Next Article in Journal
Modeling Dynamics in Land Use and Land Cover and Its Future Projection for the Amazon Biome
Previous Article in Journal
Extractable Compounds in a Birch Tree—Variations in Composition and Yield Potentials
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Forests
Volume 14
Issue 7
10.3390/f14071280
forests-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

RNA Interference-Mediated Knockdown of Tryptophan 2,3-Dioxygenase and Kynurenine-3-Monooxygenase in Monochamus Alternatus: Implications for Insect Control

by
Minghui Zhang
1,2,†,
Xiaoqian Weng
1,2,†,
Qing Li
1,2,†,
Liangjing Sheng
1,2,
Yajie Guo
1,3,
Liya Xiong
1,2,
Feiping Zhang
1,2,* and
Songqing Wu
1,2,*
1
College of Forestry, Fujian Agriculture and Forestry University, Fuzhou 350002, China
2
Key Laboratory of Integrated Pest Management in Ecological Forests, Fujian Agriculture and Forestry University, Fuzhou 350002, China
3
Laboratory of Forest Symbiology, Asian Research Center for Bioresource and Environmental Sciences, Graduate School of Agricultural and Life Sciences, The University of Tokyo, Tokyo 188-0002, Japan
*
Authors to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Forests 2023, 14(7), 1280; https://doi.org/10.3390/f14071280
Submission received: 13 March 2023 / Revised: 31 May 2023 / Accepted: 13 June 2023 / Published: 21 June 2023
(This article belongs to the Section Forest Ecophysiology and Biology)
Browse Figures
Versions Notes

Abstract

:
Monochamus alternatus Hope (Coleoptera: Cerambycidae), an invasive beetle that has caused billions of dollars in economic losses, is a serious pest of Pinus massoniana in many Asian countries. Clarifying the eye pigment gene and its knockdown phenotype of M. alternatus can provide functional gene identification and a marker for screening of gene editing, as well as help develop new control ideas. In this study, we first screened the transcriptome and found one homologous gene of tryptophan 2,3-dioxygenase (TDO) and one of kynurenine-3-monooxygenase (KMO). By measuring the expression levels of TDO and KMO in different developmental periods, it was indicated that TDO and KMO were expressed in various stages of M. alternatus. The gene expression of MaKMO was higher than MaTDO, showing high expression after pupation and decreasing at the beginning of eclosion. MaTDO and MaKMO were knocked down using RNA interference technology in different periods of expression, and the temporal expression changes were obtained using RT-qPCR technology. The results showed that the expressions of MaTDO and MaKMO were significantly inhibited by the injection of dsRNA; the expressions of MaTDO at 48 h, 96 h and after pupation were 21.9%, 32.3% and 59.2%, respectively, meanwhile, those of KMO were 23.4%, 25.0% and 69.7%, respectively. There was a significant change in eye color, and the beetles were able to pupate normally without their activity being affected. Therefore, both MaTDO and MaKMO can be used as tag genes for M. alternatus. A dominant marker system based on eye color can be developed for the genetic manipulation and control of M. alternatus.

1. Introduction

Pine wood nematode disease is a devastating pine forest disease caused by Bursaphelenchus xylophilus. Originating in North America, the disease usually only damages exotic pine species, but has spread to Canada, the United States, Mexico, Spain, Portugal, Japan, China, South Korea, North Korea and other regions. It is causing massive die-offs of native pine trees in eastern Asian countries and in countries on the western edge of Europe [1]. At present, the most important vector of this disease is considered to be Monochamus alternatus [2,3]. Among them, M. alternatus is its main vector in Asia. A single M. alternatus can carry anywhere from hundreds to more than two hundred thousand B. xylophilus, creating an invasion port for them (spawning trough, feeding wound) [4,5,6]. Therefore, the control of M. alternatus can effectively prevent the spread of pine wood nematode disease. The common control methods mainly include chemical control, trapping control and biological control. In practical application, comprehensive control methods are often used, including dead wood burning combined with traditional chemical agents and traps [7]. However, for this devastating disease, it is expected that it will take a lot of human and material resources, as well as time, to achieve a radical cure with the existing means; thus, it is still necessary to explore new, efficient and green means of control. The rapid spread of the disease is mainly dependent on the reciprocal cycle between the pine wood nematode, the pine tree and the beetle. In order to completely stop the spread of the disease and at the same time delay and prevent the evolution of the resistant population, gene editing can be used to change the beetle’s transmission habits, feeding habits or reduce its reproductive ability.
The development of targeted genome editing technology plays an important role in the study of physiological functions such as growth, development and reproduction of insects, as well as in the analysis of behavioral regulatory mechanisms. It also provides a new idea for pest control. In 1996, Kim et al. fused zinc finger protein with the cleavage domain of the restriction enzyme Fok I to create a genome-editing technology that can recognize and cut specific DNA sequences using the zinc finger structure [8]. After innovation of zinc finger nucleases (ZFNs), upon emergence of transcription activator-like effecter nuclease (TALENs) and CRISPR/Cas, targeted genome-editing technology is becoming more efficient, less costly and easier to operate. In addition to common model insects such as D. melanogaster, Culex and Tribolium castaneum, the most updated targeted genome-editing technology, the CRISPR/Cas system, is also widely used in Diptera [9,10], Hymenoptera [11], Hemiptera [12], Lepidoptera [13,14,15] and Orthoptera [16]. Therefore, it is urgent to develop a feasible gene-editing system for M. alternatus and expand the system’s prevention methods. However, a major limitation in the establishment of editing systems is the availability of marker genes [17]. Marker genes play a specific role in gene editing, which can be directly verified in target insects. More important than their function is the means of clarifying insects’ mutant characters, and the uniqueness and safety of the products. Thus, the search for non-lethal phenotypic genes to verify the efficiency of the knockout system in M. alternatus is vital.
Currently, marker genes are divided into five types: (1) Morphological markers, which refer to visual or measurable external features such as appearance, color and body shape. (2) Cytometric labeling, which refers to the individual chromosome number and morphology differences analyzed. (3) Biochemical genetic markers, which refer to biochemical traits in organisms, such as blood type and isozyme, most of which are protein variants. (4) Immune markers, which are genetic markers that are based on immune proteins such as red blood cell antigens and thymic cell antigens. (5) Molecular genetic markers, which are used to make molecular markers on DNA through the variation of nucleotide sequences in genetic material between individuals and directly reflect genetic polymorphisms at the DNA level [18,19]. In practical applications, morphological markers are mostly used in larger animals, and biochemical genetic markers are mostly used in microorganisms. As early as 1997, ommochromes have been applied to Aedes aegypti as a genetic marker; then in T. castaneum, B. mori, Nilaparvata lugens and Helicoverpa zea (Boddie) as tag genes of editing systems; and in Anisopteromalus calandrae for the study of reproductive capacity [17,20,21,22,23,24,25].
In this study, we used M. alternatus as a model to identify the KMO and TDO transcriptome sequences, using RNA interference technology to obtain knockdown phenotypes, and examined the feasibility of disrupting eye color; the expression levels of the treated genes were obtained with RT-qPCR, which verified the feasibility of ocular pigment-related genes as tag genes for subsequent gene editing. This study will provide available target genes for the establishment of the gene-editing system for M. alternatus, provide technical reference for the application of RNA interference technology in M. alternatus and provide a basis for the biological control of M. alternatus.

2. Materials and Methods

2.1. Insect Husbandry

M. alternatus was captured in the wild and cultured in the laboratory. The feeding environment included a temperature of 27 ± 1 °C, relative humidity of 80% and a photoperiod of 14 h/10 h (D/N). The larvae were fed with homemade artificial feed (main ingredients: wheat bran, wood chips, sucrose, dry yeast, shrimp shell powder, water, preservatives). From pupa to sexually matured adult, the M. alternatus specimens were kept isolated from each other to prevent biting, and the adults were fed with fresh pine twigs that were one–two years old. The embryos were collected using sturdy pine trees that were 15–20 cm high.

2.2. Synthesis of cDNA

Pupal-stage M. alternatus specimens with obvious eye discoloration were selected from the test M. alternatus; their heads were taken, and the total RNA was extracted using a Trizol method. The concentration was measured using an ultraviolet spectrophotometer, and the integrity of the RNA was determined with gel electrophoresis. RNA samples with an OD 260/280 of about 2.0 and a complete band were selected as the reverse transcription template. The cDNA was synthesized using a HiScript 1st Strand cDNA Synthesis Kit (Vazyme, Nanjing, China) according to its instructions for subsequent experiments.

2.3. Synthesis and Purification of dsRNA

Complete sequences of the KMO and TDO genes were obtained via M. alternatus whole genome sequencing, and primers were designed online by NCBI (http://www.ncbi.nlm.nih.gov/ (accessed on 21 December 2021)).
The cDNA was amplified using PCR with specific primers (Table 1), then purified with an E. Z. N. A.® Gel Extraction Kit (Omega, Norcross, GA, USA). The concentration and purity were detected via a UV spectrophotometer, and the band size was detected via gel electrophoresis to obtain pure dsDNA. dsRNA was synthesized in vitro using the T7 RNAi Transcription Kit (Vazyme, Nanjing, China): 500 ng of dsDNA template was added to the reaction system at 20 μL, incubated at 37 °C for 4 h or overnight, then added with 1 μL of DNase 1, 2 μL of RNase T1 and 17 μL of nuclease-free water and incubated at 37 °C for 30 min. The dsDNA template in the solution was eliminated. The dsRNA was purified using VAHTS RNA Clean Beads (Vazyme, Nanjing, China) according to the manufacturer’s instructions and stored at −80 °C for future use. Considering the requirements of injection volume and the effect of RNAi for subsequent injection, the OD 260/280 of dsRNA after purification was about 2.0, and the concentration was not less than 500 ng/uL.

2.4. Injection

The injection objects were M. alternatus at the beginning of the pupa stage, with injection of the abdominal site between two quarters. Each insect was injected with 1 μg of dsRNA (MaTDO, MaKMO, GFP), and the control group was injected with the same amount of nuclease-free water. After injection, the pupae were placed in a damp paper towel for breeding, with an ambient temperature of 27 °C, relative humidity of 80% and dark treatment. Each treatment required 60 pupae. A total of 5 pupae were collected at 48 h, 96 h and eclosion as a biological replicate, and 3 biological replicates were collected at each time point and frozen at −80 °C for the subsequent RT-qPCR analysis.

2.5. Expression Analysis of RT-qPCR

RNA was extracted using Trizol and then reverse-transcribed into cDNA using the same procedure as in Section 2.2. The RT-qPCR reaction system was configured according to the manual (Table 2), using ChamQ Universal SYBR qPCR Master Mix (Vazyme, Nanjing, China). The cDNA was diluted by a factor of 10 for the RT-qPCR pre-experiments. A concentration of “S” type whose amplification curve was smooth was selected as the final addition amount. In iQ SYBR Green Supermix-validated primers used for testing, using the ΔΔC (t) analysis method, the reaction conditions were as follows: predenaturation at 95 °C for 30 s, 95 °C for 10 s, 60 °C for 30 s, for a total of 40 cycles. The results of the treatment group and the experimental group were compared after reference gene correction. The data were processed using the 2−ΔΔCt method and analyzed using GraphPad.

3. Results

3.1. Phylogenetic Tree and Sequence Analysis of MaKMO and MaTDO Genes

Through whole genome sequencing, the results showed that there was a KMO gene on the seventh autosome and a TDO gene on the fifth autosome. In order to compare the homology of the KMO and TDO genes among other insects, a phylogenetic analysis was conducted, and a molecular phylogenetic tree was constructed. MaKMO was compared with Athalia rosae, Chrysoperla carnea, Lygus hesperus, Bradysia coprophila, Anopheles sinensis, Helicoverpa armigera, Spodoptera litura, Arctia plantaginis, T. castaneum and Anoplophora glabripennis; MaTDO was compared with Pararge aegeria, H. zea, Ctenocephalides felis, A. albimanus, A. merus, A. glabripennis, Asbolus verrucosus, T. castaneum and Leptinotarsa decemlineata. The reliability of each branch was above 70%. By comparing the genetic distances of other organisms, MaKMO and MaTDO were the most similar to A. glabripennis, the same member of Coleoptera. The MaTDO gene was clustered in the same clade with A. verrucosus and T. castaneum, and the similarity was more than 70%. However, in KMO, there was no high similarity between M. alternatus and T. castaneum (Figure 1B).
The MaKMO and MaTDO protein domains were predicted using SMART website (http://smart.embl-heidelberg.de/ (accessed on 15 February 2022)).
The results showed that the transmembrane region of MaKMO was mainly concentrated in the amino terminus and the carboxyl terminus, while there was no transmembrane region of MaTDO; only a small transmembrane region existed at about 100 bp. Therefore, MaKMO genes are likely to act as membrane receptors, or as membrane-anchoring proteins or ion channel proteins that are localized to membrane proteins (Figure 2).

3.2. Expressions of MaKMO and MaTDO at Different Stages

The expression levels of MaKMO and MaTDO in different stages were measured. It was found that MaKMO was expressed in the growth of M. alternatus, and that the expression level of MaKMO was higher than that of MaTDO. The expression levels of MaTDO appear to be almost absent except at the second instar larval stage. MaKMO also showed a small peak in the second instar larval stage, and the expression level was also low in the larval and pupal stages. However, it was observed that the expression level of MaKMO in both female and male larvae began increasing from the pupal stage, then began decreasing in the adult stage, suggesting that the MaKMO gene may be expressed in large quantities in the pupal stage. At the same time, the difference in expression between male and female may indicate that MaKMO is also involved in other expression pathways related to gender, which needs to be verified (Figure 3).

3.3. The Effect of RNA Interference of MaKMO and MaTDO on Eye Color

The synthesized dsKMO and dsTDO were injected into M. alternatus one day after pupation. The phenotypes on days 7, 9, 11 and eclosion after injection are shown in Figure 4. The results showed that there were obvious differences in eye color between the treatment group and the control group from the seventh day. When increasing time, this difference became more obvious, and it was observed that the eyes of M. alternatus showed a gradual deepening from the center to the edge. It was found that the knockdown of MaKMO or MaTDO alone significantly reduced eye color. This may be because tryptophan is first oxidized by MaTDO to produce formyl kynurenine, which provides a reaction substrate for MaKMO. The formation of xanthommatin is regulated by both of these genes, and the deletion of either gene will lead to interruption of the pathway. At the same time, there was no dilution of the color of the body surface, indicating that the gene did not affect the formation of pigment other than eye pigment. After injection, the surviving M. alternatus were reared until sexual maturity, with the eggs being collected using pine wood. There was no difference in appearance between the knockdown individuals and the wild-type, and they could mate normally and produce hatchable eggs. Therefore, the MaKMO and MaTDO can be used as potential marker genes without affecting reproduction.

3.4. RT-qPCR after MaKMO and MaTDO RNA Interference

The expression levels of MaKMO and MaTDO in the pupae of the treated group and the control group were significantly decreased via RT-qPCR analysis. The silencing effect decreased as development proceeded, but it could continue until the initial emergence. When injected with nuclease-free water as a negative control, dsGFP expression levels were comparable, with a difference of 1%–6% at the measured time points. The results indicated that dsGFP had no significant effect on the expression level. The expression level of MaKMO decreased by 77% after 48 h, 75% after 96 h and 30% after emergence, compared with the negative control. After injection of MaTDO, the expression level decreased by 78% after 48 h, 68% after 96 h and 41% after emergence, compared with negative control. By comparing the expression levels of MaKMO and MaTDO, there was no significant difference in the silencing effect between the two groups under the condition of injecting the same amount of dsRNA (1 μg), and both genes could effectively inhibit the synthesis of xanthommatin (Figure 5).

4. Discussion

Gene-editing technology has developed rapidly and has been widely used in recent years. Among the techniques, ZFNs have been successfully applied to D. melanogaster [26], Silkworm [27], A. aegypti [28] and Danaus plexippus [29] gene knockout and site-specific modifications. TALENs has been applied in the silkworm BmBLOS2 gene, resulting in an oily silkworm phenotype and succeeding in B. mori clock gene period e knockout; gene knockout of Ostrinia furnacalis sex pheromone related receptor gene resulted in abnormal mating behavior [30,31]. The CRISPR/Cas system has also been used in D. melanogaster [9], A. aegypti [10], Locusta migratoria [32] and Blattella germanica [33]. However, these techniques have not been applied in M. alternatus. In the development of gene-editing tools, a visible and harmless gene is needed that can be used as a marker gene to verify the success of the gene editing. This study proved that MaTDO and MaKMO were highly conserved sequences, and that their phenotypes after knockdown were defined. Moreover, knockdown of MaTDO and MaKMO did not affect the life activities of M. alternatus, and these genes can be used as safe marker genes.
Ommochromes are a kind of pigment that is unique to protostomes, and they are the only source of eye pigment in many insects. Based mainly on phenoxazone and a phenothiazine ring, they can form different hair groups so that phenothiazine can absorb different specific wavelengths by changing the side chain or REDOX state. Ommochromes present different colors, giving the insect a color range that goes from yellow to red to purple. These molecules are divided into three groups based on their permeability: ommatins, ommidins and ommins. Firstly, tryptophan 2,3-dioxygenase can accelerate the metabolism of tryptophan to form formyl kynurenine by reacting with the tryptophan indole ring, which plays an important role in the oxidation of tryptophan [34]. Kynurenine undergoes redox by kynurenine 3-monooxygenase to form 3-hydroxykynurenine. The mechanism for the conversion of 3-hydroxycanuridine into xanthommatin, ommin A, ommatin D and other monochromators has been hypothesized, but has not been proven yet. It has been established that the main pathway of tryptophan catabolism in vertebrates and invertebrates is the kynurenine pathway [35,36]. In L. hesperus Knight, the FAD-binding domain was identified as a typical lutein-dependent hydroxylase by comparing it with different eye pigments from D. melanogaster using BLASTx [37]. In Tenebrio molitor, the TDO gene was silenced as a screening label for its mutant phenotype [38]. This experiment verified that KMO and TDO are involved in the formation of ocular lutein, which is the main determinant of ocular color. The MaKMO and MaTDO can be used as potential marker genes of M. alternatus.
RNAi technology has been used in biological functional verification since as early as 2002 when it was applied to silkworm to block the expression of its pigment genes [20], and then, through the injection of wing formation related genes, through the generation of defective types, to study gene function [39]. Subsequently, the function of AaGATAr, a receptor for vitellogenin in A. aegypti, was successfully identified [40]. Via RNAi silencing of the expression of dicer-1 in Blattella germanica, it was found that microRNAs play a key role in the process of abnormal behavior [41]. Expression of dsRNA designed against insect target genes in transgenic plants has been shown to give protection against pests through RNAi. However, RNAi efficiency varies greatly among different insects. Compared to other insects, RNAi is more efficient and systematic in Coleoptera insects, most notably the western corn rootworm, a devastating pest impacting corn production in the United States [42]. However, differences in intercellular transport, dsRNA-to-siRNA processing, and RNA-induced silencing complex formation can still affect RNAi efficiency [43]. Common RNAi methods in insects include microinjection, feeding, and soaking. Microinjection can be applied to different developmental stages of insects. Studies have shown that RNAi is effective for 4–5 instar larvae or 1-day-old pupae in M. alternatus, but the effective dsRNA mass span is large (10–300 ng) [44,45]. In this experiment, the injection volume was 1 μg. In the absence of supplementary injections, an obvious light-yellow color was still observed in the eyes of M. alternates after its emergence, indicating that dsRNA can exist for a long time and degrade slowly in M. alternates over a long time; and it therefore has a good silencing effect. In comparison with other eye pigment knockdowns of M. alternatus in previous studies [46], this experiment showed higher inhibitory effects on the expression level 48 h after injection, and showed slightly lower inhibitory levels at 96 h and emergence. These results indicated that the injection of dsRNA in the pupal stage of M. alternatus was feasible.

5. Conclusions

RNAi-based gene silencing studies have been attempted in many species. However, frontier research is relatively primary, and there are few studies on M.alternatus RNAi. In this study, a KMO gene and TDO gene were identified and confirmed to be involved in the synthesis of M.alternatus eye pigment via RNAi-based gene silencing. In addition, studies showed that pupal injections and gene silencing are feasible, resulting in a decrease in the expression of TDO transcripts and resulting in a white-eye phenotype in treated individuals. This study described a genetic marker in M.alternatus that can cause phenotypic changes, which can be used as markers for future transgenic strategies to genetically engineer this pest.

Author Contributions

Conceptualization, S.W. and F.Z.; methodology, M.Z. and X.W.; data curation, M.Z., X.W. and Q.L.; Project administration, F.Z.; Resources, S.W.; Supervision, Y.G. and L.S.; formal analysis, M.Z. and L.X.; writing—original draft preparation, M.Z.; writing—review and editing, L.S., X.W. and Q.L. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Key R & D Program of China (grant number 2021YFD1400900); the National Major Emergency Science and Technology Program of China (grant number ZD202001); the National Natural Science Foundation of China (grant numbers U1905201 and 32171805); the Forestry Key Program of Science and Technology in Fujian Province (grant number 2021FKJ03); the Natural Science Foundation of Fujian Province, China (grant number 2021J01056); the Forestry Programs of Science and Technology in Fujian Province (grant number Mincaizhi (2020) 601); the Science and Technology Program of Fujian Province (grant number 2018N5002); the Forestry Science Research Project of Fujian Forestry Department (grant number Minlinke (2017) 03); the Forestry Peak Discipline Construction Project of Fujian Agriculture and Forestry University (grant number 72202200205); the Forest Science Peak Project of College of Forestry, Fujian Agriculture and Forestry University (grant number 71201800720) and the Undergraduate Training Program for Innovation and Entrepreneurship of China (grant numbers 202210389029, X202210389174, and X202210389176).

Data Availability Statement

The data presented in this study are available in the article.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Ye, J.R. Analysis on epidemic status, control techniques and countermeasures of pine wilt disease in China. For. Sci. 2019, 55, 1–10. [Google Scholar]
  2. Tiao, N.Y.; Yu, L.F.; Jian, T.; Sun, J.H. Advances in research on Bursaphelenchus xylophilus and its key vector Monochamus spp. Entomol. Knowl. 2004, 41, 97–104. [Google Scholar]
  3. Kim, D.S.; Lee, S.M.; Chung, Y.J.; Chung, Y.J.; Moon, Y.S.; Park, C.G. Emergence Ecology of Japanese Pine Sawyer, Monchamus alternatus(Coleptera: Cerambycidae), a Vector of Pinewood Nematode, Bursaphelenchus xylophilus. Korean J. Appl. Entomol. 2003, 42, 307–313. [Google Scholar]
  4. Zhang, X.Y.; Luo, Y.Q. Major Forest Diseases and Insect Pests in China, 1st ed.; Chinese Forestry Publisher: Beijing, China, 2003; p. 386. [Google Scholar]
  5. Fielding, N.J.; Evans, H.F. The pine wood nematode Bursaphelenchus xylophilus (Steiner and Buhrer) Nickle (= B. lignicolus Mamiya and Kiyohara): An assessment of the current position. For. Int. J. For. Res. 1996, 69, 35–46. [Google Scholar] [CrossRef] [Green Version]
  6. Lee, K.S. Stability Analysis and Optimal Control Strategy for Prevention of Pine Wilt Disease. Abstr. Appl. Anal. 2014, 2014, 182680. [Google Scholar] [CrossRef] [Green Version]
  7. Zhao, B.G.; Futai, K.; Sutherland, J.R.; Takeuchi, Y. Pine Wilt Disease, 1st ed.; Springer: Tokyo, Japan, 2008; p. 459. [Google Scholar]
  8. Kim, Y.G.; Cha, J.; Chandrasegaran, S. Hybrid restriction enzymes: Zinc finger fusions to Fok I cleavage domain. Proc. Natl. Acad. Sci. USA 1996, 93, 1156–1160. [Google Scholar] [CrossRef] [Green Version]
  9. Trivedi, D. Using CRISPR-Cas9-based genome engineering tools in Drosophila melanogaster. Prog. Mol. Biol. Transl. Sci. 2021, 180, 85–121. [Google Scholar]
  10. Zulhussnain, M.; Zahoor, M.K.; Ranian, K.; Ahmad, A.; Jabeen, F. CRISPR Cas9 mediated knockout of sex determination pathway genes in Aedes aegypti. Bull. Entomol. Res. 2023, 113, 243–252. [Google Scholar] [CrossRef]
  11. Cheng, F.P.; Hu, X.F.; Pan, L.X.; Gong, Z.X.; Qin, K.X.; Li, Z.; Wang, Z.L. Transcriptome changes of Apis mellifera female embryos with fem gene knockout by CRISPR/Cas9. Int. J. Biol. Macromol. 2023, 229, 260–267. [Google Scholar] [CrossRef]
  12. Heu, C.C.; McCullough, F.M.; Luan, J.; Rasgon, J.L. CRISPR-Cas9-Based Genome Editing in the Silverleaf Whitefly (Bemisia tabaci). CRISPR J. 2020, 3, 89–96. [Google Scholar] [CrossRef] [Green Version]
  13. Zhao, J.; Tan, Y.; Jiang, Y.; Zhu-Salzman, K.; Xiao, L. CRISPR/Cas9-mediated methoprene-tolerant 1 knockout results in precocious metamorphosis of beet armyworm (Spodoptera exigua) only at the late larval stage. Insect Mol. Biol. 2023, 32, 132–142. [Google Scholar] [CrossRef] [PubMed]
  14. Guo, H.; Chen, F.; Zhou, M.; Lan, W.; Zhang, W.; Shen, G.; Lin, P.; Xia, Q.; Zhao, P.; Li, Z. CRISPR-Cas9-Mediated Mutation of Methyltransferase METTL4 Results in Embryonic Defects in Silkworm Bombyx mori. Int. J. Mol. Sci. 2023, 24, 3468. [Google Scholar] [CrossRef] [PubMed]
  15. Wang, J.; Wang, H.; Liu, S.; Liu, L.; Tay, W.T.; Walsh, T.K.; Yang, Y.; Wu, Y. CRISPR/Cas9 mediated genome editing of Helicoverpa armigera with mutations of an ABC transporter gene HaABCA2 confers resistance to Bacillus thuringiensis Cry2A toxins. Insect Biochem. Mol. Biol. 2017, 87, 147–153. [Google Scholar] [CrossRef] [PubMed]
  16. Bai, Y.; He, Y.; Shen, C.Z.; Li, K.; Li, D.L.; He, Z.Q. CRISPR/Cas9-Mediated genomic knock out of tyrosine hydroxylase and yellow genes in cricket Gryllus bimaculatus. PLoS ONE 2023, 18, e0284124. [Google Scholar] [CrossRef]
  17. Cornel, A.J.; Benedict, M.Q.; Rafferty, C.S.; Howells, A.J.; Collins, F.H. Transient expression of the Drosophila melanogaster cinnabar gene rescues eye color in the white eye (WE) strain of Aedes aegypti. Insect Biochem. Mol. Biol. 1997, 27, 993–997. [Google Scholar] [CrossRef]
  18. Tao, H.J.; Lin, L.; Su, Y.; Huang, D.N. The types and functions of genetic markers and the research status of rice genetic markers. China Rice 2011, 17, 21–24. [Google Scholar]
  19. Du, X.Y.; Huo, X.Y.; Chen, Z.W. Research progress and application of laboratory animal genetic quality monitoring technology. Lab. Anim. Sci. 2021, 38, 1–5. [Google Scholar]
  20. Quan, G.X.; Kim, I.; Kômoto, N.; Sezutsu, H.; Ote, M.; Shimada, T.; Kanda, T.; Mita, K.; Kobayashi, M.; Tamura, T. Characterization of the kynurenine 3-monooxygenase gene corresponding to the white egg 1 mutant in the silkworm Bombyx mori. Mol. Genet. Genom. MGG 2002, 267, 1–9. [Google Scholar] [CrossRef]
  21. Khanh, H.D.T.; Bressac, C.; Chevrier, C. Male sperm donation consequences in single and double matings in Anisopteromalus calandrae. Physiol. Entomol. 2005, 30, 29–35. [Google Scholar] [CrossRef]
  22. Quan, G.X.; Kobayashi, I.; Kojima, K.; Uchino, K.; Kanda, T.; Sezutsu, H.; Shimada, T.; Tamura, T. Rescue of white egg 1 mutant by introduction of the wild-type Bombyx kynurenine 3–monooxygenase gene. Insect Sci. 2007, 14, 85–92. [Google Scholar] [CrossRef]
  23. Sherry, A.; Marcé, L.; Brenda, O. Metabolic pathway interruption: CRISPR/Cas9-mediated knockout of tryptophan 2,3-dioxygenase in Tribolium castaneum. J. Insect Physiol. 2018, 107, 104–109. [Google Scholar]
  24. Perera, O.P.; Little, N.S.; Pierce, C.A. CRISPR/Cas9 mediated high efficiency knockout of the eye color gene Vermillion in Helicoverpa zea (Boddie). PLoS ONE 2018, 13, e0197567. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. Liu, S.H.; Luo, J.; Yang, B.J.; Wang, A.Y.; Tang, J. Karmoisin and cardinal ortholog genes participate in the ommochrome synthesis of Nilaparvata lugens (Hemiptera: Delphacidae). Insect Sci. 2017, 26, 35–43. [Google Scholar] [CrossRef] [Green Version]
  26. Lin, S.C.; Chang, Y.Y.; Chan, C.C. Strategies for gene disruption in Drosophila. Cell. Biosci. 2014, 4, 63. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  27. Tsubota, T.; Sakai, H.; Sezutsu, H. Genome Editing of Silkworms. Methods Mol. Biol. 2023, 2637, 359–374. [Google Scholar] [PubMed]
  28. McMeniman, C.J.; Corfas, R.A.; Matthews, B.J.; Ritchie, S.A.; Vosshall, L.B. Multimodal Integration of Carbon Dioxide and Other Sensory Cues Drives Mosquito Attraction to Humans. Cell. 2014, 156, 1060–1071. [Google Scholar] [CrossRef] [Green Version]
  29. Merlin, C.; Beaver, L.E.; Taylor, O.R.; Wolfe, S.A.; Reppert, S.M. Efficient targeted mutagenesis in the monarch butterfly using zinc-finger nucleases. Genome Res. 2013, 23, 159–168. [Google Scholar] [CrossRef] [Green Version]
  30. Ikeda, K.; Daimon, T.; Shiomi, K.; Udaka, H.; Numata, H. Involvement of the Clock Gene Period in the Photoperiodism of the Silkmoth Bombyx mori. Zool. Sci. 2021, 38, 523–530. [Google Scholar] [CrossRef]
  31. Yang, B.; Fujii, T.; Ishikawa, Y.; Matsuo, T. Targeted mutagenesis of an odorant receptor co-receptor using TALEN in Ostrinia furnacalis. Insect Biochem. Mol. Biol. 2016, 70, 53–59. [Google Scholar] [CrossRef]
  32. Guo, X.; Yu, Q.; Chen, D.; Wei, J.; Yang, P.; Yu, J.; Wang, X.; Kang, L. 4-Vinylanisole is an aggregation pheromone in locusts. Nature 2020, 584, 584–588. [Google Scholar] [CrossRef]
  33. Shirai, Y.; Piulachs, M.-D.; Belles, X.; Daimon, T. DIPA-CRISPR is a simple and accessible method for insect gene editing. Cell Rep. Methods 2022, 2, 100215. [Google Scholar] [CrossRef]
  34. Capece, L.; Arrar, M.; Roitberg, A.E.; Yeh, S.R.; Marti, M.A.; Estrin, D.A. Substrate stereo-specificity in tryptophan dioxygenase and indoleamine 2,3-dioxygenase. Proteins 2010, 78, 2961–2972. [Google Scholar] [CrossRef] [Green Version]
  35. Abdulla, A.B. Kynurenine Pathway of Tryptophan Metabolism: Regulatory and Functional Aspects. Int. J. Tryptophan Res. 2017, 10, 1178646917691938. [Google Scholar]
  36. Zhuravlev, A.V.; Vetrovoy, O.V.; Ivanova, P.N.; Savvateeva-Popova, E.V. 3-Hydroxykynurenine in Regulation of Drosophila Behavior: The Novel Mechanisms for Cardinal Phenotype Manifestations. Front. Physiol. 2020, 11, 971. [Google Scholar] [CrossRef] [PubMed]
  37. Baum, J.A.; Roberts, J.K. Chapter Five-Progress towards RNAi-Mediated Insect Pest Management. In Advances in Insect Physiology; Dhadialla, T.S., Gill, S.S., Eds.; Academic Press: Cambridge, MA, USA, 2014; Volume 47, pp. 249–295. [Google Scholar]
  38. Oppert, B.; Chu, F.C.; Reyna, S.; Pinzi, S.; Adrianos, S.; Perkin, L.; Lorenzen, M. Effects of targeting eye color in Tenebrio molitor through RNA interference of tryptophan 2,3-dioxygenase (vermilion): Implications for insect farming. Arch. Insect Biochem. Physiol. 2019, 101, e21546. [Google Scholar] [CrossRef]
  39. Kennerdell, J.R.; Carthew, R.W. Heritable gene silencing in Drosophila using double-stranded RNA. Nat. Biotechnol. 2000, 18, 896–898. [Google Scholar] [CrossRef] [PubMed]
  40. Attardo, G.M.; Higgs, S.; Klingler, K.A.; Vanlandingham, D.L.; Raikhel, A.S. RNA interference-mediated knockdown of a GATA factor reveals a link to anautogeny in the mosquito Aedes aegypti. Proc. Natl. Acad. Sci. USA 2003, 100, 896–898. [Google Scholar] [CrossRef] [Green Version]
  41. Belles, X. The endocrine regulation of insect metamorphosis and the emerging role of microRNAs. Front. Endocrinol. 2010, 1. [Google Scholar] [CrossRef]
  42. Brent, C.S.; Spurgeon, D.W. Egg Production and Longevity of Lygus hesperus (Hemiptera: Miridae) Adult Females Under Constant and Variable Temperatures. J. Entomol. Sci. 2019, 54, 69–80. [Google Scholar] [CrossRef]
  43. Zhu, K.Y.; Palli, S.R. Mechanisms, Applications, and Challenges of Insect RNA Interference. Annu. Rev. Entomol. 2020, 65, 293–311. [Google Scholar] [CrossRef] [Green Version]
  44. Niu, B.L.; Shen, W.F.; Liu, Y.; Weng, H.B.; He, L.H.; Mu, J.J.; Wu, Z.L.; Jiang, P.; Tao, Y.Z.; Meng, Z.Q. Cloning and RNAi-mediated functional characterization of MaLac2 of the pine sawyer, Monochamus alternatus. Insect Mol. Biol. 2008, 17, 303–312. [Google Scholar] [CrossRef] [PubMed]
  45. Weng, H.; Shen, W.; Liu, Y.; He, L.; Niu, B.; Meng, Z.; Mu, J. Cloning and characterization of two EcR isoforms from Japanese pine sawyer, Monochamus alternates. Arch. Insect Biochem. Physiol. 2013, 84, 27–42. [Google Scholar] [CrossRef] [PubMed]
  46. Sheng, L.J.; Weng, X.Q.; Weng, M.Q.; Guo, Y.J.; Carballar-Lejarazú, R.; Zhang, F.P.; Wu, S.Q. Function of Tryptophan 2,3-Dioxygenase in Monochamus alternatus Hope Revealed by RNA Interference. Forests 2023, 14, 215. [Google Scholar] [CrossRef]
Forests 14 01280 g001 550
Figure 1. (A). Alignment of MaKMO and MaTDO predicted protein sequences. The contrast sequences came from A. verrucosus (RZC41040. 1), A. glabripennis (XP_018573815. 1, NP_001034500. 1), T. castaneum (NP_001034499. 1, XP_018568258. 1) and C. septempunctata (XP_044756441. 1). Red background indicates conserved amino acids, blue box calculated from all groups with a global similarity score > 0.7. Gaps have been introduced to permit alignment. (B). Phylogenetic analysis of MaKMO and MaTDO. Other TDO proteins were L. decemlineata (XP_023026760. 1), H. zea (AVR75813. 1), C. felis (XP_026480894. 1), A. albimanus (XP_035795822. 1), A. merus (XP_041762204. 1), A. glabripennis (XP_018573815. 1), A. verrucosus (RZC41040. 1), T. castaneum (NP_001034499. 1) and P. aegeria (XP_039748089. 1); Other KMO proteins were A. rosae (XP_025602652. 1), C. carnea (XP_044742432. 1), L. hesperus (AZS64105. 1), B. coprophila (XP_037043194. 1), A. sinensis (KFB47145. 1), H. armigera (XP_021183543. 1), S. litura (XP_022820213. 1), A. plantaginis (CAB3251823. 1), T. castaneum (NP_001034500. 1) and A. glabripennis (XP_018568258. 1). The tree was constructed using the neighbor-joining method based on the full-length protein sequence alignments. Bootstrap analyses of 1000 replications were carried out, and bootstrap values > 50% are shown on the tree.
Figure 1. (A). Alignment of MaKMO and MaTDO predicted protein sequences. The contrast sequences came from A. verrucosus (RZC41040. 1), A. glabripennis (XP_018573815. 1, NP_001034500. 1), T. castaneum (NP_001034499. 1, XP_018568258. 1) and C. septempunctata (XP_044756441. 1). Red background indicates conserved amino acids, blue box calculated from all groups with a global similarity score > 0.7. Gaps have been introduced to permit alignment. (B). Phylogenetic analysis of MaKMO and MaTDO. Other TDO proteins were L. decemlineata (XP_023026760. 1), H. zea (AVR75813. 1), C. felis (XP_026480894. 1), A. albimanus (XP_035795822. 1), A. merus (XP_041762204. 1), A. glabripennis (XP_018573815. 1), A. verrucosus (RZC41040. 1), T. castaneum (NP_001034499. 1) and P. aegeria (XP_039748089. 1); Other KMO proteins were A. rosae (XP_025602652. 1), C. carnea (XP_044742432. 1), L. hesperus (AZS64105. 1), B. coprophila (XP_037043194. 1), A. sinensis (KFB47145. 1), H. armigera (XP_021183543. 1), S. litura (XP_022820213. 1), A. plantaginis (CAB3251823. 1), T. castaneum (NP_001034500. 1) and A. glabripennis (XP_018568258. 1). The tree was constructed using the neighbor-joining method based on the full-length protein sequence alignments. Bootstrap analyses of 1000 replications were carried out, and bootstrap values > 50% are shown on the tree.
Forests 14 01280 g001
Forests 14 01280 g002 550
Figure 2. Prediction of protein transmembrane regions of MaKMO and MaTDO.
Figure 2. Prediction of protein transmembrane regions of MaKMO and MaTDO.
Forests 14 01280 g002
Forests 14 01280 g003 550
Figure 3. Quantitative expressions of MaKMO and MaTDO in different developmental stages. NF: new female pupae, OF: old female pupae, F1: 1-day-old female, F20: 20-days-old female, NM: new male pupae, OM: old malepupae, M1: 1-day-old male, M8: 8-days-old male, L1: first instar larva, L2: second instar larva; L3: third instar larva; L4: fourth instar larva.
Figure 3. Quantitative expressions of MaKMO and MaTDO in different developmental stages. NF: new female pupae, OF: old female pupae, F1: 1-day-old female, F20: 20-days-old female, NM: new male pupae, OM: old malepupae, M1: 1-day-old male, M8: 8-days-old male, L1: first instar larva, L2: second instar larva; L3: third instar larva; L4: fourth instar larva.
Forests 14 01280 g003
Forests 14 01280 g004 550
Figure 4. The phenotypes of MaKMO and MaTDO injections in pupae at different stages.
Figure 4. The phenotypes of MaKMO and MaTDO injections in pupae at different stages.
Forests 14 01280 g004
Forests 14 01280 g005 550
Figure 5. The phenotype of MaKMO and MaTDO injections in pupae at different stages. GFP-MaKMO: MaKMO expression after injection of dsGFP. GFP-MaTDO: MaTDO expression after injection of dsGFP. Different letters indicate significant differences at a p-value < 0.05 using one-way ANOVA with the Tukey–Kramer test.
Figure 5. The phenotype of MaKMO and MaTDO injections in pupae at different stages. GFP-MaKMO: MaKMO expression after injection of dsGFP. GFP-MaTDO: MaTDO expression after injection of dsGFP. Different letters indicate significant differences at a p-value < 0.05 using one-way ANOVA with the Tukey–Kramer test.
Forests 14 01280 g005
Table
Table 1. Primers of dsRNA synthesis.
Table 1. Primers of dsRNA synthesis.
Primer NamesSequences 5′–3′Application
MaKMO-FTAATACGACTCACTATAGGGAGAAAGTAGCGGCTGGAAGATCGdsRNA synthesis primer
MaKMO-RTAATACGACTCACTATAGGGAGATGCGCCATCAGCTCCAATTA
MaTDO-FTAATACGACTCACTATAGGGAGATGGGGGAAATACCAACGAGC
MaTDO-RTAATACGACTCACTATAGGGAGACTTCTCATGGCGGAGGTGAG
Table
Table 2. Primers of RT-qPCR.
Table 2. Primers of RT-qPCR.
Primer NamesSequences 5′–3′Application
Actin-FAGCCGGTTTCGCCGGTGATGACRT-qPCR primer
Actin-RCACTTCATGATGGAGTTGTAGAC
MaKMO-qPCR-FTAAATGCGCCATCAGCTCCA
MaKMO-qPCR-RGAGACCCTCGACGATCAAGC
MaTDO-qPCR-FAGAACAAACTTGGCGTCCGA
MaTDO-qPCR-RTTGCTTGGTGGCATCCTCAT
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Zhang, M.; Weng, X.; Li, Q.; Sheng, L.; Guo, Y.; Xiong, L.; Zhang, F.; Wu, S. RNA Interference-Mediated Knockdown of Tryptophan 2,3-Dioxygenase and Kynurenine-3-Monooxygenase in Monochamus Alternatus: Implications for Insect Control. Forests 2023, 14, 1280. https://doi.org/10.3390/f14071280

AMA Style

Zhang M, Weng X, Li Q, Sheng L, Guo Y, Xiong L, Zhang F, Wu S. RNA Interference-Mediated Knockdown of Tryptophan 2,3-Dioxygenase and Kynurenine-3-Monooxygenase in Monochamus Alternatus: Implications for Insect Control. Forests. 2023; 14(7):1280. https://doi.org/10.3390/f14071280

Chicago/Turabian Style

Zhang, Minghui, Xiaoqian Weng, Qing Li, Liangjing Sheng, Yajie Guo, Liya Xiong, Feiping Zhang, and Songqing Wu. 2023. "RNA Interference-Mediated Knockdown of Tryptophan 2,3-Dioxygenase and Kynurenine-3-Monooxygenase in Monochamus Alternatus: Implications for Insect Control" Forests 14, no. 7: 1280. https://doi.org/10.3390/f14071280

APA Style

Zhang, M., Weng, X., Li, Q., Sheng, L., Guo, Y., Xiong, L., Zhang, F., & Wu, S. (2023). RNA Interference-Mediated Knockdown of Tryptophan 2,3-Dioxygenase and Kynurenine-3-Monooxygenase in Monochamus Alternatus: Implications for Insect Control. Forests, 14(7), 1280. https://doi.org/10.3390/f14071280

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop