Skip to main content
Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Flavin-Dependent Monooxygenases as a Detoxification Mechanism in Insects: New Insights from the Arctiids (Lepidoptera)

  • Sven Sehlmeyer ,

    Contributed equally to this work with: Sven Sehlmeyer, Linzhu Wang

    Affiliation Institute for Pharmaceutical Biology, TU Braunschweig, Braunschweig, Germany

  • Linzhu Wang ,

    Contributed equally to this work with: Sven Sehlmeyer, Linzhu Wang

    Affiliation Biochemical Ecology and Molecular Evolution, Botanical Institute and Botanical Garden, Christian-Albrechts-Universität, Kiel, Germany

  • Dorothee Langel,

    Affiliation Biochemical Ecology and Molecular Evolution, Botanical Institute and Botanical Garden, Christian-Albrechts-Universität, Kiel, Germany

  • David G. Heckel,

    Affiliation Department of Entomology, Max Planck Institute for Chemical Ecology, Jena, Germany

  • Hoda Mohagheghi,

    Affiliation Institute for Pharmaceutical Biology, TU Braunschweig, Braunschweig, Germany

  • Georg Petschenka,

    Affiliation Molecular Evolution, Institute of Zoology, University of Hamburg, Hamburg, Germany

  • Dietrich Ober

    Affiliation Biochemical Ecology and Molecular Evolution, Botanical Institute and Botanical Garden, Christian-Albrechts-Universität, Kiel, Germany


Insects experience a wide array of chemical pressures from plant allelochemicals and pesticides and have developed several effective counterstrategies to cope with such toxins. Among these, cytochrome P450 monooxygenases are crucial in plant-insect interactions. Flavin-dependent monooxygenases (FMOs) seem not to play a central role in xenobiotic detoxification in insects, in contrast to mammals. However, the previously identified senecionine N-oxygenase of the arctiid moth Tyria jacobaeae (Lepidoptera) indicates that FMOs have been recruited during the adaptation of this insect to plants that accumulate toxic pyrrolizidine alkaloids. Identification of related FMO-like sequences of various arctiids and other Lepidoptera and their combination with expressed sequence tag (EST) data and sequences emerging from the Bombyx mori genome project show that FMOs in Lepidoptera form a gene family with three members (FMO1 to FMO3). Phylogenetic analyses suggest that FMO3 is only distantly related to lepidopteran FMO1 and FMO2 that originated from a more recent gene duplication event. Within the FMO1 gene cluster, an additional gene duplication early in the arctiid lineage provided the basis for the evolution of the highly specific biochemical, physiological, and behavioral adaptations of these butterflies to pyrrolizidine-alkaloid-producing plants. The genes encoding pyrrolizidine-alkaloid-N-oxygenizing enzymes (PNOs) are transcribed in the fat body and the head of the larvae. An N-terminal signal peptide mediates the transport of the soluble proteins into the hemolymph where PNOs efficiently convert pro-toxic pyrrolizidine alkaloids into their non-toxic N-oxide derivatives. Heterologous expression of a PNO of the generalist arctiid Grammia geneura produced an N-oxygenizing enzyme that shows noticeably expanded substrate specificity compared with the related enzyme of the specialist Tyria jacobaeae. The data about the evolution of FMOs within lepidopteran insects and the functional characterization of a further member of this enzyme family shed light on this almost uncharacterized detoxification system in insects.


Flavin-dependent monooxygenases (FMOs) and cytochrome P450 monooxygenases (CYPs) are two prominent families of monooxygenases in eukaryotes [1], [2]. They catalyze the transfer of one atom of molecular oxygen to a substrate and reduce the other to water. FMO genes are found in all phyla [3]. In vertebrates, FMOs form a gene family of five similar genes. They provide an efficient detoxification system for xenobiotics, as they catalyze the conversion of heteroatom-containing chemicals from the animal's food to polar, readily excretable metabolites [4]. Yeast possesses, unlike mammals, only one FMO isoform, which has been shown to be involved in redox regulation and in the correct folding of proteins containing disulfide bonds [5], [6]. In plants, FMOs form a large gene family (29 genes in the model plant Arabidopsis thaliana), but information about their physiological role is sparse. For A. thaliana, individual FMO sequences have been related to auxin biosynthesis and pathogen defense [7]. FMOs oxygenate nucleophilic substrates that usually contain nitrogen or sulfur, such as amines, amides, thiols, and sulfides [8]. A unique feature of FMO is the catalytic cycle that forms a reactive 4α-hydroperoxyflavin intermediate as a potent monooxygenating agent before the substrate is bound to the enzyme. Like a cocked gun, this activated intermediate will readily react with all substrates that are able to access the active site [9], [10].

Our extensive knowledge about the structural and catalytic properties of vertebrate FMOs is contrasted by an almost complete lack of knowledge about this enzyme family in insects. CYP enzymes play the dominant role in drug and xenobiotic metabolism in insects [11], possibly compensating the need for FMOs in these processes. Of note, the genome of Drosophila melanogaster contains only two genes for FMOs [1], [12] but 90 genes for CYPs, of which 86 seem to be functional [13]. The large number of CYPs in insect genomes has been suggested to be necessary to protect the insect from the diverse array of harmful compounds in its environment [14]. This is also the case for lepidopteran species. One of the best studied examples is the CYP gene superfamily of Papilio butterflies (Papilionidae), which have adapted to furanocoumarins, the toxic components of their food plants [15], [16]. These toxins are degraded by inducible CYPs that are expressed in the midgut and also in the fat body [17], [18].

Adaptation to host plant-derived toxins has also been described for the cinnabar moth, Tyria jacobaeae (Arctiidae). Larvae of this species feed exclusively on tansy ragwort (Jacobaea vulgaris, syn. Senecio jacobaea), which contains toxic pyrrolizidine alkaloids (PAs) and sequester these plant toxins for their own chemical defense. PAs can exist in two transmutable forms: the pro-toxic free base (tertiary amine) and its non-toxic N-oxide [19], [20]. In the plant, PAs usually occur as N-oxides, which are easily converted to their respective free base in the reducing gut milieu of any herbivorous vertebrate or insect feeding on these plants [21], [22]. The toxicity of PAs for non-adapted insects has been shown by feeding experiments [23] and is attributable to cytochrome P450-mediated bioactivation [24]. For Tyria, senecionine N-oxygenase (SNO), a soluble enzyme located in the hemolymph, has been shown to convert the pro-toxic free base efficiently into its non-toxic N-oxide [22] (Figure 1). This enzyme with high substrate specificity for toxic 1,2-unsaturated PAs, is the prerequisite for sequestration of the plant-derived alkaloids by the insect. Recently, SNO has been shown to be a FMO and, until now, the only functionally characterized FMO of insect origin [1].

Figure 1. N-oxygenation of PAs by SNO, a flavin-dependent monooxygenase (A) and structures of selected PAs and of atropine (B).

A: PAs are present in the plant mainly as N-oxide. After uptake in the diet, they are reduced in the gut of the herbivore to the their respective tertiary form, which is lipophilic and easily permeates membranes. In PA-adapted insects, this pro-toxic PA is efficiently converted to the respective PA N-oxide in the hemolymph to prevent bioactivation. B: Structures of PAs and atropine tested as substrates with recombinant SNO and PNO.

PA sequestration is known for many species of the tiger moth (Lepidoptera, Arctiidae), some of which use these alkaloids as a precursor for pheromone synthesis [25]. As shown for Utetheisa ornatrix, the PAs are acquired during the course of larval feeding and are transferred through metamorphosis to the adult stage [26]. At mating, the male advertises his PA load to the female by the PA-derived pheromone, hydroxydanaidal. Males with the highest PA load have the highest mating success and transfer a portion of their PAs via the spermatophore to the female [27], [28]. Together with the female's own load of PA, these alkaloids are passed to the eggs, protecting them against insect predators, including beetles, ants, and parasitoids [27], [29], [30].

In contrast to the specialist T. jacobaeae, the PA-sequestering arctiid species studied in this project are polyphagous. For Estigmene acrea, PAs have been shown to be important for development, as these alkaloids are used as precursors for the biosynthesis of the sex pheromone hydroxydanaidal [31], [32]. Grammia geneura is not known to synthesize PA-derived pheromones but benefits from sequestered PAs as defense compounds against herbivores and parasitoids [33].

Using an alignment of SNO of T. jacobaeae and of FMO-like sequences of the dipteran species D. melanogaster and Anopheles gambiae, we have been able to identify several new FMO sequences of arctiids and other Lepidoptera. With regard to the polyphagous species G. geneura, we have identified and expressed a PA-specific FMO in E. coli and compared its substrate specificity with the SNO of monophagous T. jacobaeae. Phylogenetic analysis shows that, in Lepidoptera, FMOs form three distinct sequence clusters. In one of these clusters, PA-specific FMO (PA N-oxygenases, PNO) originated by gene duplication early in the lineage of arctiids. These results allow a first glimpse into the hitherto untouched area of evolution and functionality of the FMO gene family in insects.

Materials and Methods


Larvae of Tyria jacobaeae were collected in The Netherlands and in the vicinity of Kiel, Germany. Larvae of Arctia caja, Arctia villica, and Diacrisia sannio were obtained from private breeders. Larvae of Estigmene acrea and Grammia geneura came from cultures established by Elisabeth A. Bernays and Michael S. Singer from specimens of field populations collected in southeastern Arizona, USA. All larvae were reared in the lab on a food plant mixture of Taraxacum officinale, Plantago lanceolata, and Rubus fruticosus or on an artificial diet [34], except for those of T. jacobaeae, which were exclusively reared on leaves of Senecio jacobaea.

Design of degenerate primers for identification of cDNAs coding for FMO-like sequences

cDNA sequences homologous to FMOs were identified in arctiid insects by a polymerase chain reaction (PCR) approach with degenerate primers. Primers P2, P3, and P8 (Table S1) were designed based on the alignment of amino acid sequences of the SNO of T. jacobaeae [1] and of four putative FMOs, two each of the genome of Drosophila melanogaster (Acc. No. AAF47118 and AAF57364) and of Anopheles gambiae (Acc. No. XP_311551 and XP_311550). Later in this project, the primers P12, P17, und P29 (Table S1) were designed according to alignments that resulted from the inclusion of the amino acid sequences of the FMO of A. caja and A. villica and of the PNO of Grammia geneura, as identified in this project.

Identification of cDNAs encoding FMO-like sequences of A. caja (AcFMO), A. villica (AvFMO), and T. jacobaeae (TjFMO)

The fat bodies of larvae of A. caja, A. villica, and T. jacobaeae were prepared and quickly frozen in liquid nitrogen. Total RNA was extracted by using the RNeasy Mini Kit (Qiagen) in combination with QIAshredder (Qiagen) mini spin columns. An aliquot containing 1 µg total RNA was reverse-transcribed with oligo-dT primer P1 (Table S1) by using Superscript III reverse transcriptase (Invitrogen). For identification of FMO-encoding cDNAs of A. caja and A. villica, a semi-nested PCR approach was used to amplify specific DNA fragments of ca. 900 bp in length with a constant annealing temperature of 52°C and Taq DNA polymerase (Invitrogen) in a total volume of 25 µl. For the first PCR, primer pair P2/P1 was used. The resulting reaction mix was diluted 1∶100 with 10 mM Tris/HCl buffer, pH 8, and used as a template for a PCR with primer pair P2/P3. The FMO-coding cDNA of T. jacobaeae was amplified in a nested PCR approach by using a touch-down temperature program with decreasing annealing temperature from 60°C to 45°C (0.5°C per cycle). After the first PCR with primer pair P2/P3, the reaction mix was diluted 1∶100 with 10 mM Tris/HCl buffer, pH 8, and used as template for the second PCR with primer pair P12/P29 resulting in a fragment of ca. 550 bp. For identification of the missing cDNA-ends, 3′-RACE (rapid amplification of cDNA ends) and 5′-RACE techniques were applied as described previously [35], [36] with primers P4-P7, P8-P11, and P30/P33 (Table S1) for the cDNA sequences of A. caja, A. villica, and T. jacobaeae, respectively. The proteins encoded by the resulting full-length sequences were denominated AcFMO, AvFMO, and TjFMO, respectively. For identification of the cDNA encoding PNO of A. caja (AcPNO), the same strategy was used as that described for the AcFMO with the following modifications: the primer pairs P2/P17 and P12/P17 were used with annealing temperatures of 54°C and 58°C for the first and second reaction of the semi-nested PCR approach, respectively. The 3′-RACE and 5′-RACE of the resulting fragment of ca. 600 bp were performed with the primers P18–P21. For identification of a cDNA encoding PNO of G. geneura, cDNA was prepared as described for FMO-like sequences of A. caja and used as template in a PCR with primer pair P12/P3 and a touch-down program with decreasing annealing temperature from 60°C to 45°C within 30 cycles. The cDNA fragment of ca. 800 bp was completed by 3′-RACE and 5′-RACE with primers P13–P16.

Partial identification of further FMO-like cDNAs of various Arctiids

For the identification of further FMO-like cDNA sequences of A. villica, Diacrisia sannio, and Estigmene acrea, the same strategies were used as those described above for the identification of the full-length cDNAs of various arctiids. The PCR parameters are given in Table S2.

Identification of FMO-like sequences of Helicoverpa armigera and Bombyx mori

FMO sequences of D. melanogaster and SNO of T. jacobaeae were used in tblastn searches of expressed sequence tag (EST) libraries of H. armigera and B. mori, and the genome sequence of B. mori at NCBI (, KaikoBase (, and SilkDB ( Sequences identified from B. mori included FMO1 (GenBank Accession No. GU564654), FMO2 (GU564656), and FMO3 (GU564657). Genomic locations of B.mori FMOs were determined by using the gbrowse genome browser of Kaikobase. Coding sequences of FMO1 from clone HA-GN-M-03-libF_K16 (GU564659) and FMO3 from clone HA-PAN-384-04-libF-F3_A05 (GU564662) were obtained from cDNA libraries constructed from H. armigera larvae by Heiko Vogel, Max Planck Institute for Chemical Ecology. The sequence of FMO2 from clone 28d18 (GU564660) was obtained from a cDNA library constructed from H. armigera larval midgut by Vladimir D. Grubor, University of Melbourne [37].

Sequence analysis

cDNA sequences were analyzed by using the ProtParam tool [38] for prediction of various protein properties, by using the signalP3.0 server [39] for prediction of signal peptides and their cleavage sites, by using the PSORT II and the TargetP1.1 server [40], [41] for the detection of sorting signals and subcellular localizations, and by using the TMHMM2.0 Server [42] for the prediction of transmembrane helices. To generate an alignment of amino acid sequences, ClustalX [43] was used before phylogenies were estimated with the following software of the PHYLIP program package [44]: PROTDIST with the Jones-Taylor-Thornton matrix in combination with NEIGHBOR for neighbor-joining analyses [45] and SEQBOOT and CONSENSE to estimate bootstrap values. Accession numbers of sequences taken from the database are given in Table S3.

Heterologous expression of cDNA encoding PNO of Grammia geneura

The open reading frame (ORF) of the PNO of Grammia geneura was analyzed for the presence of an N-terminal signal peptide by using the SignalP3.0 server [39]. For amplification of the whole ORF without the signal peptide, a pair of gene-specific primers was constructed that contained restriction sites for subcloning in addition to an artificial initiation codon (P34, NdeI) and the stop codon behind an artificial sequence encoding six histidine residues (P35, BamHI). The fragment resulting from amplification with AccuTaq LA DNA Polymerase (Sigma) of the oligo(dT)17 primed cDNA of G. geneura as the template was NdeI/BamHI-digested and ligated into a NdeI/BamHI-linearized pET3a vector (Novagen) for expression with the T7 polymerase system [46]. Positive clones were identified as described previously for SNO of T. jacobaeae [1]. Expression in E. coli BL21(DE3) (Stratagene) harboring the pRDKJG plasmid [47], was achieved in a modified lysogeny broth (LB) medium pH 7.5 containing 2.5 mM betaine hydrochloride and 0.8 M sorbitol at 4°C for 96 h.

Semiquantitative reverse transcription PCR

Per sample, 1 µg total RNA was used as a template for cDNA synthesis with an oligo-dT primer (P1) by using Superscript III RT (Invitrogen, Carlsbad, CA). PCR was performed with Taq DNA polymerase (Invitrogen) and the following temperature program: 4 min at 95°C initial denaturation, 46 cycles with an annealing temperature of 65°C, and elongation at 72°C for 2 min. The primer pairs specific for TjSNO (P36/P37) and TjFMO (P38/P39) resulted in fragments of ca. 1400 bp. Aliquots were taken at intervals of 3 cycles starting after cycle 28.

Enzyme assay

Enzyme activity of PNOs was assayed photometrically or by qualitative tracer assays as described previously [1]. Briefly, for the photometric assay, reactions were set up in a total volume of 300 µl 10 mM potassium phosphate pH 8 and 120 µM NADPH. After preincubation for 3 min at 37°C, the reaction was started by addition of 170 µM PA substrate and incubated at 37°C. The reaction was followed by the decrease of NADPH absorption at 340 nm with an Ultrospec 2100 pro UV/Visible Spectrophotometer (GE Healthcare). For the tracer assays, the volume of the reaction mixture was reduced to 50 µl. The amount of radioactively labeled PA N-oxide formed from tertiary PA was analyzed by thin-layer chromatography (TLC). Aliquots of 12.5 µl were therefore taken from the reaction mixture at time intervals between 3 to 30 min to ensure linearity of product formation and applied to silica gel 60 F254 TLC plates (Merck, Darmstadt). Enzyme activity was calculated from the substrate (PA)/product (PA N-oxide) ratio after separation by using the solvent system dichloromethane∶methanol∶ammonium hydroxide (25%) at a ratio of 80∶20∶3. Radioscans were performed by means of a radioactivity thin-layer analyzer (RITA, Raytest, Straubenhardt, Germany). PAs applied as enzyme substrates were obtained as described [22].


Identification of cDNA sequences encoding FMO-like proteins from Arctiids

For identification of cDNA sequences encoding FMOs from Arctiids, species were selected that were closely related to the specialist T. jacobaeae. These species included the polyphagous species Arctia caja, Diacrisia sannio, Estigmene acrea, and Grammia geneura, for all of which an association with PA-containing plants was previously described [48]. For the species Arctia villica, the PA association was confirmed by the ability to N-oxygenize tertiary PAs. Senecionine N-oxide was detected after incubation of the hemolymph with [14C]senecionine as the sole substrate in an assay for PNO activity (data not shown). For cDNA identification by a reverse transcription (RT)-PCR approach, an alignment of FMO-like sequences from insects was used to design degenerate primers. Two sequences identified within each of the genomes of Anopheles gambiae and Drosophila melanogaster, respectively, and the SNO of T. jacobaeae (TjSNO) were the only FMO-like sequences of insect origin that were available in this phase of our project. The degree of identity between these sequences at the amino acid level was between 33% and 56%. Conserved sequence motifs that were appropriate for primer design encompassed the FAD- and NADPH-binding sites (motifs 1 and 3 in Figure 2) and an additional conserved sequence stretch in the C-terminal part of the sequence (amino acid 323–329 of TjSNO). Later in this project, after identification of three FMO-like sequences from A. caja, A. villica, and G. geneura, we were able to improve the quality of the alignment and of the degenerate primers. Of all species, except for G. geneura, we identified two distinct cDNA sequences of which the derived amino acid sequences showed a high degree of identity to other FMO sequences in the database. The complete ORF was determined for the cDNAs encoding the PNO of G. geneura (GgPNO), a putative PNO of Arctia caja (AcPNO), and a FMO-like sequence of unknown function of each, T. jacobaeae (TjFMO) A. caja (AcFMO), and A. villica (AvFMO) (Table 1). In addition, partial sequences were isolated from Diacrisia sannio (DsFMO and DsPNO) and Estigmene acrea (EaFMO and EaPNO) and an additional sequence A. villica (AvPNO). Of those sequences for which we were able to identify the complete coding region, the properties of the cDNA and the encoded proteins are summarized in Table 1. Figure 2 shows that the ORFs of these sequences possess all three characteristic motifs of FMO sequences, i.e., the nucleotide-binding domains that stabilize the binding of FAD (consensus: GxGxxG) and NADPH (consensus: GxGxx(A/G)), respectively, and the FMO-identifying sequence (consensus: FxGxxxHxxx(F/Y)). Computer-based sequence analyses predicted N-terminal signal peptides for the vesicular pathway and a lack of a C-terminal membrane anchor, properties previously identified for the senecionine N-oxygenase of T. jacobaeae [1].

Figure 2. Alignment of the amino acid sequences of FMOs of various arctiid species.

Motif 1, FAD-binding site; motif 2, FMO-identifying sequence; motif 3, NADPH-binding site; motif 4, insertion of six amino acids characteristic for sequences belonging to the putative PNO cluster; TjSNO, T. jacobaeae SNO; GgPNO, G. geneura PNO; AcPNO, A. caja PNO; TjFMO, T. jacobaeae FMO; AcFMO, A. caja FMO; AvFMO, A. villica FMO.

Table 1. Characteristics of FMO-like sequences identified from three Arctiid species.

Heterologous expression of recombinant proteins

The heterologous expression of the SNO of T. jacobaeae resulted in the formation of an insoluble and inactive protein [1]. Despite the finding that protein solubilization and subsequent renaturation resulted in an active protein, the specific activity remained low. Therefore, we tried to improve the expression system by using Sf9 insect cells, a system based on pupal ovarian cells of Spodoptera frugiperda, a noctuid lepidopteran species related to the Arctiids. Using this system, we were able to detect active SNO in the medium supporting the functionality of the predicted N-terminal signal peptide, but the yield of protein was too low for further biochemical characterization. Insufficient protein yield was also the problem when using a yeast expression system (data not shown). Finally, by modifying a method described by Blackwell and Horgan [49], we succeeded in expressing PNO of G. geneura without the signal peptide in at least a partially soluble and active form. Therefore, we added betaine to the medium and promoted its uptake by osmotic stress by sorbitol. Further improvements were achieved by coexpression with the E. coli chaperones DnaK/DnaJ/GrpE by using the plasmid pRDKJG [47] and by reducing the expression temperature to 4°C. The purified PNO of G. geneura showed a specific activity of 55 nkat/mg with senecionine as substrate.

Substrate specificity of recombinant PNO of G. geneura

The substrate specificity of the PNO of the generalist G. geneura was characterized to compare it with the data described previously for SNO of the specialist T. jacobaeae [1], [22]. Therefore, PAs of the various structural types were tested in addition to some other alkaloids and to substrates of mammalian and yeast FMO (Table 2). The data show that all tested PAs, with exception of the otonecine derivative of senecionine, viz., senkirkine, were substrates for the PNO of G. geneura. The most obvious differences from the SNO of T. jacobaeae was the ability of the PNO to N-oxygenize phalaenopsine, a 1,2-saturated PA, and atropine. Dimethylaniline, as a typical substrate for mammalian FMO, necine bases, and nicotine was neither accepted by the specialist's enzyme of T. jacobaeae, nor by the generalist's enzyme of G. geneura. Of note, the enzyme of G. geneura showed a low but unequivocal activity with glutathione as substrate.

Table 2. Substrate specificity of nativeTjSNO and recombinant GgPNO.

Tissue-specific expression of FMOs in insects

The SNO of T. jacobaeae is a soluble protein present in the hemolymph [22]. As a signal peptide was identified at the N-terminus of the respective cDNA [1], a semiquantitative RT-PCR approach was used to identify the tissue expressing the transcript of PA-specific SNO in comparison with the FMO of unknown function. The larvae were dissected, and the various tissues used separately for total RNA extraction. As shown in Figure 3, the N-oxygenase transcript was detectable in the fat body, the tissue that synthesizes and secretes proteins of the hemolymph [50]. In addition, a signal was detectable in the integument that might have been attributable to contamination of the sample tissue by adhering fat body tissue, and a signal was detected in the head of the larvae. Cloning and sequencing of the PCR products of the head and the fat body sample revealed that both transcripts were identical at the nucleic acid level.

Figure 3. Tissue-specific expression of SNO (A) and FMO (B) of T. jacobaeae.

Semiquantitative reverse transcription PCR was performed with total RNA of various tissues of T. jacobaeae larvae. Aliquots were taken after 34 and 43 cycles of amplification with primers specific for TjSNO (A) and TjFMO (B), respectively. M, 100-bp DNA ladder (Fermentas) with the 1500-bp fragment labeled by an arrowhead; lane 1, head; lane 2, integument; lane 3, fat body; lane 4, hemolymph; lane 5, gut; lane 6, negative control (H2O); lane 7 and lane 8, positive controls (0.5 pg of plasmid carrying the full-length cDNA encoding TjSNO and TjFMO, respectively).

PA-specific N-oxygenases form a separate cluster within lepidopteran FMO1

For phylogenetic analysis, we used the arctiid sequences identified in this project in combination with selected FMO-like sequences from Lepidoptera available in the databases. EST sequences of Bicyclus anynana (Nymphalidae) and Plodia interpunctella (Pyralidae) were assembled to deduce the encoded amino acid sequences. Three FMO-like sequences were taken from the Bombyx mori (Bombycidae) genome database (SilkDB, Additionally, we included three sequences from Helicoverpa armigera (Noctuidae) that were identified from a cDNA library of larval midgut [37] and of entire larvae (H. Vogel, unpublished results), respectively. To avoid misleading results attributable to missing N-terminal and C-terminal ends of sequences, only the central part of the alignment was used in which gaps of unidentified sequences had to be filled with replacement characters within the sequences of P. interpunctella, B. anynana, and the FMO of E. acrea (EaFMO). The alignment is available as Figure S1. A neighbor-joining tree with two FMO-like sequences from the genome of Drosophila melanogaster as outgroup is shown in Figure 4. The branching pattern shows that lepidopteran FMO-like sequences occur in three well-supported clusters. The presence of one of the three FMO-like sequences of Bombyx mori in each of these clusters suggested that FMO in Lepidoptera formed a gene family of three members that we termed FMO1, FMO2, and FMO3 in analogy to the mammalian FMO gene family consisting of five members (FMO1 to FMO5) [51]. FMO3 sequences are only distantly related to lepidopteran FMO1 and FMO2. The closer relationship between FMO1 and FMO2 suggested by the tree is supported by the observation that both genes sit next to each other (tail-to-tail) on chromosome 25 of the Bombyx genome, indicating that they originated by a gene duplication event. All arctiid FMOs were identified within this project group with FMO1 and split into two distinct and well-supported clusters of paralogous sequences. In both clusters, the sequences identified from T. jacobaeae, i.e., TjSNO and TjFMO, respectively, were each sisters to all other sequences of the respective cluster. This branching pattern is supported by the classification of T. jacobaeae into the tribe Callimorphini (subfamily Arctiinae), in contrast to the other arctiid species of this study, which belong to the tribe Arctiini [52]. One of the two arctiid FMO clusters contained the functionally characterized SNO of T. jacobaeae [1] and the PNO of G. geneura.

Figure 4. Rooted neighbor-joining tree of amino acid sequences derived from cDNA encoding lepidopteran FMOs with two sequences of D. melanogaster as the outgroup.

The framed sequences were experimentally characterized as SNO and PNO, respectively, whereas the others should be regarded as putative FMO-coding cDNA. Branch lengths are proportional to the number of substitutions per site (scale: 0.1 substitutions per site). Bootstrap proportions resulted from 1000 replicates. Ac, Arctia caja; Av, Arctia villica; Ba, Bicyclus anynana; Bm, Bombyx mori; Dm, Drosophila melanogaster; Ds, Diacrisia sannio; Ea, Estigmene acrea; Gg, Grammia geneura; Ha, Helicoverpa armigera; Pi, Plodia interpunctella; Tj, Tyria jacobaeae.


Insects experience a wide array of chemical pressures from plant allelochemicals and pesticides and have developed several effective counterstrategies to cope with these toxins [53]. Among these, CYPs appear to have a key role in plant-insect interactions [11]. In mammals, the P450 system of xenobiotic detoxification is supplemented by the FMO gene family, which consists of five genes [54], [55]. Only recently, we have been able to identify and to characterize functionally the first FMO of insects. The cinnabar moth Tyria jacobaeae (Lepidoptera, Arctiidae), an insect specialized for plants containing toxic PAs, has evolved an FMO for the modification and storage of these plant-derived toxins [1]. Here, we describe the identification of several members of the FMO-gene family in lepidopteran species with a focus on sequences of the tiger moth family (Arctiidae), which recruited FMO-encoding genes for adaptation to PA-containing plants by means of a gene duplication early in its lineage. The invention of this new class of FMO was the prerequisite for these insects (i) to feed, unrivaled, on PA-containing plants, (ii) to convert these plant toxins to pro-toxins and to sequester them for their own chemical defense, and (iii) to use them in certain cases for the biosynthesis of sex pheromones.

Optimization of heterologous expression of soluble and active PNO in E. coli

Using an alignment of FMO sequences of D. melanogaster, A. gambiae, and T. jacobaeae for degenerate primer design, we have been able to identify five full-length cDNA sequences of arctiid FMOs (Table 1) in addition to several sequence fragments. All sequences have been classified as FMO, because of their sequence similarities and characteristic sequence motifs, i.e., two dinucleotide-binding signatures (Rossman folds) for FAD and NADP and the FMO-identifying motif FxGxxxHxxx(Y/F) [2], [56]. In addition to these three fingerprint sequences for FMO (motifs 1 to 3 in Figure 2), we have identified another motif (motif 4 in Figure 2) representing an insertion of six amino acids that is most characteristic for these sequences in the PNO cluster of Arctiid sequences (Figure 4). A primer constructed on this sequence motif (primer P17, Figure 2) has enabled us to restrict the amplification of FMO homologs to sequences belonging to this sequence cluster. We have used this strategy successfully for the identification of sequence fragments of the putative PNO of A. caja, A. villica, D. sannio, and E. acrea. The PNO identified from the generalist G. geneura is also characterized by this sequence insertion and has been selected for heterologous expression and functional characterization. In a previous study, attempts to express the SNO of T. jacobaeae heterologously in E. coli were hampered by the formation of inclusion bodies. For biochemical analysis, the inclusion bodies had to be solubilized under denaturing conditions followed by a renaturation procedure requiring several dialysis steps [1]. Nevertheless, the yield of active enzyme was low in comparison with activities observed for the native enzyme (0.5 nkat/mg in comparison to 77.6 nkat/mg [1]). As expression of GgPNO in E. coli also results in inclusion body formation, we have optimized the expression system. The supplementation of the medium with betaine in the presence of sorbitol, the coexpression of E. coli chaperons that have proved to be helpful previously [57], and a drastic reduction of the expression temperature to 4°C have all led to the expression of soluble and active protein. The specific activity of about 55 nkat/mg almost equals the value of 77.6 nkat/mg that has been described for the native SNO of Tyria jacobaeae [1].

GgPNO encodes a PA N-oxygenase with extended substrate specificity

The SNO of the specialist T. jacobaeae (TjSNO) is characterized by a high substrate specificity for PAs that are toxic because of the following structural features: (i) an 1,2-double bond in the ring system, (ii) an allylic esterified hydroxyl group at C9, and (iii) a free or esterified hydroxyl group at C9 [22]. The same high specificity has been established for the recombinant SNO that is heterologously expressed in E. coli [1]. In contrast, the PNO of G. geneura accepts a wider range of substrates. In addition to the alkaloids accepted by TjSNO, the 1,2-saturated PA phalaenopsine is N-oxygenized. The only PA that is not accepted by Grammia PNO and Tyria SNO is senkirkine, an otonecine derivative that cannot be N-oxygenized because of a methyl group at the ring-bound nitrogen. Feeding experiments of G. geneura have shown that senkirkine cannot be detoxified by N-oxidation and is neither sequestered nor metabolized [58]. 1,2-saturated PAs are devoid of the characteristic double bond, and therefore, they are regarded as non-toxic at least as far as bioactivation-mediated toxicity is concerned. However, the observations that 1,2-saturated PAs are accumulated in the plant preferentially in reproductive and young tissues [59], [60], [61] and that these structures might mediate antifeedant activity and neurotoxic effects [62], [63] suggest an ecological role for these alkaloids. The finding that phalaenopsine is N-oxygenized by Grammia PNO is in good agreement with the described ability of the larvae of G. geneura to sequester and metabolize these 1,2-saturated PAs into insect PAs. The observation that these insect PAs are transferred via metamorphosis to the adult stage has been interpreted as support for their ecological role for the insect, most probably in the chemical defense of the insect [58]. Of note, atropine, a tropane alkaloid produced by certain solanacous plants as part of their chemical defense against herbivores [64] is also accepted by Grammia PNO as a substrate. This wide substrate specificity is in accord with the finding that Grammia feeds as a generalist on a wide variety of food plants. Approximately 80 different species of about 50 taxonomically disparate families have been counted by Singer et al. [65], of which several are avoided by other generalist insects because of their toxicity, including species containing various types of PAs. Differences with respect to substrate specificity have also been described for CYPs as counterdefense enzymes of the specialist Papilio polyxenes, which feeds exclusively on furanocoumarin-containing plants and of the generalist Helicoverpa zea, which feeds on hundreds of types of host plant [66]. The toxicological challenge of generalized feeding is considerable with respect to the tremendous diversity of plant defense compounds. Specialization on a narrow range of host plants is interpreted as an adaptative strategy to plant toxins, involving more specialized detoxification enzymes [66]. Future research has to show the number and kind of enzymes that are involved, in addition to PNO, in the detoxification of plant-derived toxins in polyphagous Arctiids.

Remarkably, Grammia PNO also accepts glutathione, a substrate described for yeast FMO but that is not accepted by the SNO of Tyria. By oxidation of glutathione and other biological alcohols, yeast FMO provides the oxidizing equivalents that are essential for the proper folding of disulfide-containing proteins at the endoplasmic reticulum [67], [68]. Currently, we do not know whether this activity is a unique feature of the PNO of Grammia, or whether this conversion is an inherent activity of lepidopteran FMOs, suggesting a similar physiological role for lepidopteran FMOs as described for yeast and postulated for the FMOs of mammals and of Trypanosoma cruzi [69], [70]. The observation that the SNO of T. jacobaeae is expressed not only in the fat body, but also in the head of the larvae, suggests that the respective genes are pleiotropic and not only involved in PA N-oxygenation.

Duplication of a FMO-encoding gene as a key innovation in the Arctiids for adaptation to PA-containing plants

The identification of several lepidopteran FMO-like sequences in this project has enabled us to construct a neighbor-joining tree of this gene family present in this order of insects. The branching pattern of this tree shows three well-supported clusters that we have named FMO1 to FMO3, each containing one of the three FMO-coding sequences present in the Bombyx mori genome. Incorporation of EST sequence data of other lepidopteran species available in the databases supports these three clusters (data not shown). Two of these EST sequences that have shown the lowest degree of missing sequence in our alignment have been included in the phylogenetic analysis. The finding that the genomes of D. melanogaster and A. gambiae contain only two FMO-like sequences that do not cluster with FMO sequences of the Lepidoptera suggests that lepidopteran FMO form a lineage-specific group. A similar observation is described by Hao et al. [3] who has reconstructed a phylogeny of 104 FMO sequences of 34 species belonging to various metazoan phyla. The mammalian FMOs encompassing the five types FMO1–FMO5 show a monophyletic origin, well separated from the clades containing the fish FMO or the invertebrate FMO-like sequences. Therefore, the different lineages of animals do not have truly orthologous genes. Instead, diversification of the fmo genes occurred independently in the lineages by gene duplications, resulting in gene copies some of which were lost again, with others evolving different functions and metabolizing different substrates [3], [71]. Within the Lepidoptera, the gene duplication that resulted in the origin of FMO1 and FMO2 is well supported by the position of both genes close to each other on the chromosome 25 in the B. mori genome. Another gene duplication seems to be specific for the lineage of the Arctiids and has resulted in a cluster of FMO sequences of unknown function and in a separate cluster, encompassing sequences of which two have been shown to be involved in the N-oxygenation of plant-derived PAs, i.e., the SNO of T. jacobaeae and the PNO of G. geneura. Therefore, this gene duplication can be interpreted as a “key innovation” within this lineage according to the interpretation of Berenbaum et al. [72], it being the prerequisite for the evolution of the biochemical basis for the multifaceted adaptations of tiger moths to PA-containing plants. Indeed, in 1999, Weller et al. [52] postulated that the ability to sequester PAs from the larval diet should have arisen at an ancestral node early in the Arctiid family.

Of note, all FMO-like sequences that have been identified from arctiid species can be grouped into the two arctiid-specific clusters within the FMO1 group. No single sequence has been identified that clusters with FMO2 and FMO3 of other lepidopteran species, although the degenerate primers used for our approach were at least at the beginning of our study, not specific for lepidopteran FMO. For the design of the degenerate primers, we have used an alignment of only one lepidopteran FMO (TjSNO) and of four FMOs from two dipteran genomes. Work is in progress to test whether this arises from using mainly fat body tissues for cDNA preparations or whether the arctiids are indeed devoid of any sequences orthologous to FMO2 and FMO3 of B. mori because of a loss of the respective genes. In this regard, the branch length of the arctiid FMO sequences are notably longer than those of the PNO-sequence cluster or of other lepidopteran FMO1, suggesting a higher substitution rate within these FMO-coding sequences. A challenge for the future will be to assign a specific function to this arctiid-specific sequence cluster and to compare it with the FMO1 sequences of other lepidopteran species.

Supporting Information

Figure S1.

Amino acid alignment of flavin-dependent monooxygenases of various lepidopteren species.

(0.04 MB PDF)

Table S1.

Sequences of primers used for the identification and cloning of cDNAs of flavin-dependent monooxygenases of the Lepidoptera.

(0.02 MB PDF)

Table S2.

PCR-based strategy for identification of partial FMO-like sequences of various lepidopteren species.

(0.01 MB PDF)

Table S3.

Accession numbers of all nucleotide sequences that have been identified within this project and that have been taken from the databases.

(0.01 MB PDF)


We thank K. Vrieling (Van der Klaauw Laboratorium, Leiden, The Netherlands) for his help in collecting T. jacobaeae larvae at the beginning of this project, E. Bernays (University of Arizona, Tucson, USA) and M. Singer (Wesleyan University, Middletown, USA) for providing the larvae of G. geneura and E. acrea, and H. Vogel (Max Planck Institute for Chemical Ecology, Jena, Germany) for cDNA sequences of H. armigera libraries. We are also grateful to T. Hartmann (Technical University Braunschweig, Germany) for helpful discussions and to B. Schemmerling, M. Doose, C. Theuring, and A. Backenköhler for their excellent technical assistance.

Author Contributions

Conceived and designed the experiments: SS LW DL HM GP DO. Performed the experiments: SS LW DL HM. Analyzed the data: SS LW DL DGH DO. Contributed reagents/materials/analysis tools: DGH GP. Wrote the paper: DGH DO.


  1. 1. Naumann C, Hartmann T, Ober D (2002) Evolutionary recruitment of a flavin-dependent monooxygenase for the detoxification of host plant-acquired pyrrolizidine alkaloids in the alkaloid-defended arctiid moth Tyria jacobaeae. Proc Natl Acad Sci U S A 99: 6085–6090.
  2. 2. Alfieri A, Malito E, Orru R, Fraaije MW, Mattevi A (2008) Revealing the moonlighting role of NADP in the structure of a flavin-containing monooxygenase. Proc Natl Acad Sci U S A 105: 6572–6577.
  3. 3. Hao D, Chen S, Mu J, Xiao P (2009) Molecular phylogeny, long-term evolution, and functional divergence of flavin-containing monooxygenases. Genetica 137: 173–187.
  4. 4. Cashman JR (2002) Flavin monooxygenases. In: Ioannides C, editor. Enzyme Sytems that Metabolize Drugs and Other Xenobiotics. Chichester: John Wiley & Sons, Ltd. pp. 67–93.
  5. 5. Suh JK, Poulsen LL, Ziegler DM, Robertus JD (2000) Redox regulation of yeast flavin-containing monooxygenase. Arch Biochem Biophys 381: 317–322.
  6. 6. Zhang M, Robertus JD (2002) Molecular cloning and characterization of a full-length flavin-dependent monooxygenase from yeast. Arch Biochem Biophys 403: 277–283.
  7. 7. Schlaich NL (2007) Flavin-containing monooxygenases in plants: looking beyond detox. Trends Plant Sci 12: 412–418.
  8. 8. Ziegler DM (2002) An overview of the mechanism, substrate specificities, and structure of FMOs. Drug Metab Rev 34: 503–511.
  9. 9. Ziegler DM (1993) Recent studies on the structure and function of multisubstrate flavin-containing monooxygenases. Annu Rev Pharmacol Toxicol 33: 179–199.
  10. 10. Cashman JR (2005) Some distinctions between flavin-containing and cytochrome P450 monooxygenases. Biochem Biophys Res Commun 338: 599–604.
  11. 11. Feyereisen R (2005) Insect Cytochrome P450. In: Gilbert LI, Iatrou K, Gill SS, editors. Comprehensive Molecular Insect Science. Amsterdam: Elsevier. pp. 1–77.
  12. 12. Scharf ME, Scharf DW, Bennett GW, Pittendrigh BR (2004) Catalytic activity and expression of two flavin-containing monooxygenases from Drosophila melanogaster. Arch Insect Biochem Physiol 57: 28–39.
  13. 13. Tijet N, Helvig C, Feyereisen R (2001) The cytochrome P450 gene superfamily in Drosophila melanogaster: Annotation, intron-exon organization and phylogeny. Gene 262: 189–198.
  14. 14. Claudianos C, Ranson H, Johnson RM, Biswas S, Schuler MA, et al. (2006) A deficit of detoxification enzymes: pesticide sensitivity and environmental response in the honeybee. Insect Mol Biol 15: 615–636.
  15. 15. Hung CF, Berenbaum MR, Schuler MA (1997) Isolation and characterization of CYP6B4, a furanocoumarin-inducible cytochrome P450 from a polyphagous caterpillar (Lepidoptera: Papilionidae). Insect Biochem Mol Biol 27: 377–385.
  16. 16. Li W, Schuler MA, Berenbaum MR (2003) Diversification of furanocoumarin-metabolizing cytochrome P450 monooxygenases in two papilionids: Specificity and substrate encounter rate. Proc Natl Acad Sci U S A 100: Suppl 214593–14598.
  17. 17. Harrison TL, Zangerl AR, Schuler MA, Berenbaum MR (2001) Developmental variation in cytochrome P450 expression in Papilio polyxenes in response to xanthotoxin, a hostplant allelochemical. Arch Insect Biochem Physiol 48: 179–189.
  18. 18. Petersen RA, Zangerl AR, Berenbaum MR, Schuler MA (2001) Expression of CYP6B1 and CYP6B3 cytochrome P450 monooxygenases and furanocoumarin metabolism in different tissues of Papilio polyxenes (Lepidoptera: Papilionidae). Insect Biochem Mol Biol 31: 679–690.
  19. 19. Hartmann T, Ober D (2000) Biosynthesis and metabolism of pyrrolizidine alkaloids in plants and specialized insect herbivores. In: Leeper FJ, Vederas JC, editors. Topics in Current Chemistry. Berlin, Heidelberg: Springer. pp. 207–244.
  20. 20. Hartmann T, Ober D (2008) Defense by pyrrolizidine alkaloids: developed by plants and recruited by insects. In: Schaller A, editor. Induced Plant Resistance to Herbivory. Berlin: Springer Science+Business Media B.V.. pp. 213–231.
  21. 21. Mattocks AR (1986) Chemistry and Toxicology of Pyrrolizidine Alkaloids. London: Academic Press.
  22. 22. Lindigkeit R, Biller A, Buch M, Schiebel HM, Boppré M, et al. (1997) The two faces of pyrrolizidine alkaloids: The role of the tertiary amine and its N-oxide in chemical defense of insects with acquired plant alkaloids. Eur J Biochem 245: 626–636.
  23. 23. Narberhaus I, Zintgraf V, Dobler S (2005) Pyrrolizidine alkaloids on three trophic levels - evidence for toxic and deterrent effects on phytophages and predators. Chemoecology 15: 121–125.
  24. 24. Fu PP, Xia Q, Lin G, Chou MW (2004) Pyrrolizidine alkaloids - genotoxicity, metabolism enzymes, metabolic activation, and mechanisms. Drug Metab Rev 36: 1–55.
  25. 25. Conner WE, Eisner T, van der Meer RK, Guerrero A, Meinwald J (1981) Precopulatory sexual interactions in an arctiid moth (Utetheisa ornatrix): Role of a pheromone derived from dietary alkaloids. Behav Ecol Sociobiol 9: 227–235.
  26. 26. Eisner T, Meinwald J (1995) The chemistry of sexual selection. Proc Natl Acad Sci U S A 92: 50–55.
  27. 27. Dussourd DE, Ubik K, Harvis C, Resch J, Meinwald J, et al. (1988) Biparental defensive endowment of eggs with acquired plant alkaloid in the moth Utetheisa ornatrix. Proc Natl Acad Sci U S A 85: 5992–5996.
  28. 28. Conner WE, Roach B, Benedict E, Meinwald J, Eisner T (1990) Courtship pheromone production and body size as correlates of larval diet in males of the arctiid moth Utetheisa ornatrix. J Chem Ecol 16: 543–552.
  29. 29. Hare JF, Eisner T (1993) Pyrrolizidine alkaloid deters ant predators of Utetheisa ornatrix eggs: Effects of alkaloid concentration, oxidation state, and prior exposure of ants to alkaloid-laden prey. Oecologia 96: 9–18.
  30. 30. Bezzerides A, Yong T-H, Bezzerides J, Husseini J, Ladau J, et al. (2004) Plant-derived pyrrolizidine alkaloid protects eggs of a moth (Utetheisa ornatrix) against a parasitoid wasp (Trichogramma ostriniae). Proc Natl Acad Sci U S A 101: 9029–9032.
  31. 31. Krasnoff SB, Roelofs WL (1989) Quantitative and qualitative effects of larval diet on male scent secretions of Estigmene acrea, Phragmatobia fuliginosa, and Pyrrharctia isabella (Lepidoptera: Arctiidae). J Chem Ecol 15: 1077–1093.
  32. 32. Schulz S (2009) Alkaloid-derived male courtship pheromones. In: Conner WE, editor. Tiger Moth and Woolly Bears. New York: Oxford University Press. pp. 145–153.
  33. 33. Singer MS, Mace KC, Bernays EA (2009) Self-medication as adaptive plasticity: increased ingestion of plant toxins by parasitized caterpillars. PLoS ONE 4: e4796.
  34. 34. Bergomaz R, Boppré M (1986) A simple instant diet for rearing Arctiidae and other moths. Journal of the Lepidopterists' Society 40: 131–137.
  35. 35. Ober D, Hartmann T (1999) Homospermidine synthase, the first pathway-specific enzyme of pyrrolizidine alkaloid biosynthesis, evolved from deoxyhypusine synthase. Proc Natl Acad Sci U S A 96: 14777–14782.
  36. 36. Ober D, Hartmann T (1999) Deoxyhypusine synthase from tobacco: cDNA isolation, characterization, and bacterial expression of an enzyme with extended substrate specificity. J Biol Chem 274: 32040–32047.
  37. 37. Grubor VD (2003) The roles of cytochrome P450s in pyrethroid resistance in the AN02 strain of Helicoverpa armigera [PhD Thesis]. Melbourne, Australia: University of Melbourne.
  38. 38. Gasteiger E, Hoogland C, Gattiker A, Duvaud S, Wilkins MR, et al. (2005) Protein Identification and Analysis Tools on the ExPASy Server. In: Walker JM, editor. The Proteomics Protocols Handbook. Totowa, , NJ: Humana Press.. 988 p.
  39. 39. Bendtsen JD, Nielsen H, von Heijne G, Brunak S (2004) Improved prediction of signal peptides: SignalP 3.0. J Mol Biol 340: 783–795.
  40. 40. Horton P, Nakai K (1997) Better prediction of protein cellular localization sites with the κ nearest neighbors classifier. Proc Int Conf Intell Syst Mol Biol 5: 147–152.
  41. 41. Emanuelsson O, Nielsen H, Brunak S, von Heijne G (2000) Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300: 1005–1016.
  42. 42. Krogh A, Larsson B, von Heijne G, Sonnhammer ELL (2001) Predicting transmembrane protein topology with a hidden Markov model: Application to complete genomes. J Mol Biol 305: 567–580.
  43. 43. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882.
  44. 44. Felsenstein J (2001) (2001) PHYLIP, Phylogeny Inference Package (Univ. of Washington, Seattle), Version 3.6(alpha2).
  45. 45. Saitou N, Nei M (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425.
  46. 46. Studier FW, Rosenberg AH, Dunn JJ, Dubendorff JW (1990) Use of T7 RNA polymerase to direct expression of cloned genes. Methods Enzymol 185: 60–89.
  47. 47. Caspers P, Stieger M, Burn P (1994) Overproduction of bacterial chaperones improves the solubility of recombinant protein tyrosine kinases in Escherichia coli. Cell Mol Biol 40: 635–644.
  48. 48. Conner WE, Weller SJ (2004) A quest for alkaloids: the curious relationship between tiger moth and plants containing pyrrolizidine alkaloids. In: Cardé RT, Millar JG, editors. Advances in Insect Chemical Ecology. Cambridge: Cambridge University Press. pp. 248–282.
  49. 49. Blackwell JR, Horgan R (1991) A novel strategy for production of a highly expressed recombinant protein in an active form. FEBS Lett 295: 10–12.
  50. 50. Jean D, Monique L, Jean-Antoine L, Florence M, Francine P, et al. (1989) Larval fat body-specific gene expression in D. melanogaster. Dev Genet 10: 220–231.
  51. 51. Lawton MP, Cashman JR, Cresteil T, Dolphin CT, Elfarra AA, et al. (1994) A nomenclature for the mammalian flavin-containing monooxygenase gene family based on amino acid sequence identities. Arch Biochem Biophys 308: 254–257.
  52. 52. Weller SJ, Jacobson NL, Conner WE (1999) The evolution of chemical defences and mating systems in tiger moths (Lepidoptera: Arctiidae). Bot J Linn Soc 68: 557–578.
  53. 53. Despres L, David J-P, Gallet C (2007) The evolutionary ecology of insect resistance to plant chemicals. Trends Ecol Evol 22: 298–307.
  54. 54. Guengerich FP (2002) Cytochrome P450. In: Ioannides C, editor. Enzyme Systems that Metabolize Drugs and Other Xenobiotics. Chichester: John Wiley & Sons, Ltd. pp. 33–65.
  55. 55. Cashman JR (1995) Structural and catalytic properties of the mammalian flavin-containing monooxygenase. Chem Res Toxicol 8: 165–181.
  56. 56. Eswaramoorthy S, Bonanno JB, Burley SK, Swaminathan S (2006) Mechanism of action of a flavin-containing monooxygenase. Proc Natl Acad Sci U S A 103: 9832–9837.
  57. 57. Michalski C, Mohagheghi H, Nimtz M, Pasteels J, Ober D (2008) Salicyl alcohol oxidase of the chemical defense secretion of two chrysomelid leaf beetles: molecular and functional characterization of two new members of the glucose-methanol-choline oxidoreductase gene family. J Biol Chem 283: 19219–19228.
  58. 58. Hartmann T, Theuring C, Beuerle T, Bernays EA, Singer MS (2005) Acquisition, transformation and maintenance of plant pyrrolizidine alkaloids by the polyphagous arctiid Grammia geneura. Insect Biochem Mol Biol 35: 1083–1099.
  59. 59. Anke S, Gonde D, Kaltenegger E, Hansch R, Theuring C, et al. (2008) Pyrrolizidine alkaloid biosynthesis in Phalaenopsis orchids: developmental expression of alkaloid-specific homospermidine synthase in root tips and young flower buds. Plant Physiol 148: 751–760.
  60. 60. Frölich C, Hartmann T, Ober D (2006) Tissue distribution and biosynthesis of 1,2-saturated pyrrolizidine alkaloids in Phalaenopsis hybrides (Orchidaceae). Phytochemistry 67: 1493–1502.
  61. 61. Nurhayati N, Gondé D, Ober D (2009) Evolution of pyrrolizidine alkaloids in Phalaenopsis orchids and other monocotyledons: Identification of deoxyhypusine synthase, homospermidine synthase and related pseudogenes. Phytochemistry 70: 508–516.
  62. 62. Reina M, Gonzalez-Coloma A, Gutierrez C, Cabrera R, Henriquez J, et al. (1997) Bioactive saturated pyrrolizidine alkaloids from Heliotropium floridum. Phytochemistry 46: 845–853.
  63. 63. Becker DP, Flynn DL, Moormann AE, Nosal R, Villamil CI, et al. (2006) Pyrrolizidine esters and amides as 5-HT4 receptor agonists and antagonists. J Med Chem 49: 1125–1139.
  64. 64. Rhoades DF (1979) Evolution of plant chemical defense against herbivores. In: Rosenthal GA, Janzen DH, editors. Herbivores: Their Interaction with Secondary Plant Metabolites. New York: Academic Press. pp. 1–55.
  65. 65. Singer MS, Bernays EA, Carrière Y (2002) The interplay between nutrient balancing and toxin dilution in foraging by a generalist insect herbivore. Anim Behav 64: 629–643.
  66. 66. Li X, Baudry J, Berenbaum MR, Schuler MA (2004) Structural and functional divergence of insect CYP6B proteins: From specialist to generalist cytochrome P450. Proc Natl Acad Sci U S A 101: 2939–2944.
  67. 67. Suh JK, Robertus JD (2000) Yeast flavin-containing monooxygenase is induced by the unfolded protein response. Proc Natl Acad Sci U S A 97: 121–126.
  68. 68. Suh JK, Poulsen LL, Ziegler DM, Robertus JD (1999) Yeast flavin-containing monooxygenase generates oxidizing equivalents that control protein folding in the endoplasmic reticulum. Proc Natl Acad Sci U S A 96: 2687–2691.
  69. 69. Ziegler DM, Poulsen LL (1977) Protein disulfide bond synthesis: a possible intracellular mechanism. Trends Biochem Sci 2: 79–81.
  70. 70. Agosin M, Ankley GT (1987) Conversion of N,N-dimethylaniline to N,N-dimethylaniline-N-oxide by a cytosolic flavin-containing enzyme from Trypanosoma cruzi. Drug Metab Dispos 15: 200–203.
  71. 71. Overby LH, Buckpitt AR, Lawton MP, Atta AAE, Schulze J, et al. (1995) Characterization of flavin-containing monooxygenase 5 (FMO5) cloned from human and guinea pig: Evidence that the unique catalytic properties of FMO5 are not confined to the rabbit ortholog. Arch Biochem Biophys 327: 275–284.
  72. 72. Berenbaum MR, Favret C, Schuler MA (1996) On defining “key innovations” in an adaptive radiation: cytochrome P450s and Papilionidae. Am Nat 148: S139.