Open access peer-reviewed chapter

Vector Control: Insights Arising from the Post-Genomics Findings on Insects’ Reproductive Biology

Written By

Isabela Ramos and Fabio Gomes

Submitted: 27 May 2022 Reviewed: 04 July 2022 Published: 18 August 2022

DOI: 10.5772/intechopen.106273

From the Edited Volume

New Advances in Neglected Tropical Diseases

Edited by Márcia Aparecida Sperança

Chapter metrics overview

235 Chapter Downloads

View Full Metrics

Abstract

The high prevalence of neglected vector-borne diseases, such as Chagas disease and dengue fever, imposes enormous health and financial burdens in developing countries. Historically, and still, to this day, the main effective methods to manage those diseases rely on vector population control. Although early efforts in understanding vector-specific biology resulted in important advancements in the development of strategies for the management of vector-borne diseases, studies regarding the complex physiology of local vector species were weakened by the expanding use of insecticide-based tools, which were, at the time, proven simpler and effective. The rising threat of insecticide resistance and climate change (which can expand endemic areas) has reemphasized the need to rely on thorough species-specific vector biology. One approach to controlling vector populations is to disrupt molecular processes or antagonize the metabolic targets required to produce viable eggs. Here, we discuss new findings arising from post-genomics molecular studies on vector reproductive biology and discuss their potential for the elaboration of new effective vector control interventions.

Keywords

  • reproductive biology
  • post-genomics
  • evidence-based vector control

1. Introduction

1.1 Vector-borne diseases

Vector-borne diseases remain among the deadliest and most prevalent infectious diseases worldwide. Mosquito-transmitted arboviral diseases inflict an enormous burden in tropical areas of the world. A previous study estimated that almost 400 million people are infected by the dengue virus (DENV) every year [1]. While tremendous progress on vector control has been made over the twentieth and twenty-first centuries, we are now facing a critical moment. Record-breaking numbers of dengue cases were detected in the Americas in 2019 [2]. This is aggravated by the fact that multiple serotypes circulate in these regions, maximizing the risk of hemorrhagic fever and other severe complications. Other arboviruses are also of concern. For example, the Zika virus (ZIKV), transmitted by Aedes aegypti, rapidly spread in the Americas in 2014–2015, where it was linked with a surge of newborn malformations, including microcephaly [3, 4], inflicting a lifelong impact on children, their parents, and the health public system. Other viruses, like Mayaro (MAYV), chikungunya (CHKV), and yellow fever (YFV), continue to circulate and periodically reemerge. In addition to the arboviruses, vector-borne parasitic diseases, such as Chagas disease, are also medically important. The triatomine bug Rhodnius prolixus is a primary vector of Trypanosoma cruzi, the causative agent of Chagas disease, a neglected disease endemic to Central and South America. Chagas disease remains the main cause of death related to neglected infectious diseases in the Americas. Currently, Chagas affects approximately 8 million people and migration among endemic and non-endemic regions has expanded its occurrence to approximately 350,000 infected carriers around the globe [5].

Overall, these diseases not only result in a high number of deaths and hospitalizations but also generate a huge economic impact due to the disability of people during their learning and working ages [6, 7]. While most restricted to tropical areas, where vectors meet the perfect conditions for mating and reproduction, models of climate change predict an expansion of the global areas suitable for vector reproduction. Under these models, areas of Europe and America have already seen an increase in the suitability of vector populations and this trend is going to accelerate in the following decades [8, 9]. This is of special concern as these pathogens meet an immunologically naive local population lacking any previous exposure to such diseases.

Advertisement

2. Reproduction as a target for vector control

The interference in insect vector natural populations has remained one of the key strategies for the control of vector-borne diseases. For the past 50 years, vector control policies have relied on the utilization of insecticide-based tools. With the rising of resistance spreading across populations, a major threat to the ongoing success of control programs has been acknowledged [10].

Although insects are the largest and most diverse group of animals on the planet, most species are regarded as species with a high reproductive capacity. Females can generally produce a large number of eggs in a short period, and the high rates of embryo viability boost their natural populations. Blood-feeding (hematophagy) is necessary for most human disease vectors to obtain the energy and nutrients required for efficient oogenesis, enabling the abovementioned high rates of oviposition [11]. Within a vector reproductive cycle, the overall process of converting protein from the blood meal into yolk protein precursors (YPPs), as well as coordinating their delivery to developing oocytes is the most complex stage of reproduction and requires the coordination of intricate metabolic and neuroendocrine pathways in the adult female. As a result, a comprehensive understanding of the complexity of egg production is the most promising approach to designing safe tools for interference in vector reproduction (Figure 1).

Figure 1.

Targets for intervention within the reproductive cycle of vectors. After digestion, adult females are able to lay a large number of highly viable eggs, thus contributing to the increase and maintenance of vector natural populations. The complex physiology process of transforming the contents of the blood meal into mature fertilized eggs requires intricate coordination to accomplish vitellogenesis, delivery of the yolk to the oocytes (yolk uptake), eggshell biogenesis (choriogenesis), and fertilization (mating habits). Interference in any of those stages directly impairs vectors' egg production capacity and embryo viability, rendering drastically reduced reproduction rates.

The molecular physiology of oogenesis is highly conserved within the different insect vectors [11, 12, 13]. In brief, oogenesis is triggered by signals from nutritional status and the blood meal. The levels of the sesquiterpene juvenile hormone (JH) [14], secreted by the corpora allata in the brain, increase over the early periods of insect maturity triggering changes in the fat body that become sensitive to the ovary-producing steroid hormone ecdysone [15]. After the blood meal, the brain stops JH synthesis and releases the ovarian ecdysiotropic hormone, signaling to the ovaries to produce ecdysone. In the fat body, ecdysone is hydroxylated to 20-hydroxyecdysone (20E) and binds to the 20E receptor EcR/USP to trigger vitellogenesis, that is, the production of the YPPs (yolk protein precursors). YPPs are secreted to the hemolymph and delivered to the developing oocytes in the ovaries via receptor-mediated endocytosis. Apart from the huge metabolic challenge of transforming the blood meal into a large number of eggs, the maximum capacity of egg production is also dependent on successful mating, fertilization, and proper conditions for embryo development [12, 16, 17, 18, 19, 20, 21, 22, 23].

Advertisement

3. Potential targets for intervention within vectors reproduction

3.1 The molecular mechanisms of vitellogenesis and oogenesis

The conversion of protein from the blood meal into YPPs for eggs biogenesis is a vital component of the reproductive cycle and understanding how this process is regulated is necessary to design safe, specific, and effective ways to block reproduction in vectors. Mosquito 20E has been shown to play multiple roles in Anopheles.Apart from regulating monogamy in Anopheles, the male-transferred 20E was shown to be important in maintaining sperm viability over the female lifetime through induction of the heme peroxidase 15 (HPX15) [24]. Accordingly, HPX15 knockdown was shown to dramatically increase the ratio of infertile eggs. Upon insemination, the male 20E further interacts with the female Mating-Induced Stimulator of Oogenesis (MISO) and induces an increase in fecundity by increasing the expression of LP and oocyte numbers [25]. Finally, 20E was also shown to be necessary for Anopheles egg-laying [26].

On that note, several genes that are somehow essential for oogenesis and generate unviable embryos have been identified and functionally tested in R. prolixus. The orthologue of Bicaudal C (BicC), a gene originally identified in D. melanogaster involved in embryonic patterning has shown to be maternally expressed and essential for the arrangement of the follicle cells [27]. The control of iron and heme homeostasis is particularly critical for hematophagous insects, especially for the strictly hematophagous triatomines, such R. prolixus. In this model, the silencing of multiple iron-related genes, namely, ferritin, iron responsive protein 1 (IRP1), heme oxygenase (HO), and heme exporter feline leukemia virus C receptor (FLVCR), impairs oogenesis and embryo viability [28, 29].

The role of receptor-mediated endocytosis in yolk uptake has been investigated in oocytes of many insect species. The internalization of yolk proteins through the presence of a specialized endocytic cortex in the oocytes, which includes prominent microvilli, coated pits, coated vesicles, and endosomes have been shown in several species, including Aedes [30, 31, 32, 33, 34, 35, 36, 37, 38] However, the regulations encompassing the recruitment of the endocytic machinery to specific sites of the oocyte cortex and the signals that govern the oocyte endocytic pathways and endosomal maturation are yet to be addressed. In R. prolixus, ATG6/Beclin1 class-III PI3K complexes I and II were shown to be essential for YPP uptake. Insects silenced for the genes present in both complexes produce yolk-deficient eggs generating unviable embryos due to the lack of generated phosphatidylinositol-3P (PI3P) to recruit the endocytic machinery in vitellogenic oocytes [39, 40].

3.2 Choriogenesis as an emerging target for safe interventions

The final checkpoint of oogenesis, before fertilization, is the triggering of the choriogenesis program, in which the multiple layers of the chorion are secreted by the follicle cells that envelop the developing oocytes. Remarkably, while the chorion’s primary protective function is conserved in insects, its general composition and structure have evolved in a highly species-specific manner, giving rise to a wide range of morphologies and functional adaptations. The main chorion proteins in insects have been identified in models, such as the silk moth Bombyx mori, the fruit fly D. melanogaster, and the mosquito Aedes Aegypti, and revealed to be broadly unrelated to their counterparts in each of these species [41, 42]. Proteins that are conserved in a wide variety of organisms are not ideal target molecules as vector control agents because of deleterious effects on non-target organisms, such as vertebrates, pollinating agricultural insects, and beneficial predators. As a result, studies on the molecular biology of the chorion biogenesis in insect vectors are biotechnology strategic as they are likely to unravel safe molecular targets that are at the same time essential for reproduction and highly specific to one species.

The A. aegypti eggshell is composed of different structural proteins, enzymes, odorant-binding proteins, as well as many uncharacterized proteins of unknown function. Melanization proteins and enzymes of the eggshell have been identified [43, 44, 45, 46, 47, 48, 49], and proteomics studies have been performed [42]. Isoe and colleagues [50] designed in silico analyses to identify mosquito-specific genes that are essential for successful embryo development. After systematic RNAi functional screening of over 40 selected genes, the authors identified a chorion-related protein named eggshell organizing factor 1 (EOF1), which is essential for eggshell biogenesis and embryo development. The EOF1 sequence includes an F-box functional motif, which is characterized by the interaction with the SKP1 protein in the SCF E3 ubiquitin ligase complex [51]. Although its exact function in the eggshell has not been elucidated, such findings are very promising in terms of designing safe strategies for vector control.

In R. prolixus, some aspects of the chorion ultrastructure and permeability properties were previously explored [52, 53, 54] and the identification of the specific chorion proteins Rp30 and Rp45, the latter associated with antifungal activity, was also described [55]. In this model, the cell biology of the follicle cells (FCs), the tissue that synthesizes and secretes the chorion components, has been explored. Early transcriptome analysis showed that the FCs are committed to transcription, translation, and vesicular traffic [56]. Accordingly, electron microscopy evidenced the FC’s typical secretory cell morphology with a high content of vesiculated rough endoplasmic reticulum [57, 58]. Systemic RNAi experiments targeting the autophagy-related genes ULK1/ATG1, the autophagy-dedicated E2-conjugating enzyme ATG3 [57, 59], and E1-activating and E2-conjugating ubiquitin enzymes [60] generated particular phenotypes of chorion malformations due to alterations in the general protein homeostasis of the FCs during choriogenesis, resulting in extremely lower rates of embryo viability. Taken together, the data points to a high degree of complexity in the chorion biogenesis program in R. prolixus, rendering the process extremely sensitive to changes in proteostasis of the FCs, and, thus, an interesting target for slight but effective interventions.

Resistance to desiccation is another potential intervention target. Although mosquito eggs are laid in water, they are susceptible to dehydration in the first hours of development. Thus, this property directly affects mosquito reproduction. In A. Aegypti, the serosal cuticle secretion (an inner layer of the chorion secreted during embryogenesis) coincides with an increase in dry resistance and the presence of chitin as one of the serosal cuticle components has been detected [61, 62, 63]. In R. prolixus, chitin was detected in the ovaries and the embryonic cuticle [64, 65]. Additionally, exposure to lufenuron (a chitin synthesis inhibitor) and chitin synthase RNAi experiments reduced oviposition and embryo viability [66]. Therefore, the synthesis and deposition of chitin or chitin-like components in the eggshells are also promising targets for reproduction interventions.

Altogether, and combined with the above-mentioned high degree of species-specificity of the chorion proteins, choriogenesis has the potential to emerge as the foremost target for the generation of new and environmentally safe strategies to achieve vector control.

3.3 Molecular neuroendocrine control of egg production

Major advancements have been achieved in the understanding of the neurohormonal control of egg production. In R. prolixus, a detailed model depicting the control of oogenesis, ovulation, and oviposition has been designed and elegantly reviewed by Lange and colleagues [13]. Post-genomics has allowed the identification and functional characterization of dozens of reproduction-related neuropeptide receptor families, processing enzymes, and neurochemicals. Historically, R. prolixus has been an important model, wherein the basics of insect physiology have been determined [67, 68, 69, 70]. Interestingly, the integration of the post-genomics findings with the smartly designed early physiology experiments has allowed the elucidation and depiction of many aspects of the global endocrinal integration in this vector.

3.4 Crosstalk between reproduction and immunity

A relationship between reproductive potential and immune status has been long established. Collectively, these studies suggest a tradeoff between immune activation and egg production, reflected by the identification of follicular atresia and other cell death pathways [71, 72]. Recent advances are now highlighting the role of nutrient-sensing pathways and vector immunometabolism [73]. Future studies will provide further insight into how signaling pathways, such as TOR and Insulin pathways, well-known vitellogenic and immune regulators, coordinate energetic balance during infection. Interestingly, previous work using natural combinations of vector-parasite has suggested that coevolution might have minimized the impact of infection [74], possibly by the fact that immune tolerance can induce a less-energetic costly immune response.

Rerouting of yolk components can be used as a nutritional factor for parasite development. Mosquito LP has been incorporated into Plasmodium oocyst as a lipid source [75]. While parasite development was accelerated by LP delivery, it did not induce any detectable reproductive cost [76]. Interestingly, mosquito lipids influenced not only total parasite numbers, but also Plasmodium sporozoite virulence upon transmission to vertebrate hosts [77]. Similarly, VG is a key component for Plasmodium survival. An interplay between YPPs and immune response has been demonstrated. Both LP and VG were shown to reduce the efficiency of the binding of the major parasite-killing TEP1 [78], increasing parasite survival following mosquito infection.

3.5 Interventions on mosquito mating and insemination

Mosquitoes are thought to use a set of sonorous, visual, and chemical cues to identify and attract their partners. While the manipulation of such signals used to guide mosquitoes is an interesting target to prevent mosquito mating, the molecular identity of its components, such as sex pheromones and their odorant-binding receptors are scant. In that sense, both Anopheles and Aedes mosquitoes can adopt a swarming behavior during mating. Aggregation pheromones have been identified in Anopheles [79]. Such compounds can be used to manipulate mating behavior in wild vector populations and are a likely target of vector control strategies. More recently, genes regulating cuticular hydrocarbon productions and the circadian cycle have been described to be coordinated with light and temperature to guide swarming in Anopheles [80]. Aggregation pheromones have also been described in A. aegypti (Fawaz et al., 2014). Interestingly, Aedes swarming does not require swarming before mate and Aedes mosquitoes have been shown to mate in pairs throughout the day [81].

Upon mating, male sperm is transferred to a spermatheca (one in Anopheles, two in Aedes) where it is stored for the lifetime of the female mosquito. The role of odorant receptors in activating spermatozoa flagella has been previously shown [82]. While several candidate agonists were shown to activate flagellar beating, its physiological ligand remains to be further defined. Upon insemination, females are thought to mate once in their lifetime. In most Anopheles species, this is enforced by the formation of a mating plug that forms a barrier to prevent further female insemination [83]. The mating plug is composed of seminal secretions produced by the male accessory glands [84], and 20-hydroxyecdysone (20E) embedded is thought to play a signaling role in inducing monogamy in the female [26]. While a mating plug is not formed in Aedes, a physical barrier is temporarily formed by components of the male sperm produced at the male accessory gland [85]. Later, bioactive proteins collectively known as matrone can modulate female behavior at the neuronal levels and induce monogamy [86, 87]. Nevertheless, the exact molecular composition of matrone remains to be defined. A further understanding of the molecular basis for male-induced monogamy is of great importance, as it could potentially identify chemicals that could be used (e.g., in aerosols) to prevent virgin female mating.

3.6 Sterile insect technique (SIT)

General models from the middle of the twentieth century had already predicted the potential of releasing sterile male releases to suppress insect populations [88]. This approach known as the sterile insect technique (SIT) has been originally accomplished by irradiation of mosquitoes. Sterile animals have been released, mostly by preliminary investigations, in several locations around the world with varied success, as previously discussed [89]. While claims of reduced competitive rates of irradiated mosquitoes are still a matter of debate [90], the biggest issues facing SIT seem to rely on the scalability and sustainability of such efforts [89]. An alternative approach has been the release of transgenic-induced sterile mosquitoes. Still, the logistic challenges of such practices remain a major challenge for field implementations in large geographic areas, with wild-type mosquito populations rapidly returning after release interruption [91]. At present, the development of efficient and flexible gene drive techniques, such as CRISPR/Cas9, remains a promising approach to the development of efficient cost-effective SIT implementations independent of continuous mosquito release [92].

3.7 Wolbachia-induced cytoplasmic incompatibility

The utilization of Wolbachia-induced cytoplasmic incompatibility remains one of the most promising alternatives for insecticide-independent strategies of vector control. Wolbachia is an arthropod-specific bacteria that establish a systemic infection and can be vertically transmitted by infecting the host oocytes [93]. Several strains of Wolbachia are known to induce a phenomenon known as cytoplasmic incompatibility (CI) where the progeny from infected males and uninfected females are turned nonviable [94]. While recent reports have identified populations of A. aegypti carrying native Wolbachia infections [95], these seem to be deviations from a general rule where Wolbachia strains are not naturally able to infect A. aegypti. Nevertheless, Wolbachia strains have previously been adapted to infect A. aegypti by trans-infection in the lab, and CI has been shown to manifest in this model. In that sense, the release of CI-carrying Wolbachia-infected males has been proposed as a strategy to suppress Aedes populations, and field trials have been implemented [96, 97]. The molecular mechanisms mediating cytoplasmic incompatibility started to be elucidated and two key genes linked with the prophage WO have been identified [98, 99]. Transgenesis of such genes would provide alternatives to induce CI in the absence of Wolbachia infections [100]. Such strategies would be beneficial for the cases where stable Wolbachia trans-infections have not been achieved, as is the case of many anophelines.

Advertisement

4. Concluding remarks

The control of vector populations has shrunk the map of many vector-borne diseases [10, 101], but new strategies will need to be developed to continue this process. Although the fundamental biology behind oocyte development is known and mostly conserved, its molecular mechanisms are still to be explored. The recent completion of multiple genome sequencing projects will allow comparative genomics studies that not only increase our knowledge about reproductive processes but also facilitate the identification of novel species-specific targets for vector control. Research directed to understanding how this process is regulated and being able to manipulate the female’s capacity to produce so many viable eggs will lead to safe and effective ways to block reproduction in blood-feeding insects. To accomplish this, there is an urgent need to integrate the post-genomics findings with the species-specific vectors’ physiology. Such tactics are the safest path to unravel evidence-based information and design customized tools to manage vector populations in different endemic areas.

Advertisement

Acknowledgments

This work was funded by the following grants. Fundação Carlos Chagas Filho De Amparo À Pesquisa Do Estado Do Rio De Janeiro (FAPERJ) (JCNE E-26/2031802017; http://www.faperj.br/) to I.R.; Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) (INCT-EM 16/2014; http://cnpq.br/) to I.R.; Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) (https://www.gov.br/capes/pt-br) to I.R. The authors thank Vinicius Torrão for the excellent technical assistance with the illustration design.

Advertisement

Conflict of interest

The funders had no role in study design, data collection, analysis, and decision to publish, or preparation of the manuscript.

References

  1. 1. Bhatt S, Gething PW, Brady OJ, Messina JP, Farlow AW, Moyes CL, et al. The global distribution and burden of dengue. Nature. 2013;496:504. DOI: 10.1038/nature12060
  2. 2. Pan American Health Organization. Annual epidemiological update for Dengue, Chikungunya and Zika in 2020. 2020. Available from: https://www.paho.org/en
  3. 3. Mlakar J, Korva M, Tul N, Popović M, Poljšak-Prijatelj M, Mraz J, et al. Zika virus associated with microcephaly. The New England Journal of Medicine. 2016;374:951-958. DOI: 10.1056/NEJMOA1600651/SUPPL_FILE/NEJMOA1600651_DISCLOSURES.PDF
  4. 4. de Araújo TVB, de Ximenes RA, de Miranda-Filho D, Souza WV, Montarroyos UR, de Melo AP, et al. Association between microcephaly, zika virus infection, and other risk factors in Brazil: Final report of a case-control study. The Lancet Infectious Diseases. 2018;18:328-336. DOI: 10.1016/S1473-3099(17)30727-2
  5. 5. Álvarez-Hernández D-A, García-Rodríguez-Arana R, Ortiz-Hernández A, Álvarez-Sánchez M, Wu M, Mejia R, et al. A systematic review of historical and current trends in Chagas disease. Therapeutic Advanced Infectious Diseases. 2021;8:20499361211033716. DOI: 10.1177/20499361211033715
  6. 6. Sachs J, Malaney P. The economic and social burden of malaria. Nature. 2002;415:680-685
  7. 7. Shepard DS, Coudeville L, Halasa YA, Zambrano B, Dayan GH. Economic impact of dengue illness in the Americas. The American Journal of Tropical Medicine and Hygiene. 2011;84:200-207. DOI: 10.4269/AJTMH.2011.10-0503
  8. 8. Caminade C, McIntyre KM, Jones AE. Impact of recent and future climate change on vector-borne diseases. Annals of the New York Academy of Sciences. 2019;1436:157-173. DOI: 10.1111/NYAS.13950
  9. 9. Iwamura T, Guzman-Holst A, Murray KA. Accelerating invasion potential of disease vector Aedes aegypti under climate change. Nature Communication. 2020;111:1-10
  10. 10. Wilson AL, Courtenay O, Kelly-Hope LA, Scott TW, Takken W, Torr SJ, et al. The importance of vector control for the control and elimination of vector-borne diseases. PLoS Neglected Tropical Diseases. 2020;14:1-31. DOI: 10.1371/journal.pntd.0007831
  11. 11. Attardo GM, Hansen IA, Raikhel AS. Nutritional regulation of vitellogenesis in mosquitoes: Implications for anautogeny. Insect Biochemistry and Molecular Biology. 2005;35:661-675. DOI: 10.1016/j.ibmb.2005.02.013
  12. 12. Shaw WR, Attardo GM, Aksoy S, Catteruccia F. A comparative analysis of reproductive biology of insect vectors of human disease. Current Opinion in Insect Science. 2015;10:142-148. DOI: 10.1016/j.cois.2015.05.001
  13. 13. Lange AB, Leyria J, Orchard I. The hormonal and neural control of egg production in the historically important model insect , Rhodnius prolixus : A review , with new insights in this post-genomic era. General and Comparative Endocrinology. 2022:322
  14. 14. Rivera-Pérez C, Clifton ME, Noriega FG, Jindra M. Juvenile hormone regulation and action. Advances in invertebrate (neuro) endocrinology. Apple Academic Press; 2020:1-76
  15. 15. Zou Z, Saha TT, Roy S, Shin SW, Backman TWH, Girke T, et al. Juvenile hormone and its receptor, methoprene-tolerant, control the dynamics of mosquito gene expression. Proceedings of the National Academy of Sciences of the United States of America. 2013;110:E2173-E2181. DOI: 10.1073/pnas.1305293110
  16. 16. Davey KG. Some consequences of copulation in Rhodnius prolixus. Journal of Insect Physiology. 1967;13:1629-1636. DOI: 10.1016/0022-1910(67)90158-8
  17. 17. Davey KG. Effect of the brain and corpus cardiacum on egg production in Rhodnius prolixus. Archives of Insect Biochemistry and Physiology. 1987;4:243-249
  18. 18. Davey KG, Maimets I-K, Ruegg RP. The relationship between crop size and egg production in Rhodnius prolixus. Canadian Journal of Zoology. 1986;64:2654-2657. DOI: 10.1139/z86-385
  19. 19. Davey KG. Inputs to the hormonal control of egg development in Rhodnius prolixus. Memórias do Instituto Oswaldo Cruz. 1987;82:103-108
  20. 20. Borovsky D, Carlson DA, Hancock RG, Rembold H, Van Handel E. De novo biosynthesis of juvenile hormone III and I by the accessory glands of the male mosquito. Insect Biochemistry and Molecular Biology. 1994;24:437-444. DOI: 10.1016/0965-1748(94)90038-8
  21. 21. Clifton ME, Correa S, Rivera-Perez C, Nouzova M, Noriega FG. Male Aedes aegypti mosquitoes use JH III transferred during copulation to influence previtellogenic ovary physiology and affect the reproductive output of female mosquitoes. Journal of Insect Physiology. 2014;64:40-47. DOI: 10.1016/j.jinsphys.2014.03.006
  22. 22. Klowden MJ. Mating and nutritional state affect the reproduction of Aedes albopictus mosquitoes. Journal of American Mosquito Control Association. 1993;9:169-173
  23. 23. Klowden MJ, Chambers GM. Male accessory gland substances activate egg development in nutritionally stressed Aedes aegypti mosquitoes. Journal of Insect Physiology. 1991;37:721-726. DOI: 10.1016/0022-1910(91)90105-9
  24. 24. Shaw WR, Teodori E, Mitchell SN, Baldini F, Gabrieli P, Rogers DW, et al. Mating activates the heme peroxidase HPX15 in the sperm storage organ to ensure fertility in Anopheles gambiae. Proceedings of the National Academy of Sciences. USA. 2014;111:5854-5859. DOI: 10.1073/pnas.1401715111
  25. 25. Baldini F, Gabrieli P, South A, Valim C, Mancini F, Catteruccia F. The interaction between a sexually transferred steroid hormone and a female protein regulates oogenesis in the malaria mosquito Anopheles gambiae. PLoS Biology. 2013;11:e1001695-e1001695. DOI: 10.1371/JOURNAL.PBIO.1001695
  26. 26. Gabrieli P, Kakani EG, Mitchell SN, Mameli E, Want EJ, Mariezcurrena Anton A, et al. Sexual transfer of the steroid hormone 20E induces the postmating switch in Anopheles gambiae. Proceedings of the National Academy of Sciences of the United States of America. 2014;111:16353-16358. DOI: 10.1073/pnas.1410488111
  27. 27. Pascual A, Vilardo ES, Taibo C, Sabio García J, Pomar RR. Bicaudal C is required for the function of the follicular epithelium during oogenesis in Rhodnius prolixus. Development Genes and Evolution. 2021;231:33-45. DOI: 10.1007/s00427-021-00673-0
  28. 28. Walter-Nuno AB, Oliveira MP, Oliveira MF, Gonçalves RL, Ramos IB, Koerich LB, et al. Silencing of maternal heme-binding protein causes embryonic mitochondrial dysfunction and impairs embryogenesis in the blood sucking insect rhodnius prolixus. The Journal of Biological Chemistry. 2013;288:29323-29332. DOI: 10.1074/jbc.M113.504985
  29. 29. Walter-Nuno AB, Taracena ML, Mesquita RD, Oliveira PL, Paiva-Silva GO. Silencing of iron and heme-related genes revealed a paramount role of iron in the physiology of the hematophagous vector Rhodnius prolixus. Frontiers in Genetics. 2018;9:1-21. DOI: 10.3389/fgene.2018.00019
  30. 30. Raikhel AS, Dhadialla TS. Accumulation of yolk proteins in insect oocytes. Annual Review of Entomology. 1992;37:217-251. DOI: 10.1146/annurev.en.37.010192.001245
  31. 31. Roy S, Saha TT, Zou Z, Raikhel AS. Regulatory pathways controlling female insect reproduction. Annual Review of Entomology. 2018;63:489-511. DOI: 10.1146/annurev-ento-020117-043258
  32. 32. Anderson KL, Woodruff RI. A gap junctionally transmitted epithelial cell signal regulates endocytic yolk uptake in Oncopeltus fasciatus. Developmental Biology. 2001;239:68-78. DOI: 10.1006/dbio.2001.0433
  33. 33. Postlethwait JH, Giorgi F. Vitellogenesis in insects. Developmental Biology. 1985;1985(1):85-126
  34. 34. Raikhel AS, Dhadialla TS, Cho W-L, Hays AR, Koller CN. Biosynthesis and endocytosis of yolk proteins in the mosquito, in molecular insect science. Hagedorn HH, Hildebrand JG, Kidwell MG, Law JH, editors. Boston, MA: Springer US; 1990:147-154. DOI: 10.1007/978-1-4899-3668-4_18
  35. 35. Brooks RA, Woodruff RI. Calmodulin transmitted through gap junctions stimulates endocytic incorporation of yolk precursors in insect oocytes. Developmental Biology. 2004;271:339-349
  36. 36. Ferenz H-J. Receptor-mediated endocytosis of insect yolk proteins. In: Hagedorn HH, Hildebrand JG, Kidwell MG, Law JH, editors. Molecular Insect Science. Boston, MA: Springer US; 1990. pp. 131-138. DOI: 10.1007/978-1-4899-3668-4_16
  37. 37. Richard DS, Gilbert M, Crum B, Hollinshead DM, Schelble S, Scheswohl D. Yolk protein endocytosis by oocytes in Drosophila melanogaster: Immunofluorescent localization of clathrin, adaptin and the yolk protein receptor. Journal of Insect Physiology. 2001;47:715-723. DOI: 10.1016/S0022-1910(00)00165-7
  38. 38. Raikhel AS, Dhadialla TS, Cho W-L, Hays AR, Koller CN. Biosynthesis and endocytosis of yolk proteins in the mosquito. In: Hagedorn HH, Hildebrand JG, Kidwell MG, Law JH, editors. Molecular Insect Science. Boston, MA: Springer US; 1990. pp. 147-154. DOI: 10.1007/978-1-4899-3668-4_18
  39. 39. Vieira PH, Benjamim CF, Atella GC, Ramos I. VPS38/UVRAG and ATG14, the variant regulatory subunits of the ATG6/Beclin 1-PI3K complexes, are crucial for the biogenesis of the yolk organelles and are transcriptionally regulated in the oocytes of the vector Rhodnius prolixus. PLoS Neglected Tropical Diseases. 2021;1:1
  40. 40. Vieira PH, Bomfim L, Atella GC, Masuda H, Ramos I. Silencing of RpATG6 impaired the yolk accumulation and the biogenesis of the yolk organelles in the insect vector R. prolixus. PLoS Neglected Tropical Diseases. 2018;12:1-19. DOI: 10.1371/journal.pntd.0006507
  41. 41. Papantonis A, Swevers L, Iatrou K. Chorion genes: A landscape of their evolution, structure, and regulation. Annual Review of Entomology. 2015;60:177-194. DOI: 10.1146/annurev-ento-010814-020810
  42. 42. Marinotti O, Ngo T, Kojin BB, Chou SP, Nguyen B, Juhn J, et al. Integrated proteomic and transcriptomic analysis of the Aedes aegypti eggshell. BMC Developmental Biology. 2014;14:1-11. DOI: 10.1186/1471-213X-14-15
  43. 43. Han Q , Li G, Li J. Purification and characterization of chorion peroxidase from Aedes aegypti eggs. Archives of Biochemistry and Biophysics. 2000;378:107-115. DOI: 10.1006/abbi.2000.1821
  44. 44. Li JS, Li J. Major chorion proteins and their crosslinking during chorion hardening in Aedes aegypti mosquitoes. Insect Biochemistry and Molecular Biology. 2006;36:954-964. DOI: 10.1016/j.ibmb.2006.09.006
  45. 45. Fang J, Han Q , Johnson JK, Christensen BM, Li J. Functional expression and characterization of Aedes aegypti dopachrome conversion enzyme. Biochemical and Biophysical Research Communications. 2002;290:287-293. DOI: 10.1006/bbrc.2001.6200
  46. 46. Johnson JK, Li J, Christensen BM. Cloning and characterization of a dopachrome conversion enzyme from the yellow fever mosquito, Aedes aegypti. Insect Biochemeical and Moecularl Biology. 2001;31:1125-1135. DOI: 10.1016/s0965-1748(01)00072-8
  47. 47. Kim SR, Yao R, Han Q , Christensen BM, Li J. Identification and molecular characterization of a prophenoloxidase involved in Aedes aegypti chorion melanization. Insect Molecular Biology. 2005;14:185-194. DOI: 10.1111/j.1365-2583.2004.00547.x
  48. 48. Li JS, Li J. Characterization of N-linked oligosaccharides in chorion peroxidase of Aedes aegypti mosquito. Protein Science. 2005;14:2370-2386. DOI: 10.1110/ps.051419105
  49. 49. Ferdig MT, Li J, Severson DW, Christensen BM. Mosquito dopa decarboxylase cDNA characterization and blood-meal-induced ovarian expression. Insect Molecular Biology. 1996;5:119-126. DOI: 10.1111/j.1365-2583.1996.tb00046.x
  50. 50. Isoe J, Koch LE, Isoe YE, Rascón AA, Brown HE, Massani BB, et al. Identification and characterization of a mosquito-specific eggshell organizing factor in Aedes aegypti mosquitoes. PLoS Biology. 2019;17:1-23. DOI: 10.1371/journal.pbio.3000068
  51. 51. Wang Z, Liu P, Inuzuka H, Wei W. Roles of F-box proteins in cancer. Nature Reviews. Cancer. 2014;14:233-247. DOI: 10.1038/nrc3700
  52. 52. Dias FA, Gandara AC, Carolin P, Queiroz-Barros FG, RLL O, MHF S, et al. Ovarian dual oxidase (Duox) activity is essential for insect eggshell hardening and waterproofing. The Journal of Biological Chemistry. 2013;288:35058-35067. DOI: 10.1074/jbc.M113.522201
  53. 53. Bomfim L, Vieira P, Fonseca A, Ramos I. Eggshell ultrastructure and delivery of pharmacological inhibitors to the early embryo of R . prolixus by ethanol permeabilization of the extraembryonic layers. PLoS One. 2017;12:e0185770. DOI: 10.1371/journal.pone.0185770
  54. 54. Beament JWL. The waterproofing process in eggs of Rhodnius prolixus Stähl. Proceedings of the Royal Society of London. 1946;133:407-418
  55. 55. Bouts DMD, Melo AC, Andrade AL, Silva-Neto MA, de Paiva-Silva SMH, et al. Biochemical properties of the major proteins from Rhodnius prolixus eggshell. Insect Biochemistry and Molecular Biology. 2007;37:1207-1221. DOI: 10.1016/j.ibmb.2007.07.010
  56. 56. Medeiros MN, Logullo R, Ramos IB, Sorgine MHF, Paiva-Silva GO, Mesquita RD, et al. Transcriptome and gene expression profile of ovarian follicle tissue of the triatomine bug rhodnius prolixus. Insect Biochemistry and Molecular Biology. 2011;41:823-831. DOI: 10.1016/j.ibmb.2011.06.004
  57. 57. Bomfim L, Ramos I. Deficiency of ULK1/ATG1 in the follicle cells disturbs ER homeostasis and causes defective chorion deposition in the vector Rhodnius prolixus. FASEB Journal of Official Publication. 2020;34:13561-13572. DOI: 10.1096/fj.202001396R
  58. 58. Rios T, Bomfim L, Ramos I. The transition from vitellogenesis to choriogenesis triggers the downregulation of the UPR sensors IRE1 and PERK and alterations in the ER architecture in the follicle cells of the vector Rhodnius prolixus. Cell and Tissue Research. 2022;387:63-74. DOI: 10.1007/s00441-021-03547-z
  59. 59. Santos A, Ramos I. ATG3 is important for the chorion ultrastructure during oogenesis in the insect vector Rhodnius prolixus. Frontiers in Physiology. 2021;12:1-11. DOI: 10.3389/fphys.2021.638026
  60. 60. Pereira J, Dias R, Ramos I. Knockdown of E1- and E2-ubiquitin enzymes triggers defective chorion biogenesis and modulation of autophagy-related genes in the follicle cells of the vector Rhodnius prolixus. Journal of Cellular Physiology. 2022;1:12
  61. 61. Farnesi LC, Menna-Barreto RFS, Martins AJ, Valle D, Rezende GL. Physical features and chitin content of eggs from the mosquito vectors Aedes aegypti, Anopheles aquasalis and Culex quinquefasciatus: Connection with distinct levels of resistance to desiccation. Journal of Insect Physiology. 2015;83:43-52. DOI: 10.1016/j.jinsphys.2015.10.006
  62. 62. Rezende GL, Martins AJ, Gentile C, Farnesi LC, Pelajo-Machado M, Peixoto AA, et al. Embryonic desiccation resistance in Aedes aegypti: Presumptive role of the chitinized serosal cuticle. BMC Developmental Biology. 2008;8:1-14
  63. 63. Moreira MF, dos Santos AS, Marotta HR, Mansur JF, Ramos IB, Machado EA, et al. A chitin-like component in Aedes aegypti eggshells, eggs and ovaries. Insect Biochemistry and Molecular Biology. 2007;37:1249-1261. DOI: 10.1016/j.ibmb.2007.07.017
  64. 64. Mansur JF, Figueira-Mansur J, Santos AS, Santos-Junior H, Ramos IB, de Medeiros MN, et al. The effect of lufenuron, a chitin synthesis inhibitor, on oogenesis of Rhodnius prolixus. Pesticide Biochemistry and Physiology. 2010;98:59-67. DOI: 10.1016/j.pestbp.2010.04.013
  65. 65. Souza-Ferreira PS, Mansur JF, Berni M, Moreira MF, Dos Santos RE, Araújo HMM, et al. Chitin deposition on the embryonic cuticle of Rhodnius prolixus: The reduction of CHS transcripts by CHS-ds RNA injection in females affects chitin deposition and eclosion of the first instar nymph. Insect Biochemistry and Molecular Biology. 2014;51:101-109. DOI: 10.1016/j.ibmb.2013.12.004
  66. 66. Mansur JF, Alvarenga ESL, Figueira-Mansur J, Franco TA, Ramos IB, Masuda H, et al. Effects of chitin synthase double-stranded RNA on molting and oogenesis in the Chagas disease vector Rhodnius prolixus. Insect Biochemistry and Molecular Biology. 2014;51:110-121. DOI: 10.1016/j.ibmb.2013.12.006
  67. 67. Wigglesworth VB. Source of moulting hormone in Rhodnius. Nature. 1951;168:558. DOI: 10.1038/168558b0
  68. 68. Wigglesworth VB. Factors controlling moulting and ‘metamorphosis’ in an insect. Nature. 1934;133:725-726. DOI: 10.1038/133725b0
  69. 69. Buxton PA. The biology of a blood-sucking bug, Rhodnius prolixus. Transaction on Entomological Society of London. 1930;78
  70. 70. Wigglesworth VB. The Principles of Insect Physiology. 7th ed. London: Chapman and Hall ltda; 1979. DOI: 10.1007/978.94.009.5973-6
  71. 71. Ahmed AM, Hurd H. Immune stimulation and malaria infection impose reproductive costs in Anopheles gambiae via follicular apoptosis. Immun. Infect. 2006;8:308-315. DOI: 10.1016/j.micinf.2005.06.026
  72. 72. Medeiros MN, Ramos IB, Oliveira DMP, da Silva RCB, Gomes FM, Medeiros LN, et al. Microscopic and molecular characterization of ovarian follicle atresia in Rhodnius prolixus Stahl under immune challenge. Journal of Insect Physiology. 2011;57:945-953. DOI: 10.1016/j.jinsphys.2011.04.010
  73. 73. Samaddar S, Marnin L, Butler LR, Pedra JHF. Immunometabolism in arthropod vectors: Redefining interspecies relationships. Trends in Parasitology. 2020;36:807-815. DOI: 10.1016/J.PT.2020.07.010
  74. 74. Mitchell SN, Catteruccia F. Anopheline reproductive biology: Impacts on Vectorial capacity and potential avenues for malaria control. Cold Spring Harbor Perspectives in Medicine. 2017;7:a025593-a025593. DOI: 10.1101/CSHPERSPECT.A025593
  75. 75. Atella GC, Bittencourt-Cunha PR, Nunes RD, Shahabuddin M, Silva-Neto MAC. The major insect lipoprotein is a lipid source to mosquito stages of malaria parasite. Acta Tropica. 2009;109:159-162. DOI: 10.1016/J.ACTATROPICA.2008.10.004
  76. 76. Werling K, Shaw WR, Itoe MA, Westervelt KA, Marcenac P, Paton DG, et al. Steroid hormone function controls non-competitive plasmodium development in Anopheles. Cell. 2019;177:315-325. DOI: 10.1016/J.CELL.2019.02.036
  77. 77. Costa G, Gildenhard M, Eldering M, Lindquist RL, Hauser AE, Sauerwein R, et al. Non-competitive resource exploitation within mosquito shapes within-host malaria infectivity and virulence. NatureCommunication. 2018;9:1-11
  78. 78. Rono MK, Whitten MMA, Oulad-Abdelghani M, Levashina EA, Marois E. The major yolk protein vitellogenin interferes with the anti-plasmodium response in the malaria mosquito Anopheles gambiae. PLoS Biology. 2010;8:e1000434-e1000434. DOI: 10.1371/JOURNAL.PBIO.1000434
  79. 79. Mozūraitis R, Hajkazemian M, Zawada JW, Szymczak J, Pålsson K, Sekar V, et al. Male swarming aggregation pheromones increase female attraction and mating success among multiple African malaria vector mosquito species. Nature Ecological and Evolution. 2020;410:1401-1395
  80. 80. Wang G, Vega-Rodríguez J, Diabate A, Liu J, Cui C, Nignan C, et al. Clock genes and environmental cues coordinate Anopheles pheromone synthesis, swarming, and mating. Science. 2021;371:411-415. DOI: 10.1126/SCIENCE.ABD4359
  81. 81. Hartberg WK. Observations on the mating behaviour of Aedes aegypti in nature. Bulletin World Health Organ. 1971;45:847
  82. 82. Pitts RJ, Liu C, Zhou X, Malpartida JC, Zwiebel LJ. Odorant receptor-mediated sperm activation in disease vector mosquitoes. Proceedings of the National Academy of Sciences. 2014;111:2566-2571. DOI: 10.1073/pnas.1322923111
  83. 83. Mitchell SN, Kakani EG, South A, Howell PI, Waterhouse RM, Catteruccia F. Evolution of sexual traits influencing vectorial capacity in anopheline mosquitoes. Science. 2015;347:985-988. DOI: 10.1126/science.1259435
  84. 84. Rogers DW, Baldini F, Battaglia F, Panico M, Dell A, Morris HR, et al. Transglutaminase-mediated semen coagulation controls sperm storage in the malaria mosquito. PLoS Biology. 2009;7:e1000272-e1000272. DOI: 10.1371/JOURNAL.PBIO.1000272
  85. 85. Spielman A Sr, Leahy MG, Skaff V. Seminal loss in repeatedly mated female Aedes aegypti. The Biological Bulletin. 1967;132:404-412
  86. 86. Fuchs MS, Craig GB, Hiss EA. The biochemical basis of female monogamy in mosquitoes: I. extraction of the active principle from Aedes aegypti. Life Sciences. 1968;7:835-839. DOI: 10.1016/0024-3205(68)90114-8
  87. 87. Gwadz RW. Neuro-hormonal regulation of sexual receptivity in female Aedes aegypti. Journal of Insect Physiology. 1972;18:259-266. DOI: 10.1016/0022-1910(72)90126-6
  88. 88. Knipling EF. Possibilities of insect control or eradication through the use of sexually sterile males. Journal of Economic Entomology. 1955;48:459-462. DOI: 10.1093/JEE/48.4.459
  89. 89. Benedict MQ. Sterile insect technique: Lessons from the past. Journal of Medical Entomology. 2021;58:1974-1979. DOI: 10.1093/JME/TJAB024
  90. 90. Bouyer J, Vreysen MJB. Yes, irradiated sterile male mosquitoes can Be sexually competitive! Trends in Parasitology. 2020;36:877-880. DOI: 10.1016/J.PT.2020.09.005
  91. 91. Garziera L, Pedrosa MC, de Souza FA, Gómez M, Moreira MB, Virginio JF, et al. Effect of interruption of over-flooding releases of transgenic mosquitoes over wild population of Aedes aegypti: Two case studies in Brazil. Entomologia Experimentalis et Applicata. 2017;164:327-339. DOI: 10.1111/EEA.12618
  92. 92. Macias VM, Ohm JR, Rasgon JL. Gene drive for mosquito control: Where did it come from and where are we headed? International Journal of Environmental Research and Public Health. 2017;14:1006
  93. 93. Kaur R, Shropshire JD, Cross KL, Leigh B, Mansueto AJ, Stewart V, et al. Living in the endosymbiotic world of Wolbachia: A centennial review. Cell Host & Microbe. 2021;29:879-893. DOI: 10.1016/j.chom.2021.03.006
  94. 94. Sicard M, Bonneau M, Weill M. Wolbachia prevalence, diversity, and ability to induce cytoplasmic incompatibility in mosquitoes. Current Opinion in Insect Science. 2019;34:12-20. DOI: 10.1016/J.COIS.2019.02.005
  95. 95. Balaji S, Jayachandran S, Prabagaran SR. Evidence for the natural occurrence of Wolbachia in Aedes aegypti mosquitoes. FEMS Microbiology Letters. 2019;366:55. DOI: 10.1093/FEMSLE/FNZ055
  96. 96. Beebe NW, Pagendam D, Trewin BJ, Boomer A, Bradford M, Ford A, et al. Releasing incompatible males drives strong suppression across populations of wild and Wolbachia-carrying Aedes aegypti in Australia. Proceedings of the National Academy of Sciences of the United States of America. 2021;118:2106828118. DOI: 10.1073/PNAS.2106828118/SUPPL_FILE/PNAS.2106828118.SAPP.PDF
  97. 97. Crawford JE, Clarke DW, Criswell V, Desnoyer M, Cornel D, Deegan B, et al. Efficient production of male Wolbachia-infected Aedes aegypti mosquitoes enables large-scale suppression of wild populations. Nature Biotechnology. 2020;38:482
  98. 98. Beckmann JF, Ronau JA, Hochstrasser M. A Wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility. Nature Microbiology. 2017;25(2):1-7. DOI: 10.1038/nmicrobiol.2017.7
  99. 99. Le Page DP, Metcalf JA, Bordenstein SR, On J, Perlmutter JI, Shropshire JD, et al. Prophage WO genes recapitulate and enhance Wolbachia-induced cytoplasmic incompatibility. Nature. 2017;543:243-274
  100. 100. Adams KL, Abernathy DG, Willett BC, Selland EK, Itoe MA, Catteruccia F. Wolbachia cif B induces cytoplasmic incompatibility in the malaria mosquito vector. Natural Microbiology. 2021;6:1575-1582
  101. 101. Shaw WR, Catteruccia F. Vector biology meets disease control: Using basic research to fight vector-borne diseases. Nature Microbiology. 2019;4:20-34. DOI: 10.1038/s41564-018-0214-7

Written By

Isabela Ramos and Fabio Gomes

Submitted: 27 May 2022 Reviewed: 04 July 2022 Published: 18 August 2022