Abstract
The gut microbiota serves as a critical “organ” in the life cycle of animals, particularly in the intricate interplay between herbivorous pests and plants. This review summarizes the pivotal functions of the gut microbiota in mediating the insect–plant interactions, encompassing their influence on host insects, modulation of plant physiology, and regulation of the third trophic level species within the ecological network. Given these significant functions, it is plausible to harness these interactions and their underlying mechanisms to develop novel eco-friendly pest control strategies. In this context, we also outline some emerging pest control methods based on the intestinal microbiota or bacteria-mediated interactions, such as symbiont-mediated RNAi and paratransgenesis, albeit these are still in their nascent stages and confront numerous challenges. Overall, both opportunities and challenges coexist in the exploration of the intestinal microbiota-mediated interactions between insect pests and plants, which will not only enrich the fundamental knowledge of plant–insect interactions but also facilitate the development of sustainable pest control strategies.
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Introduction
Arthropoda is one of the most diverse and successful animal phyla, boasting an estimated 5.5 million insect species, of which only one million species have been named thus far1. More than half of these species are herbivorous insects, which have caused significant economic losses and serious ecological problems to agricultural and forestry production. It has been reported that global crop production suffers a loss of more than 15% annually due to these pests2. In recent years, the issue of pests has been compounded by factors such as climate change, rapid globalization, and urbanization, as well as the spread of invasive species3,4. While chemical pesticides have been widely employed to combat pests, their injudicious use has led to various problems, including the disruption of natural ecosystems, the emergence of pesticide-resistant pests, and adverse health effects on humans5,6,7. Hence, it is crucial to identify eco-friendly methods for pest control in agriculture and forestry.
Insects harbor a diverse and abundant microbial population in their gut system8. Some gut bacteria play a positive role in adapting pests to host plants, including providing nutrition, aiding digestion, detoxification, and directing pest behavior9,10,11. The symbiotic relationships between gut bacteria and insect pests have partly contributed to the success and diversification of insects, but have also increased the difficulty of pest control. Interestingly, some studies have indicated that the function of gut bacteria can change and become detrimental to the host insects when a third species participates in the pest–plant interaction or due to other abiotic factors12,13,14. Therefore, comprehensively understanding the function of gut bacteria in multispecies cascading interactions can not only unveil novel resources for biocontrol but also facilitate the development of new biopesticides.
Many reviews have summarized the functions of gut bacteria in insects, but there have been few reviews that have highlighted the diversified functions of gut bacteria in the context of insect–plant interactions, especially in multispecies cascading interactions7,9,15,16,17. Drawing on previous studies, we will describe the role of gut bacteria in this system from multiple perspectives, and discuss how to screen key bacteria and apply these theories to pest control in combination with other techniques.
Gut bacteria affect the insect–plant relationship by affecting the host insect
Herbivorous insects have adapted to plants that provide heterogeneous resources, which can be considered as the “center of activity” for the insects18. These adaptations may be facilitated by the direct and indirect interactions of insect gut bacteria with host plants19. Among these interactions, certain gut bacteria possess ecological functions that directly influence insect behavior and physiology, which can be generally categorized as follows (Table 1 and Fig. 1); firstly, gut bacteria have been shown to mediate the plant selection preference of insects, assisting them in finding suitable host plants for survival and reproduction (Fig. 1a). Once herbivorous insects successfully locate and attack the selected plants, they often encounter substantial challenges in feeding, including low nutrient content, indigestibility, and toxicity of many plant tissues11. Some insect gut bacteria confer the ability to overcome these feeding obstacles, allowing insects to develop and reproduce on plants (Fig. 1b). Conversely, chemicals or inadequate nutrition in plant tissue cause dysbiosis in the gut microbiota of herbivores insects, which often have adverse effects on their biology (Fig. 1c).
a Gut bacteria mediate insects’ plant selection preference; b Gut bacteria assist insects in overcoming feeding obstacles; c Plant defense-induced dysbiosis of gut microbiota is lethal to insects.
Gut bacteria mediate host plant preferences of insects
Host plant preference enables herbivorous insects to choose a suitable host in terms of the fitness of themselves and their descendants20, primarily relying on their chemosensory system to analyze different plant volatile organic compounds (VOCs) and make behavioral decisions20,21. Indeed, this selection preference is often influenced by microorganisms. While fungi, endophytes, and plant pathogens have been shown to influence insect selection preferences for host plants, a small group of gut bacteria also has a similar effect22,23. For instance, western flower thrips prefer thrips-damaged leaves over fresh leaves, which have more gut bacteria on the leaf surface24.
Gut bacteria are first transferred to host plants by insects through foraging, excretion, or laying eggs. Once inside the plant, gut bacteria use plant nutrients to spread throughout the plant and produce VOCs that attract or repel intraspecific individuals. In one study, the gut bacteria (Citrobacter sp.) of the oriental fruit fly Bactrocera dorsalis attracted other female flies to lay eggs on the host fruit by producing 3-hexenyl acetate (3-HA)10. The gut bacteria of the bark beetle Dendroctonus valens can produce a multifunctional pheromone, verbenone, in its frass, which assists subsequent beetles in determining the suitability of the pine tree for colonization25,26. Gut bacteria produce VOCs not only for behavioral attraction but also to highlight the mutual benefit of interacting organisms. While insects can find suitable places to lay their eggs, their gut bacteria have a means of spreading. Although there have been relatively few reports of gut bacteria directly regulating host plant selection, given that this strategy benefits both insects and gut bacteria, we believe this phenomenon will be uncovered in more insect species.
Gut bacteria digest stubborn plant polymers or provide nutrients lacking in plants
Herbivorous insects tend to feed on all parts of the plant, but digesting stubborn plant polymers presents a challenge as most insects lack the ability to synthesize the necessary enzymes, such as cellulase, pectinase, ligninase, to effectively breakdown these polymers27. The gut microbiome has been shown to aid insects’ digestion by enhancing digestive proficiency via enzymatic potentiality. Cellulose is an abundant source of carbon, but it is not readily available to the insect as it exists in the form of crystalline or amorphous microfibers in plant cell walls28. It needs to be hydrolyzed into small molecules of sugar by cellulase, a process in which gut bacteria are typically involved9. Researchers have identified Streptomyces and Pantoea in the gut of an invasive wood-eating wasp (Sirex noctilio) that provides cellulase to the host insect, enabling the breakdown of cellulose and facilitating nutrient acquisition29. Similar functional gut bacteria (e.g., Provedencia sp., Bacillus spp., and Klebsiella spp.) are also found in Blattodea (Macrotermes gilvus), Coleoptera (Lepidiota mansueta, Odontotaenius disjunctus), and Lepidoptera (Cossus cossus, Diatraea saccharalis)30,31,32,33,34.
The cellulose and hemicellulose fibers within plant cell walls are embedded in the pectin matrix, necessitating the initial digestion of pectin to provide substrates for the enzymes35. Research has demonstrated that the degradation of pectin could also rely on the activity of pectinase produced by insect gut bacteria. For instance, the gut symbiont Stammera of a leaf beetle Cassida rubiginosa possesses genes involved in pectin digestion36. Similarly, in insects that heavily rely on pollen as their primary nutrient source, accessing the nutrients within pollen requires overcoming the barrier of the pollen wall. The chemical composition of the pollen exine layers consists mainly of sporopolysin, while the inner layer contains pectin37,38. Metagenomic studies have revealed the presence of genes associated with pectin degradation in the gut bacteria Gilliamella apicola of honey bees, and in vitro culturing tests have confirmed its pectin degradation activity39. Lignin is another kind of common natural polymer with abundant content and complex structure that is difficult for insects to metabolize. Although fungi might be the main players for lignin degradation in insects, some gut bacteria have the ability to breakdown lignin40. Several studies have isolated and identified bacteria with lignin-degrading potential from the gut of termites40,41,42. Transcriptomic approaches also revealed that midgut microorganisms participate in lignin degradation in the longhorn beetle Anoplophora glabripennis43,44. These studies highlight the ability of various gut bacteria species to assist their hosts in the degradation of resilient plant polymers.
Meanwhile, a plant-based diet is often lacking in some of the nutrients necessary for herbivorous insect survival and development. To alleviate nutrient limitations, insects have evolved various gut bacteria-mediated strategies that allow them to take advantage of plants lacking certain nutrients11. Nitrogen is generally considered to be the limiting factor in the diet of herbivorous insects45. Apart from this, plant proteins lack essential amino acids (EAAs), so the growth and development of insects may be severely limited46. The gut bacteria of herbivorous insects can use atmospheric nitrogen sources for biological nitrogen fixation, such as the passalid beetle Odontotaenius disjunctus and the medfly Ceratitis capitata47,48. In addition, herbivores insects’ gut bacteria employ nitrogenous waste recycling (NWR) as a key method to acquire supplemental nitrogen sources, utilizing these wastes to synthesize essential amino acids (EAAs) that can be reabsorbed by the insect hosts. For example, the symbiotic bacteria of the oriental fruit fly B. dorsalis can drive NWR with the help of Morganella morganii and Klebsiella oxytoca49. Similarly, gut bacteria have been found in leafcutter ants to recycle urea (and possibly uric acid) and use the recycled nitrogen to make large amounts of EAAs50. Besides, insects that feed on plant xylem and phloem sap have limited vitamin availability51. Gut bacteria have been shown to provide vitamins to insect hosts. Symbiotic bacteria colonizing the bean bug Riptortus pedestris midgut produce B vitamins that are scarce in the host insect’s soy diet52. Through genomic analysis, it was demonstrated that the gut bacteria (Bartonellaceae) of herbivorous ants (Dolichoderus) encode genes for several vitamins and all essential amino acid biosynthetic pathways53. The biosynthetic capability of vitamins exhibited strain-specific variability among different bacterial species. For example, while the thiE gene responsible for vitamin B1 synthesis was present in all members of the Bartonellaceae family, only one strain possessed a complete set of genes for the synthesis of vitamin B3.
Gut bacteria degrade plant toxins
In addition to the aforementioned stubborn plant polymers that hinder insect feeding, herbivorous insect attacks can induce phytochemical resistance through the production of secondary metabolites, which often serve as defensive compounds to deter herbivores from feeding54. Aside from the insects’ inherent detoxifying metabolic ability, gut microbes significantly contribute to the degradation of the ingested phytoallelopathic substances55,56,57. For example, gut bacteria of the pine weevil (Hylobius abietis) can help weevils metabolize diterpene acids of Norway spruce, and the gut bacteria can even utilize diterpenoids as a carbon source and may produce nutrients to increase insect fitness58. For the primary pest of Chinese tea plants—Curculio chinensis, Acinetobacter species in the gut was identified to be involved in the degradation of tea saponin59. Some studies have found that caffeine is degraded in the gut of the coffee berry borer (Hypothenemus hampei), and have shown that Pseudomonas species play an important role in the degradation of caffeine60.
Furthermore, the gut microbiota exposed to challenging compounds, and defensive substances can dynamically adjust their structure or remain stable so as to better assist insects in adapting to such environmental stressors61,62,63. The viewpoint has been validated in research systems involving pine sawyers and bark beetles, among others. Gut-associated microbiota of the pine sawyer Monochamus saltuarius can shift in response to different dietary stimuli, which correlates with its diverse ability to metabolize secondary plant metabolites64. In contrast, the gut bacterial community of the bark beetle D. valens maintains resilience, enabling the beetle to catabolize pine defense chemicals65. Both pests are pine feeders, but the two present distinct and opposite patterns of gut microbiota changes, which remain unclear but are worth further investigation. The above examples collectively demonstrate that microorganisms in insect guts can aid in the rapid adaptation of phytophagous insects to plant secondary metabolites. Furthermore, insects utilize gut bacteria to degrade plant defense compounds, thereby allowing them to have more energy available for their own growth, development, and reproduction.
Dysbiosis of gut microbiota is detrimental to insect survival
Plants have various defense mechanisms to resist herbivorous insects66, and the insect gut is frequently targeted by plant defenses. Both physical and chemical plant defenses can disrupt the protective peritrophic matrix (PM) of the gut, resulting in increased permeability and potential invasion by gut bacteria, leading to septicemia67,68,69. One study found that plant defenses against the fall armyworm Spodoptera frugiperda facilitated gut microbes to penetrate gut barriers, invade the body cavity, and worsen the negative impacts of plant defenses on the insect70. Similar observations were reported for the diamondback moth Plutella xylostella, where plant defensive substance (3E)-4,8-dimethyl-1,3,7-nonatriene (DMNT) disrupted midgut microbiota populations and PM, and midgut microbes helped plant toxins kill insects13. Even if the structure of the gut structure is not damaged, disturbance of the gut microbiota can still negatively affect insects. A recent study found that tomato plants colonized by Trichoderma negatively affected the development and survival of another lepidopteran pest Spodoptera littoralis by altering the structure and function of the gut microbiota of S. littoralis larvae71.
Consequently, apart from the beneficial effects of gut bacteria, certain gut bacteria can turn pathogenic when insects are exposed to plant defenses64. The composition and structure of the gut microbiota are crucial for the functional attributes of each gut bacterium. Hence, we are compelled to contemplate whether the gut microbiota influenced by plant defense impacts insect development or if alterations in insect physiology induced by plant defense lead to subsequent modifications in the gut microbiota, thereby further influencing the physiological processes of the insect. Addressing the former hypothesis could be achieved through direct transplantation of the plant-altered microbiota into axenic insects under conditions without plant defense. However, investigating the latter scenario poses more intricate challenges in research design and interpretation. Overall, gut bacteria primarily act as facilitators for insects when the plant’s defense is relatively weak. However, in situations where plants exhibit strong defenses or when insects are in poor physiological condition, gut bacteria often transform into pathogenic bacteria and accelerate the mortality of the pests, as supported by the examples we described above.
Gut bacteria influence the insect–plant relationship by impacting plant physiology
Gut microbiota not only impact the interaction between insects and plants by means of direct effects on the former, but they also shape this association by exerting an influence on the latter (Fig. 2). Insects are able to transmit their gut bacteria to plants through various means such as saliva, reflux, feces, eggs, and honeydew72,73,74, which significantly influence the plants’ physiology in diverse ways. Firstly, some gut bacteria can act as promoters of plant growth like plant-associated microbes have shown. Furthermore, gut bacteria affect the adaptability of insects and plants by inducing plant defenses, including direct defenses mediated by plant hormones, and indirect defenses that attract natural enemies through volatile compounds.
The purple circle represents an example of gut bacteria promoting plant growth directly. The yellow one represents an example of gut bacteria regulating SA/JA signaling pathways of plants. And the brown circle shows an example of gut bacteria inducing the production of plant VOCs to attract/repel natural enemies of insects.
Gut bacteria promote plant growth directly
Plants harbor a myriad of bacteria, including those capable of promoting plant growth75,76,77. Meanwhile, herbivorous insects constantly feed, lay eggs, and traverse plants, providing ample opportunities for plant-associated bacteria to colonize their guts78. The colonization of plant leaf-associated bacteria in the gut of leaf-eating insects is logical and comprehensible, exemplified by the ability of the poplar-associated bacterium Pseudomonas putida to colonize the gut of the leaf beetle Plagiodera versicolora79. Even soil bacteria can be transmitted to aboveground insects (foliar-feeding insects) and colonize their gut78.
Moreover, nutrient cycling rates in the insect gut environment are much higher than in soil and plants, leading to a greater potential for disturbance. Consequently, insect gut bacteria are more metabolically versatile than bacteria found in other environments80, providing a basis for them to influence plant functions. For example, Pseudomonas putida, the gut bacterium of P. versicolora, benefits the host plant by promoting growth and reducing trichloroethane phytotoxicity81. Previous research has demonstrated that eight gut bacteria of diamondback moth larvae (P. xylostella) possess plant growth-promoting traits, including those that fix atmospheric nitrogen, produce indole acetic acid (IAA) and salicylic acid (SA), solubilize phosphates, promote zinc absorption, and produce glucanases, chitinases and 1-aminocyclopropane-1-carboxylate (ACC) deaminase80. Another study found that gut bacteria in Diabrotica virgifera can promote the growth of tomato plants82. Based on the reasons mentioned above and these examples, we have sufficient reasons to believe that more and more discoveries of insect gut bacteria that affect host plant growth will be reported.
Gut bacteria regulate SA/JA signaling pathways of plants
Plant defense is triggered when the plant detects various insect-derived cues, such as tissue damage, oral secretions, oviposition, and frass, and is mediated by phytohormones to elicit resistance83,84. In response to chewing insects, plants activate jasmonic acid (JA)-mediated defenses, while biotrophic pathogens and some piercing–sucking insects elicit salicylic acid (SA)-mediated defenses85,86. The antagonistic relationship between JA and SA signaling pathways enables plants to fine-tune their defense responses to specific organisms87,88.
Herbivorous insect gut bacteria can alter the JA/SA pathways and inhibit plant defense by secreting gut bacteria from oral secretions and frass onto plants during feeding. For example, the Colorado potato beetle (Leptinotarsa decemlineata) carried gut endosymbiotic bacteria that were deposited on plants through the beetle’s oral secretions, and the authors have shown that associated bacteria enhanced the SA signaling pathway and suppressed JA-mediated defenses in tomato89. Similarly, oral secretion-associated bacteria of the cotton leafworm Spodoptera litura contributed to the ability of host insects to manipulate plant defenses by promoting the Arabidopsis SA signaling pathway and inhibiting the JA signaling pathway90. Nevertheless, in some cases, insect gut bacteria can induce the expression of defense-related enzymes and JA regulatory genes. For instance, the gut bacteria of the corn earworm Helicoverpa zea (Enterobacter ludwigii) induced the expression of polyphenol oxidase and JA regulatory genes, while inhibiting SA disease-related genes, providing potential benefit to the tomato plant’s fitness74.
The current research on gut bacteria that can influence JA and SA pathways is mostly focused on chewing insects, while there are fewer studies on the gut bacteria of piercing–sucking insects. Overall, the interaction between insect gut bacteria and plant SA/JA signaling is complex and can involve both synergistic and antagonistic effects, depending on the specific insect species and bacterial strains involved, as well as the host plant57,91. Furthermore, on one hand, gut bacteria of herbivorous insects can promote plant growth, while on the other hand, they can also modulate plant defense responses through JA or SA signaling pathways. In fact, a trade-off exists between plant defense and growth81. Therefore, it is worth exploring how plants balance these seemingly contradictory signals derived from gut bacteria.
Gut bacteria induce the production of plant VOCs to attract/ repel natural enemies of insects
Plants respond to herbivore attacks by emitting specific blends of herbivore-induced plant volatiles (HIPVs), which can not only directly affect the herbivores but also indirectly impact them by attracting their natural enemies92,93. Without surprise, the induction of the indirect herbivore-defense mechanism can be affected by insect gut bacteria. For instance, bacteria present in the honeydew of the rice brown planthopper (Nilaparvata lugens; BPH) strongly elicit indirect defenses in rice, and the release of volatile organic compounds from the leaves serve as attractants for natural enemies of the herbivores. Almost all of the microbes isolated from the honeydew were previously reported to be gut symbiotic bacteria of BPH72. On the contrary, the bacterial symbiont Hamiltonella defensa has been shown to reduce the recruitment of parasitic wasps and improve pea aphid (Acyrthosiphon pisum) fitness by reducing the amount of volatile compounds produced by plants94.
It seems that whether the gut bacteria-stimulated chemical compounds attract or repel natural enemies depends on the insect, natural enemy, and plant species involved, and this complex interaction mediated by gut bacteria is worth further investigation. Regardless of whether the gut-bacteria-mediated HIPVs attract or repel natural enemies, we should carefully consider the ecological significance of gut bacteria possessing these functions in further research.
Gut bacteria influence the insect–plant relationship by modulating the third trophic level species within the ecological network
As mentioned earlier, the insect–plant interaction process involves the participation of other organisms from different trophic levels, either through gut bacteria-mediated HIPVs or through intentional interventions, or other means. Interestingly, the involvement of organisms from another trophic level can alter the existing pairwise interactions between insects and plants (Fig. 3). For example, when pathogenic microorganisms are employed for insect pest control, gut bacteria may play a synergistic or antagonistic role in influencing the virulence of the microbial insecticide. Additionally, different gut microbiota can stimulate plants to produce varied HIPVs and further elicit varying behavioral responses in insect natural enemies, thereby altering the insect–plant interaction dynamics.
a Some gut bacteria show synergistic effects with entomopathogens to kill insects; b gut bacteria in several insects possess antagonistic effects against pathogens; c the VOCs produced by insects’ gut bacteria can attract natural enemies; d VOCs produced by gut bacteria of insects may also repel their natural enemies.
Gut bacteria alter the virulence of microbial insecticides
Microbial insecticides, which typically consist of entomopathogenic bacteria, fungi, or viruses95, have developed various pathogenic factors and toxins in the evolutionary arms race with host insects, making them powerful weapons to control insect pests. However, the pathogenicity of these pathogenic microorganisms can be easily influenced by a range of biological factors. Among them, gut bacteria often become involved in the process by promoting or inhibiting the infection. For instance, a previous study discovered that Beauveria bassiana infection in bark beetle D. valens LeConte caused dysbiosis of gut microbiota and overgrowth of the bacterium Erwinia sp. in the gut, thereby accelerating beetle mortality12. Another study showed that the Cry1Ac protoxin produced by Bacillus thuringiensis (Bt) induced a dynamic change in the midgut microbiota of the diamondback moth P. xylostella, and that the Bt Cry1Ac protoxin interacts with the gut microbiota to accelerate the mortality of larvae96. In contrast, the loss of gut microbiota significantly decreased the Bt susceptibility of P. xylostella larvae96. Similar insecticidal mechanisms of synergistic action between pathogenic microorganisms and gut bacteria were also reported in the European gypsy moth (Lymantria dispar asiatica) and the tobacco hornworm (Manduca sexta)97,98.
Apart from the synergistic effect between pathogens and insect associated bacteria interactions, antagonistic effects between these two agents were also widely reported. Following Nosema bombycis infection, the abundance of Enterococcus in the silkworm Bombyx mori’s gut increased, and Enterococcus faecalis LX10 reduced the N. bombycis spore germination rate and the infection efficiency in vitro and in vivo99. In the spruce beetle, D. rufipennis bacteria present in oral secretions inhibited the growth of entomopathogenic fungi100. Similarly, gut bacteria of the locust Schistocerca gregaria and Scarabs (Holotrichia oblita, Holotrichia parallela, and Anomala corpulenta) have also been demonstrated to exert antimicrobial activity similar to the above study101,102. Indeed, the interactions between pathogenic microorganisms and gut bacteria do not adhere to a consistent pattern of synergy or antagonism. Even the same bacterial species, such as Enterococcus faecalis, can exert completely opposite effects when confronted with different pathogens within various host insects. A better understanding of pathogenic microorganisms–insect–gut bacteria interactions is critical for the development of more effective microbial insecticide.
Gut bacteria produce VOCs to attract or repel natural enemies of the host
Insect natural enemies, as another important trophic level similar to entomopathogens, can also intervene and alter the insect–plant relationships103. Natural enemies of herbivorous insects, including predators and parasitoids, typically locate their concealed prey in structurally complex environments using volatile chemical cues104. In most cases, the behavior of natural enemies is mediated by plant or host volatiles, which can either repel or attract them. Studies have shown that volatile compounds produced by gut bacteria in insects can also affect the behavior of natural enemies. For instance, researchers have demonstrated that volatile compounds produced by bacteria in an aphid (Acyrthosiphon pisum) honeydew can attract predators and certain chemicals produced by Staphylococcus sciuri have been identified as attractants and ovipositional stimulants for the predator hoverfly (Episyrphus balteatus)14. However, other studies have shown a negative relationship between honeydew bacteria and the attraction of the aphid parasitoid Aphidius colemani105. Additionally, Thibout et al. found that the volatiles used by the parasitoid Diadromus pulchellus to locate its host, the leek moth (Acrolepiopsis assectella), are produced by the bacteria developing in the frass of larvae106.
Insect gut bacteria often serve as a reservoir, being present in the honeydew and feces of the insects through excretion, feeding, and other activities. The aforementioned bacteria, which have the ability to influence natural enemies, are closely related to the gut bacteria and can be broadly classified as such. Moreover, as previously discussed, gut bacteria can produce volatile compounds that influence various behaviors of insects, including feeding and oviposition. It is plausible to consider that over the course of long-term evolution, insect natural enemies may have acquired the ability to perceive and interpret these “chemical messages” to locate and target their prey. Therefore, besides exploring whether natural enemies can decode the chemical information produced by insect gut bacteria, we should also pay attention to how insects fine-tune their gut microbiota to prevent natural enemies from deciphering this information.
The leverage of using gut bacteria for pest management
Entomopathogenic bacteria and their toxins have been successfully developed and utilized against a wide range of pests. However, the susceptibility of insect pests to microbial insecticides varies, and some pests have developed resistance to pathogenic bacteria107,108,109, necessitating the development of more active insecticidal bacteria. In this context, gut bacteria have received increasing attention as potential sources of insecticidal bacteria (Fig. 4 and Table 2). Besides, with the advancement of modern molecular biology techniques, microbial control technology has expanded beyond the use of a single bacteria, with RNAi, sterile insect technology, and paratransgenesis integrated with gut bacteria for pest control (Fig. 4 and Table 2).
The vast potential of gut microbiota for pest management.
Direct insecticidal effects and potential of gut bacteria
Gut bacteria become virulent under specific physiological or environmental conditions or due to disruption of normal microbial composition71,79,110. Two main mechanisms of insect-killing by gut bacteria include toxin-induced starvation and sepsis caused by the microbiome111. Enterobacter cloacae, for example, can induce pathogenicity in its host the cotton leafworm S. litura by causing starvation and disrupting the normal gut microbiota when fed orally112. In another instance, the presence of Enterococcus faecalis in the midgut of the tobacco hornworm M. sexta larvae does not cause obvious disease, but it dies quickly when injected directly into the larval hemolymph98. Other pathogenic gut bacteria include Serratia marcescens, Bacillus licheniformis, Pseudomonas aeruginosa, Proteus vulgaris, Alcaligenes faecalis, and Planococcus sp.113. As all of these bacteria have the potential to kill pests, they can be considered as potential biological control agents for insects in pest management.
Sterile insect technology (SIT) involves mass-rearing and δ-irradiation sterilization of male insects, which are then released into the target area to compete with wild males for mating with wild females, and has been successfully used in pest control of Trypetidae114,115,116,117. Surprisingly, studies have shown that the mating competitiveness of sterile medfly male (C. capitata) obtained by radiation treatment in the field was significantly reduced, with a significant decrease in the abundance of Klebiella in the male gut and an increase in the conditional pathogenic Pseudomonas118. Feeding K. xytoca was able to enhance the mating competitiveness of infertile males118. Therefore, by manipulating the gut microbiota through certain methods (e.g., increasing Klebsiella or reducing Pseudomonas abundance), we may improve the competitive ability of sterile medfly males, thus leading to better pest control.
Collectively, given the direct role of gut bacteria in enhancing the adaptability of insects119,120,121, manipulating the insect gut microbiome has the potential to either increase or decrease insect fitness. We can utilize this knowledge for pest control purposes. Previous research has been conducted to leverage plant-mediated expression of antimicrobial peptides as a potential strategy to regulate plant-associated microbiota122. This method holds promise for regulating the gut microbiota of herbivorous insects, wherein plants could be engineered to express antimicrobial compounds or facilitate the transmission of beneficial microorganisms based on pest control requirements. However, further comprehensive studies are necessary to assess their efficacy and safety in practical applications.
Indirect insecticidal effects and potential of gut bacteria
Apart from directly influencing the fitness of herbivorous, insect gut bacteria can also produce or induce plant VOCs to attract other pests or natural enemies10,14,72. However, despite their potential, few examples of using VOCs associated with gut bacteria for pest control currently exist, e.g., while the protective effects of the multifunctional pheromone (verbenone) on Pinus contorta trees from the bark beetle D. valens have been well-documented, the discovery of its synthesis by gut bacteria came at a later stage123,124. We emphasize the consideration of VOCs associated with gut microbes in integrated pest management strategies, based on two primary reasons. Firstly, research in this area has the potential to reveal novel VOCs for pest management. Secondly, it provides an opportunity to utilize insect gut bacteria as “fermentation factories” to produce these VOCs.
Symbiont-mediated RNAi (SMR), which involves genetically engineering gut symbionts to continuously produce and deliver dsRNA within pests, has been successfully applied to a variety of pests125,126,127. For example, Taracena et al. genetically modified the symbiotic bacterium (Rhodococcus rhodnii) of a blood-sucking bug Rhodnius prolixus to express dsRNA targeting antioxidant function, resulting in a reduction in its oviposition rate128. Similar examples of modifying insect gut bacteria have also been reported in honeybees129. As orally ingested dsRNA must pass through the gut and enter epithelial cells and/or hemolymph to function, there is a high likelihood of interaction between gut bacteria and dsRNA. This hypothesis was confirmed in a study on a leaf beetle P. versicolora. Ingestion of dsRNA by P. versicolora led to the disturbance of gut microbiota, and the degradation products of dsRNA by insects preferentially promoted the growth of insect pathogenic bacteria, thereby increasing the insecticidal efficiency of RNAi79. These results indicate that it is the degradation products of dsRNA, rather than the knock-down of the targeted gene, that influenced the gut microbiota. Moreover, the altered microbiota subsequently led to different RNAi effects. Although there are currently limited studies on the combination of gut bacteria and RNAi for pest management, this provides a promising approach for developing pest control mechanisms based on gut bacteria.
The fields of synthetic biology and symbiotic insect bacteria have merged to create a new pest control strategy called paratransgenesis. This technique involves using symbiotic bacteria as gene expression vectors to introduce target genes into insects130. The expression of foreign genes interferes with pathogen development or insect fitness traits (e.g., proteotoxoids) for pest control purposes131,132. Unlike host-dependent endosymbiosis, gut bacteria are often culturable and easier to manipulate genetically and therefore more suitable as transgenic vectors. They can be easily reintroduced to host insects by oral ingestion and spread through the environment by horizontal transfer9. Another advantage of using gut bacteria as transgenic vectors is that they have the ability to colonize a variety of different insects, and can be passed horizontally and vertically from one generation to another133. For example, Serratia sp. (AS1) and Pantoea agglomerans, which are present in the gut of Anopheles mosquitoes, have been genetically engineered to secrete anti-Plasmodium effector protein and feeding the recombinant strain can inhibit the development of Plasmodium falciparum in mosquitoes134,135. Recently, the feasibility of paratransgenesis has also been demonstrated in agricultural pests. Serratia symbiotica CWBI-2.3 T, a culturable enteric-associated bacterium isolated from the black bean aphid and which can be genetically engineered136. Although paratransgenesis has not been extensively used in agriculture and forestry pest control, it holds promise as a potential approach for developing more scientifically based pest control methods.
Future perspectives and challenges
Here, we discuss the diverse functions of gut bacteria in the context of insect–plant interactions, which includes altering insect adaptability to host plants, influencing insects’ perference on different plants, regulating plant growth and defense, changing pathogenic virulence of microorganisms, and attracting/repelling natural enemies. Thus, gut bacteria is crucial for the development of novel tools in pest control methods. With advances in RNAi and paratransgenesis, the use of gut bacteria for pest control has become more diverse. Moreover, compared to chemical pesticides, biological control agents are less likely to cause environmental pollution. Therefore, rational utilization of these multispecies cascading interactions can be considered as an eco-friendly and novel approach for pest control.
Despite the promising potential of using gut bacteria for pest control, significant challenges still exist in their application. Firstly, our understanding of the role of gut bacteria in insect–plant interactions and behind mechanisms is currently incomplete due to the complexity of these interactions. Obtaining axenic insects is a major challenge in studying these interactions, as only a small fraction of agricultural and forestry pests have been successfully reared under axenic conditions. Axenic rearing approaches allow for the deconstruction and reconstruction of insect–plant–microbe interactions to identify the functions of specific gut bacteria in insect pest–plant interactions137,138. Secondly, the vast majority of gut bacteria are unculturable, which makes functional and practical studies difficult. Although recent advances in large-scale culturing methods, particularly culturomics, have made it possible to culture some gut bacteria, the challenge of large-scale cultivation and application remains139,140.Thirdly, in addition to gut bacteria, insect–plant interaction in the field often involves multiple other species, and conclusions based on laboratory studies may be varied when applied in real-world conditions, particularly with the addition of abiotic factors such as climate change. Therefore, further research is needed to integrate the specific mechanisms by which insect, plant, and gut microbial communities interact under different climatic and ecological variables. This is undoubtedly a significant challenge for researchers.
Additionally, when considering the use of other biological technologies such as RNAi or SIT in conjunction with gut bacteria or targeting the pest–gut bacteria interaction for pest control, it is necessary to take into account the efficiency and cost of these techniques51. Similar to the bottleneck encountered in developing RNAi-based insect pest control technologies, gut bacteria, and RNAi-based insect control technology also require consideration of insect uptake, degradation of dsRNA, and mode of delivery141,142. Furthermore, the large-scale production of dsRNA and its application in the field is a cumbersome and expensive process143. Similarly, SIT technology requires a substantial population of males to compete with wild males, making field application difficult and costly144.
Despite these challenges, recent advances in macrogenomics and transcriptomics have greatly increased our understanding of the functions of gut bacteria in insect–plant interactions145, providing a theoretical basis for the development of new pest management strategies. In the face of rampant pest resistance to chemical pesticides, Bt toxin, and dsRNA, it is strategically important to innovate and develop new pest management strategies that target the symbiotic relationship between pests and their gut microbiota, or gut bacteria-based pest management. Therefore, further research on the molecular mechanisms of gut bacteria in insect–plant interactions is not only of great scientific value but also has far-reaching practical implications.
References
Stork, N. E. How many species of insects and other terrestrial arthropods are there on earth? Annu. Rev. Entomol. 63, 31–45 (2018).
Basit, A. et al. Do microbial protein elicitors PeaT1 obtained from alternaria tenuissima and PeBL1 from Brevibacillus laterosporus enhance defense response against tomato aphid (Myzus persicae)? Saudi J. Biol. Sci. 28, 3242–3248 (2021).
Deutsch, C. A. et al. Increase in crop losses to insect pests in a warming climate. Science 361, 916–919 (2018).
Harwood, J. D. & Parajulee, M. N. Global impact of biological invasions: transformation in pest management approaches. Biol. Invasions 12, 2855–2856 (2010).
Yang, Y., Wu, N. & Wang, C. Toxicity of the pyrethroid bifenthrin insecticide. Environ. Chem. Lett. 16, 1377–1391 (2018).
Thompson, D. A. et al. A critical review on the potential impacts of neonicotinoid insecticide use: current knowledge of environmental fate, toxicity, and implications for human health. Environ. Sci. Process Impacts 22, 1315–1346 (2020).
Siddiqui, J. A. et al. Role of insect gut microbiota in pesticide degradation: a review. Front. Microbiol. 13, 870462 (2022).
Paniagua Voirol, L. R. et al. Bacterial symbionts in Lepidoptera: their diversity, transmission, and impact on the host. Front. Microbiol. 9, 556 (2018).
Engel, P. & Moran, N. A. The gut microbiota of insects-diversity in structure and function. FEMS Microbiol. Rev. 37, 699–735 (2013).
He, M. et al. Gut bacteria induce oviposition preference through ovipositor recognition in fruit fly. Commun. Biol. 5, 973 (2022).
Hansen, A. K. & Moran, N. A. The impact of microbial symbionts on host plant utilization by herbivorous insects. Mol. Ecol. 23, 1473–1496 (2014).
Xu, L. et al. Gut microbiota in an invasive bark beetle infected by a pathogenic fungus accelerates beetle mortality. J. Pest Sci. 92, 343–351 (2018).
Chen, C. et al. Volatile DMNT directly protects plants against Plutella xylostella by disrupting the peritrophic matrix barrier in insect midgut. Elife 10, e63938 (2021).
Leroy, P. D. et al. Microorganisms from aphid honeydew attract and enhance the efficacy of natural enemies. Nat. Commun. 2, 348 (2011).
Zhang, X., Zhang, F. & Lu, X. Diversity and functional roles of the gut microbiota in lepidopteran insects. Microorganisms 10, 1234 (2022).
Jaffar, S., Ahmad, S. & Lu, Y. Contribution of insect gut microbiota and their associated enzymes in insect physiology and biodegradation of pesticides. Front. Microbiol. 13, 979383 (2022).
Salem, H. & Kaltenpoth, M. Beetle-bacterial symbioses: endless forms most functional. Annu. Rev. Entomol. 67, 201–219 (2022).
Raghu, S., Clarke, A. R. & Bradley, J. Microbial mediation of fruit fly-host plant interactions: is the host plant the “centre of activity”? Oikos 97, 319–328 (2002).
Frago, E., Dicke, M. & Godfray, H. C. Insect symbionts as hidden players in insect-plant interactions. Trends Ecol. Evol. 27, 705–711 (2012).
Gripenberg, S., Mayhew, P. J., Parnell, M. & Roslin, T. A meta-analysis of preference-performance relationships in phytophagous insects. Ecol. Lett. 13, 383–393 (2010).
Gadenne, C., Barrozo, R. B. & Anton, S. Plasticity in insect olfaction: to smell or not to smell? Annu. Rev. Entomol. 61, 317–333 (2016).
Zhu, H. et al. Fitness consequences of oviposition choice by an herbivorous insect on a host plant colonized by an endophytic entomopathogenic fungus. J. Pest Sci. 96, 745–758 (2023).
Tack, A. J. M., Dicke, M. & Bennett, A. Plant pathogens structure arthropod communities across multiple spatial and temporal scales. Funct. Ecol. 27, 633–645 (2013).
De Vries, E. J., Vos, R. A., Jacobs, G. & Breeuwer, H. A. J. Western flower thrips (Thysanoptera: Thripidae) preference for thrips-damaged leaves over fresh leaves enables uptake of symbiotic gut bacteria. Eur. J. Entomol. 103, 779–786 (2006).
Xu, L. et al. Sexual variation of bacterial microbiota of Dendroctonus valens guts and frass in relation to verbenone production. J. Insect. Physiol. 95, 110–117 (2016).
Lu, M., Hulcr, J. & Sun, J. The role of symbiotic microbes in insect invasions. Annu. Rev. Ecol. Evol. Syst. 47, 487–505 (2016).
Corinne, V., Bastien, C., Emmanuelle, J. & Heidy, S. Trees and insects have microbiomes: consequences for forest health and management. Curr. For. Rep. 7, 81–96 (2021).
Watanabe, H. & Tokuda, G. Cellulolytic systems in insects. Annu. Rev. Entomol. 55, 609–632 (2010).
Adams, A. S. et al. Cellulose-degrading bacteria associated with the invasive woodwasp Sirex noctilio. ISME J. 5, 1323–1331 (2011).
Barbosa, K. L. et al. Bacterial cellulase from the intestinal tract of the sugarcane borer. Int. J. Biol. Macromol. 161, 441–448 (2020).
Arfah, R. A. et al. Isolation and characterization of Soil Termites (Macrotermes gilvus) cellulolytic bacteria and activity determination of cellulase enzyme on newsprint substrates. J. Phys. Conf. Ser. 1341, 032037 (2019).
Dantur, K. I., Enrique, R., Welin, B. & Castagnaro, A. P. Isolation of cellulolytic bacteria from the intestine of Diatraea saccharalis larvae and evaluation of their capacity to degrade sugarcane biomass. AMB Express 5, 15 (2015).
Baharuddin, M. Cellulase enzyme activity of bacillus Circulans from Larvae Cossus Cossus in Lignocellulosic Substrat. Am. J. Biomed. Life Sci. 4, 21 (2016).
Ceja-Navarro, J. A. et al. Gut anatomical properties and microbial functional assembly promote lignocellulose deconstruction and colony subsistence of a wood-feeding beetle. Nat. Microbiol. 4, 864–875 (2019).
Kirsch, R. et al. Metabolic novelty originating from horizontal gene transfer is essential for leaf beetle survival. Proc. Natl Acad. Sci. USA 119, e2205857119 (2022).
Salem, H. et al. Symbiont digestive range reflects host plant breadth in herbivorous beetles. Curr. Biol. 30, 2875–2886.e2874 (2020).
Ma, X., Wu, Y. & Zhang, G. Formation pattern and regulatory mechanisms of pollen wall in Arabidopsis. J. Plant Physiol. 260, 153388 (2021).
Blackmore, S., Wortley, A. H., Skvarla, J. J. & Rowley, J. R. Pollen wall development in flowering plants. N. Phytol. 174, 483–498 (2007).
Engel, P., Martinson, V. G. & Moran, N. A. Functional diversity within the simple gut microbiota of the honey bee. Proc. Natl Acad. Sci. USA 109, 11002–11007 (2012).
Zhou, H. et al. Screening and identification of lignin-degrading bacteria in termite gut and the construction of LiP-expressing recombinant Lactococcus lactis. Micro. Pathog. 112, 63–69 (2017).
Suman, S. K. et al. Investigation of lignin biodegradation by Trabulsiella sp. isolated from termite gut. Int. Biodeterior. Biodegrad. 112, 12–17 (2016).
Tsegaye, B., Balomajumder, C. & Roy, P. Isolation and characterization of novel lignolytic, cellulolytic & hemicellulolytic bacteria from wood-feeding termite Cryptotermes brevis. Int. Microbiol. 22, 29–39 (2019).
Scully, E. D. et al. Functional genomics and microbiome profiling of the Asian longhorned beetle (Anoplophora glabripennis) reveal insights into the digestive physiology and nutritional ecology of wood feeding beetles. BMC Genomics 15, 1096 (2014).
Scully, E. D. et al. Host-plant induced changes in microbial community structure and midgut gene expression in an invasive polyphage (Anoplophora glabripennis). Sci. Rep. 8, 9620 (2018).
Mattson, W. J. Herbivory in relation to plant nitrogen content. Annu. Rev. Ecol. Syst. 11, 119–161 (1980).
Jha, B., Singh, N. P. & Mishra, A. Proteome profiling of seed storage proteins reveals the nutritional potential of Salicornia brachiata Roxb., an extreme halophyte. J. Agric. Food Chem. 60, 4320–4326 (2012).
Behar, A., Yuval, B. & Jurkevitch, E. Enterobacteria-mediated nitrogen fixation in natural populations of the fruit fly Ceratitis capitata. Mol. Ecol. 14, 2637–2643 (2005).
Ceja-Navarro, J. A. et al. Compartmentalized microbial composition, oxygen gradients and nitrogen fixation in the gut of Odontotaenius disjunctus. ISME J. 8, 6–18 (2014).
Ren, X. et al. Gut symbiotic bacteria are involved in nitrogen recycling in the tephritid fruit fly Bactrocera dorsalis. BMC Biol. 20, 201 (2022).
Hu, Y. et al. Herbivorous turtle ants obtain essential nutrients from a conserved nitrogen-recycling gut microbiome. Nat. Commun. 9, 964 (2018).
Rupawate, P. S. et al. Role of gut symbionts of insect pests: a novel target for insect-pest control. Front. Microbiol. 14, 1146390 (2023).
Ohbayashi, T. et al. Comparative cytology, physiology and transcriptomics of Burkholderia insecticola in symbiosis with the bean bug Riptortus pedestris and in culture. ISME J. 13, 1469–1483 (2019).
Bisch, G. et al. Genome evolution of Bartonellaceae symbionts of ants at the opposite ends of the trophic scale. Genome Biol. Evol. 10, 1687–1704 (2018).
Stauber, E. J. et al. Turning the ‘mustard oil bomb’ into a ‘cyanide bomb’: aromatic glucosinolate metabolism in a specialist insect herbivore. PLoS ONE 7, e35545 (2012).
Li, X., Schuler, M. A. & Berenbaum, M. R. Molecular mechanisms of metabolic resistance to synthetic and natural xenobiotics. Annu. Rev. Entomol. 52, 231–253 (2007).
Hammer, T. J. & Bowers, M. D. Gut microbes may facilitate insect herbivory of chemically defended plants. Oecologia 179, 1–14 (2015).
Mason, C. J. Complex relationships at the intersection of insect gut microbiomes and plant defenses. J. Chem. Ecol. 46, 793–807 (2020).
Berasategui, A. et al. Gut microbiota of the pine weevil degrades conifer diterpenes and increases insect fitness. 26, 4099–4110 (2017).
Zhang, S. et al. Soil-derived bacteria endow Camellia weevil with more ability to resist plant chemical defense. Microbiome 10, 97 (2022).
Ceja-Navarro, J. A. et al. Gut microbiota mediate caffeine detoxification in the primary insect pest of coffee. Nat. Commun. 6, 7618 (2015).
Javal, M. et al. Does host plant drive variation in microbial gut communities in a recently shifted pest? Microb. Ecol. 86, 636–646 (2022).
Lv, D. et al. Comparison of gut bacterial communities of fall armyworm (Spodoptera frugiperda) reared on different host plants. Int. J. Mol. Sci. 22, 11266 (2021).
Meng, L., Xia, C., Jin, Z. & Zhang, H. Investigation of gut bacterial communities of Asian Citrus Psyllid (Diaphorina citri) reared on different host plants. Insects 13, 694 (2022).
Ge, S. X. et al. Gut bacteria associated with Monochamus saltuarius (Coleoptera: Cerambycidae) and their possible roles in host plant adaptations. Front. Microbiol. 12, 687211 (2021).
Xu, L. et al. Pine Defensive monoterpene alpha-pinene influences the feeding behavior of Dendroctonus valens and its gut bacterial community Structure. Int. J. Mol. Sci. 17, 1734 (2016).
War, A. R. et al. Mechanisms of plant defense against insect herbivores. Plant Signal. Behav. 7, 1306–1320 (2012).
Kariyat, R. R. et al. Non-glandular trichomes of Solanum carolinense deter feeding by Manduca sexta caterpillars and cause damage to the gut peritrophic matrix. Proc. Biol. Sci. 284, 20162323 (2017).
Konno, K. & Mitsuhashi, W. The peritrophic membrane as a target of proteins that play important roles in plant defense and microbial attack. J. Insect Physiol. 117, 103912 (2019).
Mohan, S. et al. Degradation of the S. frugiperda peritrophic matrix by an inducible maize cysteine protease. J. Insect Physiol. 52, 21–28 (2006).
Mason, C. J. et al. Plant defenses interact with insect enteric bacteria by initiating a leaky gut syndrome. Proc. Natl Acad. Sci. USA 116, 15991–15996 (2019).
Di Lelio, I. et al. A soil fungus confers plant resistance against a phytophagous insect by disrupting the symbiotic role of its gut microbiota. Proc. Natl Acad. Sci. USA 120, e2216922120 (2023).
Wari, D. et al. Honeydew-associated microbes elicit defense responses against brown planthopper in rice. J. Exp. Bot. 70, 1683–1696 (2019).
Acevedo, F. E. et al. Fall armyworm-associated gut bacteria modulate plant defense responses. Mol. Plant Microbe Interact. 30, 127–137 (2017).
Wang, J. et al. Helicoverpa zea gut-associated bacteria indirectly induce defenses in tomato by triggering a salivary elicitor(s). N. Phytol. 214, 1294–1306 (2017).
Hameeda, B., Rupela, O. P., Reddy, G. & Satyavani, K. Application of plant growth-promoting bacteria associated with composts and macrofauna for growth promotion of Pearl millet (Pennisetum glaucum L.). Biol. Fertil. Soils 43, 221–227 (2006).
Ruiu, L. Plant-growth-promoting bacteria (PGPB) against insects and other agricultural pests. Agronomy 10, 861 (2020).
Stegelmeier, A. A., Rose, D. M., Joris, B. R. & Glick, B. R. The use of PGPB to promote plant hydroponic growth. Plants (Basel) 11, 2783 (2022).
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).
Xu, L. et al. Synergistic action of the gut microbiota in environmental RNA interference in a leaf beetle. Microbiome 9, 98 (2021).
Indiragandhi, P., Anandham, R., Madhaiyan, M. & Sa, T. M. Characterization of plant growth-promoting traits of bacteria isolated from larval guts of diamondback moth Plutella xylostella (lepidoptera: plutellidae). Curr. Microbiol. 56, 327–333 (2008).
He, Z., Webster, S. & He, S. Y. Growth-defense trade-offs in plants. Curr. Biol. 32, R634–R639 (2022).
Krawczyk, K. et al. Insect gut bacteria promoting the growth of tomato plants (Solanum lycopersicum L.). Int. J. Mol. Sci. 23, 13548 (2022).
Acevedo, F. E. et al. Phytohormones in fall armyworm saliva modulate defense responses in plants. J. Chem. Ecol. 45, 598–609 (2019).
Felton, G. W. & Tumlinson, J. H. Plant-insect dialogs: complex interactions at the plant-insect interface. Curr. Opin. Plant Biol. 11, 457–463 (2008).
Erb, M., Meldau, S. & Howe, G. A. Role of phytohormones in insect-specific plant reactions. Trends Plant Sci. 17, 250–259 (2012).
Glazebrook, J. Contrasting mechanisms of defense against biotrophic and necrotrophic pathogens. Annu. Rev. Phytopathol. 43, 205–227 (2005).
Thaler, J. S., Humphrey, P. T. & Whiteman, N. K. Evolution of jasmonate and salicylate signal crosstalk. Trends Plant Sci. 17, 260–270 (2012).
Costarelli, A. et al. Salicylic acid induced by herbivore feeding antagonizes jasmonic acid mediated plant defenses against insect attack. Plant Signal. Behav. 15, 1704517 (2020).
Chung, S. H. et al. Herbivore exploits orally secreted bacteria to suppress plant defenses. Proc. Natl Acad. Sci. USA 110, 15728–15733 (2013).
Yamasaki, Y. et al. Phytohormone-dependent plant defense signaling orchestrated by oral bacteria of the herbivore Spodoptera litura. N. Phytol. 231, 2029–2038 (2021).
Wielkopolan, B., Frackowiak, P., Wieczorek, P. & Obrepalska-Steplowska, A. The impact of oulema melanopus-associated bacteria on the wheat defense response to the feeding of their insect hosts. Cells 11, 2342 (2022).
Dicke, M. & Baldwin, I. T. The evolutionary context for herbivore-induced plant volatiles: beyond the ‘cry for help’. Trends Plant Sci. 15, 167–175 (2010).
Du, Y. W. et al. Chinese cabbage changes its release of volatiles to defend against Spodoptera litura. Insects 13, 73 (2022).
Frago, E. et al. Symbionts protect aphids from parasitic wasps by attenuating herbivore-induced plant volatiles. Nat. Commun. 8, 1860 (2017).
Qiao, J., Du, Y., Yu, J. & Guo, J. MicroRNAs as potential biomarkers of insecticide exposure: a review. Chem. Res. Toxicol. 32, 2169–2181 (2019).
Li, S. et al. Gut microbiota mediate Plutella xylostella susceptibility to Bt Cry1Ac protoxin is associated with host immune response. Environ. Pollut. 271, 116271 (2021).
Bai, J. et al. Gut bacterial microbiota of Lymantria dispar asiatica and its involvement in Beauveria bassiana infection. J. Invertebr. Pathol. 197, 107897 (2023).
Mason, K. L. et al. From commensal to pathogen: translocation of Enterococcus faecalis from the midgut to the hemocoel of Manduca sexta. mBio 2, e00065–00011 (2011).
Zhang, X. et al. The gut commensal bacterium Enterococcus faecalis LX10 contributes to defending against Nosema bombycis infection in Bombyx mori. Pest Manag. Sci. 78, 2215–2227 (2022).
Cardoza, Y. J., Klepzig, K. D. & Raffa, K. F. Bacteria in oral secretions of an endophytic insect inhibit antagonistic fungi. Ecol. Entomol. 31, 636–645 (2006).
Dillon, R. J. & Charnley, A. K. Inhibition of Metarhizium anisopliae by the gut bacterial flora of the desert locust: characterisation of antifungal toxins. Can. J. Microbiol. 34, 1075–1082 (1988).
Shan, Y. et al. Cultivable gut bacteria of scarabs (Coleoptera: Scarabaeidae) inhibit Bacillus thuringiensis multiplication. Environ. Entomol. 43, 612–616 (2014).
Tariq, M., Wright, D. J., Bruce, T. J. & Staley, J. T. Drought and root herbivory interact to alter the response of above-ground parasitoids to aphid infested plants and associated plant volatile signals. PLoS ONE 8, e69013 (2013).
Wang, G. Y. et al. Identification and expression analysis of putative chemoreception genes from Cyrtorhinus lividipennis (Hemiptera: Miridae) antennal transcriptome. Sci. Rep. 8, 12981 (2018).
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).
Thibout, E. et al. Origin and identification of bacteria which produce kairomones in the frass of Acrolepiopsis assectella (Lep., Hyponomeutoidea). Experientia 51, 1073–1075 (1995).
Yang, J. et al. Profiling of microRNAs in midguts of plutella xylostella provides novel insights into the bacillus thuringiensis resistance. Front. Genet. 12, 739849 (2021).
Pickett, B. R., Gulzar, A., Ferre, J. & Wright, D. J. Bacillus thuringiensis Vip3Aa toxin resistance in Heliothis virescens (Lepidoptera: Noctuidae). Appl. Environ. Microbiol. 83, e03506–16 (2017).
Jurat-Fuentes, J. L., Heckel, D. G. & Ferre, J. Mechanisms of resistance to insecticidal proteins from bacillus thuringiensis. Annu. Rev. Entomol. 66, 121–140 (2021).
Ma, M. et al. Metabolic and immunological effects of gut microbiota in leaf beetles at the local and systemic levels. Integr. Zool. 16, 313–323 (2021).
Sina Rahme, B. et al. The fliR gene contributes to the virulence of S. marcescens in a Drosophila intestinal infection model. Sci. Rep. 12, 3068 (2022).
Thakur, A., Dhammi, P., Saini, H. S. & Kaur, S. Pathogenicity of bacteria isolated from gut of Spodoptera litura (Lepidoptera: Noctuidae) and fitness costs of insect associated with consumption of bacteria. J. Invertebr. Pathol. 127, 38–46 (2015).
Osborn, F. et al. Pathogenic effects of bacteria isolated from larvae of Hylesia metabus Crammer (Lepidoptera: Saturniidae). J. Invertebr. Pathol. 80, 7–12 (2002).
Fezza, T. J., Follett, P. A. & Shelly, T. E. Effect of the timing of pupal irradiation on the quality and sterility of oriental fruit flies (Diptera: Tephritidae) for use in Sterile Insect Technique. Appl. Entomol. Zool. 56, 443–450 (2021).
Aceituno-Medina, M. et al. Mass rearing, quality parameters & bioconversion in Drosophila suzukii (Diptera: Drosophilidae) for sterile insect technique purposes. J. Econ. Entomol. 113, 1097–1104 (2020).
Ikegawa, Y., Ito, K., Himuro, C. & Honma, A. Sterile males and females can synergistically suppress wild pests targeted by sterile insect technique. J. Theor. Biol. 530, 110878 (2021).
Rathnayake, D. N., Lowe, E. C., Rempoulakis, P. & Herberstein, M. E. Effect of natural predators on Queensland fruit fly, Bactrocera tryoni (Froggatt) (Diptera: Tephritidae) control by sterile insect technique (SIT). Pest Manag. Sci. 75, 3356–3362 (2019).
Ben Ami, E., Yuval, B. & Jurkevitch, E. Manipulation of the microbiota of mass-reared Mediterranean fruit flies Ceratitis capitata (Diptera: Tephritidae) improves sterile male sexual performance. ISME J. 4, 28–37 (2010).
Liu, G., Cao, L. & Han, R. Plant quercetin degradation by gut bacterium Raoultella terrigena of ghost moth Thitarodes xiaojinensis. Front. Microbiol. 13, 1079550 (2022).
Gandotra, S. et al. Screening of nutritionally important gut bacteria from the Lepidopteran insects through qualitative enzyme assays. Proc. Natl Acad. Sci. India Sect. B: Biol. Sci. 88, 329–337 (2016).
Li, J. et al. Gut Microbiota of Ostrinia nubilalis larvae degrade maize cellulose. Front. Microbiol. 13, 816954 (2022).
Weinhold, A. et al. Antimicrobial peptide expression in a wild tobacco plant reveals the limits of host-microbe-manipulations in the field. Elife 7, e28715 (2018).
Xu, L. et al. Gut-associated bacteria of Dendroctonus valens and their involvement in verbenone production. Micro. Ecol. 70, 1012–1023 (2015).
Gillette, N. E. et al. Verbenone-releasing flakes protect individual Pinus contorta trees from attack by Dendroctonus ponderosae and Dendroctonus valens (Coleoptera: Curculionidae, Scolytinae). Agric. For. Entomol. 8, 243–251 (2006).
Dyson, P., Figueiredo, M., Andongma, A. A. & Whitten, M. M. A. Symbiont-mediated RNA interference (SMR): using symbiotic bacteria as vectors for delivering RNAi to insects. Methods Mol. Biol. 2360, 295–306 (2022).
Li, T. et al. Facultative symbionts are potential agents of symbiont-mediated RNAi in aphids. Front. Microbiol. 13, 1020461 (2022).
Lariviere, P. J. et al. Honey bee functional genomics using symbiont-mediated RNAi. Nat. Protoc. 18, 902–928 (2023).
Taracena, M. L. et al. Genetically modifying the insect gut microbiota to control Chagas disease vectors through systemic RNAi. PLoS Negl. Trop. Dis. 9, e0003358 (2015).
Lang, H. et al. Engineered symbiotic bacteria interfering Nosema redox system inhibit microsporidia parasitism in honeybees. Nat. Commun. 14, 2778 (2023).
Huang, W. et al. Combining transgenesis with paratransgenesis to fight malaria. Elife 11, e77584 (2022).
Coutinho-Abreu, I. V., Zhu, K. Y. & Ramalho-Ortigao, M. Transgenesis and paratransgenesis to control insect-borne diseases: current status and future challenges. Parasitol. Int. 59, 1–8 (2010).
Lopez-Ordonez, T. et al. Cultivable bacterial diversity in the gut of the chagas disease vector Triatoma dimidiata: identification of possible bacterial candidates for a paratransgenesis approach. Front. Ecol. Evol. 5, 174 (2018).
Ratcliffe, N. A. et al. Overview of paratransgenesis as a strategy to control pathogen transmission by insect vectors. Parasit. Vectors 15, 112 (2022).
Wang, S. et al. Fighting malaria with engineered symbiotic bacteria from vector mosquitoes. Proc. Natl Acad. Sci. USA 109, 12734–12739 (2012).
Wang, S. et al. Driving mosquito refractoriness to Plasmodium falciparum with engineered symbiotic bacteria. Science 357, 1399–1402 (2017).
Elston, K. M. et al. Engineering a culturable serratia symbiotica strain for aphid paratransgenesis. Appl. Environ. Microbiol. 87, e02245–20 (2020).
Wu, J. et al. Axenic and gnotobiotic insect technologies in research on host-microbiota interactions. Trends Microbiol. 31, 858–871 (2023).
Ma, M. et al. Preparing and rearing axenic insects with tissue cultured seedlings for host-gut microbiota interaction studies of the leaf beetle. J. Vis. Exp. 176, e63195 (2021).
Diakite, A. et al. Extensive culturomics of 8 healthy samples enhances metagenomics efficiency. PLoS ONE 14, e0223543 (2019).
Hongoh, Y. Toward the functional analysis of uncultivable, symbiotic microorganisms in the termite gut. Cell Mol. Life Sci. 68, 1311–1325 (2011).
Zhu, K. Y. & Palli, S. R. Mechanisms, applications & challenges of insect RNA interference. Annu. Rev. Entomol. 65, 293–311 (2020).
Yan, S., Ren, B., Zeng, B. & Shen, J. Improving RNAi efficiency for pest control in crop species. Biotechniques 68, 283–290 (2020).
Hough, J. et al. Strategies for the production of dsRNA biocontrols as alternatives to chemical pesticides. Front. Bioeng. Biotechnol. 10, 980592 (2022).
Simon, S., Otto, M. & Engelhard, M. Synthetic gene drive: between continuity and novelty: crucial differences between gene drive and genetically modified organisms require an adapted risk assessment for their use. EMBO Rep. 19, e45760 (2018).
Xiong, W. Intestinal microbiota in various animals. Integr. Zool. 17, 331–332 (2022).
Li, Y. et al. Genome sequencing of gut symbiotic Bacillus velezensis LC1 for bioethanol production from bamboo shoots. Biotechnol. Biofuels 13, 34 (2020).
Hatefi, A., Makhdoumi, A., Asoodeh, A. & Mirshamsi, O. Characterization of a bi-functional cellulase produced by a gut bacterial resident of Rosaceae branch borer beetle, Osphranteria coerulescens (Coleoptera: Cerambycidae). Int. J. Biol. Macromol. 103, 158–164 (2017).
Sheng, P. et al. Isolation, screening & optimization of the fermentation conditions of highly cellulolytic bacteria from the hindgut of Holotrichia parallela larvae (Coleoptera: Scarabaeidae). Appl. Biochem. Biotechnol. 167, 270–284 (2012).
Zhang, Q. et al. Enhanced biogas production by ligninolytic strain enterobacter hormaechei KA3 for anaerobic digestion of corn straw. Energies 14, 2990 (2021).
Evangelista, D. E., de Paula, F. F., Rodrigues, A. & Henrique-Silva, F. Pectinases from Sphenophorus levis Vaurie, 1978 (Coleoptera: Curculionidae): putative accessory digestive enzymes. J. Insect Sci. 15, 168 (2015).
Alonso-Pernas, P. et al. In vivo isotopic labeling of symbiotic bacteria involved in cellulose degradation and nitrogen recycling within the gut of the forest cockchafer (Melolontha hippocastani). Front. Microbiol. 8, 1970 (2017).
Salem, H. et al. Vitamin supplementation by gut symbionts ensures metabolic homeostasis in an insect host. Proc. Biol. Sci. 281, 20141838 (2014).
Berasategui, A. et al. The gut microbiota of the pine weevil is similar across Europe and resembles that of other conifer-feeding beetles. Mol. Ecol. 25, 4014–4031 (2016).
Zhang, S. et al. The gut microbiota in Camellia Weevils are influenced by plant secondary metabolites and contribute to Saponin Degradation. mSystems 5, e00692–19 (2020).
Cheng, C. et al. Bacterial microbiota protect an invasive bark beetle from a pine defensive compound. Microbiome 6, 132 (2018).
Shukla, S. P. & Beran, F. Gut microbiota degrades toxic isothiocyanates in a flea beetle pest. Mol. Ecol. 29, 4692–4705 (2020).
Consuelo, G. C.-M. et al. The gut bacteria symbionts from the monophagous insect Acrobasis nuxvorellaproduce tannase for the digestion of Carya illinoinensis tannins. J. Asia-Pac. Entomol. 25, 102005 (2022).
Leite-Mondin, M. et al. The gut microbiota composition of Trichoplusia ni is altered by diet and may influence its polyphagous behavior. Sci. Rep. 11, 5786 (2021).
Wang, W. et al. Glutamicibacter halophytocola-mediated host fitness of potato tuber moth on Solanaceae crops. Pest Manag. Sci. 78, 3920–3930 (2022).
Mason, C. J., Peiffer, M., Hoover, K. & Felton, G. Tomato chemical defenses intensify corn earworm (Helicoverpa zea) mortality from opportunistic bacterial pathogens. J. Chem. Ecol. 49, 313–324 (2023).
Aggarwal, C. et al. A modified semi-synthetic diet for bioassay of non-sporeforming entomopathogenic bacteria against Spodoptera litura. Biocontrol Sci. Technol. 24, 1202–1205 (2014).
Devi, S., Saini, H. S. & Kaur, S. Insecticidal and growth inhibitory activity of gut microbes isolated from adults of Spodoptera litura (Fab.). BMC Microbiol. 22, 71 (2022).
Zhang, P., Zhao, Q., Ma, X. & Ma, L. Pathogenicity of Serratia marcescens to hazelnut weevil (Curculio dieckmanni). J. For. Res. 32, 409–417 (2020).
Vesga, P. et al. Phylogenetically closely related pseudomonads isolated from arthropods exhibit differential insect-killing abilities and genetic variations in insecticidal factors. Environ. Microbiol. 23, 5378–5394 (2021).
Mashtoly, T. A. et al. Phylogenetic characteristics of novel Bacillus weihenstephanensis and Pseudomonas sp. to desert locust, Schistocerca gregaria Forskål (Orthoptera: Acrididae). Egyptian J. Biol. Pest Cont. 29, 85 (2019).
Luo, J. et al. Variation of gut microbiota caused by an imbalance diet is detrimental to bugs’ survival. Sci. Total Environ. 771, 144880 (2021).
Msaad Guerfali, M. et al. Evaluation of Providencia rettgeri pathogenicity against laboratory Mediterranean fruit fly strain (Ceratitis capitata). PLoS ONE 13, e0196343 (2018).
Kyritsis, G. A. et al. Enterobacter sp. AA26 gut symbiont as a protein source for Mediterranean fruit fly mass-rearing and sterile insect technique applications. BMC Microbiol. 19, 288 (2019).
Zhang, Q. et al. Manipulation of gut symbionts for improving the sterile insect technique: quality parameters of Bactrocera dorsalis (Diptera: Tephritidae) genetic sexing strain males after feeding on bacteria-enriched diets. J. Econ. Entomol. 114, 560–570 (2021).
Li, Y. et al. Bt GS57 interaction with gut microbiota accelerates spodoptera exigua mortality. Front. Microbiol. 13, 835227 (2022).
Lei, X., Zhang, F. & Zhang, J. Gut microbiota accelerate the insecticidal activity of plastid-expressed bacillus thuringiensis Cry3Bb to a leaf beetle, Plagiodera versicolora. Microbiol. Spectr. 11, e0504922 (2023).
Elston, K. M., Maeda, G. P., Perreau, J. & Barrick, J. E. Addressing the challenges of symbiont-mediated RNAi in aphids. PeerJ 11, e14961 (2023).
Acknowledgements
This research was supported by the National Natural Science Foundation of China (31971663), the Natural Science Foundation of Hubei Province (2022CFA061) and the Young Elite Scientists Sponsorship Program by CAST (2020QNRC001).
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L.X. and S.Z. conceptualized and planned the review. L.X., S.Z., and Y.Z. wrote and revised the paper. Y.Z. and L.X. drew pictures. All of the authors have read and approved the manuscript.
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Zhang, Y., Zhang, S. & Xu, L. The pivotal roles of gut microbiota in insect plant interactions for sustainable pest management. npj Biofilms Microbiomes 9, 66 (2023). https://doi.org/10.1038/s41522-023-00435-y
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DOI: https://doi.org/10.1038/s41522-023-00435-y
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