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Impact of Insecticide Resistance on Vector Competence

The capacity of insects to transmit pathogens is known as vector competence (VC). Evidence indicates that insecticide exposure and resistance increase the risk of pathogen transmission. Under this context, adverse effects such as epidemics in human populations or economic repercussions on crops will increase while current vector control efforts become entirely ineffective. However, studies also point to opposite effects where IR or exposure to insecticides reduces VC. To determine the impact of insecticide resistance (IR) on VC precisely, it is essential to establish reproducible experimental designs to reduce the presence of confusing variables that make the interpretation of results difficult. Therefore, although there is evidence related to the influence of IR on VC, more research is necessary.

  • insecticide resistance
  • insecticide exposure
  • vector competence
  • pathogen transmission

1. Introduction

Insects have a close relationship with humans. They participate in activities that benefit human well-being (i.e., pollination) and exert adverse effects such as those observed in public health, crops, hygiene, and other sectors [1]. On the side of unfavorable impacts, the researchers can highlight the insects’ role as vectors of diseases. For example, mosquitoes (i.e., Culicidae), triatomine bugs (Reduviidae), blackflies (Simuliidae), and lice (Pediculidae) affect human health by transmitting arboviruses, parasites, or bacteria [2]. Additionally, aphids, whiteflies, and thrips transmit pathogens to economically important crops [3]. Together, vectors of human and crop pathogens cause considerable economic losses due to human health costs and lower agricultural production [4].
This capacity of insects to transmit pathogens is known as vector competence (VC). This trait defines the intrinsic capacity of an organism to acquire, maintain replication, disseminate, and transmit a pathogen. VC is a complex trait influenced by factors such as the genetic background of hosts and insects, strain and genotype of pathogens, and other aspects associated with environmental variables such as temperature [5][6]. For example, many studies have evaluated the VC of Aedes aegypti. It has been determined that the extrinsic incubation period (the time needed for a mosquito to become infectious) is shorter at higher temperatures [7]. Additionally, bacterial symbionts affect VC by shaping immune responses [8]. Furthermore, Souza-Neto et al. [9], in a systematic review of VC literature in different populations of Ae. aegypti did not find any record of a fully refractory natural population to virus infection; however, there are populations completely susceptible to Zika, dengue, and chikungunya viruses, demonstrating differential regulation of VC.
Studies of VC are heterogeneous (e.g., pathogen challenges performed through intrathoracic injections vs. membrane feeding), and experimental limitations are present (e.g., lack of model animals that mimic human pathogenesis). There are several reports giving a detailed description of past and current knowledge of VC in mosquitoes [9][10][11][12][13][14].
The primary strategy to avoid adverse impacts of insects on public health or agriculture is the use of pesticides. Pesticides are molecules used to destroy, prevent, or repel insects that are a nuisance to humans [15]. Nowadays, various chemicals are applied to control insect populations; according to their structure and synthesis, the diversity of insecticides includes chlorinated hydrocarbon compounds, organophosphates, carbamates, pyrethroids, neonicotinoids, formamidines, and other molecules, plus botanical and microbial agents [16]. The continued use of insecticides has had unintended consequences, particularly the emergence of resistant populations in both human and crop insect vectors. Given many reports on the ineffectiveness of chemical agents in controlling insect populations, the researchers can conceptualize insecticide resistance (IR). This phenomenon is defined as the decrease in the susceptibility of an insect population to a previously effective insecticide caused by its continued use and/or possible cross-selection with other chemical substances, which arises through genetic, physiological, or behavioral changes and is also a hereditary trait [17][18][19].
IR has been described in vectors regarding a broad spectrum of chemical compounds, including organophosphates [20][21][22], among others. Four mechanisms have been determined to reduce the efficacy of pesticides: changes in insect behavior, thickening of the insect cuticle, increased activity of detoxifying enzymes, and modification of the target site [23][24][25]. The more studied IR mechanisms are target site modification and detoxifying enzyme alteration. For example, there is a vast amount of literature concerning mutations in the voltage-gated sodium channel (VGSC), known together as knockdown resistant mutations (kdr mutations, hereafter), as well as mutations in the acetylcholinesterase gene (referred to as Ace mutations hereafter). These mutations are related to pyrethroid and DDT resistance and organophosphate and carbamate resistance, respectively [26][27]. On the other hand, different enzymes participate in detoxification events leading to metabolic resistance. For instance, mixed-function oxidases are greatly involved in pyrethroid resistance [28][29], along with glutathione S transferases and esterases [30][31]. It has been shown that IR affects current efforts in vector control to prevent the emergence of epidemics of emerging or re-emerging diseases such as chikungunya [32][33]. The same has been observed in pest vectors of important crops such as soybeans and tomatoes [34][35][36].
On the other hand, little is known about the impact of IR on VC. Given the actual scenario in which IR affects vector control, the researchers aimed to systematically synthesize and analyze the research on the effect of IR on VC in vector species that impact human health or crops. It is important to note that the researchers only reviewed research that describes experimental procedures that directly link IR and VC.

2. Effects of Insecticide Exposure on Pathogen Transmission

One key aspect that must be considered while exploring current control efforts’ effectiveness is determining if insecticide exposure could impair or enhance VC. It is possible that exposure to sub-lethal doses of insecticides could decrease insects’ capacity to acquire a pathogen, limiting infection capacity. As this factor remains to be elucidated completely, a contrary enhancing pattern may be present, as reported by Muturi and Alto [37], Moltini-Conclois et al. [38], and Knecht et al. [39]. In this section, the researchers show evidence supporting either of the two scenarios.
Regarding enhancement of VC after exposure to insecticides, Muturi and Alto [40] found increased viral infection and dissemination of the Sindbis virus in Ae. aegypti when larvae were exposed to malathion. However, heat treatment at 30 °C was also applied to immature stages; results must be interpreted with caution as temperature may influence VC [41]. The same effect was found in Ae. aegypti exposed to Bti. Here, the larval exposure augmented dengue infection and dissemination in two Bti-resistant strains of Ae. aegypti [38].
Insecticide susceptibility could interact with physiological characteristics such as insect age. For example, older Ae. albopictus mosquitoes exposed to sublethal doses of bifenthrin develop higher dissemination viral titers of Zika virus than unexposed old (11–12 days old) and younger mosquitoes (6–7 days old). Remarkably, older mosquitoes exposed to bifenthrin exhibited greater viral dissemination to other tissues outside the midgut, even when compared with younger exposed mosquitoes [39].
On the other hand, studies that show an impairment of VC after insecticide exposure are scarcer. Oral consumption of bifenthrin has been shown to reduce dengue infection rates and body titers (dissemination) at 14 dpi in Ae. albopictus, but no effect was observed at 7 dpi [42]. The same trend was consistently found in An. gambiae s. s. collected in Uganda. Homozygous mosquitoes for the kdr mutation L1014S had a reduction in prevalence and infection intensity by P. falciparum after exposure to deltamethrin in contrast to the unexposed control group [43]. Finally, Hauser et al. [44] determined that exposing insects at the larval or adult stage or both stages to permethrin diminished VC of An. gambiae s. s. for P. berghei.
As reported in the previous section, a neutral trend is found in the relationship between insecticide exposure and VC. Alomar et al. [45] reported that exposure to pyriproxyfen had no impact on the infection, dissemination, or transmission rates of the Zika virus in Ae. aegypti.
In this section, the researchers have aimed to review all information regarding the exposure of insects to pesticides; nonetheless, few heterogeneous studies are published. At the time of conducting this research, only seven studies strictly adhered to the direct exposure of immature stages or adult insects to any pesticide and the further evaluation of any component of VC. Besides the small number of studies, there is variation in experimental settings that limits the researchers' ability to establish clear conclusions about the influence of insecticide exposure on VC. For example, only one study was performed in the Anopheles-Plasmodium association [43]; the remaining were conducted in Ae. aegypti or Ae. albopictus [37][38][39][42][45]. To expand this variation, insecticides used for bioassays were from different toxicological groups possessing different modes of action (malathion, an organophosphate [37]; Bti, a biological insecticide [38]; pyriproxyfen, a juvenile hormone analog [45]; bifenthrin [39][42]; permethrin [44]; and deltamethrin [43]). Three studies focused on insecticide exposure in larvae [40][37][45], while the other four were performed in adults [38][41][42][44]; finally, there is also variation in the pathogen used for infections; two studies involved the dengue virus [37][41] (Table 1).
Table 1. Studies aimed at the relationship between insecticide resistance (IR) or exposure to insecticides and vector competence (VC).
Species Pathogen Insecticide Exposure Metabolic Resistance Target Site Modifications Phenotypic Resistance Type of Association Location Additional Treatments Reference
Anopheles gambiae Plasmodium berghei DDT GST   DDT Positive Lab   [46]
An.gambiae s.s. Plasmodium falciparum     L1014S Deltamethrin Positive Field   [47]
Plasmodium falciparum DDT   L1014F DDT Positive Field   [48]
Plasmodium falciparum     L1014F, G119S OP, CAR, and PYR-DDT Negative 1 Lab   [49]
Plasmodium falciparum     L1014F, G119S OP, CAR, and PYR-DDT Positive 2 Lab   [49]
Metarhizium 3 anisopliae     L1014F PYR Positive Lab   [50]
Beauveria bassiana 3     L1014F PYR Positive Lab   [50]
Plasmodium falciparum Deltamethrin   L1014S   Negative Field   [43]
Plasmodium berghei Permethrin       Negative Lab Larval competition [44]
An.s gambiae s. l. Plasmodium falciparum α-Cypermethrin, Deltamethrin, Permethrin   N1575Y, I1527T, L1014F, G119S PYR Neutral Field   [51]
Plasmodium sp.     L1014F, G119S   Neutral Field   [52]
Plasmodium falciparum     L1014F, L1014S   Positive Lab   [53]
An. funestus Plasmodium falciparum     L119F-GSTe2   Negative 1 Lab   [54]
Plasmodium falciparum     L119F-GSTe2   Positive 2 Lab   [54]
Plasmodium sp.     L119F-GSTe2   Neutral Field   [55]
Plasmodium sp.     L119F-GSTe2   Positive Field   [55]
Plasmodium sp.     A296S (GABA)   Negative Field   [55]
Culex gelidus Japanese Encephalitis Virus       Deltamethrin, Malathion Neutral Field   [56]
Cx. pipiens Plasmodium relictum     Ester, AceR   Neutral Field   [57]
Plasmodium relictum     Ester, AceR   Neutral Lab   [57]
Cx. quinquefasciatus Wuchereria bancrofti   Esterase activity     Negative Field   [58]
Wuchereria bancrofti   Esterase activity     Negative Lab   [58]
WNV     G119S, Ester OP Positive Lab   [59]
RVV     G119S, Ester OP Neutral Lab   [59]
Aedes aegypti DENV-2 DDT       Neutral Lab Heat shock [60]
DENV-1 Bti       Neutral Lab Larval densities [61]
Zika     V1016I, F1534C PYR Positive Lab   [62]
Sindbis Malathion       Positive Lab Heat treatment [37]
DENV Bti     Bti Positive Lab   [38]
ZIKV Pyriproxyfen       Neutral Lab   [45]
DENV-1   CYP and GST V1016I, F1534C PYR Positive Lab   [63]
DENV     V1016I, F1534C   Negative Field   [64]
Ae. albopictus DENV-2       Deltamethrin Negative Lab   [65]
Zika Bifenthrin       Positive Lab   [39]
DENV Bifenthrin       Negative Lab   [42]
Frankliniella occidentalis Tomato spotted wilt orthotospovirus       Spinosad Positive Lab   [66]
Tomato spotted wilt orthotospovirus       Spinosad Neutral Lab   [67]
Myzus persicae Potato Virus Y λ-Cyhalothrin   Ace, M918L Diethyl carbamates PYR Positive Lab   [68]
OP, organophosphates; CAR, carbamates; PYR, pyrethroids. 1 Indicates that the negative influence of IR on VC was detected in the prevalence of infection. 2 Indicates that the positive impact of IR or exposure to insecticides was detected in the pathogen’s burden (intensity of infection. 3 They were included to show the relationship between infection susceptibility and IR; these are independent of VC.

3. Impacts of Insecticide Resistance on Pest Vectors in Crops

It has been noted that patterns found in vectors of human pathogens are also present in crop pests. To illustrate this relationship, Wan et al. [66] found that spinosad-resistant Frankliniella occidentalis had higher viral replication, dissemination, and transmission of tomato spotted wilt orthotospovirus (TSWV), contrasting with its susceptible counterpart. These results were obtained only in scenarios where short acquisition periods of the pathogen were evaluated. Regarding life parameters, it is remarkable that resistant F. occidentalis showed a prolonged pre-adult stage, a reduced life span, and lower pupation and sex ratios than its susceptible counterpart. Concerning F. occidentalis IR mechanisms, it has been pointed out that several of these mechanisms resemble those found in human-associated vectors, including decreased penetration of insecticides, detoxifying enzymes, and target resistance. Zhang et al. [70] reported evidence of these similarities, who described genes associated with insecticide resistance using metagenomic approaches and alteration of immune pathways in TSWV-infected F. occidentalis.
Another alteration in VC is observed in Myzus persicae, the peach potato aphid. Here, pyrethroid-susceptible individuals (Type J) displayed less acquisition of potato virus Y in λ-cyhalothrin sprayed leaves than the control (non-sprayed) leaves even three days after spraying. When repeating this experiment using a resistant strain (Type O) characterized by an Ace and a kdr mutation in M918L, the spraying of leaves did not produce a reduction in acquisition of potato virus Y by M. persicae type O. It can be hypothesized that the presence of IR mechanisms affects viral acquisition [68]. In contrast to these findings, no association was determined between IR and VC. Zhao et al. [67] found that, after 48 h, there was no difference in transmission rates between susceptible and spinosad-resistant F. occidentalis individuals. This difference could arise given that IR alters only acquisition events and not the molecular machinery related to the transmission of pathogens [66].
All the studies mentioned earlier were conducted in laboratory settings, and less is known regarding how field variants (e.g., different biotypes) could affect this relationship. Studies conducted in whiteflies (Bemisia tabaci) have reported variations in IR as well as genes related to the transmission of tomato yellow leaf curl virus (TYLCV) and tomato crinivirus (TC) among Middle East-Asia Minor 1 (MEAM 1) and Asia II 1 species. It is important to specify the variation of genes involved in IR, such as acetylcholinesterase like protein and cytochrome P450, proteins acting as target sites for carbamates/organophosphates and detoxifying enzymes for pyrethroids, respectively [71]. On the other hand, protein variants involved in virus transmission were also detected for TYLCV, such as an aldo-ketoreductase and elicitin-like protein 6, in addition to variants for viral transmission of tomato crinivirus such as 70 Da heat shock protein, AAA-ATPase-like domain-containing protein, alpha-glucosidase, and others [71]. Given this example, it is essential to evaluate how this could enhance or hamper VC in the light of IR. Considering these small number of research advances, the researchers can look at fine-scale variation in the relationship between VC and IR in non-human associated vectors. First, it is apparent that, in controlled situations, IR alters the VC of two vectors of crops [66][68][67]. Second, this alteration is complex and remains to be investigated and expanded to other insect species as well as factors such as immune system interplay and evaluation of individuals exposed to insecticides but with one or several resistance mechanisms (Table 1).

This entry is adapted from 10.3390/insects13040377


  1. Eggleton, P. The state of the World’s insects. Annu. Rev. Environ. Resour. 2020, 45, 61–82.
  2. WHO. Vector-Borne Diseases; WHO: Geneva, Switzerland, 2020. Available online: (accessed on 5 December 2021).
  3. Heck, M. Insect transmission of plant pathogens: A systems biology perspective. MSystems 2018, 3, e00168-17.
  4. Savary, S.; Willocquet, L.; Pethybridge, S.J.; Esker, P.; McRoberts, N.; Nelson, A. The global burden of pathogens and pests on major food crops. Nat. Ecol. Evol. 2019, 3, 430–439.
  5. Beerntsen, B.T.; James, A.A.; Christensen, B.M. Genetics of mosquito vector competence. Microbiol. Mol. Biol. Rev. 2000, 64, 115–137.
  6. Viglietta, M.; Bellone, R.; Blisnick, A.A.; Failloux, A.B. Vector specificity of arbovirus transmission. Front. Microbiol. 2021, 12, 773211.
  7. Chan, M.; Johansson, M.A. The incubation periods of dengue viruses. PLoS ONE 2012, 7, e50972.
  8. Gabrieli, P.; Caccia, S.; Varotto-Boccazzi, I.; Arnoldi, I.; Barbieri, G.; Comandatore, F.; Epis, S. Mosquito trilogy: Microbiota, immunity and pathogens, and their implications for the control of disease transmission. Front. Microbiol. 2021, 12, 630438.
  9. Souza-Neto, J.A.; Powell, J.R.; Bonizzoni, M. Aedes aegypti vector competence studies: A review. Infect. Genet. Evol. 2019, 67, 191–209.
  10. Chamberlain, R.W.; Sudia, W.D. Mechanism of transmission of viruses by mosquitoes. Annu. Rev. Entomol. 1961, 6, 371–390.
  11. Hardy, J.L.; Houk, E.J.; Kramer, L.D.; Reeves, W.C. Intrinsic factors affecting vector competence of mosquitoes for arboviruses. Annu. Rev. Entomol. 1983, 28, 229–262.
  12. Franz, A.W.; Kantor, A.M.; Passarelli, A.L.; Clem, R.J. Tissue barriers to arbovirus infection in mosquitoes. Viruses 2015, 7, 3741–3767.
  13. Kramer, L.D.; Ciota, A.T. Dissecting vectorial capacity for mosquito-borne viruses. Curr. Opin. Virol. 2015, 15, 112–118.
  14. Wu, P.; Yu, X.; Wang, P.; Cheng, G. Arbovirus lifecycle in mosquito: Acquisition, propagation and transmission. Expert. Rev. Mol. Med. 2019, 21, e1.
  15. Manyilizu, W.B. Pesticides, Anthropogenic Activities, History and the Health of Our Environment: Lessons from Africa. In Pesticides—Use and Misuse and Their Impact in the Environment; Larramendy, M., Soloneski, S., Eds.; IntechOpen: London, UK, 2019; pp. 1–13. Available online: (accessed on 15 December 2021).
  16. Yu, S.J. The Toxicology and Biochemistry of Insecticides; CRC Press: Boca Raton, FL, USA, 2015; pp. 31–100. ISBN 9781482210606.
  17. WHO. Test Procedures for Insecticide Resistance Monitoring in Malaria Vector Mosquitoes, 2nd ed.; WHO: Geneva, Switzerland, 2016. Available online: (accessed on 15 December 2021).
  18. Oppold, A.M.; Müller, R. Epigenetics: A hidden target of insecticides. In Advances in Insect Physiology; Verlinden, H., Ed.; Academic Press: Cambridge, MA, USA, 2017; pp. 313–324.
  19. Insecticide Resistance Action Committee (IRAC). Resistance. 2019. Available online: (accessed on 15 December 2021).
  20. Hayd, R.L.N.; Carrara, L.; de Melo Lima, J.; de Almeida, N.C.V.; Lima, J.B.P.; Martins, A.J. Evaluation of resistance to pyrethroid and organophosphate adulticides and kdr genotyping in Aedes aegypti populations from Roraima, the northernmost Brazilian State. Parasites Vectors 2020, 13, 264.
  21. Chanda, J.; Saili, K.; Phiri, F.; Stevenson, J.C.; Mwenda, M.; Chishimba, S.; Mulube, C.; Mambwe, B.; Lungu, C.; Earle, D.; et al. Pyrethroid and carbamate resistance in Anopheles funestus Giles along Lake Kariba in Southern Zambia. Am. J. Trop. Med. Hyg. 2020, 103 (Suppl. 2), 90–97.
  22. Chen, M.; Du, Y.; Wu, S.; Nomura, Y.; Zhu, G.; Zhorov, B.S.; Dong, K. Molecular evidence of sequential evolution of DDT- and pyrethroid-resistant sodium channel in Aedes aegypti. PLoS Negl. Trop. Dis. 2019, 13, e0007432.
  23. Dang, K.; Doggett, S.L.; Veera Singham, G.; Lee, C.T. Insecticide resistance and resistance mechanisms in bed bugs, Cimex spp. (Hemiptera: Cimicidae). Parasites Vectors 2017, 10, 318.
  24. Zalucki, M.P.; Furlong, M.J. Behavior as a mechanism of insecticide resistance: Evaluation of the evidence. Curr. Opin. Insect Sci. 2017, 21, 19–25.
  25. Ndiath, M.O. Insecticides and insecticide resistance. Methods Mol. Biol. 2019, 2013, 287–304.
  26. Chen, M.; Du, Y.; Nomura, Y.; Zhorov, B.S.; Dong, K. Chronology of sodium channel mutations associated with pyrethroid resistance in Aedes aegypti. Arch. Insect Biochem. Physiol. 2020, 104, e21686.
  27. Guo, D.; Luo, J.; Zhou, Y.; Xiao, H.; He, K.; Yin, C.; Xu, J.; Li, F. ACE: An efficient and sensitive tool to detect insecticide resistance-associated mutations in insect acetylcholinesterase from RNA-Seq data. BMC Bioinform. 2017, 18, 330.
  28. Aïzoun, N.; Aïkpon, R.; Padonou, G.G.; Oussou, O.; Oké-Agbo, F.; Gnanguenon, V.; Ossè, R.; Akogbéto, M. Mixed-function oxidases and esterases associated with permethrin, deltamethrin and bendiocarb resistance in Anopheles gambiae s.l. in the south-north transect Benin, West Africa. Parasites Vectors 2013, 6, 223.
  29. Pethuan, S.; Jirakanjanakit, N.; Saengtharatip, S.; Chareonviriyaphap, T.; Kaewpa, D.; Rongnoparut, P. Biochemical studies of insecticide resistance in Aedes (Stegomyia) aegypti and Aedes (Stegomyia) albopictus (Diptera: Culicidae) in Thailand. Trop. Biomed. 2007, 24, 7–15.
  30. Che-Mendoza, A.; Penilla, R.P.; Rodriguez, D.A. Insecticide resistance and glutathione S-transferases in mosquitoes: A review. Afr. J. Biotech. 2009, 8, 1386–1397.
  31. Bhatt, P.; Bhatt, K.; Huang, Y.; Lin, Z.; Chen, S. Esterase is a powerful tool for the biodegradation of pyrethroid insecticides. Chemosphere 2020, 244, 125507.
  32. Barbosa, S.; Kay, K.; Chitnis, N.; Hastings, I.M. Modelling the impact of insecticide-based control interventions on the evolution of insecticide resistance and disease transmission. Parasites Vectors 2018, 11, 482.
  33. Shanmugaraj, B.; Malla, A.; Ramalingam, S. Epidemiology, clinical features and transmission of re-emerging arboviral infection chikungunya. Asian Pac. J. Trop. Biomed. 2019, 9, 135–139.
  34. Boudh, S.; Singh, J.S. Pesticide Contamination: Environmental Problems and Remediation Strategies. In Emerging and Eco-Friendly Approaches for Waste Management; Bharagava, R.N., Chowdhary, P., Eds.; Springer: Singapore, 2019; pp. 245–269.
  35. Koch, R.L.; Hodgson, E.W.; Knodel, J.J.; Varenhorst, A.J.; Potter, B.D. Management of insecticide-resistant soybean aphids in the upper midwest of the United States. J. Integr. Pest Manag. 2018, 9, 23.
  36. Roditakis, E.; Vasakis, E.; García-Vidal, L.; Martínez-Aguirre, M.D.R.; Rison, J.L.; Haxaire-Lutun, M.O.; Nauen, R.; Tsagkarakou, A.; Bielza, P. A four-year survey on insecticide resistance and likelihood of chemical control failure for tomato leaf miner Tuta absoluta in the European/Asian region. J. Pest Sci. 2017, 91, 421–435.
  37. Muturi, E.J.; Alto, B.W. Larval environmental temperature and insecticide exposure alter Aedes aegypti competence for arboviruses. Vector-Borne Zoonotic Dis. 2011, 11, 1157–1163.
  38. Moltini-Conclois, I.; Stalinski, R.; Tetreau, G.; Després, L.; Lambrechts, L. Larval exposure to the bacterial insecticide bti enhances dengue virus susceptibility of adult Aedes aegypti mosquitoes. Insects 2018, 9, 193.
  39. Knecht, H.; Richards, S.L.; Balanay, J.A.G.; White, A.V. Impact of mosquito age and insecticide exposure on susceptibility of Aedes albopictus (Diptera: Culicidae) to infection with Zika virus. Pathogens 2018, 7, 67.
  40. Cornet, S.; Gandon, S.; Rivero, A. Patterns of phenoloxidase activity in insecticide resistant and susceptible mosquitoes differ between laboratory-selected and wild-caught individuals. Parasites Vectors 2013, 6, 315.
  41. Bellone, R.; Failloux, A.B. The Role of Temperature in shaping mosquito-borne viruses transmission. Front. Microbiol. 2020, 11, 584846.
  42. Richards, S.L.; White, A.V.; Balanay, J.A.G. Potential for sublethal insecticide exposure to impact vector competence of Aedes albopictus (Diptera: Culicidae) for dengue and Zika viruses. Res. Rep. Trop. Med. 2017, 8, 53–57.
  43. Kristan, M.; Lines, J.; Nuwa, A.; Ntege, C.; Meek, S.R.; Abeku, T.A. Exposure to deltamethrin affects development of Plasmodium falciparum inside wild pyrethroid resistant Anopheles gambiae s.s. mosquitoes in Uganda. Parasites Vectors 2016, 9, 100.
  44. Hauser, G.; Thiévent, K.; Koella, J.C. Consequences of larval competition and exposure to permethrin for the development of the rodent malaria Plasmodium berghei in the mosquito Anopheles gambiae. Parasites Vectors 2020, 13, 107.
  45. Alomar, A.A.; Eastmond, B.H.; Alto, B.W. The effects of exposure to pyriproxyfen and predation on Zika virus infection and transmission in Aedes aegypti. PLoS Negl. Trop. Dis. 2020, 14, e0008846.
  46. Saddler, A.; Burda, P.C.; Koella, J.C. Resisting infection by Plasmodium berghei increases the sensitivity of the malaria vector Anopheles gambiae to DDT. Malar. J. 2015, 14, 134.
  47. Kabula, B.; Tungu, P.; Rippon, E.J.; Steen, K.; Kisinza, W.; Magesa, S.; Mosha, F.; Donnelly, M.J. A significant association between deltamethrin resistance, Plasmodium falciparum infection and the Vgsc-1014S resistance mutation in Anopheles gambiae highlights the epidemiological importance of resistance markers. Malar. J. 2016, 15, 289.
  48. Alout, H.; Yameogo, B.; Djogbénou, L.S.; Chandre, F.; Dabiré, R.K.; Corbel, V.; Cohuet, A. Interplay between Plasmodium infection and resistance to insecticides in vector mosquitoes. J. Infect. Dis. 2014, 210, 1464–1470.
  49. Alout, H.; Ndam, N.T.; Sandeu, M.M.; Djégbe, I.; Chandre, F.; Dabiré, R.K.; Djogbénou, L.S.; Corbel, V.; Cohuet, A. Insecticide resistance alleles affect vector competence of Anopheles gambiae s.s. for Plasmodium falciparum field isolates. PLoS ONE 2013, 8, e63849.
  50. Howard, A.F.; Koenraadt, C.J.; Farenhorst, M.; Knols, B.G.; Takken, W. Pyrethroid resistance in Anopheles gambiae leads to increased susceptibility to the entomopathogenic fungi Metarhizium anisopliae and Beauveria bassiana. Malar. J. 2010, 9, 168.
  51. Collins, E.; Vaselli, N.M.; Sylla, M.; Beavogui, A.H.; Orsborne, J.; Lawrence, G.; Wiegand, R.E.; Irish, S.R.; Walker, T.; Messenger, L.A. The relationship between insecticide resistance, mosquito age and malaria prevalence in Anopheles gambiae s.l. from Guinea. Sci. Rep. 2019, 9, 8846.
  52. Wolie, R.; Koffi, A.A.; Ahoua Alou, L.P.; Sternberg, E.D.; N’Nan-Alla, O.; Dahounto, A.; Yapo, F.H.A.; Kanh, K.M.H.; Camara, S.; Oumbouke, W.A.; et al. Evaluation of the interaction between insecticide resistance-associated genes and malaria transmission in Anopheles gambiae sensu lato in central Côte d’Ivoire. Parasites Vectors 2021, 14, 581.
  53. Ndiath, M.O.; Cailleau, A.; Diedhiou, S.M.; Gaye, A.; Boudin, C.; Richard, V.; Trape, J.F. Effects of the kdr resistance mutation on the susceptibility of wild Anopheles gambiae populations to Plasmodium falciparum: A hindrance for vector control. Malar. J. 2014, 13, 340.
  54. Ndo, C.; Kopya, E.; Irving, H.; Wondji, C. Exploring the impact of glutathione S-transferase (GST)-based metabolic resistance to insecticide on vector competence of Anopheles funestus for Plasmodium falciparum. Wellcome Open Res. 2019, 4, 52.
  55. Tchouakui, M.; Chiang, M.C.; Ndo, C.; Kuicheu, C.K.; Amvongo-Adjia, N.; Wondji, M.J.; Tchoupo, M.; Kusimo, M.O.; Riveron, J.M.; Wondji, C.S. A marker of glutathione S-transferase-mediated resistance to insecticides is associated with higher Plasmodium infection in the African malaria vector Anopheles funestus. Sci. Rep. 2019, 9, 5772.
  56. Anbalagan, R.; Shukla, A.; Subramanian, V.; Srivastava, P.K.; Krishnan, J. Monitoring of insecticide resistance and exploring the presence of virus in field populations of Culex gelidus at Thiruvarur District of Tamil. J. Commun. Dis. 2021, 53, 76–83.
  57. Vézilier, J.; Nicot, A.; Gandon, S.; Rivero, A. Insecticide resistance and malaria transmission: Infection rate and oocyst burden in Culex pipiens mosquitoes infected with Plasmodium relictum. Malar. J. 2020, 9, 379.
  58. McCarroll, L.; Hemingway, J. Can insecticide resistance status affect parasite transmission in mosquitoes? Insect Biochem. Mol. Biol. 2002, 32, 1345–1351.
  59. Atyame, C.M.; Alout, H.; Mousson, L.; Vazeille, M.; Diallo, M.; Weill, M.; Failloux, A.B. Insecticide resistance genes affect Culex quinquefasciatus vector competence for West Nile virus. Proc. Biol. Sci. 2019, 286, 20182273.
  60. Yadav, P.; Barde, P.V.; Gokhale, M.D.; Vipat, V.; Mishra, A.C.; Pal, J.K.; Mourya, D.T. Effect of temperature and insecticide stresses on Aedes aegypti larvae and their influence on the susceptibility of mosquitoes to dengue-2 virus. SE Asian J. Trop. Med. Public Health 2005, 36, 1139–1144.
  61. Alto, B.W.; Lord, C.C. Transstadial Effects of Bti on traits of Aedes aegypti and infection with dengue virus. PLoS Negl. Trop. Dis. 2016, 10, e0004370.
  62. Parker-Crockett, C.; Connelly, C.R.; Siegfried, B.; Alto, B. Influence of pyrethroid resistance on vector competency for Zika virus by Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 2021, 58, 1908–1916.
  63. Chen, T.Y.; Smartt, C.T.; Shin, D. Permethrin resistance in Aedes aegypti affects aspects of vectorial capacity. Insects 2021, 12, 71.
  64. Stephenson, C.J.; Coatsworth, H.; Waits, C.M.; Nazario-Maldonado, N.M.; Mathias, D.K.; Dinglasan, R.R.; Lednicky, J.A. Geographic partitioning of dengue virus transmission risk in Florida. Viruses 2021, 13, 2232.
  65. Deng, J.; Guo, Y.; Su, X.; Liu, S.; Yang, W.; Wu, Y.; Wu, K.; Yan, G.; Chen, X.G. Impact of deltamethrin-resistance in Aedes albopictus on its fitness cost and vector competence. PLoS Negl. Trop. Dis. 2021, 15, e0009391.
  66. Wan, Y.; Zheng, X.; Xu, B.; Xie, W.; Wang, S.; Zhang, Y.; Zhou, X.; Wu, Q. Insecticide resistance increases the vector competence: A case study in Frankliniella occidentalis. J. Pest Sci. 2021, 94, 83–91.
  67. Zhao, W.; Wan, Y.; Xie, W.; Xu, B.; Zhang, Y.; Wang, S.; Wei, G.; Zhou, X.; Wu, Q. Effect of spinosad resistance on transmission of tomato spotted wilt virus by the western flower thrips (Thysanoptera: Thripidae). J. Econ. Entomol. 2015, 109, 62–69.
  68. Fenton, B.; Salter, W.T.; Malloch, G.; Begg, G.; Anderson, E. Stopped in its tracks: How λ-cyhalothrin can break the aphid transmission of a potato potyvirus. Pest Manag. Sci. 2015, 71, 1611–1616.
  69. Pigeault, R.; Nicot, A.; Gandon, S.; Rivero, A. Mosquito age and avian malaria infection. Malar. J. 2015, 14, 383.
  70. Zhang, Z.; Zhang, P.; Li, W.; Zhang, J.; Huang, F.; Yang, J.; Bei, Y.; Lu, Y. De novo transcriptome sequencing in Frankliniella occidentalis to identify genes involved in plant virus transmission and insecticide resistance. Genomics 2013, 101, 296–305.
  71. Hussain, S.; Farooq, M.; Malik, H.J.; Amin, I.; Scheffler, B.E.; Scheffler, J.A.; Liu, S.S.; Mansoor, S. Whole genome sequencing of Asia II 1 species of whitefly reveals that genes involved in virus transmission and insecticide resistance have genetic variances between Asia II 1 and MEAM1 species. BMC Genom. 2019, 20, 507.
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