Skip to main content
NCBI home page
Journal of Insect Science logoLink to Journal of Insect Science
. 2023 Apr 19;23(2):14. doi: 10.1093/jisesa/iead019

Isolation and characterization of gut bacteria associated with the degradation of host-specific terpenoids in Pagiophloeus tsushimanus (Coleoptera: Curculionidae) larvae

Heng Qiao 1,2,b, Han Zhu 3,4,b, Hui Li 5,6, Hongjian Chen 7,8, Shouyin Li 9,10, Cong Chen 11,12, Dejun Hao 13,14,✉
Editor: Xiao-Qiang Yu
PMCID: PMC10114288  PMID: 37074003

Abstract

Insect intestinal bacteria play an important role in resisting defensive substances of host plants. Pagiophloeus tsushimanus (Coleoptera: Curculionidae) feeds exclusively on camphor trees (Cinnamomum camphora, Laurales: Lauraceae) in China, causing substantial economic and ecological losses. It is unclear how the larvae of P. tsushimanus outcome the main secondary metabolites of C. camphora such as D-camphor, eucalyptol, and linalool. In this study, we isolated terpenoid-degrading bacteria from the gut of P. tsushimanus larvae by using selective culture medium. Maximum likelihood phylogenetic analyses were performed with 16S rDNA sequences to identify the bacteria, and results showed ten strains belonged to four genera, including Pseudomonas, Enterobacter, Serratia, and Corynebacterium. Then, gas chromatography was employed to determine the degradability of D-camphor, eucalyptol, and linalool by the isolated strains, results showed that Z5 strain (i.e., Corynebacterium variabile, Actinomycetales: Corynebacteriaceae), F1 strain (i.e., Pseudomonas aeruginosa, Pseudomonadales: Pseudomonaceae), and A3 strain (i.e., Serratia marcescens, Enterobacterales: Enterobacteriaceae) had the highest degradation rates of D-camphor, linalool, and eucalyptol, respectively. The intestinal bacteria were capable of terpenoid degradation in vitro, which suggested that these gut bacteria associated with P. tsushimanus play an important role in overcoming host plant secondary metabolite defense, thereby facilitating the host specialization of this pest.

Keywords: Pagiophloeus tsushimanus, intestinal bacteria, plant secondary metabolite, Cinnamomum camphora, functional analysis

The diversity and evolutionary success of insects depends partly on their inextricable relationships with beneficial microorganisms (Engel and Moran 2013). Insects associate with many microorganisms, and in their guts there are many symbiotic bacteria that have a certain impact on the insect growth and development, food digestion, and immunity (Zhang et al. 2021). Insect microorganisms participate in upgrading nutrient-poor diets; providing essential amino acids, vitamins, and other nutrients (Moran et al. 2003); detoxifying insecticides and protecting the host from poisoning (Kikuchi et al. 2012); protecting from predators, parasites, and pathogens (Kaltenpoth and Engl 2014); contributing to inter- and intra-specific communication; and inhibiting the defense of host plants (Berasategui et al. 2017, Paniagua Voirol et al. 2018).

Host plants can produce a large class of secondary metabolites, which repel or prevent insect feeding, cause direct intoxication and growth inhibition, and eventually produce the insect death (Jamwal et al. 2018, Divekar et al. 2022). Terpenoids are the most extensive plant secondary metabolites and have direct antifeedant and toxic effects on natural enemies (Bennett and Wallsgrove 1994). For example, in previous studies, volatile terpenoids extracted from Solanum nigrum, such as 7-epizingiberene (i.e., sesquiterpene), have been shown to be toxic to the silver leaf whitefly Bemisia tabaci, thereby reducing insect feeding on plants (Rosen et al. 2015). Correspondingly, herbivorous insects have evolved different mechanisms in the long-term interaction between plants and herbivorous insects in order to overcome the noxious effects of plant defense (Hammer and Bowers 2015). The gut bacteria of some insects can play an essential role in digesting and detoxifying foods containing plant secondary metabolites (Hehemann et al. 2010, Paniagua Voirol et al. 2018). Some reports have shown that bacteria associated with the mountain pine beetle Dendroctonus ponderosae (i.e., Pseudomonas and Rahnella) were proven to be able to reduce concentrations of terpenes, which directly or indirectly contribute to the capacity of the mountain pine beetle D. ponderosae to overcome host tree defenses (Adams et al. 2013, Boone et al. 2013). Currently, knowledge of the intestinal bacteria assisting host insects in detoxifying secondary metabolites is limited to some easily manipulated model insects, such as the diamondback moth Plutella xylostella, cabbage root fly Delia radicum, coffee berry borer Hypothenemus hampei, and pine weevils Hylobius abietis (Petersson and Örlander 2003, Jaramillo et al. 2006, Dosdall et al. 2012, Xia et al. 2017).

Pagiophloeus tsushimanus (Coleoptera: Curculionidae) is a phloem-feeding weevil that attacks camphor trees, Cinnamomum camphora (Laurales: Lauraceae) (Morimoto 1982). This weevil was first reported in Shanghai in 2014 and identified as a new record species in China (Huang et al. 2014). Over the past few years, this weevil pest has rapidly reached epidemic numbers, causing widespread damage to camphor plantations in our study areas (Songjiang District, Shanghai Province, China). From late April to early June, males and females mating often occurs on the sunny side of the crown, and it is observed that adults repeatedly mate and lay eggs. After mating, females crawl along the trunk below the canopy to find a suitable oviposition site and form an egg chamber in the cambium, and all females only lay one egg at a time, with one in each chamber. Pagiophloeus tsushimanus has one life cycle per year in Shanghai. Adults and 3–5 instar larvae overwinter from early November to early April. Adults overwinter in grooves on the underside of branches or branch nodes, and larvae overwinter in tunnels. Adults feed on the bark of twigs, and occasionally on new buds. Larvae (1st–2nd instar) feed on the phloem, while 3rd–5th instar can bore into the phloem and the cambium (Chen et al. 2021). Therefore, the larvae were supposed to have the ability to overcome toxic secondary metabolites produced by camphor trees when feeding on the phloem. We previously analyzed the phloem and cambium of the trunk at the feeding site of P. tsushimanus larvae by gas chromatography–mass spectrometry (GC–MS) and found 19 main secondary metabolites. Among them, the relative content of the top three substances were eucalyptol (31.89%), D-camphor (19.22%), and linalool (10.16%) (Li et al. 2022). Studies have shown that the secondary metabolites of C. camphora are considered to have a high degree of resistance to a variety of herbivorous insects (Jiang et al. 2016, Wu et al. 2020). In our previous study, however, low doses of exogenous camphor can instead promote growth and development of P. tsushimanus (Li et al. 2023), suggesting that this specialist weevil appears to have evolved a strategy adapted to its host-specific chemical defenses. Thus, this deserves in-depth studies to investigate whether the symbiotic gut bacteria in P. tsushimanus larvae play an important role in detoxifying and metabolizing these hosts’ toxins.

In this study, we isolated the gut bacteria of P. tsushimanus larvae and identified the strains by combining morphological characteristics as well as phylogenetic analysis for the first time. We further tested the terpenoids degradability of isolated strains by GC measurements. Our results may further elucidate interactions between P. tsushimanus and its gut bacteria in terms of host plant adaptations.

Materials and Methods

Insect Collection and Dissection

Pagiophloeus tsushimanus larvae were collected from a camphor plantation in Songjiang District, Shanghai, China. The larvae were subjected to starvation in an incubator at a temperature of 28 ± 0.5 °C and a relative humidity of 70 ± 5% in the dark for 24 h. Thirty 4th-instar larvae were randomly selected, and the surface of the larvae was disinfected with 70% alcohol and soaked in phosphate buffer solution (PBS), which was repeated three times to avoid external contamination (Morales-Jiménez et al. 2012). After dissection under sterile conditions, the guts were immediately transferred to a 2 ml sterile centrifuge tube with 0.5 ml of PBS for subsequent grinding.

Isolation and Purification of Terpenoid-Degrading Bacteria

Eucalyptol [99.5% purity, Chemical Abstracts Service (CAS) number: 470-82-6] (Aladdin, China), D-camphor (96% purity, CAS number: 464-49-3) (Aladdin, China), and linalool (98% purity, CAS number: 78-70-6) (Aladdin, China) were added to 50 ml of Bushnell Haas broth liquid medium at a ratio of 1% (w/v), and the liquid BHB medium consisted of 0.20 g/l MgSO4, 0.02 g/l CaCl2, 1.0 g/l KH2PO4, 1.0 g/l K2HPO4, 1.0 g/l (NH4)2SO4, 0.05 g/l FeCl3 (pH 7.0–7.2). Then, the larval intestinal abrasive was inoculated into the medium at a ratio of 3% (v/v), without intestinal abrasive medium as a control. After inoculation, the medium was incubated at 28 °C and 150 rpm. When the bacterial liquid was cloudy, the culture was serially diluted from 10−1 to 10−6, and 100 μl of dilution was coated on LB, NA, and TYA solid media, with three replicates per medium. The LB, NA, and TYA solid media consisted of 10 g/l peptone, 5 g/l yeast extract, 10 g/l NaCl, and 18 g/l agar (pH 7.2); 10 g/l peptone, 3 g/l beef extract, 5 g/l NaCl, and 18 g/l agar (pH 7.2); and 2.0 g/l yeast extract, 2.0 g/l beef paste, 6.0 g/l peptone, 40 g/l glucose, 3.0 g/l ammonium acetate, 0.5 g/l KH2PO4, 0.2 g/l neutral red, 0.2 g/l MgSO4·7H2O, 0.02 g/l FeSO4·7H2O, and 15 g/l agar (pH 6.5), respectively. After coating, the Petri dishes were incubated at 28 °C for 48 h in a bacteriological incubator. After selecting single colonies, the isolated and purified strains were preserved in 25% glycerol at −80 °C.

Morphological Identification of Terpenoid-Degrading Bacteria

Referring to Shen and Chen (2018), morphological characteristics such as color, size, shape, edges, bulges, transparency, and gloss of individual colonies were observed and described. And the morphology of the strain was observed by Gram staining, spore, capsule, and flagella staining.

Molecular Identification of Terpenoid-Degrading Bacteria

The purified bacterial strains were inoculated in LB liquid medium. After inoculation, these media were incubated at 28 °C, with shaking at 220 rpm for 12 h, and 1 ml of the cloudy bacterial solution was taken. Then, the bacterial genomic DNA was extracted by the E.Z.N.A.Soil DNA Kit (Omega Bio-Tek, USA) following the manufacturer’s instructions.

PCR amplification was performed using the extracted total bacterial DNA as a template with Taq DNA polymerase (Accurate, China) and 16S rDNA universal primers 27F (5ʹ-AGAGTTTGATCCTGG CTCAG-3ʹ) and 1492R (5ʹ-GGTTACCTTGTTACGACTT-3ʹ). The PCR reaction volume was 25 μl: 9.5 μl ddH2O, 12.5 μl 2 × Accurate Taq Master Mix, 1.0 μl gDNA (50 ng/μl), 1 μl primer 27F (10 μmol/l), and 1 μl primer 1492R (10 μmol/l). The PCR program included 30 s at 94 °C; 30 cycles of 98 °C for 10 s, 55 °C for 30 s, 72 °C for 2 min; and 72 °C for 2 min. Then, the PCR products were separated on a 1% agarose gel and sequenced by Jie Li Biology Co. Ltd (Shanghai, China).

The sequences of different bacterial isolates were blasted by online NCBI Blastn program (http://www.ncbi.nih.gov/blast). Multiple sequence alignments of 16S rDNA gene sequences of terpenoids-degrading bacteria and their closest relatives or members were performed by MAFFT version 7 software (https://www.ebi.ac.uk/Tools/msa/mafft/). Phylogenetic analyses were conducted using Bayesian inference (BI) and Maximum likelihood (ML). ModelFinder (Kalyaanamoorthy et al. 2017) was used to select the best-fit model using AIC criterion in PhyloSuite (Zhang et al. 2020). Best-fit model according to AIC: TIM3+F+R3 for BI and ML. ML analyses were conducted with IQ-TREE (Nguyen et al. 2015) using aligned sequences. Clade support was assessed by 5,000 bootstrap pseudoreplicates with the standard bootstrap of 1,000 replicates of the aligned data set. Bayesian inference (BI) analyses were analyzed in MrBayes v. 3.2.6 (Ronquist et al. 2012). Two million generations were run, with 25% of the generations as burn-in. The potential scale reduction factor (PSRF) close to 1.0 and the average standard deviation of split frequencies below 0.01 were accepted.

The DNA sequences of the A1, A2, A3, F1, F2, Z1, Z2, Z3, Z4, and Z5 strains were uploaded to GenBank with the numbers of ON153202, ON153203, ON153204, ON153205, ON153206, ON153207, ON153208, ON153209, ON153210, and ON153211, respectively.

Degradation Capacity of Bacterial Isolations

Sample collection and processing for GC.

The purified and preserved strain was inoculated in 30 ml of LB liquid medium for activation. The activated bacterial liquid was centrifuged at 10,000 rpm for 10 min at room temperature. The medium was discarded, and the bacteria were retained. Then, the bacteria were continuously washed with sterile water three times, and the concentration of bacterial liquid was diluted to an OD600 value of 1.5. After activation, 100 μl bacterial liquid was inoculated into 3 ml BHB medium containing 1% (w/v) eucalyptol, and the BHB medium without bacterial liquid was used as the blank control group. After shaking culture at 28 °C and 150 r/min for 10 days, 2 ml of chromatographically pure n-hexane (98% purity, CAS number: 110-54-3) (Aladdin, China) was added, treated in a sealed shaker for 3 h, placed upright, and treated overnight at −20 °C. After thawing at room temperature, 1 ml of hexane extract was transferred into a 2 ml autosampler vial (Figure S1). There were five replicates in the experimental group and the control group.

Sample processing.

Ten microlitres of n-hexane solution with 50% isobutylbenzene (99.5% purity, CAS number: 538-93-2) (Aladdin, China) was added as the internal standard (IS) for each sample, GC was used for quantitative analysis, and the sample was stored in a −20 °C refrigerator before analysis. Standard treatment: eucalyptol, D-camphor, and linalool standards with different masses were added to the n-hexane solution. Then, 10 μl n-hexane solution with 50% isobutylbenzene was added as the IS to prepare the standard with mass ratios of 1:1, 2:1, 3:1, 4:1, and 5:1. After mixing evenly, the standard was placed in a refrigerator at −20 °C to be tested.

Gas chromatography.

The degradation capacity of each strain for D-camphor, eucalyptol, and linalool was determined by a GC system (6890N, Agilent Technologies, USA) equipped with a flame ionization detector with an ionization energy of 70 eV and a fused silica capillary column (DB-5, 30 m × 0.32 mm × 0.25 μm, Agilent J & W GC columns, USA). The carrier gas was nitrogen, with a flow rate of 1 ml/min. The injector and mass transfer line temperatures were both 250 °C. The injection volume was 1 μl, with a hydrogen flow of 40 ml/min, air flow of 450 ml/min, and nitrogen flow of 15 ml/min. The starting temperature of the heating program was kept at 40 °C for 1 min, 20 °C/min to 110 °C for 1 min, 4 °C/min to 120 °C for 1 min, and 20 °C/min to 200 °C. The total running time was 13 min.

Calculation of the Degradation Rate. A 1 μl standard sample was taken for GC analysis. The peak retention time of each component in the sample to be measured is compared with that in the standard sample to determine the component to be tested. The standard samples with different mass ratios were determined by GC. According to the peak area ratio of standard samples to IS, the standard curve fitting was plotted by Origin 2021 (OriginLab, UK) to obtain the linear regression equation. The quantifications were carried out using the internal standard method (Deng and Zhang 2019), the formula of the relative correction factor f is as follows:

f=Asms/Armr

As: peak area of internal standard sample; Ar: peak area of standard sample; ms: internal standard mass; mr: standard sample mass.

According to the peak area (Ai) of the tested component, the peak area ratio of the internal standard, and the relative correction factor, the mass (mi) of the tested component was calculated as follows:

mi=f×(Ai/Asms)

Finally, the degradation rate of the strain (DE) was calculated according to the following formula:

DE(%)=(miC−miE)/miC

mi C : the mass of tested components in the control group; miE: the mass of tested components in the experimental group.

Differences in the degradation rates of D-camphor and eucalyptol by different D-camphor degrading strains and eucalyptol-degrading strains were determined using one-way analysis of variance (ANOVA), followed by Tukey’s test (P < 0.05). Difference in the linalool degradation rate between F1 strain and F2 strain was determined by independent sample t-test (P < 0.05). All statistical analyses were performed using SPSS 22.0 (IBM SPSS Statistics, USA) and plotted with Origin 2021 (OriginLab, UK).

Results

Isolation and Morphological Characterization of Terpenoid-Degrading Bacteria in the Gut of P. tsushimanus Larvae

A total of ten strains of bacteria were isolated from the intestines of 4th-instar larvae. Three eucalyptol-degrading bacteria were coded as A1, A2, and A3; two linalool-degrading bacteria were coded as F1 and F2; and four D-camphor-degrading bacteria were coded as Z1, Z2, Z3, and Z4 (Fig. 1). The morphological characteristics of the ten strains are shown in Table 1. The bacterial isolates exhibited diverse colony sizes, colors, margins, texture, and so on. The sizes of six isolates (i.e., A1, A2, A3, F1, Z1 and Z2) were medium, and four isolates (i.e., Z2, Z3, Z4, and Z5) were small. The colors of A2, F2, and Z3 were milky; A1, F1, Z1, Z2, and Z5 were yellowish; and A3 and Z4 were red. The margins of A2, F2, Z2, and Z3 were neat; A1, F1, and Z1 were irregular margins; and A3, Z4, and Z5 were regular margins. The shapes of the ten isolates were rod-like, and none had spores; four isolates (i.e., A1, F1, Z1, and Z2) had capsulatum, and nine isolates had flagellum. Regarding Gram staining, only the strain F5 was Gram positive, and the remaining isolates were Gram negative.

Fig. 1.

Fig. 1.

Open in a new tab

The terpenoid-degrading bacteria in the gut of P. tsushimanus larvae (A1, A2, A3, F1, F2, Z1, Z2, Z3, Z4, and Z5). D-camphor-degrading bacteria: Z1, Z2, Z3, Z4, and Z5; eucalyptol-degrading bacteria: A1, A2, and A3; and linalool-degrading bacteria: F1 and F2.

Table 1.

Morphological characteristics of terpenoid-degrading bacteria in the gut of P. tsushimanus larvae

Strain Bacterium shape Gram staining Spores Capsule Flagella Colony morphology
A1 Rod Gram −ve – + + Yellowish, medium-size, round, irregular margin, opaque, smooth
A2 Rod Gram −ve – – + Milky, medium-size, round, neat margin, convex, opaque, smooth
A3 Rod Gram −ve – – + Red, medium-size, round, regular margin, convex, opaque, smooth
F1 Rod Gram −ve – + + Yellowish, medium-size, round, irregular margin, opaque, smooth
F2 rod Gram −ve – – + Milky, medium-size, round, neat margin, convex, opaque, smooth
Z1 rod Gram −ve – + + Yellowish, medium-size, round, irregular margin, opaque, smooth
Z2 rod Gram −ve – + + Yellowish, small-size, round, neat margin, convex, opaque, sticky
Z3 rod Gram −ve – – + Milky, small-size, round, neatly margin, convex, opaque, smooth
Z4 rod Gram −ve – – + Red, small-size, round, regular margin, convex, opaque, smooth
Z5 rod Gram +ve – – – Yellowish, small-size, round, regular margin, convex, translucent, smooth
Open in a new tab

Molecular Identification of Terpenoid-Degrading Bacteria in the Gut of P. tsushimanus Larvae

It was found that the identity between the sequences of 10 strains and the known 16S rDNA sequences in the database ranged from 99.57% to 100% by BLAST. As shown in Table 2, the BLAST results showed that the eucalyptol-degrading strains A1, A2, and A3 had the highest identity with Pseudomonas aeruginosa, Enterobacter sp., and Serratia marcescens, respectively; the linalool-degrading strains F1 and F2 had the highest identity with P. aeruginosa and Enterobacter sp., respectively; and the D-camphor-degrading strains Z1, Z2, Z3, Z4, and Z5 had the highest identity with P. aeruginosa, Pseudomonas sp., Enterobacter sp., S. marcescens, and Corynebacterium variabile, respectively.

Table 2.

16S rDNA identification results of degrading bacteria in larval gut of P. tsushimanus

Strain Strain with the highest identity in Blast GenBank accession number Identity (%)
A1 Pseudomonas aeruginosa MF872727.1 100.00
F1 Pseudomonas aeruginosa OM865894.1 100.00
Z1 Pseudomonas aeruginosa KF483133.1 99.93
Z2 Pseudomonas sp. AY456708.1 99.86
A3 Serratia marcescens AY043386.1 99.79
Z4 Serratia marcescens AY043386.1 99.72
A2 Enterobacter sp. JQ765425.1 99.72
F2 Enterobacter sp. JQ765425.1 99.72
Z3 Enterobacter sp. JQ765425.1 99.72
Z5 Corynebacterium variabile MT573863.1 99.57
Open in a new tab

The BI and ML analyses based on 16S rDNA gene sequences of terpenoids-degrading bacteria produced very similar tree topologies. The topology of the ML phylogram was shown as Fig. 2. All strains respectively clustered in two major clades, including Actinomycetota and Pseudomonadota clades. Among them, Z5 clustered together with the strains of C. variabile, which belonged to Actinomycetota; and other strains all clustered together with the strains of Pseudomonadota. To be specific, the phylogenetic tree showed that A1, F1, and Z1 clustered together with the strains of P. aeruginosa; Z2 clustered together with the strains of P. chlororaphis, and A1, F1, Z1, and Z2 both belonged to the genus Pseudomonas; A2, F2, and Z3 all clustered together with the strains of the genus Enterobacter, and A2 and Z3 both clustered together with the strains of Pluralibacter gergovlae; A3 and Z4 both clustered together with the strains of S. marcescens.

Fig. 2.

Fig. 2.

Open in a new tab

Phylogenetic analysis of the terpenoid-degrading strains inferred using 16S rDNA gene sequences. The phylogram was constructed using maximum likelihood by 5,000 bootstrap pseudoreplicates with the standard bootstrap of 1,000 replicates. ML (bootstrap values) and BI (posterior probabilities) support values ≥80% are reported above and below nodes, respectively. Granulosicoccus antarcticus (EF495228.1) and G. antarcticus (NR_044255.1) were included as outgroups.

The above molecular identification results showed that A1, F1, and Z1 are the same species, which belonged to P. aeruginosa; Z2 belonged to P. chlororaphis; A2, F2, and Z3 all belong to Enterobacter sp.; A3 and Z4 belonged to S. marcescens; Z5 belonged to C. variabile. Although A1, F1, and Z1 showed little difference in morphological and molecular phylogenetic traits, we initially experimented with them as different strains due to the lack of in-depth analysis, which does not exclude the possibility that they are consistent. Likewise, A2, F2, and Z3 are also that.

Analysis of the Degradation Capacity of Terpenoid-Degrading Bacteria in the Gut of P. tsushimanus Larvae

The terpenoid-degrading bacteria were added to BHB inorganic salt medium containing D-camphor, eucalyptol, and linalool, respectively, and then the residues of D-camphor, eucalyptol, and linalool in BHB were determined by GC, which could determine the degradation rates of D-camphor, eucalyptol, and linalool by each bacterium. The standard curve for calculating the degradation rate is shown in Figure S2. The results showed that the retention time of the absorption peaks of D-camphor, eucalyptol, and linalool in the chromatograms of the samples supplemented with the ten strains was consistent with that of the control chromatograms, but the content of substances as determined from the absorption peaks area was significantly reduced. For example, the retention time of the D-camphor absorption peak was 9.7 min, the response values of Z1, Z2, Z3, Z4, and Z5 decreased to different degrees compared with that of the control samples (Fig. 3A). Among the five D-camphor-degrading bacteria (Z1-5), the degradation rate of Z5 was the highest (11.44 ± 2.88%), which was significantly different from that of the other four bacteria (F = 10.709; df = 4, 20; P < 0.001); the degradation rates of Z4, Z1, and Z2 were 6.07 ± 3.27%, 5.59 ± 1.89%, and 3.61 ± 2.19%, respectively. The degradation rate of Z3 was the lowest, which was 2.00 ± 1.59% (Fig. 3Aʹ). When the retention time of the eucalyptol absorption peak was 7.7 min, and the response value of A1, A2, and A3 all decreased to different degrees, compared with that of the control group (Fig. 3B). Among the three strains of eucalyptol-degrading bacteria (A1-3), the degradation rate of A3 was 17.08 ± 2.74%, which was significantly higher than that of A1 and A2 (F = 16.996; df = 2, 12; P < 0.001). Namely, the degradation rates of A1 and A2 were 9.49 ± 1.94% and 11.80 ± 1.45%, respectively (Fig. 3Bʹ). As shown in Fig. 3C, the retention time of the linalool absorption peak was 8.7 min, and the response values of F1 and F2 both decreased to different degree compared with that of the control samples. The degradation rates of F1 and F2 were 16.03 ± 2.75% and 12.05 ± 2.04%, respectively, but there was no significant difference in the degradation rates of the two strains of linalool-degrading bacteria (t = 2.594; P = 0.780) (Fig. 3Cʹ).

Fig. 3.

Fig. 3.

Open in a new tab

Degradation capacity of terpenoid-degrading bacteria in the gut of P. tsushimanus larvae. (A, Aʹ) The chromatographic diagram and degradation rate of D-camphor-degrading bacteria; significance was determined by Tukey’s one-way-ANOVA (P < 0.05); the different letters indicate significant differences, and the same letters indicate no differences. (B, Bʹ) The chromatographic diagram and degradation rate of eucalyptol-degrading bacteria; significance was determined by Tukey’s one-way-ANOVA (P < 0.05); the different letters indicate significant differences, and the same letters indicate no differences. (C, Cʹ) The chromatographic diagram and degradation rate of linalool-degrading bacteria; Significance was determined by independent sample t test (P < 0.05); NS indicates nonsignificant. All bars and error bars represent mean and standard error.

Discussion

Herbivorous insects mitigate the effects of toxic plant secondary metabolites by seeking help from symbiotic gut microbes to adapt to plant secondary metabolites (Afroz et al. 2021). The fact that symbiotic bacteria of some weevils can help host to overcome plant secondary metabolites has been confirmed (Adams et al. 2013, Boone et al. 2013). Pagiophloeus tsushimanus is an emerging insect pest exclusively infesting camphor trees. Its outbreak is largely due to its evolution of a high degree of adaptability to the chemical defense of specific host plants (i.e., camphor trees). In this context, clarifying the biological metabolic function of microbial symbioses in P. tsushimanus is not only helpful to understand the multiple interactions of insect–plant–microbes, but also provides a new insight for the comprehensive control of this pest. So, it’s of great interest to explore the microbial symbionts of P. tsushimanus that contribute to the detoxification of plant secondary metabolites. In this study, we first isolated and identified terpenoid-degrading bacteria in the gut of P. tsushimanus larvae. These bacteria were proven to degrade terpenoids when cultured separately on medium containing one terpenoid (i.e., eucalyptol, linalool, or D-camphor). Although A1, F1, and Z1 strain have almost no differences in morphology (including size, color, margin, shape, and so on) and molecular phylogenetic traits, but since there is no in-depth analysis, we initially treated them as different strains for the GC experiments, which does not rule out the possibility that they are consistent. Similarly, A2, F2, and Z3 are also that. After determining the terpenoid degradation rate of each bacterial strain by GC, we found that each strain could efficiently degrade on terpenoids (i.e., eucalyptol, linalool, and D-camphor), and that the degradation efficiencies of Pseudomonas and Serratia were relatively significant. In summary, these results revealed that the intestinal bacteria of P. tsushimanus have the capacity in detoxifying the host’s defensive substances and sequentially help P. tsushimanus adapt to host plants.

The fact that monoterpenes are utilized as energy and carbon sources by several aerobic microorganisms has been known since the 1960s (Lüddeke et al. 2012). Most reports dealt with Pseudomonas and Enterobacter species, for example P. aeruginosa (Förster-Fromme et al. 2006), P. putida (Tudroszen et al. 1977), P. delhiensis (Prakash et al. 2007), E. cowanii (Yang et al. 2007), and so on. In this research, our findings are similar to the previous studies. The bacterial identification showed that most monoterpenes-degradating strains belonged to Pseudomonas and Enterobacter species, for example, P. aeruginosa, P. chlororaphis, and Enterobacter sp., and S. marcescens and C. variabile also metabolize these terpenoids.

Some intestinal bacteria can help host insects defend against toxic secondary metabolites of plants and play an important role in the co-evolution of phytophagous insects and host plants (Engel and Moran 2013). In particular, symbiotic bacteria have been previously demonstrated to break down terpenoids, including sesquiterpenes, monoterpenes, and diterpenes. For example, Pseudomonas spp. and Rahnella aquatilis associated with D. ponderosae are capable of α-pinene degradation (Mason et al. 2014). The genera Bacillus, Burkholderia, Enterobacter, Klebsiella, and Pseudomonas that were isolated from leaf-cutter ants could degrade β-pinene, β-caryophyllene, or linalool (Francoeur et al. 2020). Pseudomonas and Brevundimonas isolated from the gut of bark beetles D. ponderosae and D. valens were found to degrade monoterpenes and diterpenes (Boone et al. 2013, Xu et al. 2016). Pseudomonas abietaniphila and Burkholderia xenovorans also degrade diterpenes, and they are able to utilize diterpenes as their sole carbon source (Martin and Mohn 2000, Smith et al. 2007). The intestinal bacteria of P. tsushimanus larvae in this study were also capable of utilizing monoterpenes as the sole carbon source. Despite the widespread ability of bacteria to degrade diterpenes as described in the literature, our contribution only test the monoterpene degradation (i.e., D-camphor, eucalyptol, and linalool) capacity of P. tsushimanus gut bacteria and have not yet been tested with other terpenoids, such as sesquiterpenes, and diterpenes.

Certainly, the degradation of plant secondary metabolites by intestinal symbiotic bacteria is not limited to terpenes (Berasategui et al. 2016). For instance, a strain of bacteria isolated from the foregut of Spodoptera exigua larvae can produce N-acyl amino acid hydrolases, which helps to decompose these defense-induced substances, thereby reducing the toxic effects of secondary metabolites on larvae (Ping et al. 2007). Acinetobacter in the gut of L. dispar larvae can degrade phenolic glycosides secreted by Populus tremuloides to protect the host from toxicity (Mason et al. 2016). Klebsiella and Corynebacterium in the gut of Brithys crini can degrade alkaloids in food, and Nocardioides, Gordonia, and Curtobacterium in the gut of Hyles euphorbiae can tolerate latex secreted by plants and protect the gut of insects (Vilanova et al. 2016). Likewise, intestinal bacteria that degrade plant secondary metabolites also exist in other organisms. Gut microbes of the desert woodrat Neotoma lepida degrade phenolic compounds produced by the toxic creosote bush Larrea tridentata (Kohl et al. 2014, 2016). In addition to bacteria, symbiotic fungi can also degrade plant secondary metabolites for hosts. For example, the yeast-like symbiont of the Lasioderma serricome has 1-naphthyl acetate esterase activity, which can metabolize exogenous toxic substances (Shen and Dowd 1991).

Interestingly, some intestinal microorganisms can detoxify and reutilize toxic plant secondary metabolites produced by host plants. The terpenes degraded by intestinal microorganisms might be reused by insects (DiGuistini et al. 2011). For instance, the members of the gut community of D. ponderosae (i.e., Rahnella sp., Pantoea sp., and Stenotrophomonas sp.) and D. rhyzophagus (i.e., Pseudomonas sp., Rahnella sp., and Klebsiella sp.) can fix nitrogen and provide nutrition (Morales-Jiménez et al. 2012). Our results showed that the absorption peaks of these secondary metabolites were significantly reduced and no new peaks were detected in these chromatograms. Therefore, we suspected that these gut bacteria can not only degrade D-camphor, eucalyptol, and linalool, but also reutilize decomposition products for their own growth. However, other researchers speculated that no breakdown products were detected via GC, which could be a result of complete degradation of the compounds into components of central metabolism (Marmulla and Harder 2014). Therefore, these results need to be further verified. Another interesting thing is that, the same or taxonomically related bacteria may play different roles in different insect systems and different microenvironments. In the paper, the detected bacteria included S. marcescens and Pseudomonas were considered as highly efficient terpene degrading bacteria. However, in many insect systems, some S. marcescens and Pseudomonas show extremely high insect pathogenicity. For instance, S. marcescens strain SRM and EML-SE1 showed high pathogenicity to Helicoverpa armigera and P. xylostella larvae, respectively (Jeong et al. 2010, Mohan et al. 2011). Pseudomonas fluorescens CHA0 and P. chlororaphis PCL1391 could kill insects via oral infection (Ruffner et al. 2013). We speculate that these strains may produce different proteins, including enzymes, toxin proteins, etc.

Bacterial communities are multifunctional, with multiple members performing different roles within the same species, such as bacterial communities associated with bark beetles (Six 2013). The degradation rates of eucalyptol, D-camphor, and linalool showed that the bacterial communities associated with P. tsushimanus had different degradation efficiencies in degrading terpenoids. Among the ten isolated terpenoid degradation strains, P. aeruginosa and Enterobacter sp. are effective in degradation of eucalyptol, D-camphor, and linalool, but C. variabile had the highest degradation rate of D-camphor, and the rate of linalool degradation by P. aeruginosa was the highest. S. marcescens had the highest decomposition rate of eucalyptol. This was consistent with the findings of other insect gut bacteria, such as Pseudomonas sp. and Serratia sp. of D. valens, which perform main roles in degrading α-pinene and tolerating high levels of α-pinene in vitro, and helping other microorganisms and hosts overcome the α-pinene defense of pine trees (Xu et al. 2016). Thus, it can be seen that each genus of bacteria plays different roles in detoxification. However, we only tested the degradation of D-camphor, eucalyptol, and linalool by intestinal bacteria in vitro by GC. It was also necessary to perform re-infected bioassays to a further evaluation of the biological activity of isolated monoterpene degradation strains to P. tsushimanus and verify the metabolic activities of plant secondary metabolites in vivo.

In conclusion, similar to other herbivorous insects, the intestinal microbiota of P. tsushimanus is obviously important for determining palatable plant substrates. These bacteria could degrade secondary metabolites of C. camphora to various extents, especially Pseudomonas aeruginosa with the ability to degrade these three secondary metabolites (i.e., D-camphor, eucalyptol, and linalool), which highlighted the important role of intestinal bacteria in degrading secondary metabolites of C. camphora. It must be noted that our current work is not sufficient, and in-depth studies should focus on testing the physiological role in vivo and clarifying the metabolic pathways of these bacteria in degrading secondary metabolites.

Supplementary Material

iead019_suppl_Supplementary_Material
Click here for additional data file. (343.6KB, docx)

Acknowledgments

We thank Dr Sheng-chang Lai for the help of software operation and image modification.

Contributor Information

Heng Qiao, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Han Zhu, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Hui Li, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Hongjian Chen, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Shouyin Li, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Cong Chen, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Dejun Hao, Co-Innovation Center for Sustainable Forestry in Southern China, Nanjing Forestry University, Nanjing 210037, China; College of Forestry, Nanjing Forestry University, Nanjing 210037, China.

Funding

This research was financially supported by the Science and Technology Commission of Shanghai Municipality (Grant number 18391903200), and the Shanghai Landscaping & City Appearance Administrative Bureau (Grant number G161206), Postgraduate Research & Practice Innovation Program of Jiangsu Province (Grant number KYCX23_1236), and National natural science foundation of China (Grant number 32271884).

Authors Contributions

Heng Qiao (Writing – original draft-Equal), Han Zhu (Writing – original draft-Equal), Hui Li (Data curation-Equal), Hongjian Chen (Visualization-Equal), Shouyin Li (Investigation-Equal), Cong Chen (Investigation-Equal), De-Jun Hao (Writing – review & editing-Equal)

Data Availability

The gene sequences of the A1, A2, A3, F1, F2, Z1, Z2, Z3, Z4 and Z5 strains have been deposited on Genbank database with accession numbers: ON153202 to ON153211. And these sequences can be found here: https://www.ncbi.nlm.nih.gov/nuccore/?term=ON153202:ON153211[accn].

References Cited

  1. Adams  AS, Aylward  FO, Adams  SM, Erbilgin  N, Aukema  BH, Currie  CR, Suen  G, Raffa  KF.  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. 2013:79(11):3468–3475. 10.1128/AEM.00068-13 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Afroz  M, Rahman  M, Amin  R.  Insect plant interaction with reference to secondary metabolites: a review. Agric Rev. 2021:42:427–433. [Google Scholar]
  3. Bennett  RN, Wallsgrove  RM.  Secondary metabolites in plant defence mechanisms. New Phytol. 1994:127(4):617–633. 10.1111/j.1469-8137.1994.tb02968.x [DOI] [PubMed] [Google Scholar]
  4. Berasategui  A, Axelsson  K, Nordlander  G, Schmidt  A, Borg-Karlson  AK, Gershenzon  J, Terenius  O, Kaltenpoth  M.  The gut microbiota of the pine weevil is similar across Europe and resembles that of other conifer-feeding beetles. Mol Ecol. 2016:25:4014–4031. [DOI] [PubMed] [Google Scholar]
  5. Berasategui  A, Salem  H, Paetz  C, Santoro  M, Gershenzon  J, Kaltenpoth  M, Schmidt  A.  Gut microbiota of the pine weevil degrades conifer diterpenes and increases insect fitness. Mol Ecol. 2017:26(15):4099–4110. 10.1111/mec.14186 [DOI] [PubMed] [Google Scholar]
  6. Boone  CK, Keefover-Ring  K, Mapes  AC, Adams  AS, Bohlmann  J, Raffa  KF.  Bacteria associated with a tree-killing insect reduce concentrations of plant defense compounds. J Chem Ecol. 2013:39(7):1003–1006. 10.1007/s10886-013-0313-0 [DOI] [PubMed] [Google Scholar]
  7. Chen  C, Zhang  C, Li  S, Zhu  H, Fan  B, Wang  Y, Su  P, Han  Y, Hao  D.  Biological traits and life history of Pagiophloeus tsushimanus (Coleoptera: Curculionidae), a weevil pest on camphor trees in China. J For Res. 2021:32:1979–1988. [Google Scholar]
  8. Deng  H, Zhang  J.. Analytical chemistry experiments. Wuhan: Huazhong University of Science and Technology Press. (In Chinese); 2019. [Google Scholar]
  9. Diguistini  S, Wang  Y, Liao  NY, Taylor  G, Tanguay  P, Feau  N, Henrissat  B, Chan  SK, Hesse-Orce  U, Alamouti  SM.  Genome and transcriptome analyses of the mountain pine beetle-fungal symbiont Grosmannia clavigera, a lodgepole pine pathogen. Proc Natl Acad Sci. 2011:108:2504–2509. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Divekar  PA, Narayana  S, Divekar  BA, Kumar  R, Gadratagi  BG, Ray  A, Singh  AK, Rani  V, Singh  V, Singh  AK, et al. Plant secondary metabolites as defense tools against herbivores for sustainable crop Protection. Int J Mol Sci. 2022:23(5):2690. 10.3390/ijms23052690 [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Dosdall  L, Harker  K, O’donovan  J, Blackshaw  R, Kutcher  H, Gan  Y, Johnson  E.  Crop sequence effects on root maggot (Diptera: Anthomyiidae: Delia spp.) infestations in canola. J Econ Entomol. 2012:105:1261–1267. [DOI] [PubMed] [Google Scholar]
  12. Engel  P, Moran  NA.  The gut microbiota of insects-diversity in structure and function. FEMS Microbiol Rev. 2013:37(5):699–735. 10.1111/1574-6976.12025 [DOI] [PubMed] [Google Scholar]
  13. FöRster-Fromme  K, HöSchle  B, Mack  C, Bott  M, Armbruster  W, Jendrossek  D.  Identification of genes and proteins necessary for catabolism of acyclic terpenes and leucine/isovalerate in Pseudomonas aeruginosa. Appl Environ Microbiol. 2006:72(7):4819–4828. 10.1128/AEM.00853-06 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Francoeur  CB, Khadempour  L, Moreira-Soto  RD, Gotting  K, Book  AJ, Pinto-Tomás  AA, Keefover-Ring  K, Currie  CR.  Bacteria contribute to plant secondary compound degradation in a generalist herbivore system. MBio. 2020:11(5):e02146–e02120. 10.1128/mBio.02146-20 [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Hammer  TJ, Bowers  MD.  Gut microbes may facilitate insect herbivory of chemically defended plants. Oecologia. 2015:179(1):1–14. 10.1007/s00442-015-3327-1 [DOI] [PubMed] [Google Scholar]
  16. Hehemann  J-H, Correc  G, Barbeyron  T, Helbert  W, Czjzek  M, Michel  G.  Transfer of carbohydrate-active enzymes from marine bacteria to Japanese gut microbiota. Nature. 2010:464(7290):908–912. 10.1038/nature08937 [DOI] [PubMed] [Google Scholar]
  17. Huang  J, Wu  S, Gao  L, Yang  L.  Diagnosis and damage of weevil pest Pagiophloeus tsushimanus on camphor tree. J Zhejiang A F Univ. 2014:31:764–767. In Chinese. [Google Scholar]
  18. Jamwal  K, Bhattacharya  S, Puri  S.  Plant growth regulator mediated consequences of secondary metabolites in medicinal plants. J Appl Res Med Aromat Plants. 2018:9:26–38. [Google Scholar]
  19. Jaramillo  J, Borgemeister  C, Baker  P.  Coffee berry borer Hypothenemus hampei (Coleoptera: Curculionidae): searching for sustainable control strategies. Bull Entomol Res. 2006:96(3):223–233. 10.1079/ber2006434 [DOI] [PubMed] [Google Scholar]
  20. Jeong  HU, Mun  HY, Oh  HK, Kim  SB, Yang  KY, Kim  I, Lee  HB.  Evaluation of insecticidal activity of a bacterial strain, Serratia sp. EML-SE1 against diamondback moth. J Microbiol. 2010:48(4):541–545. 10.1007/s12275-010-0221-9 [DOI] [PubMed] [Google Scholar]
  21. Jiang  H, Wang  J, Song  L, Cao  X, Yao  X, Tang  F, Yue  Y.  GC× GC-TOFMS analysis of essential oils composition from leaves, twigs and seeds of Cinnamomum camphora L. Presl and their insecticidal and repellent activities. Molecules. 2016:21(4):423. 10.3390/molecules21040423 [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Kaltenpoth  M, Engl  T.  Defensive microbial symbionts in Hymenoptera. Funct Ecol. 2014:28:315–327. [Google Scholar]
  23. Kalyaanamoorthy  S, Minh  BQ, Wong  TK, Von Haeseler  A, Jermiin  LS.  ModelFinder: fast model selection for accurate phylogenetic estimates. Nat Methods. 2017:14:587–589. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Kikuchi  Y, Hayatsu  M, Hosokawa  T, Nagayama  A, Tago  K, Fukatsu  T.  Symbiont-mediated insecticide resistance. Proc Natl Acad Sci. 2012:109:8618–8622. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Kohl  KD, Stengel  A, Dearing  MD.  Inoculation of tannin-degrading bacteria into novel hosts increases performance on tannin-rich diets. Environ Microbiol. 2016:18(6):1720–1729. 10.1111/1462-2920.12841 [DOI] [PubMed] [Google Scholar]
  26. Kohl  KD, Weiss  RB, Cox  J, Dale  C, Denise Dearing  M.  Gut microbes of mammalian herbivores facilitate intake of plant toxins. Ecol Lett. 2014:17(10):1238–1246. 10.1111/ele.12329 [DOI] [PubMed] [Google Scholar]
  27. Li  S, Li  H, Wang  J, Chen  C, Hao  D.  Hormetic response and co-expression of cytochrome P450 and cuticular protein reveal the tolerance to host-specific terpenoid defences in an emerging insect pest, Pagiophloeus tsushimanus (Coleoptera: Curculionidae). J Pest Sci. 2023:96:141–160. [Google Scholar]
  28. Li  S, Wang  J, Chen  C, Li  H, Hao  D.  Tolerance, biochemistry and related gene expression in Pagiophloeus tsushimanus (Coleoptera: Curculionidae) exposed to chemical stress from headspace host-plant volatiles. Agric For Entomol. 2022:24:189–203. [Google Scholar]
  29. Lüddeke  F, Dikfidan  A, Harder  J.  Physiology of deletion mutants in the anaerobic β-myrcene degradation pathway in Castellaniella defragrans. BMC Microbiol. 2012:12:1–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Marmulla  R, Harder  J.  Microbial monoterpene transformations—a review. Front Microbiol. 2014:5:346. 10.3389/fmicb.2014.00346 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Martin  VJ, Mohn  WW.  Genetic investigation of the catabolic pathway for degradation of abietane diterpenoids by Pseudomonas abietaniphila BKME-9. J Bacteriol. 2000:182(13):3784–3793. 10.1128/JB.182.13.3784-3793.2000 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Mason  CJ, Couture  JJ, Raffa  KF.  Plant-associated bacteria degrade defense chemicals and reduce their adverse effects on an insect defoliator. Oecologia. 2014:175(3):901–910. 10.1007/s00442-014-2950-6 [DOI] [PubMed] [Google Scholar]
  33. Mason  CJ, Lowe-Power  TM, Rubert-Nason  KF, Lindroth  RL, Raffa  KF.  Interactions between bacteria and aspen defense chemicals at the phyllosphere–herbivore interface. J Chem Ecol. 2016:42(3):193–201. 10.1007/s10886-016-0677-z [DOI] [PubMed] [Google Scholar]
  34. Mohan  M, Selvakumar  G, Sushil  SN, Bhatt  JC, Gupta  HS.  Entomopathogenicity of endophytic Serratia marcescens strain SRM against larvae of Helicoverpa armigera (Noctuidae: Lepidoptera). World J Microbiol Biotechnol. 2011:27(11):2545–2551. 10.1007/s11274-011-0724-4 [DOI] [Google Scholar]
  35. Morales-Jiménez  J, Zúñiga  G, Ramírez-Saad  HC, Hernández-Rodríguez  C.  Gut-associated bacteria throughout the life cycle of the bark beetle Dendroctonus rhizophagus Thomas and Bright (Curculionidae: Scolytinae) and their cellulolytic activities. Microb Ecol. 2012:64(1):268–278. 10.1007/s00248-011-9999-0 [DOI] [PubMed] [Google Scholar]
  36. Moran  NA, Plague  GR, Sandström  JP, Wilcox  JL.  A genomic perspective on nutrient provisioning by bacterial symbionts of insects. Proc Natl Acad Sci. 2003:100:14543–14548. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Morimoto  K.  The family curculionidae of Japan. I. Subfamily hylobxinae. ESAKIA.  1982:19:51–121. [Google Scholar]
  38. Nguyen  L-T, Schmidt  HA, Von Haeseler  A, Minh  BQ.  IQ-TREE: a fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Mol Biol Evol. 2015:32(1):268–274. 10.1093/molbev/msu300 [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Paniagua Voirol  LR, Frago  E, Kaltenpoth  M, Hilker  M, Fatouros  NE.  Bacterial symbionts in Lepidoptera: their diversity, transmission, and impact on the host. Front Microbiol. 2018:9:556. 10.3389/fmicb.2018.00556 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Petersson  M, Örlander  G.  Effectiveness of combinations of shelterwood, scarification, and feeding barriers to reduce pine weevil damage. Can J For Res. 2003:33(1):64–73. 10.1139/x02-156 [DOI] [Google Scholar]
  41. Ping  L, Büchler  R, Mithöfer  A, Svatoš  A, Spiteller  D, Dettner  K, Gmeiner  S, Piel  J, Schlott  B, Boland  W.  A novel Dps-type protein from insect gut bacteria catalyses hydrolysis and synthesis of N-acyl amino acids. Environ Microbiol. 2007:9(6):1572–1583. 10.1111/j.1462-2920.2007.01279.x [DOI] [PubMed] [Google Scholar]
  42. Prakash  O, Kumari  K, Lal  R.  Pseudomonas delhiensis sp. nov., from a fly ash dumping site of a thermal power plant. Int J Syst Evol Microbiol. 2007:57(Pt 3):527–531. 10.1099/ijs.0.64456-0 [DOI] [PubMed] [Google Scholar]
  43. Ronquist  F, Teslenko  M, Van Der Mark  P, Ayres  DL, Darling  A, Höhna  S, Larget  B, Liu  L, Suchard  MA, Huelsenbeck  JP.  MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol. 2012:61(3):539–542. 10.1093/sysbio/sys029 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Rosen  R, Kanakala  S, Kliot  A, Pakkianathan  BC, Farich  BA, Santana-Magal  N, Elimelech  M, Kontsedalov  S, Lebedev  G, Cilia  M.  Persistent, circulative transmission of begomoviruses by whitefly vectors. Curr Opin Virol. 2015:15:1–8. [DOI] [PubMed] [Google Scholar]
  45. Ruffner  B, Péchy-Tarr  M, Ryffel  F, Hoegger  P, Obrist  C, Rindlisbacher  A, Keel  C, Maurhofer  M.  Oral insecticidal activity of plant-associated pseudomonads. Environ Microbiol. 2013:15:751–763. [DOI] [PubMed] [Google Scholar]
  46. Shen  P, Chen  X.. Microbiology experiments. 5th ed. Beijing: Higher Education Press. (In Chinese); 2018. [Google Scholar]
  47. Shen  SK, Dowd  PF.  1-Naphthyl acetate esterase activity from cultures of the symbiont yeast of the cigarette beetle (Coleoptera: Anobiidae). J Econ Entomol. 1991:84(2):402–407. 10.1093/jee/84.2.402 [DOI] [Google Scholar]
  48. Six  DL.  The bark beetle holobiont: why microbes matter. J Chem Ecol. 2013:39(7):989–1002. 10.1007/s10886-013-0318-8 [DOI] [PubMed] [Google Scholar]
  49. Smith  DJ, Park  J, Tiedje  JM, Mohn  WW.  A large gene cluster in Burkholderia xenovorans encoding abietane diterpenoid catabolism. J Bacteriol. 2007:189(17):6195–6204. 10.1128/JB.00179-07 [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Tudroszen  NJ, Kelly  DP, Millis  NF.  alpha-Pinene metabolism by Pseudomonas putida. Biochem J. 1977:168(2):315–318. 10.1042/bj1680315 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Vilanova  C, Baixeras  J, Latorre  A, Porcar  M.  The generalist inside the specialist: gut bacterial communities of two insect species feeding on toxic plants are dominated by Enterococcus sp. Front Microbiol. 2016:7:1005. 10.3389/fmicb.2016.01005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Wu  M, Ni  L, Lu  H, Xu  H, Zou  S, Zou  X.  Terpenoids and their biological activities from Cinnamomum: a review. J Chem. 2020:2020:1–14. [Google Scholar]
  53. Xia  X, Gurr  GM, Vasseur  L, Zheng  D, Zhong  H, Qin  B, Lin  J, Wang  Y, Song  F, Li  Y, et al. Metagenomic sequencing of diamondback moth gut microbiome unveils key holobiont adaptations for herbivory. Front Microbiol. 2017:8:663. 10.3389/fmicb.2017.00663 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Xu  LT, Lu  M, Sun  JH.  Invasive bark beetle-associated microbes degrade a host defensive monoterpene. Insect Sci. 2016:23(2):183–190. 10.1111/1744-7917.12255 [DOI] [PubMed] [Google Scholar]
  55. Yang  E-J, Park  Y-J, Chang  H-C.  Cloning of four genes involved in limonene hydroxylation from Enterobacter cowanii 6L. J Microbiol Biotechnol. 2007:17:1169–1176. [PubMed] [Google Scholar]
  56. Zhang  D, Gao  F, Jakovlić  I, Zou  H, Zhang  J, Li  WX, Wang  GT.  PhyloSuite: an integrated and scalable desktop platform for streamlined molecular sequence data management and evolutionary phylogenetics studies. Mol Ecol Resour. 2020:20(1):348–355. 10.1111/1755-0998.13096 [DOI] [PubMed] [Google Scholar]
  57. Zhang  Y, Lyu  B, Yang  F, Tu  Y, Jiang  F, Qi  K, Li  Z.  Community structure analyses of the intestine cultivable bacteria of Clostera restitura larvae. J Trop Crops  2021:42:1066–1070. In Chinese. [Google Scholar]

Associated Data

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

Supplementary Materials

iead019_suppl_Supplementary_Material
Click here for additional data file. (343.6KB, docx)

Data Availability Statement

The gene sequences of the A1, A2, A3, F1, F2, Z1, Z2, Z3, Z4 and Z5 strains have been deposited on Genbank database with accession numbers: ON153202 to ON153211. And these sequences can be found here: https://www.ncbi.nlm.nih.gov/nuccore/?term=ON153202:ON153211[accn].

Articles from Journal of Insect Science are provided here courtesy of University of Wisconsin Libraries

RESOURCES