Abstract
The insect’s microbiota is well acknowledged as a “hidden” player influencing essential insect traits. The gut microbiome of butterflies and moths (Lepidoptera) has been shown to be highly variable between and within species, resulting in a controversy on the functional relevance of gut microbes in this insect order. Here, we aim to (i) review current knowledge on the composition of gut microbial communities across Lepidoptera and (ii) elucidate the drivers of the variability in the lepidopteran gut microbiome and provide an overview on (iii) routes of transfer and (iv) the putative functions of microbes in Lepidoptera. To find out whether Lepidopterans possess a core gut microbiome, we compared studies of the microbiome from 30 lepidopteran species. Gut bacteria of the Enterobacteriaceae, Bacillaceae, and Pseudomonadaceae families were the most widespread across species, with Pseudomonas, Bacillus, Staphylococcus, Enterobacter, and Enterococcus being the most common genera. Several studies indicate that habitat, food plant, and age of the host insect can greatly impact the gut microbiome, which contributes to digestion, detoxification, or defense against natural enemies. We mainly focus on the gut microbiome, but we also include some examples of intracellular endosymbionts. These symbionts are present across a broad range of insect taxa and are known to exert different effects on their host, mostly including nutrition and reproductive manipulation. Only two intracellular bacteria genera (Wolbachia and Spiroplasma) have been reported to colonize reproductive tissues of Lepidoptera, affecting their host’s reproduction. We explore routes of transmission of both gut microbiota and intracellular symbionts and have found that these microbes may be horizontally transmitted through the host plant, but also vertically via the egg stage. More detailed knowledge about the functions and plasticity of the microbiome in Lepidoptera may provide novel leads for the control of lepidopteran pest species.
Introduction
Bacterial symbionts inhabiting insects can significantly impact the biology of their host (). These symbionts can be distinguished as intra- and extracellular based on whether they live within insect cells, or colonize the lumen or lining of insect cavities and body surface (; ; ). Symbionts are considered as primary or secondary, depending on whether they are needed by the host to survive or provide non-essential benefits ().
Obligatory symbionts are commonly harbored in specialized cells (bacteriocytes) and play important roles for nutrition in certain insect groups, particularly in phloem feeding taxa. For example, intracellular Buchnera bacteria associated with aphids provide essential amino acids and vitamins (; ). The benefits provided by secondary symbionts, on the other hand, are often context-dependent. In aphids, for example, secondary symbionts can provide a range of ecological benefits including resistance to pathogens and parasitoids, and heat tolerance, but they can be costly under benign conditions (). Some Wolbachia strains living intracellularly can manipulate host reproduction to favor their own spread in the population (; ), while others can be facultative () or even become obligatory in certain species ().
The composition and robustness of gut bacterial communities varies extensively across the animal kingdom ranging from more than 1,000 phylotypes in humans (), over several hundreds in termites (; ), and a few tens in lepidopterans (; ; ), to an almost complete absence of bacteria in aphid guts (; ). In insects, the best studied and most diverse gut bacterial communities are those belonging to groups feeding on wood, decaying matter, or detritus such as termites, cockroaches, crickets, and some beetles (; ). Gut bacterial communities often deliver metabolic benefits to their hosts by the provision of digestive enzymes and production of vitamins, thus improving nutrient uptake on deficient diets (; ; ; ). Furthermore, they can provide protection against pathogens () and support detoxification of pesticides or harmful plant secondary metabolites (; ; ).
Lepidoptera comprise the second most diverse insect order with some of the most devastating agricultural pests worldwide (). Yet, clear evidence for bacterial associates playing a fundamental role in lepidopteran biology is scarce. The functional role of the gut microbiome of Lepidoptera has been challenged by a recent study reporting that caterpillars harbor no or only few resident bacteria when compared to other insect orders (). The authors of this study argue that this is probably due to caterpillars being rough environments for bacterial colonization, because they possess an unusually alkaline gut with a rapid food passage of approximately two hours. In addition lepidopterans undergo a holometabolous metamorphosis which entirely re-shapes their body structures (). In spite of this harsh environment for the gut microbiota, several studies have shown that bacteria do affect essential physiological functions in Lepidoptera, i.e., facilitation of nutrient acquisition and digestion (; ; ), overcoming plant anti-herbivore defenses (; ), or strengthening of immune responses for protection against pathogens ().
Out of the 157,424 recognized lepidopteran species (), <0.1% have been screened for bacterial associates, which reveals that our knowledge on bacterial associates in Lepidoptera is still limited. Many of these studies focused on specific endosymbionts known to be widespread in arthropods, such as Wolbachia and Spiroplasma. Many other studies are mostly descriptive and focused on larvae, while only a few have addressed the potential impact on their host traits. These studies screened bacteria from specimens of several of the ∼43 lepidopteran superfamilies (), i.e., Hepialoidea, Yponomeutoidea, Tortricoidea, Cossoidea, Papilionidea, Gelechioidea, Pyralioidea, Depranoidea, Noctuoidea, Geometroidea, and Bombycoidea (Supplementary Table 1). Knowledge on whether certain bacteria taxa are persistent across the Lepidoptera order is limited, as well as information on how Lepidoptera transfer symbiotic bacteria among individuals of a population and between generations.
Our review aims to find hints for answering these questions and points to future studies by screening the current literature on microbial associates in Lepidoptera. We surveyed the literature to assess which bacterial taxa were detected in independent studies comprising 30 different lepidopteran species, and asked which ones are ubiquitous in these taxa. We further considered potential drivers explaining the variability found in the composition of the lepidopteran gut microbiome. These drivers include ecological, morphological, and developmental traits of Lepidoptera. These features significantly impact the way by which bacterial symbionts are transmitted between individuals and through generations. Understanding the role of symbiotic bacteria in such an economically important insect order may provide novel leads for improving current integrated pest management techniques. This knowledge is also important from a fundamental perspective to understand the role that symbiotic bacteria play in helping lepidopteran larvae to cope with challenges such as diet deficiencies, host plant switches, and natural enemy attacks.
Composition of the Gut Microbiota in Lepidoptera
In order to elucidate whether some bacterial taxa are ubiquitous in the gut of Lepidoptera, we screened independent studies comprising 30 different lepidopteran species. Despite the differences in the methodology used in the different studies, such as differences in the life stage, insect diet, and screening technique (culture-based, cloning/sequencing, or high-throughput amplicon) (see Supplementary Table 1), our survey based on presence/absence shows that certain bacteria taxa are widespread across lepidopterans. Most of the detected gut bacterial families belong to the Proteobacteria phylum (42%) (Figure 1). Within this group, those families belonging to the α- and γ-Proteobacteria classes are the most common (72%) (Figure 1). Bacteria belonging to the Enterobacteriaceae, Bacillaceae, Pseudomonadaceae, Staphylococcaceae, and Enterococcaceae families are present in >60% of the screened lepidopteran species (Figure 2). At the genus level, the most widespread bacteria belong to Pseudomonas, Bacillus, Staphylococcus, Enterobacter, and Enterococcus, each being present in >70% of the studied lepidopteran species (Figure 3). Persistence of some gut bacterial species occurs regardless of the diet the insects fed upon, indicating the presence of a core community (; ; ; ; ). Despite this, various studies also indicate that the gut microbiome shows great variability across lepidopteran species and even within a species, a question that will be discussed in detail in the following section.
FIGURE 1
FIGURE 2
FIGURE 3
Drivers of Variability in Bacterial Gut Communities
The high variability of the lepidopteran gut microbiome could be promoted by different drivers, which may act alone or in concert and include the environment, insect diet, insect developmental stage, and gut physiology. Firstly, the environment where insects live affects the composition of the insects’ microbiome. Insects reared in the laboratory or collected in the field show different microbial communities even if they feed on the same host plant (e.g., ). The habitat may thus significantly affect the bacteria associated with lepidopteran species (). Secondly, diet can have a major influence on bacterial community variability. Recent studies could not () or hardly () detect any resident, host insect-specific, and food-independent bacteria in Lepidoptera. The bacterial community can therefore be expected to differ significantly between oligophagous and polyphagous species, or between herbivorous and carnivorous species. A comparative study on microbial communities associated with herbivorous and carnivorous Lycaenidae larvae, however, did not find consistent patterns in community profiles that could relate them to the diet of the insect (). By contrast, an assessment on the influence of diet and host taxonomy on gut bacterial communities across several insect orders found that, depending on the insect taxon, either factor was significant (). Insects feeding on decaying matter presented the richest communities, while bees and wasps had the lowest. While host taxonomy was an important driver of bacterial communities in hymenopterans and termites, diet was important in insects feeding on lignocellulose-derived components. Non-conclusive patterns of clustering among lepidopterans were found, based on a rather small number of species studied ().
In addition to diet and environment, the developmental stage can influence the host’s gut microbiota. Concordantly, instar-specific bacterial communities were detected in larvae of the moth Spodoptera littoralis (). As in all other holometabolous insect orders, metamorphosis in Lepidoptera entails major morphological rearrangements and is usually accompanied by a change in diet, which can have a strong impact on gut microbiota composition. While almost all lepidopteran species feed upon plant tissue during their larval stage (with a few notable carnivorous and fungivorous exceptions), the adult stage of most species feeds on nectar (). With the proviso that gut communities depend on the diet, it is not surprising that bacterial communities differ considerably between larvae and adults of the same species (; ). Nevertheless, certain taxa may persist throughout the entire insect life cycle as shown for bacteria species belonging to the families Acetobacteraceae, Moraxellaceae, Enterobacteriaceae, Enterococcaceae, Streptococcaceae, and unclassified Bacteroidetes, which dominate the gut of the larval, pupal, and adult stages of the red postman (Heliconius erato) (). Some bacteria like Enterococcus mundtii may even survive and propagate in the digestive tract of S. littoralis across its life cycle, and persist up to two consecutive generations (). Such persistence of some bacterial symbionts across the entire development is also found in other holometabolous insects that inhabit different ecological niches during the larval and adult stage, like the emerald ash borer beetle Agrilus planipennis (), the fruitfly Ceratitis capitata (), or the scarabaeid beetle Melolontha hippocastani (). Thus, while some core bacteria persist in holometabolous insects including Lepidoptera, a considerable change in the bacterial community composition from larvae to adults is common. This is probably due to the physiological changes occurring during metamorphosis, and also due to the change in diet from larvae to adults, which is particularly dramatic in Lepidoptera.
Persistence of bacteria in the gut of lepidopteran larvae is further impeded by the lack of intricate pouches-like gut structures that are known to harbor bacterial symbionts in other insect taxa (; ). A complex anatomy of the gut with a high number of pouches (diverticula, caeca) might favor the establishment of a robust bacterial community, as seen in other non-lepidopteran insects with extremely rich bacterial gut communities. For example, termites with their complex gut structures harbor highly robust gut communities that vary across the gut compartments (; ; ; ).
Main Transmission Routes of Gut Bacteria: Vertically or Horizontally?
How Lepidoptera gain and retain gut bacterial members is a largely unresolved question. It is still unclear to which extent gut symbiotic bacteria are transmitted either (a) vertically from one generation to the next or (b) horizontally between individuals directly via contact among individuals or indirectly via uptake from the diet (; ). Detection of core bacterial associates in the gut suggests a potential vertical transmission or a consistent horizontal acquisition of gut symbionts in some species, while environmental uptake of transient associates appears to be likely in others ().
Since gut bacteria live extracellularly, they are probably not transmitted inside insect oocytes, although translocation of gut bacteria to the oocytes was reported in Galleria mellonella (). However, these authors did not test whether such bacteria remain viable in the following generation. In some insect orders like Heteroptera, extracellular bacteria are added to the egg surface by the females in secretions or feces, which are later acquired by the hatching nymphs (). Transmission of extracellular symbionts via the egg stage requires that bacteria remain alive before colonizing the newly emerged larvae. This would be possible if symbionts are deposited in an inactive stage, or if active bacteria are nourished through the egg shell or egg-associated secretions. In either case, transmission via the egg stage requires that larvae take up these bacteria when hatching. Since neonate lepidopteran larvae bite through their egg shell while hatching and often fully ingest it after hatching (Figure 4), infection of neonate larvae with bacteria on the outer egg surface is possible.
FIGURE 4
The presence of insect gut bacteria associated with the eggs has been shown for some lepidopterans (; ; ; ). Yet, only for the tobacco hornworm (Manduca sexta), bacterial (Enterococcus) metabolic activity has been confirmed during the egg stage (). The fact that these bacteria are metabolically active when associated with M. sexta eggs suggests a high metabolic adaptability which allows them to survive both in association with the eggs and later in the gut of larvae (). However, metabolic activity is not strictly necessary for successful vertical transmission, since some inactive egg-associated bacteria may also re-activate their metabolism once reaching the gut of the neonate. Active bacterial growth during the egg stage may increase their chances to colonize not only the hatching neonates but also their environment (e.g., soil or host plant), which could ultimately promote horizontal transmission. It would be relevant to quantify bacterial growth during the egg incubation phase to elucidate bacterial proliferation since the beginning of insect development.
Intra- and extracellular symbionts in herbivorous insects are known to be also acquired through their host plant (; ; ; ), which is another example that indicates an enormous adaptability of the bacteria to various habitats. Despite these evidences, bacteria present on the host plant may also reflect the bacteria community present in the habitat, and studies of bacterial associates in Lepidoptera should always include the appropriate controls to disentangle whether the detected bacteria originate from the environment or are specific for the insect.
As outlined above, many gut bacteria found in Lepidoptera are ubiquitous. For example, Enterococcus has been detected in many lepidopteran species and other insect orders (; ; ; ; ). A comparative study of Heliothis virescens butterflies obtained from either the laboratory or the field revealed that Enterococcus was dominant in individuals of the laboratory strain, but absent in those from the field, suggesting that the presence of this bacterium is a laboratory artifact in this species (). Due to the high prevalence of Enterococcus in different habitats and conditions (), these bacteria might be simply acquired from the laboratory environment, and their presence on the eggs might not represent a natural condition.
With such scarce evidence of insect-specific gut bacteria being present on eggs, it remains speculative to consider eggs as a vehicle for vertical symbiont transmission in Lepidoptera. More studies are therefore needed to determine to what extent the bacterial members persist across generations to pinpoint those bacterial members that potentially share a long co-evolutionary history with their lepidopteran host. Bacterial symbionts that share a longer history with their host and that are vertically transmitted are most likely playing fundamental roles in the host biology. However, symbionts acquired de novo from the environment every generation could also be functionally important as was found in the bean bug (Riptortus pedestris), which developed insecticide resistance after acquiring insecticide-degrading bacteria (Burkholderia) from the soil ().
Gut Bacteria and Their Impact on Digestion and Nutrient Acquisition in Lepidoptera
Dietary nitrogen is particularly limiting in the diet of phloem-feeding insects which have evolved associations with amino acid- and vitamin-supplementing intracellular symbionts. Considering the high carbon-to-nitrogen ratio in leaves, it is likely that chewing insects also need to cope with limited dietary nitrogen (; ). Symbiotic bacteria may be beneficial by fixing nitrogen and converting it into physiologically relevant nitrogen-containing compounds (). Rhizobacteria fixing molecular dinitrogen into ammonium are well known to be associated with plant roots (; ). Some insects like termites harbor bacteria which also have the ability to fix dinitrogen into ammonia which is then assimilated by gut endosymbionts that biosynthesize vitamins and amino acids needed for insect development (; ; ; ; ). A study on the moth Plutella xylostella demonstrated that the same may occur in Lepidoptera, as many bacteria isolated from the gut were found to be able to fix nitrogen in vitro ().
Nitrogen present in hairs and feathers in the form of keratin, a cysteine-rich polypeptide, is the nutritional, nitrogen-rich resource of a few lepidopteran species like the clothes’ moth Tineola bisselliella. This species can break the cysteine-bonds in keratin thus making the polypeptide more accessible to proteases for digestion. Since keratinases are only known from bacteria and fungi (), it is possible that a microbial partner is involved in this digestion. Despite this, the mechanism by which T. bisselliella uses keratin is yet to be clarified, firstly because did not find any culturable bacterial colonies in the gut that could play this role, and secondly because the complete microbiome of this species has not been characterized. In addition, genes that resemble bacterial sequences encoding keratinolytic enzymes were not found in the genome of this moth in a later study ().
Plant cell wall degrading enzymes (PCWDEs) include cellulases, hemicellulases, and pectinases that originate from insects and are responsible for the uptake of carbohydrates via the break-up of plant cell walls. The genes encoding for some of these enzymes are found in crickets (Orthoptera), stick insects (Phasmatodea), cockroaches and termites (Dictyoptera), lice (Phthiraptera), aphids (Homoptera), beetles (Coleoptera), and bees and wasps (Hymenoptera) (). Phylogenetic analyses revealed that in many cases, these genes encoding PCWDEs were originally horizontally acquired from microorganisms and integrated into the host genome (; ; ,). However, the tortoise leaf beetle Cassida rubiginosa still maintains a bacterial symbiont that degrades pectin (), a ubiquitous recalcitrant component of all plant tissues. So far, evidence for PCWDEs-encoding genes or symbionts in Lepidoptera is scarce (), although found evidence for hypothetical cellulose-encoding genes in the hesperiid butterfly Lerema accius. These genes, however, were not detected in a close hesperiid relative, Achalarus lyciades (). The presence of cellulase-encoding genes in ancestral insect taxa and its lack in most of the studied Lepidoptera may suggest that lepidopteran species lost these genes and may rely on symbionts for cellulose digestion. One of these examples may include P. xylostella the gut microbiome of which has thousands of genes from six families that encode carbohydrate-active enzymes, including cellulases ().
Numerous other enzymes may be provided by bacteria inhabiting the lepidopteran gut. For example, bacteria present in caterpillars and pupae of the saturniid moths Automeris zugana and Rothschildia lebeau provide gelatinase, caseinase, lipase, esterase, and chitinase activity (). These bacterial enzymatic activities might become especially important for efficient food digestion by the host insect during periods of food shortage or after host plant shifts (; ). Since these bacteria were detected in pupae, it is also possible that they assist during metamorphosis, which is a very active event in terms of metabolic activity. For example, symbionts may provide chitinases for cuticle digestion, or aromatic amino acids for the synthesis of the new cuticle as suggested by .
Protection Against Entomopathogens by Lepidopteran Gut Bacteria
In addition to helping with nutrient acquisition, resident gut bacteria can provide protection against pathogens (; ). One way to achieve this is by outcompeting pathogens, the so-called colonization resistance (), as found in Homona magnanima whose caterpillars are more susceptible to Bacillus thuringiensis bacteria when they are reared aseptically than when they are not (). Enterococcus faecalis found in the gypsy moth is known to acidify its local environment so that it can colonize alkaline niches. This probably protects the gut against pathogenic toxins that are activated in alkaline conditions, such as those from B. thuringiensis (). The gut bacterial communities in some insects produce bactericidal substances that selectively target foreign bacteria, but they do not affect autochthonous ones (). E. mundtii, for instance, is a highly abundant bacterium in the gut of S. littoralis, which produces an antimicrobial compound against Gram-positive pathogens like Listeria, but not against resident gut bacteria ().
Some common gut bacterial inhabitants may be detrimental or beneficial depending on the community composition of the gut. One of such bacteria may be Serratia spp., a genus of bacteria commonly reported in lepidopterans and known to be pathogenic in many animals (; ). It would be relevant to evaluate if these bacteria provide benefits under certain conditions and whether they switch to a virulent phase when the structure of the bacterial community is altered. If the virulence of Serratia spp. depends on the composition of the entire insect microbial community, this would indicate a community-wide role in preventing pathogenic outbursts (). The role of gut bacteria in protecting lepidopterans against pathogens should be thus studied in a community context. For instance, while aseptic rearing in some cases results in less susceptibility to B. thuringiensis (), for Lymantria dispar it was found that the midgut community is actually required for the activity of B. thuringiensis toxin ().
Several studies have shown that exposure of insects to certain pathogens or parasites can boost the immune system of their offspring, an effect known as transgenerational immune priming (; ; ; ). For example, oral uptake of pathogenic bacteria by female caterpillars of the greater wax moth G. mellonella resulted in the increased expression of immunity-relevant genes in the caterpillars but also in the eggs laid by adult females developing from these caterpillars (). Transgenerational immune priming by orally ingested bacteria has also been observed in the moth Trichoplusia ni (). It has been proposed that such priming in Lepidoptera occurs by means of maternal transfer of bacteria or bacteria-associated compounds to the developing eggs (), which reinforces the idea that gut microbes can play an important role in insect defense against natural enemies.
Lepidopteran Gut Bacteria That Counteract Anti-herbivore Plant Defenses
Plants defend themselves by a plethora of physical and chemical weapons against insect herbivory (; ; ; ). Numerous plant secondary compounds with deterrent, anti-digestive, or toxic effects on insect herbivores are known. Some microbial symbionts can play a fundamental role in promoting the pest status of certain insect species by detoxifying plant allelochemicals (). Bacterial symbionts of insects can also influence insect–plant interactions to a greater extent than previously thought by helping their hosts manipulate the induction of plant defenses (, ; ).
A general strategy to detoxify plant lipophilic toxins is to convert them into water-soluble compounds which can easily be excreted. To achieve this, lipophilic toxins are usually functionalized (e.g., oxidized) and then conjugated with a highly polar compound, like glutathione. Bacteria in the gut of the diamondback moth (P. xylostella) are known to provide glutathione-S-transferase, an enzyme involved in this conjugation process ().
Phenolic compounds, almost ubiquitously present in plants, can impair digestion of proteins through interactions with plant proteins and insect digestive enzymes. These compounds promote the production of reactive oxygen species (ROS) when they are oxidized into quinones, especially at alkaline pH values as those present in the lepidopteran gut (; ). High concentrations of ROS, which are highly aggressive compounds interacting with almost all cellular components, may significantly damage cells (e.g., ). Bacteria in the lepidopteran gut, e.g., Enterobacter spp., can provide ROS-detoxifying enzymes like superoxide dismutase or catalase ().
Terpenes, another class of secondary metabolites that are widespread in the plant kingdom, are toxic to many insects and bacteria because they may disturb chemiosmosis when lipophilic non-oxidized terpenes promote interactions with cell membranes (). For conifer-feeding bark beetles and weevils, gut symbionts have been implicated in the detoxification of terpenes that allow the insects to subsist on the terpene-rich diets (; ; ). The gypsy moth can tolerate diets enriched with monoterpenes (; ) and this is likely due to association with Rhodococcus gut bacteria which are able to degrade monoterpenes at high alkalinity ().
Biosynthesis of protease inhibitors is a further means of defense employed by plants that impairs plant protein digestion by herbivorous insects (; ). Bacteria-derived proteases may counter-balance the inhibition of insect-derived proteases sensitive to plant protease inhibitors. For example, the velvetbean caterpillar (Anticarsia gemmatalis) feeds on soy bean, which is known to possess high amounts of protease inhibitors that act as anti-herbivory defenses (; ; ). This moth may use gut bacteria from the genera Bacillus, Enterococcus, and Staphylococcus to overcome these defenses as they have protease activity (). Characterizing gut bacterial proteases and confirming their resistance against plant-derived protease inhibitors may allow the development of analogs of protease inhibitors that target bacterial proteases, which could be used for pest control (; ).
Induction of plant defenses by chewing insects may be counteracted by orally released bacteria into plant wounds, as found in the Colorado potato beetle (Leptinotarsa decemlineata) (). So far, no lepidopteran species are known to orally release bacteria that inhibit induced plant defenses. On the contrary, a recent study by showed that gut-associated Enterococcus ludwigii of field-collected Helicoverpa zea caterpillars indirectly increase anti-herbivore defense of tomato plants attacked by this insect. As suggested by these authors, this difference in effects of coleopteran- and lepidopteran-associated bacteria on a plant’s anti-herbivore defense may be due to differences in the feeding modes between beetle and lepidopteran larvae. The latter ones release saliva into the wound, but hardly any bacteria-containing regurgitant, whereas the former ones release regurgitants that can contain oral and/or foregut bacteria. Another interesting example revealed that Enterococcus bacteria in H. zea larvae were found to promote increased release of salivary elicitors such as glucose oxidase leading to enhanced anti-herbivore defenses and a decrease in weight of H. zea larvae. Wang and co-authors suggest that field-collected H. zea larvae harbor this bacterium, because it might facilitate metabolism of plant toxins, but comes at the cost of triggering plant defenses.
In summary, although it is well known that insects have evolved many adaptations to counteract plant toxic defenses (; ; ), the composition of the bacterial community is increasingly recognized as having a great effect on how an herbivorous insect can cope with plant toxins (). Since gut bacteria can either withstand or detoxify plant toxins (), it seems logical to speculate that the bacterial community in an insect’s gut can determine whether the insect can survive by feeding on plant tissues that are marinated with potentially toxic chemical compounds ().
Intracellular Symbiotic Bacteria
Primary endosymbionts living intracellularly in specialized host cells (bacteriocytes) are well-known to establish mutualistic relationships with insects feeding on phloem, blood, or other diets with severe deficiencies in essential amino acids and/or vitamins (). However, vertically transmitted secondary endosymbionts are present across a broad range of insect taxa with diverse feeding habits, and they are known to exert different effects on their host (). Persistent maternal transmission of bacterial endosymbionts in lepidopterans has been demonstrated only for two symbionts: Wolbachia and Spiroplasma (; ; ). Several moth and butterfly species are colonized by these endosymbionts that infect the reproductive tissue and manipulate the host’s physiology to enhance their own transmission (; ; ; ; ; ).
About 80% of lepidopteran species have been estimated to be infected by Wolbachia (). A recent study detected a high diversity of Wolbachia strains in Lepidoptera comprising a total of 90 different strains (). The mean infection prevalence (proportion of infection within a population) has been reported to be about 27% in Lepidoptera (). A co-phylogenetic analysis of lepidopteran species and their associated Wolbachia strains revealed weak congruence, as closely related lepidopteran host taxa harbored distantly related Wolbachia strains (). This finding suggests horizontal transmission, not only between individuals of the same species but also between different lepidopteran groups and probably between Lepidoptera and other insect orders.
The most common effect of Wolbachia infection is reproductive as it often shifts the sex ratio in favor of females. In Lepidoptera, sex ratio distortion associated to this symbiont is achieved by (1) male-killing in Hypolimnas bolina (killing of male embryo) (; ), Acraea encedon, and A. encedana (; ,), (2) feminization of genetic males in Eurema hecabe (), Ostrinia scapulalis (), and O. furnacalis (), and (3) cytoplasmic incompatibility (gametes being unable to form viable offspring) in Cadra cautella, Ephestia kuehniella, and H. bolina (; ). In Danaus chrysippus, sex ratio distortion is caused by the intracellular bacterium Spiroplasma that induces male-killing ().
From the bacterial point of view, sex ratio distortion is an efficient strategy that promotes rapid spread in insect populations, because it results in a higher number of female offspring in Wolbachia-infected vs. uninfected females, thereby driving the infection into the population. The spread of male-killing Wolbachia in a population can be very successful, as has been shown in a population of the moth A. encedana, in which 95% of females were infected with Wolbachia ().
Although Wolbachia is mostly known for its role in sex ratio distortion, other effects are also known. Wolbachia infection can, in certain cases, lead to increased longevity and fecundity of the host. This may be caused, for example, through provisioning of riboflavin (). Wolbachia may impact on the host’s behavior and the host’s immunity against entomopathogens (), and it can even manipulate the physiology of the plant in favor of its lepidopteran host (). The leaf miner moth Phyllonorycter blancardella possesses a Wolbachia strain affecting cytokinin levels of apple tree leaf tissues. The cytokinin manipulation promotes plant nutrient mobility and delays senescence in those patches where the caterpillars feed on (). These effects result in the formation of photosynthetically active patches that allow larval development in otherwise decaying leaves (“green island phenotype”).
Bacterial Symbionts and the Control of Lepidopteran Pests
Many pests have developed resistance to a great variety of pesticides, and although insect pesticide resistance is believed to be based on the genetic repertoire of the insect, recent studies have shown the potential role of bacterial symbionts in developing such resistance (). The bean bug R. pedestris harbors Burkholderia bacteria which can degrade an organophosphate pesticide (fenitrothion), thus conferring resistance to the host (). Studying bacterial symbionts in important lepidopteran pests such as P. xylostella should be of special relevance. P. xylostella is a worldwide pest known to have developed insecticide resistance, causing 4–5 billion dollars of damage per year (; ; ). A link between insecticide resistance and abundance of certain bacteria in the larval midgut of P. xylostella was found (). The biological control agent B. thuringiensis (Bt) has been efficiently used for more than a decade against important lepidopteran pests, but resistance against the toxin produced by this microbe has also recently evolved. A potential role of gut bacteria in reducing larval mortality after exposure to Bt was reported for P. xylostella (). Also, mortality of the cotton bollworm, H. armigera, declined across generations exposed to Bt, whereas antibiotic-treated lines did not develop resistance ().
These studies suggest that gut bacteria are important in the evolution of Bt resistance, and thus elucidating the role of bacterial symbionts of lepidopterans in this context might help developing improved methods of biological control. Increasing evidence suggests that manipulation of the microbiome could reduce the abundance of pest insects in agriculture and forestry or limit disease-vectoring activity of insects (; ). However, all the manipulative strategies suggested so far might not only target the pest insect’s microbiome, but may also significantly affect the microbiota associated with other organisms present in the system. Genetic transformation of bacteria to impact the phenotype of the host (paratransgenesis) () entails the risk of horizontal gene transfer to other bacteria in the ecosystem, because of the easy exchange of DNA sequences among environmental microbiota (e.g., ). Hence, development of such techniques and their use other than in laboratory research will first require very careful ecological risk assessment studies.
Conclusions and Open Questions for Future Research
In many insect taxa, coevolution between hosts and their beneficial symbionts has been shown to broaden the ecological niches that can be colonized by the host. In Lepidoptera, such host–symbiont coevolution has not been demonstrated, because most studies have found little evidence of a core bacterial community with functional relevance in this order. However, the acquisition and transfer of some persistent bacterial members has been reported in several species, in spite of the harsh physiological conditions of the lepidopteran larval gut and the change of ecological niches between juvenile and adult stage. The high variability of the lepidopteran gut microbiome implies on the one hand that Lepidopterans do not rely on a fixed beneficial microbiota that is present in each generation. On the other hand, such variability may also imply the chance of harboring a very dynamic microbiome that allows their hosts to adapt to changing conditions including changes in abiotic conditions, food resources, and risk of natural enemy attack.
The factors leading to the evolutionary success of the highly diverse lepidopteran taxon are still unclear. According to , independence of microbes may have resulted in high diversification rates and lead to an extraordinary diversity and abundance of Lepidoptera. A lack of a vertically transmitted core microbiome that dictates host plant use, in combination with an ancient horizontal transfer of genes originating from bacteria () could be thus possible reasons for their success. The latter explanation has been shown to precede the diversification of phasmids (stick and leaf insects) (). While microbial symbiosis can provide novel ecological functions, Hammer and co-workers argue that dependence on symbionts might increase extinction risks because insects are constraint in their diet breadth and less able to switch to new food plants. It is therefore possible that independence of symbiosis might have facilitated switching to different host plants and promote diversification. In fact, the most species-rich superfamily of Lepidoptera, Noctuoidea, consists of many polyphagous species, among them numerous agricultural pests (). Each host plant switch confronts a lepidopteran individual with a novel environment and novel microbiome present on the host plant, which may lead to the symbiont community of caterpillars being often dominated by leaf-associated bacteria that are taken up from the host plant (). More studies are needed to understand the relationship between plant and lepidopteran microbiomes and their role in host plant shifts, and diversification. These studies should also consider plant-induced defenses and explore how bacteria that originate from caterpillar frass or salivary regurgitants affect plant physiology as found in beetles by .
The microbiome of moths and butterflies may not only be shaped by their interactions with plants, but also by interactions with antagonists like pathogens, predators, and parasites. Defensive symbioses are known in many animal taxa (), and it is likely that they also exist in Lepidoptera. Defensive symbionts are often facultative, and under laboratory conditions they are likely to get lost because the pressure imposed by natural enemies is lacking. Detection of defensive symbionts in natural lepidopteran populations is therefore a challenge for future research.
Studies on individuals from the field are of great significance in order to distinguish between ecologically important bacteria colonizing lepidopterans in their natural habitats and bacteria that are the product of laboratory rearing conditions (compare: , ; ). There is a strong selection pressure when animals are reared under laboratory conditions. This can result in the loss of traits, including relationships with facultative symbiotic bacteria, which may have an effect that is only beneficial in natural populations.
In a nutshell, to gain a deeper understanding of the mechanisms by which Lepidoptera-associated microbes affect host traits, ecological, microbiological, and molecular approaches are needed. This knowledge will provide fundamental insights into host–microbe interactions in one of the most speciose animal groups on the planet, and may ultimately lead to a better control of important agricultural pests.
Statements
Author contributions
LRPV designed the first draft of this review article and performed the data analyses presented in the figures. NEF and MH contributed to the concept and structure of the review, and all authors contributed to the writing, revising, and editing of the paper.
Funding
LRPV and MH are supported by a grant of the German Research Foundation (DFG-CRC 973, project B4; www.sfb973.de). EF was funded by the Regional Council of Réunion, the Departmental Council of the Région Réunion, the European Union (EAFRD), and CIRAD. NEF was supported by a Vidi grant of the Netherlands Organization for Scientific Research (NWO/TTW Vidi 14854). MK was supported by a grant of the German Research Foundation (KA2846/2-2).
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2018.00556/full#supplementary-materila
References
1
AdamsA. S.AylwardF. O.AdamsS. M.ErbilginN.AukemaB. H.CurrieC. R.et al (2013). Mountain pine beetles colonizing historical and naive host trees are associated with a bacterial community highly enriched in genes contributing to terpene metabolism.Appl. Environ. Microbiol.793468–3475. 10.1128/AEM.00068-13
2
AhmedM. Z.Araujo-JnrE. V.WelchJ. J.KawaharaA. Y. (2015). Wolbachi in butterflies and moths: geographic structure in infection frequency.Front. Zool.12:16. 10.1186/s12983-015-0107-z
3
AhmedM. Z.BreinholtJ. W.KawaharaA. Y. (2016). Evidence for common horizontal transmission of Wolbachia among butterflies and moths.BMC Evol. Biol.16:118. 10.1186/s12862-016-0660-x
4
Alonso-PernasP.Arias-CorderoE.NovoselovA.EbertC.RybakJ.KaltenpothM.et al (2017). Bacterial community and PHB-accumulating bacteria associated with the wall and specialized niches of the hindgut of the forest cockchafer (Melolontha hippocastani).Front. Microbiol.8:291. 10.3389/fmicb.2017.00291
5
AnandA. A.VennisonS. J.SankarS. G.PrabhuD. I. G.VasanP. T.RaghuramanT.et al (2010). Isolation and characterization of bacteria from the gut of Bombyx mori that degrade cellulose, xylan, pectin and starch and their impact on digestion.J. Insect Sci.10:107. 10.1673/031.010.10701
6
AppelH. M. (1993). Phenolics in ecological interactions: the importance of oxidation.J. Chem. Ecol.191521–1552. 10.1007/BF00984895
7
Arias-CorderoE.PingL.ReichwaldK.DelbH.PlatzerM.BolandW. (2012). Comparative evaluation of the gut microbiota associated with the below-and above-ground life stages (larvae and beetles) of the forest cockchafer, Melolontha hippocastani.PLoS One7:e51557. 10.1371/journal.pone.0051557
8
BarbehennR. V.Peter ConstabelC. (2011). Tannins in plant-herbivore interactions.Phytochemistry721551–1565. 10.1016/j.phytochem.2011.01.040
9
BaumannP.MoranN. A.BaumannL. (1997). The evolution and genetics of aphid endosymbionts.Bioscience4712–20. 10.2307/1313002
10
BaxterS. W.ZhaoJ. Z.GahanL. J.SheltonA. M.TabashnikB. E.HeckelD. G. (2005). Novel genetic basis of field-evolved resistance to Bt toxins in Plutella xylostella.Insect Mol. Biol.14327–334. 10.1111/j.1365-2583.2005.00563.x
11
BerasateguiA.SalemH.PaetzC.SantoroM.GershenzonJ.KaltenpothM.et al (2017). Gut microbiota of the pine weevil degrades conifer diterpenes and increases insect fitness.Mol. Ecol.264099–4110. 10.1111/mec.14186
12
BerasateguiA.ShuklaS.SalemH.KaltenpothM. (2016). Potential applications of insect symbionts in biotechnology.Appl. Microbiol. Biotechnol.1001567–1577. 10.1007/s00253-015-7186-9
13
BiJ.FeltonG. (1995). Foliar oxidative stress and insect herbivory: primary compounds, secondary metabolites, and reactive oxygen species as components of induced resistance.J. Chem. Ecol.211511–1530. 10.1007/BF02035149
14
BodyM.KaiserW.DubreuilG.CasasJ.GironD. (2013). Leaf-miners co-opt microorganisms to enhance their nutritional environment.J. Chem. Ecol.39969–977. 10.1007/s10886-013-0307-y
15
BrinkmannN.MartensR.TebbeC. C. (2008). Origin and diversity of metabolically active gut bacteria from laboratory-bred larvae of Manduca sexta (Sphingidae, Lepidoptera, Insecta).Appl. Environ. Microbiol.747189–7196. 10.1128/AEM.01464-08
16
BroderickN. A.RaffaK. F.GoodmanR. M.HandelsmanJ. (2004). Census of the bacterial community of the gypsy moth larval midgut by using culturing and culture-independent methods.Appl. Environ. Microbiol.70293–300. 10.1128/AEM.70.1.293-300.2004
17
BroderickN. A.RaffaK. F.HandelsmanJ. (2006). Midgut bacteria required for Bacillus thuringiensis insecticidal activity.Proc. Natl. Acad. Sci. U.S.A.10315196–15199. 10.1073/pnas.0604865103
18
BruneA. (2014). Symbiotic digestion of lignocellulose in termite guts.Nat. Rev. Microbiol.12168–180. 10.1038/nrmicro3182
19
BruneA.DietrichC. (2015). The gut microbiota of termites: digesting the diversity in the light of ecology and evolution.Annu. Rev. Microbiol.69145–166. 10.1146/annurev-micro-092412-155715
20
CarliniC. R.Grossi-de-SáM. F. (2002). Plant toxic proteins with insecticidal properties. A review on their potentialities as bioinsecticides.Toxicon401515–1539. 10.1016/S0041-0101(02)00240-4
21
Caspi-FlugerA.InbarM.Mozes-DaubeN.KatzirN.PortnoyV.BelausovE.et al (2012). Horizontal transmission of the insect symbiont Rickettsia is plant-mediated.Proc. R. Soc. Lond. B Biol. Sci.2791791–1796. 10.1098/rspb.2011.2095
22
ChadwickJ. S.CaldwellS. S.ChadwickP. (1990). Adherence patterns and virulence for Galleria mellonella larvae of isolates of Serratia marcescens.J. Invertebr. Pathol.55133–134. 10.1016/0022-2011(90)90044-7
23
CharlatS.HornettE. A.FullardJ. H.DaviesN.RoderickG. K.WedellN.et al (2007). Extraordinary flux in sex ratio.Science317:214. 10.1126/science.1143369
24
ChenB.TehB. S.SunC.HuS.LuX.BolandW.et al (2016). Biodiversity and activity of the gut microbiota across the life history of the insect herbivore Spodoptera littoralis.Sci. Rep.6:29505. 10.1038/srep29505
25
ChrostekE.Pelz-StelinskiK.HurstG. D. D.HughesG. L. (2017). Horizontal transmission of intracellular insect symbionts via plants.Front. Microbiol.8:2237. 10.3389/fmicb.2017.02237
26
ChungS. H.RosaC.ScullyE. D.PeifferM.TookerJ. F.HooverK.et al (2013). Herbivore exploits orally secreted bacteria to suppress plant defenses.Proc. Natl. Acad. Sci. U.S.A.11015728–15733. 10.1073/pnas.1308867110
27
ColmanD. R.ToolsonE. C.Takacs-VesbachC. (2012). Do diet and taxonomy influence insect gut bacterial communities?Mol. Ecol.215124–5137. 10.1111/j.1365-294X.2012.05752.x
28
CongQ.BorekD.OtwinowskiZ.GrishinN. V. (2015). Skipper genome sheds light on unique phenotypic traits and phylogeny.BMC Genomics16:639. 10.1186/s12864-015-1846-0
29
CoxC. R.GilmoreM. S. (2007). Native microbial colonization of Drosophila melanogaster and its use as a model of Enterococcus faecalis pathogenesis.Infect. Immun.751565–1576. 10.1128/IAI.01496-06
30
CrewtherW. G.McquadeQ. A. (1955). The intestinal microflora of the clothes moth larva Tineola bisselliella in relation to wool digestion.J. Gen. Microbiol.12311–313. 10.1099/00221287-12-2-311
31
da Silva FortunatoF.de Almeida OliveiraM. G.BrumanoM. H. N.SilvaC. H. O.GuedesR. N. C.MoreiraM. A. (2007). Lipoxygenase-induced defense of soybean varieties to the attack of the velvetbean caterpillar (Anticarsia gemmatalis Hübner).J. Pest Sci.80241–247. 10.1007/s10340-007-0179-4
32
DespresL.DavidJ. P.GalletC. (2007). The evolutionary ecology of insect resistance to plant chemicals.Trends Ecol. Evol.22298–307. 10.1016/j.tree.2007.02.010
33
DillonR. J.DillonV. M. (2004). The gut bacteria of insects: nonpathogenic interactions.Annu. Rev. Entomol.4971–92. 10.1146/annurev.ento.49.061802.123416
34
DoblerS.PetschenkaG.PankokeH. (2011). Coping with toxic plant compounds-the insect’s perspective on iridoid glycosides and cardenolides.Phytochemistry721593–1604. 10.1016/j.phytochem.2011.04.015
35
DouglasA. (1988). On the source of sterols in the green peach aphid, Myzus persicae, reared on holidic diets.J. Insect Physiol.34403–408. 10.1016/0022-1910(88)90110-2
36
DouglasA. (1998). Nutritional interactions in insect-microbial symbioses: aphids and their symbiotic bacteria Buchnera.Annu. Rev. Entomol.4317–37. 10.1146/annurev.ento.43.1.17
37
DouglasA. E. (2007). Symbiotic microorganisms: untapped resources for insect pest control.Trends Biotechnol.25338–342. 10.1016/j.tibtech.2007.06.003
38
DouglasA. E. (2009). The microbial dimension in insect nutritional ecology.Funct. Ecol.2338–47. 10.1111/j.1365-2435.2008.01442.x
39
DouglasA. E. (2015). Multiorganismal insects: diversity and function of resident microorganisms.Annu. Rev. Entomol.6017–34. 10.1146/annurev-ento-010814-020822
40
DysonE. A.HurstG. D. (2004). Persistence of an extreme sex-ratio bias in a natural population.Proc. Natl. Acad. Sci. U.S.A.1016520–6523. 10.1073/pnas.0304068101
41
EngelP.MoranN. A. (2013). The gut microbiota of insects - diversity in structure and function.FEMS Microbiol. Rev.37699–735. 10.1111/1574-6976.12025
42
FisherK.PhillipsC. (2009). The ecology, epidemiology and virulence of Enterococcus.Microbiology1551749–1757. 10.1099/mic.0.026385-0
43
FlorezL. V.BiedermannP. H.EnglT.KaltenpothM. (2015). Defensive symbioses of animals with prokaryotic and eukaryotic microorganisms.Nat. Prod. Rep.32904–936. 10.1039/c5np00010f
44
FlórezL. V.ScherlachK.GaubeP.RossC.SitteE.HermesC.et al (2017). Antibiotic-producing symbionts dynamically transition between plant pathogenicity and insect-defensive mutualism.Nat. Commun.8:15172. 10.1038/ncomms15172
45
FragoE.DickeM.GodfrayH. C. J. (2012). Insect symbionts as hidden players in insect–plant interactions.Trends Ecol. Evol.27705–711. 10.1016/j.tree.2012.08.013
46
FragoE.MalaM.WeldegergisB. T.YangC.McLeanA.GodfrayH. C. J.et al (2017). Symbionts protect aphids from parasitic wasps by attenuating herbivore-induced plant volatiles.Nat. Commun.8:1860. 10.1038/s41467-017-01935-0
47
FreitakD.HeckelD. G.VogelH. (2009). Dietary-dependent trans-generational immune priming in an insect herbivore.Proc. R. Soc. B Biol. Sci.2762617–2624. 10.1098/rspb.2009.0323
48
FreitakD.SchmidtbergH.DickelF.LochnitG.VogelH.VilcinskasA. (2014). The maternal transfer of bacteria can mediate trans-generational immune priming in insects.Virulence5547–554. 10.4161/viru.28367
49
FrohlichJ.KoustianeC.KampferP.Rossello-MoraR.ValensM.BerchtoldM.et al (2007). Occurrence of rhizobia in the gut of the higher termite Nasutitermes nigriceps.Syst. Appl. Microbiol.3068–74. 10.1016/j.syapm.2006.03.001
50
FujiiY.KageyamaD.HoshizakiS.IshikawaH.SasakiT. (2001). Transfection of Wolbachia in Lepidoptera: the feminizer of the adzuki bean borer Ostrinia scapulalis causes male killing in the Mediterranean flour moth Ephestia kuehniella.Proc. R. Soc. Lond. B Biol. Sci.268855–859. 10.1098/rspb.2001.1593
51
GentaF. A.DillonR. J.TerraW. R.FerreiraC. (2006). Potential role for gut microbiota in cell wall digestion and glucoside detoxification in Tenebrio molitor larvae.J. Insect Physiol.52593–601. 10.1016/j.jinsphys.2006.02.007
52
GershenzonJ.DudarevaN. (2007). The function of terpene natural products in the natural world.Nat. Chem. Biol.3408–414. 10.1038/nchembio.2007.5
53
GrenierA.-M.DuportG.PagèsS.CondemineG.RahbéY. (2006). The phytopathogen Dickeya dadantii (Erwinia chrysanthemi 3937) is a pathogen of the pea aphid.Appl. Environ. Microbiol.721956–1965. 10.1128/AEM.72.3.1956-1965.2006
54
HammerT. J.JanzenD. H.HallwachsW.JaffeS. P.FiererN. (2017). Caterpillars lack a resident gut microbiome.Proc. Natl. Acad. Sci. U.S.A.1149641–9646. 10.1073/pnas.1707186114
55
HammerT. J.McMillanW. O.FiererN. (2014). Metamorphosis of a butterfly-associated bacterial community.PLoS One9:e86995. 10.1371/journal.pone.0086995
56
HansenA. K.MoranN. A. (2014). The impact of microbial symbionts on host plant utilization by herbivorous insects.Mol. Ecol.231473–1496. 10.1111/mec.12421
57
HeckelD. G. (2014). “Insect detoxification and sequestration strategies,” inAnnual Plant Reviews: Plant Insect InteractionsVol. 47edsVoelckelC.JanderG. (Chichester: Wiley-Blackwell) 77–114. 10.1002/9781118829783.ch3
58
HilkerM.FatourosN. E. (2016). Resisting the onset of herbivore attack: plants perceive and respond to insect eggs.Curr. Opin. Plant Biol.329–16. 10.1016/j.pbi.2016.05.003
59
HirokiM.KatoY.KamitoT.MiuraK. (2002). Feminization of genetic males by a symbiotic bacterium in a butterfly, Eurema hecabe (Lepidoptera: Pieridae).Naturwissenschaften89167–170. 10.1007/s00114-002-0303-5
60
HongohY.DeevongP.InoueT.MoriyaS.TrakulnaleamsaiS.OhkumaM.et al (2005). Intra- and interspecific comparisons of bacterial diversity and community structure support coevolution of gut microbiota and termite host.Appl. Environ. Microbiol.716590–6599. 10.1128/AEM.71.11.6590-6599.2005
61
HornettE. A.DuplouyA. M.DaviesN.RoderickG. K.WedellN.HurstG. D.et al (2008). You can’t keep a good parasite down: evolution of a male-killer suppressor uncovers cytoplasmic incompatibility.Evolution621258–1263. 10.1111/j.1558-5646.2008.00353.x
62
HosokawaT.KogaR.KikuchiY.MengX. Y.FukatsuT. (2010). Wolbachia as a bacteriocyte-associated nutritional mutualist.Proc. Natl. Acad. Sci. U.S.A.107769–774. 10.1371/journal.pbio.1000002
63
HughesJ.VoglerA. P. (2006). Gene expression in the gut of keratin-feeding clothes moths (Tineola) and keratin beetles (Trox) revealed by subtracted cDNA libraries.Insect Biochem. Mol. Biol.36584–592. 10.1016/j.ibmb.2006.04.007
64
HuntJ.CharnleyA. (1981). Abundance and distribution of the gut flora of the desert locust, Schistocerca gregaria.J. Invertebr. Pathol.38378–385. 10.1016/0022-2011(81)90105-1
65
HurstG. D. D. (2017). Extended genomes: symbiosis and evolution.Interface Focus7:20170001. 10.1098/rsfs.2017.0001
66
IndiragandhiP.AnandhamR.MadhaiyanM.SaT. M. (2008). Characterization of plant growth-promoting traits of bacteria isolated from larval guts of diamondback moth Plutella xylostella (Lepidoptera: Plutellidae).Curr. Microbiol.56327–333. 10.1007/s00284-007-9086-4
67
International Silkworm Genome Consortium (2008). The genome of a lepidopteran model insect, the silkworm Bombyx mori.Insect Biochem. Mol. Biol.381036–1045. 10.1016/j.ibmb.2008.11.004
68
IshiiK.AdachiT.HamamotoH.SekimizuK. (2014). Serratia marcescens suppresses host cellular immunity via the production of an adhesion-inhibitory factor against immunosurveillance cells.J. Biol. Chem.2895876–5888. 10.1074/jbc.M113.544536
69
JigginsF. M.HurstG. D. D.DolmanC. E.MajerusM. E. N. (2000a). High-prevalence male-killing Wolbachia in the butterfly Acraea encedana.J. Evol. Biol.13495–501. 10.1046/j.1420-9101.2000.00180.x
70
JigginsF. M.HurstG. D.JigginsC. D.MajerusM. (2000b). The butterfly Danaus chrysippus is infected by a male-killing Spiroplasma bacterium.Parasitology120439–446.
71
JigginsF. M.HurstG. D.MajerusM. E. (1998). Sex ratio distortion in Acraea encedon (Lepidoptera: Nymphalidae) is caused by a male-killing bacterium.Heredity8187–91. 10.1046/j.1365-2540.1998.00357.x
72
JongsmaM. A.BolterC. (1997). The adaptation of insects to plant protease inhibitors.J. Insect Physiol.43885–895. 10.1016/S0022-1910(97)00040-1
73
KageyamaD.NishimuraG.HoshizakiS.IshikawaY. (2002). Feminizing Wolbachia in an insect, Ostrinia furnacalis (Lepidoptera: Crambidae).Heredity88444–449. 10.1038/sj.hdy.6800077
74
KaiserW.HuguetE.CasasJ.ComminC.GironD. (2010). Plant green-island phenotype induced by leaf-miners is mediated by bacterial symbionts.Proc. R. Soc. Lond. B Biol. Sci.2772311–2319. 10.1098/rspb.2010.0214
75
KesslerA.BaldwinI. T. (2002). Plant responses to insect herbivory: the emerging molecular analysis.Annu. Rev. Plant Biol.53299–328. 10.1146/annurev.arplant.53.100301.135207
76
KikuchiY.HayatsuM.HosokawaT.NagayamaA.TagoK.FukatsuT. (2012). Symbiont-mediated insecticide resistance.Proc. Natl. Acad. Sci. U.S.A.1098618–8622. 10.1073/pnas.1200231109
77
KirschR.GramzowL.TheissenG.SiegfriedB. D.Ffrench-ConstantR. H.HeckelD. G.et al (2014). Horizontal gene transfer and functional diversification of plant cell wall degrading polygalacturonases: key events in the evolution of herbivory in beetles.Insect Biochem. Mol. Biol.5233–50. 10.1016/j.ibmb.2014.06.008
78
KneipC.LockhartP.VossC.MaierU.-G. (2007). Nitrogen fixation in eukaryotes–new models for symbiosis.BMC Evol. Biol.7:55. 10.1186/1471-2148-7-55
79
LauzonC.McCombsS.PotterS.PeabodyN. (2009). Establishment and vertical passage of Enterobacter (Pantoea) agglomerans and Klebsiella pneumoniae through all life stages of the Mediterranean fruit fly (Diptera: Tephritidae).Ann. Entomol. Soc. Am.10285–95. 10.1603/008.102.0109
80
LehmanR. M.LundgrenJ. G.PetzkeL. M. (2009). Bacterial communities associated with the digestive tract of the predatory ground beetle, Poecilus chalcites, and their modification by laboratory rearing and antibiotic treatment.Microb. Ecol.57349–358. 10.1007/s00248-008-9415-6
81
LiS.-J.AhmedM. Z.LvN.ShiP.-Q.WangX.-M.HuangJ.-L.et al (2016). Plant–mediated horizontal transmission of Wolbachia between whiteflies.ISME J.111019–1028. 10.1038/ismej.2016.164
82
LozuponeC. A.StombaughJ. I.GordonJ. I.JanssonJ. K.KnightR. (2012). Diversity, stability and resilience of the human gut microbiota.Nature489220–230. 10.1038/nature11550
83
MasonC. J.CoutureJ. J.RaffaK. F. (2014). Plant-associated bacteria degrade defense chemicals and reduce their adverse effects on an insect defoliator.Oecologia175901–910. 10.1007/s00442-014-2950-6
84
MasonC. J.RaffaK. F. (2014). Acquisition and structuring of midgut bacterial communities in gypsy moth (Lepidoptera: Erebidae) larvae.Environ. Entomol.43595–604. 10.1603/EN14031
85
MitchellA.MitterC.RegierJ. C. (2006). Systematics and evolution of the cutworm moths (Lepidoptera: Noctuidae): evidence from two protein-coding nuclear genes.Syst. Entomol.3121–46. 10.1111/j.1365-3113.2005.00306.x
86
MithoferA.BolandW. (2012). Plant defense against herbivores: chemical aspects.Annu. Rev. Plant Biol.63431–450. 10.1146/annurev-arplant-042110-103854
87
MitterC.DavisD. R.CummingsM. P. (2017). Phylogeny and evolution of Lepidoptera.Annu. Rev. Entomol.62265–283. 10.1146/annurev-ento-031616-035125
88
MoreauJ.MartinaudG.TroussardJ. P.ZanchiC.MoretY. (2012). Trans-generational immune priming is constrained by the maternal immune response in an insect.Oikos1211828–1832. 10.1111/j.1600-0706.2011.19933.x
89
MoriyamaM.NikohN.HosokawaT.FukatsuT. (2015). Riboflavin provisioning underlies Wolbachia’s fitness contribution to its insect host.mBio6:e01732-15. 10.1128/mBio.01732-15
90
NardiJ. B.MackieR. I.DawsonJ. O. (2002). Could microbial symbionts of arthropod guts contribute significantly to nitrogen fixation in terrestrial ecosystems?J. Insect Physiol.48751–763. 10.1016/S0022-1910(02)00105-1
91
NaritaS.NomuraM.KageyamaD. (2007). Naturally occurring single and double infection with Wolbachia strains in the butterfly Eurema hecabe: transmission efficiencies and population density dynamics of each Wolbachia strain.FEMS Microbiol. Ecol.61235–245. 10.1111/j.1574-6941.2007.00333.x
92
OliveiraM. G.De SimoneS. G.XavierL. P.GuedesR. N. (2005). Partial purification and characterization of digestive trypsin-like proteases from the velvet bean caterpillar, Anticarsia gemmatalis.Comp. Biochem. Physiol. B Biochem. Mol. Biol.140369–380. 10.1016/j.cbpc.2004.10.018
93
OliverK. M.DegnanP. H.BurkeG. R.MoranN. A. (2010). Facultative symbionts in aphids and the horizontal transfer of ecologically important traits.Annu. Rev. Entomol.55247–266. 10.1146/annurev-ento-112408-085305
94
ParamasivaI.SharmaH. C.KrishnayyaP. V. (2014). Antibiotics influence the toxicity of the delta endotoxins of Bacillus thuringiensis towards the cotton bollworm, Helicoverpa armigera.BMC Microbiol.14:200. 10.1186/1471-2180-14-200
95
PauchetY.HeckelD. G. (2013). The genome of the mustard leaf beetle encodes two active xylanases originally acquired from bacteria through horizontal gene transfer.Proc. R. Soc. Lond. B Biol. Sci.28020131021. 10.1098/rspb.2013.1021
96
PilonF. M.VisottoL. E.GuedesR. N.OliveiraM. G. (2013). Proteolytic activity of gut bacteria isolated from the velvet bean caterpillar Anticarsia gemmatalis.J. Comp. Physiol. B183735–747. 10.1007/s00360-013-0744-5
97
Pinto-TomásA.Uribe-LoríoL.BlancoJ.FontechaG.RodríguezC.MoraM.et al (2007). Actividades enzimáticas en aislamientos bacterianos de tractos digestivos de larvas y del contenido de pupas de Automeris zugana y Rothschildia lebeau (Lepidoptera: Saturniidae).Rev. Biol. Trop.55401–415. 10.15517/rbt.v55i2.6020
98
Pinto-TomásA. A.SittenfeldA.Uribe-LorioL.ChavarriaF.MoraM.JanzenD. H.et al (2011). Comparison of midgut bacterial diversity in tropical caterpillars (Lepidoptera: Saturniidae) fed on different diets.Environ. Entomol.401111–1122. 10.1603/EN11083
99
PowellJ. S.RaffaK. F. (1999). Effects of selected Larix laricina terpenoids on Lymantria dispar (Lepidoptera: Lymantriidae) development and behavior.Environ. Entomol.28148–154. 10.1093/ee/28.2.148
100
RaymondB.JohnstonP. R.WrightD. J.EllisR. J.CrickmoreN.BonsallM. B. (2009). A mid-gut microbiota is not required for the pathogenicity of Bacillus thuringiensis to diamondback moth larvae.Environ. Microbiol.112556–2563. 10.1111/j.1462-2920.2009.01980.x
101
RobinsonC. J.SchlossP.RamosY.RaffaK.HandelsmanJ. (2010). Robustness of the bacterial community in the cabbage white butterfly larval midgut.Microb. Ecol.59199–211. 10.1007/s00248-009-9595-8
102
RothO.JoopG.EggertH.HilbertJ.DanielJ.Schmid-HempelP.et al (2010). Paternally derived immune priming for offspring in the red flour beetle, Tribolium castaneum.J. Anim. Ecol.79403–413. 10.1111/j.1365-2656.2009.01617.x
103
SaddB. M.Schmid-HempelP. (2007). Facultative but persistent trans-generational immunity via the mother’s eggs in bumblebees.Curr. Biol.17R1046–R1047. 10.1016/j.cub.2007.11.007
104
SalemH.BauerE.KirschR.BerasateguiA.CrippsM.WeissB.et al (2017). Drastic genome reduction in an herbivore’s pectinolytic symbiont.Cell1711520.e13–1531.e13. 10.1016/j.cell.2017.10.029
105
SalemH.FlorezL.GerardoN.KaltenpothM. (2015). An out-of-body experience: the extracellular dimension for the transmission of mutualistic bacteria in insects.Proc. R. Soc. Lond. B Biol. Sci.282:20142957. 10.1098/rspb.2014.2957
106
SalminenJ. P.KaronenM. (2011). Chemical ecology of tannins and other phenolics: we need a change in approach.Funct. Ecol.25325–338. 10.1111/j.1365-2435.2010.01826.x
107
SasakiT.KuboT.IshikawaH. (2002). Interspecific transfer of Wolbachia between two lepidopteran insects expressing cytoplasmic incompatibility: a Wolbachia variant naturally infecting Cadra cautella causes male killing in Ephestia kuehniella.Genetics1621313–1319.
108
SchmidR. B.LehmanR. M.BrözelV. S.LundgrenJ. G. (2014). An indigenous gut bacterium, Enterococcus faecalis (Lactobacillales: Enterococcaceae), increases seed consumption by Harpalus pensylvanicus (Coleoptera: Carabidae).Fla. Entomol.97575–584. 10.1653/024.097.0232
109
SchoonhovenL. M.Van LoonJ. J.DickeM. (2005). Insect-plant Biology.Oxford: Oxford University Press.
110
ShaoY.ChenB.SunC.IshidaK.HertweckC.BolandW. (2017). Symbiont-derived antimicrobials contribute to the control of the lepidopteran gut microbiota.Cell Chem. Biol.2466–75. 10.1016/j.chembiol.2016.11.015
111
ShelomiM.DanchinE. G.HeckelD.WipflerB.BradlerS.ZhouX.et al (2016a). Horizontal gene transfer of pectinases from bacteria preceded the diversification of stick and leaf insects.Sci. Rep.6:26388. 10.1038/srep26388
112
ShelomiM.HeckelD. G.PauchetY. (2016b). Ancestral gene duplication enabled the evolution of multifunctional cellulases in stick insects (Phasmatodea).Insect Biochem. Mol. Biol.711–11. 10.1016/j.ibmb.2016.02.003
113
ShenJ.CongQ.BorekD.OtwinowskiZ.GrishinN. V. (2017). Complete genome of Achalarus lyciades, the first representative of the eudaminae subfamily of skippers.Curr. Genomics18366–374. 10.2174/1389202918666170426113315
114
SreeK. S.VarmaA. (2015). Biocontrol of Lepidopteran Pests: Use of Soil Microbes and their Metabolites.Berlin: Springer. 10.1007/978-3-319-14499-3
115
StaudacherH.KaltenpothM.BreeuwerJ. A.MenkenS. B.HeckelD. G.GrootA. T. (2016). Variability of bacterial communities in the moth Heliothis virescens indicates transient association with the host.PLoS One11:e0154514. 10.1371/journal.pone.0154514
116
StrongD. R.LawtonJ. H.SouthwoodS. R. (1984). Insects on Plants. Community Patterns and Mechanisms.London: Blackwell Scientific Publications.
117
SudakaranS.KostC.KaltenpothM. (2017). Symbiont acquisition and replacement as a source of ecological innovation.Trends Microbiol.25375–390. 10.1016/j.tim.2017.02.014
118
TagamiY.MiuraK. (2004). Distribution and prevalence of Wolbachia in Japanese populations of Lepidoptera.Insect Mol. Biol.13359–364. 10.1111/j.0962-1075.2004.00492.x
119
TakatsukaJ.KunimiY. (2000). Intestinal bacteria affect growth of Bacillus thuringiensis in larvae of the oriental tea tortrix, Homona magnanima diakonoff (Lepidoptera: Tortricidae).J. Invertebr. Pathol.76222–226. 10.1006/jipa.2000.4973
120
TangX.FreitakD.VogelH.PingL.ShaoY.CorderoE. A.et al (2012). Complexity and variability of gut commensal microbiota in polyphagous lepidopteran larvae.PLoS One7:e36978. 10.1371/journal.pone.0036978
121
TehB. S.ApelJ.ShaoY.BolandW. (2016). Colonization of the intestinal tract of the polyphagous pest Spodoptera littoralis with the GFP-tagged indigenous gut bacterium Enterococcus mundtii.Front. Microbiol.7:928. 10.3389/fmicb.2016.00928
122
TeixeiraL.FerreiraA.AshburnerM. (2008). The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster.PLoS Biol.6:e2. 10.1371/journal.pbio.1000002
123
Trauer-KizilelmaU.HilkerM. (2015). Insect parents improve the anti-parasitic and anti-bacterial defence of their offspring by priming the expression of immune-relevant genes.Insect Biochem. Mol. Biol.6491–99. 10.1016/j.ibmb.2015.08.003
124
van BormS.BuschingerA.BoomsmaJ. J.BillenJ. (2002). Tetraponera ants have gut symbionts related to nitrogen–fixing root–nodule bacteria.Proc. R. Soc. Lond. B Biol. Sci.2692023–2027. 10.1098/rspb.2002.2101
125
van den BoschT. J. M.WelteC. U. (2017). Detoxifying symbionts in agriculturally important pest insects.Microb. Biotechnol.10531–540. 10.1111/1751-7915.12483
126
van der VlugtC. J.van der WerfM. J. (2001). Genetic and biochemical characterization of a novel monoterpene 𝜀-lactone hydrolase from Rhodococcus erythropolis DCL14.Appl. Environ. Microbiol.67733–741. 10.1128/AEM.67.2.733-741.2001
127
VanceC. P. (2001). Symbiotic nitrogen fixation and phosphorus acquisition. Plant nutrition in a world of declining renewable resources.Plant Physiol.127390–397. 10.1104/pp.010331
128
VasanthakumarA.HandelsmanJ.SchlossP. D.BauerL. S.RaffaK. F. (2008). Gut microbiota of an invasive subcortical beetle, Agrilus planipennis Fairmaire, across various life stages.Environ. Entomol.371344–1353. 10.1093/ee/37.5.1344
129
VigneronA.MassonF.VallierA.BalmandS.ReyM.Vincent-MonégatC.et al (2014). Insects recycle endosymbionts when the benefit is over.Curr. Biol.242267–2273. 10.1016/j.cub.2014.07.065
130
VisottoL. E.OliveiraM. G.RibonA. O.Mares-GuiaT. R.GuedesR. N. (2009). Characterization and identification of proteolytic bacteria from the gut of the velvetbean caterpillar (Lepidoptera: Noctuidae).Environ. Entomol.381078–1085. 10.1603/022.038.0415
131
VogelH.MusserR. O.Celorio-ManceraM.-L. (2014). Transcriptome responses in herbivorous insects towards host plant and toxin feeding.Annu. Plant Rev.47197–233. 10.1002/9781118472507.ch6
132
VosM.HesselmanM. C.Te BeekT. A.van PasselM. W.Eyre-WalkerA. (2015). Rates of lateral gene transfer in prokaryotes: high but why?Trends Microbiol.23598–605. 10.1016/j.tim.2015.07.006
133
WaldbauerG.FriedmanS. (1991). Self-selection of optimal diets by insects.Annu. Rev. Entomol.3643–63. 10.1146/annurev.en.36.010191.000355
134
WangJ.PeifferM.HooverK.RosaC.ZengR.FeltonG. W. (2017). Helicoverpa zea gut-associated bacteria indirectly induce defenses in tomato by triggering a salivary elicitor(s).New Phytol.2141294–1306. 10.1111/nph.14429
135
WangY.LimL.DiGuistiniS.RobertsonG.BohlmannJ.BreuilC. (2013). A specialized ABC efflux transporter GcABC-G1 confers monoterpene resistance to Grosmannia clavigera, a bark beetle-associated fungal pathogen of pine trees.New Phytol.197886–898. 10.1111/nph.12063
136
WatanabeH.TokudaG. (2010). Cellulolytic systems in insects.Annu. Rev. Entomol.55609–632. 10.1146/annurev-ento-112408-085319
137
WeinertL. A.Araujo-JnrE. V.AhmedM. Z.WelchJ. J. (2015). The incidence of bacterial endosymbionts in terrestrial arthropods.Proc. R. Soc. Lond. B Biol. Sci.282:20150249. 10.1098/rspb.2015.0249
138
WernerG. D.CornwellW. K.SprentJ. I.KattgeJ.KiersE. T. (2014). A single evolutionary innovation drives the deep evolution of symbiotic N2-fixation in angiosperms.Nat. Commun.5:4087. 10.1038/ncomms5087
139
WerrenJ. H.BaldoL.ClarkM. E. (2008). Wolbachia: master manipulators of invertebrate biology.Nat. Rev. Microbiol.6741–751. 10.1038/nrmicro1969
140
WhitakerM. R.SalzmanS.SandersJ.KaltenpothM.PierceN. E. (2016). Microbial communities of lycaenid butterflies do not correlate with larval diet.Front. Microbiol.7:1920. 10.3389/fmicb.2016.01920
141
WybouwN.DermauwW.TirryL.StevensC.GrbicM.FeyereisenR.et al (2014). A gene horizontally transferred from bacteria protects arthropods from host plant cyanide poisoning.eLife3:e02365. 10.7554/eLife.02365
142
XiaX.GurrG. M.VasseurL.ZhengD.ZhongH.QinB.et al (2017). Metagenomic sequencing of diamondback moth gut microbiome unveils key holobiont adaptations for herbivory.Front. Microbiol.8:663. 10.3389/fmicb.2017.00663
143
XiaX.ZhengD.ZhongH.QinB.GurrG. M.VasseurL.et al (2013). DNA sequencing reveals the midgut microbiota of diamondback moth, Plutella xylostella (L.) and a possible relationship with insecticide resistance.PLoS One8:e68852. 10.1371/journal.pone.0068852
144
XiangH.WeiG. F.JiaS.HuangJ.MiaoX. X.ZhouZ.et al (2006). Microbial communities in the larval midgut of laboratory and field populations of cotton bollworm (Helicoverpa armigera).Can. J. Microbiol.521085–1092. 10.1139/w06-064
145
ZaluckiM. P.ShabbirA.SilvaR.AdamsonD.Shu-ShengL.FurlongM. J. (2012). Estimating the economic cost of one of the world’s major insect pests, Plutella xylostella (Lepidoptera: Plutellidae): just how long is a piece of string?J. Econ. Entomol.1051115–1129. 10.1603/EC12107
146
Zhu-SalzmanK.ZengR. (2015). Insect response to plant defensive protease inhibitors.Annu. Rev. Entomol.60233–252. 10.1146/annurev-ento-010814-020816
147
ZugR.HammersteinP. (2015). Bad guys turned nice? A critical assessment of Wolbachia mutualisms in arthropod hosts.Biol. Rev. Camb. Philos. Soc.9089–111. 10.1111/brv.12098
Summary
Keywords
gut bacteria, endosymbionts, moth, butterfly, symbiosis, horizontal transfer, maternal transfer
Citation
Paniagua Voirol LR, Frago E, Kaltenpoth M, Hilker M and Fatouros NE (2018) Bacterial Symbionts in Lepidoptera: Their Diversity, Transmission, and Impact on the Host. Front. Microbiol. 9:556. doi: 10.3389/fmicb.2018.00556
Received
22 December 2017
Accepted
12 March 2018
Published
27 March 2018
Volume
9 - 2018
Edited by
Sébastien Duperron, National Museum of Natural History, France
Reviewed by
Amparo Latorre, Universitat de València, Spain; Irene Lucile Garcia Newton, Indiana University Bloomington, United States
Updates
Copyright
© 2018 Paniagua Voirol, Frago, Kaltenpoth, Hilker and Fatouros.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Nina E. Fatouros, nina.fatouros@wur.nl
This article was submitted to Microbial Symbioses, a section of the journal Frontiers in Microbiology
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