) 97–105 101

more than 35% compared to control diet after 7 days of feeding(t(55) = 3.30, P = 0.002). On the other hand, MBOA had no effecton the growth of S. littoralis (Mann–Whitney-U = 353, P = 0.297)and S. frugiperda (Mann–Whitney-U = 304, P = 0.108) (Fig. 6a andb).

MBOA-N-Glc is not toxic to herbivores

To determine whether MBOA-N-Glc itself has any effects on thegrowth of S. frugiperda or O. nubilalis, no-choice feeding assayswere conducted using synthetic MBOA and purified MBOA-N-Glc.Whereas no significant effects of either molecule on S. frugiperdawere observed (ANOVA, F2,80 = 0.64, P = 0.532), unexpectedly,MBOA-N-Glc increased O. nubilalis growth by more than 60%(Fig. 7). As in the previous assay, MBOA caused a significant reduc-tion of O. nubilalis weight gain (ANOVA, F2,43 = 32.79, P < 0.001).

Discussion

Following the observation of a potentially novel BX derivative incaterpillar frass (Glauser et al., 2011) this study aimed at elucidat-ing its structure and determining its significance for plant-herbivore interactions. Particular attention was paid to thethorough and unambiguous characterisation of the MBOA

102 D. Maag et al. / Phytochemistry 102 (2014) 97–105

glucoside (1) detected, as a closely related carbamate (2) had beenreported before. In this previous study (Hofmann et al., 2006),maize plants that were grown in presence of MBOA accumulateda methoxy glucoside carbamate (2) in their roots, which washypothesised to derive from a spontaneous intramolecular rear-rangement of (1) and to be a product of MBOA detoxification.The original structure elucidation of (2) was performed by NMRusing the compound dissolved in CD3OD. However, as theexchangeable NMR resonances were missing, the obtained datadid not allow unambiguous discrimination between (1) and (2).Therefore, in the present study, data from specific HR-MS andNMR experiments were combined with UV spectral data of thepurified compound itself and a synthesised model carbamate (3)to ensure efficient discrimination between the two possible struc-tures. All analytical data obtained were in good agreement with theMBOA-N-Glc (1). Definite proof was provided by the observation ofthe four OH proton resonances on the glucoside moiety and theirscalar connectivities in the DQF COSY spectrum in DMSO-d6(Fig. 3). This allowed for the unambiguous identification of themolecule as 3-b-D-glucopyranosyl-6-methoxy-2-benzoxazolinone(1). It should however be kept in mind that in the present studyan insect derivative of MBOA has been characterised. Thus, it can-not be excluded that plant roots are able to produce (2) under cer-tain conditions, as previously proposed (Hofmann et al., 2006).

To understand the significance of MBOA glucosylation, the toxiceffects of the BX breakdown product MBOA were examined inthree different insect species that frequently infest maize plantsin the field and correlated with their capacity to form MBOA-N-Glc. Exposure of O. nubilalis larvae to MBOA significantly reducedtheir growth. At the same time, these insects only excreted minoramounts of MBOA-N-Glc in their faeces. By contrast, S. littoralis andS. frugiperda were not susceptible to MBOA, which was associatedwith a substantial accumulation of MBOA-N-Glc in their frass andan almost complete depletion of MBOA. As MBOA-N-Glc did nothave any negative effect on the growth of S. frugiperda or O.

(a) S. frugiperda

(b) O. nubilalis

aa a

a

b

c

Control MBOA MBOA-N-Glc

Control MBOA MBOA-N-Glc

Fig. 7. MBOA-N-Glc is not toxic to S. frugiperda or O. nubilalis. (a and b) Average(±standard error) relative weight gains of S. frugiperda (a) and O. nubilalis (b) larvaeafter 7 days of feeding on different artificial diets containing either 300 lg MBOA/gFW diet (S. frugiperda), 400 lg MBOA/g FW diet (O. nubilalis), or 500 lg MBOA-N-Glc/g FW diet. Caterpillars in the control group were kept on a BX-free diet (N = 26–29 for S. frugiperda; N = 15–16 for O. nubilalis). Different letters indicate statisticallysignificant differences between treatments at P < 0.05.

nubilalis, we propose that the glucosylation of MBOA is an effectivestrategy of insect herbivores to escape the negative effects ofMBOA. Interestingly, O. nubilalis caterpillars grew considerably fas-ter when feeding on MBOA-N-Glc-containing diet compared tocontrol diet. The reason for this effect remains elusive. MBOA-N-Glc may either serve as a feeding stimulant or increase the nutri-tive value of the diet. However, due to its high stability (see below)and retention through the gut, it is unlikely that MBOA-N-Glc hasany nutritional value for O. nubilalis. Further experiments will benecessary to understand how MBOA-N-Glc positively influencesO. nubilalis growth.

In vitro assays with gut homogenate from S. frugiperda suggestthat the addition of the sugar moiety to MBOA is facilitated enzy-matically in the insect gut, as heat inactivation of the gut suspen-sion led to a loss of the glucosylation activity. In addition,glucosylation activity of S. frugiperda gut homogenate was pH-dependent. The observed optimum at physiological pH suggestsan intracellular localisation of the respective enzyme, presumablyin gut epithelial cells. Enzymatic glycosylation of toxic compoundsis a widespread counter-adaptation of insect herbivores to deacti-vate the chemical defence systems of their host plants (Ahn et al.,2012; Despres et al., 2007). The transfer of a sugar moiety from adonor to an acceptor molecule is typically catalysed by UDP-glu-cosyltransferases (UGTs). In insects, UDP-glucose acts as the mainsugar donor (Ahn et al., 2012). It has been shown that larvae of therice armyworm M. separata are able to re-glucosylate DIMBOA inan UDP-glucose-dependent manner suggesting the involvementof UGTs in this process (Sasai et al., 2009). Recently, the differentialtolerance levels of several Ostrinia species to plant-derived BXswere also related to their ability to degrade these molecules(Kojima et al., 2010). Whereas the in vitro degradation of BXs bya gut suspension from the Asian corn borer Ostrinia furnacalis alsorequired the presence of UDP-glucose, no de novo formation of anyBX glucosides was observed in these assays. It was hypothesisedthat newly formed DIMBOA-Glc may have been converted furtherand thus escaped detection. However, it cannot be excluded thatBX breakdown products were glucosylated by O. furnacalis instead.Generally, glycosylation may stabilise the target molecule, therebylowering its reactivity and simultaneously increase its water solu-bility, thus facilitating excretion (Pentzold et al., 2013). In ourexperiments, MBOA-N-Glc was stable under extreme conditions:The N-glycosidic bond was not prone to hydrolysis under strongacidic or alkaline conditions and elevated temperature in vitro(data not shown). Furthermore, MBOA-N-Glc could not be hydroly-sed in vitro by maize b-glucosidases (data not shown). Therefore,the glucosylation step may result in a largely inactive and stablemolecule compared to the aglycone MBOA.

The three insect species that were used in this study differremarkably in their feeding behaviours. Whereas S. littoralis andS. frugiperda can infest and defoliate young maize seedlings, adultfemales of O. nubilalis deposit their eggs on older plants (Turlingset al., 1998). After hatching, O. nubilalis larvae preferably feed inthe whorl or on the tassel before they move downwards and drillinto the stem to establish a feeding site (Beck, 1987). In maizestems, no effect of inducible defences on O. nubilalis larvae wasfound although a local induction of HDMBOA-Glc was observed(Dafoe et al., 2011). It was therefore hypothesised that O. nubilalismay have adapted to BXs and thus may be able to actively detoxifythese plant defence metabolites. Compared to levels in young seed-lings, however, BX concentrations in the stems of older maizeplants are more than three orders of magnitude lower and thusmay have been too low to exert any effect on the insects (Dafoeet al., 2011; Glauser et al., 2011). Even though young O. nubilalislarvae may forage on leaf tissue as well, they are unlikely toencounter high amounts of BXs, as concentrations decreasedramatically with plant age (Cambier et al., 2000). Therefore, our

D. Maag et al. / Phytochemistry 102 (2014) 97–105 103

finding that O. nubilalis is not able to process MBOA and is there-fore susceptible to it can be interpreted in two ways. First, becauseO. nubilalis rarely encounters high levels of BXs, there may simplynot have been a strong selection pressure for BX detoxificationmechanisms in O. nubilalis. Second, the inability of O. nubilalis todetoxify MBOA may have restricted its feeding niche and may haveled to a temporal adaptation within which the adults colonisemaize fields later and colonise older plants only. Comparing uni-and bivoltine O. nubilalis strains that colonise maize plants at dif-ferent phenologies may shed light on the second hypothesis inthe future. It is also noteworthy that S. littoralis, a herbivore thatis not adapted to maize chemical defences, displayed such highglucosylation capabilities. While S. littoralis is not able to success-fully stabilise and detoxify high amounts of DIMBOA in vivo(Glauser et al., 2011), glucosylation of MBOA may represent analternate strategy for this less adapted feeder.

Conclusion

Herbivores not only experience defence compounds producedby their host plant, but also have to cope with the breakdown prod-ucts of these compounds, which may still be active or even more ac-tive than the precursor molecules. We conclude that glucosylationof MBOA is a potentially significant detoxification mechanism of BXbreakdown products that occurs at the metabolic interface betweenplants and insects in Spodoptera spp. Future work should focus onthe identification of the genes and enzymes that are involved inMBOA glucosylation as well as on the adaptive value of this detox-ification strategy in generalist and specialist insect herbivores.

Experimental

Chemicals

Formic acid (FA) p.a. (P98%) and 6-methoxy-2-benzoxazoli-none (MBOA) were obtained from Sigma–Aldrich. HPLC grademethanol (MeOH) and acetonitrile (ACN) were obtained fromVWR International and Merck, respectively. Water, ACN and FAfor UHPLC-MS (ULC-MS grade) were obtained from Biosolve. Etha-nol (EtOH, 5% IPA) was obtained from Thommen-Furler. DMSO-d6(D, 99.9%) and CD3OD-d4 (D, 99.8%) were obtained fromCambridge Isotope Laboratories and UDP-Glucose was obtainedfrom Santa Cruz Biotechnology.

Insects and plants

Larvae of S. frugiperda (J.E. Smith, 1797) were obtained from acolony at the University of Neuchâtel. They were reared on a chick-pea flour-based artificial diet in plastic boxes under artificial lightconditions and at ambient temperature (Methods S1 for artificialdiets). Eggs of S. littoralis (Boisduval, 1833) were provided bySyngenta (Stein, Switzerland) and eggs of univoltine O. nubilalis(Hübner, 1825) were obtained from the Agroscope Changins-Wädenswil (Switzerland) and the two species were reared ontwo different wheatgerm-based artificial diets under the sameconditions as above. Maize (Zea mays L. var. Delprim) seeds wereobtained from Delley semences et plantes SA (DSP, Switzerland).Maize plants (Z. mays L. var. Delprim) were sown in plastic pots(11 cm height, 4 cm diameter) in commercial soil (Aussaaterde,Ricoter, Switzerland) and grown in a climatised green house withadditional lights (12:12 h l:d) during autumn season.

Frass analysis

Five fourth instar larvae of S. frugiperda were starved for a six-hour period and then transferred to freshly excised 25 d old maize

plants (3 fully developed leaves) inside a plastic box(15 � 13 � 6 cm3). Frass was collected after 16 h, pooled andlyophilised over night for chemical analysis. For quantitative frassanalyses fourth instar larvae of S. frugiperda, S. littoralis and O.nubilalis were starved for 6 h before they were transferred to arti-ficial diet containing 400 lg MBOA/g FW diet. After 24 h all insectswere transferred to clean Petri dishes and supplied with fresh diet.Caterpillars were allowed to feed for another 24 h before their frasswas collected. The frass from three individuals was pooled andconsidered as one biological replicate. The lyophilised frass wasthen ground to a fine powder using a mortar. An amount of approx-imately 4 mg was weighed into 1.5 ml tubes and 1 ml of MeOH/H2O (50:50, v/v; 0.5% FA) and 3–5 glass beads were added. Afterextraction in a beadmill for 3 min (30 Hz), the samples were centri-fuged at 14000 rpm for 5 min and 500 ll of the supernatants wererecovered. Benzoxazinoid derivatives were analysed using aWaters Acquity UPLC™ system coupled to a Waters Synapt G2QTOF mass spectrometer according to Meihls et al. (2013). Stan-dard curves were obtained from commercially available MBOA(Sigma–Aldrich) and purified compound (1).

Extraction and isolation of (1) from frass

Compound (1) was extracted from frass of fourth instar larvaeof S. frugiperda, which were kept on artificial diet containing500 lg MBOA/g FW. In total, 16.56 g dry weight (DW) of caterpillarfrass was collected and ground to a fine powder using a beadmill. Avolume of 700 ml of MeOH/H2O (50:50, v/v) was added to the pow-der and the suspension was sonicated in a bath for 30 min at ambi-ent temperature. Following centrifugation at 3500 rpm for 5 minthe supernatant was passed through a filter paper to obtain a clearextract. The solvent was partially evaporated using a rotary evap-orator and residual water was removed in a freeze dryer overnightyielding 5.8 g of crude extract. This extract was then subjected to asolid phase extraction (SPE). Briefly, it was re-dissolved in 100 mlof MeOH/H2O (5:95, v/v) and passed through 40 g of a C18 phase(Zeoprep 40–63 lm) inside a Buchner filtration funnel (porosity4). After a washing step with MeOH/H2O (5:95, v/v), the solidphase was eluted with 200 ml of methanol and the eluate wasevaporated to dryness in vacuo yielding 0.5 g of dry matter.

Compound (1) was further purified by semi-preparative HPLC(Waters 1525 EF) on an XTerra MS C18 column (150 � 19 mmi.d., 5 lm, Waters) connected to a dual wavelength UV detector(Waters 2487) equipped with a semi-preparative flow cell (volume2.6 ll, length 3 mm). The mobile phase consisted of 0.05% FA inwater (phase A) and 0.05% FA in ACN (phase B). The gradient wasas follows: 5% B over 3.15 min, 5–12.6% B over 41.85 min, 12.6–100% B over 10 min, held at 100% B for 15 min. The flow rate was8 ml/min. Before subsequent injections the column was re-equili-brated at initial conditions (5% B) during 15 min at a flow rate of12 ml/min. A total of 10 injections were executed. Compound (1)was detected at 285 and 305 nm and fractions were collected ata rate of 1 min�1 using a Gilson FC203B fraction collector. Presenceof (1) in the fractions was verified by UHPLC-QTOFMS and twofractions, which contained over 90% of the compound, werecollected, unified and stored at 4 �C until all injections had beenconducted. Solvents were partially evaporated using a rotaryevaporator and the remaining water was removed via freeze-drying. A total amount of 58.2 mg of (1) was isolated as a whitepowder.

Structure elucidation of compound (1)

UHPLC-HRMS analyses were carried out on a Waters AcquityBEH C18 column (50 � 2.1 mm i.d., 1.7 lm) using a Waters AcquityUPLC™ system coupled to a Waters Synapt G2 QTOF mass

104 D. Maag et al. / Phytochemistry 102 (2014) 97–105

spectrometer according to Meihls et al. (2013). MS2 and pseudo-MS3 experiments on MBOA and (1), respectively, were performedas follows: partial in source fragmentation of (1) was induced byapplying a relatively high cone voltage (25 V) while keeping MBOAintact, and the ion at m/z 166 was selected in the first quadrupoleand further fragmented in the transfer region of the collision cellusing a ramped collision energy of 10–30 eV and argon as collisiongas (2.2 ml/min). The resulting fragments were monitored by TOF-MS in the product ion scan mode over a mass range of m/z 40–600 Da.

For NMR analysis, a total of 2 mg of (1) was dissolved in 750 llof DMSO-d6 or CD3OD-d4. NMR experiments were performed on aBruker Avance II 400 NMR spectrometer equipped with a 5 mm BB/1H/19F probe. One dimensional 1H and 13C and two dimensional1H DQF COSY, 1H ROESY, 13C-1H HSQC and 13C-1H HMBC experi-ments were recorded at T = 25 �C. The signals of DMSO-d6 (1H:d = 2.50 ppm; 13C: d = 39.52 ppm) and CD3OD-d4 (1H:d = 3.31 ppm; 13C: d = 49.00 ppm) were used as internal references.The 2D 1H DQF COSY experiment was recorded with 8 scans foreach of the 256 t1 increments and with a SW of 7.5 ppm. The 2DROESY experiment was recorded with 8 scans for each of the256 t1 increments and with a SW of 8 ppm. The spin lock (lengthof 0.3 s) was performed with a train of 180 (x)–180 (�x) pulsesfor reducing the TOCSY magnetization transfer (Hwang and Shaka,1992). The 2D HSQC experiment was recorded with 8 scans foreach of the 128 t1 increments and with a SW of 8 and 110 ppmin the 1H and 13C dimensions, respectively and proton decouplingduring the acquisition period. For all three experiments (2D 1HDQF COSY, 2D ROESY, 2D HSQC), the data were multiplied with acosine square window function in both dimensions and zero-filledto 2048 and 1024 points in x2 and x1 dimensions, respectively,prior to Fourier transformation. The 2D HMBC experiment was re-corded with 96 scans for each of the 128 t1 increments and with aSW of 8 and 150 ppm in the 1H and 13C dimension, respectively.The data were multiplied with a sine window function in bothdimensions and zero-filled to 2048 and 1024 points in x2 and x1

dimensions, respectively, prior to Fourier transformation. The 1D13C spectrum was recorded with 2048 scans, a SW of 219 ppmand proton decoupling during the acquisition period.

UV measurements were carried out on a 8452A HewlettPackard diode array spectrophotometer with the different com-pounds dissolved in 0.1 M acetate buffer (pH 4) at the followingconcentrations: [MBOA] = 10 lg/ml; [Compound (1)] = 10 lg/ml;[carbamate (3)]: 50 lg/ml. IR analysis was performed on a Ther-mo Scientific Nicolet iS5 FTIR in the attenuated total reflection(ATR) mode. Specific optical rotation was measured on a JascoP-1030 polarimeter.

3-b-D-glucopyranosyl-6-methoxy-2-benzoxazolinone (1):Amorphous powder; [a]D

25 +16.3 (c = 0.6, MeOH); UV (Na acetatebuffer pH 4) kmax nm (log e): 286 (4.12); IR (ATR) mmax 3371,1754, 1639, 1499, 1454; HRESIMS m/z 328.1034 [M+H+] (calcu-lated mass for C14H18NO8 328.1032).

1H NMR (DMSO-d6, T = 25 �C) d 7.20 (d, 3J = 8.7 Hz, 1H, H-4),7.09 (d, 4J = 2.4 Hz, 1H, H-7), 6.79 (dd, 3J = 8.7 Hz, 4J = 2.4 Hz, 1H,H-5), 5.43 (d, 3J = 5.1 Hz, 1H, 20-OH), 5.17 (d, broad, 3J � 4.1 Hz,1H, 30-OH), 5.11 (d, 3J = 5.4 Hz, 1H, 40-OH), 5.08 (d, 3J = 9.2 Hz, 1H,H-10), 4.60 (t, 3J = 5.3 Hz, 1H, 60-OH), 3.77 (m, 1H, 20-H), 3.75 (s,3H, OCH3), 3.72 (m, 1H, 60-H), 3.47 (m, 1H, 60-H), 3.38 (m, 1H, 50-H), 3.33 (m, 1H, 30-H), 3.25 (m, 1H, 40-H); 13C NMR (DMSO-d6) d155.7 (C-6), 153.1 (C-2), 142.7 (C-7a), 122.3 (C-4a), 111.7 (C-4),109.4 (C-5), 97.1 (C-7), 83.9 (C-10), 79.9 (C-50), 76.8 (C-30), 69.7(C-40), 69.2 (C-20), 61.0 (C-60), 55.9 (OCH3).

1H NMR (CD3OD-d4, T = 23 �C) d 7.24 (d, 3J = 8.7 Hz, 1H, H-4),6.94 (d, 4J = 2.4 Hz, 1H, H-7), 6.79 (dd, 3J = 8.7 Hz, 4J = 2.4 Hz, 1H,H-5), 5.25 (d, 3J = 9.3 Hz, 1H, H-10), 3.99 (t, 3J = 9.0 Hz, 1H, 20-H,),3.91 (dd, 2J = 12.1 Hz, 3J = 1.9 Hz, 1H, 60-H,), 3.80 (s, 3H, OCH3),

3.72 (dd, 2J = 12.1 Hz, 3J = 5.6 Hz, 1H, 60-H), 3.53 (m, 1H, 30-H),3.51 (m, 1H, 50-H), 3.48 (m, 1H, 40-H); 13C NMR (CD3OD-d4) d158.0 (C-6), 155.8 (C-2), 144.8 (C-7a), 123.7 (C-4a), 112.8 (C-4),110.6 (C-5), 98.1 (C-7), 85.6 (C-10), 81.1 (C-50), 78.7 (C-30), 71.3(C-40), 71.1 (C-20), 62.7 (C-60), 56.4 (OCH3).

Synthesis of tert-butyl-(2-hydroxy-4-methoxyphenyl) carbamate (3)

Tert-butyl-(2-hydroxy-4-methoxyphenyl) carbamate (3) wassynthesised from 5-methoxy-2-nitrophenol via a non-optimisedtwo step procedure (for an optimised protocol see Sicker (1989)),and further purified by semi-preparative HPLC to yield 3.2 mg ofa yellow substance. A detailed protocol of the synthesis is givenin the supporting information (Methods S2).

In vitro UGT assays

Gut homogenates for in vitro assays were recovered from thirdto fourth instar larvae of S. frugiperda that were kept on artificialdiet. The caterpillars were dissected in cold 10 mM phosphate buf-fer (pH 7.0) and the gut tissue was isolated and its contents re-moved. The washed gut tissues were then transferred to a freshtube and homogenised with 100 ll of 10 mM phosphate buffer(pH 7.0) per gut. For the in vitro assays 10 ll of gut suspensionwere incubated with 75 nmol of MBOA and 150 nmol of UDP-glu-cose in 100 mM phosphate buffer at pH 7.0 (final assay volume:50 ll). After an incubation period of 30 min at 30 �C the reactionwas stopped by adding 50 ll of MeOH/FA (50:50, v/v). The sampleswere centrifuged at 7000 rpm for 5 min prior to analysis by HPLC-MS/MS. For the assay with boiled tissue an aliquot of the guthomogenate was heated at 100 �C for 15 min. To determine thepH optimum of in vitro MBOA-N-Glc formation homogenised gutsuspensions of S. frugiperda were incubated with MBOA andUDP-glucose as described above at five different pH values: 4.0(citrate), 5.4 (citrate), 7.0 (phosphate), 8.4 (Tris–HCl), 10.0 (carbon-ate; all buffers at 100 mM).

MBOA and MBOA-N-Glc toxicity assays

To assess the effects of MBOA and MBOA-N-Glc on insect per-formance, artificial diets were prepared (Methods S1). MBOA andpurified MBOA-N-Glc were dissolved in EtOH/H2O (50:50, v/v) toa concentration that was ten times higher than the desired finalconcentration in the artificial diet ([MBOA] = 400 lg/g FW diet;[MBOA-N-Glc] = 792 lg/g FW diet) and were added during thecooling process. An identical volume of EtOH/H2O (50:50, v/v)was added to the control diet. Larvae of the different herbivore spe-cies were kept individually in 6-well plates and supplied with dietcubes. The diet cubes were exchanged regularly to prevent themfrom drying and to guarantee ad libitum feeding. Any residualethanol was allowed to evaporate for approximately 15 min atambient temperature before the diet was presented to the caterpil-lars. Larval weights were determined at the beginning of the exper-iment and after 7 days of feeding and relative weight gain wascalculated according to the following formula: relative weightgain = weight gain/start weight. During the assays, the caterpillarswere kept in an air-conditioned room at 20 �C. The actual concen-trations of the compounds in the various diets were monitored byUHPLC-QTOFMS during the assay. To do so, 1 ml of MeOH/H2O(50:50, v/v; 0.5% FA) and 5–8 glass beads were added to approxi-mately 10–15 mg of fresh diet and the samples were extractedfor 10 min in a beadmill (30 Hz). After centrifugation at14,000 rpm for 5 min 500 ll of the supernatants were recoveredfor analysis. Quantification was done as described above. The ac-tual measured concentration was 330 lg MBOA/g FW diet in thefirst assay for both S. frugiperda and O. nubilalis. In the second assay,

D. Maag et al. / Phytochemistry 102 (2014) 97–105 105

actual concentrations were 300 lg MBOA/g FW diet (S. frugiperda),400 lg MBOA/g FW diet (O. nubilalis) and 500 lg MBOA-N-Glc/gFW diet (both insects). The concentrations of both compoundswere stable over the course of the experiments.

Statistics

All statistics were performed in SigmaPlot for Windows Version12.0 (Systat software). Insect growth reduction data were calcu-lated by dividing the relative weight gain of individual caterpillarsfeeding on MBOA-containing diet by the average relative weightgain of the control and expressed as percentage growth reduction.Statistically significant differences were detected by means of Stu-dent’s t-tests or Mann–Whitney rank sum tests using relativeweight gain as variable and treatment (‘Control’ vs. ‘MBOA’) as fac-tor. Statistical analysis of MBOA-N-Glc/MBOA ratios in the frass ofthe different insect species was conducted using a one-way ANOVAfollowed by a Holm-Sidak post hoc test. Prior to the ANOVA all datawere log-transformed to obtain equal variances. A one-way ANO-VA was also used to determine overall significant effects of MBOAand MBOA-N-Glc on caterpillar growth. Holm-Sidak post hoc testswere used for subsequent pairwise comparisons of the treatments.

Acknowledgements

Supply with eggs of Ostrinia nubilalis by Noémie Chervet fromthe Agroscope Changins-Wädenswil (Switzerland), Spodopteralittoralis eggs by Syngenta (Switzerland) and rearing of Spodopterafrugiperda by Yves Borcard (University of Neuchâtel) are acknowl-edged by the authors. We wish to thank Dr. Dieter Sicker and Dr.Bernd Schneider for useful comments on the manuscript. Thiswork was supported by a Sinergia grant from the Swiss NationalScience Foundation (CRSII3_136184).

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.phytochem.2014.03.018.

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