Introduction

As obligate intracellular parasites, most animal and plant viruses lack autonomous motility and usually rely on various insect vectors for transmission. For instance, dengue virus (DENV) and Zika virus (ZIKV) induce N-acetyl-L-tyrosine accumulation in mosquito heads, activating the tyrosine–dopamine pathway and thereby elevating dopamine levels to enhance locomotion and blood-feeding propensity. They also disrupt the circadian rhythm of Aath, which encodes tyrosine hydroxylase, sustaining dopamine biosynthesis [1]. Similar to animal viruses, more than 70% of plant viruses are transmitted by various insect vectors [2]. Through long-term coevolution, plant viruses have selected the most efficient vector species, a process that appears almost active and purposeful [3]. Most plant viruses have evolved to exploit the feeding behavior and morphology of their insect vectors as an efficient transmission strategy, effectively “hitchhiking” insect vectors to enhance dispersal and increase epidemic potential [4]. Plant viruses are transmitted by insect vectors via nonpersistent, semi-persistent, and persistent modes, determined by the duration of viral particles within the vectors. Nonpersistent viruses such as cucumber mosaic virus (CMV) and semi-persistent viruses such as cauliflower mosaic virus (CaMV) are transmitted in a noncirculative manner, during which virions bind to specific cuticular structures within the insect stylet or foregut [5–7]. These viruses are acquired and transmitted within a short period. In contrast, persistent transmission occurs in a circulative manner, during which the viruses cross multiple biological barriers by entering the epithelial cells of the insect midgut, traversing the hemolymph, and ultimately reaching the salivary glands [2,7]. Persistent viruses are further classified into two types, propagative viruses which replicate within insect tissues, such as the rhabdovirus barley yellow striate mosaic virus (BYSMV) and the tenuivirus rice stripe virus (RSV) [8–10]; and nonpropagative viruses, which circulate through the vector body without replication, such as the luteovirus barley yellow dwarf virus (BYDV) [4,6,11].

For successful transmission and epidemic spread, plant viruses must not only trigger attraction of appropriate insect vectors to infected plants to maximize viral acquisition, but also enhance vector settling and feeding, as well as subsequently facilitate efficient release and inoculation into new host plants. During these processes, viruses actively manipulate both plant hosts and insect vectors to facilitate their transmission. Increasing evidence indicates that plant viruses indirectly influence vector behavior by reprogramming host phytohormone signaling and secondary metabolism, which can increase vector attraction, improve host suitability, and promote virus acquisition. For instance, in plants infected with Begomovirus, the βC1 protein encoded by the associated β-satellite DNA interacts with the plant transcription factor MYC2 to suppress the expression of terpene synthase genes, thereby reducing plant resistance to the whitefly vector [12]. βC1 also interacts with the vascular-specific transcription factor WRKY20, disrupting its dimerization and suppressing its transcriptional activity. This interaction leads to increased accumulation of aliphatic glucosinolates in nonvascular leaf tissues, accompanied by a reduction in indole glucosinolates within leaf veins [13]. Such metabolic reprogramming inhibits nonvector insects but benefits vector whiteflies, thereby facilitating virus transmission [13]. Similarly, tomato chlorosis virus (ToCV) infection promotes whitefly attraction and feeding by altering host physiology. The viral P9 protein interacts with light-harvesting complex I subunit Lhca4 (Lhca4), causing chlorophyl degradation and increased accumulation of the volatile neophytadiene for whitefly attractiveness. In addition, ToCV suppresses jasmonic acid (JA) signaling, further facilitating vector feeding and virus spread [14]. These mechanisms have been extensively summarized in recent reviews, which demonstrate that viruses indirectly modulate their insect vectors through the manipulation of plant hosts [15–21].

Although it has long been recognized that plant viruses can directly manipulate their insect vectors, the molecular mechanisms underlying these interactions have remained largely obscure [17]. Persistent-propagative plant viruses include negative-sense RNA viruses, such as rhabdoviruses, orthotospoviruses, and tenuiviruses, double-stranded RNA viruses in the family Reoviridae [22], and geminiviruses such as TYLCV, whose replication in insect vectors was previously debated but has recently been supported by experimental evidence [23,24]. Unlike nonpropagative viruses, these viruses can replicate within insect vectors and synthesize viral RNAs and proteins, as they do in plant cells, thereby establishing more active and intimate interactions with their vectors [25]. Recent studies have revealed that several of these viruses can directly modulate vector behavior by altering wing morphology, neural signaling, olfactory perception, and feeding activity [26–28]. In this review, we summarize recent advances in understanding how persistent-propagative plant viruses manipulate insect vectors through both indirect plant-mediated and direct vector-targeted mechanisms, and discuss the implications for plant virus–vector interactions and disease epidemiology.

Plant rhabdoviruses coordinate neurotropic infection–mediated circadian regulation and jasmonate signaling to manipulate vector behavior

Plant rhabdoviruses, members of the subfamily Betarhabdovirinae within the family Rhabdoviridae, currently comprise 253 classified species [29,30], and possess negative-sense RNA genomes that may be nonsegmented, bipartite, or tripartite [31]. Most plant rhabdoviruses replicate in their arthropod vectors and are transmitted in a persistent-propagative manner [32]. Increasing evidence indicates that these viruses establish neurotropic infections in their insect hosts. Early studies showed that maize mosaic virus (MMV) and maize fine streak virus (MFSV) systemically infect the nervous systems of their insect vectors, suggesting that a neurotropic route may help these viruses overcome transmission barriers [33,34]. Subsequent work further demonstrated that the matrix protein of rice yellow stunt virus (RYSV) interacts with axonal microtubules and facilitates the transport of nonenveloped viral structures through the central nervous system, thereby promoting viral dissemination to the salivary glands and efficient transmission [35].

Animal rhabdoviruses such as rabies virus usually cause significant cytopathological damage to host nervous systems [36]. By contrast, plant rhabdoviruses appear to have evolved a more balanced relationship with their insect vectors. Recent research has revealed that the Hikaru genki homolog (NcHig), a neural factor in leafhoppers, binds to the M proteins of both RYSV and rice stripe mosaic virus (RSMV), suppressing viral infection in neural tissues [37]. This interaction likely represents a co-evolutionary adaptation, whereby a stealth strategy maintains vector fitness to promote viral persistence. Thus, neurotropism suggests that plant rhabdoviruses can disseminate through the insect nervous system to reach the salivary glands, providing an alternative to the classical midgut–hemolymph–salivary gland route and resembling neuroinvasive routes used by some animal rhabdoviruses [38].

In addition to facilitating systemic spread within insect vectors, viral infection of the nervous system can also influence vector circadian rhythm and feeding behavior. BYSMV, an economically important cytorhabdovirus, is dependent on the small brown planthopper (SBPH) for transmission and infects more than 26 crop species in the field [39]. The accessory protein P6 of BYSMV interacts with the barley COP9 signalosome subunit 5 (HvCSN5), and thereby suppresses LsCSN5-regulated de-neddylation of Cullin 1 (CUL1). Consequently, inhibition of CUL1-based E3 ligase-mediated degradation of JAZ proteins suppresses jasmonate signaling, thereby enhancing vector attraction and virus acquisition [40]. Consistent with other animal and plant rhabdoviruses, BYSMV has been shown to infect the brains of its insect vectors [8]. Interestingly, BYSMV P6 also interacts with the insect COP9 signalosome subunit 5 (LsCSN5), a homolog of HvCSN5, and suppresses LsCSN5-regulated de-neddylation of Cullin 1 (CUL1). This inhibition prevents CUL1-based E3 ligase-mediated degradation of the circadian clock protein Timeless (TIM), thereby disrupting circadian rhythm and enhancing locomotor activity in infected insects, which increases the probability of viral transmission [28]. Thus, BYSMV uses the same viral effector to target homologous CSN5–CUL1 ubiquitin-system components in both plants and insects, indirectly promoting vector attraction through plant jasmonate signaling and directly enhancing vector transmission activity through disruption of the insect circadian clock (Fig 1).

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Fig 1. BYSMV manipulates insect vector behavior through indirect and direct mechanisms.

In BYSMV-infected barley plants, the viral P6 protein interacts with HvCSN5, a subunit of the COP9 signalosome, thereby stabilizing the jasmonate (JA) signaling repressor JAZ and suppressing JA-mediated anti-insect defense. This plant-mediated effect facilitates feeding by small brown planthoppers (SBPHs) and promotes virus acquisition (indirect manipulation). In viruliferous insects, BYSMV replicates in the central nervous system, where P6 binds to LsCSN5, the insect homolog of HvCSN5, disrupts the rhythmic expression of the circadian clock protein TIM, and enhances vector locomotor activity and feeding behavior (direct manipulation). Together, these processes form a dual-host manipulation strategy that maximizes BYSMV transmission. This figure was created in BioRender, https://BioRender.com/gydn1yh.

https://doi.org/10.1371/journal.ppat.1014368.g001

Similarly, RSMV also exploits the host ubiquitin-system to create favorable ecological conditions for viral persistence and spread. The RSMV P6 protein interacts with the rice heading-related E3 ubiquitin ligase Heading date Associated Factor 1 (HAF1), impairing its function and delaying rice heading. This virus-induced developmental change creates a more favorable environment for leafhopper vectors and viruses to overwinter [41], thereby facilitating long-term virus maintenance and subsequent transmission.

Plant bunyaviruses manipulate stylet permeability, hormone signaling, and wing development to enhance vector-mediated transmission

Beyond plant rhabdoviruses, several other persistent-propagative plant negative-sense RNA viruses are also major threats to crop production, including rice RSV and rice grassy stunt virus (RGSV) in the genus Tenuivirus, and tomato spotted wilt virus (TSWV) in the genus Orthotospovirus, all of which belong to the class Bunyaviricetes [42]. Recent studies, particularly those on RSV, have begun to reveal how these viruses manipulate vector behavior and transmission through both indirect plant-mediated and direct vector-targeted mechanisms.

The stylet of insect vectors serves as both the entry and exit conduit for plant viruses during acquisition and inoculation. It consists of a food canal and a salivary canal, which merge at the tip into a common duct. This architecture is mainly composed of cuticular chitin [43]. Recent studies have revealed that plant chitin-binding lectins (ChtBLs) function as defensive factors against insect feeding. In rice, SBPH feeding strongly induces OsChtBL1, which encodes a protein with four tandem chitin-binding domains. OsChtBL1 binds to stylet chitin and forms large aggregates, producing dense mesh-like deposits within the food canal that act as a physical barrier to virus acquisition and inoculation. To overcome the feeding barrier, RSV has evolved a specific suppression mechanism through inhibiting the accumulation of plant OsChtBL1. Mechanistically, the RSV P2 protein links OsChtBL1 to the host RING-type E3 ubiquitin ligase OsRING18, thereby promoting the ubiquitination and subsequent proteasome-dependent degradation of OsChtBL1. This degradation decreases OsChtBL1 deposition within the stylet food canal, ultimately facilitating viral acquisition and transmission [44] (Fig 2A). These findings reveal a plant-mediated strategy by which RSV remodels the vector feeding interface, suggesting that the insect stylet is not only a passive conduit for virus movement but also a critical target in virus–plant–vector interactions.

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Fig 2. Plant bunyavirus RSV manipulates stylet permeability and wing development to enhance vector-mediated transmission.

(A) During feeding, SBPHs induce expression of ChtBL1, which enters the stylet food canal, binds chitin, and forms inclusion bodies that restrict stylet permeability and suppress feeding. Upon RSV infection, the viral P2 protein mediates E3 ubiquitin ligase-dependent degradation of ChtBL1, increasing stylet permeability and facilitating viral transmission (indirect manipulation). (B) RSV replication in SBPHs upregulates the insulin pathway kinase Akt, which in turn induces expression of Encounter, a planthopper-specific protein. An RSV-derived siRNA also acts as a microRNA targeting the 5′untranslated region of Encounter, enhancing its transcription. These two synergistic effects elevate Encounter expression and drive the development of long-winged male morphs that favor viral spread (direct manipulation). This figure was created in BioRender, https://BioRender.com/1txyyud.

https://doi.org/10.1371/journal.ppat.1014368.g002

RSV can also directly manipulate vector morphology to facilitate transmission. Insect wings exhibit typical phenotypic plasticity under different environmental conditions. For instance, brown planthoppers develop either short-winged or long-winged morphs depending on host plant quality, a process controlled by insulin receptor-mediated nutrient-sensing pathways [45]. Virus infection can also alter wing morph determination in insect vectors to facilitate long-distance transmission. RSV specifically induces the development of long-winged morphs in male insects. Mechanistically, an RSV-derived small interfering RNA (vsiRNA) targets the 5′ untranslated region (UTR) of the Encounter gene and improves its expression. Encounter functions as a downstream effector of Akt within the insulin signaling pathway. RSV-induced upregulation of Akt enhances Encounter expression and promotes the development of the long-winged phenotype [26] (Fig 2B).

Modulation of wing development has also been observed in the nonpersistent virus CMV. CMV infection is often associated with satellite RNAs (satRNAs) [46], among which Y-satellite RNA (Y-sat) induces leaf yellowing by producing a vsiRNA that targets magnesium protoporphyrin chelatase subunit I (ChlI), a key enzyme in chlorophyl biosynthesis. Degradation of ChlI mRNA reduces chlorophyl accumulation and enhances the visual attraction of aphid vectors [47,48]. During feeding, Y-sat double-stranded RNA can enter aphids and be processed into siRNAs. One Y-sat-derived 24-nt siRNA mimics aphid miR-9b and competitively binds the mRNA of the wing development-related gene ABCG4, relieving miR-9b-mediated repression and upregulating ABCG4 expression to promote wing formation [48]. This dual “pull–push” strategy enables CMV/Y-sat to attract aphid vectors and promote their dispersal, thereby enhancing transmission efficiency and epidemic spread. Unlike RSV-derived vsiRNAs, which are produced during viral replication within insect vectors, CMV/Y-sat-derived siRNAs originate from plant-derived dsRNAs that are ingested and processed by aphids. Nevertheless, both mechanisms ultimately regulate vector wing development to facilitate virus transmission.

Plant bunyaviruses also target hormone signaling to manipulate vector behavior and transmission ecology. RSV NS2 interacts with OsMYC2 and sequesters it in the cytoplasm, thereby blocking OsMYC2-mediated expression of OsBSMT1 and methyl salicylate (MeSA) biosynthesis, thereby impairing the recruitment of parasitoids that attack insect vectors. By reducing natural enemy attraction, RSV promotes vector persistence on infected plants and indirectly supports viral transmission from a broader ecological niche perspective [49]. MYC transcription factors are also targeted by TSWV. TSWV NSs protein directly interacts with MYC2 and its close homologs MYC3 and MYC4, thereby suppressing JA-mediated activation of terpene synthase genes. This reduces the biosynthesis of repellent volatile monoterpenes, increasing plant attractiveness to thrips vectors and promoting virus spread [50]. Together, these studies suggest that plant viruses manipulate insect vector behavior through complex, multilayered regulatory networks rather than a single linear pathway, thereby reshaping both vector preference and the broader ecological context of transmission.

Plant reoviruses reshape vector behavior by reprogramming plant volatiles and modulating vector olfaction

Several major rice viral diseases are caused by plant reoviruses, including rice dwarf virus (RDV), rice gall dwarf virus (RGDV), rice black-streaked dwarf virus (RBSDV), and southern rice black-streaked dwarf virus (SRBSDV). These viruses possess 10–12 segmented double-stranded RNA genomes and are transmitted by leafhopper or planthopper vectors in a persistent-propagative manner [51]. RDV, a member of the genus Phytoreovirus, is transmitted by green rice leafhoppers (GRLHs, Nephotettix cincticeps) and manipulates vector behavior by altering rice volatile emissions. RDV-infected plants emit increased levels of specific volatiles, including E-β-caryophyllene and 2-heptanol. E-β-caryophyllene attracts nonviruliferous GRLHs to settle on infected plants, thereby facilitating virus acquisition, but has little effect on viruliferous GRLHs. In contrast, 2-heptanol repels viruliferous GRLHs from infected plants while having little effect on nonviruliferous insects [52]. Similarly, SRBSDV, a member of the genus Fijivirus, is transmitted by the white-backed planthopper (WBPH, Sogatella furcifera). Nonviruliferous WBPHs preferentially settle on SRBSDV-infected rice plants to acquire the virus, whereas viruliferous individuals shift their preference toward healthy plants, thereby promoting viral inoculation and spread [53,54].

In plants, during early infection, the SRBSDV P6 protein localizes to the cytoplasm, where it interacts with ethylene-response repressor OsRTH2 to enhance ethylene signaling and promote SRBSDV replication. At the later stage, P6 translocates into the nucleus and interacts with the key ethylene signaling component OsEIL2, thereby suppressing ethylene signaling and enhancing the attractiveness of infected plants to vectors [55]. Olfactory perception plays a central role in shaping insect development and behavior [56]. In insects, SRBSDV infection shifts vector preference from infected to healthy rice plants for efficient virus transmission. Mechanistically, the viral P8 protein competitively binds to the kinase Pelle, suppressing Toll signaling and consequently increasing viral infection. Moreover, this inhibition prevents phosphorylation and nuclear translocation of DIF, an important transcription factor of Toll signaling, leading to downregulation of the olfactory receptor Or86 and upregulation of the receptor Or127 in the insect olfactory system. Consequently, nonviruliferous planthoppers exhibit enhanced antennal responses to E-β-farnesene by Or86, whereas viruliferous individuals were more sensitive to β-caryophyllene by Or127. By perturbing the Toll-DIF regulatory pathway, SRBSDV reprograms olfactory receptor gene expression and reshapes vector sensory perception, thereby driving a behavioral shift that favors viral acquisition and dissemination [27,57] (Fig 3). Future work should investigate how ethylene signaling interacts with volatile biosynthesis pathways and how the two volatiles E-β-farnesene and β-caryophyllene differ between healthy and virus-infected plants to clarify their roles in mediating vector behavior.

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Fig 3. Plant reovirus SRBSDV reshapes vector behavior by reprogramming plant volatiles and modulating vector olfaction.

In SRBSDV-infected rice plants, the viral P6 protein exhibits stage-specific localization and function. During early infection, P6 localizes to the cytoplasm, where it interacts with the ET negative regulator OsRTH2, thereby activating the ET signaling pathway and promoting viral replication. At later stages, P6 translocates into the nucleus and binds to the ET positive regulator OsEIL2, suppressing ET signaling, which may enhance the production of the volatile E-β-farnesene that may be favored by healthy WBPHs (indirect manipulation). In SRBSDV-infected vectors, the viral P8 protein interacts with Pelle, a key kinase in the Toll pathway, inhibiting the phosphorylation and nuclear translocation of the transcription factor DIF. DIF suppression perturbs odorant receptor gene regulation, leading to the downregulation of Or86 (responsive to E-β-farnesene) and the upregulation of Or127 (β-caryophyllene). Consequently, viruliferous insects display altered olfactory sensitivity, enabling them to distinguish between infected and healthy rice plants and thereby promoting efficient virus transmission (direct manipulation). This figure was created in BioRender, https://BioRender.com/mh8up2p.

https://doi.org/10.1371/journal.ppat.1014368.g003

Although SRBSDV-induced changes in rice volatile profiles have not yet been directly demonstrated, RDV studies show that viral infection increases (E)-β-caryophyllene emission, attracting nonviruliferous GRLHs but not viruliferous ones. By contrast, SRBSDV-infected WBPHs become more sensitive to β-caryophyllene through Or127 upregulation, suggesting that β-caryophyllene-related cues are differentially perceived across vector species and infection statuses.

Geminiviruses coordinate plant volatile cues, insect olfactory receptors, and neural signaling to reshape vector behavior

Geminiviruses are economically important plant single-stranded DNA viruses that are traditionally considered to be transmitted by whiteflies, leafhoppers, or aphids in a circulative-persistent manner [58–60]. However, recent evidence that TYLCV, a representative member of the genus Begomovirus, can replicate within whiteflies suggests that its transmission may also exhibit propagative features [23]. TYLCV-free Bemisia tabaci MED (Mediterranean genetic group) preferentially settle on TYLCV-infected tomato plants rather than healthy plants, likely attributable to virus infection-mediated modifications of plant volatiles [61]. By contrast, TYLCV-infected whiteflies lose this preference and no longer discriminate between infected and healthy tomatoes, indicating that the virus directly modulates vector feeding behavior. TYLCV employs a dual-level manipulation strategy that integrates host volatile modulation with vector sensory reprogramming to enhance transmission. TYLCV-infected plants significantly increase the emission of the monoterpene β-myrcene, which enhances their attractiveness to virus-free Bemisia tabaci MED through the odorant receptor BtMEDOR6. Silencing BtMEDOR6 abolishes the innate preference of naïve MED for infected plants, confirming its essential role in volatile detection. Intriguingly, after virus acquisition, BtMEDOR6 expression is markedly downregulated, resulting in loss of the preference for different host plants to facilitate insect vector dispersal from infected plants to healthy hosts [62] (Fig 4).

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Fig 4. Geminivirus TYLCV reshapes vector behavior through plant volatile cues, olfactory receptor modulation, and neural signaling.

TYLCV-infected tomato plants emit higher levels of the volatile β-myrcene, which increases their attractiveness to the primary vector, the whitefly Bemisia tabaci MED (indirect manipulation). In viruliferous whiteflies, TYLCV suppresses the expression of the β-myrcene receptor BtOR6 while simultaneously activating caspase-dependent neuronal apoptosis. Together, these processes abolish host preference in viruliferous insects, driving their dispersal from infected to healthy plants and thereby facilitating TYLCV transmission (direct manipulation). This figure was created in BioRender. https://BioRender.com/gydn1yh.

https://doi.org/10.1371/journal.ppat.1014368.g004

As a direct vector-targeted mechanism, TYLCV infection also triggers apoptotic neurodegeneration in the whitefly brain, characterized by vacuolar neuropathological lesions reminiscent of those described in Alzheimer’s and Parkinson’s disease models [63]. This neurodegeneration neutralizes host preference after virus acquisition, thereby biasing viruliferous vectors toward healthy plants and completing the transmission cycle. Mechanistically, TYLCV infection induces Caspase 3b cleavage into its active homodimer, resulting in apoptotic neurodegeneration, whereas silencing Caspase 1 or Caspase 3b mitigates TYLCV-induced loss of host preference in free-choice assays [62] (Fig 4). Collectively, TYLCV first attracts naïve vectors to infected plants through altered plant volatiles and then neutralizes host preference after acquisition by altering olfactory receptor expression and inducing neurodegeneration, thereby optimizing viruliferous vector behavior for efficient transmission. Whether replication within insect vectors also occurs in other geminiviruses remains unresolved, but this question may shift the field from viewing geminiviruses as circulative-persistent viruses toward recognizing potential persistent-propagative features.

Knowledge gaps and future perspectives

Accumulating evidence supports the conclusion that persistent-propagative plant viruses reshape their vectors at multiple levels, from morphology to behavior, to optimize transmission. Over the past decade, the molecular mechanisms underlying these phenomena have begun to be elucidated. Despite these advances, several challenges remain in experimental systems, technological platforms, and field-relevant ecological complexity.

Expanding reverse genetics systems for plant NSR and dsRNA viruses

Most plant negative-strand RNA (NSR) viruses and double-stranded RNA (dsRNA) viruses are transmitted by insect vectors in a persistent-propagative manner. However, reverse genetics systems have been established for only a few plant NSR viruses, and no infectious reverse genetics system is yet widely available for plant dsRNA reoviruses in either plant hosts or insect vectors. This limitation restricts the use of gene-deletion or site-directed viral mutants to rigorously define the functions of viral genes during infection and transmission. The recent development of reverse genetics systems for plant rhabdoviruses, including BYSMV, northern cereal mosaic virus (NCMV), RSMV, and MMV [9,64–67], has greatly advanced studies of tripartite interactions among viruses, host plants, and insect vectors. GFP- and/or RFP-expressing BYSMV, NCMV, RSMV, and MMV clones enable real-time and unbiased monitoring of viral infection dynamics in both plants and insect vectors [9,65,66,68,69]. More importantly, individual viral genes can be mutated to uncover their authentic functions in genetically defined systems [68,70,71]. Reverse genetics systems have also been established for some plant bunyaviruses, such as TSWV, providing valuable tools for studying virus–plant–vector interactions [72]. For RSV, infectious clones remain unavailable, although minireplicon systems have been developed [73,74]. Reverse genetics systems remain unavailable for plant dsRNA reoviruses such as RDV, RBSDV, and SRBSDV. However, the successful rescue of several animal dsRNA viruses provides valuable conceptual and technical templates for developing comparable reverse genetics platforms for plant dsRNA viruses [75]. These advances in reverse genetics will present new opportunities for extensive investigations of virus–insect interactions.

Efficient CRISPR-based systems are needed for the genetic manipulation of insect vectors

Most nonmodel insect vectors still lack reliable genetic transformation systems, which limits transgenic overexpression and gene knockout approaches for functional validation of vector genes. As a result, dsRNA-mediated RNA interference is widely used to silence vector genes, whereas CRISPR-based knockouts, although achieved in planthoppers and aphids, remain labor-intensive, technically demanding, and often inefficient [76–78]. A major challenge is that many insect vectors have small bodies and fragile embryos, making transformation methods established in model insects such as Drosophila difficult to apply directly.

Virus-mediated expression and genome editing systems may provide a powerful alternative strategy for functional studies in insect vectors. Plant rhabdovirus-based vectors, including BYSMV and RSMV, have been developed to express foreign proteins in several planthopper and leafhopper vectors, including Nilaparvata lugens, Laodelphax striatellus, Sogatella furcifera, and Recilia dorsalis [65,79]. The BYSMV vector has been developed into a delivery system for Virus-induced Genome Editing in Tiller (ViGET) for heritable and transgene-free genome editing in monocot crops [80]. Extending similar virus-mediated genome editing strategies to insect vectors would be highly valuable, as it could substantially lower the technical barrier for functional genetic studies of vector genes involved in virus acquisition, circulation, replication, and transmission, as well as for controlling insect-borne viral diseases. For example, knockout of the aphid acrostyle receptor Stylin-01, required for CaMV P2 binding, severely impaired CaMV transmission, demonstrating that vector genome editing can disrupt viral receptor sites and limit virus spread [81].

From single virus–vector systems to multidimensional interaction networks

Current understanding of virus-mediated manipulation of insect vector behavior is still largely derived from laboratory studies using single virus–plant–vector systems. However, under field conditions, host plants and insect vectors are frequently challenged by mixed infections involving multiple plant viruses, insect-specific viruses, and microbial symbionts. For example, several rice-infecting viruses share both host plants and insect vectors. RDV and RYSV are transmitted by the green rice leafhopper Nephotettix cincticeps [51,35], whereas rice gall dwarf virus (RGDV) and RSMV are transmitted by the leafhopper Recilia dorsalis [82]. Such overlap in host and vector ranges raises an important question: do co-occurring plant viruses cooperate, compete, or independently manipulate the same plant–vector interface under mixed-infection conditions?

Beyond plant virus–plant virus interactions, insect-associated symbiotic viruses can also modulate plant virus transmission. The leafhopper symbiotic virus Recilia dorsalis bunyavirus (RdBV) enhances RSMV transmission by suppressing the insect E3 ubiquitin ligase RdSina-mediated degradation of the RSMV phosphoprotein, thereby promoting RSMV accumulation [83]. Similarly, the leafhopper symbiotic virus Recilia dorsalis filamentous virus (RdFV) promotes paternal vertical transmission of RGDV, with its capsid protein interacting with RGDV P8 to facilitate viral movement through the male reproductive pathway [84]. Bacterial symbionts add another layer of complexity to plant virus–vector interactions. RDV exploits the obligate symbiont Sulcia in green rice leafhoppers, with its outer capsid protein P2 directly interacting with a Sulcia outer membrane protein to access the symbiont-associated oocyte entry route for transovarial transmission [85]. Conversely, Wolbachia suppresses rice ragged stunt virus (RRSV) infection and transmission in Nilaparvata lugens [86], indicating that bacterial symbionts can either facilitate or restrict plant virus transmission. Together, these findings suggest that virus-mediated vector manipulation is not a simple one-virus–one-vector process, but rather a multidimensional network shaped by co-infecting plant viruses, insect-associated viruses, bacterial symbionts, and host plant responses.

Concluding remarks

Insect vector transmission is essential for the infection cycles of most plant viruses. Accumulating evidence has revealed that plant viruses are not passive passengers but actively modulate the feeding behavior, physiology, and development of insect vectors to promote efficient transmission through both indirect plant-mediated and direct vector-targeted mechanisms. Deciphering the molecular basis of these interactions will not only advance our understanding of virus–plant–insect coevolution but also facilitate the development of novel strategies to control insect vectors and interrupt viral spread. Future studies should establish robust viral reverse genetics systems and efficient insect genetic manipulation tools, while extending research from simplified laboratory models to field-relevant ecological contexts involving mixed infections, vector microbiota, environmental variation, and multitrophic interactions.

Acknowledgments

We are grateful for the helpful discussion from our colleagues J. Yu, D. Li, and Y. Zhang.