Geographical, seasonal, and growth-related dynamics of gut microbiota in a grapevine pest, Apolygus spinolae (Heteroptera: Miridae)

doi:10.21203/rs.3.rs-3957390/v1

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Geographical, seasonal, and growth-related dynamics of gut microbiota in a grapevine pest, Apolygus spinolae (Heteroptera: Miridae)

https://doi.org/10.21203/rs.3.rs-3957390/v1

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A number of insects are associated with gut microorganisms, wherein symbionts play pivotal metabolic roles such as nutrient supplementation, diet degradation, and pesticide detoxification. Despite the ecological and evolutionary importance of gut microbial communities in insects, their diversity and dynamics need to be better investigated in many insect species. The green plant bug Apolygus spinolae, a notorious grapevine pest in Japan, damages grape shoots and severely reduces grape berry yield and quality. The plant bug possesses a simple tubular gut housing ~ 10⁴ bacteria. Here we investigated geographic, seasonal, and growth-related dynamics of gut microbiota by high-throughput sequencing in the grapevine pest. In Japanese populations of the plant bug, gut bacterial community was dynamically altered depending on geographic region, season, and developmental stage, in which particularly Serratia, Lactococcus and Caballeronia were abundantly detected. In one vineyard population, Spiroplasma was prevalent. Among the gut bacterial microorganisms, Serratia was consistently and abundantly detected, but significantly affected by seasonal change. In addition, Caballeronia, known as a specific symbiont of stinkbugs, was abundantly detected, especially in insects collected in late summer. The increase of Caballeronia was negatively correlated with Serratia, strongly suggesting an antagonistic interaction between gut bacterial members in A. spinolae.

Grapevine pest

Apolygus spinolae

Gut microbiota

Bacterial diversity

Many insects harbor diverse microorganisms inside their body, wherein they play essential metabolic roles for the insect hosts [1–3]. In particular, recent studies have revealed that gut microbiota provides various benefits to the host insects by, for example, activation of immune systems, provision of essential nutrients, digestion of indigestible foods, and/or degradation of plant toxins and insecticides [2–6]. Despite the accumulation of knowledge, the gut microbial community of several insect species remains unclear, and the biological functions of gut symbiotic microbes remain under investigation.

Hemiptera is a well-known insect group that possesses proboscis and straw-like mouthparts. and The insect group includes a number of agricultural pests, many of which are notorious not only because of their damage to crops, but also the transmission of serious plant pathogens [7, 8]. Homopterans, such as aphids and plant hoppers, commonly possess intracellular symbionts that provide essential amino acids scarcely contained in plant sap [9]. Meanwhile, heteropterans, such as stinkbugs of the infraorder Pentatomomorpha, commonly harbor specific extracellular gut symbionts in the crypt-bearing posterior region of their midgut called M4 [9, 10]. There are thus many examples in which stinkbugs retain symbiotic bacteria in their gut.

In contrast to the well-studied stinkbug groups, the family Miridae of the infraorder Cimicomomorpha has long been neglected in terms of microbial symbiosis. Miridae includes more than 11,000 species worldwide [11], and they cause serious damage to agricultural crops [12]. Based on their anatomical features, members of the Miridae, or plant bugs, are not thought to be associated with any specific gut symbionts. Recent studies, however, revealed that plant bugs possess diverse microorganisms in their gut [13–17], some of which are commonly found in or even specific to host plant bug species [16]. Considering the diverse functions of gut bacteria in insects, elucidating the diversity and biological functions of gut microbiota in plant bugs is pivotal for controlling the pest insect group.

The green pale plant bug Apolygus spinolae, distributed in a wide range of temperate zones in Eurasia, is a notorious pest of various crops including sunflowers, asparagus, tea trees, apples, and vines [18, 19]. In addition to these crops, reports of feeding damage to wine grapes by A. spinolae have increased in recent years and are becoming a problem, especially in northern Japan (personal communication). In Europe, the occurrence of the green pale plant bug in vineyards has also been reported in Slovenia [20]. While adults suck and damage grapevine leaves, nymphs cause more severe damage because they feed on shoots during their budding period, resulting in ragged leaves with numerous holes (Fig. 1A and B), and eventually leading to the loss of grape quantity and quality. Although A. spinolae has been recognized as an important agricultural pest of grapevines, its physiology and ecology, as well as symbiotic association with microbes, are still under investigation.

Here we investigated geographical, seasonal, and growth-related dynamics of gut microbiota of A. spinolae populations in vineyards of Hokkaido, northern Japan, by high-throughput sequencing of bacterial 16S rRNA gene. This study reveals community change in A. spinolae gut microbiota. Our results suggest that environments affect the gut microbiota of the pest plant bug species. This fundamental information potentially contributes to controlling this serious pest of wine grapes.

Insects

A. spinolae were collected at various sites in Hokkaido, Japan (Table S1). The five sites were Tsurunuma, Furano, Obihiro, Niseko, and Hakodate (Table S1). Captured insect samples were dipped in acetone during storage for DNA preservation. At Tsurunuma, the plant bugs were caught from June to September 2022. Other sites were sampled in September 2022. Insects were captured by sweeping more than 20 times between the six hedges of grapevine, targeting vineyards growing in and around the survey area. To clarify basic ecological and anatomical information of this plant bug, some adults and hatchlings were reared in a laboratory by feeding on sunflower seeds, buckwheat, wheat seeds, and distilled water containing 10% honey at 25℃ under long-day condition (light:dark = 18h:6h). To investigate gut morphology, adults were dissected in phosphate-buffered saline (PBS: 0.14 M NaCl, 0.0027 M KCl, 0.01 M PO₄³⁻, pH 7.4) using forceps under a binocular (Leica S8AP0 stereo microscope; Leica, Germany).

DNA extraction

After volatilizing the acetone, the entire body of the stinkbug was homogenized in sterile water before performing DNA extraction. In addition to the insects collected at the above sites, DNA was also extracted from 3rd, 4th, and 5th instar larvae. To perform DNA extraction on collected insects, Qiagen DNA extraction kits (Qiagen, Netherlands) were used according to company protocols. Nanodrop (ThermoFisher MA, USA) was used to check the quality of extracted DNA. The CO1 region of the stinkbug and V4 region of bacterial 16S were amplified by PCR, and the extraction of the target DNA was checked by electrophoresis using 1.5% agarose gel.

Amplification was performed on CO1 and 16S V4 genes using Amplitaq Gold polymerase (Applied Biosystems CA, USA). The 16S rRNA variable region V4 of the bacterial gene was amplified with the degenerate PCR primers 515F (5'-GTGCCAGCMGCCGCGGTAA-3') and 806R (5'-GGACTACHVGGGTWTCTAAT-3'). The PCR temperate program for the 16S rRNA V4 region is as follows: initial denaturation at 95 ℃ for 10 min, followed by 35 cycles of 95℃ for 30s, 55℃ for 60s, and 72℃ for 90s, and a final extension at 72℃ for 120s. Moreover, the CO1 region of the stink bug was amplified by the primers LCO1490 (5'-GGTCAACAAATCATAAAGATATTGG-3') and HCO2198 (5'-TAAACTTCAGGGTGACCAAAAAATCA-3'). PCR cycling conditions for the CO1 region were as follows: initial denaturation at 95℃ for 10 min, followed by 30 cycles of 95℃ for 30s, 48℃ for 60s, and 72℃ for 90s, and a final extension at 72℃ for 10 min. The PCR products were purified with Wizard SV Gel and PCR Clean-Up system (Promega, WI, USA).

Quantitative PCR

Whole homogenized adult and nymph bodies were used to estimate their bacterial amount. To indirectly estimate the number of bacterial cells the copy number of a bacterial gene, 16S rRNA variable region V4, was measured by quantitative PCR (qPCR). The qPCR was performed using a reaction master mix including the following: primers 515F and 806R, the KAPA SYBR FAST qPCR Master Mix Kit (KAPA Biosystems, MA, USA), and the Roche LightCycler 96 System (Roche, Switzerland). The qPCR temperature profile was as follow: 45 cycles at 95℃ for 15s, 55℃ for 30s, 72℃ for 30s. The gene copy number was calculated based on the standard curve for the 16S V4 region gene containing 10, 10², 10³, 10⁴, 10^5, and 10⁶ copies per reaction of the PCR product.

Amplicon sequencing

Prepared DNA was subjected to PCR amplification of 16S rRNA V4 gene for deep sequencing. The variable region was amplified using universal primer 515F and 806R [21]. The reaction master mix was comprised of 50 µM each dNTP, 0.4 µM 515F with Illumina P5 sequences, 0.4 µM 806R with Illumina P7 sequences (Illumina, CA, USA) [21], Q5 High-Fidelity DNA polymerase (New England Biolabs, MA, USA) and the extracted DNA template. The PCR cycling was as follow: initial denaturation at 98 ℃ for 30s, followed by 35 cycles of 98 ℃ for 10s, 55 ℃ for 20s, and 72 ℃ for 20s, and a final extension at 72 ℃ for 2 min. The DNA library containing tagged amplicons and the internal control phiX were used for sequencing on an Illumina iSeq100 sequencer (Illumina) using iSeq100 Reagent kit V2 (Illumina). The library generated 2 × 151 bp paired-end reads. All sequence data obtained in this study were deposited in the DDBJ/GenBank/EBI, under accession number PRJDB17462.

The paired-end reads were uploaded to the EzBioCloud 16S-based MTP app (ChunLab, Inc., Seoul, South Korea) [22] to check the data quality. The EzBioCloud software app filtered out the low-quality sequences with averaged Q values of less than 30. Moreover, the low-quality amplicons were removed from filtrated total amplicons, and low valid read samples (less than 35%) were avoided for analysis. The EzBioCloud 16S chimera-free database applied with the UCHIME algorithm [23] removed the chimera sequences. The EzBioCloud 16S database used with the VSEARCH program [24] performed the taxonomic assignment to detect and calculate the sequence similarities of the query paired-ends reads. EzBioCloud sequencing reads were clustered into the operational taxonomic units (OTUs) at 97% sequence similarity by the UPARSE algorithm [25].

Data analysis

Microbial richness was measured by ACE and Chao1 by using OTUs generated in this microbiome taxonomic profile. The diversity values of Shannon and Simpson α-diversity indices were applied to estimate the evenness for each group. Statistical analysis for ACE, Chao1, Shannon, and Simpson was assessed with the Kruskal-Wallis test, One-way analysis of variance (ANOVA), and Mann-Whitney U test, and these tests were conducted by GraphPad Prism v. 9. 5. 1 (Graph Pad Software, MA, US). Bata-diversity was assessed using Bray-Curtis distance between groups and their ordination visualized with principal coordinate analysis (PCoA) by vegdist function of vegan package in R [26]. Statistical differences in community structure between groups were tested with permutational multivariate analysis of variance (PERMANOVA) generated prcomp function of ape package in R programming language [26]. Generalized liner model (GLM) and correlation analysis were conducted using R language [26]. Climatic factors performed by GLM were obtained from the Automated Meteorological Data Acquisition System (AMeDAS) administered by the Japan Meteorological Agency (https://www.jma.go.jp)

Phylogenetic analysis

More than 2% of bacterial OTUs were used to construct a phylogenetic tree of the insect-associated bacteria using the MAFFT program [27] which aligned the V4 regions (250 bp) of the OTUs acquired in this study. In addition, bacterial 16S rRNA sequences of related genera were obtained from the NCBI nucleotide (nr) database and subsequently aligned together with the OTUs. The phylogenetic analysis was conducted using the maximum likelihood method (1,000 bootstraps) through the MEGA X software [28].

Basic information about A. spinolae and its gut

The field collection and laboratory rearing confirmed that A. spinolae has five nymphal stages before reaching adulthood (Fig. 1C). The midgut of A. spinolae was anatomically divided into three sections, M1-M3. The boundaries of each section are unclear and M4, the crypt-bearing gut symbiotic organ in other stinkbugs, is completely absent (Fig. 1D).

Geographic diversity of gut bacterial community

In the late summer season (September), A. spinolae were collected from five geographically different vineyards (Table S1) and their gut bacterial communities were investigated by high-throughput sequencing. The size of the bacterial population in A. spinolae was estimated by qPCR targeting bacterial 16S rRNA gene. The gene copy sizes of the gut and the remaining body parts excluding the gut, indicated here as mean ± SD

were of 1.6×10⁴ ± 1.7×10⁴ gene copies insect^− 1 and 1.1×10¹ ± 1.5×10¹ gene copies insect^− 1, for both groups respectively. To analyze the gut microbiota, we decided to investigate the whole body of A. spinolae due to the small amount of 16S rRNA gene copies in body parts other than the gut. The size of the bacterial population from whole homogenized adult bodies varied among samples with means of 2.4×10⁴ ± 2.5×10⁴ gene copies per insect (Table S2).

We obtained raw reads of adults (mean: 13,273 ± 8,031 reads) from Illumina iSeq100 (Table S2). Moreover, Good’s coverage of all clean reads was above 99%, which indicates that the sequencing effort was enough to capture total diversity (Table S2). At the phylum level, the Beta-diversity analysis showed that no significant differences were found among vineyard samples (Fig. S1). However, the genus level diversities among the vineyard showed significant differences by PERMANOVA using a PCoA plot based on Bray-Cutis distance (Fig. 2B). Microbiota of all vineyard samples included the genera Serratia, Lactococcus, and Caballeronia in different proportions (Fig. 2A). Among the five populations, microbiota of plant bugs collected in Tsurunuma, Niseko, and Hakodate were remarkably similar with each other (Fig. 2B), wherein, all three microbiota include a high % of Caballeronia (Fig. 2). In Furano and Obihiro vineyards, Serratia showed the highest proportional abundance in adult insects guts (Fig. 2). Turning to the Hakodate site, the A. spinolae microbiota had a unique composition in which Spiroplasma was most abundant (Fig. 2A). The phylogenic analysis indicated that the OTUs generated from Hakodate samples were relatively close to the insect-associated Spiroplasma platyhelix group and the plant pathogen S. citri group (Fig. S2). Alpha-diversity analysis indicated significant differences in the richness and evenness scores (Fig. S3), while the diversity analysis between females and males showed no significant difference (Fig. S4).

Growth-related diversity of gut bacterial community

The growth-related diversity of the microbial community was investigated in A. spinolae nymphs collected in the Tsurunuma vineyard from August to September. For the homogenized nymph’s body, the gene copy numbers were generated as 6.7×10³ ± 4.3×10³ gene copies insect^− 1 (Table S2). We obtained raw reads of nymphs stages (mean: 18,473 ± 8,594 reads) from deep sequencing (Table S2). At the phylum level, the Beta-diversity analysis didn’t show significant differences among each stage sample (Fig. S5). Among the detected bacterial genera, Hymenobacter, Methylobacterium, and Pseudomonas were consistently present from the 3rd to 5th instar, while in adults, these bacterial communities were negligible (Fig. 3A and Fig. S6). In 3rd instar nymphs, Caballeronia was not detected. Still, Caballeronia increased gradually from 4th instar stage to adult (Fig. 3A). Focusing on the adult stage, the microbiota changed in the bacterial proportion, in which Caballeronia, Acinetobacter, Cutibacterium, Flavobacterium, and Aquabacterium were

relatively increased. Serratia was consistently detected in high proportion through the developmental stages. To compare the bacterial community among different stages, a PCoA plot was generated and analyzed by PERMANOVA, which results indicated that bacterial community structure significantly differed among the growth stages (3rd instar nymph vs. 5th instar nymph: F = 3.01, r² = 0.333, P = 0.036; 3rd instar nymph vs adult: F = 4.013, r² = 0.223, P = 0.018; 4th instar nymph vs 5th instar nymph: F = 4.049, r² = 0.405, P = 0.072; 4th instar nymph vs adult: F = 3.12, r² = 0.194, P = 0.048; 5th instar nymph vs

Figure 3 Monthly change in bacterial diversity at each stage, 3rd instar (n = 3), 4th instar (n = 2), 5th instar (n = 6), and Adult (n = 13). (A) Relative abundance of dominant OTUs with more than 2.0%. (B) Beta diversity comparisons at the genus level between the different growth stages. Principal coordinate analysis (PCoA), based on Bray-Curtis dissimilarity, was tested using PERMANOVA with 999 permutations. The confidence ellipsoids of different stages provided by lines indicates 95% confidence intervals.

adult: F = 7.52, r² = 0.307, P < 0.0001) (Fig. 3B). Alpha-diversity analysis showed that the richness was not significantly different, but evenness showed a significant difference among the samples by month (Fig. S7).

Seasonal diversity of gut bacterial community

To reveal the seasonal population dynamics of A. spinolae and associated gut bacterial dynamics, we monitored the seasonal occurrence of adult A. spinolae from May to September in the Tsurunuma vineyard and investigated their gut microbiota (Fig. 4). The

vineyard has hedge-style viticulture, located on a rich vegetation mountain. Insect sweeping were done at six ridges between the grapevines. In May, A. spinolae adults did not occur, while they reached over 100 individuals in June; they then decreased in number in the midsummer season, but drastically increased with more than 300 individuals in September (Fig. 4A). Although we collected only a small number of nymphs and adults of A. spinolae inside vineyards in the midsummer season, our preliminary investigation suggests this plant bug proliferates outside vineyards and migrates into vineyards in late summer (data not shown).

To reveal the seasonal diversity of microbiota of adult A. spinolae, we examined and compared community structure for adults collected in June, July, and September (not enough adults were collected in August for analysis). PCoA plots based on Bray-Curtis showed that microbiota at the phylum level of monthly samples was significantly different between seasons (June vs July: F = 19.918, r² = 0.454, P < 0.0001; July vs September: F = 3.599, r² = 0.141, P = 0.0049; June vs September: F = 4.018, r² = 0.134, P = 0.0002) (Fig. S8). In the bacterial genera, the Beta-diversity analysis result indicated that the microbiota changed over the months (Fig. 3B and Fig. S9). In June, Methylobacterium, Pseudomonas, Sphingomonas, Enterobacterales, Acinetobacter, and Serratia appeared in many of the insect samples. However, the proportion of Enterobacterales and Acinetobacter decreased in July. Instead of Enterobacterales and Acinetobacter, the proportions of genus Caballeronia and Flavobacterium increased in the samples. Finally, the microbiota of insects collected in September included a large proportion of the genus Caballeronia, Aquabacterium, Acinetobacter, Cutibacterium, Flavobacterium, and Serratia. However, the proportion of Methylobacterium, Pseudomonas, Sphingomonas, and Enterobacterales decreased gradually from June to September. Otherwise, the genera Caballeronia, Aquabacterium, Acinetobacter, and Cutibacterium increased over the months. Beta-analysis by PCoA plot generated from Bray-Curtis dissimilarity indicated that monthly samples were significantly differed by PERMANOVA (June vs July: F = 5.3452, r² = 0.185, P < 0.0001; July vs September: F = 7.427, r² = 0.252, P < 0.0001; June vs September: F = 16.124, r² = 0.383, P < 0.0001) (Fig. 3C). Alpha-diversity analysis based on generated OTUs showed that the richness and evenness were not significantly different among the samples by month (Fig. S10).

Environmental factors affect Caballeronia and Serratia proportions in A. spinolae

Caballeronia and Serratia were consistently highly detected in most investigated individuals (Fig. 2, Fig. 3, and Fig. 4), but most bacterial groups seem negatively correlated with Serratia following GLM analysis (Table S3). To investigate important factors influencing Serratia proportions, ANOVA was performed and indicated that the proportion of Caballeronia and Serratia from June to September differed significantly (Fig. S9 and Table S4). Focusing on the environmental factor, GLM indicated that temperature, precipitation, humidity, and day length had a significant impact on the OTUs numbers of Serratia (Table S5). On the other hand, the climatic conditions did not influence Caballeronia (Table S5).

Phylogenetic diversity of Caballeronia and Serratia associated with A. spinolae

To analyze the phylogenetic diversity, more than 2% proportion of bacterial OTUs were selected. Notably, not all but many A. spinolae individuals collected in vineyards consistently possess a certain proportion of Caballeronia (Fig. 2, Fig. 3, and Fig. 4). The genus Caballelonia is well characterized as highly-specialized gut symbiotic bacteria of the bean bug Riptortus pedestris and allied stinkbug species [29]. Caballelonia includes over 60 described species and is currently grouped into four clades based on phylogenetic analyses: SBE-α, SBE-β, SBE-γ, and Coreoidea-clade [29]. Phylogenetic analysis of Caballeronia OTUs detected from the Tsurunuma population revealed that they belong to the SBE-β clade of Caballeronia (Fig. S11). In addition, OTUs detected from A. spinolae collected at other vineyards were placed into the clades SBE-α and SBE-β (Fig. S11).

With respect to Serratia, the genus is classified into omnivorous insect-associated group and insect-symbiont group [30–32]. Since the omnivorous insect-associated group includes entomopathogens and phytopathogens, it is often known as an important bacterial group in agriculture [33]. The phylogenetic analysis indicated that the OTUs detected in A. spinolae were placed in the omnivorous insect-associated group (Fig. S12).

To clarify community dynamics of associated microbiota in a grapevine pest Apolygus spinolae, we performed high-throughput sequencing of bacterial 16S rRNA gene. The microbiota of the plant bug was dynamically changed depending on geographic region, season, and insect developmental stage (Fig. 2, Fig. 3, and Fig. 4). Considering that A. spinolae lacks a gut symbiotic organ (Fig. 1D) and the remarkable diversity of their microbiota, it is plausible that the plant bug has no specific symbionts and acquires its microbiota from the environment every generation, although the insect looks likely to have an affinity for certain bacterial group such as Caballeronia and Serratia. Although the whole body of insects was homogenized and subjected to high-throughput sequencing in this study, most of the detected bacteria are thought to be gut bacteria, because almost no PCR amplification was detected in the insect tissues after removing the gut. However, some bacterial species such as Spiroplasma, detected only in Hakodate samples (Fig. 2 and Fig. S2), are probably associated with body tissues but not the luminal region of the gut, as reported previously in other insects [34]. There are few studies to comprehensively investigate the geography, season, and insect developmental stages that affect the gut bacterial composition of insect pests. Therefore, this report is a good example that integrated factors affect insect microbiota.

In recent years, the gut microbiota has been actively investigated in some members of the heteropteran family Miridae, including the sorghum plant bug Stenotus rubrovittatus, the small green mirid Nesidiocoris tenuis, the tropical plant bug Monalonion velezangeli, Adelphocoris suturatis, and Apolygus lucorum [14–17, 35, 36]. These studies have led to findings that the composition of the gut microbiota varies with climate, habitat, diet, and developmental stages, as well as results suggesting that some plant bugs may be vectors of phytopathogen and endophytes [15–17, 35, 37]. As shown here in A. spinolae, Proteobacteria consistently dominate the gut bacterial community in Miridae species, although the composition and dominant bacterial genus are remarkably diverse. While the feeding habits of A. spinolae are not thoroughly investigated, feeding habits of the Miridae are generally diverse from phytophagous to carnivorous through omnivorous, and change throughout their life cycle [12], which may be a factor causing their diverse gut community. Indeed, rearing experiments in Adelphocoris suturatis confirmed the diet-dependent effects on gut microbiota composition in this plant bug species [35].

In the rearing experiments on Adelphocoris suturatis, the proportion of Serratia increased when the insect was reared under carnivorous or omnivorous conditions [35], suggesting a metabolic function of this bacterial group when plant bugs feed on other insects. In addition, Serratia species are commonly detected in the gut of carnivorous and omnivorous insects, including mosquitoes (Aedes aegypti), Anopheles stephensi, fruit fly (Drosophila), and ladybugs (Harmonia axyridis) [38–41]. Indeed, Serratia species are known to have urea metabolism and secrete highly active proteases and chitinases in insect bodies, suggesting that Serratia support host digestion [42]. If so, it may be notable that Serratia shows seasonal changes in A. spinolae (Fig. 4, Table S4, and Table S5). Although speculative, the dynamics of Serratia suggest the seasonal changes in the feeding habits of A. spinolae in grapevine fields.

Alternatively, considering the Serratia OTUs detected in this study were closely related to S. marcescens, a well-known entomopathogenic bacterium [43, 44] (Fig. S12), this bacterial group is possibly a seasonal gut pathogen of A. spinolae. In A. suturatis, dietary change from herbivorous to omnivorous/carnivorous enhanced S. marcescens in the gut, which increased mortality [35]. The number of Serratia OTUs from A. spinolae appeared to decrease due to the increase of many other bacteria (Table S3). Therefore, it is hypothesized that dietary alterations in A. spinolae leads to an increase in the number of other bacteria and regulates the intestinal microflora to prevent proliferation of S. marcescens.

In addition to Serratia, members of Caballeronia were also consistently detected in A. spinolae. The genus Caballeronia is well known as the gut symbiont of several stinkbug species [10, 29, 45], wherein the symbiont plays important metabolic roles in recycling metabolic wastes and insecticide resistance [46–48]. Phylogenetic analysis showed that OTUs from Tsurunuma and Niseko were classified as SBE-β, while OTUs detected from Obihiro and Hakodate were classified as SBE-α (Fig. S11). Considering that not all but some Caballeronia strains can degrade insecticides [46], this region-dependent difference may be due not only to vegetation and diet but also to vineyard management. As discussed above, the variation may simply be related to changes in the bug’s feeding behavior. Alternatively, the bacterial groups probably inhibit each other inside the gut, and the power balance could be influenced by the insect’s habitat, environmental changes, and physiological changes. These hypotheses should be clarified in future studies.

A. spinolae collected in Hakodate had a high percentage of Spiroplasma (Fig. 2). S. citri, causing citrus stubborn disease (CSD), and S. kunkelii, causing corn stunt in maize, are known as typical phytopathogenic bacteria in the Spiroplasma family [49, 50]. Some of the Spiroplama infecting insects cause the well-known phenomenon of Drosophila male-killer [34]. Among the Spiroplasma species detected in this study, there were not only insect parasites but also OTUs classified in the same group as S. citri and S. kunkelii, which are known plant pathogens (Fig. S4). These results suggest that A. spinolae is a vector of the plant pathogen Spiroplasma. Although there are very few known cases of Spiroplasma transmission in the Miridae, the findings of this study may provide an opportunity to shed new light on how to manage phytopathogenic Spiroplasma.

In this study, we comprehensively analyzed the gut microbiota of vineyard populations of A. spinolae. To develop an integrated pest management (IPM) strategy for the pest plant bug, it is necessary to elucidate this pest's detailed developmental status and life cycle in vineyards. The clarification of the relationship and function between insects and gut symbiotic bacteria from this study not only leads to elucidating the ecological basis of the insects, but also has great potential to be developed into agricultural technology.

Supplementary Information The online version contains supplementary material available at https://doi.org/******resistlation_number

Acknowledgments We thank the researchers at the National Institute of Advanced Industrial Science and Technology (AIST) and local vineyard farmers for their help with collecting A. spinolae samples. Hideomi Ito and Kazumori Mise gave valuable advice about bioinformatic analysis. Madoka Miyazaki contributed insect survey and PCR assay. Antoine-Olivier Lirette checked English usage in this paper.

Author Contribution HM, T. Sato, T. Sone, and YK designed and developed the study. HM, KI, T. Sato, T. Sone, and YK collected insect samples. HM and KI performed 16S rRNA amplicon sequencing and data analysis. HM and KI performed statistical analysis. HM and YK wrote the manuscript with input from all authors.

Funding This study was supported by NOASTEC (S-3-7) to YK, JSPS KAKENHI to T. Sato (21K20579), and JSPS research fellowship for Young Scientists to KI (22KJ0057).

Conflict of Interest　The authors declare no competing interests.

Engel P, Moran NA (2013) The gut microbiota of insects–diversity in structure and function. FEMS Microbiol Rev 37: 699-735. https://doi.org/10.1111/1574-6976.12025
Douglas AE (2015) Multiorganismal insects: diversity and function of resident microorganisms. Annu Rev Entomol, Vol 60 60: 17-34. https://doi.org/10.1146/annurev-ento-010814-020822
Douglas AE (2022) Insects and their beneficial microbes. Princeton University Press
Dillon RJ, Dillon V (2004) The gut bacteria of insects: nonpathogenic interactions. Annu Rev Entomol 49: 71-92. https://doi.org/10.1146/annurev.ento.49.061802.123416
Kikuchi Y, Hosokawa T, Fukatsu T (2007) Insect-microbe mutualism without vertical transmission: a stinkbug acquires a beneficial gut symbiont from the environment every generation. Appl Environ Microb 73: 4308-4316. https://doi.org/10.1128/AEM.00067-07
Horak RD, Leonard SP, Moran NA (2020) Symbionts shape host innate immunity in honeybees. Proc Royal Soc B 287: 20201184. https://doi.org/10.1098/rspb.2020.1184.
Wang H, Wu K, Liu Y, Wu Y, Wang X (2015) Integrative proteomics to understand the transmission mechanism of barley yellow dwarf virus-GPV by its insect vector Rhopalosiphum padi. Sci Rep 5: 10971. https://doi.org/10.1038/srep10971
Liu W, Wang X (2018) New insights on the transmission mechanism of tenuiviruses by their vector insects. Curr Opin Virol 33: 13-17. https://doi.org/10.1016/j.coviro.2018.07.004
Kikuchi Y (2009) Endosymbiotic bacteria in insects: their diversity and culturability. Microbes Environ 24: 195-204. https://doi.org/10.1264/jsme2.ME09140S
Takeshita K, Kikuchi Y (2017) Riptortus pedestris and Burkholderia symbiont: an ideal model system for insect-microbe symbiotic associations. Res Microbiol 168: 175-187. https://doi.org/10.1016/j.resmic.2016.11.005
Cassis G, Schuh R (2012) Systematics, biodiversity, biogeography, and host associations of the Miridae (Insecta: Hemiptera: Heteroptera: Cimicomorpha). Annu Rev Entomol 57: 377-404. https://doi.org/10.1146/annurev-ento-121510-133533
Lu Y, Wyckhuys KA, Wu K (2023) Pest status, bio-ecology, and area-wide management of Mirids in east asia. Annu Rev Entomol 69. https://doi.org/10.1146/annurev-ento-121322-015345
Xue H, Zhu X, Wang L, Zhang K, Li D, Ji J, Niu L, Wu C, Gao X, Luo J (2021) Gut bacterial diversity in different life cycle stages of Adelphocoris suturalis (Hemiptera: Miridae). Front Microbiol 12: 670383. https://doi.org/10.3389/fmicb.2021.670383
Navarro-Escalante L, Benavides P, Acevedo FE (2023) Diversity of bacterial symbionts associated with the tropical plant bug Monalonion velezangeli (Hemiptera: Miridae) revealed by high-throughput 16S-rRNA sequencing. Research Square https://doi.org/10.21203/rs.3.rs-2022560/v7
Owashi Y, Minami T, Kikuchi T, Yoshida A, Nakano R, Kageyama D, Adachi-Hagimori T (2023) Microbiome of zoophytophagous biological control agent Nesidiocoris tenuis. Microb Ecol 86: 2923-2933. https://doi.org/10.1007/s00248-023-02290-y
Sato Y, Akao T, Takeshita K (2023) High prevalence of Pantoea spp. in microbiota associated with the sorghum plant bug Stenotus rubrovittatus (Heteroptera: Miridae). Microbes Environ 38: ME22110. https://doi.org/10.1264/jsme2.ME22110
Yang ZW, Luo JY, Men Y, Liu ZH, Zheng ZK, Wang YH, Xie Q (2023) Different roles of host and habitat in determining the microbial communities of plant-feeding true bugs. Microbiome 11: 244. https://doi.org/10.1186/s40168-023-01702-y
Kasiwara T, Asakawa K (1986) Hand book of crop pests, Tokyo
Kim DS, Jeon HY, Yiem MS, Lee JH, Na SY, Lee JO (2000) Damage patterns caused by Lygocoris spinolae(Hemiptera: Miridae) on ‘campbell early’ grapes. J Asia Pac Entomol 3: 95-101. doi: https://doi.org/10.1016/S1226-8615(08)60062-X
Beber K, Vrabl S, Matis G, Beber M (2001) First experiences with occurrence and control of green grape capsid (Lygocoris spinolae Meyer-Duer).Proccedings of the 5^th Slovenian Conference on Plant Protection: 350-355.
Caporaso JG, Lauber CL, Walters WA, Berg-Lyons D, Lozupone CA, Turnbaugh PJ, Fierer N, Knight R (2011) Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. PNAS 108: 4516-4522. https://doi.org/10.1073/pnas.1000080107
Yoon SH, Ha SM, Kwon S, Lim J, Kim Y, Seo H, Chun J (2017) Introducing EzBioCloud: a taxonomically united database of 16S rRNA gene sequences and whole-genome assemblies. Int J Syst Evol Microbiol 67: 1613. https://doi.org/10.1099/ijsem.0.001755
Edgar RC, Haas BJ, Clemente JC, Quince C, Knight R (2011) UCHIME improves sensitivity and speed of chimera detection. Bioinform 27: 2194-2200. https://doi.org/10.1093/bioinformatics/btr381
Rognes T, Flouri T, Nichols B, Quince C, Mahé F (2016) VSEARCH: a versatile open source tool for metagenomics. Peer J 4: e2584. https://doi.org/10.7717/peerj.2584
Edgar RC (2013) UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat Methods 10: 996-998. https://doi.org/10.1038/nmeth.2604
R Core Team R (2023) R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vinna, Austria https://www.r-project.org
Katoh K, Standley DM (2013) MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol Biol Evol 30: 772-780. https://doi.org/10.1093/molbev/mst010
Kumar S, Stecher G, Li M, Knyaz C, Tamura K (2018) MEGA X: Mol Evol Genet Analysis across Computing Platforms. Mol Biol Evol 35: 1547-1549. https://doi.org/10.1093/molbev/msy096
Ohbayashi T, Cossard R, Lextrait G, Hosokawa T, Lesieur V, Takeshita K, Tago K, Mergaert P, Kikuchi Y (2022) Intercontinental diversity of Caballeronia gut symbionts in the conifer pest bug Leptoglossus occidentalis. Microbes Environ 37: ME22042. https://doi.org/10.1264/jsme2.ME22042
Fukatsu T, Nikoh N, Kawai R, Koga R (2000) The secondary endosymbiotic bacterium of the pea aphid Acyrthosiphon pisum (Insecta: Homoptera). Appl Environ Microbiol 66: 2748-2758. https://doi.org/10.1128/aem.66.7.2748-2758.2000
Iguchi A, Nagaya Y, Pradel E, Ooka T, Ogura Y, Katsura K, Kurokawa K, Oshima K, Hattori M, Parkhill J, Sebaihia M, Coulthurst SJ, Gotoh N, Thomson NR, Ewbank JJ, Hayashi T (2014) Genome evolution and plasticity of Serratia marcescens, an important multidrug-resistant nosocomial pathogen. Genome Biol Evol 6: 2096-2110. https://doi.org/10.1093/gbe/evu160
da Mota FF, Castro DP, Vieira CS, Gumiel M, de Albuquerque JP, Carels N, Azambuja P (2019) In vitro trypanocidal activity, genomic analysis of isolates, and in vivo transcription of type VI secretion system of Serratia marcescens belonging to the microbiota of Rhodnius prolixus digestive tract. Front Microbiol 9. https://doi.org/10.3389/fmicb.2018.03205
Soenens A, Imperial J (2020) Biocontrol capabilities of the genus Serratia. Phytochem Rev 19: 577-587. https://doi.org/10.1007/s11101-019-09657-5
Harumoto T, Anbutsu H, Lemaitre B, Fukatsu T (2016) Male-killing symbiont damages host's dosage-compensated sex chromosome to induce embryonic apoptosis. Nat Commun 7. https://doi.org/10.1038/ncomms12781
Luo J, Cheng Y, Guo L, Wang A, Lu M, Xu L (2021) Variation of gut microbiota caused by an imbalance diet is detrimental to bugs' survival. Sci Total Environ 771: 144880. https://doi.org/10.1016/j.scitotenv.2020.144880
Xue H, Zhu X, Wang L, Zhang K, Li D, Ji J, Niu L, Gao X, Luo J, Cui J (2023) Dynamics and diversity of symbiotic bacteria in Apolygus lucorum at different developmental stages. J Cotton Res 6: 1-11. https://doi.org/10.1186/s42397-023-00142-1
Galambos N, Compant S, Wäckers F, Sessitsch A, Anfora G, Mazzoni V, Pertot I, Perazzolli M (2021) Beneficial insects deliver plant growth-promoting bacterial endophytes between tomato plants. Microorganisms 9: 1294. https://doi.org/10.3390/microorganisms9061294
Chandler JA, Lang JM, Bhatnagar S, Eisen JA, Kopp A (2011) Bacterial communities of diverse Drosophila species: ecological context of a host-microbe model system. PLoS Genet 7. https://doi.org/10.1371/journal.pgen.1002272
Chen SC, Walker ED (2017) Genomic and physiologic characterization of Serratia marcescens isolated from the gut of Anophles stephensi. Am J Trop Med Hyg 97: 455-455. https://doi.org/10.3389/fmicb.2017.01483
Wu P, Sun P, Nie KX, Zhu YB, Shi MY, Xiao CG, Liu H, Liu QY, Zhao TY, Chen XG, Zhou HN, Wang PH, Cheng G (2019) A gut commensal bacterium promotes mosquito permissiveness to arboviruses. Cell Host & Microbe 25: 101. https://doi.org/10.1016/j.chom.2018.11.004
Du LE, Xue H, Hu FM, Zhu XZ, Wang L, Zhang KX, Li DY, Ji JC, Niu L, Luo JY, Cui JJ, Gao XK (2022) Dynamics of symbiotic bacterial community in whole life stage of Harmonia axyridis (Coleoptera: Coccinellidae). Front Microb 13. https://doi.org/10.3389/fmicb.2022.1050329
Jupatanakul N, Pengon J, Selisana SMG, Choksawangkarn W, Jaito N, Saeung A, Bunyong R, Posayapisit N, Thammatinna K, Kalpongnukul N, Aupalee K, Pisitkun T, Kamchonwongpaisan S (2020) Serratia marcescens secretes proteases and chitinases with larvicidal activity against Anopheles dirus. Acta Trop 212. https:/doi.org/10.1016/j.actatropica.2020.105686
Li B, Yu RR, Liu BP, Tang QM, Zhang GQ, Wang YL, Xie GL, Sun GC (2011) Characteriazation and comparison of Serratia marcescens isolated from edible cactus and from silkworm for virulence potential and chitosnan susceptibility. Braz J Microbiol 42: 96-104. https://doi.org/10.1590/s1517-83822011000100013
Miller TA, Lauzon CR, Lampe DJ (2008) Technological advances to enhance agricultural pest management. Transgene Manag Vector-Borne Dis 627: 141-150.
Kikuchi Y, Hosokawa T, Fukatsu T (2011) Specific developmental window for establishment of an insect-microbe gut symbiosis. Appl Environ Microbiol 77: 4075-4081. https://doi.org/10.1128/aem.00358-11
Kikuchi Y, Hayatsu M, Hosokawa T, Nagayama A, Tago K, Fukatsu T (2012) Symbiont-mediated insecticide resistance. PNAS 109: 8618-8622. https://doi.org/10.1073/pnas.1200231109
Ohbayashi T, Futahashi R, Terashima M, Barrière Q, Lamouche F, Takeshita K, Meng XY, Mitani Y, Sone T, Shigenobu S, Fukatsu T, Mergaert P, Kikuchi Y (2019) Comparative cytology, physiology and transcriptomics of Burkholderia insecticola in symbiosis with the bean bug Riptortus pedestris and in culture. ISME J 13: 1469-1483. https://doi.org/10.1038/s41396-019-0361-8
Sato Y, Jang S, Takeshita K, Itoh H, Koike H, Tago K, Hayatsu M, Hori T, Kikuchi Y (2021) Insecticide resistance by a host-symbiont reciprocal detoxification. Nat Commun 12. https://doi.org/10.1038/s41467-021-26649-2
Whitcomb RF, Chen TA, Williamson DL, Liao C, Tully JG, Bove JM, Mouches C, Rose DL, Coan ME, Clark TB (1986) Spiroplasma kunkell sp. nov characteriztion of the etiologic agent of corn stunt disease. Int J Syst Bacteriol 36: 170-178. https://doi.org/10.1099/00207713-36-2-170
Gasparich GE (2010) Spiroplasmas and phytoplasmas: microbes associated with plant hosts. Biologicals 38: 193-203. https://doi.org/10.1016/j.biologicals.2009.11.007

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Geographical, seasonal, and growth-related dynamics of gut microbiota in a grapevine pest, Apolygus spinolae (Heteroptera: Miridae)

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Abstract

Figures

Introduction

Materials and Methods

DNA extraction

Quantitative PCR

Amplicon sequencing

Data analysis

Phylogenetic analysis

Results

Geographic diversity of gut bacterial community

Growth-related diversity of gut bacterial community

Seasonal diversity of gut bacterial community

Discussion

Declarations

References

Additional Declarations

Supplementary Files

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