1. Introduction
Sogatella furcifera (Horváth) (Hemiptera: Delphacidae) is a typical hemimetabolous rice pest found in many East Asian countries [1,2,3]. It is known to damage rice through sucking sap directly from the phloem of rice plants and through ovipositing on rice [4,5,6,7]. Moreover, it is known to transmit rice viruses, such as southern rice black-streaked dwarf virus. Once rice is infected with the virus, it can cause severe stunting and reduce the setting rate, thus leading to rice yield losses [8,9,10]. The growth and development of S. furcifera progresses through three sequential stages: egg, nymph, and adult stages. Moreover, the nymph stage is divided into five instars. Thus, S. furcifera must undergo five molting processes to develop from a nymph to an adult [11,12,13,14]. Molting and wing expansion are therefore the key processes for the growth and migration of S. furcifera. In insects, both molting and wing development require the production of chitin [15,16,17].
Chitin, a linear polysaccharide homopolymer of N-acetylglucosamines, is an essential structural composition of the insect cuticle and therefore a pivotal target for controlling pest insects [18,19]. Chitin biosynthesis in insects is a complex and dynamic process involving at least eight enzymes [20,21]. Chitin synthesis begins with the hydrolysis of trehalose to glucose by trehalase (Tre). Next, glucose enters a sequential catalysis reaction involving hexokinase (HK), glucose-6-phosphate isomerase (G6PI), glutamine: fructose-6-phosphate aminotransferase (GFAT), glucosamine-6-phosphate N-acetyltransferase (GNA), phosphoacetylglucosamine mutase (PAGM), and UDP-N-acetylglucosamine pyrophosphorylase (UAP). Finally, a chitin polymer is synthesized by chitin synthase (CHS) [22]. To date, only three of the enzymes involved in the chitin synthesis pathway (i.e., Tre, UAP, and CHS) have been widely studied [23,24,25,26,27]. In contrast, data regarding the molecular and functional characterization of other chitin synthetic enzymes in insects are limited.
GFAT (EC 2.6.1.16), a pivotal rate-limiting enzyme in the hexosamine pathway, specifically catalyzes the conversion of fructose-6-phosphate and glutamine into glucosamine-6-phosphate [28]. This reaction product is then further processed by several enzymes to produce uridine diphosphate-N-acetylglucosamine, a vital precursor molecule for chitin synthesis in insects [29]. To date, GFAT has been identified in relatively few insect species, including Drosophila melanogaster [30], Aedes aegypti [31], Nilaparvata lugens [32], and Hyphantria cunea [33]. In D. melanogaster, GFAT activity can be inhibited by UDP-N-acetylglucosamine, which acts to regulate the rate of chitin formation [30]. In A. aegypti, GFAT expression is upregulated in the midgut after blood feeding, and RNAi knockdown of AeGFAT-1 was found to severely impair the formation of a peritrophic matrix [31,34]. Furthermore, the silencing of GFAT in N. lugens led to a decrease in the expression of genes related to chitin metabolism and caused very high levels of malformation and mortality [32]. In addition, knockdown of GFAT caused the downregulation of other genes, including GNA, PAGM, UAP, and CHSA, thereby resulting in decreased chitin content in the epidermis [33]. Overall, data from previous studies suggest that GFAT plays an essential role in the regulation of insect growth and metamorphosis.
In this research, a GFAT gene (SfGFAT) was identified for the first time in S. furcifera. In addition, structural molecular characteristics and a phylogenetic tree including SfGFAT were determined via bioinformatic analyses. Moreover, the biological function of SfGFAT was determined using RNA interference (RNAi) knockdown. These results can help us understand the biological function of SfGFAT in planthopper chitin synthesis and may provide a potential target for the development of new chitin synthesis inhibitors.
2. Materials and Methods
2.1. Insect Rearing and Sample Collection
The S. furcifera were raised in a growth chamber at the institute of entomology, Guizhou University, Guiyang, China. The insects rearing was kept in mesh cages with fresh Taichung Native-1 rice seedlings at 25 °C ± 1 °C, 70% ± 10% relative humidity, and a 16:8 h (L:D) photoperiod [15].
As described previously [35], samples were collected during the feeding process. The sample collection schedule included 18 time points from the egg stage to the adult stage, including 1–2-day-old egg (EG1–EG2), 1–2-day-old 1st instar nymph (1L1–1L2), 1–2-day-old 2nd instar nymph (2L1–2L2), 1–3-day-old 3rd instar nymph (3L1–3L3), 1–3-day-old 4th instar nymph (4L1–4L3), 1–3-day-old 5th instar nymph (5L1–5L3), and 1–3-day-old adult (AD1–AD3). Samples of different tissues were collected from the head, integument, fat body, and gut of 1-day-old 5th instar nymphs and from the ovary of 3-day-old adults. During sampling, three biological replicates of each sample were fleetly frozen in liquid nitrogen and stored at −80 °C prior to use.
2.2. Primer Design
Primers were generated based on transcriptome sequencing data of S. furcifera (SRR116252). Primer design was performed using Primer Premier version 6.0 (Palo Alto, CA, USA). The sequences of all primers used in this study are shown in Table 1. All primers were synthesized by Sangon Biotech Co., Ltd. (Shanghai, China).
2.3. RNA Isolation and cDNA Synthesis
Whole bodies of S. furcifera nymphs or adults were used to isolate total RNA for the cloning of SfGFAT. First, total RNA was extracted using an HP Total RNA Kit (Omega Bio-Tek, Norcross, GA, USA) with genomic DNA removal columns following the manufacturer’s instructions. The integrity of the extracted RNA was verified via 1% agarose gel electrophoresis. Further analysis using a Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA) was performed to estimate the concentration and purity of RNA. The purified RNA was then stored at −80 °C for future use. An AMV First Strand cDNA Synthesis Kit with an oligo(dT) primer (Sangon Biotech, Shanghai, China) was used to synthesize first-strand complementary DNA (cDNA) following the manufacturer’s instructions. All cDNA samples were then stored at −20 °C for future experiments.
2.4. Cloning of SfGFAT
Based on the results of transcriptome sequencing of S. furcifera (SRR116252), two cDNA fragments encoding GFAT were obtained using Geneious 2020.0.5 (Biomatters, Inc., Auckland, New Zealand). We then amplified these sequences by polymerase chain reaction (PCR) using prosynthetic cDNA and gene-specific primers (GSPs, Table 1). PCR was performed using a Bio-Rad T100 Thermal Cycler PCR System (Bio-Rad, Hercules, CA, USA). In brief, 25 μL reaction mixtures contained 2 μL dNTP (2.5 mM), 2.5 μL 10× LA PCR Buffer (Mg2+ plus), 1 μL of each primer (10 mM), l μL cDNA template, 0.25 μL LA Taq polymerase (TaKaRa, Dalian, China), and double-distilled water up to 25 μL. PCR reaction conditions were as follows: one cycle of pre-denaturation at 94 °C for 3 min; followed by 30 cycles of denaturation at 94 °C for 30 s, annealing at 53 °C for 30 s, and extension at 72 °C for 2 min; and a final extension at 72 °C for 10 min. Finally, the amplified products were examined by 1% agarose gel electrophoresis. The target bands for the desired products were then purified using an EasyPure® Quick Gel Extraction Kit (TransGen Biotech, Beijing, China). The purified DNA was then ligated to a pMD18-T vector (TaKaRa, Dalian, China) and sequenced by Sangon Biotech (Shanghai, China).
Next, we amplified the ends of SfGFAT via rapid amplification of cDNA end PCR (RACE-PCR) using a SMARTer RACE 5′/3′ Kit (Clontech, Mountain View, CA, USA). In particular, we used long universal primers and GSPs to perform the primary RACE-PCR. Here, the reaction conditions were as follows: 30 cycles of denaturation at 94 °C for 30 s, annealing at 55 °C–57 °C (according to the primer annealing temperature) for 30 s, and final extension at 72 °C for 60 s. For the nested RACE-PCR reaction, the primary PCR product was initially diluted 100 times before being used as a template with a short universal primer and GSPs. The reaction conditions were the same as those used for the primary PCR reaction. All RACE-PCR products were then purified and sequenced as previously described.
2.5. Bioinformatics Analyses
All obtained sequencing fragments were assembled using SeqMan version 5.0 (DNASTAR, Inc., Madison, WI, USA). The nucleotide sequence was first edited using DNAMAN version 7.0 (Lynnon Biosoft, California, CA, USA). NCBI BLAST (
2.6. Real-Time Quantitative PCR (RT-qPCR) Analysis of SfGFAT Expression Levels
All primers used for RT-qPCR analyses were designed using Primer Premier version 6.0 and are listed in Table 1. S. furcifera 18S rRNA was used as an internal reference gene. RT-qPCR was performed in a CFX-96 real-time quantitative PCR system (Bio-Rad, Hercules, CA, USA) with 20 μL mixtures containing 10 μL FastStart Essential DNA Green Master (Roche, Diagnostics, Shanghai, China), 1 μL (10 μM) of each primer, 1 μL cDNA, and 7 μL hyper-pure water. The PCR amplification conditions were as follows: pre-denaturation at 95 °C for 10 min, 40 cycles of 95 °C for 30 s, and annealing at 55 °C for 30 s. After the reaction, a melting curve analysis was performed from 60 °C to 95 °C to verify the specificity of RT-qPCR products. The relative expression levels of SfGFAT were then calculated using the 2−ΔΔCt method [36].
2.7. Functional Analysis of SfGFAT Using RNAi
To investigate the biological functions of SfGFAT, we used unique primers for SfGFAT and added a T7 RNA polymerase promoter (Table 1) for dsRNA synthesis. Templates for in vitro transcription reactions were synthesized by PCR from a plasmid containing SfGFAT DNA using these primers. The PCR products were then subcloned and sequenced to ascertain its specificity. Next, the expected fragments were purified using an EasyPure® Quick Gel Extraction Kit (TransGen Biotech, Beijing, China). The concentrations of the purified products were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA), and the products were then used for in vitro transcription reactions.
Double-stranded RNA (dsRNA) was synthesized using a MEGAscript® RNAi Kit (Ambion, Carlsbad, CA, USA) with all procedures performed according to the user manual. In vivo SfGFAT gene silencing in S. furcifera nymphs was conducted as previously described [15,35,37]. Briefly, first day 5th instar nymphs were anesthetized with CO2 for approximately 90 s, then placed on a 1% agarose gel plate with grooves. A Nanoliter 2010 Injector (World Precision Instruments, Sarasota, FL, USA) was then used to inject 100 ng of dsGFAT into the junction of the prothorax and mesothorax of S. furcifera subjects; in some instars, dsGFP was injected as a negative control. Each experimental treatment (n = 50) contained three biological replicates. Subsequently, injected nymphs were maintained on fresh rice seedlings and abnormality and mortality rates were assessed daily. Photographs of abnormal insects were captured using a Keyence VH-Z20R stereoscopic microscope (Keyence, Osaka, Japan). In addition, 10 injected insects were selected randomly at 72 h after injection for evaluation of their mRNA levels. The RNAi efficiency was then evaluated via RT-qPCR performed using primers listed in Table 1.
2.8. Statistical Analysis
All data were statistically analyzed using Microsoft Excel 2003 and SPSS version 13 (IBM SPSS Inc., Chicago, IL, USA). The relative expression of SfGFAT at different stages and in different tissues of S. furcifera was determined using the 2−ΔΔCt method. All data were expressed as the mean ± standard error (SE) of three replicates. Differences in gene expression at different stages and in different tissues were calculated using one-way analysis of variance (ANOVA) followed by Duncan’s multiple range test (p < 0.05). Finally, an independent sample t-test was used to evaluate the statistical significance of gene silencing.
3. Results
3.1. Identification and Sequence Analysis of SfGFAT
The entire cDNA sequence of SfGFAT was identified from the DNA fragments amplified using PCR and 5′/3′ RACE (GenBank registration number: MF964939). The SfGFAT sequence was 3162 bp long and included a 5′ noncoding region of 83 bp and a 3′ noncoding region of 1012 bp. The ORF of SfGFAT was 2067 bp long and encoded 688 amino acid residues. The 3′ end of the cDNA sequence of SfGFAT contained a typical AATAAA tail and a poly-A structure (Figure 1). The theoretical molecular weight of this protein was 76.89 kDa, and its theoretical isoelectric point (pI) value was 6.33. Next, we used the NetNGlyc version 1.0 Server to predict potential N-glycosylation sites and found two sites at residues 159 and 327 (Figure 1). Further analysis revealed that the SfGFAT protein did not contain signal peptide and transmembrane helices.
We then used the SWISS-MODEL online tool to model the homology of SfGFAT. We found that the SfGFAT protein consisted of three domains (Figure 2). These included a glutamine aminotransferase class 2 domain (GAT2) in the N-terminus of the protein, which may be responsible for catalyzing the transfer of an amino group from glutamine, as well as two sugar isomerase domains (SIS) in the C-terminus, which function as phosphosugar isomerases or phosphosugar binding proteins.
We then used BLAST to query for homologous sequences of the amino acid sequence encoded by SfGFAT. These results revealed that the amino acid sequence of SfGFAT shared the highest identity with the Hemipteran N. lugens (KU556833.1, 89.63% identity) followed by the Lepidopteran Plutella xylostella (XM_011570168.3, 74% identity). To explore the evolutionary relationships of GFAT and homologous proteins, a phylogenetic tree was constructed using the neighbor-joining method as implemented in MEGA version 6.06 (Figure 3). The tree indicated that SfGFAT has a close evolutionary relationship with other Hemipteran insects, especially N. lugens.
3.2. Spatiotemporal Expression Profile of SfGFAT in S. furcifera
Next, we examined the spatiotemporal expression profiles of SfGFAT at various developmental stages ranging from the egg to adult stages using RT-qPCR. We observed that SfGFAT was continuously expressed across the 18 examined developmental points. Moreover, the relative SfGFAT mRNA expression levels were found to increase just before molting days, reach their highest levels immediately after molting, and then decrease afterward. Finally, peak SfGFAT mRNA expression was found to occur on the first day of adulthood (Figure 4A).
Next, we determined the expression levels of SfGFAT for the integument, fat body, gut, head, and ovary (Figure 4B). SfGFAT expression was significantly higher in the integument than in the other tissues followed by the fat body and ovary. The lowest level of SfGFAT expression was detected in the gut. The relative expression level of SfGFAT in the integument was 72.38, which was 2.51, 2.63, and 14.77 times higher than that in the fat body, ovary, and head, respectively.
3.3. Functional Analysis of SfGFAT
3.3.1. Analysis of SfGFAT mRNA Levels and Survival after RNAi Exposure
To explore the functional significance of SfGFAT, dsRNAs prepared in vitro were injected into newly molted fifth instar nymphs. After 72 h, we collected surviving insects and determined the mRNA levels of SfGFAT present (Figure 5). RT-qPCR results indicated that the levels of SfGFAT mRNA were significantly inhibited following dsGFAT injection (p < 0.01).
The survival rates of the tested S. furcifera individuals were continuously monitored following injection to ascertain whether their development was altered in response to SfGFAT gene silencing (Figure 6). These results showed that the cumulative survival rate declined gradually over time. In particular, no change in survival between the dsGFAT and dsGFP groups 12 h after injection was detected. However, from 24 h onward, the survival of individuals injected with dsGFAT decreased sharply, with only 49.3% of the individuals surviving to eclosion. After eclosion, the survival rate was only 27.3%.
3.3.2. Phenotype Analysis after RNAi
After the successful injection of dsGFAT, the tested insects exhibited four different lethal phenotypes (Figure 7). Approximately 43% of the malformed individuals exhibited a “double-skin” phenotype (I); 8% displayed a “wasp-waisted” phenotype in which the body of the injected nymphs was significantly longer, and the junction between the chest and abdomen was narrower (II); 16% did not shed their old cuticle normally and showed shrunken wings (III); and 9% with a smaller body and misshaped wings died (IV).
4. Discussion
Chitin is the second most abundant biological polysaccharide matrix and provides structural support to the insect exoskeleton, tracheal system, and alimentary canal [38]. Insect chitin remodeling is a highly complex process that is regulated by several enzymes. Previous studies have suggested that the suppression of specific insect chitin remodeling enzymes may be a useful strategy for developing pest control treatments [23,35,37,39]. GFAT is a crucial enzyme that catalyzes the rate-limiting step of the chitin biosynthesis pathway. However, the molecular mechanisms and functions of GFAT underlying the regulation of chitin biosynthesis in S. furcifera remain unknown.
In the present study, we characterized SfGFAT, the GFAT gene from S. furcifera. Our results revealed that the SfGFAT cDNA sequence was 3162 bp in length and encoded a protein containing 688 amino acids. A previous study on A. aegypti showed that the sequence of its GFAT gene contained a 3′ noncoding region that was 770 bp in length [31]. This finding is consistent with the findings of our study since we found that SfGFAT also contains a long 3′ noncoding region (i.e., even longer than AeGfat-1), suggesting the possibility of complex regulation on the translation level. Next, BLAST analysis revealed that the N. lugens GFAT shared 89.63% identity with the S. furcifera GFAT. Phylogenetic analysis indicated that the GFATs of S. furcifera and the hemipteran N. lugens were more closely related than SfGFAT and the GFATs of other eukaryotes and bacteria. Furthermore, structural domain analysis showed that SfGFAT contained a GAT2 domain and two SIS domains, which is similar to the structures of other previously described GFATs. The N-terminal glutamine aminotransferase class 2 domain hydrolyzes glutamine to release an amino group, which then transfers to a new substrate. In addition, the two C-terminal sugar isomerase domains are involved in phosphosugar isomerization and are able to bind phosphosugars [30,31,40,41,42].
Based on the observed mRNA levels of SfGFAT at different developmental stages, it is noteworthy that SfGFAT expression is periodically upregulated just before each molting cycle. More specifically, the relative mRNA level of SfGFAT increased significantly just before the molting days, reached its highest expression level immediately after each molting, and decreased thereafter. One reasonable explanation for this phenomenon is that the formation and hardening of the new cuticle of S. furcifera requires a large amount of chitin and therefore an active chitin biosynthesis pathway. Moreover, the expression trends observed for SfGFAT are highly analogous to trends found in our previous studies of S. furcifera [15,35,37]. In addition, a recent study of N. lugens revealed that NlGFAT was continuously expressed in all developmental stages after the fourth instar and also showed relatively higher expression levels during molting [32]. Previous studies have also verified that the expression of GFAT is tissue-specific. In N. lugens, GFAT was widely detected in a variety of tissues, but NlGFAT was most highly expressed in the wing bud and cuticle [32]. Our tissue-specific expression experiment indicates that SfGFAT was also ubiquitously expressed in the tissues of S. furcifera but showed the highest mRNA levels in the integument, which contain a great deal of chitin. This finding is therefore consistent with the hypothesis that GFAT expression is closely linked to chitin biosynthesis. Similarly, a GFAT in Haemaphysalis longicornis was found to be present in various tissues, including the cuticle, midgut, salivary gland, and ovary [40]. This was consistent with our finding that SfGFAT showed relatively high expression in the ovary. Another previous study of A. aegypti also demonstrated that chitin material is present in the ovaries [43]. We therefore speculate that SfGFAT plays an essential role in chitin biosynthesis and insect reproduction.
Since chitin biosynthesis and degradation pathways are unique to insects, they have been identified as potential targets for controlling pest populations using RNAi methods [44]. Knockdown of chitin metabolic pathway genes using RNAi has been reported to be a practical method for controlling planthoppers. For example, dsCHS1 injection causes a significant decrease in the transcript levels of CHS1 and a remarkable increase in the malformation rates and mortality of both S. furcifera and N. lugens [15,45]. Moreover, the knockdown of N. lugens Tre was found to inhibit the relative expression of other genes involved in the chitin metabolic pathway (e.g., HK, G6PI, chitinase, and CHS) and cause severe molting deformities and mortality [46]. In addition, dsNlTPS injection can reduce the mRNA levels of TPS and thereby induce a lethal response in N. lugens nymphs [47]. Similarly, RNAi-mediated downregulation of SfUAP has been found to seriously affect the growth and metamorphosis of S. furcifera [37]. In our experiment, the transcript levels of SfGFAT significantly decreased following RNAi injection. After eclosion, the survival rate decreased to 27.3%, which further indicated that dsRNA successfully suppressed the expression of GFAT. Finally, dsGFAT injection led to a significant increase in abnormality rates and lethality rates. These findings were consistent with those obtained in N. lugens [32].
5. Conclusions
In summary, we successfully identified the GFAT cDNA of S. furcifera and assessed the normal dynamic changes induced by SfGFAT at several different developmental stages and in several different tissues. Moreover, using RNAi, we found that the knockdown of SfGFAT severely inhibited the expression of the target gene and caused severe molting difficulty and wing malformation in S. furcifera. Overall, our findings demonstrate that SfGFAT plays a vital role in chitin synthesis as well as indicate that SfGFAT may serve as a promising candidate gene for future planthopper control treatments.
Methodology, D.J., Z.W. and G.L.; resources, D.J. and H.Y.; software, C.Z.; funding acquisition, D.J. and Z.W.; investigation, Z.W., G.L. and H.Z.; data curation, D.J. and Z.W.; writing—original draft, Z.W. and G.L. All authors have read and agreed to the published version of the manuscript.
Not applicable.
Not applicable.
The data were deposited in GenBank under accession number MF964939.
The authors declare no conflict of interest.
Footnotes
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Enlarge this image.Figure 1. Nucleotide sequence of SfGFAT cDNA and deduced amino acid sequence. The start codon is highlighted in bold, and the stop codon is indicated in bold with an asterisk. The two putative N-glycosylation sites are shown in red. The polyadenylation signal is shadowed in green. Moreover, the glutamine aminotransferase class 2 domain is underlined, and the sugar isomerase domains are indicated with dotted lines.
Enlarge this image.Figure 3. Phylogenetic analysis of GFAT proteins from S. furcifera and other organisms.
Enlarge this image.Figure 4. Developmental stages (A) and tissues (B) mRNA levels of SfGFAT in S. furcifera. Relative mRNA levels of SfGFAT were measured using qRT-PCR. Data were normalized using S. furcifera 18S rRNA and are shown as the mean ± SE of three independent tests. Different letters imply statistically significant differences in mean expression (p < 0.05, Duncan’s multiple range test in one-way ANOVA).
Enlarge this image.Figure 5. Relative mRNA levels of SfGFAT after injection with an interfering dsRNA (dsGFAT) and a negative control (dsGFP). Values are shown as the mean ± SE of three independent tests. ** indicates extremely significant differences (p < 0.01).
Enlarge this image.Figure 6. Survival rates of S. furcifera after dsRNA injection. Shown are the survival rates of insects injected with 100 ng dsGFAT and dsGFP dsRNAs on the first day of fifth instar nymphs. Insect age in days is displayed on the X-axis; e.g., 5L1, first day of fifth instar nymphs; 5L2 and 5L2′ represent the two 12 h halves of a single day; AD, adults. Values show the mean ± SE of three independent tests.
Primer sequence information.
| Gene | Notes | Primer Name | Primer Sequence (5′–3′) |
|---|---|---|---|
| SfGFAT | SfGFAT | SfGFAT-F | CGAGCAAGTCATCCAACA |
| cloning | SfGFAT-R | GGTCAACAAGAGCCAGAG | |
| 5′GFAT-R1 | TTTGGTGGGTTCCTCTTTAC | ||
| 5′GFAT-R2 | ACTTCCTCTCCTTGTTGCT | ||
| 3′GFAT-F1 | TGCCAGTGATAATGATTGTC | ||
| 3′GFAT-F2 | GAAGATGGAGACACTGAGAC | ||
| RT-qPCR | qGFAT-F | CGAAGATGGAGACACTGAG | |
| for SfGFAT | qGFAT-R | CGGCAATGTGATAGGAGAG | |
| dsGFAT | dsGFAT-F | TAATACGACTCACTATAGGGGTAGCAACAAGGAGAGGAAG | |
| synthesis | dsGFAT-R | TAATACGACTCACTATAGGGACAGCCAATCAGCATCAAG | |
| Sf18S | RT-qPCR for | q18S-F | CGGAAGGATTGACAGATTGAT |
| rRNA | reference gene | q18S-R | CACGATTGCTGATACCACATAC |
| GFP | dsGFP | dsGFP-F | TAATACGACTCACTATAGGGAAGGGCGAGGAGCTGTTCACCG |
| synthesis | dsGFP-R | TAATACGACTCACTATAGGGCAGCAGGACCATGTGATCGCGC |
Note: The underlined sequence represents the T7 promoter.
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; Zhu, Huan 1 ; Jin, Daochao 3
; Yang, Hong 3 ; Cao, Zhou 4 1 College of Environment and Life Sciences, Kaili University, Kaili 556011, China;
2 School of Ethnic-Minority Medicine, Guizhou Minzu University, Guiyang 550025, China;
3 Provincial Key Laboratory for Agricultural Pest Management of Mountainous Regions and Scientific Observation and Experimental Station of Crop Pests in Guiyang, Ministry of Agriculture and Rural Affairs of the People’s Republic of China, Institute of Entomology, Guizhou University, Guiyang 550025, China
4 College of Life Sciences, Chongqing Normal University, Chongqing 401331, China;