Cold Spring Harbor protocols最新文献

To understand the circuitry of the brain, it is often advantageous to visualize the processes of a single neuron or population of neurons. Identifying sites where a neuron, or neurons, originates and where it projects can allow a researcher to begin to map the circuitry underlying various processes, including sensory-guided behaviors. Furthermore, neural tracing allows one to map locations where processes terminate onto regions of the brain that may have known functions and sometimes to identify candidate upstream or downstream connections, based on proximity. Many methods of neural tracing are available; here, we focus on loading fluorescent dyes into a neuron (fluorescent dye filling). Different options for dyes exist to label neurites. Among the most versatile and easy to use are dextran amine-conjugated dyes. They fill neurons bidirectionally, not discriminating between anterograde or retrograde loading direction. Dye filling must be done in unfixed tissue, as the dye needs to move through the neurons; however, dextran amine conjugates are aldehyde-fixable and once cells have been fully loaded with dye the tissue can be fixed and subjected to immunostaining. Coupling neural tracing with immunofluorescence is a useful way to determine specific brain or ventral nerve cord (VNC) regions where a neuron projects. This protocol describes methods for loading dextran amine conjugated dyes into a sensory tissue in the mosquito to visualize sites of sensory neuron innervation in the central nervous system, as well as efferent projections to these structures. This protocol is described for Aedes aegypti, for which it was optimized, but it also works across a variety of insects.

Mosquito-borne disease is a major global public health issue. One path toward the development of evidence-based strategies to limit mosquito biting is the study of the mosquito nervous system-in particular, the sensory systems that drive biting behavior. The central nervous system of insects consists of the brain and the ventral nerve cord. Here, we describe a protocol for dissecting, immunofluorescent labeling, and imaging both of these structures in the mosquito. This protocol was optimized for Aedes aegypti and works well on Anopheles gambiae tissue. It has not been tested in other mosquito species, but we anticipate that it would work on a range of mosquitoes, and, if not, our protocol will provide a starting point from which to optimize. Notably, a limited number of antibodies cross-react with Ae. aegypti proteins. This protocol is intended for use with validated antibodies and can also be used to test new antibodies as they are generated. It has been successfully used to visualize protein tags, such as green fluorescent protein, that have been introduced into the mosquito to amplify or detect their presence.

A researcher may have many reasons for wanting to establish new laboratory colonies from field-collected mosquitoes. In particular, the ability to study the diversity found within and among natural populations in a controlled laboratory environment opens up a wide range of possibilities for understanding how and why burdens of vector-borne disease vary over space and time. However, field-collected mosquitoes are often more difficult to work with than established laboratory strains, and considerable logistical challenges are involved in safely transporting field-collected mosquitoes into the laboratory. Here, we provide advice for researchers working with Aedes aegypti, Anopheles gambiae, and Culex pipiens, as well as notes on other closely related species. We provide guidance on each stage of the life cycle and highlight the life stages for which it is easiest to initiate new laboratory colonies for each species. In accompanying protocols, we provide methods detailing Ae. aegypti egg collection and hatching as well as how to transport larvae and pupae from the field.

Mosquitoes transmit deadly pathogens from person to person as they obtain the blood meal that is essential for their life cycle. Female mosquitoes of many species are unable to reproduce without consuming protein that they obtain from blood. This developmental stage makes them highly efficient disease vectors of deadly pathogens. They can transmit pathogens between members of the same species and different species that can provide a route for evolving zoonotic viruses to jump from animals to humans. One possible way to develop novel strategies to combat pathogen transmission by mosquitoes is to study the sensory systems that drive mosquito reproductive behaviors, in particular the neural architecture and circuits of mosquito sensory afferent neurons, the central circuits that process sensory information, and the downstream circuits that drive reproductive behaviors. The study of mosquito neuroanatomy and circuitry also benefits basic neuroscience, allowing for comparative neuroanatomy in insect species, which has great value in the current model species-heavy landscape of neuroscience. Here, we introduce two important techniques that are used to study neuroanatomy and neural circuitry-namely, immunofluorescent labeling and neural tracing. We describe how to apply these approaches to study mosquito neuroanatomy and describe considerations for researchers using the techniques.

Laboratory study of field-collected mosquitoes can allow researchers to better understand the ways variation within and among mosquito populations shapes burdens of mosquito-borne disease. The Anopheles gambiae complex comprises the most important vectors of malaria, but it can be challenging to keep in the laboratory. For some species of mosquitoes, especially An. gambiae, it is very difficult to bring viable eggs into the laboratory. Instead, it is preferable to collect larvae or pupae and then transport them as carefully as possible back to the laboratory. This simple protocol allows a researcher to start new laboratory colonies from larvae or pupae collected from natural breeding sites or proceed directly to their planned experiments. The use of natural breeding sites provides additional reassurance that the resulting colonies are representative of natural populations.

Genetically modified (GM) mosquitoes are an important tool in the fight against mosquito-borne disease, both indirectly through their use in research investigating host-pathogen interaction, mosquito olfaction, and anthropomorphic behavior and in future direct uses for suppression and possibly eradication through sterile insect technique (SIT) and/or gene-drive programs. Successful creation of GM mosquitoes depends on microinjection procedures that precisely deliver injection materials while causing as little damage to mosquito embryos as possible. Genetic modification reagents, such as transposon system components (vector plasmids, helper plasmids, and helper mRNA), and CRISPR-Cas9 components (guide RNAs, Cas9 protein, plasmids expressing Cas9 and/or guide RNAs, and donor plasmids used in homology-directed repair [HDR]), must be delivered into the preblastoderm embryo at the posterior end where the pole cells will form before cellularization occurs. Sharp needles that pierce the embryo easily are important tools in this procedure and work best when the embryos are not desiccated. The two main procedures for mosquito embryo microinjection involve injecting embryos under halocarbon oil or under aqueous solution.

In a closed-loop experimental paradigm, an animal experiences a modulation of its sensory input as a function of its own behavior. Tools enabling closed-loop experiments are crucial for delineating causal relationships between the activity of genetically labeled neurons and specific behavioral responses. We have recently developed an experimental platform known as "Raspberry Pi Virtual Reality" (PiVR) that is used to perform closed-loop optogenetic stimulation of neurons in unrestrained animals. PiVR is a system that operates at high temporal resolution (>30-Hz) and with low latencies. Larvae of the fruit fly Drosophila melanogaster are ideal to study the role of individual neurons in modulating behavior to aid the understanding of the neural pathways underlying various guided behaviors. Here, we introduce larval chemotaxis as an example of a navigational behavior in which an animal seeks to locate a target-in this case, the attractive source of an odor-by tracking a concentration gradient. The methodologies that we describe here combine the use of PiVR with the study of larval chemotaxis in real and virtual odor gradients, but these can also be readily adapted to other sensory modalities.

The process of genetically modifying mosquitoes requires skilled delivery of reagents for modification. Plasmids, RNA, DNA, and/or protein must be transported into the developing embryo during an appropriate time in development when these agents will have access to the genome. Embryo microinjection has been the main method by which such modifying agents have been delivered. Ideally the microinjection process will deliver these modifying agents in sufficient quantity to effect the genetic modification without severely damaging or killing the injected embryo in the process. As semiaquatic insects, mosquitoes have embryos that are susceptible to desiccation and the degree to which embryos are susceptible is based on species. Two microinjection methods are outlined here. The first method describes embryo microinjections performed under Halocarbon-27 oil. The oil is used to reduce desiccation during the injection process. A second method limits desiccation by injecting the mosquito embryos in water. In both procedures, the embryos are first aligned and then injected before the embryos cellularize, ∼1 h and 45 min after oviposition.