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Review

RNA-Interference-Mediated Aphid Control in Crop Plants: A Review

by
Jiahui Zhang
1,2,
Huiyuan Li
1,2,
Xue Zhong
1,2,
Jinfu Tian
1,2,
Arnaud Segers
2,
Lanqin Xia
1,3,* and
Frédéric Francis
2,*
1
Institute of Crop Sciences (ICS), Chinese Academy of Agricultural Sciences (CAAS), Beijing 100081, China
2
Functional and Evolutionary Entomology, Gembloux Agro-Bio Tech, University of Liège, B-5030 Gembloux, Belgium
3
Hainan Yazhou Bay Seed Laboratory, National Nanfan Research Institute (Sanya), Chinese Academy of Agricultural Sciences, Sanya 572024, China
*
Authors to whom correspondence should be addressed.
Agriculture 2022, 12(12), 2108; https://doi.org/10.3390/agriculture12122108
Submission received: 26 October 2022 / Revised: 2 December 2022 / Accepted: 7 December 2022 / Published: 9 December 2022
(This article belongs to the Special Issue Sustainable Pest Management in Agriculture)
Browse Figure
Review Reports Versions Notes

Abstract

:
Crop plants suffer severe yield losses due to the significant damages caused by aphids. RNA interference (RNAi) technology is a versatile and environmentally friendly method for pest management in crop protection. Transgenic plants expressing siRNA/dsRNA and non-transformative methods such as spraying, microinjection, feeding, and a nanocarrier-delivery-mediated RNAi approach have been successfully applied for agricultural insect pest management. In this review, we summarize the application of host-induced gene silencing (HIGS)-mediated RNAi, spray-induced gene silencing (SIGS)-mediated RNAi, and other delivery-method-mediated RNAi methods for aphid control. We further discuss the challenges in RNAi application and propose potential solutions to enhance RNAi efficiency.

1. Introduction

Cereal plants are frequently attacked sequentially or simultaneously by different aphid species, significantly reducing the quality and quantity of grain. Although chemical control could successfully suppress aphid populations, it has accelerated insecticide resistance development and led to pest resurgence. The overuse of chemical pesticides has led to severe environmental problems and threatens human health [1]. Therefore, to guarantee food safety and security, it is important and imperative to develop effective pest management approaches to control aphid damage to cereals. Extensive research in recent decades has typically concentrated on further understanding crop–aphid interactions, which has significantly facilitated the development of sustainable aphid management strategies [2].
RNA interference (RNAi) is a biological process that can be triggered by endogenously expressed or exogenously applied double-stranded RNAs (dsRNAs). In this process, transcriptional silencing is induced by directing inhibitory chromatin modifications, and post-transcriptional silencing is induced by decreasing the stability or translation capability of the targeted mRNA [3,4,5,6,7,8]. The RNAi technique has enormous potential applications in agricultural practices, extending to viruses, bacteria, fungi, nematodes, insects, and plants. RNAi-mediated control has been exploited for several phloem-feeding aphids via targeting essential genes involved in ingestion, molting, development, and fecundity [9]. With applications in crop protection and production, host-induced gene silencing (HIGS), which employs transgenic plants that have been precisely engineered to produce dsRNA, and spray-induced gene silencing (SIGS), which uses topically applied dsRNA molecules, are being exploited. Here, we summarize the RNAi-based protection against different aphid species in crop plants, discuss the challenges associated with RNAi application, and propose potential solutions to improve RNAi efficiency.

2. RNA-Interference-Based Aphid Control in Crop Plants

The first evidence of RNA-induced gene silencing was described in pigmented petunia petals when they attempted to overexpress a key gene involved in flavonoid biosynthesis named chalcone synthase (CHS) but blocked anthocyanin biosynthesis via a post-transcriptional mechanism [10]. A subsequent investigation demonstrated that dsRNA resulted in the decreased or eliminated expression of a target transcript in Caenorhabditis elegans. This discovery established that dsRNA was more effective than single-stranded RNA (ssRNA), which represented an extraordinary milestone in the RNAi revolution [11]. Since the discovery that dsRNA induces effective target gene silencing, a variety of techniques have been investigated to deliver dsRNA in insect species. In laboratory or agricultural practice, exogenous RNAs are applied through surface treatments or invasive methods such as spraying, soaking, injection, infiltration, soil/root drenching, and petiole absorption [12,13,14,15,16]. Plant-mediated and insect-mediated RNAi have been exploited as promising alternative strategies for pest management [17,18,19] (Figure 1). The application of RNAi through expressing dsRNA in transgenic crop plants or utilizing dsRNA directly as an insecticide appears promising for agricultural pest control, which can be achieved by host-induced gene silencing (HIGS) and spray-induced gene silencing (SIGS) [20,21].

2.1. Host-Induced Gene Silencing

Host-induced gene silencing is known as a plant-mediated transgenic strategy in which plants are genetically engineered to produce pest- or pathogen-gene-targeting sRNAs or dsRNAs. Subsequently, these RNAs are transported into the pest or pathogen to silence target genes [22,23].
The HIGS molecular mechanisms in insects may differ from those in fungi. In herbivorous insects, long dsRNAs (including hpRNAs) appear to be absorbed directly from the host. Then, gene silencing is induced via RNAi machinery. In fungi, the existing evidence indicates that gene silencing is induced through taking up siRNAs and microRNAs (miRNAs) produced by the host plant [24].
Host-induced gene silencing (HIGS) was first reported in Arabidopsis thaliana. With an expressed hpRNA of a nematode 16D10 gene, transgenic plants exhibited significant resistance against four main root-knot nematode species [25]. The first proof-of-concept research on plant-mediated dsRNA delivery for insect pest management was reported in western corn rootworm (WCR). In a growth chamber assay, transgenic maize plants expressing WCR dsRNAs significantly reduced the damage caused by WCR feeding [26]. Subsequently, numerous studies have been reported using HIGS in crop plants to protect against various plant pathogens and pests, including fungi [27,28], oomycetes [29,30], and insects [31,32].

2.2. Host-Induced Gene Silencing Based Protection of Crop Plants from Aphids

HIGS has great potential to manage insects from the order Hemiptera that feed on plants, especially aphids. The application of HIGS has been exploited in different aphid species, including the peach aphid (Myzus persicae) and grain aphid (Sitobion avenae) (Table 1).
Many studies of HIGS focused on M. persicae through various transgenic plants, for example, Arabidopsis thaliana, Nicotiana benthamiana, and Solanum lycopersicon. Some salivary effectors have been identified in aphids, such as MpC002, MpPIntO1 (Mp1), MpPIntO2 (Mp2), and Mp55. The knockdown of these genes reduced the reproduction of aphids, which indicated that these effectors could be selected as potential RNAi targets [17,33,34]. Rack-1 is a conserved multifunctional scaffold protein that was identified as a luteovirus-binding protein in peach aphids. The knockdown of Rack-1 reduced the fecundity of peach aphids [17]. Based on previous studies of Rack1, MpC002, and MpPIntO2, the persistence and transgenerational effects of plant-mediated RNAi were also investigated through transgenic Arabidopsis [35]. Transgenic tomato plant mediated RNAi has been shown to effectively silence the Acetylcholinesterase 1 (Ace1) gene and reduce the fecundity of peach aphids when fed transgenic plants [36]. A study reported that the knockdown of the cysteine protease Cathepsin B3 (CathB3) gene improved the performance of a non-tobacco-adapted (NTA) aphid lineage on tobacco. CathB3 elicited host defenses to suppress phloem sap ingestion by the aphid [37]. Plastid-mediated RNA interference (RNAi) was successfully employed to silence MpDhc64C. Both transgenic and transplastomic tobacco plants exhibited significant resistance to peach aphids, as demonstrated by decreased survival fecundity and survivor weight [38].
Most of the studies on S. avanae were applied by wheat-mediated HIGS. A particle-bombardment-mediated wheat transformation method was used to obtain stable transgenic wheat plants. Feeding on transgenic wheat expressing the carboxylesterase (CbE E4) gene could suppress the expression level of CbE E4 in grain aphids and impair larval tolerance to phoxim insecticides [18]. Silencing the lipase maturation factor 2-like (lmf2-like) gene reduced the molting number and decreased the survival and reproduction of aphids [39]. Similarly, the knockdown of the Chitin synthase 1 (CHS1) gene reduced the molting and survival of aphids [40]. Silencing the G protein (Gqα) gene could also reduce reproduction and molting in grain aphids [41]. Silencing the zinc finger protein (SaZFP) gene led to high mortality and decreased fecundity of grain aphids. The transgenerational silencing effect was investigated in the successive first to fourth generations [42].

2.3. Spray-Induced Gene Silencing

Although transgenes are convenient, they are not required for ectopic gene silencing activation in pathogens or pests. According to some research, eukaryotic pests and pathogens, including fungi and nematodes, are able to take up RNAs from the environment [104,105,106]. This phenomenon was defined as ‘Environmental RNAi’, in which the transferred RNAs complemented to the sequence of target genes in the organism can induce highly effective target gene silencing [104,107]. These studies prompted the development of spray-induced gene silencing (SIGS). In spray-induced gene silencing, dsRNAs or sRNAs that target pathogen or pest genes are sprayed directly onto plants. Then, these RNAs move into the pest or pathogen cells and silence target genes [106,108].
The first evidence of the exogenous application of dsRNA for pest control was in citrus and grapevine trees, in which dsRNA targeting the arginine kinase gene was used to control psyllids and sharpshooter pests [109]. Fusarium graminearum development in barley leaves was suppressed by spraying dsRNA to target the fungal cytochrome P450, establishing the feasibility of spray-induced gene silencing (SIGS) [110]. Moreover, the potential non-transgenic, spray-based exogenous dsRNA or sRNA (SIGS) application has been widely used to decrease disease in crop plants [111,112,113,114].

2.4. Spray-Induced Gene Silencing Based Aphid Control

The delivery of siRNA and dsRNA via nanoparticle carriers is a novel strategy that has been successfully applied in some insect systems [115,116,117]. The majority of SIGS-based studies employed nanocarrier delivery systems for aphid control (Table 1).
tor is a carotene dehydrogenase gene that plays an important role in pigmentation in A. pisum. The branched-chain amino acid transaminase (bcat) gene is important in branched-chain amino acid metabolism in aphids. An aerosolized siRNA-nanoparticle delivery strategy induced a modest tor gene knockdown in A. pisum and a bcat gene knockdown in Aphis glycines as well as the associated phenotype. These results indicated that the aerosolized siRNA-nanoparticle method was an effective RNAi delivery system [44].
According to previous studies, Yan et al. [45] selected the soluble trehalase (TREH), V-type proton ATPase subunit D (ATPD), V-type proton ATPase subunit E (ATPE), and chitin synthase 1 (CHS1) genes as RNAi target genes to test the silencing effect in A. glycines [66,115,118,119]. This study indicated that A. glycines exhibited higher mortality when it fed on soybean seedlings sprayed with a dsATPD + dsCHS1 nanoparticle formulation. They also demonstrated that a water-soluble cationic dendrimer (nanocarrier) was an efficient gene carrier [45].
Biedenkopf et al. [46] reported that the application of dsRNA to detached barley leaves resulted in the effective SIGS of the sheath protein (Shp) gene in grain aphids. Systemic RNAi was also observed in Hordeum vulgare after a spray treatment in which sprayed dsRNA moved from barley leaves to stems and root tissues. This research contributed significantly to understanding the mechanism of RNA spray technology, especially for SIGS. However, another study in barley suggested that grain aphids fed barley seedlings sprayed with naked SaMIF-dsRNAs did not affect the survival of nymphs, which indicated that aphids were unable to absorb dsRNA from these plants [47]. A recently published paper reported that the SIGS-based nanocarrier-mediated dsRNA delivery system effectively silenced the putative salivary effector Sg2204 in Schizaphis graminum and its homologs from four other aphid species. Aphids with silenced Sg2204 exhibited a stronger defense response, and the treatment induced a negative impact on aphid survival, fecundity, and feeding behavior [48].

2.5. Other Delivery-Method-Mediated Gene Silencing for Aphid Control

Microinjection is an efficient and widely used research method for delivering dsRNAs. The first evidence of successful dsRNA microinjection was applied to silence the frizzled and frizzled 2 genes in Drosophila melanogaster embryos by injecting their corresponding dsRNAs [120]. Since then, microinjection has become a potential method for delivering dsRNA into various insect species. This method was reported to apply in many aphid species, namely A. gossypii, A. pisum, M. persicae, and S. avenae (Table 1). The injection of siRNA-C002 into pea aphids decreased the transcription level of C002 [63]. Injections of dsRNAs of different aphid genes that play important roles in aphid sheath formation (SHP) [73], cuticular waterproofing (CYP4G51) [75], (E)-b-farnesene (EβF) reception (ApisOR5, ApisOBP3, and ApisOBP7) [76], chitin biosynthesis (CHS) [80], molting (ApCCAP and ApCCAPR) [62], flight musculature formation, and wing extension (flightin) [85] induced effective target gene silencing.
Feeding was another basic delivery method for aphids because of its less laborious and easier operation. Aphids fed a diet containing synthetic dsRNA were more appliable for target gene knockdown. It was first reported that feeding on E. coli bacteria expressing dsRNA in C. elegans conferred silencing effects on the nematode larvae [121]. In Aphis citricidus, RNAi was performed by feeding dsRNAs of target genes with citrus leaf through stem dipping. Acetylcholinesterase (AChE) is an important gene targeted by insecticides based on organophosphates and carbamates. The silencing of two aphid AChE genes, Tcace1 and Tcace2, increased susceptibility to malathion and carbaryl insecticides. Furthermore, Tcace1 silencing resulted in higher aphid mortality than Tcace2 silencing, which indicated that TcAChE1 was essential for A. citricidus postsynaptic neurotransmission [50]. A knockdown of Vitellogenin (Vg) and its receptor (VgR) had a negative impact on embryonic and postembryonic development, which led to nymph–adult transition delay, a longer pre-reproductive period, and a shorter reproductive period [51]. Cuticle protein is a primary target in insect development and molting. The silencing of the cuticle protein 19 (CP19) gene in A. citricidus led to aphid mortality [52]. Similarly, aphids fed dsRNA of a Gram-negative binding protein gene (AcGNBP1) caused target gene silencing and high mortality [53]. The same delivery strategy was applied in A. pisum. Different dsRNAs were fed with bean leaves through stem dipping. The silencing of the CP19 gene in pea aphids also led to high mortality [52]. Parental silencing of the carotenoid desaturase gene (CdeB) reduced the intensity of the body color in vivo in the treated aphids and subsequent generations and negatively affected aphid performance [82]. The silencing of ApGNBP1 but not ApGNBP2 in A. pisum decreased immune-related phenoloxidase activity [53]. Feeding on Brassica leaves inserted into a solution containing MpCP19 and MpGNBP1 dsRNAs also induced effective target gene silencing [52,53]. With the aim of decreasing insecticide use and eliminating pesticide-resistant evolved populations, RNAi has also been used to increase the susceptibility of aphids to insecticides. A study reported that RpAce1 suppression increased the susceptibility to pirimicarb and malathion in Rhopalosiphum padi. Silencing SaAce1 also increased S. avenae susceptibility to pirimicarb [94].
It has also been demonstrated that mechanical inoculation can help deliver dsRNA and induce RNAi by spreading dsRNA with soft sterile brushes and gentle rubbing inoculation [122,123]. The molecules were rapidly absorbed by tomato plants and were ingested by peach aphids (M. persicae) when the tomato leaves were gently rubbed with dsRNA solution [88]. With the use of a nanocarrier and detergent, a novel dsRNA formulation was exploited, which can quickly penetrate through the body wall of A. glycines and effectively suppress gene expression. This suggests that transdermal dsRNA delivery could be developed as a potential SIGS-based aphid control strategy. Hemocytin (Hem) is an important factor in the hemocytes and fat bodies of insects, which might regulate aphid population density. When spreading a dsRNA-HEM nanocarrier/detergent formulation on A. glycines, the expression level of hemocytin was efficiently silenced, which impaired the survival and fecundity of aphids and suppressed aphid population growth [54]. Another study also investigated the RNAi efficacy of the ATPD gene in woolly apple aphids (Eriosoma lanigerum) via a nanocarrier-mediated transdermal dsRNA delivery system. Their results suggested that the interference efficiency was greatly increased using nanocarriers and induced high aphid mortality [86]. The nanocarrier-delivered RNAi method was also used to silence the flightin, vestigial (vg), and Ultrabithorax (Ubx) genes, which suppressed the wing development in M. persicae [85,91]. In Sitobion miscanthi, the putative salivary effector Sm9723 was effectively silenced via a nanocarrier-mediated transdermal dsRNA delivery system. The fecundity and survival of S. miscanthi dramatically decreased after Sm9723 silencing, and the aphid feeding behavior was also impaired [103].

3. Challenges for Enhancing RNA Interference Efficiency

3.1. Target Gene Selection

The selection of the target gene is essential to the successful application of RNAi-based insect control. RNAi efficiency varies considerably among different insect species for the same transcripts [26,124]. The efficiency can vary in the same species with different transcripts, genotypes, and tissues, even among the same transcript from different areas [26,125,126,127,128,129]. The ideal RNAi gene target must be essential for insect survival and highly expressed and should not have functional redundancy [130,131]. Therefore, potential target genes should be thoroughly investigated for the capacity to suppress specific transcripts and the ability to cause mortality to enhance the efficiency of RNAi-based pest control.
RNAi targets are initially selected based on the discovery of key genes in other organisms or by cDNA library screening. Numerous studies have indicated that genome-wide screens of high-sensitivity target genes are effective in RNAi. Other high-throughput approaches, such as RNA-seq and digital gene expression tag profiles (DGE-tag), were used in the Asian corn borer (ACB; Ostrinia furnacalis) to identify potential RNAi targets [132]. The expression profiling and transcriptome reconstruction of an increasing number of insects have been made possible by second-generation sequencing. High-throughput screens such as feeding assays [66] and the topical application of dsRNA [44,54,132] are also powerful tools to identify potential RNAi targets. With the available databases growing, tissue-specific and developmental-stage-specific expression profiles of insects may narrow down candidate pools for target gene selection. After identifying candidate genes, screening for dsRNA-induced mortality is necessary to evaluate the capacity of specific dsRNAs to induce the desirable phenotype. The potential for the candidate dsRNA sequences to cause mortality at various stages of life can be examined in further experiments. Targeting multiple genes, dsRNA concatemerization, or using different dsRNA structures can all be performed to improve the efficiency of RNAi [133,134,135,136].

3.2. Length of dsRNA

In some insect species, the uptake and silencing efficiency of RNAi are determined by the length of the expressed dsRNA. Different insect species require different minimum lengths of dsRNA to achieve maximal RNAi silencing [137]. In Tribolium castaneum, an analysis revealed that the dsRNA length had a significant impact on the effectiveness of the RNAi response. Longer dsRNA is proving to be more effective at suppressing gene expression. The desired interference requires a minimum length of 70 nucleotides [138]. The length of dsRNA sequences between 139 bp and 773 bp was used in the majority of the aphid feeding experiments to obtain successful RNAi (Table 1).
As we described above, siRNA injections were able to suppress the target gene (C002) expression in pea aphids, which dramatically reduced aphid survival [63]. In grain aphids, RNAi targeting the sheath protein (SHP) gene with transgenic barley plants expressing a 491 bp shp-dsRNA strongly inhibited the feeding and reproductive behavior of grain aphids and negatively impacted their survival [31]. Gq proteins play critical roles in insect cellular signal transduction. The downregulation of the Gqα gene with a 540 bp fragment of dsRNA resulted in decreases in the fecundity and molting rate [41]. A 198 bp dsSaZFP fragment could induce target gene silencing in grain aphids when feeding on transgenic wheat plants, resulting in decreased reproduction and survival rates [42].
Therefore, both short and long dsRNAs effectively induce gene silencing, depending on the target pests and genes. Longer dsRNAs may increase the possibility of off-target effects on beneficial organisms due to the generation of potentially large siRNA pools. Accordingly, RNAi efficiency will be improved by selecting the optimal lengths of target-specific RNAi targets combined with effective siRNA analysis [9].

3.3. Delivery of dsRNA

Various dsRNA delivery methods, including microinjection, feeding, soaking, HIGS mediated by transgenic plants, and SIGS mediated by spraying, have been applied in pest management. As we discussed before, microinjection and feeding are the two basic delivery methods. The soaking delivery method was usually applied in insect cell lines via adding dsRNA directly into the cell culture medium [139,140], and some studies have investigated topically applied dsRNA/siRNA formulations penetrating into the insect cuticles to induce mortality [13,141,142,143,144,145]. Transgenic plants expressing dsRNA or siRNA have lots of advantages for pest control [146]. The SIGS-mediated delivery method does not require plant genetic engineering. dsRNAs/siRNAs are applied topically to the plant surface via spraying in this silencing type [106].
To improve dsRNA delivery efficiency, various new technologies have been exploited, such as cationic-liposome-assisted and nanoparticle-enabled methods. The application of RNAi in conjunction with nanotechnology may develop as a more environmentally friendly approach to pest control. In the first investigation of nanoparticle-mediated dsRNA delivery, chitosan was used to silence the chitin synthase genes in Anopheles gambiae, and the RNAi effectiveness was found to be enhanced [115]. Short interfering RNA (siRNA)–nanoparticle complexes, peptide nanomaterial branched amphiphilic peptide capsules (BAPCs), and nanocarrier-based transdermal dsRNA delivery systems were demonstrated to be successful for aphid RNAi, which could efficiently silence gene expression [44,45,54,147].

3.4. The Stability of dsRNA

RNAi stability and efficiency vary drastically depending on the length and concentration of the dsRNA, the delivery method and technique, plant-organ-specific processes, insect life stage, target gene selection, and adverse environmental conditions [145,148,149]. Environmental microorganisms can degrade dsRNA before it is consumed by pathogens or pests. Nucleases in pest saliva, the gut lumen, and hemolymph may also rapidly degrade dsRNA [19,127,150,151,152,153].
The stability of dsRNA in the insect gut is critical for a successful RNAi response, and increased nuclease expression can result in dsRNA degradation and subsequent RNAi failure [154]. The activity of gut nucleases can be impacted by the high or low pH present in the gut lumena of particular pests, which can directly or indirectly decrease dsRNA stability [155]. Some strategies have already been exploited to improve the stability of dsRNA. For example, the nanoparticle-mediated dsRNA delivery system was demonstrated to be efficient in increasing dsRNA stability and efficacy, and has been applied to improve the penetration and persistence of dsRNA into plants or insects [9,149,156,157].

3.5. Nontarget and Off-Target RNAi Effects

Silencing nontarget genes in the same or nontarget organisms has resulted in off-target effects [158,159,160]. To improve RNAi efficiency and minimize off-target effects, species-specific or tissue-specific dsRNA could be selected. A study reported that the silencing of V-ATPase genes in A. pisum, D. melanogaster, M. sexta, and T. castaneum was observed without affecting nontarget species using species-specific dsRNA [66].
To design efficient and potent RNAi targets, various web-based computational design approaches have been exploited to minimize potential off-target effects. For example, pssRNAit was developed to design specific and effective siRNAs [161]. Further assessments were applied to selected sequences using software, for example, ERNAi [162], dsCheck [163], and basic local alignment search tool (BLAST) [164] analysis against the transcriptomic datasets of human and beneficial insects [9].

4. Conclusions and Perspectives

During the past few years, RNAi has developed as a promising, valuable, and effective technique for functional genomic studies. Various RNAi-based approaches have been applied in crop protection for species-specific and ecofriendly pest management. In this review, we summarize the present studies on numerous strategies exploited against different aphid species.
Growing evidence suggests that HIGS-based and SIGS-based crop protection against pests is effective. Transgenic plants appear to be a more beneficial approach to enhancing RNAi effects. Nevertheless, a lack of transformation technology in several crop species has restricted the widespread application of HIGS. Furthermore, they are still regarded as genetically modified (GM) products in many countries, requiring a thorough assessment of the plants before being licensed. The development of transplastomic technology was also restricted by the extensive regulatory process. Global applications of HIGS are limited by public concern over the biosafety of genetically modified organisms (GMOs) [165,166]. Using optimized target gene and fragment selection strategies, more effective transformation constructs, and stable transgenic systems, the major challenges for the HIGS strategy will be overcome [167]. SIGS, in comparison to HIGS, does not produce GMOs. However, it has become clear that the instability of naked dsRNA is a significant limitation of SIGS, resulting in a relatively short period of protection. In order to address this issue and improve the insecticidal activity of non-transformative RNAi products, SIGS-based dsRNAs affiliated with different types of nanoparticles would be an efficient technique [168,169,170,171,172,173,174]. These prospective strategies may decrease the cost and improve the dependability of the present delivery techniques. They may also create new opportunities to study the roles of important genes. Another consideration for RNAi application is to exclude potential off-target effects and effects on nontarget organisms. To support the biosafety claims of RNAi applications, a combination of bioinformatics and ecological bioassays using selected target species is essential.
With the development of new technology, clustered regularly interspaced short palindromic repeat/CRISPR-associated endonuclease Cas9 (CRISPR/Cas9)-based genome editing had been reported in Spodoptera exigua [175], Helicoverpa armigera [176,177,178], S. litura [179,180], and Nilaparvata lugens [181,182]. However, many of these studies have focused on insect genomic functions. Further study is needed to exploit genome editing as a viable strategy to create resistant varieties against numerous insect pests and enhance pest resistance in crops [183].
Overall, by obtaining a deep understanding of the RNAi machinery and the development of various dsRNA delivery strategies, RNAi will be more effectively used in aphid control for crop protection.

Author Contributions

J.Z., H.L., X.Z., J.T. and A.S. wrote the manuscript. F.F. and L.X. revised the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by the Shennong Laboratory, Zhengzhou Henan 450002, China (SN01-2022-01); the Innovation Program of the Chinese Academy of Agricultural Sciences (ZDXM03 and S2021ZD03); and the National Engineering Laboratory of Crop Molecular Breeding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Acknowledgments

J.Z. was supported by the China Scholarship Council (No. 202003250096) and the GSCAAS-ULg Joint PhD Program.

Conflicts of Interest

The authors declare no conflict of interest.

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Agriculture 12 02108 g001 550
Figure 1. Schematic of RNA interference (RNAi) delivery strategies, the RNAi mechanism in aphid cells, and challenges affecting RNAi efficiency in aphids. HIGS: host-induced gene silencing, SIGS: spray-induced gene silencing, dsRNA: double-stranded RNA, Dicer: Dice-like, siRNA: short interfering RNA, Ago: Argonaute, RISC: RNA-induced silencing complex.
Figure 1. Schematic of RNA interference (RNAi) delivery strategies, the RNAi mechanism in aphid cells, and challenges affecting RNAi efficiency in aphids. HIGS: host-induced gene silencing, SIGS: spray-induced gene silencing, dsRNA: double-stranded RNA, Dicer: Dice-like, siRNA: short interfering RNA, Ago: Argonaute, RISC: RNA-induced silencing complex.
Agriculture 12 02108 g001
Table
Table 1. Summary of RNAi application for aphid control.
Table 1. Summary of RNAi application for aphid control.
Types of Gene SilencingAphid SpeciesPlant SpeciesDelivery StrategyTarget GenesMoleculeSizeMain EffectsReference
HIGSMyzus persicaeNicotiana benthamiana and A. thalianaTransgenic N. benthamiana and A. thalianaMpC002, Rack1dsRNA710 bp, 309 bpKnockdown of target genes.[17]
Myzus persicaeN. benthamiana and A. thalianaTransgenic N. benthamiana and A. thalianaMpC002, MpPIntO1 (Mp1), MpPIntO2 (Mp2)dsRNA710 bp, 263 bp, 254 bpSilencing of MpC002 and MpPIntO2 reduced nymph production.[33]
Myzus persicaeN. tabacum, A. thaliana, and N. benthamianaTransgenic N. tabacum, A. thaliana, and N. benthamianaMp55dsRNA>900 bpReduced aphid reproduction.[34]
Myzus persicaeA. thalianaTransgenic A. thalianaRack1, MpC002, MpPIntO2 (Mp2)dsRNA309 bp, 710 bp, 254 bpReduced aphid reproduction.[35]
Myzus persicaeA. thalianaTransgenic A. thalianaCuticular protein MyCPdsRNA327 bpAttenuation of fecundity in aphids.[43]
Myzus persicaeTomatoAgrobacterium-mediated transformation and transgenic tomatoAcetylcholinesterase 1 (Ace 1)dsRNA571 bpSilenced the target gene (Ace 1) and inhibited fecundity.[36]
Myzus persicaeTobaccoInjection and feeding on transgenic tobaccoCysteine protease Cathepsin B3 (CathB3)dsRNA230 bpImproved the performance of non-tobacco-adapted lineages on tobacco.[37]
Myzus persicaeTobaccoPlastid-mediated RNA interference and transgenic tobaccoMpDhc64CdsRNA269 bpReduced insect survival, impaired fecundity, and decreased weight of survivors.[38]
HIGSSitobion avenaeWheatParticle bombardment method and transgenic wheatCarboxylesterase
(CbE E4)		dsRNA350 bpSuppressed CbE E4 expression impaired S. avenae larval tolerance of phoxim insecticides.[18]
Sitobion avenaeWheatParticle bombardment method and transgenic wheatLipase maturation factor 2-like gene, lmf2-likedsRNA543 bpReductions in molting number, survival, and reproduction.[39]
Sitobion avenaeWheatParticle bombardment method and transgenic wheatChitin synthase 1 (CHS1)dsRNA550 bpDecreased CHS1 expression level and reduced total and molting aphid numbers.[40]
Sitobion avenaeWheatParticle bombardment method and transgenic wheatGq protein alpha subunit (Gqα)dsRNA517 bpReduced reproduction and molting in aphids.[41]
Sitobion avenaeWheatParticle bombardment method and transgenic wheatZinc finger protein (SaZFP)dsRNA198 bpHigh mortality and decreased fecundity.[42]
SIGSAphis glycines Aerosolized
siRNA-nanoparticle delivery method		Carotene dehydrogenase (tor), branched-chain amino acid transaminase (bcat)siRNA25 ntKnockdown of target genes.[44]
Aphis glycines Nanocarrier-based dsRNA delivery systemTREH, ATPD, ATPE, and CHS1dsRNA431 bp, 504 bp, 536 bp, 429 bpSilenced target gene expression and led to high mortality.[45]
Acyrthosiphon pisum Aerosolized
siRNA-nanoparticle delivery method		Carotene dehydrogenase (tor), branched-chain amino acid transaminase (bcat)siRNA25 ntKnockdown of target genes.[44]
Sitobion avenaeBarleySprayingStructural sheath protein (SHP)dsRNA491 bpReduced shp expression level.[46]
Sitobion avenaeBarleySpraying and feedingMacrophage migration inhibitory factors, SaMIF1, SaMIF2, and SaMIF3dsRNA223 bp, 323 bp, 212 bpFeeding on artificial diet led to high mortality rates; feeding from barley seedlings sprayed with naked SaMIF-dsRNAs did not alter nymph survival.[47]
Schizaphis graminum Aerosolized
siRNA-nanoparticle delivery method		Carotene dehydrogenase (tor) and branched-chain amino acid transaminase (bcat)siRNA25 ntKnockdown of target genes.[44]
SIGSSchizaphis graminumWheatNanocarrier-mediated transdermal dsRNA delivery systemSg2204dsRNA/Induced a stronger wheat defense response and resulted in negative impacts on aphid feeding behavior, survival, and fecundity.[48]
Other delivery methodAphis citricidus Feeding and citrus stem dippingInsulin receptor genes AcInR1 and AcInR2dsRNA511 bp, 609 bpDevelopmental defects and co-silencing of AcInR1 and AcInR2 resulted in high mortality.[49]
Aphis citricidus Feeding and citrus stem dippingAcetylcholinesterase, TcAChE1, and TcAChE2dsRNA435 bp, 421 bpHigh mortality and increased the susceptibility of A. citricidus to malathion and carbaryl.[50]
Aphis citricidus Feeding and citrus stem dippingVitellogenin (AcVg), Vitellogenin receptor (AcVgR)dsRNA557 bp, 577 bpSlower embryonic development and fewer newborn nymphs.[51]
Aphis citricidus Feeding and citrus stem dippingAcCP19dsRNA183 bpInduced target gene silencing and high mortality.[52]
Aphis citricidus Feeding and citrus stem dippingAcGNBP1dsRNA431 bpDecreased the activity of immune-related phenoloxidase.[53]
Aphis glycines Topical application, nanocarrier, and
detergent-mediated transdermal delivery system		Hemocytin, HemdsRNA555 bpReduced the target gene expression and aphid population density.[54]
Aphis gossypii FeedingCarboxylesterase CarEdsRNA686 bpDecreased resistance to organophosphorus insecticides.[55]
Aphis gossypii FeedingCytochrome P450 monooxygenase gene CYP6A2dsRNA773 bpIncreased sensitivity to spirotetramat and alpha-cypermethrin.[56]
Aphis gossypii FeedingOdorant-binding proteins AgOBP2dsRNA434 bpInterfered with the odorant perception of aphids.[57]
Aphis gossypii FeedingCYP6CY14dsRNA459 bpIncreased the resistant aphid’s susceptibility to thiamethoxam.[58]
Other delivery methodAphis gossypii FeedingCYP380C6dsRNA436 bpIncreased the sensitivity of the resistant adults and nymphs to spirotetramat.[59]
Aphis gossypii FeedingdsCYP6DC1, dsCYP6CY14, and dsCYP6CZ1dsRNA494 bp, 499 bp, 499 bpIncreased the Ace-R strain’s sensitivity to acetamiprid.[60]
Aphis gossypii FeedingEcdysone receptor (EcR)dsRNA486 bpIncreased mortality rates and decreased longevity and fecundity.[61]
Aphis gossypii InjectionCrustacean cardioactive peptide (ApCCAP), crustacean cardioactive peptide receptor (ApCCAPR)dsRNA339 bp, 519 bpDevelopmental failure during nymph–adult ecdysis.[62]
Acyrthosiphon Pisum InjectionC002siRNA21-23 ntDecreased C002 transcript level.[63]
Acyrthosiphon Pisum InjectionCalreticulin, cathepsin-LdsRNA434 bp, 353 bpInduced target gene silencing.[64]
Acyrthosiphon Pisum FeedingApAQP1dsRNA451 bpKnocked down the ApAQP1 expression level, resulting in elevated hemolymph osmotic pressure.[65]
Acyrthosiphon Pisum injectionvATPasedsRNA185 bpInduced high levels of mortality.[66]
Acyrthosiphon Pisum FeedingHunchbackdsRNA524 bp, 497 bpReduced Aphb transcripts and increased insect lethality.[67]
Acyrthosiphon pisum Injection and feedingEnzyme Cathepsin-LdsRNA357 bpInduced lethal effects.[68]
Acyrthosiphon pisum InjectionACYPI39568dsRNA246 bpReduced ACYPI39568 expression level but did not affect the survival rate.[69]
Acyrthosiphon Pisum InjectionAngiotensin-converting enzymes ACE1, ACE2dsRNA313 bp, 468 bpKnockdown of ACE1 and ACE2 caused a higher mortality rate.[70]
Acyrthosiphon Pisum InjectionApMIF1dsRNA213 bpDisturbed their ability to feed from phloem sap.[71]
Acyrthosiphon Pisum InjectionArmetdsRNA286 bpDisturbed feeding behavior and led to a shortened life span.[72]
Acyrthosiphon Pisum InjectionStructural sheath protein (SHP)dsRNA491 bpDisrupted sheath formation, prevented efficient long-term feeding from sieve tubes, and had a silencing effect on reproduction but not survival.[73]
Other delivery methodAcyrthosiphon Pisum InjectionPeroxiredoxins, ApPrx1dsRNA206 bpDecreased survival rate.[74]
Acyrthosiphon Pisum Injection and ingestionCytochrome P450 gene, CYP4G51dsRNA310 bp, 325 bpReduced CYP4G51 expression, caused reductions in internal and external long-chain hydrocarbons (HCs), and increased mortality.[75]
Acyrthosiphon Pisum InjectionOdorant receptors, ApisOR5,
odorant-binding proteins, ApisOBP3, and ApisOBP7		dsRNA/The repellent behavior of A. pisum to EBF disappeared.[76]
Acyrthosiphon Pisum FeedingCuticular protein,
Stylin-01,
Stylin-02		siRNA19 bpSilencing stylin-01 decreased the efficiency of cauliflower mosaic virus transmission by M. persicae.[77]
Acyrthosiphon Pisum InjectionNeuropeptide F (NPF), NPF receptor (NPFR)dsRNA232 bp, 354 bpReduced aphid food intake and indicated a lower appetite for food after NPF silencing.[78]
Acyrthosiphon Pisum FeedingamiD, ldcA1dsRNA311 bp, 353 bpReduction in Buchnera abundance and activity was accompanied by depressed aphid growth rates.[19]
Acyrthosiphon Pisum InjectionGap gene HunchbackdsRNA448 bpKnockdown of target gene.[79]
Acyrthosiphon Pisum Injection and ingestionChitin synthase, CHSdsRNA364 bpInduced mortality and development deformity.[80]
Acyrthosiphon Pisum InjectionApHRCdsRNA263 bpSerratia-infected aphids displayed shorter phloem-feeding durations and caused Ca2+ elevation and ROS accumulation in plants.[81]
Acyrthosiphon pisum Feeding and bean stem dippingCuticle protein gene, ApCP19dsRNA216 bpInduced target gene silencing and high mortality.[52]
Other delivery methodAcyrthosiphon pisum Feeding and bean stem dippingCarotenoid desaturase, CdeBdsRNA431 bpReduced aphid performance and altered the age structure of the population.[82]
Acyrthosiphon pisum Feeding and bean stem dippingGram-negative binding proteins, ApGNBP1, ApGNBP2dsRNA550 bp,
518 bp		Decreased the activity of immune-related phenoloxidase.[53]
Acyrthosiphon Pisum InjectionCCHamide-2 receptor (CCHa2-R)dsRNA478 bpReduced CCHa2-R expression, food intake in adult aphids, and reproduction but not survival.[83]
Acyrthosiphon Pisum InjectionFatty acid synthase 1 (FASN1) and diacylglycerol-o-acyltransferase 2 (DGAT2)dsRNA609 bp, 388 bpProlonged the nymphal growth period and decreased the aphid body weight.[84]
Acyrthosiphon pisum Injection and nanocarrier delivery flightindsRNA374 bpMalformed wings, deformed dorsal longitudinal muscle (DLM) shapes, and wider and looser dorsoventral flight muscles (DVMs) were observed.[85]
Eriosoma lanigerum Hausmann Topical application and nanocarrier-mediated transdermal dsRNA delivery systemV-ATPase subunit D (ATPD)dsRNA/Induced target gene silencing and led to high mortality.[86]
Myzus nicotianae FeedingTRV-ALY, TRV-EphdsRNA182 bp, 249 bpInhibition of target genes.[87]
Myzus persicae InjectionMpMIF1dsRNA 205 bpDisturbed their ability to feed from phloem sap.[71]
Myzus persicae Foliar applicationZYMV HC-ProdsRNA588 bpInsects successfully took up dsRNA; the dsRNA was processed into siRNA by the insect RNAi machinery.[88]
Myzus persicae FeedingCuticular protein,
Stylin-01,
Stylin-02		siRNA19 bpSilencing stylin-01 decreased the efficiency of cauliflower mosaic virus transmission by Myzus persicae.[77]
Other delivery methodMyzus persicae FeedingVoltage-gated sodium channel MpNavdsRNA289 bpInduced high mortality and lower fecundity and longevity.[89]
Myzus persicae Feeding and Brassica stem dippingMpCP19dsRNA139 bpInduced target gene silencing and high mortality.[52]
Myzus persicae Feeding and Brassica stem dippingMpGNBP1dsRNA450 bpDecreased the activity of immune-related phenoloxidase.[53]
Myzus persicae FeedingMp58, OBP2dsRNA423 bp, 428 bpInduced high mortality.[90]
Myzus persicae Topical and root applications and nanocarrier-mediated delivery systemVestigial (vg), Ultrabithorax (Ubx)dsRNA489 bp, 359 bpDownregulated target genes and caused wing aberration.[91]
Myzus persicae InjectionATP-binding cassette transporter gene (ABCG4), DnaJ homolog subfamily C member 1 (DnaJC1) dsRNA~400 bpIncreased mortality rate.[92]
Megoura viciae InjectionTyrosine hydroxylase MV-THdsRNA400 bpReduced the L-DOPA level in aphids and a slight decrease in exuvia tanning.[93]
Rhopalosiphum padi InjectionAcetylcholinesterase gene RpAce1 dsRNA383 bpIncreased susceptibilities to pirimicarb and malathion in R. padi and reduced fecundity.[94]
Sitobion avenae FeedingCatalase CATdsRNA471 bpReduced survival rate and ecdysis index.[95]
Sitobion avenae FeedingUnigenes DSR8, DSR32, DSR33, DSR48dsRNA162 bp, 411 bp, 439 bp, 397 bpDownregulation of target genes and aphid mortality.[96]
Sitobion avenae InjectionAcetylcholinesterase gene SaAce1dsRNA400 bpIncreased susceptibility to pirimicarb in S. avenae and reduced fecundity.[94]
Sitobion avenae FeedingEcdysone receptor (SaEcR), ultraspiracle protein (SaUSP)dsRNA469 bp, 411 bpSignificantly decreased the survival of aphids.[97]
Other delivery methodSitobion avenae FeedingLaccase 1, SaLac 1dsRNA613 bpInhibited the transcript levels of SaLac 1 and decreased the survival rate.[98]
Sitobion avenae FeedingOdorant-binding protein (SaveOBP9)dsRNA501 bpReduced SaveOBP9 expression and induced a nonsignificant response in S. avenae to tetradecane, octanal, decanal, and hexadecane.[99]
Sitobion avenae FeedingOdorant-binding protein (SaveOBP10)dsRNA432 bpAphids exhibited nonattraction towards β-caryophyllene and a nonsignificant behavioral response to pentadecane, butylated hydroxytoluene, and tetradecane.[100]
Schizaphis graminum FeedingSgC002siRNA476 bpFeeding on artificial diet for 3 days followed by transfer to aphid-susceptible wheat suppressed SgC002 expression and led to lethality.[101]
Schizaphis graminum FeedingMRA, GAT, TLPdsRNA376 bp, 433 bp, 422 bpIncreased susceptibility to imidacloprid.[102]
Sitobion miscanthi Topical application and nanocarrier-mediated transdermal dsRNA delivery systemSm9723dsRNA/Decreased fecundity and survival and negatively affected the feeding behavior. [103]
Note: HIGS: host-induced gene silencing. SIGS: spray-induced gene silencing.
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MDPI and ACS Style

Zhang, J.; Li, H.; Zhong, X.; Tian, J.; Segers, A.; Xia, L.; Francis, F. RNA-Interference-Mediated Aphid Control in Crop Plants: A Review. Agriculture 2022, 12, 2108. https://doi.org/10.3390/agriculture12122108

AMA Style

Zhang J, Li H, Zhong X, Tian J, Segers A, Xia L, Francis F. RNA-Interference-Mediated Aphid Control in Crop Plants: A Review. Agriculture. 2022; 12(12):2108. https://doi.org/10.3390/agriculture12122108

Chicago/Turabian Style

Zhang, Jiahui, Huiyuan Li, Xue Zhong, Jinfu Tian, Arnaud Segers, Lanqin Xia, and Frédéric Francis. 2022. "RNA-Interference-Mediated Aphid Control in Crop Plants: A Review" Agriculture 12, no. 12: 2108. https://doi.org/10.3390/agriculture12122108

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