Transgenic Manipulation of Arthropod Vectors: Tools to Study Vector-Borne Diseases

Blood-feeding arthropods such as mosquitoes, ticks, and sandflies transmit several medically relevant pathogens, many of which are understudied and lack adequate therapeutics. Therefore, preventing disease transmission often requires targeting the arthropod¹. For example, integrated vector management (IVM) is a comprehensive framework that aims to increase the efficacy of traditional vector control methods, such as insecticides, by augmenting them with more innovative methods such as genetic control². This special collection highlights a diverse array of transgenic arthropods being generated to study and control vector-borne diseases, and the technical and creative skills necessary to develop such genetically modified organisms.

Arthropod-pathogen-host relationships evolve over many years to shape the genetics of all affected species^3,4,5. Model systems are typically inadequate to study blood-feeding arthropods because their biology and behaviors are species-specific. Molecular entomologists must recreate the complex environments that allow for blood feeding and reproduction so the individual species can be routinely maintained and studied in the laboratory under controlled conditions. Rearing hematophagous arthropods requires both biological expertise and the “meticulous attention of skilled individuals”⁶. For this reason, it is no wonder that vector biology is at the forefront of genetic innovation.

Although this special issue covers generating transgenic arthropods across diverse genera, one central theme emerges: the process requires a keen attention to detail, a sharp understanding of the organism’s biology, and an inventive mindset. Louradour et al.⁷ highlight the challenges of adapting embryonic microinjection techniques to the vectors of Leishmania parasites – namely, sandflies – that lay very small eggs. Freshly laid eggs are transferred to individual pieces of black filter paper on a microscope slide with a coverslip and are carefully lined up for microinjection. The humidity levels, the quality of the prepared materials, and the care with which the process is performed are just a few variables that determine whether the technique succeeds. Hard tick egg microinjection presents a distinct set of challenges – a thick external wax layer on the embryos, hard chorion, and high intra-oval pressure⁸. Sharma et al. describe procedures in Ixodes scapularis that overcome these obstacles by removing the wax layer, softening the chorion, and desiccating the eggs⁸. These include ablating the Gené's organ (wax gland) from egg-laying females, collecting the eggs within 24 h of laying, and then placing the eggs in a 1% (w/v) sodium chloride solution for approximately 1 h before microinjecting⁸.

It is possible to circumvent embryonic microinjections by injecting pupa or adults with the machinery that edits or silences genes. Benetta et al. describe this technique using two ovary delivery methods – Receptor-Mediated Ovary Transduction of Cargo (ReMOT Control) or Branched Amphiphilic Peptide Capsules (BAPC) – as well as two injection strategies (either a Femtojet or an aspirator tube) in the parasitic jewel wasp Nasonia vitripennis⁹. Pupae were lined up and attached to a microscope slide with glue to facilitate the injection process and reduce mortality⁹. Adults, on the other hand, were anesthetized on ice so that the user could support the female with forceps during the abdomen injection.

In addition to CRISPR/Cas9-mediated gene editing, this issue covers other genetic engineering technologies used in arthropods, including site-directed φC31 recombinase and GAL4-UAS. The former, described by Adolfi et al., achieves site-directed integration of attB-flanked transgenic cargo for single integration or recombinase-mediated cassette exchange in the malaria vectors Anopheles gambiae and Anopheles stephensi¹⁰. This technology allows for site-directed insertion into docking lines that have been previously established, which simplifies validation by avoiding positional effects and downstream mating schemes necessary for a stable genetic line¹⁰. The other technique, the bipartite GAL4-UAS system, is used by Poulton et al. to generate transgenic Anopheles gambiae that express transgene cassettes in a spatiotemporal, tissue-specific manner, even at the expense of severe fitness costs¹¹. Because of its flexibility, this system lets investigators screen phenotypes and infer gene functions that may be impossible otherwise.

Indeed, transgenic cargo often confers fitness costs to the arthropod; therefore, Williams et al. laid out simple protocols for measuring fitness costs in transgenic Aedes aegypti mosquitoes¹². These included fecundity, wing size and shape, fertility, sex ratio, viability, development times, male contribution, and adult longevity¹². Such fitness measurements can be used to compare the health of transgenic mosquitoes generated across studies or to model the rate of transgene fixation in mathematical models.

The methodological details in these manuscripts are visually illustrated to facilitate reproduction. A variety of molecular and entomological techniques are presented: sexing and screening mosquito pupae for fluorescent markers^11,12; combining and purifying donor and helper φC31 plasmid mixes¹⁰; injecting adult jewel wasps intra-abdominally⁹; lining up sand fly eggs for microinjection⁷; and ablating tick wax glands⁸. Given the breadth of techniques covered in this special issue, both amateurs and experts are invited to glimpse the underpinnings of generating genetically modified arthropods. These techniques are the foundation for studying arthropod physiology and vector-pathogen interactions, which are essential for developing novel strategies that reduce vector-borne disease.

The authors have nothing to disclose.

This work was funded by the NIH, grant number R01 AI130085, and the NIH Division of Intramural Research Program: AI001246.