Insect Biochemistry and Molecular Biology 124 (2020) 103431

Contents lists available at ScienceDirect

Insect Biochemistry and Molecular Biology

journal homepage: www.elsevier.com/locate/ibmb

Novel glucosinolate metabolism in larvae of the leaf beetle Phaedon cochleariae

Jeanne Friedrichs a, Rabea Schweiger a, Svenja Geisler a, Andreas Mix b, Ute Wittstock c, Caroline Müller a,* a Department of Chemical Ecology, Bielefeld University, Universitatsstr.€ 25, 33615, Bielefeld, Germany b Department of Inorganic and Structural Chemistry, Bielefeld University, Universitatsstr.€ 25, 33615, Bielefeld, Germany c Institute of Pharmaceutical Biology, Technische Universitat€ Braunschweig, Mendelssohnstr. 1, 38106, Braunschweig, Germany

ARTICLE INFO ABSTRACT

Keywords: Plants of the Brassicales are defended by a binary system, in which glucosinolates are degraded by myrosinases, Phaedon cochleariae forming toxic breakdown products such as isothiocyanates and nitriles. Various detoxification pathways and Glucosinolate avoidance strategies have been found that allow different herbivorous insect taxa to deal with the glucosinolate- Detoxification myrosinase system of their host plants. Here, we investigated how larvae of the leaf beetle species Phaedon Metabolism cochleariae (Coleoptera: Chrysomelidae), a feeding specialist on Brassicaceae, cope with this binary defence. We Leaf beetle performed feeding experiments using leaves of watercress (Nasturtium officinale, containing 2-phenylethyl glu­ cosinolate as major glucosinolate and myrosinases) and pea (Pisum sativum, lacking glucosinolates and myr­ osinases), to which benzenic glucosinolates (benzyl- or 4-hydroxybenzyl glucosinolate) were applied. Performing comparative metabolomics using UHPLC-QTOF-MS/MS, N-(phenylacetyl) aspartic acid, N-(benzoyl) aspartic acid and N-(4-hydroxybenzoyl) aspartic acid were identifiedas major metabolites of 2-phenylethyl-, benzyl- and 4-hydroxybenzyl glucosinolate, respectively, in larvae and faeces. This suggests that larvae of P. cochleariae metabolise isothiocyanates or nitriles to aspartic acid conjugates of aromatic acids derived from the ingested benzenic glucosinolates. Myrosinase measurements revealed activity only in second-instar larvae that were fed with watercress, but not in freshly moulted and starved second-instar larvae fed with pea leaves. Our results indicate that the predicted pathway can occur independently of the presence of plant myrosinases, because the same major glucosinolate-breakdown metabolites were found in the larvae feeding on treated watercress and pea leaves. A conjugation of glucosinolate-derived compounds with aspartic acid is a novel metabolic pathway that has not been described for other herbivores.

1. Introduction the resulting aglucons are further metabolised into often toxic com­ pounds (Kissen et al., 2009; Morant et al., 2008; Thangstad et al., 2004). Plants have evolved a broad spectrum of defence mechanisms that Nevertheless, many insects can feed on plants expressing these binary restrict damage by insect antagonists. In addition to physical defences, defence mechanisms, because they possess various morphological, plants produce various classes of specialised (or secondary) metabolites behavioural or physiological adaptations (Pentzold et al., 2014b). (Mithofer€ and Boland, 2012) that act as repellents, deterrents or toxins Intriguingly, particularly with regard to physiological means of detoxi­ to herbivores (Agnihotri et al., 2018; Bruce, 2014; Kim and Jander, fication, multiple mechanisms have evolved to circumvent the same 2007). Several of these compounds are stored in the plants as glucosides, class of plant defences (Heckel, 2014; Jeschke et al., 2016; Pentzold such as benzoxazinoid, cyanogenic and iridoid glucosides as well as et al., 2014a, 2014b), but not all of these mechanisms are yet fully glucosinolates (Dobler et al., 2011; Morant et al., 2008; Winde and explored. Wittstock, 2011). Upon plant tissue disruption due to herbivore attack, The Brassicaceae are an agriculturally important plant family, which these metabolites come into contact with plant β-glucosidases, which are is well known for the glucosinolate-myrosinase system, also frequently usually stored separately. These β-glucosidases cleave the glucose and referred to as the ‘mustard oil bomb’ (Matile, 1980). Glucosinolates are

* Corresponding author. E-mail address: [email protected] (C. Müller). https://doi.org/10.1016/j.ibmb.2020.103431 Received 1 May 2020; Received in revised form 18 June 2020; Accepted 22 June 2020 Available online 10 July 2020 0965-1748/© 2020 Elsevier Ltd. All rights reserved. J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

anions composed of thiohydroxymates with an S-linked β-glucopyr­ feeding generalists as well as the chrysomelid P. chrysocephala conjugate anosyl residue and an O-linked sulfate residue in addition to a variable isothiocyanates with the pseudo-tripeptide glutathione via a glutathione side chain, which is derived from different amino acids (Halkier, 2016; S-transferase and excrete the resulting non-toxic conjugates (Beran Bla�zevi�c et al., 2020). Myrosinases are thioglucoside glucohydrolases et al., 2018; Schramm et al., 2012; Wadleigh and Yu, 1988). These ex­ generally stored separately from glucosinolates. When coming into amples highlight the huge diversity and species-specificity of glucosi­ contact, the S-glycosidic bond of the glucosinolate is hydrolysed and an nolate metabolism by insect herbivores. unstable aglucon formed, which is further re-arranged. Depending on In the present study, we investigated the glucosinolate metabolism in the pH, the structural type of the glucosinolate side chain and the larvae of the mustard leaf beetle Phaedon cochleariae (Coleoptera: presence of specifier proteins, different biologically active metabolites Chrysomelidae), which is a feeding specialist on plants of the Brassica­ arise, including isothiocyanates, thiocyanates, nitriles or other classes of ceae. The beetles rely on glucosinolates for host plant recognition hydrolysis products (Lambrix et al., 2001; Wittstock et al., 2016) (Nielsen, 1978; Reifenrath and Müller, 2008), but it was unclear how (Fig. 1A). However, there are also instances of endogenous catabolism of this species deals with the glucosinolate-myrosinase system upon glucosinolates without prior tissue disruption (Bednarek et al., 2009; feeding. To study which metabolites result from glucosinolate meta­ Petersen et al., 2002). Moreover, microbiota localised in the guts of bolism in P. cochleariae, larvae were offered leaves of a Brassicaceae various organisms can metabolise plant glucosinolates (Hammer and host, watercress (Nasturtium officinale), or a non-host, pea (Pisum sat­ Bowers, 2015; van den Bosch and Welte, 2017). ivum, Fabaceae). Pea lacks glucosinolates and myrosinases but is Numerous herbivorous insect species have evolved physiological accepted by P. cochleariae when glucosinolates are applied on the leaves counter-adaptations to the glucosinolate-myrosinase system of the (Reifenrath et al., 2005). The benzenic glucosinolates benzyl glucosi­ plants (Abdalsamee et al., 2014; Jeschke et al., 2016; Winde and Witt­ nolate (glucotropaeolin) and 4-hydroxybenzyl glucosinolate (glucosi­ stock, 2011). To circumvent the ‘mustard oil bomb’ some species nalbin), which do not naturally occur in watercress (Reifenrath et al., sequester glucosinolates into body tissues, thereby separating them from 2005), were administered on leaves of both plant species and larvae and plant myrosinases. This mechanism has been found in insects with their faeces analysed for potential glucosinolate breakdown metabolites. piercing-sucking feeding style such as Brevicoryne brassicae (Hemiptera, Due to the highly species-specific detoxification pathways for the Aphididae) (Francis et al., 2001), Murgantia histrionica (Hemiptera, glucosinolate-myrosinase system known from different insects, we Pentatomidae) (Aliabadi et al., 2002) as well as in species with anticipated that P. cochleariae evolved yet another way to cope with this chewing-biting mouthparts such as Athalia rosae (Hymenoptera, Ten­ defence. Furthermore, if the metabolism of glucosinolates in thredinidae) (Müller et al., 2001), Phyllotreta striolata and Psylliodes P. cochleariae depends on plant myrosinases, we predicted that gluco­ chrysocephala (Coleoptera, Chrysomelidae) (Beran et al., 2014, 2018). sinolates are metabolised in larvae feeding on treated watercress leaves Aphids of B. brassicae and beetles of P. striolata express their own myr­ but remain intact in larvae fed with treated pea leaves. osinases, forming a binary defence together with the sequestered glu­ cosinolates (Beran et al., 2014; Kazana et al., 2007). In larvae of Pieris 2. Materials and methods rapae (Lepidoptera, Pieridae), a nitrile-specifier protein (NSP) redirects the hydrolysis of glucosinolates to less toxic nitriles (Wittstock et al., 2.1. Plant and insect rearing 2004). The feeding generalist Schistocerca gregaria (Orthoptera, Acridi­ dae) and the specialist Plutella xylostella (Lepidoptera, Plutellidae) pro­ A breeding stock of P. cochleariae was kept in the laboratory under � duce a sulfatase that hydrolyses the glucosinolates to constant climate conditions (20 C, 65% r.h., 16:8 h light:dark). The desulfoglucosinolates (Falk and Gershenzon, 2007; Ratzka et al., 2002), stock consisted of 500–600 individuals and every year about 100 wild which cannot be cleaved by myrosinases. Likewise, in other insect taxa individuals were collected from watercress plants at the river Ems in � 0 00 � 0 00 glucosinolate sulfatase activity has been found (Beran et al., 2018; Germany (51 51 21 N, 8 41 37 E) and crossed into the stock. Beetles Malka et al., 2016; Opitz et al., 2011). In contrast, some Lepidopteran were reared in plastic boxes (20 � 20 � 6.5 cm; 50–100 individuals per

Fig. 1. A) Glucosinolate–myrosinase system of plants. Upon tissue damage myrosinases catalyse the hydrolysis of glucosinolates, yielding glucose and unstable aglucons. The aglucons rearrange spontaneously to form isothiocyanates, nitriles, epithionitriles, organic thiocyanates and other products. B) Photo of a second-instar larva of Phaedon cochleariae, placed on metric graph paper. One minor square corresponds to 1 mm2.

2 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

box) and were provided ad libitum with 7–8 weeks old non-flowering range that is not too far from concentrations larvae experience in nature. watercress plants (seeds from Volmary GmbH, Münster, Germany). Under these rearing conditions, the larvae take about 15–17 d for their 2.3. Recovery of glucosinolates from treated leaves entire larval development with three larval instars and the pupal dura­ tion lasts for 6 d (Kühnle and Müller, 2011). The glucosinolate profileof To quantify the glucosinolate amounts that could be recovered from watercress leaves is dominated by 2-phenylethyl glucosinolate (gluco­ treated leafletsafter an experimental period of 23 h without any impact nasturtiin) (>85% of the total glucosinolate concentration) as well as by larvae, leafletsof watercress and pea were treated with glucosinolate two indole glucosinolates, 4-methoxyindol-3-ylmethyl glucosinolate solutions as described above (2.2. Feeding experiments). After evapora­ and indol-3-ylmethyl glucosinolate, while a few other glucosinolates tion of the solvents, the leafletswere kept in Petri dishes lined with wet occur only in minor quantities (Reifenrath et al., 2005). For feeding filter paper for 23 h without larvae and then frozen in liquid nitrogen experiments (see below), seven-to-eight-week-old non-flowering (six replicates per plant species and per glucosinolate solution). As watercress and six-week-old flowering pea plants (seeds from Kie­ controls, 50 μl of the 40 mM solutions of benzyl glucosinolate as well as penkerl, Bruno Nebelung GmbH, Konken, Germany) were used. All 4-hydroxybenzyl glucosinolate (one replicate per glucosinolate) were plants were grown in pots (12 cm diameter) on composted soil in a put in Eppendorf tubes and directly frozen in liquid nitrogen. All sam­ � greenhouse (60% r.h., 16:8 h light:dark). ples were stored at À 80 C and freeze-dried. Glucosinolates were extracted from this material by a threefold 2.2. Feeding experiments extraction in 80% methanol (v:v) and trapped on ion exchange columns [diethylaminoethyl (DEAE) Sephadex A-25 (Sigma Aldrich, St. Louis, To investigate the metabolism of benzenic glucosinolates in MO, USA) swelled in acetic acid buffer, for details see Abdalsamee and P. cochleariae, larvae of the second instar (about 6 mm long; Fig. 1B) Müller (2012)]. Allyl glucosinolate (sinigrin, Phytoplan GmbH) was were offered leaflets of either watercress or pea treated with 40 mM added as an internal standard at the first extraction. After incubation solutions of benzyl glucosinolate (glucotropaeolin, 1, Fig. 2) or 4- with a sulfatase overnight, desulfoglucosinolates were eluted with water hydroxybenzyl glucosinolate (glucosinalbin, 3, both glucosinolates and analysed using a high performance liquid chromatograph coupled to >99%, Phytoplan GmbH, Heidelberg, Germany) in a mixture of meth­ a diode array detector (1200 Series, Agilent Technologies, Inc., Santa μ anol, millipore water and dichloromethane (64:30:6 v:v:v; organic sol­ Clara, CA, USA) and equipped with a Supelcosil LC-18 column (3 m, � vents from Fisher Scientific, Loughborough, UK). Single leaflets of 150 3 mm, Supelco, Bellefonte, PA, USA) at a gradient from 5% to watercress or pea with a diameter of around 2.2 cm were treated with 95% methanol in water (as in Abdalsamee and Müller, 2012). Peaks of 50 μl of either benzyl glucosinolate or 4-hydroxybenzyl glucosinolate desulfoglucosinolates were quantified at 229 nm and response factors solution (resulting in 2 μmol per leaflet) or the respective solvent taken into account (0.95 for benzyl glucosinolate, 0.5 for 4-hydroxyben­ mixture only and offered in a Petri dish (55 mm diameter) lined with zyl glucosinolate and 1 for allyl glucosinolate). Recovery was calculated moistened filterpaper. The solutions or solvent were pipetted on the leaf as percentage of exogenous glucosinolate extracted from treated plant surface and the solvent allowed to evaporate for 40 min. The applied material relative to the amount detected in control samples (set to glucosinolate amount was chosen, because it equals about ten times the 100%). Recovery from watercress versus pea leaves was compared for mean concentration of benzyl glucosinolate in young leaves of white each glucosinolate with Mann-Whitney U-tests (with continuity mustard (Sinapis alba) (roughly 2 μmol/g dry weight) (Martin and correction due to ties in the data) in RStudio v. February 1, 5033 with Müller, 2007; Neubauer et al., 2014), a potential host of P. cochleariae required R v. 3.6.3. (R Developmental Core Team, 2020). (Reifenrath and Müller, 2008). We aimed to provoke sufficientamounts of glucosinolate breakdown metabolites for identificationbut to stay in a

Fig. 2. Glucosinolates applied in feeding assays [benzyl glucosinolate (1) and 4-hydroxybenzyl glucosinolate (3)] and the main glucosinolate of watercress [2- phenylethyl glucosinolate (7)] as well as their major metabolites in larvae of Phaedon cochleariae. Molecular formulas, molecular weights (MW) and main ions found using UHPLC-ESI--QTOF-MS/MS (MS mode) are shown.

3 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

2.4. Analysis of metabolites in larvae and faeces (i. e., their mean intensities in samples of larvae fed with glucosinolate- treated leafletsdivided by their mean intensities in samples of larvae fed To identify potential metabolites of ingested glucosinolates in with control leaflets) were computed. To screen for potential degrada­ P. cochleariae, larvae were taken directly after moult to the second instar tion products of glucosinolates, intensities of features in the different (11–12 days old), starved for 8 h and then offered leafletsof watercress treatment groups (plant species * treatment of the leaflet* type of larval or pea treated with glucosinolate solution or solvent control in groups of sample) were inspected and only features that fulfilledall the following three larvae in Petri dishes (as described in 2.2. Feeding experiments; n ¼ criteria were left in the dataset: 1) their mean intensity in at least one 6 replicate Petri dishes per plant species and glucosinolate or solvent treatment group was at least 50 x higher than their mean intensity in the control). After a feeding period of 23 h, three groups of three larvae per blanks, 2) they occurred in at least two of the three replicate samples of treatment combination were dissected on ice into guts and remaining at least one treatment group, 3) their fold change was >2 or they only body tissues and the guts were shortly rinsed in millipore water to clean occurred in samples of larvae fed with glucosinolate-treated leaflets(but them from haemolymph. Body parts were then put in 2 ml Eppendorf not in the corresponding control samples), 4) their mean intensity in at tubes and frozen in liquid nitrogen. The remaining three groups of three least one group of samples (of glucosinolate-fed larvae) was >1000 larvae were transferred into 2 ml Eppendorf tubes and left for 3 h counts. without food to collect their faeces. Afterwards these larvae with For the identification of relevant metabolites that differed between emptied guts were transferred to new tubes and frozen (“whole larvae”). samples taken from larvae fed with glucosinolate- versus solvent-treated In addition, watercress and pea leaflets treated with benzyl glucosino­ leaves (metabolites 1–8, Tables 1 and S1), samples in which these me­ late or 4-hydroxybenzyl glucosinolate solution or the solvent control (n tabolites were abundant were again analysed by UHPLC-QTOF-MS/MS. ¼ 3 per treatment combination) were taken at the beginning of the To further trace the ions formed in MS mode and their fragments in MS/ � experiment. All samples were frozen in liquid nitrogen, stored at À 80 C MS mode, lower spectra rates (2 Hz) were used and multiple reaction and lyophilised. monitoring (with isolation widths and collision energies adjusted for For further processing, samples were extracted in 300 μl 90% (v:v) each parent ion m/z) was applied both in negative (parameter settings as ice-cold methanol containing hydrocortisone (>98%, Sigma-Aldrich described above) and positive (capillary voltage: 4500 V) ESI mode. Chemie GmbH, Steinheim Germany) as internal standard and centri­ Molecular formulas of parent and daughter ions were created with Smart � fuged for 5 min at 4 C. The supernatants were collected, filtered with Formula (3D) in DataAnalysis and ranked according to their m/z devi­ 0.2 μm polytetrafluoroethylenemembrane syringe filters(Phenomenex, ation and isotopic pattern fit.Moreover, MetFrag (Ruttkies et al., 2016) � Torrance, CA, USA) and stored at À 80 C. Metabolome analyses were was used for in-silico fragmentation and matching of spectra against the carried out using an ultra-high performance liquid chromatograph MassBank of North America (https://mona.fiehnlab.ucdavis.edu/) to coupled to a quadrupole time of flight mass spectrometer (UHPLC- obtain suggestions for structural formulas. Retention times, UV/VIS QTOF-MS/MS; UHPLC: Dionex UltiMate 3000, Thermo Fisher Scientific, spectra as well as MS and MS/MS spectra were compared to those of San Jose,� CA, USA; Q-TOF: compact, Bruker Daltonics, Bremen, Ger­ reference standards included in an in-house database that were many) equipped with a Kinetex XB-C18 column (1.7 μm, 150 � 2.1 mm, measured under the same conditions. To specifically search for iso­ with guard column, Phenomenex, Torrance, CA, USA). Chromato­ thiocyanates and nitriles that could have been formed from phenylethyl graphic separation was done following Schrieber et al. (2019) with glucosinolate (in watercress) and the glucosinolates applied on the slight modifications, using a gradient of water with 0.1% (v:v) formic leaflets, several standards (2-phenylethylisothiocyanate, 3-phenylpro­ acid (FA; eluent additive for LC-MS, ~98%, Sigma-Aldrich) (eluent A) to panenitrile, phenylacetonitrile, 4-hydroxyphenylacetonitrile) were acetonitrile (LC-MS grade, Th. Geyer, Renningen, Germany) with 0.1% measured under the same conditions and larval samples screened for À (v:v) FA (eluent B) with a flowrate of 0.5 ml min 1, with 2% B to 30% B these metabolites. within 20 min, then to 75% B within 9 min, followed by column cleaning and equilibration. Line spectra (50–1300 m/z) were acquired in MS and 2.5. Purification and structure elucidation of major metabolite (2) of À MS/MS mode using negative electrospray ionisation (ESI ) at 8 Hz with benzyl glucosinolate end plate offset: 500 V, capillary voltage: 3000 V, nebuliser (N2) pres­ À 1 � sure: 3 bar, dry gas (N2) flow and temperature: 12 l min at 275 C, To purify the main metabolite (2) of benzyl glucosinolate (1), quadrupole ion energy: 4 eV, low mass: 90 m/z, collision energy in MS additional feeding experiments were carried out. Groups of three third mode: 7 eV, transfer time: 75 μs, pre-pulse storage: 6 μs. Fragmentation instar larvae (16 days old, to gain more material) were fed with of the most intense ions was performed using N2 as collision gas in watercress leaves treated with benzyl glucosinolate solution for 23 h in Auto-MS/MS mode, applying a ramping of collision energies and isola­ Petri dishes. After the feeding period, larvae were transferred into 2 ml tion widths along with increasing m/z. To enable mass axis recalibra­ Eppendorf tubes and allowed to defaecate for 4 h. The faeces left in the tion, a sodium formate calibration solution was introduced to the ESI corresponding Petri dishes were also added to the tubes. The extraction sprayer preceding each sample. Ten blanks were measured using the of these pooled larvae and their faeces was performed from lyophilised same method. samples as described in 2.4. The metabolite (2) was purified by semi- Mass axis recalibration and picking of molecular features (each preparative HPLC-DAD (1200 Series, Agilent Technologies) equipped described by a retention time and m/z) were done in Compass Data­ with a Kinetex XB-C18 column (5 μm, 250 � 4.6 mm, with guard col­ Analysis 4.4 (Bruker Daltonics) using the Find Molecular Features al­ umn, Phenomenex). Per run 25 μl of a sample were separated using a gorithm including spectral background subtraction. The settings used gradient of eluent A (water with 0.1% v:v FA) and eluent B (acetonitrile À were signal-to-noise threshold: 3, correlation coefficientthreshold: 0.75, with 0.1% v:v FA) at a flow rate of 1 ml min 1 with 2% B to 20% B minimum compounds length: 25 spectra, smoothing width: 10. For within 20 min, then 20% B to 100% B within 5 min, followed by column bucket generation (i. e., sorting of features belonging to the same cleaning and equilibration. When metabolite (2) was visible at 229 nm - - metabolite together) the ion types allowed were: [M-H] , [M–H2O–H] , (at approximately 16 min), it was collected manually for 1 s in a 5 ml - - - - [MþCl] , [M þ HCOOH–H] , [M þ CH3COOH–H] , [2M-H] , [2M þ glass flask.In total, collections of 56 runs were pooled and freeze-dried - - - HCOOH–H] , [2M þ CH3COOH–H] , [3M-H] . Compass ProfileAnalysis in between collection bouts. 2.3 (Bruker Daltonics) was used for alignment of buckets across samples, For analysis by nuclear magnetic resonance (NMR), metabolite (2) allowing deviations of 0.1 min (retention time) and 5 mD (m/z), was dissolved in 0.5 ml of D2O (Deutero GmbH, Kastellaun, Germany). respectively. The intensities (peak heights) of the most intense metabolic NMR spectra were recorded on a Bruker Avance FT NMR spectrometer features within each bucket were related to the intensities of the [M þ operating at a proton resonance frequency of 600.13 MHz. The instru­ - HCOOH–H] ion produced by hydrocortisone. Fold changes of features ment was equipped with a 5 mm BBO SmartProbe. Proton NMR data

4 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

were collected at room temperature using a pulse sequence including comparison to reference standards. Besides the dominant deprotonated presaturation of the water resonance. Chemical shifts are given in parts molecular ions in MS mode (see m/z values above), the glucosinolates per million (ppm). produced characteristic fragments derived from the sulfate residues, i. - - - e., [SO3] , [SO4] and [HSO4] ions (Table S1). The glucosinolates were þ not visible in ESI MS mode. 2.6. Measurements of myrosinase activities À One main metabolite with 236.0556 m/z in ESI mode (at 5.75 min, To test for active myrosinases in samples, larvae were starved for 2; Table 1) was found exclusively in larvae fed with added benzyl glu­ different durations and subsequently fed with leaflets of watercress or cosinolate (1). In larvae fed with leaves treated with 4-hydroxybenzyl pea as described in 2.2. (Feeding experiments). For the first batch, mid glucosinolate (3), three abundant metabolites with 252.0512 m/z (at second-instar larvae were starved for 3.5 h and then either further 3.20 min, 4), 137.0242 m/z (at 4.13 min, 5) and 121.0294 m/z (at 5.45 processed or fed untreated watercress or pea leaves for 23 h. In the min, 6) were detected. Exclusively in samples of larvae that had fed on second batch, freshly moulted second-instar larvae were starved for 8.5 watercress with 2-phenylethyl glucosinolate (7) as dominant glucosi­ h and then either further processed or fed untreated watercress or pea nolate, a metabolite with 250.0730 m/z (at 6.78 min, 8) was detected. For all these metabolites, the m/z difference of ca. 2 Da between the leaves for 24 h. Material from groups of three larvae was pooled directly À þ after the starvation phase (n ¼ 1) or after the corresponding feeding time dominant ions in the ESI and the ESI mode indicated that the m/z on watercress or pea leaves (n ¼ 3 replicates per treatment combina­ values given above represent deprotonated ions (Table 1). tion). The larvae were dissected and the guts rinsed in millipore water to The m/z difference between the glucosinolates (1, 3 and 7) and the clean them from haemolymph. Guts as well as the remaining bodies main metabolites found in the respective larval samples (2, 4 and 8) was were collected in separate 2 ml Eppendorf tubes. All dissection steps consistently about À 172 m/z (Table 1). The molecular formulas were performed on ice. In addition, leaves of watercress and pea (n ¼ 3 assigned to these main metabolites revealed that they contained, in per species) as well as pupae developed from larvae kept on watercress contrast to the glucosinolates, no sulfur (Table 1). Analyses of the (n ¼ 1, pooled from 3 individuals) were taken. All samples were stored at fragmentation patterns revealed that these three metabolites contained � À 20 C until activity measurements of myrosinases. aspartic acid residues, as indicated by the common fragments with about - - To test for activities of soluble myrosinases, samples were macerated 115 m/z ([C4H3O4] ) (2, 4 and 8) and 132 m/z ([C4H6NO4] ) (for 4 and in extraction buffer (200 mM Tris, 10 mM ethylenediaminetetraacetic 8) (Table S1). The compounds were putatively identifiedas N-(benzoyl) acid, pH 5.5) on ice and centrifuged. Glucosinolates present in the su­ aspartic acid (2) derived from benzyl glucosinolate (1), N-(4-hydrox­ pernatants were removed using DEAE Sephadex A-25 columns, as ybenzoyl) aspartic acid (4) derived from 4-hydroxybenzyl glucosinolate described above. The purified extracts were then measured in 96-well (3) and N-(phenylacetyl) aspartic acid (8) derived from 2-phenylethyl microplates on a microplate photometer (Multiskan EX, Thermo Elec­ glucosinolate (7) (Table 1, Fig. 2). tron, China) at 492 nm for 45 min at room temperature after addition of No reference standards were available for these three metabolites. 1 mM benzyl glucosinolate dissolved in phosphate buffer (pH 6.5) as However, the identity of N-(benzoyl) aspartic acid (2) was further substrate or phosphate buffer as controls. The kinetics of glucose release confirmedby NMR spectroscopy after purificationof the metabolite. The 1 due to myrosinase activity were measured by adding a mixture of H NMR spectrum showed corresponding signals for the two subunits of 3 ¼ glucose oxidase, peroxidase, 4-aminoantipyrine and phenol as colour the molecule (Fig. S1). The resonances at δH 7.72 [d, J(H,H) 7.5 Hz, 3 ¼ 3 ¼ reagent, following the protocol described in Travers-Martin et al. 2H], δH 7.53 [t, J(H,H) 7.5 Hz, 1H] and δH 7.45 [pseudo-t, J(H,H) (2008). A calibration curve of glucose was measured on each plate and 7.5 Hz, 2H] could be assigned to fiveprotons bound to a phenyl ring. The used for quantification. Soluble protein concentrations of the purified signals of the protons at ortho and meta positions were shifted downfield, extracts were determined according to Bradford (1976) using bovine indicating the presence of an electron-withdrawing group. For the serum albumin as standard and myrosinase activities related to those. aspartic acid moiety, three resonances were obtained in the spectrum exhibiting relative intensities of 1:1:1, as expected. The resonance at δH 3 ¼ 3. Results 4.53 [dd, J(H,H) 3.7 and 9.6 Hz, 1H] could be attributed to H–C(α). For the diastereotopic protons at C(β) two separate resonances were 2 ¼ 3 ¼ 3.1. Main metabolites identified in samples of larvae and faeces obtained at δH 2.72 [dd, J(H,H) 15.6 Hz, J(H,H) 3.7 Hz, 1H] and 2 3 δH 2.57 [dd, J(H,H) ¼ 15.6 Hz, J(H,H) ¼ 9.6 Hz, 1H]. Besides the Across all samples, eight compounds (1–8, Table 1) were identified signals of metabolite (2), further resonances were obtained which have via UHPLC-QTOF-MS/MS that differed between samples of the different to be attributed to impurities. treatments. The identities of the applied benzyl glucosinolate (1; eluting Benzyl glucosinolate (1) and 4-hydroxybenzyl glucosinolate (3), À at 5.35 min and producing a major ion with 408.0436 m/z in ESI MS which had been applied to the leaves, were found in several samples of mode) and 4-hydroxybenzyl glucosinolate (3; at 2.30 min, 424.0386 m/ the different respective insect parts (Table 2). The leaf-endogenous 2- z) as well as of the major glucosinolate of watercress, 2-phenylethyl phenylethyl glucosinolate (7) was found in four out of six faecal sam­ glucosinolate (7; at 7.90 min, 422.0583 m/z) were confirmed by ples and in one out of six bodies as well as one whole larva that had been

Table 1 Glucosinolates (benzyl glucosinolate, 4-hydroxybenzyl glucosinolate, 2-phenylethyl glucosinolate) and their breakdown metabolites found in samples of larvae of À þ Phaedon cochleariae, with molecular formulas, retention times (RT), ions (with types, m/z and formulas indicated) in negative (ESI ) and positive (ESI ) electrospray ionisation mode detected by UHPLC-QTOF-MS/MS. For further details on metabolites see Table S1. þ ID Metabolite Molecular formula RT [min] average ESI - ESI

Ion type m/z Ion formula Ion type m/z Ion formula

- - 1 Benzyl glucosinolate C14H19NO9S2 5.35 [M-H] 408.0436 [C14H18NO9S2] - - þ þ 2 N-(Benzoyl) aspartic acid C11H11NO5 5.75 [M-H] 236.0556 [C11H10NO5] [MþH] 238.0717 [C11H12NO5] - - 3 4-Hydroxybenzyl glucosinolate C14H19NO10S2 2.30 [M-H] 424.0386 [C14H18NO10S2] - - þ þ 4 N-(4-Hydroxybenzoyl) aspartic acid C11H11NO6 3.20 [M-H] 252.0512 [C11H10NO6] [MþH] 254.0657 [C11H12NO6] - - þ þ 5 4-Hydroxybenzoic acid C7H6O3 4.13 [M-H] 137.0242 [C7H5O3] [MþH] 139.0389 [C7H7O3] - - þ þ 6 4-Hydroxybenzaldehyde C7H6O2 5.45 [M-H] 121.0294 [C7H5O2] [MþH] 123.0441 [C7H7O2] - - 7 2-Phenylethyl glucosinolate C15H21NO9S2 7.90 [M-H] 422.0583 [C15H20NO9S2] - - þ þ 8 N-(Phenylacetyl) aspartic acid C12H13NO5 6.78 [M-H] 250.0730 [C12H12NO5] [MþH] 252.0855 [C12H14NO5]

5 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

of fed with differently treated watercress leaves (Table 2). The main me­ tabolites 2, 4 and 8 were found in almost all samples of the corre­ Faeces leaves sponding bodies and faeces of P. cochleariae larvae which had ingested

on glucosinolates 1, 3, or 7, respectively (Table 2), but were not found in any of the plant samples (data not shown). 1 Body In addition to (4), metabolites with 137.0242 m/z (at 4.13 min, 5) applied and 121.0294 m/z (at 5.45 min, 6) were detected in samples of larvae

were fed with leaves treated with 4-hydroxybenzyl glucosinolate (3). These ) À 3 Gut ( compounds showed a common characteristic fragment in ESI mode - [93.0346 m/z ([C6H5O] ) (5, 6)]. Moreover, fragmentation of the major þ ion in ESI mode (i. e., 139.0389 m/z) of metabolite (5) revealed a þ dominant fragment [121.0284 m/z ([C H O ] )]. By comparison to 3 3 Pea Whole 7 5 2

glucosinolate reference standards, these metabolites were identifiedas 4-hydroxyben­ zoic acid (5) and 4-hydroxybenzaldehyde (6) (Tables 1 and S1). These compounds were detected in whole bodies and compound (5) also in the 2 2 3 Faeces gut, but they were not found in faeces (Table 2). Moreover, (5) was also

(3) detected in leaf samples of watercress and pea to which 4-hydroxybenzyl glucosinolate had been applied and traces of (6) were detected in 4-hydroxybenzyl 1 1 3 Body or watercress leaves with applied glucosinolates. ) 1 ( glucosinolate

b 3.2. Stability of applied glucosinolates on leaves 1 2 2 3 (1) Gut Glucosinolates applied to leaves were still intact after 23 h under the

glucosinolate conditions of the feeding tests, but without larvae. Recovery of the two b glucosinolates was high on both plant species, but somewhat lower for 3 2 3 1 3 2 4-Hydroxybenzyl Watercress Whole (1) benzyl glucosinolate (1) on leaves of watercress [89.7% (83.9–92.4), Benzyl .

a median (IQR), n ¼ 6)] compared to pea [100% (100–100); p ¼ 0.003, Mann-Whitney U-test]. 4-Hydroxybenzyl glucosinolate (3) was recov­

Faeces >

h. ered at 90% from leaves of both plant species [watercress: 100% detected – – ¼ 23 (100 100); pea: 100% (90.02 100), p 0.176]. were for 3 Body 3 3.3. Myrosinase activities in leaves and larval parts larvae As expected, leaves of watercress had a high myrosinase activity, metabolites Gut 3 whereas leaves of pea had no myrosinace activity (Table 3). In guts of mid second-instar larvae starved and fed subsequently with watercress, which likewise a high myrosinase activity was detectable. In starved mid second-instar in b second-instar larvae and larvae fed with pea leaves instead of watercress present. 3 Pea Whole 3 (2) after starvation, a rather low myrosinase activity was detected (Table 3). not faeces,

moulted In contrast, no myrosinase activity was detectable in guts of larvae that was

their had starved directly after moulting to the second instar or were treated it or 2 3 3 Faeces 3 freshly or

to Table 3 low Myrosinase activity in leaves of watercress and pea as well as in guts and

too remaining bodies of the second-instar (L2) larvae or in pupae of Phaedon coch­ 3 Body 3 replicates) offered 3 (1) leariae (three individuals pooled per sample). All larvae were fed with watercress been

and until moulting to the L2 stage. Freshly moulted L2 larvae were offered water­

(total cress or pea leaves after 8.5 h of starvation. Mid L2 larvae fed on watercress were 3 Gut 3 pea have starved for 3.5 h and then fed with either watercress or pea leaves. The plant or ) glucosinolate species in the columns refer to the final food provided for 24 h after the star­ may 7 vation period. cochleariae 3 3 Benzyl Watercress Whole 3 Sample Myrosinase activity, mean � SD (n ¼ 3) [nmol glucose μg À À protein 1 * min 1]

Phaedon Starved Watercress Pea concentration of glucosinolate, acid � a its Leaf 0.083 0.034 n.d. acid larvae

aspartic Mid L2 larvae (starved for 3.5 h) of) acid Guts 0.014b 0.038 � 0.032 0.012 � 0.009 acid detected, glucosinolate aspartic Bodies (without gut) n.d. n.d. 2-phenylethyl glucosinolate (parts not aspartic of Freshly-moulted L2 larvae (starved for 8.5 h) was b � contaminations. glucosinolate Guts n.d. 0.052 0.032 n.d. Bodies (without gut) n.d. n.d. (containing (Benzoyl) samples -(4-Hydroxybenzoyl) -(Phenylacetyl) likely N 4-Hydroxybenzoic 4-Hydroxybenzaldehyde 2-Phenylethyl N Benzyl N- 4-Hydroxybenzyl of b

2 Pupae n.d. compound Most If a n.d. – no activity detectable. 4 5 6 7 8 1 Treatments Metabolites 2 3 a b b watercress Table Number Only 1 replicate measured.

6 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431 the same but allowed to feed on pea leaves after the starvation period. Likewise, in remaining larval bodies without guts as well as in pupae no myrosinase activity could be detected (Table 3).

4. Discussion

Feeding experiments with P. cochleariae larvae and subsequent chemical analysis support our hypothesis that P. cochleariae evolved a way that is different from the known glucosinolate metabolism path­ ways in other insect species (Abdalsamee et al., 2014; Jeschke et al., 2016; Winde and Wittstock, 2011) to deal with the ‘mustard oil bomb’. To investigate the metabolism of glucosinolates in P. cochleariae larvae, we offered larvae leaves to which glucosinolates had been added. After almost one day, most of the glucosinolates that had been applied to the leaves were still intact in a control experiment without larvae. Thus, the glucosinolate solutions were quite stable and metabolites of gluco­ sinolates found in larval and faecal samples can predominantly be assigned to processes that occurred in the larvae. Slightly lower recovery from leaves of watercress than from pea may be related to the presence of myrosinases only in the former. Due to different feeding amounts of leaves by the individual larvae, we had to pool three larvae per sample. Therefore, we focus on qualitative rather than quantitative analyses of the glucosinolate metabolism in P. cochleariae. Comparative metabolomics analysis of insect samples from our feeding experiments revealed metabolites whose presence depended on the kind of glucosinolate ingested by the larvae. For each of the tested glucosinolates (1, 3, 7), one specific major metabolite (2, 4, 8) was detected in almost all samples derived from the corresponding larvae and faeces, but none of those were found in any of the plant samples. Hence, these metabolites are likely formed by larval metabolism of glucosinolates or of plant-derived glucosinolate breakdown products. The common difference of about À 172 m/z between the ions of the glucosinolates and their main metabolites suggests that these three benzenic glucosinolates undergo the same metabolic pathway in larvae of P. cochleariae (Fig. 2). The main metabolites were identified as N- (benzoyl) aspartic acid (2) derived from benzyl glucosinolate (1), N-4- (hydroxybenzoyl) aspartic acid (4) derived from 4-hydroxybenzyl glu­ cosinolate (3) and N-(phenylacetyl) aspartic acid (8) derived from 2- phenylethyl glucosinolate (7). The structure of N-(benzoyl) aspartic acid (2) was confirmed by NMR spectroscopy (Fig. S1). Thus, larvae of P. cochleariae obviously conjugate the amino acid aspartic acid to aro­ matic acids derived from ingested benzenic glucosinolates. A similar route of benzenic glucosinolate metabolism was identifiedin caterpillars of P. rapae, where glycine serves as conjugation partner (Vergara et al., 2006, 2007). A conjugation of glucosinolate-derived metabolites with aspartic acid, as found in the current study in larvae of P. cochleariae, is, to our knowledge, not known from any other species feeding on Brassicaceae. Fig. 3. Postulated metabolism of 4-hydroxybenzyl glucosinolate (3) in larvae of While in feeding experiments with benzyl- and 2-phenylethyl glu­ Phaedon cochleariae. Possible routes of metabolism were deduced from experi­ cosinolate (1, 7) only one major metabolite for each glucosinolate was ments in which larvae ingested leaflets of watercress or pea to which this found, two further metabolites were detected in bodies, but not faeces, glucosinolate had been applied. The initial breakdown of the glucosinolate to of P. cochleariae, when larvae had ingested 4-hydroxybenzyl glucosi­ the nitrile or isothiocyanate is followed by reactions leading to formation of 4- nolate (3) (Table 2). These metabolites were identifiedas 4-hydroxyben­ hydroxybenzoic acid, which is conjugated with aspartic acid. Metabolites that were found in the current study are shown in black. Hypothetic intermediates zoic acid (5) and 4-hydroxybenzaldehyde (6) and might represent are shown in grey. pathway intermediates (Tables 1 and S1; Fig. 3). Resulting from hy­ drolysis of 4-hydroxybenzyl glucosinolate, 4-hydroxybenzyl isothiocy­ Strikingly, neither isothiocyanates nor nitriles were detected in any of anate may be hydrolysed in P. cochleariae into 4-hydroxybenzyl alcohol the samples, indicating rapid conversion to other products. and subsequently converted into 4-hydroxybenzaldehyde (6) and then In caterpillars of P. rapae, ingestion of benzyl glucosinolate- into 4-hydroxybenzoic acid (5). Further steps of amino acid conjugation containing plant material leads from phenylacetonitrile, the product of may then take place. Alternatively, 4-hydroxybenzyl glucosinolate may joint action of plant myrosinases and larval NSP, via α-hydroxylation react to 4-hydroxyphenylacetonitrile, 4-hydroxymandelonitrile and and cyanide release to benzaldehyde. After oxidation to benzoic acid, a finally to 4-hydroxybenzylaldehyde by splitting off a cyanide, followed conjugation with glycine takes place (Stauber et al., 2012; Wittstock by the above-described pathway (Fig. 3). Thus, the metabolites detected et al., 2004). Formation of nitriles upon myrosinase-catalysed glucosi­ in larval samples of P. cochleariae after ingestion of benzenic glucosi­ nolate hydrolysis has been observed at low pH (<5) or high iron con­ nolates on watercress or pea leaves could arise from both iso­ centrations (>0.01 mM) as well as upon catalysis by plant or insect NSPs thiocyanates or nitriles as initial glucosinolate breakdown metabolites.

7 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

(Burow et al., 2006; Lambrix et al., 2001; Vaughn and Berhow, 2005; samples without gut (Table 2). In the present experiments, large Wittstock et al., 2016). However, when watercress leaves are homoge­ amounts of the glucosinolates were applied on the leaves, which may not nised in water, 2-phenylethyl glucosinolate (7) is hydrolysed to the have been fully converted by P. cochleariae. Thus, there is no clear evi­ corresponding isothiocyanate (Stauber et al., 2012). In leaf homoge­ dence that larvae of P. cochleariae sequester these glucosinolates. In nates of Sinapis species (Brassicaceae) with endogenous 4-hydroxyben­ contrast, two other chrysomelid beetle species that are specialists on zyl glucosinolate, this glucosinolate is converted to Brassicaceae, P. striolata and P. chrysocephala, selectively sequester 4-hydroxybenzylalcohol via spontaneous decomposition of the corre­ various glucosinolates from their host plants (Beran et al., 2014, 2018). À sponding isothiocyanate under release of a thiocyanate ion (SCN ) This illustrates nicely the diversity of insect mechanisms directed at À (Agerbirk et al., 2008). However, at least in plant homogenates, SCN coping with the plant glucosinolate-myrosinase system, even within one formation has not been observed with e.g. benzyl glucosinolate, as this beetle family. and similar glucosinolates give rise to more stable isothiocyanates than In summary, P. cochleariae has evolved another way of dealing with 4-hydroxybenzyl glucosinolate (Agerbirk et al., 2008). Future research the ‘mustard oil bomb’. Larvae metabolise glucosinolate-derived com­ is needed to test whether benzenic isothiocyanates may be decomposed pounds, likely isothiocyanates or nitriles, to aromatic acids, which they À accompanied by SCN release upon ingestion of the corresponding then conjugate with aspartic acid. The glucosinolate metabolism seems glucosinolates by P. cochleariae. thereby not necessarily to be dependent on plant myrosinases. Future The pathways discussed above imply that the ingested glucosinolates studies are needed to test whether the proposed metabolic pathway also are hydrolysed by myrosinases from either plant or insect. Unexpect­ applies to other glucosinolates and to further elucidate the postulated edly, the major metabolites 2, 4 and 8 were also found in freshly- pathway. moulted larvae of P. cochleariae fed with glucosinolate-treated leaves of pea (Table 2) that lack myrosinases. A low myrosinase activity was Declaration of competing interest found in guts of second-instar larvae that were fed with watercress, starved and then fed with pea. In contrast, no myrosinase activity was None. found in guts of second-instar larvae that had starved directly after moult and then fed with pea. Insects appear to empty their guts entirely Acknowledgements before each moult (Beament, 1968). This may explain the lack of detectable myrosinase activity in freshly moulted larvae feeding on pea We thank the gardeners of Bielefeld University for help in plant and indicate that the myrosinase activity found in mid second-instar rearing, Laura Lachenicht and Martin Zumbrunnen for their help in in­ larvae that were switched from watercress to pea may be due to some sect rearing, Leon Brüggemann and Sarah Rhode for their help in con­ remaining myrosinase activity from watercress tissue. ducting some of the experiments. This work was funded by the grant MU Based on these data, plant myrosinases likely contributed to gluco­ 1829/20–1 to CM of the Deutsche Forschungsgemeinschaft. sinolate hydrolysis when P. cochleariae larvae fed on watercress leaves. In addition, the larvae may induce an own myrosinase once they feed on Appendix A. Supplementary data glucosinolate-containing plants, as known for the induction of a gluco­ sinolate sulfatase upon feeding on Brassicaceae in S. gregaria (Falk and Supplementary data to this article can be found online at https://doi. Gershenzon, 2007). Alternatively, larvae of P. cochleariae may carry org/10.1016/j.ibmb.2020.103431. microorganisms in their digestive system that contribute to glucosino­ late hydrolysis and may be responsible for the formation of iso­ thiocyanates and/or nitriles. In fact, bacteria that convert glucosinolates References to isothiocyanates or nitriles have been found in the guts of humans and Abdalsamee, M.K., Giampa, M., Niehaus, K., Müller, C., 2014. Rapid incorporation of rats (Liou et al., 2020; Liu et al., 2017; Mullaney et al., 2013). Microbial glucosinolates as a strategy used by a herbivore to prevent activation by symbionts offer several benefitsfor insects, because they improve insect myrosinases. Insect Biochem. Mol. Biol. 52, 115–123. nutrition by producing essential amino acids, vitamins and digestive Abdalsamee, M.K., Müller, C., 2012. Effects of indole glucosinolates on performance and sequestration by the sawfly Athalia rosae and consequences of feeding on the plant enzymes and are involved in the detoxificationor modificationof plant defence system. J. Chem. Ecol. 38, 1366–1375. allelochemicals (Beran and Gershenzon, 2016; Douglas, 2009; Genta Agerbirk, N., Warwick, S.I., Hansen, P.R., Olsen, C.E., 2008. Sinapis phylogeny and et al., 2006; van den Bosch and Welte, 2017). Larvae of Delia radicum evolution of glucosinolates and specific nitrile degrading enzymes. Phytochemistry 69, 2937–2949. (Diptera, Anthomyiidae) harbour gut bacteria that are able to hydrolyse Agnihotri, A.R., Hulagabali, C.V., Adhav, A.S., Joshi, R.S., 2018. Mechanistic insight in isothiocyanates (Welte et al., 2016). Whether and which microorgan­ potential dual role of sinigrin against Helicoverpa armigera. Phytochemistry 145, isms are involved in glucosinolate metabolism of P. cochleariae needs 121–127. Aliabadi, A., Renwick, J.A.A., Whitman, D.W., 2002. Sequestration of glucosinolates by further investigation. harlequin bug Murgantia histrionica. J. Chem. Ecol. 28, 1749–1762. Noticeably, the glucosinolate-derived metabolites did not only occur Beament, J.W.L., 1968. Advances in Insect Physiology. Academic Press Inc., London. in the guts and faeces of P. cochleariae but could also be detected in Bednarek, P., Pi�slewska-Bednarek, M., Svato�s, A., Schneider, B., Doubský, J., Mansurova, M., Humphry, M., Consonni, C., Panstruga, R., Sanchez-Vallet, A., remaining body parts (Table 2). This may indicate that initial steps of Molina, A., Schulze-Lefert, P., 2009. A glucosinolate metabolism pathway in living glucosinolate metabolism happen in the gut, while resulting metabolites plant cells mediates broad-spectrum antifungal defense. Science 323, 101–106. may be taken up through the gut wall, where they may be conjugated Beran, F., Gershenzon, J., 2016. Microbes matter: herbivore gut endosymbionts play a – with aspartic acid in the fat body or Malpighian tubules. The Malpighian role in breakdown of host plant toxins. Environ. Microbiol. 18, 1306 1307. Beran, F., Pauchet, Y., Kunert, G., Reichelt, M., Wielsch, N., Vogel, H., Reinecke, A., tubules represent an excretory and osmoregulatory system in insects and Svato�s, A., Mewis, I., Schmid, D., Ramasamy, S., Ulrichs, C., Hansson, B.S., are involved in excretion of toxic plant allelochemicals (Discher et al., Gershenzon, J., Heckel, D.G., 2014. Phyllotreta striolata flea beetles use host plant 2009; Wall et al., 1975). Preliminary images of dissected larvae of defense compounds to create their own glucosinolate-myrosinase system. Proc. Natl. Acad. Sci. U.S.A. 111, 7349–7354. P. cochleariae obtained by matrix-assisted laser desorption ionisation Beran, F., Sporer, T., Paetz, C., Ahn, S.J., Betzin, F., Kunert, G., Shekhov, A., Vassao,~ D.G., mass spectrometry imaging revealed indeed increased abundances of Bartram, S., Lorenz, S., Reichelt, M., 2018. One pathway is not enough: the cabbage N-(benzoyl) aspartic acid (4) and N-(phenylacetyl) aspartic acid (8) in stem flea beetle Psylliodes chrysocephala uses multiple strategies to overcome the glucosinolate-myrosinase defense in its host plants. Front. Plant Sci. 9. the Malpighian tubules (data not shown). Bla�zevi�c, I., Montaut, S., Bur�cul, F., Olsen, C.E., Burow, M., Rollin, P., Agerbirk, N., 2020. The applied glucosinolates (1, 3) and the major glucosinolate of Glucosinolate structural diversity, identification,chemical synthesis and metabolism watercress, 2-phenylethyl glucosinolate (7), were also found unmodified in plants. Phytochemistry 169, 112100. Bradford, M.M., 1976. A rapid and sensitive method for the quantificationof microgram in different insect parts and faeces, but there was no consistent accu­ quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. mulation of the respective glucosinolates in whole larvae and body 72, 248–254.

8 J. Friedrichs et al. Insect Biochemistry and Molecular Biology 124 (2020) 103431

Bruce, T.J.A., 2014. Glucosinolates in oilseed rape: secondary metabolites that influence Nielsen, J.K., 1978. Host plant discrimination within Cruciferae: feeding responses of four interactions with herbivores and their natural enemies. Ann. Appl. Biol. 164, leaf beetles (Coleoptera: Chrysomelidae) to glucosinolates, cucurbitacins and 348–353. cardenolides. Entomol. Exp. Appl. 24, 41–54. Burow, M., Markert, J., Gershenzon, J., Wittstock, U., 2006. Comparative biochemical Opitz, S.E.W., Mix, A., Winde, I.B., Müller, C., 2011. Desulfation followed by sulfation: characterization of nitrile-forming proteins from plants and insects that alter metabolism of benzylglucosinolate in Athalia rosae (Hymenoptera: Tenthredinidae). myrosinase-catalysed hydrolysis of glucosinolates. FEBS J. 273, 2432–2446. Chembiochem 12, 1252–1257. Discher, S., Burse, A., Tolzin-Banasch, K., Heinemann, S.H., Pasteels, J.M., Boland, W., Pentzold, S., Zagrobelny, M., Roelsgaard, P.S., Moller, B.L., Bak, S., 2014a. The multiple 2009. A versatile transport network for sequestering and excreting plant glycosides strategies of an insect herbivore to overcome plant cyanogenic glucoside defence. in leaf beetles provides an evolutionary flexible defense strategy. Chembiochem 10, PloS One 9. 2223–2229. Pentzold, S., Zagrobelny, M., Rook, F., Bak, S., 2014b. How insects overcome two- Dobler, S., Petschenka, G., Pankoke, H., 2011. Coping with toxic plant compounds - the component plant chemical defence: plant beta-glucosidases as the main target for insect’s perspective on iridoid glycosides and cardenolides. Phytochemistry 72, herbivore adaptation. Biol. Rev. 89, 531–551. 1593–1604. Petersen, B.L., Chen, S., Hansen, C.H., Olsen, C.E., Halkier, B.A., 2002. Composition and Douglas, A.E., 2009. The microbial dimension in insect nutritional ecology. Funct. Ecol. content of glucosinolates in developing Arabidopsis thaliana. Planta 214, 562–571. 23, 38–47. R Developmental Core Team, 2020. A language and environment for statistical Falk, K.L., Gershenzon, J., 2007. The desert locust, Schistocerca gregaria, detoxifies the computing. R Foundation for Statistical Computing. glucosinolates of Schouwia purpurea by desulfation. J. Chem. Ecol. 33, 1542–1555. Ratzka, A., Vogel, H., Kliebenstein, D.J., Mitchell-Olds, T., Kroymann, J., 2002. Francis, F., Lognay, G., Wathelet, J.P., Haubruge, E., 2001. Effects of allelochemicals Disarming the mustard oil bomb. Proc. Natl. Acad. Sci. U.S.A. 99, 11223–11228. from first (Brassicaceae) and second (Myzus persicae and Brevicoryne brassicae) Reifenrath, K., Müller, C., 2008. Multiple feeding stimulants in Sinapis alba host plants for trophic levels on Adalia bipunctata. J. Chem. Ecol. 27, 243–256. the oligophagous leaf beetle Phaedon cochleariae. Chemoecology 18, 19–27. Genta, F.A., Dillon, R.J., Terra, W.R., Ferreira, C., 2006. Potential role for gut microbiota Reifenrath, K., Riederer, M., Müller, C., 2005. Leaf surface wax layers of Brassicaceae in cell wall digestion and glucoside detoxificationin Tenebrio molitor larvae. J. Insect lack feeding stimulants for Phaedon cochleariae. Entomol. Exp. Appl. 115, 41–50. Physiol. 52, 593–601. Ruttkies, C., Schymanski, E.L., Wolf, S., Hollender, J., Neumann, S., 2016. MetFrag Halkier, B.A., 2016. General introduction to glucosinolates. Adv. Bot. Res. 80, 1–14. relaunched: incorporating strategies beyond in silico fragmentation. J. Cheminf. 8. Hammer, T.J., Bowers, M.D., 2015. Gut microbes may facilitate insect herbivory of Schramm, K., Vassao,~ D.G., Reichelt, M., Gershenzon, J., Wittstock, U., 2012. Metabolism chemically defended plants. Oecologia 179, 1–14. of glucosinolate-derived isothiocyanates to glutathione conjugates in generalist Heckel G, David, 2014. Insect detoxification and sequestration strategies. Annual Plant lepidopteran herbivores. Insect Biochem. Mol. Biol. 42, 174–182. Reviews 47, 77–114. Schrieber, K., Schweiger, R., Kroner,€ L., Müller, C., 2019. Inbreeding diminishes Jeschke, V., Gershenzon, J., Vassao,~ D.G., 2016. Insect detoxification of glucosinolates herbivore-induced metabolic responses in native and invasive plant populations. and their hydrolysis products. Adv. Bot. Res. 80, 199–245. J. Ecol. 107, 923–936. Kazana, E., Pope, T.W., Tibbles, L., Bridges, M., Pickett, J.A., Bones, A.M., Powell, G., Stauber, E.J., Kuczka, P., van Ohlen, M., Vogt, B., Janowitz, T., Piotrowski, M., Rossiter, J.T., 2007. The cabbage aphid: a walking mustard oil bomb. Proc. Biol. Sci. Beuerle, T., Wittstock, U., 2012. Turning the ’Mustard oil bomb’ into a ’Cyanide 274, 2271–2277. bomb’: aromatic glucosinolate metabolism in a specialist insect herbivore. PloS One Kim, J.H., Jander, G., 2007. Myzus persicae (green peach aphid) feeding on Arabidopsis 7. induces the formation of a deterrent indole glucosinolate. Plant J. 49, 1008–1019. Thangstad, O.P., Gilde, B., Chadchawan, S., Seem, M., Husebye, H., Bradley, D., Kissen, R., Rossiter, J.T., Bones, A., 2009. The ‘mustard oil bomb’: not so easy to Bones, A.M., 2004. Cell specific, cross-species expression of myrosinases in Brassica assemble?! Localization, expression and distribution of the components of the napus, Arabidopsis thaliana and Nicotiana tabacum. Plant Mol. Biol. 54, 597–611. myrosinase enzyme system. Phytochemistry Rev. 8, 69–86. Travers-Martin, N., Kuhlmann, F., Müller, C., 2008. Revised determination of free and Kühnle, A., Müller, C., 2011. Responses of an oligophagous beetle species to rearing for complexed myrosinase activities in plant extracts. Plant Physiol. Biochem. 46, several generations on alternative host plant species. Ecol. Entomol. 36, 125–134. 506–516. Lambrix, V.M., Reichelt, M., Mitchell-Olds, T., Kliebenstein, D.J., Gershenzon, J., 2001. van den Bosch, T.J.M., Welte, C.U., 2017. Detoxifying symbionts in agriculturally The Arabidopsis epithiospecifierprotein promotes the hydrolysis of glucosinolates to important pest insects. Microb. Biotechnol. 10, 531–540. nitriles and influences Trichoplusia ni herbivory. Plant Cell 13, 2793–2807. Vaughn, S.F., Berhow, M.A., 2005. Glucosinolate hydrolysis products from various plant Liou, C.S., Sirk, S.J., Diaz, C.A.C., Klein, A.P., Fischer, C.R., Higginbottom, S.K., Erez, A., sources: pH effects, isolation, and purification. Ind. Crop. Prod. 21, 193–202. Donia, M.S., Sonnenburg, J.L., Sattely, E.S., 2020. A metabolic pathway for Vergara, F., Svato�s, A., Schneider, B., Reichelt, M., Gershenzon, J., Wittstock, U., 2006. activation of dietary glucosinolates by a human gut symbiont. Cell 180, 717–728. Glycine conjugates in a lepidopteran insect herbivore - the metabolism of Liu, X.J., Wang, Y.L., Hoeflinger, J.L., Neme, B.P., Jeffery, E.H., Miller, M.J., 2017. benzylglucosinolate in the cabbage white butterfly, Pieris rapae. Chembiochem 7, Dietary broccoli alters rat cecal microbiota to improve glucoraphanin hydrolysis to 1982–1989. bioactive isothiocyanates. Nutrients 9. Vergara, F., Svato�s, A., Schneider, B., Reichelt, M., Gershenzon, J., Wittstock, U., 2007. Malka, O., Shekhov, A., Reichelt, M., Gershenzon, J., Vassao,~ D.G., Morin, S., 2016. Erratum: Glycine conjugates in a lepidopteran insect herbivore - the metabolism of Glucosinolate desulfation by the phloem-feeding insect Bemisia tabaci. J. Chem. Ecol. benzylglucosinolate in the cabbage white butterfly, Pieris rapae. Chembiochem 8, 42, 230–235. 1757. Martin, N., Müller, C., 2007. Induction of plant responses by a sequestering insect: Wadleigh, R.W., Yu, S.J., 1988. Detoxification of isothiocynante allelochemicals by relationship of glucosinolate concentration and myrosinase activity. Basic Appl. glutathione transferase in 3 lepidopterous species. J. Chem. Ecol. 14, 1279–1288. Ecol. 8, 13–25. Wall, B.J., Oschman, J.L., Schmidt, B.A., 1975. Morphology and function of malpighian Matile, P., 1980. The mustard oil bomb - compartmentation of the myrosinase system. tubules and associated structures in the cockroach, Periplaneta americana. Biochem. Physiol. Pflanz. (BPP) 175, 722–731. J. Morphol. 146, 265–306. Mithofer,€ A., Boland, W., 2012. Plant defense against herbivores: chemical aspects. Welte, C.U., de Graaf, R.M., van den Bosch, T.J.M., op den Camp, H.J.M., van Dam, N.M., Annu. Rev. Plant Biol. 63, 431–450. Jetten, M.S.M., 2016. Plasmids from the gut microbiome of cabbage root fly larvae Morant, A.V., Jørgensen, K., Jørgensen, C., Paquette, S.M., Sanchez-P� erez,� R., Møller, B. encode SaxA that catalyses the conversion of the plant toxin 2-phenylethyl L., Bak, S., 2008. beta-glucosidases as detonators of plant chemical defense. isothiocyanate. Environ. Microbiol. 18, 1379–1390. Phytochemistry 69, 1795–1813. Winde, I., Wittstock, U., 2011. Insect herbivore counteradaptations to the plant Mullaney, J.A., Kelly, W.J., McGhie, T.K., Ansell, J., Heyes, J.A., 2013. Lactic acid glucosinolate-myrosinase system. Phytochemistry 72, 1566–1575. bacteria convert glucosinolates to nitriles efficiently yet differently from Wittstock, U., Agerbirk, N., Stauber, E.J., Olsen, C.E., Hippler, M., Mitchell-Olds, T., Enterobacteriaceae. J. Agric. Food Chem. 61, 3039–3046. Gershenzon, J., Vogel, H., 2004. Successful herbivore attack due to metabolic Müller, C., Agerbirk, N., Olsen, C.E., Boeve,� J.-L., Schaffner, U., Brakefield, P.M., 2001. diversion of a plant chemical defense. Proc. Natl. Acad. Sci. U.S.A. 101, 4859–4864. Sequestration of host plant glucosinolates in the defensive hemolymph of the sawfly Wittstock, U., Kurzbach, E., Herfurth, A.M., Stauber, E.J., 2016. Glucosinolate Athalia rosae. J. Chem. Ecol. 27, 2505–2516. breakdown. Adv. Bot. Res. 80, 125–169. Neubauer, C., Heitmann, B., Müller, C., 2014. Biofumigation potential of Brassicaceae cultivars to Verticillium dahliae. Eur. J. Plant Pathol. 140, 341–352.

9