Skip to main content
NPJ Biofilms and Microbiomes logoLink to NPJ Biofilms and Microbiomes
. 2026 Jan 13;12:48. doi: 10.1038/s41522-026-00914-y

From leaves to aphid honeydew: the zucchini plants enrich bacterium to recruit natural enemy to resist herbivore attacks

Yue Liu 1,2,#, Jing Sun 1,#, Boya Jiao 3, Shichang Zhang 1, Yu Peng 2,, Yao Zhao 1,
PMCID: PMC12905166  PMID: 41530187

Abstract

Plants have evolved intricate defense strategies to combat herbivorous insect attacks, including the production of toxic secondary metabolites and the attraction of natural enemies. While bacterial-mediated direct toxicity in plant defenses has been demonstrated, the ecological mechanisms by which plants utilize bacteria to indirectly modulate the behavior of natural enemies remain insufficiently explored. In this study, we observed a significant enrichment of Staphylococcus in the tissues of zucchini Cucurbita pepo following infestation by the cotton-melon aphid Aphis gossypii. These bacteria traced from the damaged plant stems and leaves were subsequently found both in aphids and in their secreted honeydew. Among the four dominant bacterial strains isolated from the honeydew, Staphylococcus sp. markedly promoted oviposition preference in mated female ladybird beetles Propylea japonica. Further investigation identified a volatile organic compound, 4-Isopropylbenzyl alcohol, released by Staphylococcus sp. that stimulated strong antennal responses and attracted P. japonica to lay eggs. Collectively, our findings demonstrate that zucchini plants can employ indirect defense against A. gossypii through the enrichment of specific bacteria, revealing a novel ecological role for bacteria in plant defense and expanding our understanding of complex plant–microbe–insect–natural enemy interactions.

graphic file with name 41522_2026_914_Figa_HTML.jpg

Subject terms: Ecology, Ecology, Microbiology, Plant sciences

Introduction

In natural ecosystems, herbivorous insects and plants have long been engaged in a co-evolutionary “arms race.” Insects rely on plants as their primary food source, and their feeding behavior causes continuous damage to plant tissues, driving the evolution of multilayered plant defense strategies. Direct plant defenses include physical barriers such as trichomes and thickened cuticles, as well as a suite of chemical compounds, including toxic secondary metabolites and defensive proteins that can effectively interfere with insect feeding behavior and physiological metabolism13. In addition, plants can release volatile organic compounds (VOCs) to recruit predatory or parasitic natural enemies, thereby establishing an indirect defense mechanism47. These VOCs, including green leaf volatiles, terpenoids, and phenolic compounds, are typically released from damaged tissues or induced by herbivore-associated cues4,5. Beyond mediating tritrophic interactions, they can also influence herbivore host selection and oviposition behavior, making them a pivotal component of plant indirect defense strategies6,7.

Recent studies have increasingly demonstrated that plants can enhance their resistance to herbivores by modulating associated microbial communities. For example, a significant enrichment of Pseudomonas was reported in the leaves, roots, and rhizosphere soil of willow Salix babylonica infested by the leaf beetle Plagiodera versicolora, and this enrichment exhibited direct toxicity against the beetle8. Similarly, rhizobacteria (Acidovorax radicis or Bacillus subtilis) could prime the activation of plant defense and nutritional responses to suppress aphid populations on barley9. In addition, the bean bug Riptortus pedestris acquires its gut microbiota from soil and plants to enhance host reproductive performance10. However, the role of microbial interactions in mediating plant–insect relationships remains poorly understood.

In the intricate network of plant–microbe–insect interactions, insects are not merely passive herbivores or simple initiators of plant defense1113. Rather, they maintain intimate and complex symbiotic relationships with diverse microbial communities. These symbionts play vital roles in insect nutrition, immune modulation, behavioral regulation, and environmental adaptation1418. The origins of insect-associated microbes are highly diverse, generally acquired through vertical and horizontal transmission. Vertical transmission ensures the persistence of symbionts across generations via maternal secretions, reproductive tissues, or egg surfaces19. Horizontal transmission, in contrast, occurs through feeding, contact with plant surfaces, or interspecific interactions, allowing insects to acquire novel microbes from the environment. Studies have shown that the composition of gut microbiota in herbivorous insects is influenced by a variety of factors, including host plant species, habitat, and the characteristics of surrounding microbial communities10,13,2022.

The zucchini Cucurbita pepo is a widely cultivated cucurbit crop of high global agricultural value, prized for its rich nutritional profile and potential medicinal benefits23,24. Among its major insect pests, the cotton-melon aphid Aphis gossypii poses a significant threat due to its rapid reproduction, strong environmental adaptability, and increasing resistance to chemical insecticides2528. In addition, the honeydew excreted by this aphid promotes the development of sooty mold on plant surfaces, exacerbating crop damage and reducing overall plant vigor29. Earlier studies primarily regarded honeydew as a nutritional resource for certain insects, such as ants and parasitic wasps30,31. However, more recent research has highlighted its important ecological function in attracting natural enemies32,33. Honeydew is rich in fermentable sugars like glucose, fructose, and sucrose, and provides a suitable environment for the growth of various bacteria33. The bacteria that colonize the honeydew can metabolize the sugars, releasing a range of VOCs32. Research has shown that the volatile compounds emitted by these bacteria serve as strong chemical signals, detectable by predatory or parasitic enemies, thereby guiding them to locate and target their hosts. For instance, Staphylococcus sciuri found in the honeydew of aphid Acyrthosiphon pisum acts as an allelochemical, guiding predators to the prey and influencing the oviposition preference of natural enemies34. Volatiles produced by bacteria in the honeydew of aphids, A. gossypii mediate prey location by the predator Hippodamia variegate35. In addition, several other honeydew-associated bacteria have also been reported to participate in the recruitment of natural enemies3639. Although the involvement of honeydew-associated bacteria in pest-natural enemy interactions has been reported, their origins and transmission pathways remain largely unexplored.

In our previous study, we identified several bacteria derived from the honeydew of aphid A. gossypii, whose volatile metabolites significantly induce oviposition behavior in ladybird beetle Propylea japonica40. However, the origin of the exact sources of these functional bacteria is still unknown. It is uncertain whether these functional bacterial strains originate from the aphid’s symbiotic microbiota or are acquired from the surrounding environment during feeding. Therefore, the main aim of this study is to investigate the sources, transmission pattern, and ecological functions of the microbiota present in aphid honeydew. Our study may contribute to the development and application of environmentally friendly pest control strategies based on behavioral manipulation of the pest's natural enemy.

Results

Microbial community composition and its variations across different treatments

The bacterial community composition of healthy and aphid-treated rhizosphere soil, zucchini roots, stems, leaves, aphids, and honeydew was analyzed using full-length 16S rRNA amplicon sequencing. The α-diversity of bacterial communities across different treatments was assessed using the Ace and Shannon indices (Fig. 1A, B). The soil microbiome exhibited the highest richness and diversity, followed by those of the roots, stems, and leaves (Fig. 1A, B). Notably, the microbial community in healthy stems showed significantly higher Ace and Shannon diversity indices compared to stems infested by aphids (P < 0.05). No significant difference was observed in the Ace index between honeydew and A10 individuals. However, the Shannon diversity index was significantly higher in honeydew samples (P < 0.05), suggesting greater species diversity within the honeydew-associated microbiota. PCoA analysis based on Bray-Curtis dissimilarity was performed to compare the differences in bacterial community composition across samples. The results revealed that bacterial communities from healthy and aphid-infested stems and leaves exhibited partial overlap but also clear separation (Fig. 1C), suggesting that aphid feeding altered the microbial community structure of zucchini stems and leaves. Further analysis showed that the microbial communities of infested stems and leaves were more closely clustered with those of A10 aphids and their secreted honeydew (Fig. 1C). In contrast, rhizosphere soil and root samples, both before and after aphid infestation, clustered separately, indicating that the communities associated with aphids and honeydew were more similar to those of infested stems and leaves, but markedly distinct from those of the rhizosphere soil and roots (Fig. 1C).

Fig. 1. α and β diversity of bacterial communities across different sample groups.

Fig. 1

A ACE richness index and B Shannon diversity. Different lowercase letters indicate significant differences among groups (Kruskal–Wallis test, n = 5, P < 0.05, followed by Tukey’s test). C Principal coordinates analysis (PCoA) plot based on Bray-Curtis dissimilarity, visualizing the bacterial community structure (n = 5). Different colors represent different groups. Statistical significance was assessed using ANOSIM (P = 0.001). HS, HR, Hst, and HL represent healthy soil, roots, stems, and leaves, respectively. DS, DR, Dst, and DL denote the corresponding aphid-infested samples. A0 refers to newly hatched nymphs (<24 h old), A10 to nymphs after 10 days of feeding, and HD to honeydew.

Microbial community analyses across the eleven sample groups revealed pronounced ecological niche differentiation at both the phylum and genus levels (Fig. 2A, B). At the phylum level, Pseudomonadota dominated the bacterial communities in soil, roots, healthy stems and leaves of zucchini, as well as in A0 aphids and honeydew (Fig. 2A). However, infestation by aphids markedly altered the microbial composition of zucchini stems and leaves, leading to a significant increase in the relative abundance of Bacillota, which subsequently replaced Pseudomonadota as the dominant phylum (Fig. 2A). Notably, the internal microbiota of A10 aphids was similarly dominated by Bacillota. At the genus level, the top ten most abundant genera across all samples were identified, ranked in order of relative abundance as follows: Staphylococcus, Arsenophonus, Chryseobacterium, Serratia, Enterobacter, Pseudomonas, Actinomadura, Rhizobium, Roseateles, and Massilia (Fig. 2B). The distribution of these genera exhibited marked habitat specificity. For instance, Actinomadura was predominantly found in soil samples, while Rhizobium and Roseateles were enriched in root samples. In plant tissues, healthy stems and leaves harbored relatively balanced bacterial communities. However, aphid feeding induced substantial shifts in community composition (Fig. 2B). In infested stems, Staphylococcus became markedly enriched and emerged as the dominant genus. Similarly, in infested leaves, Staphylococcus replaced Chryseobacterium—the former dominant taxon—becoming the most abundant genus. In insect-associated samples, Arsenophonus predominated in A0 aphids, whereas A10 aphids, which had fed on zucchini for ten days, were primarily associated with Staphylococcus (Fig. 2B). In the honeydew samples, Serratia, Enterobacter, and Staphylococcus were the dominant genera (Fig. 2B).

Fig. 2. Microbial community composition across sample groups and source tracking of Staphylococcus in honeydew.

Fig. 2

A Relative abundance of bacterial communities at the phylum level and B at the genus level. C Source model of the microbiome showing the potential sources of aphids and honeydew bacterial communities based on environmental microbiome samples. U, unknown sources. HS, HR, Hst, and HL represent healthy soil, roots, stems, and leaves, respectively; DS, DR, Dst, and DL denote the corresponding aphid-infested samples. A0 refers to newly hatched nymphs (<24 h old), A10 to nymphs after 10 days of feeding, and HD to honeydew.

Staphylococcus enrichment in aphid-damaged zucchini stems and leaves

Group-wise comparisons of the top five most abundant bacterial genera (Fig. 3A) revealed that Staphylococcus was the most dominant genus, exhibiting significant differences in distribution among sample groups (P < 0.00001). Specifically, the relative abundance of Staphylococcus was markedly higher in the stems and leaves of aphid-infested zucchini plants compared to healthy zucchini plants, while it was scarcely detected in soil and root samples (Fig. 3A). In addition, Staphylococcus was highly enriched within A10, and a low abundance of this genus was also detectable in honeydew.

Fig. 3. Staphylococcus was enriched in aphid-infested zucchini stems and leaves.

Fig. 3

A Significant differences in the relative proportions of major bacterial genera across different samples, were observed based on the relative abundances of the top five genera, as determined by the Kruskal–Wallis test followed by Tukey’s test. B The relative abundance of Staphylococcus OTU1970 in each sample. Statistical significance was assessed using one-way ANOVA (n = 5, P < 0.05, Tukey’s test). C Phylogenetic relationships of Staphylococcus OTUs. HS, HR, Hst, and HL represent healthy soil, roots, stems, and leaves, respectively; DS, DR, Dst, and DL denote the corresponding aphid-infested samples. A0 refers to newly hatched nymphs (<24 h old), A10 to nymphs after 10 days of feeding, and HD to honeydew.

To elucidate the population structure of Staphylococcus in zucchini stems and leaves, we conducted a phylogenetic analysis of operational taxonomic units (OTUs) assigned to this genus (Fig. 3C). Among the identified OTUs, OTU1970 overwhelmingly dominated the Staphylococcus community, accounting for 99.9% of its relative abundance. Phylogenetic analysis further revealed that OTU1970 and OTU3315 clustered together with S. sciuri in the phylogenetic tree (Fig. 3C), supported by a bootstrap value of 97, indicating a close evolutionary relationship. Subsequent analysis of OTU1970 abundance showed a significant increase in its relative abundance in zucchini stems and leaves following aphid feeding (P < 0.0001), with levels markedly higher than those in healthy plants (Fig. 3B). These findings suggest that aphid infestation substantially promotes the enrichment of Staphylococcus, particularly OTU1970, in zucchini stems and leaves.

Source tracking of the honeydew microbiome

To investigate the contribution of environmental microbes to the microbiota of aphids, we employed microbial source tracking (MST) using SourceTracker to quantify the origins of Staphylococcus within aphids (Fig. 2C). The analysis revealed that zucchini stems and leaves infested by aphids were the primary sources of Staphylococcus in A10 individuals, jointly accounting for 68% of the contribution, whereas stems and leaves from healthy plants contributed only 10.4%. Contributions from soil and roots were negligible. Further analysis showed that 93.9% of Staphylococcus detected in honeydew could be traced back to the microbiota of A10 aphids (Fig. 2C). These findings suggest that aphid feeding behavior markedly facilitates the rapid transmission and accumulation of microbes, particularly Staphylococcus, within aphids and their honeydew.

Mated female ladybird beetles are significantly attracted to Staphylococcus sp. in honeydew

To identify bacterial taxa in honeydew with the potential to attract ladybirds, we isolated four representative strains through culturing and 16S rRNA gene sequencing. In Fig. 4A, phylogenetic analyses assigned these strains to the genera Staphylococcus (ABH1), Pantoea (ABH2), Enterobacter (ABH3), and Stenotrophomonas (ABH4). Construction of phylogenetic trees and comparison with the NCBI database revealed that the 16S rRNA gene sequences of all isolates shared 99.9% similarity with known species (Fig. 4B and Table S1).

Fig. 4. Isolation and identification of honeydew-associated bacteria.

Fig. 4

A Colony morphology of honeydew bacteria isolated and purified on LB agar plates. B Phylogenetic tree of honeydew-isolated strains and closely related species based on 16S rRNA gene sequences.

As shown in Fig. 6A, mated female ladybird beetles P. japonica were significantly attracted by sterilized honeydew inoculated with isolate ABH1 (Staphylococcus sp.: χ 2 = 9.6, P = 0.002, n = 60). In contrast, the other three bacterial strains, ABH2 (Pantoe sp.: χ² = 1.067, P = 0.302, n = 60), ABH3 (Enterobacter sp.: χ² = 0.016, P = 0.898, n = 60), and ABH4 (Stenotrophomonas sp.: χ² = 0.267, P = 0.606, n = 60), did not attract mated females.

Fig. 6. Olfactory responses of mated female ladybird beetles to different odorants.

Fig. 6

A Olfactory responses to four sterilized honeydew treatments inoculated with individual bacterial isolates. B Olfactory responses to 4-isopropylbenzyl alcohol at varying concentrations. Control in (B) is glycerol. Asterisks indicate significant differences (χ² test): *P < 0.05, **P < 0.01, ***P < 0.001, and n.s. indicates no significant differences.

Identification of volatile compounds from Staphylococcus sp. inoculated honeydew that elicit ladybird beetles' electroantennographic responses

Using GC-EAD, the electrophysiological responses of the antennae of mated female P. japonica were analyzed in response to the volatile compounds from sterilized honeydew inoculated with the isolated strain ABH1 (Staphylococcus sp.). A significant potential fluctuation peak was observed at a retention time of 11.23 min (Fig. 5), indicating that the compound effectively activated the electrophysiological responses of the female’s antennae. Identification based on GC-MS analysis and spectral comparison with the NIST database confirmed 4-isopropylbenzyl alcohol as the active compound, with a detected concentration of 3.16 ± 0.21 μg/mL.

Fig. 5. GC-EAD analysis of the electrophysiological responses of mated female ladybird beetles’ antennae to crude extracts of honeydew inoculated with the Staphylococcus ABH1 strain.

Fig. 5

The results are shown in two parts: the upper part displays the GC-EAD electrophysiological response profile of the antennae, while the lower part shows the GC-FID chromatogram. Dashed lines indicate the regions corresponding to compounds that elicit active electrophysiological responses.

4-Isopropylbenzyl alcohol significantly attracts mated females of P. japonica and induces oviposition

As shown in Fig. 6B, mated females of P. japonica exhibited distinct olfactory responses to 4-isopropylbenzyl alcohol at both tested concentrations (3 μg/mL and 10 mg/mL), as shown in Fig. 6B. Significant attraction was observed at 3 μg/mL (χ² = 5.40, P = 0.02, n = 60), and the response became highly significant at 10 mg/mL (χ² = 11.27, P = 0.001, n = 60).

Following the observed olfactory responses of mated females to 4-isopropylbenzyl alcohol, cage assays were performed to assess the oviposition preferences of mated P. japonica (Fig. 7A, B). During a 4-h behavioral assay, 4-isopropylbenzyl alcohol elicited a significant oviposition attraction in mated females (3 μg/mL: df = 18, t = −2.64, P = 0.017; 10 mg/mL: df = 18, t = −5.19, P < 0.0001).

Fig. 7. Oviposition preference of mated female ladybird beetles in cage experiments.

Fig. 7

A Comparison of oviposition preference between the control group and the treatment group provided with 200 μL of 4-Isopropylbenzyl alcohol. Control is glycerol. Asterisks indicate significant differences (Student’s t-test): *P < 0.05, ***P < 0.001. B Schematic diagram of the cage setup. The cage size was 90 cm × 90 cm × 90 cm, enclosed on all sides with fine mesh. The number of eggs laid on the plants, pots, or the fine mesh near the plants were recorded (area size: width = 20 cm, height = 30 cm).

Discussion

Currently, research on plant-pest-natural enemy interactions has become relatively advanced. However, studies specifically focusing on the interactions among plants, microbes, pests, and natural enemies are still lacking. In this study, we found that Staphylococcus was significantly enriched in the stems and leaves of zucchini C. pepo following infestation by aphid A. gossypii. Source tracking analysis further confirmed the pivotal role of the plant microbiome in shaping the microbial communities within both the aphid and honeydew. Additionally, we successfully isolated Staphylococcus sp. from the honeydew and discovered its significant attraction to mated females of the ladybird beetle P. japonica. Volatile compound analysis identified 4-Isopropylbenzyl alcohol as the key attractant, which significantly increased the likelihood of oviposition by P. japonica.

Plants often undergo restructuring of their internal microbial communities in response to biotic stressors such as insect herbivores. This restructuring, marked by the selective enrichment of specific microbial taxa, plays a crucial role in the plant’s defense mechanisms41,42. In this study, we observed a pronounced enrichment of Staphylococcus in the stems and leaves of zucchini after infestation by aphid A. gossypii, suggesting that these bacteria may mediate the plant’s defensive responses. This result aligns with findings by a previous study reported a similar increase in the relative abundance of Pseudomonas in willow S. babylonica following herbivory by the leaf beetle P. versicolora8. Similarly, the leaf-mining insect Scaptomyza nigrita induces jasmonic acid-mediated plant defenses, reshaping the phyllospheric microbiome of the native forb Cardamine cordifolia43. Numerous studies have reported that insect herbivory can alter the composition of rhizosphere microbial communities. Both aboveground herbivores, such as cabbage aphid Brevicoryne brassicae and diamondback moth Plutella xylostella, and belowground pests, such as cabbage maggot Delia radicum, have been implicated in inducing significant changes in the rhizosphere microbiome, which in turn influences plant resistance through plant-soil feedback mechanisms44. Additionally, the aphid Macrosiphum euphorbiae alters the rhizosphere microbiome of tomato plants, with soil legacy effects influencing aphid performance in subsequent generations of plants45. However, in our study, feeding by the aphid primarily altered the bacterial diversity of zucchini stems and leaves, rather than that of the rhizosphere. We hypothesize that aphid feeding promotes the accumulation of Staphylococcus in the stem and leaf tissues. This accumulation does not appear to originate from the soil or rhizosphere but may instead result from the proliferation of Staphylococcus already residing in the aboveground parts of the plant. The underlying mechanisms of this proliferation remain to be elucidated.

The role of host plants in shaping the microbiota of herbivorous insects has been well documented across multiple studies46,47. Plant tissues and leaf surfaces harbor diverse microbial communities, and given that herbivorous insects rely on plants as their primary food source, plant-associated microbes are considered a major contributor to the insect microbiome48,49. Our study further verified and quantified this process within the aphid–plant system. The results showed that Staphylococcus sp. were barely detectable in the soil and rhizosphere but were present in measurable amounts in the stems and leaves of zucchini, suggesting that the aboveground parts of the plant may serve as an important source of this bacterium. Correspondingly, Staphylococcus sp. was nearly undetectable in newly hatched nymphs (less than 24 h old), whose microbiota was primarily composed of Arsenophonus, a symbiont known to be vertically transmitted in aphids50. In contrast, after 10 consecutive days of feeding on zucchini, the abundance of Staphylococcus sp. in the aphid microbiota increased significantly, becoming the dominant bacterial group. This dynamic shift suggests that aphids acquire substantial amounts of Staphylococcus from continuous feeding on the host plant. Moreover, a notable clustering of the microbial communities from the infested zucchini stems and leaves, aphids, and honeydew was observed. Source tracking analysis further indicated that the plant microbiome plays a pivotal role in shaping the aphid’s microbiota. However, whether these plant-derived microorganisms can stably colonize the insect and be transmitted vertically to subsequent generations remains to be explored further.

The microbial composition of insect excretion is significantly correlated with insect species and host plants13,51. In this study, we found that the dominant bacterial genera in the aphid honeydew were Serratia, Enterobacter, and Staphylococcus. Source tracking analysis revealed that some of the microbes in the honeydew originated from the adult aphid’s microbiota, a result that is consistent with the findings of a previous study52. Based on the analysis of the source track of Staphylococcus in plants, aphids, and honeydew, we suggest that aphids acquire Staphylococcus from plant tissues during feeding and subsequently release some of these bacteria into the honeydew during excretion. This suggests that plant-associated microbes may enter the environment indirectly through herbivorous insect feeding and excretion behaviors, potentially forming a “plant–microbe–insect–excretion” transmission pathway. Additionally, we observed that the microbial diversity in honeydew was significantly higher than that in the aphid microbiota, as indicated by indices such as Ace and Shannon. This phenomenon may be attributed to the chemical properties of the honeydew, which is rich in sugars, amino acids, and other nutrients, providing a suitable growth substrate for environmental microbes32,34. Therefore, in addition to the aphid’s endogenous microbes, honeydew is also likely to adsorb environmental microbes from the air, plant surfaces, or soil, resulting in a more complex and diverse microbial composition.

Staphylococcus is a common environmental microorganism, and previous studies have shown its potential to promote plant growth53. Furthermore, bacteria from this genus play multiple roles in the physiological and ecological processes of insects. For instance, S. sciuri, found in the gut of the silkworm, can effectively defend against fungal parasitic infections by secreting a chitinase-like antifungal lysozyme48. In this study, we isolated and cultured bacteria from the honeydew and identified, through olfactory behavior experiments, that Staphylococcus sp. significantly attracts mated females of the ladybird beetle P. japonica. This finding is consistent with a previous study that isolated S. sciuri from the honeydew of the pea aphid A. pisum and confirmed that this bacterium acts as an allelochemical, significantly enhancing the control efficiency of aphids by natural enemies34.

In this study, we found that 4-Isopropylbenzyl alcohol released by Staphylococcus sp. significantly attracted mated female of the ladybird beetle P. japonica and notably increased their oviposition near plants treated with this compound. This suggests that 4-Isopropylbenzyl alcohol plays a crucial role in mediating the oviposition site choice of P. japonica. Previous studies have demonstrated the broad potential of phenylmethanol and its derivatives in regulating insect behavior. For instance, benzyl alcohol and benzyl geranate derivatives were reported to exert repellent effects on pea aphid A. pisum54. A blend of benzyl alcohol and (E)-2-hexenal has been shown to exert strong attractiveness to the predatory insect Arma chinensis55, while 4-isopropylbenzyl alcohol demonstrates significant repellent activity against the yellow fever mosquito Aedes aegypti56. Although benzyl alcohol compounds have been relatively well studied for their roles in modulating insect behavior, research on 4-isopropylbenzyl alcohol remains limited. The behavioral responses of insects are often highly sensitive to the concentration of volatile compounds. In this study, we evaluated the behavioral responses to 4-isopropylbenzyl alcohol at a low concentration of 3 μg/mL (corresponding to the level detected in S. sciuri-inoculated honeydew) and a high concentration of 10 mg/mL (a level commonly used for attracting insect natural enemies in field experiments). Our results revealed that both concentrations elicited significant attraction in mated females of P. japonica.

Our study uncovers a novel microbially mediated mechanism of indirect interaction within the tritrophic relationship among plants, herbivorous insects, and their natural enemies. We demonstrate that zucchini C. pepo mounts a “call-for-help” response upon aphid infestation, facilitated by microbe-derived chemical signals. Specifically, Staphylococcus sp. become enriched on stems and leaves following aphid A. gossypii feeding, enter the aphid body during ingestion, and are subsequently excreted via honeydew. Staphylococcus sp. colonizes aphid honeydew and releases the volatile compound 4-Isopropylbenzyl alcohol, which attracts mated females of the ladybird beetle P. japonica and induces oviposition near aphid colonies, thereby enhancing the biological control efficiency of natural enemies. This work advances our understanding of plant–microbe–herbivore–predator interactions and offers a conceptual foundation for developing environmentally sustainable pest management strategies based on microbial and chemical signaling.

Methods

Plant cultivation and insect rearing

Zucchini C. pepo seeds (variety: Bixiu; four per pot) were sown in plastic seedling pots (14 cm in diameter, 10 cm in height) filled with a substrate composed of peat and vermiculite at a 3:1 ratio. In April 2023, the aphids A. gossypii and the ladybird beetles P. japonica were collected from zucchini plants at Huazhong Agricultural University (Hubei Province, China). A single aphid was transferred onto the zucchini plant (cotyledon stage) to establish a colony. During our study, the zucchini plants were replaced every 10 days to ensure a stable food supply. The ladybird beetles were individually reared on A. gossypii in glass tubes (2 cm in diameter, 6 cm in height). The laboratory conditions were maintained at 26 ± 1 °C, 65 ± 5% relative humidity, and a 16 L:8D light cycle.

Sample collection and processing

When the plants reached the cotyledon stage, 500 newly emerged adult aphids were transferred onto the leaves of four plants in each pot and allowed to feed for 10 days. Zucchini plants without aphid infestation were used as controls. After 10 days of infestation, samples from both aphid-infested and control plants were collected for 16S rRNA gene sequencing, including rhizosphere soil, roots, stems, leaves, aphids, and honeydew. All samples were collected with five biological replicates, and each replicate was obtained from a different pot. Rhizosphere soil was collected after removing large clumps and loose soil, and the remaining soil adhering to the roots was gently brushed off using a sterile brush. A total of 500 mg of soil per sample was transferred into 2 mL sterile tubes. After collecting the rhizosphere soil, the roots were placed into sterile tubes, washed with PBS to remove surface soil, and 100 mg of root per sample was taken. The roots were then sequentially rinsed three times with sterile water, disinfected with 75% ethanol for 1 min, and washed four times with sterile water. Finally, the roots were placed on sterile filter paper to remove excess moisture before being transferred into 2 mL sterile tubes. For the zucchini stems, 100 mg of stem tissue per sample was collected, washed, and disinfected in the same manner as the roots before being chopped and placed into 2 mL sterile tubes. For the zucchini leaves, the two cotyledons were excised from the zucchini plant, cut into 2 cm × 2 cm pieces, and 100 mg per sample was collected. These leaf samples were treated with the same procedure before being transferred into 2 mL sterile tubes. Aphids were collected from zucchini leaves, with 50 individuals per sample. A0 refers to newly hatched nymphs (<24 h old), while A10 refers to individuals derived from A0 after feeding on zucchini leaves for 10 days. The aphids were washed four times with sterile water, centrifuged at 10,000 rpm for 3 min, and the supernatant was removed. The centrifuge tubes were inverted on sterile filter paper to eliminate any remaining liquid.

Honeydew from aphids was collected using the aluminum foil collection method34. An aluminum foil was placed under the zucchini leaves infested with A. gossypii. After 24 h, all the aluminum foil was collected, and the crude honeydew was extracted using a 1 mm × 100 mm glass capillary tube into 2 mL sterile tubes. Subsequently, 30 μL of honeydew was transferred into 1.5 mL sterile tubes for each sample. The collected honeydew was used for subsequent experiments on the same day. Crude honeydew was used for 16S rRNA gene sequencing and bacterial isolation.

DNA extraction, PCR amplification, and high-throughput sequencing

DNA was extracted separately from A. gossypii and their honeydew using the TIANamp Genomic DNA Kit, following the manufacturer’s instructions. For the extraction of DNA from roots, stems, and leaves, the Plant Genomic DNA Kit was used. Soil DNA was extracted using the FastDNATM SPIN Kit for Soil. Bacterial 16S rDNA V5–V7 region amplification was performed using the primers 779F (5‘-AACMGGATTAGATACCCKG-3’), 1193R (5‘-ACGTCATCCCCACCTTCC-3’), and 1392 R (5‘-ACGGGCGGTGTGTRC-3’) for nested PCR amplification57. The amplification reaction system was 20 μL, including 10 μL of Pro Taq DNA polymerase (2×), 0.8 μL of forward and reverse primers (5 μM), 10 ng/μL template DNA, and 7.4 μL of ddH2O. The PCR amplification was performed in two stages: in the first stage, 799 F and 1392R primers were used with the following conditions: an initial denaturation at 95 °C for 3 min, followed by 27 cycles of 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s, with a final extension at 72 °C for 10 min. The purified first-round product was used as the template for the second stage, using the 799F and 1193R primers with 13 cycles and the same conditions as the first stage. The PCR product was excised from a 2% agarose gel, purified with the PCR Clean-Up Kit (YuHua, Shanghai, China) following the manufacturer’s protocol, and quantified using Qubit 4.0 (Thermo Fisher Scientific, USA). The purified PCR products were then used to construct sequencing libraries with the NEXTFLEX Rapid DNA-Seq Kit. Sequencing was conducted on the Illumina NextSeq 2000 platform. The sequences from this study were deposited in GenBank SRA with the accession numbers PRJNA1237543 (healthy and aphid-infested soil), PRJNA1237561 (healthy and aphid-infested roots, stems, and leaves), PRJNA1237414 (aphids), and PRJNA1237549 (honeydew).

Microbiome analysis

The QIIME pipeline was used to screen and trim raw data58, while paired-end sequences were combined with Flash. OTU clustering was performed at a 97% similarity threshold using UPARSE 7.159,60, with chimeric sequences removed. Sequences annotated as chloroplasts and mitochondria were filtered out. To ensure data comparability, all sample sequences were rarefied to 6000 reads. Based on the normalized data, taxonomic annotation of OTUs was conducted using the RDP classifier61 (confidence threshold of 70%) against the Silva 16S rRNA database (v138). Alpha diversity indices were calculated using mothur v1.30.162, and intergroup differences were assessed with the Wilcoxon rank-sum test63. Beta diversity was analyzed by principal coordinate analysis (PCoA) based on Bray-Curtis dissimilarity using Vegan v2.5.364, and statistical significance of community composition differences was assessed by PERMANOVA. Source Tracker software and its Bayesian algorithm estimated sink sample proportions based on community structures65. A phylogenetic tree was constructed using the neighbor-joining method in MEGA 11.0, incorporating both sequenced and downloaded sequences66. The tree was then visualized and annotated using the iTOL online tool67.

Isolation and identification of bacteria from A. gossypii honeydew

Bacteria from aphid honeydew were isolated and cultured on Luria-Bertani (LB) agar plates. Serial dilutions of the honeydew solution (10⁻¹ to 10⁻⁵) were prepared, and 20 μL of each dilution was spread onto 90 mm LB agar plates. The plates were sealed, inverted, and incubated at 32 °C for 72 h. Based on colony morphology, isolates were purified through five rounds of the three-zone streaking method (Fig. 4A). Distinct colonies were transferred to 60 mm plates and incubated for seven days before being inoculated into LB liquid medium and cultured at 32 °C and 220 rpm for 10 h. Finally, bacterial cultures were mixed with 50% glycerol at a 1:1 ratio and stored at −80 °C.

The isolated bacterial strains were identified based on 16S ribosomal RNA gene sequences. Genomic DNA was extracted from pure bacterial cultures using the Bacterial Genomic DNA Extraction Kit (Tiangen Biotech, Beijing, China). The full-length 16S rRNA gene was amplified using the universal primers 27F (5’-AGAGTTTGATCGGCTCAG-3’) and 1492R (5’-TACGGYTACCTTGTTACGACTT-3’)68. The PCR reaction system consisted of 1 μL bacterial solution as the template, 22 μL Golden Star T6 Super PCR Mix (Tsingke Biotech, Beijing, China), 1 μL of each primer, and a final volume of 25 μL. The PCR conditions were as follows: initial denaturation at 95 °C for 5 min, followed by 30 cycles of 95 °C for 30 s, 56 °C for 15 s, and 72 °C for 30 s, with a final extension at 72 °C for 5 min. The PCR products were verified by 1% agarose gel electrophoresis and subsequently subjected to bidirectional sequencing by Sangon Biotech (Shanghai, China). The obtained 16S rRNA gene sequences were compared against the NCBI BLAST database, confirming their identities as ABH1 (Staphylococcus sp.), ABH2 (Stenotrophomonas sp.), ABH3 (Pantoea sp.), and ABH4 (Enterobacter sp.). The 16S rRNA gene sequences of all isolated strains have been deposited in the GenBank database under accession numbers PQ844752–PQ844755. A phylogenetic tree was constructed using MEGA 7.0 to illustrate their evolutionary relationships69.

Analysis of bacterial volatiles by coupled gas chromatography-electroantennography detection (GC-EAD)

Twenty microliters of purified single bacterial strain (ABH1, ABH2, ABH3, or ABH4) was inoculated into 5 mL of LB liquid medium and cultured at 32 °C, 220 rpm until the OD600 reached 0.6–0.870. Then, 1 mL of the bacterial culture was transferred to a 2 mL sterile centrifuge tube, centrifuged at 3000 rpm for 5 min to collect the cells. The supernatant was discarded, and the cell pellet was resuspended in 1 mL of PBS and mixed thoroughly. This washing step was repeated twice. The cell pellet was then resuspended in an appropriate amount of sterile honeydew, adjusting the OD600 to 0.2, and incubated at 32 °C, 220 rpm for 24 h. The resulting sterile honeydew inoculated with a single bacterium was used for olfactometer experiments and for GC-EAD and gas chromatography-mass spectrometry (GC-MS) analyses to identify volatile compounds.

Volatile compounds from the honeydew of the ABH1 strain were collected using dynamic headspace adsorption. The process is as follows: 200 μL of the sample was transferred into a sample vial, and the airflow rate of the gas introduction device was maintained at 400 mL/min. The airflow passed through activated carbon filtration and distilled water humidification, then carried the bacterial volatiles from the vial into an adsorption tube containing 200 mg of PoraPak™ Q (80–100 mesh). The adsorption tube was connected to the gas introduction device. Each sample was collected for 10 h. After collection, a total of 400 μL of HPLC-grade n-hexane was added in three successive rinses to elute and dissolve the volatile compounds, which were then collected into a single sample vial and stored at –20 °C for subsequent GC-EAD and GC-MS analyses.

To identify the active compounds in the volatile substances of the ABH1 strain that induce electro-physiological responses in mated female ladybird beetles P. japonica (3–4 days post-mating), GC-EAD analysis was conducted. A complete antenna from the ladybird beetle was carefully removed using a sharp surgical knife, with the distal 0.5 mm cut off to enhance electrical contact. The antennae were fixed at both ends onto two glass microelectrodes, which were filled with an electrolyte solution and connected to a high-impedance amplifier to amplify the electrical signals generated by the antenna4,70,71. Before injecting the sample, baseline signals were recorded to ensure signal stability. Then, 1 μL of the volatile sample was injected through the gas chromatography (GC) injector. The sample was separated using a DB-5MS capillary column (30 m × 0.25 mm × 0.25 μm) with the following chromatography conditions: the oven temperature was initially held at 40 °C for 7 min, then increased at 4 °C/min to 250 °C and held for 5 min, followed by a second ramp of 7 °C/min to 280 °C, where it was maintained for 1 min. The carrier gas was high-purity helium (>99.99%) with a flow rate of 1.0 mL/min. The separated volatile components were simultaneously introduced into both the flame ionization detector (FID) and the electroantennogram (EAD) device using a Y-type splitter. FID was used to detect the chromatographic peaks of the volatiles, while EAD was used to record the electrophysiological response signals of the ladybird beetle antennae. By comparing the FID chromatogram and EAD signal, the retention time of the bacterial volatile compounds was correlated with the corresponding antennal response, and then the active compounds that caused the EAD response were identified.

Volatile compounds were analyzed using coupled GC-MS

Volatile compounds from the honeydew of the ABH1 strain were collected using dynamic headspace adsorption described in the GC-EAD experiment. The volatiles trapped on the adsorbent were eluted with 400 μL n-hexane, and 400 ng nonyl acetate was added to the eluent as internal standard. The volatile compounds were analyzed using an Agilent 7890B gas chromatograph coupled with a 7000D mass spectrometer (GC-MS). The GC conditions followed those described above. The MS parameters were set as follows: electron ionization (EI) source at 230 °C, ionization energy of 70 eV, transfer line temperature of 280 °C, and full scan mode (m/z 30–350, 3 scans/s) combined with selected ion monitoring (SIM). Compounds were initially identified by matching their mass spectra against the NIST 14 spectral library and verifying their retention times using authentic reference standards. Relative quantification of identified compounds was based on comparison of their peak areas with the internal standard72,73.

Olfactometer assay

A Y-tube olfactometer was used to assess the behavioral response of female ladybird beetles P. japonica (3–4 days post-mating) to different odor sources. The olfactometer was made of glass and consisted of a main arm (20 cm) and two side arms (15 cm each) at a 75° angle, with an internal diameter of 2.5 cm4,40. The odor sources utilized in the experiments were as follows: (1) each of the four isolated bacterial strains was individually inoculated into sterilized honeydew (OD₆₀₀ = 0.2), with uninoculated sterile honeydew serving as the control; (2) the selected volatile compounds were diluted to two concentrations (3 μg/mL and 10 mg/mL), with glycerol serving as the corresponding solvent control. A concentration of 10 mg/mL was selected, as it has been widely used for insect natural enemy attraction in field experiments7476. Sterile honeydew was prepared by filtration through a 0.22 μm filter using a 1 mL syringe to remove microorganisms34,35,70. The airflow system was powered by a vacuum pump, maintaining an airflow rate of 400 mL/min. Air was purified sequentially through an activated carbon filter (glass drying tower) and a distilled water bottle before being split into two 250 mL glass bottles. Each bottle had an inlet and an outlet tube, connecting the pump and the Y-tube olfactometer’s side arms. Odor sources were prepared by applying 20 μL of the test volatile compound onto a 2 cm × 5 cm filter paper strip (Solarbio, Beijing, China), which was then placed inside the glass bottle. The volatiles were delivered to the bifurcation point of the Y-tube via Teflon tubing. At the start of the experiment, a single mated female P. japonica was gently introduced into the base of the Y-tube and given 5 min to respond. A choice was recorded if the beetle moved at least one-third of the way into a side arm and remained there for at least 15 s; otherwise, it was recorded as “no choice.” To eliminate positional bias, the positions of the Y-tube arms were switched every two replicates, and the odor sources on the filter paper strips were replaced accordingly. After each experiment, the Y-tube olfactometer apparatus was thoroughly cleaned with 75% ethanol and n-hexane to eliminate residual odors, followed by rinsing with deionized water. The apparatus was then dried in a 150 °C oven for 6 h to ensure complete decontamination. Experiments were conducted in a dark room under controlled conditions (25 ± 1 °C, 65 ± 5% relative humidity). A red light was positioned above the Y-tube’s center and arm ends as the only illumination. All tests were performed between 9:00 AM and 5:00 PM. Each odor pair was tested at least 60 times, with each ladybird beetle used only once.

Cage experiments

Following the olfactometer bioassay, a two-choice cage experiment was conducted in the bioassay chamber to evaluate the oviposition preference of mated female ladybird beetles P. japonica. The experiment was performed under controlled environmental conditions (26 ± 1 °C, 65 ± 5% RH). The experimental setup consisted of a standard insect-rearing cage (90 cm × 90 cm × 90 cm), with three-week-old zucchini plants at the two-true-leaf stage. Three potted zucchini plants were placed in each of the four corners of the cage, with a filter paper positioned centrally among each group of three plants. A diagonal distribution design was employed, with the experimental and control treatments applied to filter papers at opposing diagonal positions. A 200 μL aliquot of 4-isopropylbenzyl alcohol (3 μg/mL and 10 mg/mL) was applied to filter paper in the treatment group, whereas an equal volume of glycerol was applied to the diagonally opposite position as the control7779. To maintain stable odor concentration, the filter papers were replaced every 30 min. At the start of the experiment, 15 mated female P. japonica were released at the center of the cage. After a 4-h exposure period, the number of eggs laid in each diagonal section and on the surrounding plants was recorded (Fig. 7B, egg-laying area size: Width × Height = 20 cm × 30 cm).

Statistical analysis

Data were first tested for normality and homogeneity of variances using the Kolmogorov–Smirnov and Levene’s tests, respectively. Differences in the relative abundance of Staphylococcus OTU1970 among groups were analyzed using one-way analysis of variance (ANOVA), and significant differences among means were identified using Tukey’s post hoc multiple comparison test. Behavioral responses of P. japonica in Y-tube olfactometer assays were evaluated using the χ² goodness-of-fit test, with an expected 50% response for each olfactometer arm. In the cage experiments, differences in the number of eggs laid were analyzed using Student’s t-test. Alpha diversity was assessed using the ACE index (for species richness) and the Shannon index (for species diversity), while beta diversity was evaluated by performing PCoA based on Bray–Curtis distances using QIIME2. Differentially abundant bacterial taxa at the genus level among treatments were identified using LEfSe analysis. All statistical analyses were performed using SPSS 26.0, figures were generated with GraphPad Prism 8.0, and final layouts were assembled using Adobe Illustrator 2020.

Supplementary information

Supplementary Information (26.4KB, docx)

Acknowledgements

This research was supported by the National Natural Science Foundation of China (32302339 and 31672317) and the Frontier Projects of the Applied Foundation of Wuhan Science and Technology Bureau (2019020701011464).

Author contributions

Y.Z. and Y.P. designed the research. Y.L. and J.S. performed the experiments and contributed equally to this work. Y.L., B.J., and S.Z. analyzed the data. Y.L. and Y.Z. wrote the paper. Y.Z. and Y.P. contributed to funding acquisition. All authors contributed to the final version of the manuscript.

Data availability

The sequences from this study were deposited in GenBank SRA with the accession numbers PRJNA1237543 (healthy and aphid-infested soil), PRJNA1237561 (healthy and aphid-infested roots, stems, and leaves), PRJNA1237414 (aphids), and PRJNA1237549 (honeydew).

Competing interests

The authors declare no competing interests.

Footnotes

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

These authors contributed equally: Yue Liu, Jing Sun.

Contributor Information

Yu Peng, Email: pengyu@hubu.edu.cn.

Yao Zhao, Email: zhaoyao@hubu.edu.cn.

Supplementary information

The online version contains supplementary material available at 10.1038/s41522-026-00914-y.

References

  • 1.Liu, M. et al. Sakuranetin protects rice from brown planthopper attack by depleting its beneficial endosymbionts. Proc. Natl. Acad. Sci. USA120, e2305007120 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Onkokesung, N. et al. The plastidial metabolite 2-C-methyl-D-erythritol-2, 4-cyclodiphosphate modulates defence responses against aphids. Plant Cell Environ.42, 2309–2323 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Kloth, K. J. et al. SLI1 confers broad-spectrum resistance to phloem-feeding insects. Plant Cell Environ.44, 2765–2776 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Xiu, C. et al. Volatiles from aphid-infested plants attract adults of the multicolored Asian lady beetle Harmonia axyridis. Biol. Control129, 1–11 (2019). [Google Scholar]
  • 5.Yao, C. et al. Stemborer-induced rice plant volatiles boost direct and indirect resistance in neighboring plants. N. Phytol.237, 2375–2387 (2023). [DOI] [PubMed] [Google Scholar]
  • 6.Shi, J. H. et al. Unprecedented oviposition tactics avoid plant defences and reduce attack by parasitic wasps. Plant Cell Environ.47, 308–318 (2024). [DOI] [PubMed] [Google Scholar]
  • 7.Meijer, D. et al. Effects of far-red light on the behaviour and reproduction of the zoophytophagous predator Macrolophus pygmaeus and its interaction with a whitefly herbivore. Plant Cell Environ.47, 187–196 (2024). [DOI] [PubMed] [Google Scholar]
  • 8.Wang, H. et al. Enrichment of novel entomopathogenic Pseudomonas species enhances willow resistance to leaf beetles. Microbiome12, 169 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Mbaluto, C. M. & Zytynska, S. E. Rhizobacteria prime the activation of plant defense and nutritional responses to suppress aphid populations on barley over time. N. Phytol.247, 2390–2405 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Shan, H. W. et al. The plant-sucking insect selects assembly of the gut microbiota from environment to enhance host reproduction. npj Biofilms Microbiomes10, 64 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Rering, C. C., Vannette, R. L., Schaeffer, R. N. & Beck, J. J. Microbial co-occurrence in floral nectar affects metabolites and attractiveness to a generalist pollinator. J. Chem. Ecol.46, 659–667 (2020). [DOI] [PubMed] [Google Scholar]
  • 12.Rering, C. C., Rudolph, A. B. & Beck, J. J. Pollen and yeast change nectar aroma and nutritional content alone and together, but honey bee foraging reflects only the avoidance of yeast. Environ. Microbiol.23, 4141–4150 (2021). [DOI] [PubMed] [Google Scholar]
  • 13.Pirttilä, A. M. et al. Exchange of microbiomes in plant–insect herbivore interactions. mBio14, e03210–e03222 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Zhang, B., Leonard, S. P., Li, Y. & Moran, N. A. Obligate bacterial endosymbionts limit thermal tolerance of insect host species. Proc. Natl. Acad. Sci. USA116, 24712–24718 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Salem, H. et al. Symbiont digestive range reflects host plant breadth in herbivorous beetles. Curr. Biol.30, 2875–2886 (2020). [DOI] [PubMed] [Google Scholar]
  • 16.Douglas, A. E. Multiorganismal insects: diversity and function of resident microorganisms. Annu. Rev. Entomol.60, 17–34 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Dicke, M., Cusumano, A. & Poelman, E. H. Microbial symbionts of parasitoids. Annu. Rev. Entomol.65, 171–190 (2020). [DOI] [PubMed] [Google Scholar]
  • 18.Ma, M. et al. Gut bacteria facilitate leaf beetles in adapting to dietary specialization by enhancing larval fitness. npj Biofilms Microbiomes10, 110 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Wang, D. et al. Structural diversity of symbionts and related cellular mechanisms underlying vertical symbiont transmission in cicadas. Environ. Microbiol.23, 6603–6621 (2021). [DOI] [PubMed] [Google Scholar]
  • 20.Strano, C. P., Malacrinò, A., Campolo, O. & Palmeri, V. Influence of host plant on Thaumetopoea pityocampa gut bacterial community. Microb. Ecol.75, 487–494 (2018). [DOI] [PubMed] [Google Scholar]
  • 21.Hannula, S. E., Zhu, F., Heinen, R. & Bezemer, T. M. Foliar-feeding insects acquire microbiomes from the soil rather than the host plant. Nat. Commun.10, 1254 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Powell, J. E., Martinson, V. G., Urban-Mead, K. & Moran, N. A. Routes of acquisition of the gut microbiota of the honey bee Apis mellifera. Appl. Environ. Microbiol.80, 7378–7387 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Verdone, M., Rao, R., Coppola, M. & Corrado, G. Identification of zucchini varieties in commercial food products by DNA typing. Food Control84, 197–204 (2018). [Google Scholar]
  • 24.Silva, H. C. C., dos Santos Magalhães, C. & Randau, K. P. Comparative morphoanatomic and histochemical characterization of Cucurbita pepo L. specimens. Flora315, 152510 (2024). [Google Scholar]
  • 25.Gao, X. et al. Parasitism by Lysiphlebia japonica alters the microbiome of Aphis gossypii offspring. Entomol. Gen.43, 991–999 (2023). [Google Scholar]
  • 26.Gao, X. et al. Silencing of cytochrome P450 gene AgoCYP6CY19 reduces the tolerance to host plant in cotton-and cucumber-specialized aphids, Aphis gossypii. J. Agric. Food Chem.70, 12408–12417 (2022). [DOI] [PubMed] [Google Scholar]
  • 27.Yang, Y. et al. Plant volatiles mediate Aphis gossypii settling but not predator foraging in intercropped cotton. Pest. Manag. Sci.79, 4481–4489 (2023). [DOI] [PubMed] [Google Scholar]
  • 28.Zhang, S. et al. Chromosome-level genome assemblies of two cotton-melon aphid Aphis gossypii biotypes unveil mechanisms of host adaption. Mol. Ecol. Resour.22, 1120–1134 (2022). [DOI] [PubMed] [Google Scholar]
  • 29.Jayasinghe, W. H., Akhter, M. S., Nakahara, K. & Maruthi, M. N. Effect of aphid biology and morphology on plant virus transmission. Pest. Manag. Sci.78, 416–427 (2022). [DOI] [PubMed] [Google Scholar]
  • 30.Lee, J. C., Heimpel, G. E. & Leibee, G. L. Comparing floral nectar and aphid honeydew diets on the longevity and nutrient levels of a parasitoid wasp. Entomol. Exp. Appl.111, 189–199 (2004). [Google Scholar]
  • 31.Xu, T. et al. A trail pheromone mediates the mutualism between ants and aphids. Curr. Biol.31, 4738–4747 (2021). [DOI] [PubMed] [Google Scholar]
  • 32.Álvarez Pérez, S., Lievens, B. & de Vega, C. Floral nectar and honeydew microbial diversity and their role in biocontrol of insect pests and pollination. Curr. Opin. Insect Sci.61, 101138 (2024). [DOI] [PubMed] [Google Scholar]
  • 33.de Bobadilla, M. F. et al. Honeydew management to promote biological control. Curr. Opin. Insect Sci.61, 101151 (2024). [DOI] [PubMed] [Google Scholar]
  • 34.Leroy, P. D. et al. Microorganisms from aphid honeydew attract and enhance the efficacy of natural enemies. Nat. Commun.2, 348 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Liu, P. et al. Volatiles produced by bacteria in the honeydew of cotton aphids mediate prey location by Hippodamia variegata (Coleoptera: Coccinellidae). Biol. Control202, 105728 (2025). [Google Scholar]
  • 36.Liu, J. et al. Chemical cues from honeydew-associated bacteria to enhance parasitism efficacy: from laboratory to field assay. J. Pest Sci.97, 873–884 (2024). [Google Scholar]
  • 37.Goelen, T. et al. Identification and application of bacterial volatiles to attract a generalist aphid parasitoid: from laboratory to greenhouse assays. Pest. Manag. Sci.77, 930–938 (2021). [DOI] [PubMed] [Google Scholar]
  • 38.van Neerbos, F. A. C. et al. Bacterial volatiles elicit differential olfactory responses in insect species from the same and different trophic levels. Insect Sci.30, 1464–1480 (2023). [DOI] [PubMed] [Google Scholar]
  • 39.Goelen, T. et al. Volatiles of bacteria associated with parasitoid habitats elicit distinct olfactory responses in an aphid parasitoid and its hyperparasitoid. Funct. Ecol.34, 507–520 (2020). [Google Scholar]
  • 40.Li, X. et al. Bacterial volatiles from aphid honeydew mediate ladybird beetles oviposition site choice. Pest. Manag. Sci.81, 4063–4071 (2025). [DOI] [PubMed] [Google Scholar]
  • 41.Sugio, A., Dubreuil, G., Giron, D. & Simon, J. C. Plant–insect interactions under bacterial influence: ecological implications and underlying mechanisms. J. Exp. Bot.66, 467–478 (2014). [DOI] [PubMed] [Google Scholar]
  • 42.Crowley Gall, A. et al. Volatile microbial semiochemicals and insect perception at flowers. Curr. Opin. Insect Sci.44, 23–34 (2021). [DOI] [PubMed] [Google Scholar]
  • 43.Humphrey, P. T. & Whiteman, N. K. Insect herbivory reshapes a native leaf microbiome. Nat. Ecol. Evol.4, 221–229 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Friman, J. et al. Shoot and root insect herbivory change the plant rhizosphere microbiome and affects cabbage–insect interactions through plant-soil feedback. N. Phytol.232, 2475–2490 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.French, E., Kaplan, I. & Enders, L. Foliar aphid herbivory alters the tomato rhizosphere microbiome, but initial soil community determines the legacy effects. Front. Sustain. Food Syst.5, 629684 (2021). [Google Scholar]
  • 46.Hammer, T. J. et al. Caterpillars lack a resident gut microbiome. Proc. Natl. Acad. Sci. USA114, 9641–9646 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Ge, S. X., Li, T. F., Ren, L. L. & Zong, S. X. Host-plant adaptation in xylophagous insect-microbiome systems: contributionsof longicorns and gut symbionts revealed by parallel metatranscriptome. Iscience26, 106680 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Zhao, P. et al. From phyllosphere to insect cuticles: silkworms gather antifungal bacteria from mulberry leaves to battle fungal parasite attacks. Microbiome12, 40 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Qi, J. et al. The fall armyworm converts maize endophytes into its own probiotics to detoxify benzoxazinoids and promote caterpillar growth. Microbiome12, 240 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Jousselin, E., Cœur d’Acier, A., Vanlerberghe Masutti, F. & Duron, O. Evolution and diversity of a rsenophonus endosymbionts in aphids. Mol. Ecol.22, 260–270 (2013). [DOI] [PubMed] [Google Scholar]
  • 51.Zhu, Y. G. et al. Ecosystem microbiome science. mLife2, 2–10 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Fischer, C. Y. et al. Bacteria may enhance species association in an ant–aphid mutualistic relationship. Chemoecology25, 223–232 (2015). [Google Scholar]
  • 53.Taj, Z. & Challabathula, D. Protection of photosynthesis by halotolerant Staphylococcus sciuri ET101 in tomato (Lycoperiscon esculentum) and rice (Oryza sativa) plants during salinity stress: Possible interplay between carboxylation and oxygenation in stress mitigation. Front. Microbiol.11, 547750 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Pan, S. X. et al. Rational design, synthesis and binding mechanisms of novel benzyl geranate derivatives as potential eco-friendly aphid repellents. Pest. Manag. Sci.80, 1099–1106 (2024). [DOI] [PubMed] [Google Scholar]
  • 55.Wu, H. et al. Identification and field verification of aggregation-sex pheromone from the predaceous bug, Arma chinensis. Chemoecology29, 235–245 (2019). [Google Scholar]
  • 56.Watanabe, K. et al. New mosquito repellent from Eucalyptus camaldulensis. J. Agric. Food Chem.41, 2164–2166 (1993). [Google Scholar]
  • 57.Wang, C. et al. Variations of root-associated bacterial cooccurrence relationships in paddy soils under chlorantraniliprole (CAP) stress. Sci. Total Environ.779, 146247 (2021). [DOI] [PubMed] [Google Scholar]
  • 58.Caporaso, J. G. et al. QIIME allows analysis of high-throughput community sequencing data. Nat. Methods7, 335–336 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59.Magoc, T. & Salzberg, S. L. FLASH: fast length adjustment of short reads to improve genome assemblies. Bioinformatics27, 2957–2963 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60.Edgar, R. C. UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat. Methods10, 996–998 (2013). [DOI] [PubMed] [Google Scholar]
  • 61.Stackebrandt, E. & Goebel, B. M. Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Evol. Microbiol.44, 846–849 (1994). [Google Scholar]
  • 62.Douglas, G. M. et al. PICRUSt2 for prediction of metagenome functions. Nat. Biotechnol.38, 685–688 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 63.Mann, H. B. & Whitney, D. R. On a test of whether one of two random variables is stochastically larger than the other. Ann. Math. Stat.18, 50–60 (1947). [Google Scholar]
  • 64.Wang, Q., Garrity, G. M., Tiedje, J. M. & Cole, J. R. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol.73, 5261–5267 (2007). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 65.Han, C. et al. Majorbio Cloud 2024: update single-cell and multiomics workflows. iMeta3, e217 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 66.Bravo, A. G. et al. Geobacteraceae are important members of mercury-methylating microbial communities of sediments impacted by waste water releases. ISME J.12, 802–812 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67.Letunic, I. & Bork, P. Interactive Tree Of Life (iTOL) v5: an online tool for phylogenetic tree display and annotation. Nucleic Acids Res.49, W293–W296 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 68.Frank, J. A. et al. Critical evaluation of two primers commonly used for amplification of bacterial 16S rRNA genes. Appl. Environ. Microbiol.74, 2461–2470 (2008). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 69.Kumar, S., Stecher, G. & Tamura, K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol.33, 1870–1874 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70.Zhang, G. et al. Semiochemicals produced by microbes in mealybug honeydew attract fire ants. J. Agric. Food Chem.71, 15456–15465 (2023). [DOI] [PubMed] [Google Scholar]
  • 71.Vuts, J. et al. Responses of the two-spotted oak buprestid, Agrilus biguttatus (Coleoptera: Buprestidae), to host tree volatiles. Pest. Manag. Sci.72, 845–851 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 72.Liu, Q. et al. Cooperative herbivory between two important pests of rice. Nat. Commun.12, 6772 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 73.De Lange, E. S. et al. Spodoptera frugiperda caterpillars suppress herbivore-induced volatile emissions in maize. J. Chem. Ecol.46, 344–360 (2020). [DOI] [PubMed] [Google Scholar]
  • 74.Cai, Z. et al. Attraction of adult Harmonia axyridis to volatiles of the insectary plant Cnidium monnieri. Biol. Control143, 104189 (2020). [Google Scholar]
  • 75.Zhu, J. et al. Attraction of two lacewing species to volatiles produced by host plants and aphid prey. Naturwissenschaften92, 277–281 (2005). [DOI] [PubMed] [Google Scholar]
  • 76.Yu, H. et al. Field trapping of predaceous insects with synthetic herbivore-induced plant volatiles in cotton fields. Environ. Entomol.47, 114–120 (2018). [DOI] [PubMed] [Google Scholar]
  • 77.Xu, S. et al. The threat of the fall armyworm to Asian rice production is amplified by the brown planthopper. Plant. Cell Environ.48, 1060–1072 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 78.Hu, X. et al. Caterpillar-induced rice volatiles provide enemy-free space for the offspring of the brown planthopper. Elife9, e55421 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 79.Wang, X. et al. Bt rice could provide ecological resistance against nontarget planthoppers. Plant Biotechnol. J.16, 1748–1755 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Supplementary Information (26.4KB, docx)

Data Availability Statement

The sequences from this study were deposited in GenBank SRA with the accession numbers PRJNA1237543 (healthy and aphid-infested soil), PRJNA1237561 (healthy and aphid-infested roots, stems, and leaves), PRJNA1237414 (aphids), and PRJNA1237549 (honeydew).


Articles from NPJ Biofilms and Microbiomes are provided here courtesy of Nature Publishing Group

RESOURCES