Skip to main content
NCBI home page
Journal of Insect Science logoLink to Journal of Insect Science
. 2026 Mar 12;26(2):ieag017. doi: 10.1093/jisesa/ieag017

Identification, expression profiling, and functional characterization of trehalase genes (OaTRE1 and OaTRE) in Oedaleus asiaticus (Orthoptera: Acridoidea)

Mei Liu 1, Feng Yan 2, Ruiwen Dong 3, Haiyan Gao 4, Yidan Wu 5, Kejian Lin 6,✉, Shujing Gao 7,✉
Editor: Amr Mohamed
PMCID: PMC12979043  PMID: 41814554

Abstract

To understand the roles of trehalase-encoding genes (OaTRE1 and OaTRE) in Oedaleus asiaticus, the OaTRE1 and OaTRE genes were identified and analyzed. Real-time PCR was used to analyze their spatial and temporal expression patterns, while RNA interference (RNAi) was applied to explore their functions in growth and development. OaTRE1 and OaTRE possess open reading frames of 1,587 and 1,800 bp, encoding 528 and 599 amino acids, respectively. Both genes were differentially expressed across developmental stages and tissues. RNAi-mediated knockdown of OaTRE1 and OaTRE significantly reduced the survival rate of 4th instar nymphs of Oedaleus asiaticus at 96 h post-injection. RNAi-mediated knockdown of OaTRE1 and OaTRE significantly reduced chitin content in Oedaleus asiaticus. Gene expression was upregulated by injections of 20-hydroxyecdysone and downregulated by validamycin, suggesting that trehalase genes are primarily involved in morphogenesis during metamorphosis and adulthood. These findings highlight trehalase as a critical regulator of insect development and a promising molecular target for the development of eco-friendly strategies to control the destructive grassland pest Oedaleus asiaticus.

Keywords: Oedaleus asiaticus, trehalase, RNA interference, metamorphosis

Introduction

Chitin, a nitrogen-containing polysaccharide biogenic macromolecule, is found in a wide variety of organisms, including bacteria, yeast, fungi, insects, invertebrates, and lower and higher plants. It exists in the exoskeletons of arachnids and crustaceans, the cell walls of some fungi, and is also an important component of the insect body (Merzendorfer 2006, Rabadiya and Behr 2024).

As a polymer of N-acetyl-β-d-glucosamine (Merzendorfer and Zimoch 2003), chitin is present in the insect epidermis, trachea, and intestinal epithelium, and serves a as major structural component of the exoskeleton involved in maintaining body size and protecting against external mechanical disturbances (Hegedus et al. 2009, Moussian 2010, Kelkenberg et al. 2015).

Since chitin constitutes a vital part of the periodically shed epidermal exoskeleton and peritrophic membrane, its metabolism is closely associated with cuticle renewal and peritrophic substrate remodeling during insect development, making it an ideal target for agricultural pest control (Candy and Kilby 1962, Zhu et al. 2016). Insect chitin synthesis involves complex biochemical and physiological processes with multiple enzymes, initiating with trehalose as the substrate and concluding with chitin.

Trehalose, a non-reducing disaccharide and the main hemolymph sugar in insects, provides energy (Kramer and Koga 1986), supplies carbon for chitin biosynthesis, serves as an energy source for flight, regulates feeding behavior, and enhances environmental stress tolerance (Evans and Dethier 1957, Shukla et al. 2015, Yasugi et al. 2017). Hydrolyzed by trehalase into glucose, trehalose provides energy (Argüelles 2000) and the upstream raw material for chitin synthesis. Insects express two trehalase types: soluble trehalase (TRE1), a cytosolic enzyme for endogenous trehalose catabolism mainly distributed in the circulatory and digestive systems (Takiguchi et al. 1992, Terra and Ferreira 1994, Becker et al. 1996, Thompson 2003), and membrane-bound trehalase (TRE2), an extracellular enzyme for exogenous trehalose absorption highly expressed in metabolically active tissues like fat bodies (Tang et al. 2008, Chen et al. 2010, Zhang et al. 2011).

Both 20-hydroxyecdysone (20E) and validamycin (VA) are compounds that interfere with the hormonal regulation of insects. 20E induces molting in insects and modulates chitinase synthesis, thereby exerting a pivotal influence on insect growth and development. As a key regulator throughout the insect life cycle, it drives molting and metamorphosis processes (JUDITH 2015, MIURA 2019). Validamycin an antibiotic isolated from fungi, exhibits potent competitive inhibitory activity against trehalase across a broad spectrum of organisms (Takahashi et al. 1995, Tatun et al. 2014). Notably, validamycin can suppress trehalase activity in both fungi and insects, consequently blocking the hydrolysis of trehalose into glucose (Muller et al. 1995).

The Mongolian locust, Oedaleus asiaticus, is a destructive pest distributed in northern Chinese grasslands and southern Russia’s Transbaikal region, primarily feeding on gramineous plants such as Leymus chinensis and Stipa grandis in Inner Mongolia (Ritchie 1981, Guan and Wei 1989, Kang and Chen 1994, Feng et al. 1995, Huang et al. 2017). Swarming O. asiaticus severely damages early crops, reducing yields, accelerating grassland desertification, disrupting ecological balance, and causing heavy losses to agriculture and animal husbandry (over 50% crop loss in severe cases). Current control relies heavily on chemical insecticides, leading to increasing resistance and an urgent need for environmentally friendly alternatives.

Given the potential of chitin metabolism key enzymes as novel insecticide targets, trehalase-critical for energy supply and chitin synthesis-emerges as a promising candidate. To date, trehalase genes have been successfully cloned from 40 insect species, with representative examples including Drosophila melanogaster, Bombyx mori, Tribolium castaneum, and Locusta migratoria manilensis (Chen and Zhang 2015, Xiong et al. 2016, Chen et al. 2018, Tellis et al. 2023).

This study focused on trehalase as a key component of the chitin synthesis pathway in Oedaleus asiaticus. Specific RNA interference (RNAi) was employed to silence trehalase expression, leading to disruption of trehalose metabolism, impairment of chitin biosynthesis, interference with molting, and ultimately increased insect mortality. These findings provide a theoretical basis for the development of novel, trehalase-targeted insecticides and offer new insights into effective strategies for the management and control of O. asiaticus.

Materials and Methods

Insect Rearing and Sample Preparation

Eggs of O. asiaticus were collected from the grasslands of Siziwang Banner, Inner Mongolia, China. The eggs were incubated in the laboratory (Hohhot, China) and the insects were reared in a growth chamber with fresh wheat seedlings and bran. The room temperature was maintained at 25 ± 2 °C, with a relative humidity of 40 ± 5% and a photoperiod of 14 h light and 10 h dark. Insects at different developmental stages were used for the experiments. 1st- to 5th-instar nymphs, adult males, and adult females were collected. Additionally, the fat body, Malpighian tubules, foregut, midgut, hindgut, salivary glands, trachea, and epidermal tissues of 4th-instar nymphs were dissected, immediately frozen in liquid nitrogen, and stored at −80°C for future use.

Identification and Sequencing of OaTRE1 and OaTRE cDNA

Total RNA was extracted from the 4th instar nymphs of O. asiaticus using the Eastep Super Total RNA Extraction Kit (Promega, USA), according to the manufacturer’s instructions. The concentration and integrity of the RNA were evaluated using a spectrophotometer and 1% agarose gel electrophoresis, respectively. First-strand cDNA was synthesized following the protocol provided with the reverse transcription kit (Vazyme Biotech, Nanjing, China).

Analysis of the O.asiaticus transcriptome database (NCBI BioProject PRJNA1264109) identified two cDNA sequences, designated OaTRE1 and OaTRE, which appear to originate from two distinct trehalase-encoding genes. Specific primers were designed using Primer Premier 5.0 software to amplify the coding sequences of the candidate OaTRE1 and OaTRE genes. The 25 µL PCR reaction mixture consisted of 1 µL of cDNA template, 12.5 µL of Green Tag Mix, 1 µL each of forward and reverse primers, and 9.5 µL of ddH2O. The PCR reaction protocol was as follows: initial denaturation at 94 °C for 3 min; 40 cycles of denaturation at 94 °C for 15 s, annealing at 65 °C for 30 s, and extension at 72 °C for 1 min; followed by a final extension at 72 °C for 10 min.

After evaluation using 1% agarose gel electrophoresis, the amplified products were purified by extraction of the target bands from the gel and their subsequent ligation into the pMD19-T vector. The ligation products were transformed into Escherichia coli DH5α competent cells. Following blue-white screening and PCR identification, positive clonal bacterial suspensions were selected and sent to Beijing AuGCT Biotechnology Co., Ltd. (Beijing, China) for bidirectional sequencing. The sequences were aligned and verified against the transcriptome sequences using DNAMAN 6.0 software.

Analysis of OaTRE1 and OaTRE cDNA and Prediction of the Amino Acid Sequences

The ExPASy proteomics website (http://cn.expasy.org/) was used for the prediction of the amino acid sequences from the cDNA, as well as prediction of the molecular weight, isoelectric point (pI), and presence of a signal peptide and transmembrane regions.

Open reading frames (ORFs) were identified using the National Center for Biotechnology Information (NCBI) website (http://www.ncbi.nlm.nih.gov/), and BLASTP was used to assess sequence similarities and homology of the two OaTREs with TRE sequences from other insects. The OaTRE1 and OaTRE sequences were aligned with those of L. migratoria (LmTRE1 and LmTRE) using DNAMAN 6.0 software. Phylogenetic analysis was performed using the Neighbor-Joining (NJ) method in MEGA 6.06. Bootstrap support values were calculated with 1,000 replicates. Primer sequences and amplification characteristics for cDNA cloning and qPCR analysis are provided in Supplementary Table S1, while the sequence features of the two trehalase genes (OaTRE1 and OaTRE), including ORF length, amino acid number, molecular weight, and isoelectric point, are summarized in Supplementary Table S2.

Tissue and Developmental Expression of the Two OaTRE Genes

The mRNA expression levels of OaTRE1 and OaTRE were examined across different developmental stages (1st to 5th instar nymphs, adult males and females) and in various tissues of 4th-instar nymphs, including the fat body, Malpighian tubules, foregut, midgut, hindgut, salivary glands, trachea, and epidermis. Total RNA was isolated from each sample using TRIzol Reagent (Invitrogen, Carlsbad, California, United States), treated with DNase I to eliminate genomic DNA contamination, and reverse-transcribed into cDNA using the HiScript III RT SuperMix (Vazyme Biotech, Nanjing, China). For each experimental condition, at least three independent biological replicates—each derived from a separate cohort of insects—were analyzed to account for biological variability.

Quantitative real-time PCR (qRT-PCR) was performed in a 20 µL reaction mixture containing 2 µL of diluted cDNA template, 0.4 µL each of forward and reverse primers (10 µM), 10 µL of SYBR Green qPCR Master Mix (Vazyme Biotech, Nanjing, China), and 7.2 µL of nuclease-free water.

The thermal cycling program was as follows: an initial denaturation at 95 °C for 5 min, followed by 40 cycles of denaturation at 95 °C for 15 s and combined annealing/extension at 60 °C for 30 s. A melting curve analysis was subsequently performed by heating from 65 to 95 °C with a ramp rate of 0.5 °C every 5 s to verify amplicon specificity. All primer pairs produced a single, sharp peak in the melting curve, and agarose gel electrophoresis (1.5%) confirmed a single band of the expected size.

Primer amplification efficiencies were determined using a 5-point, 10-fold serial dilution of pooled cDNA. Standard curves exhibited excellent linearity for both target genes, with R2 =0.983 for OaTRE1 (amplification efficiency = 115.152%) and R2=0.977 for OaTRE (amplification efficiency = 102.109%). Relative gene expression was calculated using the 2−ΔΔCt method, with β-actin as the internal reference gene (Zhou 2019, Zheng et al. 2025).

Functional Analysis of OaTRE1 and OaTRE Using RNAi

To investigate the roles of OaTRE1 and OaTRE in locust development, 4th-instar nymphs were selected. Prior to injection, the integrity of dsRNA was verified by agarose gel electrophoresis (Supplementary Fig. S1). Subsequently, 5 µL of dsOaTRE1 and dsOaTRE (1,000 ng/µL) were respectively injected into the intersegmental membrane between the 2nd and 3rd abdominal segments of O. asiaticus.

During injection, the syring needle was kept as parallel as possible to the abdominal cuticle to minimize tissue damage, and dsRNA or dsGFP solution was slowly injected into the hemocoel. An equal volume (5 µL) of dsGFP (1,000 ng/µL, total dose of 5 µg per insect) was injected as a negative control. Three biological replicates were set up, with 20 nymphs per treatment group. At 12, 24, 48, and 72 h post dsRNA injection, the nymphs were collected, and the expression profiles of the target genes were analyzed by RT-qPCR. Additionally, phenotypic changes of the nymphs were observed and recorded daily.

20-hydroxyecdysone and Validamycin Treatment

To assess the effects of 20-hydroxyecdysone (20E) and validamycin (VA) on the expression of chitin synthesis-related genes in O. asiaticus, batches of healthy 4th-instar nymphs with similar body size were selected. The test concentrations of 20E and validamycin were set as 2.5, 5, 10, and 20 µg/µL with reference to the experimental methodologies reported by Tang et al. (2017) and Wang (2019), and 5 µL of each concentration solution was injected into the intersegmental membrane between the 2nd and 3rd abdominal segments of O. asiaticus, consistent with the aforementioned injection site.

Gene expression levels were analyzed at 12, 24, 48, and 72 h post-20E treatment, and at 12, 24, and 48 h validamycin treatment. For the control settings, 10% ethanol was used as the solvent for 20E, and the corresponding control group was injected with 5 μL of 10% ethanol; ddH2O was used as the solvent for validamycin, with the control group injected with 5 μL of ddH2O.

Determination of Chitin Content after RNA Interference (RNAi)

Chitin content in 4th-instar nymphs O. asiaticus nymphs was quantified indirectly via a colorimetric assay based on the detection of N-acetylglucosamine (GlcNAc), the monomeric unit released upon acid hydrolysis of chitin, following a modified protocol from Su (2017). Whole-body samples were hydrolyzed in 6 M HCl at 100 °C for 2 h to release GlcNAc (molecular weight = 221.21 g/mol). After neutralization and filtration, the hydrolysates were used for GlcNAc quantification.

A standard curve was prepared using known concentrations of GlcNAc (0 to 2.5 mmol/L). For each standard or sample, 10 μL of sodium borate buffer was added to 10 μL of GlcNAc solution or sample hydrolysate, followed by heating at 99.9 °C for 10 min to ensure complete conversion. The mixture was rapidly cooled on ice, then mixed with 100 μL of 1× p-dimethylaminobenzaldehyde (DMAB) reagent (dissolved in glacial acetic acid). The reaction was incubated at 37 °C for 20 min, and absorbance was measured at 585 nm using a microplate reader. Chitin content was calculated by converting GlcNAc concentration (from the standard curve) into chitin equivalents based on molecular weights (chitin: [C8H13NO5]ₙ; GlcNAc: C8H15NO6).

Statistical Analysis

Significant differences among experimental groups were evaluated using analysis of variance (ANOVA) for datasets meeting parametric assumptions; when a significant overall effect was detected, post-hoc multiple comparisons were further conducted via Tukey’s honestly significant difference (HSD) test, while independent-samples t-test was applied for pairwise comparisons between two groups. Survival analysis was performed using Kaplan–Meier curves, with intergroup comparisons conducted via the log-rank (Mantel-Cox) test. All data are presented as mean ± standard deviation (SD) derived from at least three independent biological replicates. Statistical significance was defined as P < 0.05.). Results were plotted using GraphPad Prism 10 (GraphPad Software, San Diego, California, United States).

Results

Characterization of Two TRE cDNAs and Their Deduced Amino Acid Sequences

A comprehensive search of the transcriptome database of O. asiaticus (GeneBank accession number PRJNA1264109) in our laboratory was carried out using “TRE” as the keyword to identify the ORFs. The trehalase genes were cloned and amplified by PCR. It was found that the ORFs of OaTRE1 and OaTRE of O. asiaticus were 1,587 and 1,800 bp in length, respectively, encoding 528 and 599 amino acids (Fig. 1).

Fig. 1.

Graph showing the multiple sequence alignment of amino acid sequences for OaTRE1, OaTRE, LmTRE1, and LmTRE proteins, with residues sharing &gt;80% similarity shaded gray and identical residues in black to highlight conserved functional domains across all four sequences.

Open in a new tab

Alignment of the amino acid sequences of OaTRE1, OaTRE, LmTRE1, and LmTRE. Numbers on the right side indicate the number of last residues in a line of TRE sequence. Residues that share more than 80% amino acid similarity are shaded in grey and identical residues are shown in black.

3.2. Phylogenetic Analysis

The amino acid sequences encoded by OaTRE1 and OaTRE were compared using BLAST, and the results showed that OaTRE1 and OaTRE had strong similarity with the trehalase sequences of L. migratoria manilensis, also a member of the locust family, with 99% and 100% similarity, respectively. The sequences of TRE proteins from other insect species were downloaded from the NCBI database and were used to construct a neighbor-joining phylogenetic tree using MEGA 6.06, as shown in Fig. 2.

Fig. 2.

Phylogenetic tree showing evolutionary relationships of trehalase proteins from Oedaleus asiaticus and orthopteran insects, with color-coded bootstrap support values and genetic distances labeled on branches.

Open in a new tab

Phylogenetic analysis of trehalase (TRE) proteins from Oedaleus asiaticus and orthopteran insects. The tree was constructed in MEGA 6.06 using the Neighbor-Joining method. Branch support was evaluated through bootstrap analysis with 1,000 replicates. Colored dots at the nodes represent bootstrap support values: red (>95%), blue (70% to 94%), and green (<70%). The values beside the branches indicate genetic distances (number of substitutions per site). Key sequences included in the analysis: Oedaleus asiaticus OaTRE1 and OaTRE; Locusta migratoria LmTRE1 (GenBank: ACP28173.1) and LmTre (GenBank: AQV08185); Calliptamus italicus CiTreM1 (GenBank: UNW45414.1); Schistocerca gregaria SgTRE (GenBank: QVD39550.1); Schistocerca piceifrons SpTRE-like (GenBank: XP_047117343); Schistocerca nitens SnTRE (GenBank: XP_049815564); Schistocerca cancellata ScTRE (GenBank: XP_049787841); Schistocerca americana SaTRE (GenBank: XP_046986626) Schistocerca serialisSsTRE(GenBank: XP_049963661); Schistocerca cancellata ScTRE1 (GenBank: XP_049787841); Schistocerca piceifrons SpTRE1 (GenBank: XP_047117343.1); Schistocerca serialis SsTRE1 (GenBank: XP_049963667); Schistocerca nitens SnTRE1 (XP_049815564.1); Schistocerca gregaria SgTRE1 (GenBank: XP_049830471); Schistocerca serialis SsTRE1 (GenBank: XP_049963667.1).

The trehalase protein sequences of O. asiaticus were found to be most closely related to those of LmTre, which is also a member of the Orthoptera. Specifically, OaTRE clustered with the TREs of LmTre on a single branch, and OaTRE1 clustered on the same branch as LmTRE1 in a single branch, with more distant relationships with the proteins of other locusts, consistent with the traditional locust taxonomy.

Expression Profiles of OaTRE and OaTRE1 in Different Tissues and Ages

Expression in Different Tissues

The relative mRNA expression levels of OaTRE1 and OaTRE in different tissues of O. asiaticus were measured by qRT-PCR (Fig. 3). The results showed that expression patterns differed among the different tissues. OaTRE1 showed the highest expression in the integument (IN), followed by the Malpighian tubules (MT) and trachea (TR), and the lowest expression in the foregut (FG), hindgut (HG), and salivary glands (SG), while OaTRE was expressed most strongly in the Malpighian tubules (MT) and fat body (FB), with the lowest expression in the salivary glands (SG).

Fig. 3.

Bar graphs showing relative expression levels of TRE1 (panel A) and TRE (panel B) across different tissues of O. asiaticus, with significance markers.

Open in a new tab

Relative expression levels of TRE1 A) and TRE B) in different tissues of O. asiaticus. Data are shown as mean ± SD from three biological replicates (n = 3). Statistical significance was determined by one-way ANOVA followed by Tukey’s HSD test (P < 0.05). Bars with different lowercase letters indicate significant differences among groups. FB, fat body; MT, Malpighian tubule; FG, foregut; MG, midgut; HG, hindgut; SG, salivary gland; TR, trachea; IN, integument.

Expression at Different Developmental Stages

The expression profiles of OaTRE and OaTRE1 in O. asiaticus at different developmental stages (1st-instar, 2nd-instar, 3rd-instar, 4th-instar, 5th-instar nymphs, adult females, and adult males) were examined by qRT-PCR (Fig. 4). The results showed that OaTRE1 was expressed in different developmental stages of locusts, with the highest expression in adult female locusts, followed by 5th-instar nymphs and adult males, and lowest in 3rd-instar nymphs. OaTRE expression was highest in adult males and lowest in 3rd-instar nymphs.

Fig. 4.

Bar graphs showing relative expression levels of TRE1 (panel A) and TRE (panel B) across developmental stages of O. asiaticus, with significance markers.

Open in a new tab

Relative expression levels of TRE1 A) and TRE B) at different developmental stages of O. asiaticus. Data are shown as mean ± standard deviation (SD) from three biological replicates (n = 3). Statistical significance was determined by one-way ANOVA followed by Tukey’s honestly significant difference (HSD) test (P < 0.05). Bars labeled with different lowercase letters indicate statistically significant differences among stages. EG, egg; N1 to N5, 1st to 5th instar nymphs; AF, adult female; AM, adult male.

Knockdown of OaTRE1 and OaTRE

To further verify the functions of the genes, RNAi-mediated knockdown was utilized to investigate the roles of these genes in the growth and development of O. asiaticus. The dsRNA of these genes was injected into 4th-instar nymphs of O. asiaticus, and gene expression levels were determined at 12, 24, 48, and 72 h post-treatment using qRT-PCR. The results indicated significant reductions in the mRNA levels of both genes with no effects on the expression of other genes. After injection of OaTRE1 dsRNA, expression of the gene dropped to its lowest level, 69.98%, after 48 h, while expression of OaTRE declined to its lowest level of 56.52% at 72 h (Fig. 5A).

Fig. 5.

Graphs and data showing effects of TRE gene RNAi on Oedaleus asiaticus, including time-course expression (A), phenotypic outcomes (B), chitin content changes (C), survival curves (D), and deformity rates (E), with statistical significance markers.

Open in a new tab

Effects of TRE Gene RNAi on the Growth and Development of Oedaleus asiaticus. (A) Time-course expression profiles of OaTRE1 and OaTRE following dsRNA injection. Relative mRNA levels were quantified by qRT-PCR at 12, 24, 48, and 72 h post-injection, normalized to the dsGFP control group and presented as mean ± standard deviation (SD) from three biological replicates. Statistical significance was determined by one-way ANOVA followed by Tukey’s honestly significant difference (HSD) test. Asterisks denote significant differences between treatment groups at each time point: ****P < 0.0001. B) Representative phenotypes of Oedaleus asiaticus after dsRNA injection. Scale bars: 0.5 cm. C) Changes in chitin content in Oedaleus asiaticus before and after RNAi of OaTRE1 and OaTRE. Data are shown as mean ± SD (n = 3 independent biological replicates). Asterisks denote significant differences between pre-RNAi and post-RNAi groups (paired t-test, P < 0.0001). D) Kaplan–Meier survival analysis of Oedaleus asiaticus injected with dsGFP (blue), dsOaTRE1 (red), and dsOaTRE (green) over 96 h. Shaded areas represent 95% confidence intervals. Vertical ticks indicate censored observations (animals alive at 96 h). Numbers at risk at each time point are shown below the x-axis. Statistical significance determined by Log-rank test: x2 = 84.91, df = 2, p = 1.1 × 10−24; Gehan-Breslow test: x2 = 79.49, df = 2, p = 1.3 × 10−23. E) Proportion of malformed nymphs following dsTRE1 injection (n = 3 independent biological replicates, 20 nymphs per replicate, random sex ratio).

These severe developmental and survival defects prompted us to examine whether the observed phenotypes were linked to disruptions in chitin metabolism. To investigate the functional roles of OaTRE1 and OaTRE in chitin metabolism, 4th-instar nymphs O. asiaticus nymphs were injected with gene-specific dsRNA, and whole-body chitin content was quantified using a colorimetric assay based on GlcNAc release. As shown in Fig. 5C, RNAi-mediated knockdown of both OaTRE1 and OaTRE resulted in significant reductions in total chitin content relative to pre-injection levels. Specifically, silencing of OaTRE1 led to an approximately 83% decrease in chitin content-the most pronounced reduction among the trehalase genes examined-whereas knockdown of OaTRE caused a moderate yet substantial decline of about 67% (Fig. 5C).

Effects of dsOaTRE1 and dsOaTRE on the Development of O. asiaticus

A preliminary investigation into the function of the TRE genes in O. asiaticus was conducted using microinjection-mediated RNAi technology. This represents the first further exploration of the functional roles of these genes in O. asiaticus. To investigate the developmental effects of dsOaTRE1 and dsOaTRE, we injected gene-specific dsRNA into 4th-instar nymphs of O. asiaticus, followed by phenotypic monitoring.

Knockdown of OaTRE1 and OaTRE through RNAi resulted in significant lethal effects and developmental abnormalities, with notable differences in phenotypic severity between the two gene knockdown groups. Treated insects exhibited pronounced morphological defects (Fig. 5B). Insects injected with dsGFP (control) developed normally. In contrast, nymphs treated with dsTRE1 showed outcomes and exhibited severe malformations (Fig.5B to b), with 33.3% of locusts displaying these abnormal phenotypes (Fig. 5E). Notably, dsTRE treatment led to higher mortality due to failure to complete molting (Fig. 5B to d), while a subset of surviving individuals appeared morphologically normal (Fig. 5B to e). These results indicate that OaTRE1 and OaTRE both play critical roles in insect development.

Notably, dsOaTRE injection induced substantially more severe lethal effects, with cumulative mortality exceeding 70% at 72 h and approaching 90% at 96 h, significantly higher than that observed in the dsOaTRE1 group (Fig. 5D). Kaplan-Meier survival analysis confirmed that both experimental groups exhibited significantly reduced survival rates compared to the dsGFP control group (Log-rank test, P = 1.1 × 10−24), with the dsOaTRE group (green curve) demonstrating substantially faster mortality progression than the dsOaTRE1 group (red curve). Collectively, these findings demonstrate that TRE genes play crucial roles in the survival and development of O. asiaticus.

Treatment with Exogenous Hormones

Treatment with 20E significantly modulated the expression of OaTRE1 and OaTRE, two key genes involved in chitin biosynthesis, in O. asiaticus. Prior to assessing the effects of 20E, we confirmed that the solvent control (10% ethanol, v/v) did not induce significant changes in the expression levels of OaTRE1 and OaTRE at any time point (12 to 72 h; Fig. 6A to B). As shown in Fig. 6A, OaTRE1 expression was rapidly upregulated within 12 h of 20E exposure, reaching a peak at 24 h with a 5.6% increase under 2.5 µg/µL treatment (P < 0.05), followed by a decline at 48 h. This induction was dose-dependent: at 24 h, 5 µg/µL and 10 µg/µL treatments resulted in 3.8% and 2.5% increases, respectively, while 20 µg/µL showed minimal induction.

Fig. 6.

Bar graphs showing effects of 20E on OaTRE1 (panel A) and OaTRE (panel B) expression in O. asiaticus at different time points, with significance markers.

Open in a new tab

Effects of 20E on the expression of OaTRE1 A) and OaTRE B) genes in O. asiaticus, with 10% ethanol as the control. Data from three biological replicates are presented as mean ± standard deviation (SD). Within each time point, groups labeled with different lowercase letters above the error bars are significantly different (Tukey’s HSD test, P < 0.05). The abscissa represents time points, and the ordinate shows expression relative to the control.

Similarly, OaTRE expression exhibited a time- and dose-dependent response (Fig. 6B). After 24 h of 2.5 µg/µL 20E treatment, OaTRE expression increased by 2.9%, further rising to 10.7% at 48 h. At 5 µg/µL, the fold-change reached 12.4% at 48 h.

These results suggest that 20E acts as a potent regulator of chitin-related gene expression in O. asiaticus, with optimal induction occurring at moderate concentrations and intermediate time points.

To investigate the effect of validamycin on chitin biosynthesis-related gene expression in O. asiaticus, 4th-instar nymphs were injected with varying concentrations of validamycin (2.5 to 20 µg/µL), with ddH2O as the control. As shown in Fig. 7A and B, validamycin treatment led to a dose-and time-dependent suppression of both OaTRE1 and OaTRE expression.

Fig. 7.

Bar graphs showing expression levels of OaTRE1 (panel A) and OaTRE (panel B) in O. asiaticus after validamycin treatment across time points, with bars representing control and varying concentrations, and significance markers.

Open in a new tab

Effects of validamycin on the expression of OaTRE1 A) and OaTRE B) genes in O. asiaticus, with ddH2O as the control. Data from three biological replicates are presented as mean ± standard deviation (SD); different lowercase letters on the error bars indicate significant differences (P < 0.05). Within each time point, groups labeled with different lowercase letters above the error bars are significantly different (Tukey’s HSD test, P < 0.05). The abscissa represents time points, and the ordinate shows expression relative to the control.

For OaTRE1 (Fig. 7A), the highest concentration (20 µg/µL) induced a rapid and profound inhibition: at 12 h post-injection, expression was reduced by 90.3% compared to the control (P < 0.05) with 10 µg/µL showing approximately 70% reduction at 12 h.

Similarly, OaTRE expression was strongly suppressed by validamycin (Fig. 7B). At 20 µg/µL, expression dropped by 94.8% at 12 h and remained below control levels throughout the observation period. Even at 5 µg/µL, a 60% to 70% reduction was observed at 12 to 24 h. Notably, the control group showed no significant change in expression over time, confirming that the observed effects are due to validamycin rather than solvent toxicity. These results demonstrate that validamycin exerts a potent inhibitory effect on key chitin synthesis genes in O. asiaticus, with maximal suppression occurring within 12 to 24 h and persisting for up to 48 h.

Discussion

Chitin synthases play crucial roles in insect molting and growth and development. Trehalase, by hydrolyzing trehalose (the primary blood sugar in insects) to glucose, is important in the production of energy. It is also indispensable for chitin remodeling and pheromone biosynthesis (Qin et al. 2015, Zhang et al. 2024). By supplying glucose as a critical carbon source, trehalase indirectly supports chitin biosynthesis and other energy-dependent physiological processes (Zhu et al. 2016, Tellis et al. 2023). In the present study, two trehalase genes involved in the chitin synthesis pathway of Oedaleus asiaticus were cloned and designated OaTRE1 and OaTRE. Sequence analysis revealed that the amino acid sequences of OaTRE1 and OaTRE shared more than 90% similarity with their respective homologs (LmTRE1 and LmTRE) in Locusta migratoria manilensis. Phylogenetic analysis showed that these sequences clustered into a major branch and a distinct sub-branch, consistent with their established taxonomic classification. Notably, Tre1 and Tre within the same insect species exhibited relatively low sequence similarity, whereas each displayed high homology with its corresponding ortholog across different insect species, suggesting functional divergence between Tre1 and Tre but strong evolutionary conservation within each group. In Lepidoptera, although overall sequence similarity between Tre1 and Tre was comparatively low, Tre1 proteins from Spodoptera frugiperda, Spodoptera litura, and S. exigua clustered closely together, indicating a conserved evolutionary relationship within this lineage (Xu et al. 2022). Moreover, with the advancement of genome sequencing, multiple trehalase genes-beyond the canonical two-have been identified in several insect species, highlighting the increasing complexity of trehalose metabolism in insects (Avonce et al. 2006, Pinheiro et al. 2020).

In this study, OaTRE1 and OaTRE exhibited distinct tissue- and stage-specific expression patterns in O. asiaticus, suggesting functional differentiation between the two trehalases. OaTRE1 was predominantly expressed in the epidermis, Malpighian tubules, and trachea, whereas OaTRE showed higher expression in the Malpighian tubules and fat body, tissues closely associated with active metabolism and energy storage. Similar tissue distribution patterns have been reported for trehalase genes in L. migratoria, where Tre-1 is highly expressed in the integument and trachea, and Tre is enriched in metabolically active tissues such as the fat body and intestine (Wang 2009).

In addition, trehalase expression varied markedly across developmental stages, consistent with reports in other insects, including Calliptamus italicus, in which trehalase genes are expressed throughout development but at different levels (Guo et al. 2015, Liu and Luo 2022). Collectively, these results indicate that OaTRE1 and OaTRE not only participate in chitin biosynthesis but also play important and partially specialized roles in energy metabolism during growth and development of O. asiaticus.

Trehalases are closely associated with insect molting by regulating chitin synthesis. Silencing of chitin synthase genes in locusts has been shown to cause molting failure and ultimately death (Zhang et al. 2010). In Acyrthosiphon pisum, RNAi-mediated silencing of ApTRE resulted in distinct phenotypic abnormalities, including molting defects in both color morphs and wing deformities specifically in the red morph (Wang et al. 2021). Similarly, Tang et al. (2016) reported that the knockdown of five trehalase genes in T. castaneum, particularly TRE1 and TRE2, led to molting deformities and high mortality through disruption of the chitin biosynthesis pathway. Collectively, these results indicate that OaTRE1 and OaTRE not only participate in chitin biosynthesis but also play important and partially specialized roles in energy metabolism during growth and development of O. asiaticus.

To further validate the function of trehalase in O. asiaticus, we conducted RNAi experiments by injecting dsRNA targeting OaTRE1 and OaTRE into 4th-instar nymphs. qRT-PCR verification demonstrated that compared with the dsGFP control group, the expression levels of the target genes in the dsRNA-treated group were significantly downregulated. This gene knockdown led to decreased survival rate and phenotypic abnormalities. Meanwhile, the chitin content in nymphs of the dsRNA-treated group was significantly reduced. These results indicate that the trehalase genes OaTRE1 and OaTRE affect the growth and development of O. asiaticus.

Although our data indicate that knockdown of OaTRE1and OaTRE genes is associated with reduced chitin content and developmental defects, we acknowledge the existence of other competing explanations. First, dsRNA injection may induce non-specific immune responses or physical damage. Second, knockdown of trehalase genes may exert pleiotropic metabolic effects beyond chitin synthesis. However, our chitin quantification data, combined with the specificity of the phenotypes, establish the first functional link between trehalase and chitin formation in this species. This foundation justifies future investment in these validation methodologies.

Previous studies have shown that the activity of insect trehalases can be regulated by hormonal signals. 20E, a key ecdysteroid hormone, plays a central role in insect growth and development. Upon entering the intestine, ecdysteroids are converted into 20E, which modulates physiological processes such as molting, pupation, and metamorphosis (Liu et al. 2015). It has been reported that 20E enhances the activity of Tre1 in Omphisa fuscidentalis Hampson. (Tatun et al. 2008). Validamycin is a widely used antifungal antibiotic in Asian agriculture, produced via fermentation by Streptomyces hygroscopicus 5008 (Li et al. 2016). It not only inhibits trehalase activity in Locusta migratoria manilensis (Fan 2009) but also exerts dual insecticidal effects in S. litura by suppressing trehalose hydrolysis-leading to trehalose accumulation and glucose depletion-and concurrently disrupting chitin biosynthesis (Yu et al. 2022).

In this study, the expression levels of OaTRE1 and OaTRE were slightly elevated compared with the control group following injection of different concentrations of 20E at various time points, suggesting that 20E can induce the expression of genes encoding key enzymes in the chitin synthesis pathway. In contrast, injection of validamycin into 4th-instar nymphs resulted in a concentration-dependent reduction in the expression of both genes, indicating that validamycin inhibits the expression of trehalase-related genes involved in chitin biosynthesis. Together, these results further support a critical role of trehalase in coordinating chitin synthesis and energy metabolism in O. asiaticus and suggest that trehalase may represent a promising molecular target for the development of environmentally friendly pest management strategies in grassland ecosystems. However, we acknowledge the absence of direct biochemical validation in this study. Consequently, mechanistic interpretations regarding trehalase-mediated chitin synthesis remain hypothetical. Future research should prioritize performing trehalase enzyme activity assays and identifying downstream biochemical markers to rigorously validate the proposed mechanism linking mRNA knockdown to chitin synthesis impairment. Concurrently, the applicability of these molecular targets for sustainable pest control strategies requires thorough field-based assessment during subsequent collection seasons.

Supplementary Material

ieag017_Supplementary_Data
ieag017_supplementary_data.doc (82KB, doc)

Acknowledgements

We sincerely thank the Inner Mongolia Autonomous Region Natural Science Foundation (grant number: 2025MS03076) and the Ordos Municipal Key Research and Development Program (grant number: YF20240068) for their generous financial support.

Contributor Information

Mei Liu, Key Laboratory of Biohazard Monitoring, Green Prevention and Control for Artificial Grassland, Ministry of Agriculture and Rural Affairs, Institute of Grassland Research of Chinese Academy of Agricultural Sciences, Hohhot, China.

Feng Yan, Ordos Vocational College of Ecological Environment, Department of Ecological Engineering, Ordos, Inner Mongolia, China.

Ruiwen Dong, Grassland Station, Forestry and Grassland Bureau, Siziwang Banner, Ulanqab City, Inner Mongolia Autonomous Region, China.

Haiyan Gao, Key Laboratory of Biohazard Monitoring, Green Prevention and Control for Artificial Grassland, Ministry of Agriculture and Rural Affairs, Institute of Grassland Research of Chinese Academy of Agricultural Sciences, Hohhot, China.

Yidan Wu, Key Laboratory of Biohazard Monitoring, Green Prevention and Control for Artificial Grassland, Ministry of Agriculture and Rural Affairs, Institute of Grassland Research of Chinese Academy of Agricultural Sciences, Hohhot, China.

Kejian Lin, Key Laboratory of Biohazard Monitoring, Green Prevention and Control for Artificial Grassland, Ministry of Agriculture and Rural Affairs, Institute of Grassland Research of Chinese Academy of Agricultural Sciences, Hohhot, China.

Shujing Gao, Key Laboratory of Biohazard Monitoring, Green Prevention and Control for Artificial Grassland, Ministry of Agriculture and Rural Affairs, Institute of Grassland Research of Chinese Academy of Agricultural Sciences, Hohhot, China.

Author Contributions

Mei Liu (Conceptualization [equal], Data curation [lead], Formal analysis [equal], Writing—original draft [lead]), Feng Yan (Data curation [equal], Software [lead]), Ruiwen Dong (Resources [equal], Supervision [equal]), Haiyan Gao (Data curation [equal], Resources [lead]), Yidan Wu (Formal analysis [equal], Methodology [lead]), Kejian Lin (Funding acquisition [equal], Writing—review & editing [equal]), and Shu-jing Gao (Funding acquisition [lead], Visualization [equal], Writing—review & editing [lead])

Supplementary Material

Supplementary material is available at Journal of Insect Science online.

Funding

This research was funded by the Inner Mongolia Autonomous Region Natural Science Foundation (grant number: 2025MS03076) and the OrdosMunicipal Key Research and Development Program (grant number: YF20240068).

Conflicts of Interest

The authors declare no competing interests.

References

  1. Argüelles JC.  2000. Physiological roles of trehalose in bacteria and yeasts: a comparative analysis. Arch. Microbiol. 174:217–224. [DOI] [PubMed] [Google Scholar]
  2. Avonce N, Mendoza-Vargas A, Morett E, et al.  2006. Insights on the evolution of trehalose biosynthesis. BMC Evol. Biol. 6:1–15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Becker A, Schlöder P, Steele JE, et al.  1996. The regulation of trehalose metabolism in insects. Experientia  52:433–439. [DOI] [PubMed] [Google Scholar]
  4. Candy DJ, Kilby BA.  1962. Studies on chitin synthesis in the desert locust. J. Exp. Biol. 39:129–140. [Google Scholar]
  5. Chen J, Tang B, Chen H, et al.  2010. Different functions of the insect soluble and membrane-bound trehalase genes in chitin biosynthesis revealed by RNA interference. PLoS One. 5:e10133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chen J, Zhang DW.  2015. Molecular cloning, tissue distribution and temperature-induced expression of two trehalose-6-phosphate synthase genes in Blattella germanica (Blattodea: Blattellidae). Acta Entomol. Sinica  58:1046–1053. [Google Scholar]
  7. Chen QW, Jin S, Zhang L, et al.  2018. Regulatory functions of trehalose-6-phosphate synthase in the chitin biosynthesis pathway in Tribolium castaneum (coleoptera: Tenebrionidae) revealed by RNA interference. Bull. Entomol. Res. 108:388–399. [DOI] [PubMed] [Google Scholar]
  8. Evans DR, Dethier VG.  1957. The regulation of taste thresholds for sugars in the blowfly. J. Insect Physiol. 1:3–17. [Google Scholar]
  9. Feng GH, Fan SX, Liu QF, et al.  1995. The determination capacity for eaten of several species grasshoppers in grassland in outside cage condition. Acta Agrestia Sinica  3:230. [Google Scholar]
  10. Fan KQ.  Inhibitory Effects of Validamycin and Its Hydrolytic Products on Trehalase [D].  Zhejiang University of Technology, 2009. [Google Scholar]
  11. JUDITH K.  2015. Juvenile hormone: a Central regulator of termite caste polyphenism. Academic Press (48:): 131–161. [Google Scholar]
  12. Guan JQ, Wei ZZ.  1989. Measurement of food consumption in oedaleus asiaticus. Chin. J. Appl. Entomol. 8–10. [Google Scholar]
  13. Guo Q, Hao YJ, Li Y, et al.  2015. Gene cloning, characterization and expression and enzymatic activities related to trehalose metabolism during diapause of the onion maggot delia antiqua (diptera: Anthomyiidae). Gene  565:106–115. [DOI] [PubMed] [Google Scholar]
  14. Hegedus D, Erlandson M, Gillott C, et al.  2009. New insights into peritrophic matrix synthesis, architecture, and function. Annu. Rev. Entomol. 54:285–302. [DOI] [PubMed] [Google Scholar]
  15. Hegedus D, Erlandson M, Gillott C, et al.  2009. “New insights into peritrophic matrix synthesis, architecture, and function.” Annu. Rev. Entomol.  54:285–302. [DOI] [PubMed] [Google Scholar]
  16. Huang X, Whitman DW, Ma J, et al.  2017. Diet alters performance and transcription patterns in oedaleus asiaticus (orthoptera: Acrididae) grasshoppers. PLoS One. 12:e0186397. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Kang L, Chen Y.  1994. Trophic niche of steppe grasshoppers. Acta Ent. Sin.  37:178–189. [Google Scholar]
  18. Kelkenberg M, Odman-Naresh J, Muthukrishnan S, et al.  2015. Chitin is a necessary component to maintain the barrier function of the peritrophic matrix in the insect midgut. Insect Biochem. Mol. Biol. 56:21–28. [DOI] [PubMed] [Google Scholar]
  19. Kramer KJ, Koga D.  1986. Insect chitin: physical state, synthesis, degradation and metabolic regulation. Insect Biochem. 16:851–877. [Google Scholar]
  20. Liu X, Dai F, Guo E, et al.  2015. 20-Hydroxyecdysone (20E) primary response gene E93 modulates 20E signaling to promote bombyx larval-pupal metamorphosis. J. Biol. Chem. 290:27370–27383. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Liu Q, Luo D.  2022. Cloning of trehalase gene from calliptamus italicus and expression pattern under low temperature acclimation in eggs. Acta Entomol. Sin. 65:157–166. [Google Scholar]
  22. Liu XJ, Zhang HH, Li DQ, et al.  2012. Sequence analysis and mRNA expression characteristics of the soluble trehalase gene in the locusta migratoria manilensis. Acta Entomol. Sin. 55:1264–1271. [Google Scholar]
  23. Livak KJ, Schmittgen TD.  2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods  25:402–408. [DOI] [PubMed] [Google Scholar]
  24. Li L, Jiang Y, Liu Z, et al.  2016. Jinggangmycin increases fecundity of the brown planthopper, nilaparvata lugens (stål) via fatty acid synthase gene expression. J. Proteomics. 130:140–149. [DOI] [PubMed] [Google Scholar]
  25. MIURA T.  2019. Juvenile hormone as a physiological regulator mediating phenotypic plasticity in pancrustaceans. Dev. Growth Differ. 61:85–96. [DOI] [PubMed] [Google Scholar]
  26. Merzendorfer H.  2006. Insect chitin synthases: a review. J. Comp. Physiol. B  176:1–15. [DOI] [PubMed] [Google Scholar]
  27. Merzendorfer H, Zimoch L.  2003. Chitin metabolism in insects: Structure, function and regulation of chitin synthases and chitinases. J. Exp. Biol. 206:4393–4412. [DOI] [PubMed] [Google Scholar]
  28. Moussian B.  2010. Recent advances in understanding mechanisms of insect cuticle differentiation. Insect Biochem. Mol. Biol. 40:363–375. [DOI] [PubMed] [Google Scholar]
  29. Muller J, Boller T, Wiemken A.  1995. Effects of validamycin A, a potent trehalase inhibitor and phytohormones on trehalose metabolism in roots and root nodules of soybean and cowpe. Planta  97:362–368. [Google Scholar]
  30. Pinheiro DH, Taylor CE, Wu K, et al.  2020. Delivery of gene‐specific dsRNA by microinjection and feeding induces RNAi response in Sri Lanka weevil, myllocerus undecimpustulatus undatus marshall. Pest Manag. Sci. 76:936–943. [DOI] [PubMed] [Google Scholar]
  31. Qin JM, Luo SD, He SY, et al.  2015. Research on the characteristics and functions of trehalose and trehalase in insects. J. Environ. Entomol. 37:163–169. [Google Scholar]
  32. Rabadiya D, Behr M.  2024. The biology of insect chitinases and their roles at chitinous cuticles. Insect Biochem. Mol. Biol. 165:104071. [DOI] [PubMed] [Google Scholar]
  33. Ritchie JM.  A taxonomic revision of the genus Oedaleus Fieber (Orthoptera: Acrididae; ). 1981. [Google Scholar]
  34. Shukla E, Thorat LJ, Nath BB, et al.  2015. Insect trehalase: physiological significance and potential applications. Glycobiology  25:357–367. [DOI] [PubMed] [Google Scholar]
  35. Su CC.  Genome-Wide Identification of Chitinase Gene Family and Analysis of Chitin Content in Chilo Suppressalis (Walker) [D]. Nanjing Agricultural University, 2017. [Google Scholar]
  36. Takiguchi M, Niimi T, Su ZH, et al.  1992. Trehalase from male accessory gland of an insect, tenebrio molitor. cDNA sequencing and developmental profile of the gene expression. Biochem. J. 288 ( Pt 1):19–22. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Tang B, Chen J, Yao Q, et al.  2010. Characterization of a trehalose-6-phosphate synthase gene from Spodoptera exigua and its function identification through RNA interference. J. Insect Physiol. 56:813–821. [DOI] [PubMed] [Google Scholar]
  38. Tang B, Chen X, Liu Y, et al.  2008. Characterization and expression patterns of a membrane-bound trehalase from Spodoptera exigua. BMC Mol. Biol. 9:51. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Tang B, Wei P, Zhao L, et al.  2016. Knockdown of five trehalase genes using RNA interference regulates the gene expression of the chitin biosynthesis pathway in Tribolium castaneum. BMC Biotechnol. 16:67–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Tang B, Wei P, Zhao L, et al.  2016. Knockdown of five trehalase genes using RNA interference regulates the gene expression of the chitin biosynthesis pathway in Tribolium castaneum. BMC Biotechnol. 16:67–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Tang B, Yang M, Shen Q, et al.  2017. Suppressing the activity of trehalase with validamycin disrupts the trehalose and chitin biosynthesis pathways in the rice brown planthopper, nilaparvata lugens. Pestic. Biochem. Physiol. 137:81–90. [DOI] [PubMed] [Google Scholar]
  42. Tang B, Yang MM, Shen QD, et al.  2017. Suppressing the activity of trehalase with validamycin disrupts the trehalose and chitin biosynthesis pathways in the rice brown planthopper, nilaparvata lugens. Pestic. Biochem. Physiol.  137:81–90. [DOI] [PubMed] [Google Scholar]
  43. Tatun N, Singtripop T, Sakurai S.  2008. Dual control of midgut trehalase activity by 20-hydroxyecdysone and an inhibitory factor in the bamboo borer omphisa fuscidentalis hampson. J. Insect Physiol. 54:351–357. [DOI] [PubMed] [Google Scholar]
  44. Tatun N, Wangsantitham O, Tungjitwitayakul J, et al.  2014. Trehalase activity in fungus-growing termite, odontotermes feae (isoptera: Termitideae) and inhibitory effect of validamycin. J. Econ. Entomol. 107:1224–1232. [DOI] [PubMed] [Google Scholar]
  45. Takahashi M, Kono Y, Kurahashi H, et al.  1995. Effect of a trehalase inhibitor, validoxylamine A, on three species of flies. Appl. Entomol. Zool. 30:231–239. [Google Scholar]
  46. Tellis MB, Kotkar HM, Joshi RS.  2023. Regulation of trehalose metabolism in insects: from genes to the metabolite window. Glycobiology  33:262–273. [DOI] [PubMed] [Google Scholar]
  47. Tellis MB, Kotkar HM, Joshi RS.  2023. Regulation of trehalose metabolism in insects: from genes to the metabolite window. Glycobiology  33:262–273. [DOI] [PubMed] [Google Scholar]
  48. Terra WR, Ferreira C.  1994. Insect digestive enzymes: properties, compartmentalization and function. Comp. Biochem. Physiol. B: Comp. Biochem. 109:1–62. [Google Scholar]
  49. Thompson SN.  2003. Trehalose—the insect ‘blood’sugar. Adv. In Insect Phys. 31:205–285. [Google Scholar]
  50. Wang G, Gou Y, Guo S, et al.  2021. RNA interference of trehalose-6-phosphate synthase and trehalase genes regulates chitin metabolism in two color morphs of acyrthosiphon pisum harris. Sci. Rep. 11:948. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Wang JE.  Cloning and Expression Analysis of Trehalase Genes in Locusta Migratoria Manilensis [D].  Chongqing University, 2009. [Google Scholar]
  52. Wang XX.  Cloning, Characterization and Functional Analysis of the Chitin Synthase B cDNA Sequence from Mythimna Separata [D].  Northeast Agricultural University, 2019. [Google Scholar]
  53. Xiong KC, Wang J, Li JH, et al.  2016. RNA interference of a trehalose-6-phosphate synthase gene reveals its roles during larval-pupal metamorphosis in bactrocera minax (diptera: Tephritidae). J. Insect Physiol. 91-92:84–92. [DOI] [PubMed] [Google Scholar]
  54. Xu JJ, Li SQ, Ren MY, et al.  2022. Research progress on the functions of key enzymes in insect chitin synthesis and the application of RNAi technology in pest control. Shaanxi Agric. Sci. 68:1–10. [Google Scholar]
  55. Yasugi T, Yamada T, Nishimura T.  2017. Adaptation to dietary conditions by trehalose metabolism in Drosophila. Sci. Rep. 7:1619. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Yu HZ, Zhang Q, Lu ZJ, et al.  2022. Validamycin treatment significantly inhibits the glycometabolism and chitin synthesis in the common cutworm, Spodoptera litura. Insect Sci. 29:840–854. [DOI] [PubMed] [Google Scholar]
  57. Zhang H, Li H, Fang S, et al.  2024. Co-application of validamycin a and dsRNAs targeting trehalase genes conferred enhanced insecticidal activity against laodelphax striatellus. Pestic. Biochem. Physiol. 205:106160. [DOI] [PubMed] [Google Scholar]
  58. Zhang J, Liu X, Zhang J, et al.  2010. Silencing of two alternative splicing-derived mRNA variants of chitin synthase 1 gene by RNAi is lethal to the Oriental migratory locust, locusta migratoria manilensis (meyen). Insect Biochem. Mol. Biol. 40:824–833. [DOI] [PubMed] [Google Scholar]
  59. Zhang L, Yan YH, Wang GQ, et al.  1995. Preliminary field investigation on the epidemic of microsporidiosis in grasshoppers. Acta Agrestia Sin. 3:223. [Google Scholar]
  60. Zhang WQ, Chen XF, Tang B, et al.  2011. Insect chitin biosynthesis and its regulation. Chin. J. Appl. Entomol  48:475–479. [Google Scholar]
  61. Zhu KY, Merzendorfer H, Zhang W, et al.  2016. Biosynthesis, turnover, and functions of chitin in insects. Annu. Rev. Entomol. 61:177–196. [DOI] [PubMed] [Google Scholar]
  62. Zhu KY, Merzendorfer H, Zhang W, et al.  2016. Biosynthesis, turnover, and functions of chitin in insects. Annu. Rev. Entomol. 61:177–196. [DOI] [PubMed] [Google Scholar]
  63. Zhou YT.  2019. Identification and functional characterization of olfaction-related proteins in Oedaleus asiaticus [D]. Master’s Thesis. Inner Mongolia Agricultural University.
  64. Zheng H, Hua M, Jiang M, et al.  2025. Transgenic expression of mAChR-C dsRNA in maize confers efficient locust control. Plant Commun. 6:101316. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

ieag017_Supplementary_Data
ieag017_supplementary_data.doc (82KB, doc)

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

RESOURCES