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. 2026 Jul 8;52(4):62. doi: 10.1007/s10886-026-01730-6

The Matador Bug and Several Related Leaf-footed Bugs (Hemiptera: Coreidae) Accumulate Cyanogens From their Host Plants

Colin R Morrison 1,2,✉, Brian E Sedio 1,3, Lawrence E Gilbert 1,2, Justin C Havird 1, Ummat Somjee 1,3,4,5
PMCID: PMC13346313  PMID: 42418022

Abstract

Cyanogenic compounds provide potent defenses to herbivorous insects, yet few species are known to acquire them from their food. Matador bugs, Bitta alipes (Hemiptera: Coreidae), have brightly colored signals that deter avian predators and are specialists of cyanogenic Passiflora plants. It is unknown whether these insects accumulate cyanogenic compounds. We document the first evidence of cyanogen accumulation in matador bugs. However, the presence and concentration of cyanogens depended on the Passiflora species from which insects were feeding. Further, three related species of inconspicuous Coreidae, and their Cactaceae and Aquifoliaceae host plants, also contained cyanogenic compounds. In all coreid species, cyanogen concentrations detected would make them unpalatable, or even lethal, to vertebrate predators. We report the first evidence of cyanogenic compounds in the Coreidae, doubling the number of species known to accumulate cyanide in the Hemiptera. Cyanogen accumulation may be more common amongst hemipterans than previously thought, prompting studies to examine the mechanisms of secondary chemical accumulation and evolution of aposematism.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1007/s10886-026-01730-6.

Keywords: Bitta alipes, Aposematism, Inconspicuous coloration, Passiflora, Toxicity, Synthesis, Cyanogen accumulation

Introduction

Animals from diverse clades have evolved multi-component defenses against predators (Exnerová et al. 2023; Hamalainen et al. 2024; Kikuchi et al. 2021; Rojas et al. 2017). In aposematic species for example, both warning coloration and toxicity can act synergistically to facilitate rapid learning of prey avoidance by predators (Caro and Ruxton 2019; Speed and Ruxton 2005). However, the ability to accumulate toxic compounds and the production of conspicuous warning signals are distinct processes that are likely to evolve asynchronously (Kikuchi et al. 2021).

The accumulation of toxic compounds in the body may occur upon metabolism of diets, and may provide some level of protection from predators, yet this can occur passively in many species and may not be associated with conspicuous anti-predator signals (Lindström 1999). Indeed, recent studies from moths (Nokelainen et al. 2024) and frogs (Coleman et al. 2025; Tarvin et al. 2024) reveal that many toxic species may not have conspicuous features that signal unpalatability. However, once an organism is chemically defended, co-evolutionary processes between predators and prey may lead to enhanced anti-predator signals (Kikuchi et al. 2021), but not always (Wang 2011). For example, signals associated with Batesian mimicry complexes such as conspicuous coloration, morphology or behavior that are associated with toxic prey may facilitate predator learning to avoid those prey items (Drinkwater et al. 2022; Umbers et al. 2019). While Batesian mimicry examples abound (Kikuchi et al. 2021), there are also brightly colored, non-toxic species without sympatric models (Betzner et al. 2025; Pfennig and Mullen 2010; Winters et al. 2018). The anti-predator effects of highly conspicuous and chemically defended organisms are the focus of many studies, yet understanding the toxicity of less conspicuous related species may allow us to uncover the evolutionary processes that shape multicomponent anti-predatory behavior.

Cyanogenic chemical compounds trigger aversive taste responses. Toxic hydrogen cyanide gas is released when tissues are disrupted; this gas is considered a potent and effective signal that prevents or deters consumption (Smiley and Morrison 2020). Yet a diverse group of arthropod taxa have evolved mechanisms to detoxify cyanide from their food including mites (Dixit et al. 2022; Wybouw et al. 2014), white flies (Easson et al. 2021), butterfly and moth larvae (Morrison et al. 2023; van Ohlen et al. 2016), millipedes and centipedes (Zagrobelny et al. 2018b). Further, some arthropods have evolved the ability to defend themselves against natural enemies by accumulating cyanogenic compounds from their food into their tissues via sequestration and de novo synthesis (Engler-Chaouat and Gilbert 2007; Pinheiro de Castro et al. 2019; Zagrobelny et al. 2007b, 2018a). Despite the relatively high diversity of insects relative to other groups, cyanogenic compounds have been documented in only three insect orders, with most cases documented in Lepidoptera, and few cases in Coleoptera and Hemiptera (Table 1).

Table 1.

Cyanogenic compounds can have potent antipredator effects yet have only been documented in three insect orders, with a majority of studies focusing on Lepidoptera. This table summarizes the non-lepidopteran insects that have been documented to accumulate cyanogens. In this study we document four additional Hemiptera, all from the family Coreidae, that accumulate cyanogens

Order Family Species Cyanogenic component Host Tissue Reference
Coleoptera Carabidae Megacephala virginica Mandelonitrile leaf Davis and Nahrstedt 1985
Coleoptera Chrysomelidae Paropsis atomaria Cyanohydrin; cyanogenic glucosides leaf Nahrstedt and Davis 1986
Hemiptera Aphididae Aphis craccivora Cyanamide leaf Kamo et al. 2012
Hemiptera Rhopalidae Jadera haematoloma Cyanolipids, cyanogenic glucosides seed Aldrich et al. 1990a; Aldrich et al. 1990b
Hemiptera Rhopalidae Jadera sanguinolenta Cyanolipids, cyanogenic glucosides seed Aldrich et al. 1990a; Aldrich et al. 1990b
Hemiptera Rhopalidae Leptocoris isolata Cardiospermin seed Braekman et al. 1982; Davis and Nahrstedt 1985
Hemiptera Coreidae Bitta alipes Unknown cyanogenic compound(s) leaf, stem, flower, fruit this study
Hemiptera Coreidae Acanthocephala terminalis Undetermined cyanogenic compound(s) stem this study
Hemiptera Coreidae Narnia femorata Undetermined cyanogenic compound(s) stem, fruit, seed this study
Hemiptera Coreidae Chelinidea vittiger Undetermined cyanogenic compound(s) stem, fruit, seed this study
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A small number of insect herbivores specialize exclusively on cyanogenic glucoside-rich Passiflora (Passifloraceae) (Benson et al. 1975; Morrison 2023; Pinheiro de Castro et al. 2019; Smiley and Morrison 2020). Passiflora specialists that have been assayed for cyanide release have been found to assimilate cyanogenic glucosides via combinations of direct sequestration and de novo synthesis via precursors present in host plants (Engler-Chaouat and Gilbert 2007; Pinheiro de Castro et al. 2019). For Heliconius butterflies that specialize on Passiflora, cyanogenic glucoside sequestration reinforces their aposematic coloration as an effective signal to deter avian predators (Chai 1990; Finkbeiner et al. 2014). Yet very little is known about other Passiflora specialist insects, particularly in the order Hemiptera which have been relatively understudied. For example, only four species from three genera in the order Hemiptera (comprising approximately 100,000 species) are known to accumulate cyanogenic compounds (Aldrich et al. 1990a, b; Braekman et al. 1982; Kamo et al. 2012).

Multiple species of leaf-footed bugs (Hemiptera: Coreidae) are specialists on Passiflora, yet no studies to date have examined the presence of cyanogenic compounds in their tissues. Specifically, the matador bug, Bitta alipes (Hemiptera: Coreidae: Aniscocelini) (= Anisoscelis alipes, see Leavengood Jr et al. 2024; Fig. 1), specializes on cyanogenic Passiflora shoots, fruits, and flowers (Schaefer and Mitchell 1983). The brightly-colored flags on their hind legs serve as warning signals directed towards avian and arthropod predators, and they are an emerging model to study the dynamics of aposematism relative to other life history traits in hemimetabolous insects (Evans-Blake et al. 2026; Longbottom et al. 2022; Rubin et al. 2024). Indeed, Aldrich (1988) remarked on the aposematic nature of matador bugs and proposed that they may have cyanogenic compounds that may act in synergy with their warning coloration, but no studies have pursued these speculations.

Fig. 1.

Fig. 1

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Matador bug (Bitta alipes) on Passiflora ambigua in Costa Rica A; Chelinidea vittiger on Opuntia engelmannii in Texas B; Acanthocephala terminalis on Vitis mustangensis in Texas C; Narnia femorata on Opuntia engelmannii in Texas, note Tachinidae parasitoid fly egg on thorax D

While matador bugs are avoided by birds (Rubin et al. 2024), many Coreidae species are consumed by avian predators (Beal 1915, 1918; Chapin 1925; Schlee 1986). Moreover, many coreids do not possess obvious, or conspicuous, coloration to a human observer. Instead, they are often non-conspicuous or blend into their surroundings (Reguera and Gomendio 1999). However, dull or inconspicuous coloration does not mean that an organism lacks chemical defenses, which are common in the Coreidae (Aldrich 1988; Aldrich and Yonke 1975; Eliyahu et al. 2012; Prudic et al. 2008; Remold 1963; Waterhouse and Gilby 1964; Whitehead et al. 2014).

We assayed cyanide release as a proxy for cyanogenic compound content in the matador bug, and three closely related species from four tribes in the subfamily Coreinae to assess if the ability to accumulate cyanogenic compounds is specific to the matador bug, present more broadly in the subfamily, or absent from all species sampled. The three other species measured here are not known to be aposematic and do not specialize on known cyanogenic host plants. Thus, we hypothesized that the matador bug, as a Passiflora specialist, would accumulate cyanogenic compounds, but closely related apparently non-aposematic species would not. The three related species were Chelinidea vitteger (Chelinideini), Narnia femorata (Anisoscelini), and Acanthocephala terminalis (Acanthocephalini) (Fig. 1). Chelinidea vitteger and N. femorata specialize exclusively on prickly pear cactus (Opuntia spp., Cactaceae), except for one population of N. femorata which also feeds on thistle (Cirsium, Asteraceae; Cirino et al. 2024). Yaupon (Ilex vomitoria, Aquifoliaceae) is the host plant on which A. terminalis was collected for this study, although it feeds on many hosts from several families across its distribution and individuals can consume multiple plant species in their lifetimes (McPherson et al. 2011). All Coreidae species studied here feed by piercing host plant tissues with their rostrum to extract sap or other nutrient-containing fluids. Given that these three non-conspicuously colored species do not specialize on cyanogenic host plants, we predicted that they do not accumulate cyanogenic compounds.

Materials and Methods

Matador bug adults and nymphs were collected at the La Selva Biological Reserve throughout 2022. La Selva is a protected tropical wet forest in Heredia Province, Costa Rica operated by the Organization for Tropical Studies (10.430334, –84.007149; 35 to 150 m.a.s.l.). There are 13 species of Passiflora present at the reserve (O. Vargas, personal communication), seven of which are known matador bug host plants in Costa Rica and Panamá (P. ambigua, P. auriculata, P. biflora, P. costaricensis, P. oerstedii, P. quadrangularis, and P. pittieri) (Gilbert and Morrison personal observations). Putatively non-cyanogenic species in the Coreinae were collected in south and central Texas in 2024. All samples were collected alive, were starved for 24 h so that plant material could pass through their digestive systems, and frozen at −32 °C until extraction at The University of Texas at Austin, Brackenridge Field Laboratory. Multiple bugs were assayed for some Passiflora species. Voucher specimens from the same collection of each coreid species assayed for cyanogenic compounds were deposited in the University of Texas at Austin Insect Collection (Table S1 for accession numbers and metadata). Host plants for the non-aposematic coreid species (Opuntia engelmannii for N. femorata and C. vittiger, Ilex vomitoria for A. terminalis) were collected at the same time as the putatively non-cyanogenic coreid species, freeze dried, and assayed for cyanide release.

Extraction followed a published protocol (Arias et al. 2016) that is based on the Lambert procedure (Lambert et al. 1975) which uses cyanide release from tissue samples as a proxy for content of cyanogenic compounds in the tissue; the procedure cannot differentiate the identity of cyanogenic compounds present in samples. Whole-body bug samples were homogenized in 4 mL of 0.1 M phosphoric acid and shaken at 100 rpm for 1 h (bug dry mass range = 10–332 mg). The solution was filtered through Pasteur pipettes with cotton filters. The filtrate was mixed with 2 mL of 5 M sulfuric acid and boiled in a water bath for 1 h to hydrolyze the cyanogenic compound(s) for release of hydrogen cyanide. Next, 5 mL of ice-cold 5 M sodium hydroxide was added to the hot solutions to convert hydrogen cyanide into sodium cyanide (NaOH + HCN → NaCN + H2O). Reactions were allowed to proceed for 1 h, in an ice bath. After removing the samples from the ice bath, 125 μL of the reaction were mixed with 875 μL of 0.2 M phosphate buffer, pH 6, followed by 400 μL of N-chlorosuccinimide solution. Samples were allowed to oxidize for 20 min. Finally, 1.6 mL of a chromogenic pyridine-barbituric acid solution was added to the samples to cause color change. Sample absorbance optical density was measured at 580 nm on a DU-520 UV/Vis spectrophotometer (Beckman; Indiana, USA) in 1 mL cuvettes against a blank sample that only contained the reagents used in the reaction; four technical replicates were run per sample and averaged. Cyanogen content of the samples was quantified using a serial sodium cyanide calibration curve with a concentration gradient ranging from 0 to 5 μg NaCN/mL (R2 = 0.97–0.98); 0.01 μg NaCN was the limit of detection; sodium cyanide analytical standard was provided by Sigma Aldrich (Missouri, USA). To assay cyanogenic compound content of the non-Passiflora host plants, we analyzed ~ 50 mg tissue samples from N = 7 O. engelmannii and N = 6 I. vomitoria individuals collected from sites in Goliad and Travis County Texas where the coreids that we sampled were feeding. We express results as cyanide released per dry mass of tissue samples (µg/mg) and as molar equivalents (µmol/g) using the molecular weight of cyanide (CN⁻, 26.02 g/mol).

We use the term ‘sequester’ here to describe the passing of chemical compounds directly across the gut wall where they are stored in specialized cells, tissues or glands and the term ‘synthesis’ to define the process of internally producing chemicals, though the precursor compounds may have been acquired from the diet (Beran and Petschenka 2022; Opitz and Müller 2009); we refer to both processes generically with the term ‘accumulation’ to convey that chemical compounds are present in an organism, regardless of the mechanism(s) that led to their accumulation. We use ‘non-aposematic’ to refer to species lacking conspicuous, high-contrast coloration to human observers, rather than as a quantitatively defined signal trait. Data visualization was conducted in R version 4.1.1 (R Core Team, 2025) using the package ggplot2 (Wickham and Wickham 2016), data wrangling was done with the package dplyr (Wickham and François 2015).

Results

The whole-body cyanogen concentration of matador bugs varied considerably among individuals collected from six Passiflora host species (Fig. 2B). Notably, no cyanogenic compounds were found in any matador bug individuals collected from P. ambigua (N = 8), yet significant cyanogen concentrations were found in individuals collected from the other five Passiflora host plants. The average concentration of the matador bug on host plants where it accumulated cyanogenic compounds was 0.40 μg/mg ± 0.11 standard error (S.E.), and it was 0.17 μg/mg ± 0.08 S.E. if individuals feeding on P. ambigua were included.

Fig. 2.

Fig. 2

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Whole-body cyanogen colorimetric analysis revealed variation in concentration across four species of the Coreidae sampled: Acanthocephala terminalis (N = 3), Bitta alipes (N = 14); Chelinidea vittiger (N = 3), and Narnia femorata (N = 3) A. The presence of cyanogenic compounds in matador bugs was dependent on the Passiflora host plant on which it was collected B; Bitta alipes samples with no detectable cyanogen (collected on P. ambigua) are excluded from panel A, but are shown in panel B. Colorimetric analysis revealed cyanogens in yaupon (Ilex vomitoria), the host plant for A. terminalis and prickly pear (Opuntia engelmannii), the host plant for N. femorata and C. vittiger C. Large dots with bars represent means and standard errors

The average cyanogen concentrations for the non-Passiflora feeding species were 0.37 µg/mg ± 0.17 S.E. for A. terminalis, 0.50 µg/mg ± 0.22 S.E. for C. vittiger, and the highest concentration, 0.76 µg/mg ± 0.08 S.E., was found in N. femorata (Fig. 2A). The non-Passiflora feeding species had a higher concentration of cyanogenic compounds in their tissues, on average, compared to the matador bug (Fig. 2A). Despite small sample numbers, the large S.E. values reported above show that intraspecific variation was considerable among the non-Passiflora feeding coreids. Opuntia engelmannii, the host plant for N. femorata and C. vittiger, had an average cyanogen concentration of 0.54 µg/mg ± 0.13 S.E. (N = 7; Fig. 2C). Ilex vomitoria, the host plant that A. terminalis was collected on, had an average concentration of 1.42 µg/mg ± 0.33 S.E. (N = 6; Fig. 2C). Molar concentrations of bug and host plant tissues are available in Supplemental Table S2.

Discussion

All arthropods with cyanogenic compounds in their tissues ultimately derive it from their food and most of them are known to acquire it from cyanogenic host plants (Pinheiro de Castro et al. 2019; Zagrobelny et al. 2018a). Some insects directly sequester cyanogenic compounds from their host plants, but growing evidence suggests that many both sequester and metabolize host-derived cyanogens which are subsequently re-synthesized into novel cyanogenic compounds (Engler-Chaouat and Gilbert 2007; Pinheiro de Castro et al. 2019; Zagrobelny et al. 2008). The matador bug is the first documented case of cyanogenic compound accumulation in a species of Coreidae, supporting our hypothesis that it can accumulate the molecule from its cyanogenic Passiflora host plants. This finding is consistent with the avoidance of these insects by wild insectivorous birds and provides a biochemical basis for the observation that naïve birds learn to avoid matador bugs (Rubin et al. 2024).

Unexpectedly, all other coreid species investigated contained cyanogenic compounds in their tissues at comparable levels to matador bugs. Also unexpectedly, their host plants contained cyanogens. The I. vomitoria and O. engelmannii cyanogen concentration range of 0.3–2.4 µg/mg reported here were lower than most Passiflora species and Lotus (Fabaceae) species that are used by Heliconiinae or Zygaena caterpillars to acquire cyanogenic compounds, but present at comparable magnitudes (e.g., 0.1–10.0 µg/mg) (Engler-Chaouat and Gilbert 2007; Morrison et al. 2023; Smiley and Morrison 2020; Zagrobelny et al. 2007a). To our knowledge, this is the first report of cyanogenesis in the Cactaceae and Aquifoliaceae. These findings demonstrate that these coreid species accumulated cyanogenic compounds from their host plants by direct sequestration and/or de novo synthesis using precursor compounds that they consumed. The discovery of three non-aposematic coreids accumulating cyanogens across three tribes within a subfamily suggests that cyanogenic compound acquisition may be shared more broadly across the Coreidae phylogeny. This discovery more than doubles the number of Hemiptera genera known to accumulate cyanogenic compounds from three to seven and lays a foundation for future studies to investigate the precise mechanism(s) of cyanogen accumulation in matador bugs, other Passiflora specialist coreids including putative ‘wasp mimics’ (e.g., Holhymenia), and coreids from other subfamilies which feed on multiple host plants. Future studies employing standard liquid chromatography-mass spectrometry instruments, with internal cyanogenic compound standards (e.g., linamarin, prunasin, dhurrin), would be able to infer the mechanism of cyanogen accumulation from host plants (e.g., Sculfort et al. 2020).

Accumulation of cyanogenic compounds from host plants is generally understudied in hemipterans. The ability was only known from the Aphidae (Kamo et al. 2012), and Rhopalidae (Aldrich et al. 1990a, b; Braekman et al. 1982; Davis and Nahrstedt 1985; Kamo et al. 2012; Nahrstedt 1988) until now. Cardiospermin is a cyanogenic glucoside synthesized by Leptocoris isolata (Rhopalidae) from fruit of its host plant Allophylus cobbe (Sapindaceae), which does not contain cardiospermin (Nahrstedt 1988). Future studies may use dissection and chemical analyses of hemolymph, scent glands, and other body tissues to identify where coreids store cyanogens and which specific compounds, sequestration or synthesis pathways are used to co-opt plant chemical compounds for defense.

The matador bugs collected from P. ambigua had no detectable cyanogens, unlike those collected from five other host species. Passiflora ambigua is known to be cyanogenic, thus the lack of detectable cyanogenic compounds in matador bugs is most likely reflective of host-plant dependent accumulation. Yet, the reason for the lack of cyanogen accumulation in matador bugs on P. ambigua, and not those collected from other Passiflora species examined, remains an open question for future studies. Population-level variation in toxicity can disrupt predator foraging behavior because prey palatability is unpredictable (Hossie et al. 2021; Prudic et al. 2019). Similar intraspecific variation exists in Heliconiinae butterfly and Zygaena moth species that accumulate cyanogenic glucosides (Chai 1990; Morrison et al. 2023; Zagrobelny et al. 2007a), as well as a diversity of insects from other orders (Agrawal 2004; Pasteels et al. 1995; Sword 1999). We predict that variation in toxin accumulation is common in Coreidae species. One N. femorata population exists entirely on thistle, which is the first observation of this species consuming and reproducing on a non-Opuntia host (Cirino et al. 2024). Comparing cyanogen content of the thistle-feeding N. femorata population which we predict does not contain cyanogenic compounds, with Opuntia-feeding populations would illuminate the extent of intraspecific variation in cyanogen synthesis possible for hemipterans. Similarly, the generalist A. terminalis, which feeds on host plants in many plant families across the eastern U.S.A. (McPherson et al. 2011), offers another opportunity for population-level cyanogenic compound accumulation study. Opuntia species, I. vomitoria, and Centaurea (another thistle genus) contain N-containing alkaloids which may be precursors for coreid cyanogen synthesis (Morrison and Sedio, unpublished data), possibly in addition to sequestration. It is also possible that these species accumulate secondary defensive compounds from host plants besides cyanogens. Thus, the Coreidae provide an exciting model to understand insect chemical ecology within populations that vary in host plant usage, and across species on the generalist-specialist diet spectrum.

The cyanogen concentrations we measured in coreids (0.1–1.0 µg/mg) are relatively low compared to Heliconius butterflies, Zygaena moths, and other arthropods which accumulate cyanogenic compounds (1.0–10.0 µg/mg), but are well within a range that can be unpalatable, or lethal, to naïve vertebrate predators (Pinheiro de Castro et al. 2019; Zagrobelny et al. 2018a). Intraspecific variation in cyanogenic compound concentration and warning coloration may influence predator learning. Birds attack and consume apparently dull-colored coreid species in Leptoglossus and Acanthocephala (Beal 1915, 1918; Chapin 1925), but avoid aposematic matador bugs (Evans-Blake et al. 2026; Rubin et al. 2024). In addition, European blackbirds prefer non-aposematic Heteroptera species (Coreidae, Pentatomidae, Pyrrhocoridae) over aposematic ones (Schlee 1986). Variation in palatability among individuals with certain color or morphological phenotypes may make it difficult for some visual predators to learn which individuals to avoid attacking (Skelhorn and Rowe 2006; Taylor et al. 2016). Other chemical compounds in Coreidae which are exuded from thoracic or abdominal glands in some species may contribute to their unpalatability (Aldrich and Yonke 1975; Waterhouse and Gilby 1964).

While conspicuous warning signals are often associated with chemical defense (Kikuchi et al. 2021), our results emphasize that toxicity can occur in the absence of overt aposematic coloration (Rowland et al. 2017). We interpret cyanogen accumulation in these inconspicuous coreids as a general defensive trait that may reduce predation risk irrespective of visual signaling. The absence of strong warning coloration does not preclude defensive value, particularly if predator responses depend on learning, sampling frequency, and community context (Rowland et al. 2017; Skelhorn and Rowe 2006), rely on cues other than color (Rojas et al. 2018), or if chemical defenses function in combination with other traits (Hamalainen et al. 2024). A more detailed evaluation of the evolutionary drivers of this pattern is beyond the scope of this study, but our findings highlight the need to consider chemically defended yet visually inconspicuous species in broader discussions of predator–prey interactions.

Arthropod predator–prey interactions are rarely determined by the presence of one chemical compound, but rather complex mixtures of compounds which act synergistically to deter or mitigate natural enemies (Richards et al. 2015; Wetzel and Whitehead 2020). More characterization of non-model arthropod chemical composition will provide a deeper understanding about how dietary variation affects resource-consumer-enemy dynamics (Abdala-Roberts et al. 2019; Sedio 2017). The unexpected results reported here indicate that more hemipterans, and insects in general, may be defending themselves with food plant-derived cyanogenic compounds than thought previously. We advocate for chemical surveys of more non-aposematic arthropods, along with host plants not known to be cyanogenic, because the paradigm in which an organism’s palatability spectrum correlates with its degree of warning coloration is more nuanced than typically appreciated (Coleman et al. 2025; Despland 2005; Drinkwater et al. 2022; Dyer 1995; Lehmann et al. 2014; Medina et al. 2020; Nokelainen et al. 2024; Pinheiro de Castro et al. 2019; Tarvin et al. 2024; Umbers et al. 2017).

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 17 KB) (16.8KB, docx)

Acknowledgements

Laboratory space was provided by the UT Austin Brackenridge Field Laboratory. Greivin Salazar Bravo of the Organization for Tropical Studies assisted with field collections. Costa Rican collecting permit SINAC-ACC-PI-re-072-2022 for B. alipes was granted to CRM by MINAE-SINAC in 2022. Private property owners granted permission for the authors to collect the USA Coreidae species on their land. Fiona MacNeill assisted with cyanogen content assays. Reviews by Mike Ryan, two external reviewers, and the editor improved the clarity of this contribution significantly.

Author Contributions

CRM collected samples, analyzed the data, developed the study design, and wrote the manuscript. BES provided feedback on the study design and reviewed the intellectual content of the results. LEG provided organism images, feedback on the study design, and field natural history wisdom about the intellectual content. JCH contributed to study design development and provided critical review of the manuscript. US collected samples, created data visualizations, contributed to early manuscript drafts, and developed the study design. All authors contributed to manuscript revisions and approved of its final submission.

Funding

CRM was supported by the Stengl-Wyer Fellowship program and a Research Grant from the Graduate Program in Ecology, Evolution and Behavior at The University of Texas at Austin. Field work in Costa Rica was supported by The Texas Ecological Laboratory Program. US was supported by the NOMIS Fellowship Program at STRI. This study was supported by the NOMIS Foundation.

Data Availability

Code and datasets used for this research are archived at Zenodo via GitHub at [https://doi.org/10.5281/zenodo.18078457] (https://zenodo.org/records/18078457) (Morrison 2026).

Declarations

Competing interests

The authors declare no competing interests.

Footnotes

Justin C. Havird and Ummat Somjee are co-senior authors.

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  1. Abdala-Roberts L, Puentes A, Finke DL, Marquis RJ, Montserrat M, Poelman EH, Rasmann S, Sentis A, van Dam NM, Wimp G, Mooney K, Björkman C (2019) Tri-trophic interactions: bridging species, communities and ecosystems. Ecol Lett 22:2151–2167. 10.1111/ele.13392 [DOI] [PMC free article] [PubMed]
  2. Agrawal AA (2004) Plant defense and density dependence in the population growth of herbivores. Am Nat 164:113–120. 10.1086/420980 [DOI] [PubMed] [Google Scholar]
  3. Aldrich JR, Yonke TR (1975) Natural products of abdominal and metathoracic scent glands of coreoid bugs Ann Entomol Soc Am 68:955–960
  4. Aldrich J, Carroll S, Oliver J, Lusby W, Rudmann A, Waters R (1990a) Exocrine secretions of scentless plant bugs; Jadera,Boisea;Niesthrea species (Hemiptera: Heteroptera: Rhopalidae). Biochem Syst Ecol 18:369–376
  5. Aldrich JR, Carroll SP, Lusby WR, Thompson MJ, Kochansky JP, Waters RM (1990b) Sapindaceae, cyanolipids, and bugs. J Chem Ecol 16:199–210. 10.1007/BF01021279 [DOI] [PubMed]
  6. Aldrich JR (1988) Chemical Ecology of the Heteroptera. Annu Rev Entomol 33: 211–238. 10.1146/annurev.en.33.010188.001235 [DOI]
  7. Arias M, Meichanetzoglou A, Elias M, Rosser N, de-Silva DL, Nay B, Llaurens V (2016) Variation in cyanogenic compounds concentration within a Heliconius butterfly community: does mimicry explain everything? BMC Evol Biol 16:272. 10.1186/s12862-016-0843-5 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Beal FEL (1915) Food of the robins and bluebirds of the United States. vol 171. US Department of Agriculture.
  9. Beal FEL (1918) Food habits of the swallows: A family of valuable native birds. vol 619. US Department of Agriculture.
  10. Benson WW, Brown KS Jr, Gilbert LE (1975) Coevolution of plants and herbivores: passion flower butterflies. Evolution 29:659–680. 10.1111/j.1558-5646.1975.tb00861.x [DOI] [PubMed]
  11. Beran F, Petschenka G (2022) Sequestration of plant defense compounds by insects: from mechanisms to insect-plant coevolution. Annu Rev Entomol 67:163–180. 10.1146/annurev-ento-062821-062319 [DOI] [PubMed] [Google Scholar]
  12. Betzner J, Watterson K, Ronk S, Enginger K, Gall B (2025) Color may not serve as a deimatic mechanism in Red-eyed treefrogs (Agalychnis callidryas): evidence from an experimental study. Ethology Ecology & Evolution 37:1–18. 10.1080/03949370.2025.2563565 [DOI]
  13. Braekman JC, Daloze D, Pasteels J (1982) Cyanogenic and other glucosides in a Neo-Guinean bug Leptocoris isolata: possible precursors in its host-plant. Biochem Syst Ecol 10:355–364. 10.1016/0305-1978(82)90010-2 [DOI]
  14. Caro T, Ruxton G (2019) Aposematism: unpacking the defences. Trends Ecol Evol 34:595–604. 10.1016/j.tree.2019.02.015 [DOI] [PubMed] [Google Scholar]
  15. Chai P (1990) Relationships between visual characteristics of rainforest butterflies and responses of a specialized insectivorous bird. In: Wicksten M (ed) Adaptive coloration in invertebrates. Texas A&M University Press, College Station, Texas, pp 31–60.
  16. Chapin EA (1925) Food habits of the vireos: a family of insectivorous birds. vol 1355. US Department of Agriculture.
  17. Cirino LA, McEvoy I, Smith LC, Emberts Z (2024) The leaf-footed cactus bug is not a cactus specialist: Narnia femorata feeds, fights, and mates on thistle. Ecol Evol 14:e70257. 10.1002/ece3.70257 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Coleman JL, Wang SY, Marek PE, Morrison CR, Sedio BE, Cannatella DC (2025) Nuance in the narrative of a brown poison frog: environmental alkaloids and specialized foraging in a presumed toxin-free and diet-generalized species. J Chem Ecol 51:1–25 [DOI] [PubMed] [Google Scholar]
  19. Davis RH, Nahrstedt A (1985) Cyanogenesis in insects. In: Kerkut GA, Gilbert LI (eds) Comprehensive insect physiology, biochemistry and pharmacology, vol 11. Pharmacology. Pergamon Press, Oxford, pp 635–654.
  20. Despland E (2005) Diet breadth and anti-predator strategies in desert locusts and other orthopterans. J Orthoptera Res 14:227–233.  10.1665/1082-6467(2005)14[227:DBAASI]2.0.CO;2 [DOI]
  21. Dixit S, Widemann E, Bensoussan N, Salehipourshirazi G, Bruinsma K, Milojevic M, Shukla A, Romero LC, Zhurov V, Bernards MA, Chruszcz M, Grbic M, Grbic V (2022) β-cyanoalanine synthase protects mites against Arabidopsis defenses. Plant Physiol 189:1961–1975. 10.1093/plphys/kiac147 [DOI] [PMC free article] [PubMed]
  22. Drinkwater E, Allen WL, Endler JA, Hanlon RT, Holmes G, Homziak NT, Kang C, Leavell BC, Lehtonen J, Loeffler-Henry K, Ratcliffe JM, Rowe C, Ruxton GD, Sherratt TN, Skelhorn J, Skojec C, Smart HR, White TE, Yack JE, Young CM, Umbers KDL (2022) A synthesis of deimatic behaviour. Biol Rev Camb Philos Soc 97:2237–2267. 10.1111/brv.12891 [DOI] [PubMed]
  23. Dyer LA (1995) Tasty generalists and nasty specialists? Antipredator mechanisms in tropical lepidopteran larvae. Ecology 76:1483–1496. 10.2307/1938150 [DOI] [Google Scholar]
  24. Easson MLAE, Malka O, Paetz C, Hojná A, Reichelt M, Stein B, van Brunschot S, Feldmesser E, Campbell L, Colvin J, Winter S, Morin S, Gershenzon J, Vassão DG (2021) Activation and detoxification of cassava cyanogenic glucosides by the whitefly Bemisia tabaci. Sci Rep 11:13244. 10.1038/s41598-021-92553-w [DOI] [PMC free article] [PubMed]
  25. Eliyahu D, Ceballos RA, Saeidi V, Becerra JX (2012) Synergy versus potency in the defensive secretions from nymphs of two pentatomomorphan families (Hemiptera: Coreidae and Pentatomidae). J Chem Ecol 38:1358–1365. 10.1007/s10886-012-0200-0 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Engler-Chaouat HS, Gilbert LE (2007) De novo synthesis vs. sequestration: negatively correlated metabolic traits and the evolution of host plant specialization in cyanogenic butterflies. J Chem Ecol 33:25–42. 10.1007/s10886-006-9207-8 [DOI] [PubMed] [Google Scholar]
  27. Evans-Blake C, Rubin JJ, Somjee U (2026) Flag-waving behavior in matador bugs is an antipredatory strategy. Curr Zool 72:293–302. 10.1093/cz/zoaf047 [DOI] [PMC free article] [PubMed]
  28. Exnerová A, Kang C, Rowland HM, Kikuchi DW (2023) Evolution of multiple prey defences: from predator cognition to community ecology. J Evol Biol 36:961–966. 10.1111/jeb.14196 [DOI] [PubMed]
  29. Finkbeiner SD, Briscoe AD, Reed RD (2014) Warning signals are seductive: relative contributions of color and pattern to predator avoidance and mate attraction in Heliconius butterflies. Evolution 68:3410–3420. 10.1111/evo.12524 [DOI] [PubMed] [Google Scholar]
  30. Hämäläinen L, Binns GE, Hart NS, Mappes J, McDonald PG, O'Neill LG, Rowland HM, Umbers KDL, Herberstein ME (2024) Predator selection on multicomponent warning signals in an aposematic moth. Behav Ecol 35:arad097. 10.1093/beheco/arad097 [DOI] [PMC free article] [PubMed]
  31. Hossie TJ, Chan K, Murray DL (2021) Increasing availability of palatable prey induces predator-dependence and increases predation on unpalatable prey. Sci Rep 11:6763. 10.1038/s41598-021-86080-x [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Kamo T, Tokuoka Y, Miyazaki M (2012) Quantification of canavanine, 2-aminoethanol, and cyanamide in Aphis craccivora and its host plants, Robinia pseudoacacia and Vicia angustifolia: effects of these compounds on larval survivorship of Harmonia axyridis. J Chem Ecol 38:1552–1560. 10.1007/s10886-012-0220-9 [DOI] [PubMed] [Google Scholar]
  33. Kikuchi DW, Herberstein ME, Barfield M, Holt RD, Mappes J (2021) Why aren’t warning signals everywhere? On the prevalence of aposematism and mimicry in communities. Biol Rev Camb Philos Soc 96:2446–2460. 10.1111/brv.12760 [DOI] [PubMed]
  34. Lambert JL, Ramasamy J, Paukstelis JV (1975) Stable reagents for the colorimetric determination of cyanide by modified Koenig reactions. Anal Chem 47:916–918 [Google Scholar]
  35. Leavengood JM Jr, Brailovsky H, Barrera E (2024) Studies in the New World tribe Anisoscelini Laporte, 1832 (Heteroptera: Coreidae: Coreinae): updated distributions, nomenclatural changes, and keys to the genera of the tribe and the species of Anisoscelis Latreille, 1829 and Bitta Osuna, 1984. Zootaxa 5415:241–268. 10.11646/zootaxa.5415.2.2 [DOI] [PubMed]
  36. Lehmann KD, Goldman BW, Dworkin I, Bryson DM, Wagner AP (2014) From cues to signals: evolution of interspecific communication via aposematism and mimicry in a predator-prey system. PLoS ONE 9:e91783. 10.1371/journal.pone.0091783 [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Lindström L (1999) Experimental approaches to studying the initial evolution of conspicuous aposematic signalling. Evol Ecol 13:605–618. 10.1023/A:1011004129607 [DOI] [Google Scholar]
  38. Longbottom C, Falk JJ, Greenway EV, Johnson MG, Ramos C, Rößler DC, Somjee U (2022) Why does the matador bug, Anisoscelis alipes (Hemiptera: Coreidae), wave its brightly colored legs? J Insect Behav 35:171–182. 10.1007/s10905-022-09809-0 [DOI]
  39. McPherson J, Packauskas RJ, Sites RW, Taylor SJ, Bundy CS, Bradshaw JD, Mitchell PL (2011) Review of Acanthocephala (Hemiptera: Heteroptera: Coreidae) of America north of Mexico with a key to species. Zootaxa 2835:30–40. 10.11646/zootaxa.2835.1.2 [DOI] [Google Scholar]
  40. Medina I, Vega-Trejo R, Wallenius T, Symonds MRE, Stuart-Fox D (2020) From cryptic to colorful: evolutionary decoupling of larval and adult color in butterflies. Evol Lett 4:34–43. 10.1002/evl3.149 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Morrison CR, Nguyen C, Gilbert LE (2023) The role of host plant usage and the accumulation of toxic secondary chemical compounds across the life cycle of a passion vine specialist butterfly. Ecol Entomol 48:389–395. 10.1111/een.13229 [DOI] [Google Scholar]
  42. Morrison CR (2023) Mechanisms driving species interactions and community structure on the phytochemical landscape. https://repositories.lib.utexas.edu/items/13f5b5e9-3d08-44e1-8b59-51c50e3f4235
  43. Morrison CR (2026) cmorrison1/Coreid_CN_assimilation: v1.0.0. Zenodo. 10.5281/zenodo.18078457 [DOI]
  44. Nahrstedt A, Davis R (1986) (R) MandeIonitrile and Prunasin, the sources of hydrogen cyanide in all stages of Paropsis atomaria (Coleoptera: Chrysomelidae). Z Naturforsch C 41:928–934
  45. Nahrstedt A (1988) Cyanogenesis and the role of cyanogenic compounds in insects. In: Evered D, Harnett S (eds) Cyanide compounds in biology. Ciba Foundation Symposium 140. Wiley, Chichester, pp 131–150 [DOI] [PubMed]
  46. Nokelainen O, Silvasti SA, Strauss SY, Wahlberg N, Mappes J (2024) Predator selection on phenotypic variability of cryptic and aposematic moths. Nat Commun 15:1678. 10.1038/s41467-024-45329-5 [DOI] [PMC free article] [PubMed]
  47. Opitz SE, Müller C (2009) Plant chemistry and insect sequestration. Chemoecology 19:117–154. 10.1007/s00049-009-0018-6 [DOI] [Google Scholar]
  48. Pasteels JM, Dobler S, Rowell-Rahier M, Ehmke A, Hartmann T (1995) Distribution of autogenous and host-derived chemical defenses in Oreina leaf beetles (Coleoptera: Chrysomelidae). J Chem Ecol 21:1163–1179. 10.1007/BF02228318 [DOI] [PubMed] [Google Scholar]
  49. Pfennig DW, Mullen SP (2010) Mimics without models: causes and consequences of allopatry in Batesian mimicry complexes. Proc Biol Sci 277:2577–2585. 10.1098/rspb.2010.0586 [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Pinheiro de Castro EC, Zagrobelny M, Zurano JP, Zikan Cardoso M, Feyereisen R, Bak S (2019) Sequestration and biosynthesis of cyanogenic glucosides in passion vine butterflies and consequences for the diversification of their host plants. Ecol Evol 9:5079–5093. 10.1002/ece3.5062 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Prudic KL, Noge K, Becerra JX (2008) Adults and nymphs do not smell the same: the different defensive compounds of the giant mesquite bug (Thasus neocalifornicus: Coreidae). J Chem Ecol 34:734–741. 10.1007/s10886-008-9480-9 [DOI] [PubMed] [Google Scholar]
  52. Prudic KL, Timmermann BN, Papaj DR, Ritland DB, Oliver JC (2019) Mimicry in viceroy butterflies is dependent on abundance of the model queen butterfly. Commun Biol 2:68. 10.1038/s42003-019-0303-z [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Reguera P, Gomendio M (1999) Predation costs associated with parental care in the golden egg bug Phyllomorpha laciniata (Heteroptera: Coreidae). Behav Ecol 10:541–544. 10.1093/beheco/10.5.541 [DOI] [Google Scholar]
  54. R Core Team (2025) R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna, Austria. https://www.R-project.org/
  55. Remold H (1963) Scent-glands of land-bugs, their physiology and biological function. Nature 198:764–768 [Google Scholar]
  56. Richards LA, Dyer LA, Forister ML, Smilanich AM, Dodson CD, Leonard MD, Jeffrey CS (2015) Phytochemical diversity drives plant-insect community diversity. Proc Natl Acad Sci U S A 112:10973–10978. 10.1073/pnas.1504977112 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Rojas B, Burdfield-Steel E, Pakkanen H, Suisto K, Maczka M, Schulz S, Mappes J (2017) How to fight multiple enemies: target-specific chemical defences in an aposematic moth. Proc Biol Sci 284:20171424. 10.1098/rspb.2017.1424 [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Rojas B, Burdfield-Steel E, De Pasqual C, Gordon S, Hernández L, Mappes J, Nokelainen O, Rönkä K, Lindstedt C (2018) Multimodal aposematic signals and their emerging role in mate attraction. Front Ecol Evol 6:93. 10.3389/fevo.2018.00093 [DOI]
  59. Rowland HM, Fulford AJ, Ruxton GD (2017) Predator learning differences affect the survival of chemically defended prey. Anim Behav 124:65–74 [Google Scholar]
  60. Rubin JJ, Medina-Madrid JL, Falk JJ, Somjee U (2024) The matador bug’s elaborate flags deter avian predators. Behav Ecol 35:arae019. 10.1093/beheco/arae019 [DOI] [Google Scholar]
  61. Schaefer CW, Mitchell PL (1983) Food plants of the Coreoidea (Hemiptera: Heteroptera). Ann Entomol Soc Am 76:591–615. 10.1093/aesa/76.4.591 [DOI] [Google Scholar]
  62. Schlee MA (1986) Avian predation on Heteroptera: experiments on the European blackbird Turdus m. merula L. Ethology 73:1–18. 10.1111/j.1439-0310.1986.tb00995.x [DOI] [Google Scholar]
  63. Sculfort O, de Castro EC, Kozak KM, Bak S, Elias M, Nay B, Llaurens V (2020) Variation of chemical compounds in wild Heliconiini reveals ecological factors involved in the evolution of chemical defenses in mimetic butterflies. Ecol Evol 10:2677–2694 [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Sedio BE (2017) Recent breakthroughs in metabolomics promise to reveal the cryptic chemical traits that mediate plant community composition, character evolution and lineage diversification. New Phytol 214:952–958. 10.1111/nph.14438 [DOI] [PubMed] [Google Scholar]
  65. Skelhorn J, Rowe C (2006) Prey palatability influences predator learning and memory. Anim Behav 71:1111–1118. 10.1016/j.anbehav.2005.08.011 [DOI] [Google Scholar]
  66. Smiley J, Morrison CR (2020) Using a portable hydrogen cyanide gas meter to uncover a dynamic phytochemical landscape. Appl Plant Sci 8:e11336. 10.1002/aps3.11336 [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Speed MP, Ruxton GD (2005) Warning displays in spiny animals: one (more) evolutionary route to aposematism. Evolution 59:2499–2508 [PubMed] [Google Scholar]
  68. Sword GA (1999) Density-dependent warning coloration. Nature 397:217–217. 10.1038/16609 [DOI] [Google Scholar]
  69. Tarvin RD, Coleman JL, Donoso DA, Betancourth-Cundar M, López-Hervas K, Gleason KS, Sanders JR, Smith JM, Ron SR, Santos JC, Sedio BE, Cannatella DC, Fitch RW (2024) Passive accumulation of alkaloids in inconspicuously colored frogs refines the evolutionary paradigm of acquired chemical defenses. eLife 13:RP100011. 10.7554/eLife.100011.3 [DOI] [PMC free article] [PubMed]
  70. Taylor LA, Amin Z, Maier EB, Byrne KJ, Morehouse NI (2016) Flexible color learning in an invertebrate predator: Habronattus jumping spiders can learn to prefer or avoid red during foraging. Behav Ecol 27:520–529. 10.1093/beheco/arv182 [DOI] [Google Scholar]
  71. Umbers KD, De Bona S, White TE, Lehtonen J, Mappes J, Endler JA (2017) Deimatism: a neglected component of antipredator defence. Biol Lett 13:20160936. 10.1098/rsbl.2016.0936 [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Umbers KDL, White TE, De Bona S, Haff T, Ryeland J, Drinkwater E, Mappes J (2019) The protective value of a defensive display varies with the experience of wild predators. Sci Rep 9:463. 10.1038/s41598-018-36995-9 [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. van Ohlen M, Herfurth A-M, Kerbstadt H, Wittstock U (2016) Cyanide detoxification in an insect herbivore: Molecular identification of β-cyanoalanine synthases from Pieris rapae. Insect Biochem Mol Biol 70:99–110 [DOI] [PubMed] [Google Scholar]
  74. Wang IJ (2011) Inversely related aposematic traits: reduced conspicuousness evolves with increased toxicity in a polymorphic poison-dart frog. Evolution 65:1637–1649. 10.1111/j.1558-5646.2011.01257.x [DOI] [PubMed] [Google Scholar]
  75. Waterhouse D, Gilby A (1964) The adult scent glands and scent of nine bugs of the superfamily Coreoidea. J Insect Physiol 10:977–987 [Google Scholar]
  76. Wetzel WC, Whitehead SR (2020) The many dimensions of phytochemical diversity: linking theory to practice. Ecol Lett 23:16–32. 10.1111/ele.13422 [DOI] [PubMed] [Google Scholar]
  77. Whitehead SR, Reid E, Sapp J, Poveda K, Royer AM, Posto AL, Kessler A (2014) A specialist herbivore uses chemical camouflage to overcome the defenses of an ant-plant mutualism. PLoS ONE 9:e102604. 10.1371/journal.pone.0102604 [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Wickham H (2016) ggplot2: elegant graphics for data analysis. Springer-Verlag, New York.
  79. Wickham H, François R (2015) dplyr: a grammar of data manipulation. R package version 0.4.3. https://CRAN.R-project.org/package=dplyr
  80. Winters AE, White AM, Dewi AS, Mudianta IW, Wilson NG, Forster LC, Garson MJ, Cheney KL (2018) Distribution of defensive metabolites in nudibranch molluscs. J Chem Ecol 44:384–396. 10.1007/s10886-018-0941-5 [DOI] [PubMed]
  81. Wybouw N, Dermauw W, Tirry L, Stevens C, Grbić M, Feyereisen R, Van Leeuwen T (2014) A gene horizontally transferred from bacteria protects arthropods from host plant cyanide poisoning. Elife 3:e02365 [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Zagrobelny M, Bak S, Ekstrom CT, Olsen CE, Moller BL (2007a) The cyanogenic glucoside composition of Zygaena filipendulae (Lepidoptera: Zygaenidae) as affected by feeding on wild-type and transgenic lotus populations with variable cyanogenic glucoside profiles. Insect Biochem Mol Biol 37:10–18. 10.1016/j.ibmb.2006.09.008 [DOI] [PubMed]
  83. Zagrobelny M, Bak S, Olsen CE, Moller BL (2007b) Intimate roles for cyanogenic glucosides in the life cycle of Zygaena filipendulae (Lepidoptera, Zygaenidae). Insect Biochem Mol Biol 37:1189–1197. 10.1016/j.ibmb.2007.07.008 [DOI] [PubMed]
  84. Zagrobelny M, Bak S, Moller BL (2008) Cyanogenesis in plants and arthropods. Phytochemistry 69:1457–1468. 10.1016/j.phytochem.2008.02.019 [DOI] [PubMed] [Google Scholar]
  85. Zagrobelny M, de Castro ECP, Moller BL, Bak S (2018a) Cyanogenesis in arthropods: from chemical warfare to nuptial gifts. Insects 9:51. 10.3390/insects9020051 [DOI] [PMC free article] [PubMed]
  86. Zagrobelny M, Jensen MK, Vogel H, Feyereisen R, Bak S (2018b) Evolution of the biosynthetic pathway for cyanogenic glucosides in Lepidoptera. J Mol Evol 86:379–394. 10.1007/s00239-018-9854-8 [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary file1 (DOCX 17 KB) (16.8KB, docx)

Data Availability Statement

Code and datasets used for this research are archived at Zenodo via GitHub at [https://doi.org/10.5281/zenodo.18078457] (https://zenodo.org/records/18078457) (Morrison 2026).

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