Glucosinolate hydrolysis products suppress entomopathogenic nematodes in vitro but do not protect sequestering flea beetle larvae in vivo

Johannes Körnig; Vojtech Beneš; Christin Manthey; Michael Reichelt; Grit Kunert; Christian Paetz; Johanna Kutzschbach; Paula Lampe; Martin Kaltenpoth; Franziska Beran

doi:10.1002/ps.70482

. 2026 Jan 5;82(4):3603–3612. doi: 10.1002/ps.70482

Glucosinolate hydrolysis products suppress entomopathogenic nematodes in vitro but do not protect sequestering flea beetle larvae in vivo

Johannes Körnig ^1,², Vojtech Beneš ^1,³, Christin Manthey ¹, Michael Reichelt ⁴, Grit Kunert ⁴, Christian Paetz ⁵, Johanna Kutzschbach ¹, Paula Lampe ¹, Martin Kaltenpoth ^1,³, Franziska Beran ^1,^2,^6,^✉

PMCID: PMC12976175 PMID: 41487052

Abstract

BACKGROUND

The efficacy of entomopathogenic nematodes (EPNs) in the biological control of insect pests can be influenced by the host's chemical defenses. Phyllotreta flea beetles, among the most destructive pests of Brassica crops, deploy highly reactive glucosinolate hydrolysis products as a defense against natural enemies. Here, we investigate the susceptibility of EPNs and their symbiotic bacteria to glucosinolate hydrolysis products and assess how this defense shapes the interaction between the horseradish flea beetle, Phyllotreta armoraciae, and EPNs.

RESULTS

Glucosinolate hydrolysis products were detected in uninjured P. armoraciae larvae but not in adults, and their levels were unaffected by EPN infection. EPNs and their bacterial symbionts were susceptible to glucosinolate hydrolysis products in vitro, with EPN immotility rates ranging from 35% to 96% and bacterial growth suppression from 20% to 85% at biologically relevant concentrations. However, reducing the levels of glucosinolate hydrolysis products in larvae, either by silencing myrosinase gene expression or by feeding on different Arabidopsis genotypes, did not make them more susceptible to EPNs. Nevertheless, the food plant influenced larval susceptibility to EPNs and the relative abundance of EPN bacterial symbionts in infected larvae.

CONCLUSION

Although glucosinolate hydrolysis products are toxic to EPNs and their symbiotic bacteria, they did not protect P. armoraciae larvae from EPN infection. However, the larval food plant influenced EPN susceptibility and bacterial community composition, highlighting the role of host plant traits in shaping insect–EPN interactions. These findings provide new insights into the limitations of EPN‐based biocontrol against glucosinolate‐sequestering pests. © 2026 The Author(s). Pest Management Science published by John Wiley & Sons Ltd on behalf of Society of Chemical Industry.

Keywords: chemical defense, Heterorhabditis, isothiocyanate, myrosinase, Photorhabdus, Phyllotreta

The flea beetle's glucosinolate‐based chemical defense fails to protect larvae from nematode infection. However, the defense inhibits the nematode's symbiotic bacteria, thereby potentially impairing nematode reproduction and biocontrol success.

graphic file with name PS-82-3603-g005.jpg

1. INTRODUCTION

Phyllotreta flea beetles are among the most destructive insect pests of Brassica crops worldwide. ¹ , ² , ³ , ⁴ The two economically most important species, Phyllotreta striolata and Phyllotreta cruciferae, are estimated to cause management costs of approximately 300 million USD per year in canola (Brassica napus) production in the northern Great Plains in North America alone. ² , ⁵ Adult beetles feed on aboveground plant tissues, producing a characteristic shot‐hole damage that can delay growth, reduce yield, or even kill seedlings. ³ , ⁶ Phyllotreta larvae typically feed on roots, but their damage is considered less severe than that caused by adults. ³ To reduce pesticide use, which currently represents the main means of controlling flea beetles in Brassica crops, it is important to develop alternative management strategies. ⁴ , ⁷ Within the framework of integrated pest management, the use of biological control agents offers a promising approach by exploiting natural enemies to suppress pest populations.

One important group of biological control agents against insect pests are entomopathogenic nematodes (EPNs) of the genera Steinernema and Heterorhabditis. ⁸ , ⁹ Their mode of action involves entering the host through body openings and releasing symbiotic bacteria, which rapidly kill the host through toxemia and septicemia, thereby enabling nematode reproduction. ⁸ , ¹⁰ Several studies have investigated the potential of EPNs as biocontrol agents against Phyllotreta flea beetles and the closely related cabbage stem flea beetle, Psylliodes chrysocephala, another major European flea beetle pest of winter oilseed rape, under laboratory and field conditions. ¹¹ , ¹² EPNs were able to kill larvae and adults of both Phyllotreta and Psylliodes in laboratory assays, although flea beetle susceptibility varied considerably among EPN species and strains. ¹² , ¹³ , ¹⁴ , ¹⁵ EPN performance also varied under field conditions and in some cases failed to reduce pest pressure. ¹³ , ¹⁶ , ¹⁷ , ¹⁸ , ¹⁹

The efficacy of EPNs is strongly influenced by biotic and abiotic factors, including temperature, soil characteristics, as well as nematode and host ecology. ⁸ , ²⁰ , ²¹ Plant defense compounds are increasingly recognized as important mediators of interactions between herbivorous insects and EPNs. ²¹ , ²² , ²³ , ²⁴ , ²⁵ For instance, Steinernema riobrave showed reduced virulence and reproduction when infecting tobacco‐fed corn earworm larvae (Helicoverpa zea) compared to larvae fed on tomato or eggplant. ²⁵ Although the underlying mechanism was not fully elucidated, defense metabolites from tobacco were suggested to play a role. Many herbivorous insects are able to sequester plant defense compounds for their own protection, ²⁶ , ²⁷ which can also influence susceptibility to EPNs. A well‐known example are western corn rootworm larvae (Diabrotica virgifera virgifera), which gain protection against EPNs by sequestering and metabolizing benzoxazinoids from their maize host plants. ²² In contrast, sequestered cucurbitacins failed to protect banded cucumber beetle larvae (Diabrotica balteata) against several tested Heterorhabditis species and strains. ²⁸ This was surprising because cucurbitacin D reduced growth of several bacterial symbionts of EPNs in vitro. ²⁹

Several Phyllotreta species were shown to sequester glucosinolates (GSLs) from their brassicaceous host plants. ³⁰ , ³¹ Together with plant myrosinase enzymes, GSLs form the characteristic activated defense of Brassicales, ³² , ³³ , ³⁴ producing highly reactive isothiocyanates and other hydrolysis products upon tissue damage. These metabolites confer resistance against non‐adapted herbivores and pathogens, ³⁵ , ³⁶ and, when applied as biofumigants, suppressed plant parasitic nematodes and pathogens in the soil. ³² , ³³ However, isothiocyanates can negatively affect infectivity of EPNs and may thus interfere with biological pest control. ³⁷ , ³⁸

Phyllotreta flea beetles not only sequester GSLs, they also possess endogenous myrosinase activity, allowing them to hydrolyze GSLs into reactive isothiocyanates. ³⁰ , ³¹ In the horseradish flea beetle, Phyllotreta armoraciae, sequestered GSLs are present in all life stages, while myrosinase activity occurs mainly in larvae and adults. ³¹ , ³⁹ Although the GSL‐myrosinase system protects P. armoraciae larvae against generalist predators, ³¹ , ³⁹ its efficacy against EPNs is unknown. Here, we investigated the susceptibility of EPNs and their bacterial symbionts to GSL hydrolysis products produced by P. armoraciae in vitro and in vivo. We show that while GSL hydrolysis products negatively affect EPNs, the GSL‐myrosinase system does not protect larvae from EPN infection. However, sequestered GSLs reduced the relative abundance of bacterial symbionts, which may influence EPN reproduction.

2. MATERIALS AND METHODS

2.1. Biological resources

Phyllotreta armoracia beetles were reared on 3‐ to 4‐week‐old potted Brassica juncea cv. ‘Bau‐Sin’ plants (Known‐You Seeds, Kaohsiung, Taiwan) in a controlled‐environment chamber at 23 °C, 60% relative humidity, and a 16 h:8 h light/dark cycle. Adults were provided with new plants once a week for feeding and oviposition. Plants with eggs were kept in separate cages for larval development under the same conditions. Any remaining leaf material was removed 3 weeks later, and the soil with pupae was kept in plastic containers (9 L volume; Lock & Lock storage container). Newly emerged adults were collected every 2–3 days.

Entomopathogenic nematodes Heterorhabditis bacteriophora (nematop®), Steinernema feltiae (nemaplus®), and Steinernema carpocapsae (nemastar®) were purchased from e‐nema GmbH (Schwentinental, Germany) and stored at 4 °C until they were used for experiments. For P. armoraciae survival assays (described in Section 2.5), H. bacteriophora were reared on Galleria mellonella larvae (Aquaexot, Jena, Germany) as previously described and used within 2 weeks of emergence. ⁴⁰

Symbiotic bacteria Photorhabdus laumondii, Xenorhabdus bovienii, and Xenorhabdus nematophila were directly isolated from H. bacteriophora, S. feltiae, and S. carpocapsae, respectively, by plating surface‐sterilized and homogenized EPNs on nutrient bromothymol blue triphenyltetrazolium agar (NBTA) plates. The identities of isolated bacteria were confirmed by Sanger sequencing of polymerase chain reaction (PCR) products obtained with universal 16S rRNA primers (sequences provided in Table S1). ⁴¹

Arabidopsis thaliana Col‐0 and tgg1 × tgg2 (tgg) double knockout mutant plants lacking myrosinase activity in leaves were cultivated at 21 °C, 60% relative humidity, and a 10 h:14 h light/dark cycle in a controlled environment chamber. ⁴²

2.2. Quantification of GLS hydrolysis products in EPN‐infected and uninfected larvae and adults

To quantify the levels of 2‐propenyl GSL‐derived hydrolysis products in non‐infected and EPN‐infected beetles, we exposed P. armoraciae third‐instar larvae and 6‐ to 8‐day old, unsexed adults to H. bacteriophora. Groups of five larvae or five adults were exposed to 5000 EPNs in 480 μL tap water in Petri dishes (3 cm diameter) lined with filter paper at top and bottom (N = 11–18 for adults, N = 12 for larvae, five individuals per sample). Beetles were incubated for 16 h at 21 °C in the dark. Larvae and adults kept under the same conditions without EPNs served as control groups. For quantification of GSL hydrolysis products, larvae and adults were weighed, flash‐frozen in liquid nitrogen, and stored at −80 °C until extraction. To verify infection, we transferred a subset of larvae and adults of inoculated and control groups to new Petri dishes without EPNs and monitored survival after 48 h (N = 5–10 for adults, N = 5 for larvae).

GSL hydrolysis products were extracted from homogenized samples in ice‐cold octane containing benzonitrile as an internal standard. Products were identified by gas chromatography–mass spectrometry (GC–MS) by comparing retention times and mass spectra to authentic standards, and quantified by gas chromatography‐flame ionization detection (GC‐FID; Table S2). Details of the extraction and analytical parameters are provided in Supporting Information Method S1. Concentrations of 2‐propenyl isothiocyanate were compared between treatment groups with Mann–Whitney rank sum test in SigmaPlot 14.0.

2.3. Susceptibility of EPNs to GSL hydrolysis products

To analyze the effects of GSL hydrolysis products on H. bacteriophora, S. feltiae, and S. carpocapsae, we exposed EPNs to 2‐propenyl isothiocyanate and 3‐butenenitrile over a concentration range of 0.1 to 1000 μm in 10‐fold dilutions. Assays were carried out in 48‐well plates with 150 EPNs in 250 μL of tap water containing the test compounds (N = 6 per compound and concentration, divided over two experimental days). Pure tap water was used as a control. After 16 h incubation at 24 °C, we assessed the status of 50 randomly selected EPNs: normal motility and immotile (elongated) per assay.

To assess the effects of GSL hydrolysis products on EPN motility, we fitted generalized linear mixed‐effects models with a binomial error distribution and logit link function using the glmer() function from the lme4 package in R v4.3.1. ⁴³ , ⁴⁴ Analyses were conducted separately for each EPN species. Fixed effects included compound, their concentration (treated as ordinal variable), and their interactions.

Well position, experimental day and the interaction of both were considered as random intercepts, and the best model was selected via Akaike information criterion (AIC) comparison. Model selection to test for the influence of the fixed effects selection followed a backward sequential approach based on likelihood ratio tests. Terms were retained only if their removal significantly reduced model fit. Model assumptions were checked using the DHARMa package. ⁴⁵ Post hoc comparisons of estimated marginal means were performed using emmeans, ⁴⁶ with Tukey adjustment for multiple testing. Due to known batch effects, we did not directly compare the immotility rates between the different EPN species. The stability of the GLS hydrolysis products was estimated using proton nuclear magnetic resonance (¹H NMR) spectroscopy prior to conducting the biological experiments (see Supporting Information Method S2 and Fig. S1).

2.4. Susceptibility of bacterial symbionts to GSL hydrolysis products

To analyze the effects of GSL hydrolysis products on the bacterial symbionts P. laumondii, X. bovienii, and X. nematophila, we exposed liquid bacterial cultures to 2‐propenyl isothiocyanate and 3‐butenenitrile over a concentration range of 4.8 to 2500 μm in 3.5‐fold dilutions. Assays were conducted in 96‐well plates by adding 100 μL of an overnight culture grown in Müller–Hinton (MH) medium diluted in pure MH medium to an optical density measured at 600 nm (OD_600nm) = 0.1 to 100 μL of MH medium containing the test compound (N = 6 per bacterial strain and concentration). Pure MH medium was used as a control. The plate was sealed with adhesive foil and incubated at 30 °C for 36 h with shaking in a plate reader (Tecan, Männedorf, Switzerland). OD_600nm values were recorded every 20 min.

Data analysis was conducted using R v4.3.1. ⁴³ The growth observed in each well was quantified as the area under the growth curve (AUC). To compensate for different growth of individual bacterial strains, the AUCs for each bacterium‐compound combination were normalized on a scale from 0 (mean AUC of wells only containing media without bacteria) to 1 (mean AUC of wells with media and bacteria but without GSL hydrolysis products). A dose–response log‐logistic model was fitted to the relationship between AUC and concentration using the drc package. ⁴⁷ The 50% inhibitory concentrations (IC₅₀) were calculated based on this model.

2.5. Impact of Phyllotreta armoraciae larval defense on susceptibility to EPNs

The influence of the beetle's chemical defense on the interaction with the EPN H. bacteriophora was assessed in two experiments. In Experiment 1, we reduced the levels of myrosinase activity by injecting second instar larvae with double‐stranded RNA (dsRNA) targeting PaMyr2 or a lepidopteran metalloproteinase inhibitor from G. mellonella (IMPI; AY330624.1) as control as previously described. ³⁹ Injected larvae were provided with B. juncea petioles and kept in the laboratory under ambient conditions. Due to limited availability of beetle larvae, we determined the impact of PaMyr2 silencing on GSL hydrolysis product levels and on the susceptibility in two separate RNA‐interference (RNAi) experiments. To first determine whether PaMyr2 silencing reduces GSL hydrolysis product levels in uninjured larvae, we sampled third‐instar larvae 6 days after dsRNA injection for myrosinase assays (N = 8 per treatment, two larvae per sample) and GSL hydrolysis product analysis (N = 12–13 per treatment, five larvae per sample). Myrosinase activity was measured in crude protein extracts and compared between treatments using Welch's t‐test in SigmaPlot 14.0. ³⁹ GSL hydrolysis products were analyzed as described in Section 2.2, and compared between treatments using generalized least squares models implemented via the gls() function from the nlme package in R v4.3.1. ⁴³ , ⁴⁸ Treatment and hydrolysis product (2‐propenyl isothiocyanate and 3‐butenenitrile) served as explanatory factors. To account for heteroscedasticity between treatment groups and hydrolysis products, we specified a group specific variance structure using the varIdent() function for both factors. Model significance was assessed via model simplification and subsequent likelihood ratio tests comparing models as recommended by Zuur. ⁴⁹ Post hoc multiple comparisons of estimated marginal means were conducted using Tukey contrasts (glht() function, multcomp package). ⁵⁰

After confirming that PaMyr2 silencing reduces GSL hydrolysis product levels in larvae, we performed survival assays with dsPaMyr2‐ and dsIMPI‐injected larvae. Six days post‐injection, groups of five larvae were exposed to 100 EPNs in 480 μL tap water in Petri dishes (3 cm diameter) lined with filter paper at top and bottom (N = 10 per treatment, five larvae per replicate). Larvae kept under the same conditions without EPNs served as control groups (N = 3 per treatment, five larvae per replicate). Larval mortality was monitored after 48 h exposure at 21 °C in the dark. No mortality was observed in controls without EPNs. Dead larvae were transferred to new Petri dishes without EPNs to visually determine the red coloration rate (not red or red) after 48 h and EPN reproduction after 96 h. Larval survival and red coloration rates were compared between treatments using Chi‐squared test with Yates's correction for continuity in R v4.3.1. ⁴³ To correlate red coloration of cadavers with EPN reproduction, we collected emerging EPNs using White traps and counted them. ⁵¹

In Experiment 2, we manipulated the levels of sequestered GSLs by feeding P. armoraciae first‐instar larvae with two different A. thaliana genotypes for 8 days: Col‐0 wild‐type and myrosinase deficient tgg mutant plants. We previously showed that larvae can sequester GSLs from leaves of the tgg mutant, but not from leaves of the corresponding Col‐0 wild‐type. ⁵² As controls, we sampled a subset of Col‐0 and tgg‐fed larvae for GSL analysis (N = 8 per treatment, three larvae per sample) and for quantification of GSL hydrolysis products (N = 8–10 per treatment, four larvae per sample). The weights of Col‐0 and tgg‐fed larvae sampled for GSL and GSL hydrolysis product analyses were compared using two‐sample t‐tests or Welch's test, respectively, in SigmaPlot 14.0.

GLS were extracted and analyzed by high‐performance liquid chromatography with a diode array detector (HPLC‐DAD) as previously described. ³⁰ GSL hydrolysis products were extracted from larval samples using ice‐cold dichloromethane containing benzonitrile as an internal standard. Following homogenization and centrifugation, the products were quantified by GC–MS and GC‐FID. Details of the extraction and analytical parameters are provided in the Supporting Information Method S3. Total concentrations of GSLs and GSL hydrolysis products were compared between treatment groups with Mann–Whitney rank sum test in SigmaPlot 14.0.

Five larvae were exposed to 200 EPNs in 480 μL tap water in Petri dishes (3 cm diameter) lined with filter paper at top and bottom (N = 12–14 per treatment). Larvae kept under the same conditions without EPNs were used to determine the background mortality (N = 3 per treatment). Larval mortality was monitored after 48 h exposure at 21 °C in the dark. No mortality was observed in controls without EPNs. Dead and alive larvae were separated and transferred to new Petri dishes without EPNs and incubated for another 48 h to determine the red coloration rate of dead larvae. Larval survival and red coloration rates were compared between treatments using Chi‐squared test with Yates's correction for continuity and Fisher's Exact test, respectively, in R v4.3.1. ⁴³

2.6. Impact of the chemical defense of Phyllotreta armoraciae larvae on bacterial symbionts

Because EPN reproduction in cadavers varied strongly in Experiment 1, we instead aimed to quantify P. laumondii abundance in EPN‐exposed larvae in Experiment 2. Therefore, all larvae (dead and alive) of the same biological replicate were pooled, weighed, frozen in liquid nitrogen and stored at −20 °C until DNA extraction using the Quick‐DNA Tissue/Insect Microprep Kit (Zymo Research, Freiburg, Germany). Because several attempts to design P. laumondii‐specific primers were unsuccessful (sequences provided in Table S1), we estimated total bacterial 16S gene copy numbers using universal BactQuant primers (Table S1). ⁵³ Total bacterial abundance in DNA extracts were estimated by quantitative PCR (qPCR) in optical 96‐well plates on a CFX Connect Real‐Time‐System (Bio‐Rad, Feldkirchen, Germany) using the Biozym Blue S'Green qPCR Kit Separate ROX Kit (Biozym Scientific, Hessisch Oldendorf, Germany). Quantification was done using a serial dilution of P. laumondii 16S rDNA. The PCR program was as follows: 95 °C for 15 min, 59 cycles of 95 °C for 15 s, 60 °C for 30 s and 72 °C for 30 s. Afterwards, the melting curve was determined from 55 °C to 95 °C in 0.5 °C increments.

To estimate the relative abundance of bacterial symbionts, we performed 16S amplicon sequencing of the V3–V4 region (N = 11–12). Raw reads were processed using a pipeline incorporating DADA2 and phyloseq to generate an amplicon sequence variant (ASV) table and assign taxonomy against the SILVA database. ⁵⁴ , ⁵⁵ , ⁵⁶ , ⁵⁷ Detailed information on the 16S amplicon sequencing data analysis and the complete R script are provided in the Supporting Information Method S4 and Data S2.

The relative abundance of P. laumondii in Col‐0 and tgg‐fed larvae was compared using a two‐sample t‐test. Only samples in which P. laumondii was detected were included and data were square‐root transformed prior to analysis. The total abundance of P. laumondii in larvae was estimated by multiplying the relative abundance of Photorhabdus determined by 16S sequencing with the total bacterial abundance determined by qPCR and compared between Col‐0‐ and tgg‐fed larvae using a two‐sample t‐test. To test whether the food plant and relative abundance of P. laumondii predict the total bacterial abundance in beetle samples, we fitted a linear model. Model selection followed a forward sequential approach based on F‐tests using analysis of variance (ANOVA) and AIC. All analyses were performed using R v4.3.2. ⁴³

3. RESULTS

3.1. GSL hydrolysis products are present in Phyllotreta armoraciae larvae but not adults

To determine whether EPNs are exposed to toxic GSL hydrolysis products during infection of P. armoraciae, we quantified the levels of 2‐propenyl isothiocyanate and 3‐butenenitrile in EPN‐infected and uninfected larvae (100% infection rate) and adults (20% infection rate). In larvae, we primarily detected 2‐propenyl isothiocyanate, whereas 3‐butenenitrile was only present in trace amounts in nematode‐infected individuals (3.46 ± 8.27 pmol mg fresh weight (FW)⁻¹). Concentrations of 2‐propenyl isothiocyanate did not differ between EPN‐infected and uninfected larvae (Mann–Whitney rank sum test, N = 12, U = 52, P = 0.260). Assuming that GSL hydrolysis products are predominantly localized in the hemolymph, the detected isothiocyanate levels correspond to average hemolymph concentrations (± standard deviation (SD)) of 106.8 ± 56.7 μm in uninfected larvae and 87.5 ± 75.3 μm in EPN‐infected larvae (Fig. 1). In adults, only traces of 2‐propenyl isothiocyanate were detected in nematode‐infected individuals in two out of 18 samples (0.55 ± 1.61 pmol mg FW⁻¹).

Isothiocyanates are present in uninjured larvae and are not induced upon EPN infection. Concentrations of 2‐propenyl glucosinolate‐derived isothiocyanate (ITC) and nitrile in larvae exposed to *Heterorhabditis bacteriophora* or uninfected control larvae. Boxplots show median, interquartile range, and individual replicate values. 3‐Butenenitrile levels were not compared. The levels of 2‐propenyl isothiocyanate did not differ significantly (n.s.; Mann–Whitney rank sum test, N = 12, U = 52, P = 0.26).

3.2. EPNs and their bacterial symbionts are susceptible to GSL hydrolysis products

To assess the sensitivity of EPNs and their bacterial symbionts to 2‐propenyl isothiocyanate and 3‐butenenitrile, we exposed H. bacteriophora, S. feltiae, and S. carpocapsae as well as their bacterial symbionts P. laumondii, X. bovienii, and X. nematophila to different concentrations of these GSL hydrolysis products. In all three EPN species, immotility rates increased with GSL hydrolysis product concentration, indicating a dose‐dependent toxicity for both metabolites (Fig. 2(A)–(C)). For H. bacteriophora and S. feltiae, 2‐propenyl isothiocyanate caused higher immotility rates than 3‐butenenitrile, with post hoc tests confirming this difference at concentrations of 100 μm and 10 μm, respectively (Fig. 2(A),(B)). In contrast, for S. carpocapsae there was no significant main effect of the hydrolysis product, but a significant interaction between compound and concentration. Although 2‐propenyl isothiocyanate appeared more toxic at intermediate concentrations, post hoc tests did not confirm a significant difference at any specific concentration (Fig. 2(C)). At physiologically relevant 2‐propenyl isothiocyanate concentrations (~100 μm), we observed average EPN immotility rates of 96% for H. bacteriophora, 80% for S. feltiae, and 64% for S. carpocapsae in our in vitro assays.

Dose‐dependent toxicity of 2‐propenyl glucosinolate hydrolysis products against EPNs and their symbiotic bacteria *in vitro*. (A–C) EPNs were exposed for 24 h to increasing concentrations of 2‐propenyl isothiocyanate and 3‐butenenitrile. Data points represent the mean EPN immotility rate (± standard deviation). Dashed and dotted horizontal lines indicate 50% and 25/75% immotility rate, respectively. (A) *Heterorhabditis bacteriophora*, generalized linear mixed model (GLMM), N = 6, hydrolysis product × concentration: χ ² = 36.989, P < 0.001, hydrolysis product: χ ² = 4.382, P = 0.036, concentration: χ ² = 2146.3, P < 0.001. (B) *Steinernema feltiae*, GLMM, N = 6, hydrolysis product × concentration: χ ² = 33.655, P < 0.001, hydrolysis product: χ ² = 6.065, P = 0.014, concentration: χ ² = 1926.3, P < 0.001. (C) *Steinernema carpocapsae*, GLMM, N = 6, hydrolysis product × concentration: χ ² = 11.378, P = 0.044, hydrolysis product: χ ² = 1.4105, P = 0.235, concentration: χ ² = 523.76, P < 0.001. Asterisks indicate a significant difference in immotility between the two compounds at a given concentration (P < 0.05). Growth inhibition of the symbiotic bacteria (D) *Photorhabdus laumondii*, (E) *Xenorhabdus bovienii*, and (F) *Xenorhabdus nematophila*. Bacterial growth, measured after 36 h, is shown as the normalized area under the curve (N = 5–6). Growth was normalized on a scale from 0 (blank medium) to 1 (untreated controls, gray boxplots). Boxplots show the median, interquartile range, and individual replicate values. Dashed and dotted horizontal lines indicate 50% and 25/75% of control growth, respectively.

In growth assays performed with bacterial symbionts, 2‐propenyl isothiocyanate inhibited growth in a dose‐dependent manner, with complete growth suppression at 204 μm (Fig. 2(D)–(F)). The IC₅₀ ± SD values were 6.8 ± 0.5 μm for X. bovienii, 26.3 ± 2.4 μm for X. nematophila and 58.4 ± 2.0 μm for P. laumondii. 3‐Butenenitrile only partially suppressed growth in the three bacterial strains, precluding reliable estimation of IC₅₀ values. These results indicate that 2‐propenyl isothiocyanate is more potent against bacterial symbionts of EPNs than its corresponding nitrile and suggest that this compound has the potential to impair growth of bacterial symbionts in the hemolymph of P. armoraciae larvae.

3.3. GSL hydrolysis products do not protect Phyllotreta armoraciae larvae against EPNs

To directly test whether PaMyr2 hydrolyzes sequestered 2‐propenyl GSL in uninjured larvae, we silenced PaMyr2 expression and compared the levels of myrosinase activity and GSL hydrolysis products with those in dsIMPI‐injected control larvae. PaMyr2 silencing decreased myrosinase activity by 78% (myrosinase activity ± SD, dsIMPI, 1.01 ± 0.46 nmol glucose min⁻¹ mg FW⁻¹; dsPaMyr2, 0.23 ± 0.46 nmol glucose min⁻¹ mg FW⁻¹; Mann–Whitney rank sum test, N = 8, U = 7, P = 0.007). In this experiment, both 2‐propenyl isothiocyanate and 3‐butenenitrile were detected in larvae, with 2‐propenyl isothiocyanate accounting for approximately 90% of the total detected GSL hydrolysis products (Fig. S2). PaMyr2‐silenced larvae contained 82% less 2‐propenyl isothiocyanate and 76% less 3‐butenenitrile than control larvae (method of generalized least squares, N = 12–13, treatment × hydrolysis product, LR = 22.475, P < 0.001; treatment, LR = 15.240, P < 0.001; hydrolysis product, LR = 13.762, P < 0.001).

Next, we determined whether PaMyr2 silencing influences larval susceptibility to EPNs by exposing PaMyr2‐silenced and control larvae to H. bacteriophora. In this experiment, PaMyr2 silencing reduced myrosinase activity by 96% compared to the control (Welch's t‐test, N = 8, t = 8.371, P < 0.001; Fig. 3(A)). Neither the mortality rate nor the red coloration rate of cadavers differed between larvae with high and low myrosinase activity (mortality, Chi‐square test, N = 50, χ ² = 2.042, P = 0.153, Fig. 3(B); red coloration rate, Chi‐square test, N = 26–34, χ ² = 2.247, P = 0.134, Fig. 3(C)). EPN reproduction was only observed in replicates with red larvae, with EPN counts ranging between 71 and 945 per dead larva (Table S3).

The glucosinolate‐myrosinase system does not protect *Phyllotreta armoraciae* larvae against an EPN. (A) *PaMyr2* silencing reduced myrosinase activity in larvae (*dsPaMyr2*) compared to control larvae. Boxplots show median, interquartile range, and individual replicate values. (B) Mortality rate of *PaMyr2*‐silenced larvae and control larvae exposed to *Heterorhabditis bacteriophora*. The proportion of dead larvae is shown in gray with a skull. (C) Anthraquinone‐induced red coloration in larval cadavers (red). (D) Concentrations of glucosinolate hydrolysis products in larvae reared on *Arabidopsis thaliana* Col‐0 wild‐type and *tgg1* × *tgg2* (*tgg)* myrosinase double knock‐out mutant plants. Boxplots show median, interquartile range, and individual replicate values. (E) Mortality of larvae with high (*tgg*) and low (Col‐0) levels of glucosinolates and glucosinolate hydrolysis products exposed to *Heterorhabditis bacteriophora*. (F) Anthraquinone‐induced red coloration in larval cadavers (red).

Because PaMyr2‐silenced larvae likely still contained GSL hydrolysis products (Fig. S2), we employed an alternative strategy to obtain larvae with further reduced metabolite levels. Larvae fed with Arabidopsis Col‐0 leaves sequestered only minimal amounts of GSL compared to larvae fed with leaves from the Arabidopsis tgg mutant (Mann–Whitney rank sum test, N = 8, U = 0, P < 0.001; Table S4) and consequently harbored markedly lower levels of GSL hydrolysis products (Mann–Whitney rank sum test, N = 8–10, U = 0, P < 0.001; Fig. 3(D) and Table S4). We hypothesized that larvae without GSL are more susceptible to H. bacteriophora than larvae with GSL, but contrary to our expectation, their mortality rate was even lower than that of larvae with GSL (Chi‐square test, N = 60–70, χ ² = 4.862, P = 0.027; Fig. 3(E)). However, none of the larval cadavers with GSLs developed red coloration, whereas 43% of cadavers without GSLs turned red (Fisher's Exact test, N = 14–24, P = 0.001; Fig. 3(F)). This observation suggested that the chemical defense of P. armoraciae larvae negatively affected P. laumondii growth.

3.4. The larval food plant influences the relative abundance of bacterial symbionts

To assess P. laumondii abundance in larvae with and without GSLs, we quantified total bacterial abundance by qPCR and the relative abundance of P. laumondii by 16S amplicon sequencing. Photorhabdus laumondii was detected in only a few samples (tgg, N = 4 out of 11; Col‐0, N = 5 out of 12; Fig. S3). Although the relative abundance of P. laumondii was higher in larvae without GSLs (t‐test, N = 4–5, t = 2.81, P = 0.026; Fig. 4(A)), the absolute P. laumondii abundance did not differ between larvae with and without GSLs (t‐test, N = 4–5, t = 1.50, P = 0.177; Fig. 4(B)).

Host chemical defense reduces the relative, but not the absolute, abundance of *Photorhabdus laumondii* in *Phyllotreta* larvae. Boxplots comparing the abundance of *Photorhabdus laumondii* in EPN‐infected larvae with high (*tgg*) and low (Col‐0) levels of glucosinolates and glucosinolate hydrolysis products. (A) The relative abundance of *Photorhabdus laumondii* within the bacterial community, determined by 16S amplicon sequencing, was lower in high‐glucosinolate larvae. (B) The estimated absolute abundance of *Photorhabdus laumondii*, calculated from total bacterial DNA and relative abundance data, did not differ between the two treatments. Boxplots show median, interquartile range, and individual replicate values. n.s., not significant.

Because both GSL hydrolysis products and Photorhabdus natural products are known to have antimicrobial activity, ⁵⁸ , ⁵⁹ , ⁶⁰ we tested whether total bacterial abundance in larvae depended on the food plant and the relative abundance of P. laumondii. Total bacterial abundance in EPN‐infected larvae was not affelacted by the food plant alone, but was influenced by the relative abundance of P. laumondii and its interaction with the food plant (linear model, N = 11–12, Photorhabdus relative abundance × food plant: F = 5.88, P = 0.025, Photorhabdus relative abundance: t = 4.72, P < 0.001, adjusted R ² = 0.49; food plant: t = 1.508, P = 0.149, adjusted R ² = 0.615). Specifically, total bacterial abundance increased more strongly with increasing Photorhabdus relative abundance in tgg‐fed larvae (with GSLs) than in Col‐0‐fed larvae (without GSLs; Fig. S4).

4. DISCUSSION

The sequestration of plant defense metabolites by herbivorous insects is a widespread defense strategy against predators, but its impact on other natural enemies such as entomopathogens remains poorly understood. EPNs are an important group of biocontrol agents against crop pests, yet EPN efficacy against GSL‐sequestering Phyllotreta flea beetle pests of Brassica crops showed considerable variation under both laboratory and field conditions. ¹³ , ¹⁴ , ¹⁵ , ¹⁷ Here we demonstrate that EPNs and their bacterial symbionts are susceptible to GSL hydrolysis products produced in P. armoraciae larvae, but that GSL hydrolysis product levels do not correlate with larval susceptibility to EPNs.

To protect Phyllotreta flea beetles against EPN infection, sequestered GSL must be hydrolyzed by endogenous myrosinases to form toxic isothiocyanates. Such activation of sequestered pro‐toxins upon entomopathogen attack has been documented in other insects, for example in GLS‐sequestering cabbage aphids (Brevicoryne brassicae) infected with the entomopathogenic fungus Beauveria bassiana, and in western corn rootworm larvae, which hydrolyze sequestered 2‐hydroxy‐4,7‐dimethoxy‐1,4‐benzoxazin‐3‐one‐O‐glucoside to 6‐methoxy‐2‐benzoxazolinone upon EPN infection. ²² , ⁶¹ In P. armoraciae, EPN infection did not increase levels of GSL hydrolysis products in larvae or adults (Fig. 1), suggesting that EPNs and their bacterial symbionts either do not activate the defense or actively suppress myrosinase activity in the host. Although GSL hydrolysis was not induced upon EPN infection, we still detected 2‐propenyl isothiocyanate and traces of the corresponding nitrile in both uninjured and infected larvae (Fig. 1). In contrast, no 2‐propenyl GSL‐derived hydrolysis products were detected in adults, despite previous reports of 4‐methylsulfinylbutyl GSL hydrolysis products in adults that had fed on Arabidopsis leaves. ⁵² The differences between larvae and adults are likely explained by different biochemical properties of myrosinase enzymes in larvae and adults, as well as higher levels of myrosinase activity in larvae compared with adults. ³¹ , ³⁹ The detection of hydrolysis products derived from 4‐methylsulfinylbutyl GSL but not 2‐propenyl GSL in adults furthermore suggests that hydrolysis products derived from different GSLs are metabolized at different rates.

Due to their electrophilic properties, isothiocyanates are toxic to a broad range of organisms, including bacteria, nematodes, and insects, some of which have evolved counter‐adaptations to overcome isothiocyanate‐based defenses. ³² , ⁶² , ⁶³ , ⁶⁴ , ⁶⁵ Other GSL hydrolysis products, such as nitriles, are generally considered less toxic ⁶⁶ , ⁶⁷ , ⁶⁸ , ⁶⁹ ; however, their biological effects have been investigated far less extensively. EPNs and their symbiotic bacteria infecting P. armoraciae larvae primarily encounter 2‐propenyl isothiocyanate but also small amounts of 3‐butenenitrile (Fig. 1). Both compounds caused similar immotility rates in a dose‐dependent manner for all three EPN species, reaching up to 100% at the highest concentration (~1 mm) in our in vitro assays (Fig. 2(A)–(C)). The sensitivity of the three EPNs to 2‐propenyl isothiocyanate is comparable to that of the root knot nematodes Meloidogyne incognita and Meloidogyne hapla, ⁷⁰ , ⁷¹ whereas Heterodera spp. appear to be less susceptible to this isothiocyanate. ⁷² , ⁷³ The susceptibility of EPNs to 3‐butenenitrile was unexpected, and to our knowledge, the effects of GSL‐derived nitriles on nematodes have not yet been investigated. These findings raise the question of whether nitriles may also affect plant‐parasitic nematodes and thus contribute to direct plant defense.

In the EPNs' bacterial symbionts, 2‐propenyl isothiocyanate suppressed growth in a dose‐dependent manner, with complete inhibition at ~200 μm (Fig. 2(D)–(F)). This concentration is markedly lower than the minimum inhibitory concentration of 2‐propenyl isothiocyanate (between 0.5 and 2 mm) determined for several other bacterial isolates. ⁷⁴ , ⁷⁵ By contrast, 3‐butenenitrile had minimal impact on P. laumondii and X. nematophila growth, and a dose–response relationship was only observed in assays with X. bovienii, showing species‐specific sensitivity of bacterial symbionts to different GSL hydrolysis products, including nitriles. The minor effects of 3‐butenenitrile on bacterial growth are consistent with earlier studies using disc diffusion assays, ⁶³ and support the hypothesis that nitriles are less toxic to bacteria than isothiocyanates. The susceptibility of EPNs and their bacterial symbionts to 2‐propenyl isothiocyanate likely explains the negative effects of mustard biofumigation on the biocontrol potential and infectivity of EPNs in glasshouse and field conditions, ³⁷ , ³⁸ and highlights the potential for selecting or engineering increased isothiocyanate resistance to improve EPN performance in Brassica crops.

Although biologically relevant levels of 2‐propenyl isothiocyanate killed EPNs and inhibited growth of their bacterial symbionts in vitro (Fig. 2), 2‐propenyl GSL hydrolysis product levels did not correlate with larval susceptibility to H. bacteriophora (Fig. 3(B),(C)). Since residual isothiocyanate levels in PaMyr2‐silenced larvae might still have affected interactions with EPNs, we increased differences in hydrolysis product levels by feeding larvae with Arabidopsis Col‐0 wild‐type or tgg leaves. Despite a 99% reduction in GSL hydrolysis products in Col‐0‐fed larvae relative to tgg‐fed larvae, Col‐0‐fed larvae survived even better than tgg‐fed larvae (Fig. 3(E)), suggesting that feeding on Col‐0 alters physiological traits that influence larval susceptibility to EPNs. One possible explanation could be that GSL sequestration and hydrolysis in tgg‐fed larvae incurs physiological costs that influence the ability to prevent EPN infection. Indeed, we found that Col‐0‐fed larvae sampled for GSL analysis and DNA extraction were heavier than the corresponding tgg‐fed larvae (GSLs, t‐test, N = 8, t = 2.569, P = 0.021; DNA extraction, t‐test, N = 12–14, t = 3.008, P = 0.006), but weights of Col‐0‐ and tgg‐fed larvae sampled for GSL hydrolysis product analyses did not differ in weight (Welch's t‐test, N = 8–10, t = 0.486, P = 0.635). Further research is needed to understand the physiological costs of GSL sequestration and both plant and insect myrosinase activity in larvae.

Insects killed by P. laumondii usually turn red due to the bacterial biosynthesis of anthraquinones, which accumulate in the cadaver. Although the biological function of anthraquinone biosynthesis is not well‐understood, ⁷⁶ , ⁷⁷ , ⁷⁸ red coloration of cadavers is widely used as a visual marker for bacterial growth. The red coloration rate did not differ between larvae with high and low levels of myrosinase activity and 2‐propenyl isothiocyanate, but was higher in Col‐0‐fed larvae, which are largely devoid of GSL hydrolysis products, than in tgg‐fed larvae (Fig. 3(F)). This suggests that Arabidopsis GSL hydrolysis products suppressed the growth of P. laumondii in larvae. However, quantification of P. laumondii revealed no difference in absolute abundance between Col‐0‐ and tgg‐fed larvae, although relative abundance was higher in Col‐0‐fed larvae, indicating a potential competitive advantage when the host is not chemically defended (Figs 4 and S3). The apparent discrepancy between red coloration and bacterial abundance may reflect reduced metabolic activity of P. laumondii in tgg‐fed larvae, potentially due to GSL hydrolysis products or other host physiological factors.

While our data suggest that GSL hydrolysis products negatively affect the relative abundance of P. laumondii in larvae (Fig. 4(A)), a deeper analysis revealed a more complex interaction: the total bacterial load increased more strongly with increasing P. laumondii abundance in tgg‐fed larvae, which are rich in GSL hydrolysis products, compared to Col‐0‐fed larvae (Fig. S4). We propose that this apparent contradiction can be explained by a mechanism of competitive release, where the host's chemical defense changes the competitive landscape in favor of the adapted resident microbiota. In Col‐0‐fed larvae without GSLs, P. laumondii likely acts as a classic pathogen, using its own antimicrobial compounds to suppress competing bacteria and establish dominance. ⁵⁸ In this scenario, when P. laumondii replaces the existing community, it leads to a modest increase in the total bacterial load. By contrast, the native P. armoraciae bacteria proliferate in tgg‐fed with GSLs, as they have probably co‐evolved and adapted to these chemical defense mechanisms. When H. bacteriophora EPNs introduce P. laumondii, the GSL hydrolysis products may inhibit the non‐adapted P. laumondii more strongly, giving the adapted native bacteria a competitive advantage. This allows the native bacteria to proliferate alongside P. laumondii in the host, resulting in a steeper increase in the total bacterial load for a given increase in Photorhabdus relative abundance (Figs S3 and S4).

In conclusion, our study reveals that GSL hydrolysis products, despite their strong toxicity toward EPNs and their bacterial symbionts in vitro, do not confer direct protection to P. armoraciae larvae in vivo. Instead, the host's chemical defense appears to influence the post‐infection microbial environment in a way that alters pathogen performance. We suggest that isothiocyanate‐mediated interference with P. laumondii growth may give an advantage to resident, chemically adapted microbiota, thereby limiting nematode reproduction. This highlights that biocontrol efficacy against chemically defended herbivores cannot be understood solely through toxin–pathogen interactions, but must also consider the competitive dynamics within the host microbiome. Based on our findings, we suggest that a deeper understanding of interactions between EPN bacterial symbionts and host microbiota, together with the selection of isothiocyanate‐tolerant bacterial strains, represents a promising route to improving the reliability of EPN‐based biocontrol of Phyllotreta flea beetle pests.

CONFLICT OF INTEREST

The authors declare that they have no conflicts of interest with the contents of this article.

Supporting information

Method S1. Quantification of GLS hydrolysis products in EPN‐infected and uninfected larvae and adults.

Method S2. Stability of GSL hydrolysis products in water

Method S3. Identification and quantification of GSL hydrolysis products in larvae fed with Arabidopsis

Method S4. 16S Amplicon sequencing of bacterial communities associated with Phyllotreta armoraciae larvae.

Figure S1. 2‐Propenyl isothiocyanate decay over time. (A) ¹H NMR spectra of a 10 μL sample of 2‐propenyl isothiocyanate in 600 μL of D₂O. The spectra were recorded immediately after dilution and after 14 h and 720 h (30 days). The signal at 4.14 ppm belongs to the methylene group adjacent to the isothiocyanate moiety. Initially, the 2‐propenyl isothiocyanate sample contained an impurity, marked by an asterisk. The intensity of this signal did not increase over time, but other hydrolysis products appeared (indicated by the arrows on the top curve). (B) Estimation of the decay rate of 2‐propenyl isothiocyanate in D₂O. According to the fit function, no more than 2% of 2‐propenyl isothiocyanate should undergo spontaneous hydrolysis during the biological experiments.

Figure S2. Isothiocyanate and nitrile formation are reduced upon PaMyr2‐silencing in Phyllotreta armoraciae larvae. Concentrations of 2‐propenyl glucosinolate‐derived isothiocyanates (orange) and nitriles (blue) in control and PaMyr2‐silenced larvae. Boxplots show median, interquartile range, and individual replicate values. Different letters indicate differences between groups (P < 0.05).

Figure S3. Impact of larval diet and entomopathogenic nematode (EPN) infection on the composition of the bacterial community associated with Phyllotreta armoraciae larvae. Stacked bar chart showing the relative abundance of bacterial genera in larval samples. Each bar represents one biological replicate of five pooled larvae. Samples are organized into four panels based on larval diet and exposure to EPNs (Heterorhabditis bacteriophora): larvae with high (tgg) or low (Col‐0) levels of glucosinolates (GSLs) and GSL hydrolysis products, and control larvae (H₂O, not exposed to EPNs) and larvae inoculated with H. bacteriophora. Colors correspond to the most abundant bacterial genera, with only those representing ≥ 1% relative abundance in ≥ 10% of samples shown individually. All other taxa are grouped into the ‘Other’ category. The table below the plot indicates the number of dead larvae per replicate and the number of cadavers that developed red coloration, a marker for Photorhabdus activity. Note that Photorhabdus (dark green) is only detected in EPN‐exposed larvae.

Figure S4. The larval food plant affects the impact of Photorhabdus on the total bacterial load in Phyllotreta armoraciae larvae. Scatter plot showing the total bacterial DNA as a function of the relative abundance of Photorhabdus in Phyllotreta armoraciae larvae inoculated with Heterorhabditis bacteriophora. Data points are colored according to the larval food plant, which determines the levels of glucosinolates and glucosinolate hydrolysis products in larvae (Table S4). Solid lines represent the linear regression fit for each treatment group, and shaded areas correspond to the 95% confidence intervals. The significantly steeper slope for the tgg treatment indicates an interaction between larval food plant and Photorhabdus relative abundance in predicting the total bacterial load (Linear model, interaction term: N = 11‐12, F = 5.88, P = 0.025).

Table S1. Primers used in this study.

Table S2. FID response factors of glucosinolate hydrolysis products relative to the internal standard benzonitrile.

Table S3. Larval mortality, larval red coloration, and numbers of emerged entomopathogenic nematodes (EPNs) in PaMyr2‐silenced and control larvae inoculated with Heterorhabditis bacteriophora.

Table S4. Concentration of glucosinolates and glucosinolate hydrolysis products in Phyllotreta armoraciae larvae reared on Arabidopsis thaliana Col‐0 wild‐type and tgg1×tgg2 double‐knockout mutant plants.

PS-82-3603-s001.docx^{(457.8KB, docx)}

Data S2. Supporting Information.

PS-82-3603-s002.R^{(14.5KB, R)}

ACKNOWLEDGEMENTS

The authors thank Christelle A. M. Robert and Paul Himmighofen for providing nematodes for preliminary studies and support with setting up nematode assays, Simon C. ‘Niels’ Groen and Jessil Ann Pajar for fruitful discussions, Susanne Donnerhacke for excellent technical assistance, Domenica Schnabelrauch for sequencing service, and the MPI greenhouse team for plant cultivation. This work was funded by the Max Planck Society and by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – project number 501638262 (to FB) and EXC 2051 – project number 390713860 (to VB and MK). Open Access funding enabled and organized by Projekt DEAL.

DATA AVAILABILITY STATEMENT

The data that supports the findings of this study are available in the supplementary material of this article.

REFERENCES

1. Li Z, Costamagna AC, Beran F and You M, Biology, ecology, and management of flea beetles in Brassica crops. Annu Rev Entomol 69:199–217 (2024). [DOI] [PubMed] [Google Scholar]
2. Knodel JJ, Flea beetles (Phyllotreta spp.) and their management, in Integrated Management of Insect Pests on Canola and Other Brassica Oilseed Crops, ed. by Reddy GVP. CABI, Wallingford, Oxfordshire, UK, p. 394 (2017). [Google Scholar]
3. Lamb R, Entomology of oilseed Brassica crops. Annu Rev Entomol 34:211–229 (1989). [Google Scholar]
4. Mittapelly P, Guelly KN, Hussain A, Cárcamo HA, Soroka JJ, Vankosky MA et al., Flea beetle (Phyllotreta spp.) management in spring‐planted canola (Brassica napus L.) on the northern Great Plains of North America. GCB Bioenergy 16:e13178 (2024). [Google Scholar]
5. Tansey JA, Dosdall LM and Keddie BA, Phyllotreta cruciferae and Phyllotreta striolata responses to insecticidal seed treatments with different modes of action. J Appl Entomol 133:201–209 (2009). [Google Scholar]
6. Lamb RJ, Effects of flea beetles, Phyllotreta spp. (Chrysomelidae: Coleoptera), on the survival, growth, seed yield and quality of canola, rape and yellow mustard. Can Entomol 116:269–280 (1984). [Google Scholar]
7. Ramasamy S, Sotelo P, Lin MY, Heng CH, Kang S and Sarika S, Validation of a bio‐based integrated pest management package for the control of major insect pests on chinese mustard in Cambodia. Crop Prot 135:1–8 (2020). [Google Scholar]
8. Koppenhöfer AM, Shapiro‐Ilan DI and Hiltpold I, Entomopathogenic nematodes in sustainable food production. Front Sustainable Food Syst 4 :1–14 (2020). [Google Scholar]
9. Lacey LA and Georgis R, Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J Nematol 44:218–225 (2012). [PMC free article] [PubMed] [Google Scholar]
10. Stock SP and Hazir S, The bacterial symbionts of entomopathogenic nematodes and their role in symbiosis and pathogenesis. J Invertebr Pathol 211:108295 (2025). [DOI] [PubMed] [Google Scholar]
11. Hoarau C, Campbell H, Prince G, Chandler D and Pope T, Biological control agents against the cabbage stem flea beetle in oilseed rape crops. Biol Control 167:104844 (2022). [Google Scholar]
12. Godina G, Vandenbossche B, Schmidt M, Sender A, Tambe AH, Touceda‐González M et al., Entomopathogenic nematodes for biological control of Psylliodes chrysocephala (Coleoptera: Chrysomelidae) in oilseed rape. J Invertebr Pathol 197:107894 (2023). [DOI] [PubMed] [Google Scholar]
13. Noosidum A, Mangtab S and Lewis EE, Biological control potential of entomopathogenic nematodes against the striped flea beetle, Phyllotreta sinuata Stephens (Coleoptera: Chrysomelidae). Crop Prot 141:105448 (2021). [Google Scholar]
14. Trdan S, Vidrih M, Valič N and Laznik Ž, Impact of entomopathogenic nematodes on adults of Phyllotreta spp. (Coleoptera: Chrysomelidae) under laboratory conditions. Acta Agric Scand Sect B — Soil Plant Sci 58:169–175 (2008). [Google Scholar]
15. Xu C, De Clercq P, Moens M, Chen S and Han R, Efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) against the striped flea beetle, Phyllotreta striolata . BioControl 55:789–797 (2010). [Google Scholar]
16. Briar SS, Antwi F, Shrestha G, Sharma A and Reddy GVP, Potential biopesticides for crucifer flea beetle, Phyllotreta cruciferae (Coleoptera: Chrysomelidae) management under dryland canola production in Montana. Phytoparasitica 46:247–254 (2018). [Google Scholar]
17. Antwi FB and Reddy GVP, Efficacy of entomopathogenic nematodes and sprayable polymer gel against crucifer flea beetle (Coleoptera: Chrysomelidae) on canola. J Econ Entomol 109:1706–1712 (2016). [DOI] [PubMed] [Google Scholar]
18. Yan X, Han R, Moens M, Chen S and De Clercq P, Field evaluation of entomopathogenic nematodes for biological control of striped flea beetle, Phyllotreta striolata (Coleoptera: Chrysomelidae). BioControl 58:247–256 (2013). [Google Scholar]
19. Morris ON, Evaluation of the nematode, Steinernema feltiae Filipjev, for the control of the crucifer flea beetle, Phyllotreta cruciferae (Goeze) (Coleoptera: Chrysomelidae). Can Entomol 119:95–101 (1987). [Google Scholar]
20. Devi G, Influence of abiotic factors on efficacy of entomopathogenic nematodes. Int J Plant Soil Sci 36:283–290 (2024). [Google Scholar]
21. Machado RAR and von Reuss SH, Chemical ecology of nematodes. Chimia 76:945–953 (2022). [DOI] [PubMed] [Google Scholar]
22. Robert CAM, Zhang X, Machado RAR, Schirmer S, Lori M, Mateo P et al., Sequestration and activation of plant toxins protect the western corn rootworm from enemies at multiple trophic levels. Elife 6:17 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Achi P, Christensen P, Iglesias V, Mccarthy C, Pena R, Bavier L et al., Entomopathogenic nematode species vary in their behavior and virulence in response to cardiac glycosides within and around insect hosts. J Chem Ecol 51:12 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Gassmann AJ, Stock SP, Tabashnik BE and Singer MS, Tritrophic effects of host plants on an herbivore‐pathogen interaction. Ann Entomol Soc Am 103:371–378 (2010). [Google Scholar]
25. Hazir S, Shapiro‐Ilan DI, Hazir C, Leite LG, Cakmak I and Olson D, Multifaceted effects of host plants on entomopathogenic nematodes. J Invertebr Pathol 135:53–59 (2016). [DOI] [PubMed] [Google Scholar]
26. Opitz SEW and Müller C, Plant chemistry and insect sequestration. Chem 19:117–154 (2009). [Google Scholar]
27. Beran F and Petschenka G, Sequestration of plant defense compounds by insects: from mechanisms to insect plant coevolution. Annu Rev Entomol 67:163–180 (2022). [DOI] [PubMed] [Google Scholar]
28. Bruno P, Arce CCM, Machado RAR, Besomi G, Spescha A, Glauser G et al., Sequestration of cucurbitacins from cucumber plants by Diabrotica balteata larvae provides little protection against biological control agents. J Pest Sci 96:1061–1075 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Barbercheck ME and Wang J, Effect of cucurbitacin D on in vitro growth of Xenorhabdus and Photorhabdus spp., symbiotic bacteria of entomopathogenic nematodes. J Invertebr Pathol 68:141–145 (1996). [DOI] [PubMed] [Google Scholar]
30. Beran F, Pauchet Y, Kunert G, Reichelt M, Wielsch N, Vogel H et al., Phyllotreta striolata flea beetles use host plant defense compounds to create their own glucosinolate‐myrosinase system. Proc Natl Acad Sci U S A 111:7349–7354 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
31. Sporer T, Körnig J and Beran F, Ontogenetic differences in the chemical defence of flea beetles influence their predation risk. Funct Ecol 34:1370–1379 (2020). [Google Scholar]
32. Ntalli N and Caboni P, A review of isothiocyanates biofumigation activity on plant parasitic nematodes. Phytochem Rev 16:827–834 (2017). [Google Scholar]
33. Sarwar M, Kirkegaard JA, Wong PTW and Desmarchelier JM, Biofumigation potential of brassicas ‐ III. In vitro toxicity of isothiocyanates to soil‐borne fungal pathogens. Plant and Soil 201:103–112 (1998). [Google Scholar]
34. Wittstock U and Burow M, Glucosinolate breakdown in Arabidopsis: mechanism, regulation and biological significance, in The Arabidopsis Book, Vol. 8. The American Society of Plant Biologists, Maryland, USA, pp. 1–14 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Lamy E, Scholtes C, Herz C and Mersch‐Sundermann V, Pharmacokinetics and pharmacodynamics of isothiocyanates. Drug Metab Rev 43:387–407 (2011). [DOI] [PubMed] [Google Scholar]
36. Jeschke V, Gershenzon J and Vassão DG, A mode of action of glucosinolate‐derived isothiocyanates: detoxification depletes glutathione and cysteine levels with ramifications on protein metabolism in Spodoptera littoralis . Insect Biochem Mol Biol 71:37–48 (2016). [DOI] [PubMed] [Google Scholar]
37. Ramirez RA, Henderson DR, Riga E, Lacey LA and Snyder WE, Harmful effects of mustard bio‐fumigants on entomopathogenic nematodes. Biol Control 48:147–154 (2009). [Google Scholar]
38. Henderson DR, Riga E, Ramirez RA, Wilson J and Snyder E, Mustard biofumigation disrupts biological control by Steinernema spp. nematodes in the soil. Biol Control 48:316–322 (2009). [Google Scholar]
39. Körnig J, Ortizo K, Sporer T, Yang Z‐L and Beran F, Different myrosinases activate sequestered glucosinolates in larvae and adults of the horseradish flea beetle. Insect Biochem Mol Biol 163:104040 (2023). [DOI] [PubMed] [Google Scholar]
40. McMullen JG 2nd and Stock SP, In vivo and in vitro rearing of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae). J Vis Exp 91:52096 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
41. Weisburg WG, Barns SM, Pelletier DA and Lane DJ, 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173:697–703 (1991). [DOI] [PMC free article] [PubMed] [Google Scholar]
42. Barth C and Jander G, Arabidopsis myrosinases TGG1 and TGG2 have redundant function in glucosinolate breakdown and insect defense. Plant J 46:549–562 (2006). [DOI] [PubMed] [Google Scholar]
43. R Core Team , R: A Language and Environment for Statistical Computing. R foundation for statistical computing, Vienna, Austria: http://www.r-project.org/ (2023). [Google Scholar]
44. Bates D, Mächler M, Bolker B and Walker S, Fitting linear mixed‐effects models using lme4. J Stat Softw 67:1–48 (2015). [Google Scholar]
45. Hartig F, DHARMa: Residual diagnostics for hierarchical (multi‐level / mixed) regression models (2024).
46. Lenth RV, emmeans: Estimated marginal means, aka Least‐squares means (2025). .
47. Ritz C, Baty F, Streibig JC and Gerhard D, Dose‐response analysis using R. PLoS One 10:1–13 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
48. Pinheiro J, Bates D, DebRoy S, Sarkar D and Team RC , nlme: Linear and nonlinear mixed effects models. R package version 31–153 (2021). .
49. Zuur AF, Mixed Effects Models and Extensions in Ecology with R, Vol. xxii. Springer, New York, NY, p. 574 (2009). [Google Scholar]
50. Hothorn T, Bretz F and Westfall P, multcomp: Simultaneous inference in general parametric models. R package version 14–14 (2020). [DOI] [PubMed]
51. White GF, A method for obtaining infective nematode larvae from cultures. Science 66:302–303 (1927). [DOI] [PubMed] [Google Scholar]
52. Sporer T, Körnig J, Wielsch N, Gebauer‐Jung S, Reichelt M, Hupfer Y et al., Hijacking the mustard‐oil bomb: how a glucosinolate‐sequestering flea beetle copes with plant myrosinases. Front Plant Sci 12:645030 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
53. Liu CM, Aziz M, Kachur S, Hsueh P‐R, Huang Y‐T, Keim P et al., BactQuant: an enhanced broad‐coverage bacterial quantitative real‐time PCR assay. BMC Microbiol 12:56 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
54. Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA and Holmes SP, DADA2: high‐resolution sample inference from Illumina amplicon data. Nat Methods 13:581–583 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
55. McMurdie PJ and Holmes S, Phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS One 8:e61217 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
56. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P et al., The SILVA ribosomal RNA gene database project: improved data processing and web‐based tools. Nucleic Acids Res 41:D590–D596 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]
57. Yilmaz P, Parfrey LW, Yarza P, Gerken J, Pruesse E, Quast C et al., The SILVA and All‐species Living Tree Project (LTP) taxonomic frameworks. Nucleic Acids Res 42:D643–D648 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
58. Bode HB, Entomopathogenic bacteria as a source of secondary metabolites. Curr Opin Chem Biol 13:224–230 (2009). [DOI] [PubMed] [Google Scholar]
59. Muangpat P, Suwannaroj M, Yimthin T, Fukruksa C, Sitthisak S, Chantratita N et al., Antibacterial activity of Xenorhabdus and Photorhabdus isolated from entomopathogenic nematodes against antibiotic‐resistant bacteria. PLoS One 15:e0234129 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
60. Cimen H, Touray M, Gulsen SH and Hazir S, Natural products from Photorhabdus and Xenorhabdus: mechanisms and impacts. Appl Microbiol Biotechnol 106:4387–4399 (2022). [DOI] [PubMed] [Google Scholar]
61. Sun R, Hong B, Reichelt M, Luck K, Mai DT, Jiang X et al., Metabolism of plant‐derived toxins from its insect host increases the success of the entomopathogenic fungus Beauveria bassiana . ISME J 17:1693–1704 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
62. Dufour V, Stahl M and Baysse C, The antibacterial properties of isothiocyanates. Microbiology 161:229–243 (2015). [DOI] [PubMed] [Google Scholar]
63. Aires A, Mota VR, Saavedra MJ, Monteiro AA, Simões M, Rosa EAS et al., Initial in vitro evaluations of the antibacterial activities of glucosinolate enzymatic hydrolysis products against plant pathogenic bacteria. J Appl Microbiol 106:2096–2105 (2009). [DOI] [PubMed] [Google Scholar]
64. Shakour ZT, Shehab NG, Gomaa AS, Wessjohann LA and Farag MA, Metabolic and biotransformation effects on dietary glucosinolates, their bioavailability, catabolism and biological effects in different organisms. Biotechnol Adv 54:107784 (2022). [DOI] [PubMed] [Google Scholar]
65. Jeschke V, Gershenzon J and Vassão DG, Insect detoxification of glucosinolates and their hydrolysis products, in Advances in Botanical Research: Glucosinolates. Glucosinolates, Vol. 80, ed. by Jacquot J‐P and Gadal P. Elsevier, UK, pp. 199–245 (2016). [Google Scholar]
66. Lambrix V, Reichelt M, Mitchell‐Olds T, Kliebenstein DJ and Gershenzon J, The Arabidopsis epithiospecifier protein promotes the hydrolysis of glucosinolates to nitriles and influences Trichoplusia ni herbivory. Plant Cell 13:2793–2807 (2001). [DOI] [PMC free article] [PubMed] [Google Scholar]
67. Mumm R, Burow M, Bukovinszkine'Kiss G, Kazantzidou E, Wittstock U, Dicke M et al., Formation of simple nitriles upon glucosinolate hydrolysis affects direct and indirect defense against the specialist herbivore, Pieris rapae . J Chem Ecol 34:1311–1321 (2008). [DOI] [PubMed] [Google Scholar]
68. Wittstock U, Kliebenstein DJ, Lambrix V, Reichelt M et al., Chapter five glucosinolate hydrolysis and its impact on generalist and specialist insect herbivores, in Recent Advances in Phytochemistry, Vol. 37, ed. by Romeo JT. Elsevier, Amsterdam, pp. 101–125 (2003). [Google Scholar]
69. Wittstock U, Kurzbach E, Herfurh A‐M and Stauber EJ, Glucosinolate breakdown, in Advances in Botanical Research: Glucosinolates, Vol. 80. Elsevier, UK, pp. 125–169 (2016). [Google Scholar]
70. Lazzeri L, Curto G, Leoni O and Dallavalle E, Effects of glucosinolates and their enzymatic hydrolysis products via myrosinase on the root‐knot nematode Meloidogyne incognita (Kofoid et White) Chitw. J Agric Food Chem 52:6703–6707 (2004). [DOI] [PubMed] [Google Scholar]
71. Dahlin P and Hallmann J, New insights on the role of allyl isothiocyanate in controlling the root knot nematode Meloidogyne hapla . Plants‐Basel 9 :1–13 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
72. Lazzeri L, Tacconi R and Palmieri S, In vitro activity of some glucosinolates and their reaction products toward a population of the nematode Heterodera schachtii . J Agric Food Chem 41:825–829 (1993). [Google Scholar]
73. Schroeder NE and Macguidwin AE, Mortality and behavior in Heterodera glycines juveniles following exposure to isothiocyanate compounds. J Nematol. 42:194–200 (2010). [PMC free article] [PubMed] [Google Scholar]
74. Borges A, Abreu AC, Ferreira C, Saavedra MJ, Simões LC and Simões M, Antibacterial activity and mode of action of selected glucosinolate hydrolysis products against bacterial pathogens. J Food Sci Technol 52:4737–4748 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
75. Dufour V, Alazzam B, Ermel G, Thepaut M, Rossero A, Tresse O et al., Antimicrobial activities of isothiocyanates against Campylobacter jejuni isolates. Front Cell Infect Microbiol 2:53 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
76. Cimen H, The role of Photorhabdus‐induced bioluminescence and red cadaver coloration on the deterrence of insect scavengers from entomopathogenic nematode‐infected cadavers. J Invertebr Pathol 196:107871 (2023). [DOI] [PubMed] [Google Scholar]
77. Cassells MD, Labaude S and Griffin CT, Beware glowing cadavers: bioluminescence of nematode symbiont Photorhabdus protects nematode‐infected host cadavers from nocturnal scavengers. Front Ecol Evol 11 :1264251 (2024). [Google Scholar]
78. Zavilgelsky GB and Shakulov RS, Mechanisms and origin of bacterial biolumenescence. Mol Biol 52:812–822 (2018). [DOI] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Method S1. Quantification of GLS hydrolysis products in EPN‐infected and uninfected larvae and adults.

Method S2. Stability of GSL hydrolysis products in water

Method S3. Identification and quantification of GSL hydrolysis products in larvae fed with Arabidopsis

Method S4. 16S Amplicon sequencing of bacterial communities associated with Phyllotreta armoraciae larvae.

Table S1. Primers used in this study.

Table S2. FID response factors of glucosinolate hydrolysis products relative to the internal standard benzonitrile.

PS-82-3603-s001.docx^{(457.8KB, docx)}

Data S2. Supporting Information.

PS-82-3603-s002.R^{(14.5KB, R)}

Data Availability Statement

The data that supports the findings of this study are available in the supplementary material of this article.

[ps70482-bib-0001] 1. Li Z, Costamagna AC, Beran F and You M, Biology, ecology, and management of flea beetles in Brassica crops. Annu Rev Entomol 69:199–217 (2024). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0002] 2. Knodel JJ, Flea beetles (Phyllotreta spp.) and their management, in Integrated Management of Insect Pests on Canola and Other Brassica Oilseed Crops, ed. by Reddy GVP. CABI, Wallingford, Oxfordshire, UK, p. 394 (2017). [Google Scholar]

[ps70482-bib-0003] 3. Lamb R, Entomology of oilseed Brassica crops. Annu Rev Entomol 34:211–229 (1989). [Google Scholar]

[ps70482-bib-0004] 4. Mittapelly P, Guelly KN, Hussain A, Cárcamo HA, Soroka JJ, Vankosky MA et al., Flea beetle (Phyllotreta spp.) management in spring‐planted canola (Brassica napus L.) on the northern Great Plains of North America. GCB Bioenergy 16:e13178 (2024). [Google Scholar]

[ps70482-bib-0005] 5. Tansey JA, Dosdall LM and Keddie BA, Phyllotreta cruciferae and Phyllotreta striolata responses to insecticidal seed treatments with different modes of action. J Appl Entomol 133:201–209 (2009). [Google Scholar]

[ps70482-bib-0006] 6. Lamb RJ, Effects of flea beetles, Phyllotreta spp. (Chrysomelidae: Coleoptera), on the survival, growth, seed yield and quality of canola, rape and yellow mustard. Can Entomol 116:269–280 (1984). [Google Scholar]

[ps70482-bib-0007] 7. Ramasamy S, Sotelo P, Lin MY, Heng CH, Kang S and Sarika S, Validation of a bio‐based integrated pest management package for the control of major insect pests on chinese mustard in Cambodia. Crop Prot 135:1–8 (2020). [Google Scholar]

[ps70482-bib-0008] 8. Koppenhöfer AM, Shapiro‐Ilan DI and Hiltpold I, Entomopathogenic nematodes in sustainable food production. Front Sustainable Food Syst 4 :1–14 (2020). [Google Scholar]

[ps70482-bib-0009] 9. Lacey LA and Georgis R, Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J Nematol 44:218–225 (2012). [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0010] 10. Stock SP and Hazir S, The bacterial symbionts of entomopathogenic nematodes and their role in symbiosis and pathogenesis. J Invertebr Pathol 211:108295 (2025). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0011] 11. Hoarau C, Campbell H, Prince G, Chandler D and Pope T, Biological control agents against the cabbage stem flea beetle in oilseed rape crops. Biol Control 167:104844 (2022). [Google Scholar]

[ps70482-bib-0012] 12. Godina G, Vandenbossche B, Schmidt M, Sender A, Tambe AH, Touceda‐González M et al., Entomopathogenic nematodes for biological control of Psylliodes chrysocephala (Coleoptera: Chrysomelidae) in oilseed rape. J Invertebr Pathol 197:107894 (2023). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0013] 13. Noosidum A, Mangtab S and Lewis EE, Biological control potential of entomopathogenic nematodes against the striped flea beetle, Phyllotreta sinuata Stephens (Coleoptera: Chrysomelidae). Crop Prot 141:105448 (2021). [Google Scholar]

[ps70482-bib-0014] 14. Trdan S, Vidrih M, Valič N and Laznik Ž, Impact of entomopathogenic nematodes on adults of Phyllotreta spp. (Coleoptera: Chrysomelidae) under laboratory conditions. Acta Agric Scand Sect B — Soil Plant Sci 58:169–175 (2008). [Google Scholar]

[ps70482-bib-0015] 15. Xu C, De Clercq P, Moens M, Chen S and Han R, Efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) against the striped flea beetle, Phyllotreta striolata . BioControl 55:789–797 (2010). [Google Scholar]

[ps70482-bib-0016] 16. Briar SS, Antwi F, Shrestha G, Sharma A and Reddy GVP, Potential biopesticides for crucifer flea beetle, Phyllotreta cruciferae (Coleoptera: Chrysomelidae) management under dryland canola production in Montana. Phytoparasitica 46:247–254 (2018). [Google Scholar]

[ps70482-bib-0017] 17. Antwi FB and Reddy GVP, Efficacy of entomopathogenic nematodes and sprayable polymer gel against crucifer flea beetle (Coleoptera: Chrysomelidae) on canola. J Econ Entomol 109:1706–1712 (2016). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0018] 18. Yan X, Han R, Moens M, Chen S and De Clercq P, Field evaluation of entomopathogenic nematodes for biological control of striped flea beetle, Phyllotreta striolata (Coleoptera: Chrysomelidae). BioControl 58:247–256 (2013). [Google Scholar]

[ps70482-bib-0019] 19. Morris ON, Evaluation of the nematode, Steinernema feltiae Filipjev, for the control of the crucifer flea beetle, Phyllotreta cruciferae (Goeze) (Coleoptera: Chrysomelidae). Can Entomol 119:95–101 (1987). [Google Scholar]

[ps70482-bib-0020] 20. Devi G, Influence of abiotic factors on efficacy of entomopathogenic nematodes. Int J Plant Soil Sci 36:283–290 (2024). [Google Scholar]

[ps70482-bib-0021] 21. Machado RAR and von Reuss SH, Chemical ecology of nematodes. Chimia 76:945–953 (2022). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0022] 22. Robert CAM, Zhang X, Machado RAR, Schirmer S, Lori M, Mateo P et al., Sequestration and activation of plant toxins protect the western corn rootworm from enemies at multiple trophic levels. Elife 6:17 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0023] 23. Achi P, Christensen P, Iglesias V, Mccarthy C, Pena R, Bavier L et al., Entomopathogenic nematode species vary in their behavior and virulence in response to cardiac glycosides within and around insect hosts. J Chem Ecol 51:12 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0024] 24. Gassmann AJ, Stock SP, Tabashnik BE and Singer MS, Tritrophic effects of host plants on an herbivore‐pathogen interaction. Ann Entomol Soc Am 103:371–378 (2010). [Google Scholar]

[ps70482-bib-0025] 25. Hazir S, Shapiro‐Ilan DI, Hazir C, Leite LG, Cakmak I and Olson D, Multifaceted effects of host plants on entomopathogenic nematodes. J Invertebr Pathol 135:53–59 (2016). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0026] 26. Opitz SEW and Müller C, Plant chemistry and insect sequestration. Chem 19:117–154 (2009). [Google Scholar]

[ps70482-bib-0027] 27. Beran F and Petschenka G, Sequestration of plant defense compounds by insects: from mechanisms to insect plant coevolution. Annu Rev Entomol 67:163–180 (2022). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0028] 28. Bruno P, Arce CCM, Machado RAR, Besomi G, Spescha A, Glauser G et al., Sequestration of cucurbitacins from cucumber plants by Diabrotica balteata larvae provides little protection against biological control agents. J Pest Sci 96:1061–1075 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0029] 29. Barbercheck ME and Wang J, Effect of cucurbitacin D on in vitro growth of Xenorhabdus and Photorhabdus spp., symbiotic bacteria of entomopathogenic nematodes. J Invertebr Pathol 68:141–145 (1996). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0030] 30. Beran F, Pauchet Y, Kunert G, Reichelt M, Wielsch N, Vogel H et al., Phyllotreta striolata flea beetles use host plant defense compounds to create their own glucosinolate‐myrosinase system. Proc Natl Acad Sci U S A 111:7349–7354 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0031] 31. Sporer T, Körnig J and Beran F, Ontogenetic differences in the chemical defence of flea beetles influence their predation risk. Funct Ecol 34:1370–1379 (2020). [Google Scholar]

[ps70482-bib-0032] 32. Ntalli N and Caboni P, A review of isothiocyanates biofumigation activity on plant parasitic nematodes. Phytochem Rev 16:827–834 (2017). [Google Scholar]

[ps70482-bib-0033] 33. Sarwar M, Kirkegaard JA, Wong PTW and Desmarchelier JM, Biofumigation potential of brassicas ‐ III. In vitro toxicity of isothiocyanates to soil‐borne fungal pathogens. Plant and Soil 201:103–112 (1998). [Google Scholar]

[ps70482-bib-0034] 34. Wittstock U and Burow M, Glucosinolate breakdown in Arabidopsis: mechanism, regulation and biological significance, in The Arabidopsis Book, Vol. 8. The American Society of Plant Biologists, Maryland, USA, pp. 1–14 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0035] 35. Lamy E, Scholtes C, Herz C and Mersch‐Sundermann V, Pharmacokinetics and pharmacodynamics of isothiocyanates. Drug Metab Rev 43:387–407 (2011). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0036] 36. Jeschke V, Gershenzon J and Vassão DG, A mode of action of glucosinolate‐derived isothiocyanates: detoxification depletes glutathione and cysteine levels with ramifications on protein metabolism in Spodoptera littoralis . Insect Biochem Mol Biol 71:37–48 (2016). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0037] 37. Ramirez RA, Henderson DR, Riga E, Lacey LA and Snyder WE, Harmful effects of mustard bio‐fumigants on entomopathogenic nematodes. Biol Control 48:147–154 (2009). [Google Scholar]

[ps70482-bib-0038] 38. Henderson DR, Riga E, Ramirez RA, Wilson J and Snyder E, Mustard biofumigation disrupts biological control by Steinernema spp. nematodes in the soil. Biol Control 48:316–322 (2009). [Google Scholar]

[ps70482-bib-0039] 39. Körnig J, Ortizo K, Sporer T, Yang Z‐L and Beran F, Different myrosinases activate sequestered glucosinolates in larvae and adults of the horseradish flea beetle. Insect Biochem Mol Biol 163:104040 (2023). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0040] 40. McMullen JG 2nd and Stock SP, In vivo and in vitro rearing of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae). J Vis Exp 91:52096 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0041] 41. Weisburg WG, Barns SM, Pelletier DA and Lane DJ, 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173:697–703 (1991). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0042] 42. Barth C and Jander G, Arabidopsis myrosinases TGG1 and TGG2 have redundant function in glucosinolate breakdown and insect defense. Plant J 46:549–562 (2006). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0043] 43. R Core Team , R: A Language and Environment for Statistical Computing. R foundation for statistical computing, Vienna, Austria: http://www.r-project.org/ (2023). [Google Scholar]

[ps70482-bib-0044] 44. Bates D, Mächler M, Bolker B and Walker S, Fitting linear mixed‐effects models using lme4. J Stat Softw 67:1–48 (2015). [Google Scholar]

[ps70482-bib-0045] 45. Hartig F, DHARMa: Residual diagnostics for hierarchical (multi‐level / mixed) regression models (2024).

[ps70482-bib-0046] 46. Lenth RV, emmeans: Estimated marginal means, aka Least‐squares means (2025). .

[ps70482-bib-0047] 47. Ritz C, Baty F, Streibig JC and Gerhard D, Dose‐response analysis using R. PLoS One 10:1–13 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0048] 48. Pinheiro J, Bates D, DebRoy S, Sarkar D and Team RC , nlme: Linear and nonlinear mixed effects models. R package version 31–153 (2021). .

[ps70482-bib-0049] 49. Zuur AF, Mixed Effects Models and Extensions in Ecology with R, Vol. xxii. Springer, New York, NY, p. 574 (2009). [Google Scholar]

[ps70482-bib-0050] 50. Hothorn T, Bretz F and Westfall P, multcomp: Simultaneous inference in general parametric models. R package version 14–14 (2020). [DOI] [PubMed]

[ps70482-bib-0051] 51. White GF, A method for obtaining infective nematode larvae from cultures. Science 66:302–303 (1927). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0052] 52. Sporer T, Körnig J, Wielsch N, Gebauer‐Jung S, Reichelt M, Hupfer Y et al., Hijacking the mustard‐oil bomb: how a glucosinolate‐sequestering flea beetle copes with plant myrosinases. Front Plant Sci 12:645030 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0053] 53. Liu CM, Aziz M, Kachur S, Hsueh P‐R, Huang Y‐T, Keim P et al., BactQuant: an enhanced broad‐coverage bacterial quantitative real‐time PCR assay. BMC Microbiol 12:56 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0054] 54. Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA and Holmes SP, DADA2: high‐resolution sample inference from Illumina amplicon data. Nat Methods 13:581–583 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0055] 55. McMurdie PJ and Holmes S, Phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS One 8:e61217 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0056] 56. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P et al., The SILVA ribosomal RNA gene database project: improved data processing and web‐based tools. Nucleic Acids Res 41:D590–D596 (2013). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0057] 57. Yilmaz P, Parfrey LW, Yarza P, Gerken J, Pruesse E, Quast C et al., The SILVA and All‐species Living Tree Project (LTP) taxonomic frameworks. Nucleic Acids Res 42:D643–D648 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0058] 58. Bode HB, Entomopathogenic bacteria as a source of secondary metabolites. Curr Opin Chem Biol 13:224–230 (2009). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0059] 59. Muangpat P, Suwannaroj M, Yimthin T, Fukruksa C, Sitthisak S, Chantratita N et al., Antibacterial activity of Xenorhabdus and Photorhabdus isolated from entomopathogenic nematodes against antibiotic‐resistant bacteria. PLoS One 15:e0234129 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0060] 60. Cimen H, Touray M, Gulsen SH and Hazir S, Natural products from Photorhabdus and Xenorhabdus: mechanisms and impacts. Appl Microbiol Biotechnol 106:4387–4399 (2022). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0061] 61. Sun R, Hong B, Reichelt M, Luck K, Mai DT, Jiang X et al., Metabolism of plant‐derived toxins from its insect host increases the success of the entomopathogenic fungus Beauveria bassiana . ISME J 17:1693–1704 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0062] 62. Dufour V, Stahl M and Baysse C, The antibacterial properties of isothiocyanates. Microbiology 161:229–243 (2015). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0063] 63. Aires A, Mota VR, Saavedra MJ, Monteiro AA, Simões M, Rosa EAS et al., Initial in vitro evaluations of the antibacterial activities of glucosinolate enzymatic hydrolysis products against plant pathogenic bacteria. J Appl Microbiol 106:2096–2105 (2009). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0064] 64. Shakour ZT, Shehab NG, Gomaa AS, Wessjohann LA and Farag MA, Metabolic and biotransformation effects on dietary glucosinolates, their bioavailability, catabolism and biological effects in different organisms. Biotechnol Adv 54:107784 (2022). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0065] 65. Jeschke V, Gershenzon J and Vassão DG, Insect detoxification of glucosinolates and their hydrolysis products, in Advances in Botanical Research: Glucosinolates. Glucosinolates, Vol. 80, ed. by Jacquot J‐P and Gadal P. Elsevier, UK, pp. 199–245 (2016). [Google Scholar]

[ps70482-bib-0066] 66. Lambrix V, Reichelt M, Mitchell‐Olds T, Kliebenstein DJ and Gershenzon J, The Arabidopsis epithiospecifier protein promotes the hydrolysis of glucosinolates to nitriles and influences Trichoplusia ni herbivory. Plant Cell 13:2793–2807 (2001). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0067] 67. Mumm R, Burow M, Bukovinszkine'Kiss G, Kazantzidou E, Wittstock U, Dicke M et al., Formation of simple nitriles upon glucosinolate hydrolysis affects direct and indirect defense against the specialist herbivore, Pieris rapae . J Chem Ecol 34:1311–1321 (2008). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0068] 68. Wittstock U, Kliebenstein DJ, Lambrix V, Reichelt M et al., Chapter five glucosinolate hydrolysis and its impact on generalist and specialist insect herbivores, in Recent Advances in Phytochemistry, Vol. 37, ed. by Romeo JT. Elsevier, Amsterdam, pp. 101–125 (2003). [Google Scholar]

[ps70482-bib-0069] 69. Wittstock U, Kurzbach E, Herfurh A‐M and Stauber EJ, Glucosinolate breakdown, in Advances in Botanical Research: Glucosinolates, Vol. 80. Elsevier, UK, pp. 125–169 (2016). [Google Scholar]

[ps70482-bib-0070] 70. Lazzeri L, Curto G, Leoni O and Dallavalle E, Effects of glucosinolates and their enzymatic hydrolysis products via myrosinase on the root‐knot nematode Meloidogyne incognita (Kofoid et White) Chitw. J Agric Food Chem 52:6703–6707 (2004). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0071] 71. Dahlin P and Hallmann J, New insights on the role of allyl isothiocyanate in controlling the root knot nematode Meloidogyne hapla . Plants‐Basel 9 :1–13 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0072] 72. Lazzeri L, Tacconi R and Palmieri S, In vitro activity of some glucosinolates and their reaction products toward a population of the nematode Heterodera schachtii . J Agric Food Chem 41:825–829 (1993). [Google Scholar]

[ps70482-bib-0073] 73. Schroeder NE and Macguidwin AE, Mortality and behavior in Heterodera glycines juveniles following exposure to isothiocyanate compounds. J Nematol. 42:194–200 (2010). [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0074] 74. Borges A, Abreu AC, Ferreira C, Saavedra MJ, Simões LC and Simões M, Antibacterial activity and mode of action of selected glucosinolate hydrolysis products against bacterial pathogens. J Food Sci Technol 52:4737–4748 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0075] 75. Dufour V, Alazzam B, Ermel G, Thepaut M, Rossero A, Tresse O et al., Antimicrobial activities of isothiocyanates against Campylobacter jejuni isolates. Front Cell Infect Microbiol 2:53 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]

[ps70482-bib-0076] 76. Cimen H, The role of Photorhabdus‐induced bioluminescence and red cadaver coloration on the deterrence of insect scavengers from entomopathogenic nematode‐infected cadavers. J Invertebr Pathol 196:107871 (2023). [DOI] [PubMed] [Google Scholar]

[ps70482-bib-0077] 77. Cassells MD, Labaude S and Griffin CT, Beware glowing cadavers: bioluminescence of nematode symbiont Photorhabdus protects nematode‐infected host cadavers from nocturnal scavengers. Front Ecol Evol 11 :1264251 (2024). [Google Scholar]

[ps70482-bib-0078] 78. Zavilgelsky GB and Shakulov RS, Mechanisms and origin of bacterial biolumenescence. Mol Biol 52:812–822 (2018). [DOI] [PubMed] [Google Scholar]

PERMALINK

Glucosinolate hydrolysis products suppress entomopathogenic nematodes in vitro but do not protect sequestering flea beetle larvae in vivo

Johannes Körnig

Vojtech Beneš

Christin Manthey

Michael Reichelt

Grit Kunert

Christian Paetz

Johanna Kutzschbach

Paula Lampe

Martin Kaltenpoth

Franziska Beran

Abstract

BACKGROUND

RESULTS

CONCLUSION

1. INTRODUCTION

2. MATERIALS AND METHODS

2.1. Biological resources

2.2. Quantification of GLS hydrolysis products in EPN‐infected and uninfected larvae and adults

2.3. Susceptibility of EPNs to GSL hydrolysis products

2.4. Susceptibility of bacterial symbionts to GSL hydrolysis products

2.5. Impact of Phyllotreta armoraciae larval defense on susceptibility to EPNs

2.6. Impact of the chemical defense of Phyllotreta armoraciae larvae on bacterial symbionts

3. RESULTS

3.1. GSL hydrolysis products are present in Phyllotreta armoraciae larvae but not adults

Figure 1.

3.2. EPNs and their bacterial symbionts are susceptible to GSL hydrolysis products

Figure 2.

3.3. GSL hydrolysis products do not protect Phyllotreta armoraciae larvae against EPNs

Figure 3.

3.4. The larval food plant influences the relative abundance of bacterial symbionts

Figure 4.

4. DISCUSSION

CONFLICT OF INTEREST

Supporting information

ACKNOWLEDGEMENTS

DATA AVAILABILITY STATEMENT

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

Similar articles

Cited by other articles

Links to NCBI Databases