Cold Spring Harbor protocols最新文献

Sleep is likely a whole-brain phenomenon, with most of the brain probably benefiting from this state of decreased arousal. Recent advances in our understanding of some potential sleep functions, such as metabolite clearance and synaptic homeostasis, make it evident why the whole brain is likely impacted by sleep: All neurons have synapses, and all neurons produce waste metabolites. Sleep experiments in the fly Drosophila melanogaster suggest that diverse sleep functions appear to be conserved across all animals. Studies of brain activity during sleep in humans typically involve multidimensional data sets, such as those acquired by electroencephalograms (EEGs) or functional magnetic resonance imaging (fMRI), and these whole-brain read-outs often reveal important qualities of different sleep stages, such as changes in frequency dynamics or connectivity. Recently, various techniques have been developed that allow for the recording of neural activity simultaneously across multiple regions of the fly brain. These whole-brain-recording approaches will be important for better understanding sleep physiology and function, as they provide a more comprehensive view of neural dynamics during sleep and wake in a relevant model system. Here, we present a brief summary of some of the findings derived from sleep activity recording studies in sleeping Drosophila flies and discuss the value of electrophysiological versus calcium imaging techniques. Although these involve very different preparations, they both highlight the value of multidimensional data for studying sleep in this model system, like the use of both EEG and fMRI in humans.

Creating transgenic mosquitoes allows for mechanistic studies of basic mosquito biology and the development of novel vector control strategies. CRISPR-Cas9 gene editing has revolutionized gene editing, including in mosquitoes. This protocol details part of the gene editing process of Aedes aegypti mosquitoes via CRISPR-Cas9, through testing and validating single-guide RNAs (sgRNAs). Gene editing activity varies depending on the sequence of sgRNAs used, so validation of sgRNA activity should be done before large-scale generation of mutants or transgenics. sgRNA is designed using online tools and synthesized in <1 h. Once mutants or transgenics are generated via embryo microinjection, sgRNA activity is validated by quick genotyping polymerase chain reaction (PCR) and DNA sequencing.

CRISPR-Cas9 has revolutionized gene editing for traditional and nontraditional model organisms alike. This tool has opened the door to new mechanistic studies of basic mosquito biology as well as the development of novel vector control strategies based on CRISPR-Cas9, including gene drives that spread genetic elements in the population. Although the promise of the specificity, flexibility, and ease of deployment CRISPR is real, its implementation still requires empirical optimization for each new species of interest, as well as to each genomic target within a given species. Here, we provide an overview of designing and testing single-guide RNAs for the use of CRISPR-based gene editing tools.

Analysis of taste sensory responses has been a powerful approach for understanding principles of taste detection and coding. The shared architecture of external taste sensing units, called sensilla, in insects opened up the study of tastant-evoked responses in any model of choice using a single-sensillum tip recording method that was developed in the mid-1900s. Early studies in blowflies were instrumental for identifying distinct taste neurons based on their responses to specific categories of chemicals. Broader system-wide analyses of whole organs have since been performed in the genetic model insect Drosophila melanogaster, revealing principles of stereotypical organization and function that appear to be evolutionarily conserved. Although limited in scope, investigations of taste sensory responses in mosquitoes showcase conservation in sensillar organization, as well as in groupings of functionally distinct taste neurons in each sensillum. The field is now poised for more thorough dissections of mosquito taste function, which should be of immense value in understanding close-range chemosensory interactions of mosquitoes with their hosts and environment. Here, we provide an introduction to the basic structure of a taste sensillum and functional analysis of the chemosensory neurons within it.

In insects, gustatory neurons sense chemicals upon contact and directly inform many behaviors critical for survival and reproduction, including biting, feeding, mating, and egg laying. However, the taste sensory system is underexplored in many anthropophilic disease vectors such as mosquitoes, which acquire and transmit human pathogens during blood feeding from human hosts. This results in a big gap in vector biology-the study of organisms that spread disease by transmitting pathogens-because insect vectors closely interact with humans while selecting suitable individuals and appropriate bite sites for blood meals. Human sweat and skin-associated chemistries are rich in nonvolatile compounds that can be sensed by the mosquito's taste system when she lands on the skin. Taste sensory units, called sensilla, are distributed in many organs across the mosquito body, including the mouthparts, legs, and ovipositors (female-specific structures used to lay eggs). Each sensillum is innervated by as many as five taste neurons, which allow detection and discrimination between various tastants such as water, sugars, salts, amino acids, and plant-derived compounds that taste bitter to humans. Single-sensillum recordings provide a robust way to survey taste responsiveness of individual sensilla to various diagnostic and ecologically relevant chemicals. Such analyses are of immense value for understanding links between mosquito taste responses and behaviors to specific chemical cues and can provide insights into why mosquitoes prefer certain hosts. The results can also aid development of strategies to disrupt close-range mosquito-human interactions to control disease transmission. Here we describe a protocol that is curated for electrophysiological recordings from taste sensilla in mosquitoes and sure to yield exciting results for the field.

Sleep studies in Drosophila melanogaster rely mostly on behavioral read-outs to support molecular or circuit-level investigations in this model. Electrophysiology can provide an additional level of understanding in these studies to, for example, investigate changes in brain activity associated with sleep manipulations. In this protocol, we describe a procedure for performing multichannel local field potential (LFP) recordings in the fruit fly, with a flexible system that can be adapted to different experimental paradigms and situations. The approach uses electrodes containing multiple recording sites (16), allowing the acquisition of large amounts of neuronal activity data from a transect through the brain while flies are still able to sleep. The approach starts by tethering the fly, followed by positioning it on an air-supported ball. A multichannel silicon probe is then inserted laterally into the fly brain via one eye, allowing for recording of electrical signals from the retina through to the central brain. These recordings can be acquired under spontaneous conditions or in the presence of visual stimuli, and the minimal surgery promotes long-term recordings (e.g., overnight). Sleep and wake can be tracked using infrared cameras, which allow for the measurement of locomotive activity as well as microbehaviors such as proboscis extensions during sleep. The protocol has been optimized to promote subject survivability, which is an important factor when performing long-term (∼16-h) recordings. The approach described here uses specific recording probes, data acquisition devices, and analysis tools. Although it is expected that some of these items might need to be adapted to the equipment available in different laboratories, the overall aim is to provide an overview on how to record electrical activity across the brain of behaving (and sleeping) flies using this kind of approach and technology.

Genetically encoded calcium indicators (GECIs) allow for the noninvasive evaluation of neuronal activity in vivo, and imaging GECIs in Drosophila has become commonplace for understanding neural functions and connectivity in this system. GECIs can also be used as read-outs for studying sleep in this model organism. Here, we describe a methodology for tracking the activity of neurons in the fly brain using a two-photon (2p) microscopy system. This method can be adapted to perform functional studies of neural activity in Drosophila under both spontaneous and evoked conditions, as well as during spontaneous or induced sleep. We first describe a tethering and surgical procedure that allows survival under the microscopy conditions required for long-term recordings. We then outline the steps and reagents required for optogenetic activation of sleep-promoting neurons while simultaneously recording neural activity from the fly brain. We also describe the procedure for recording from two different locations-namely, the top of the head (e.g., to record mushroom body calyx activity) or the back of the head (e.g., to record central complex activity). We also provide different strategies for recording from GECIs confined to the cell body versus the entire neuron. Finally, we describe the steps required for analyzing the multidimensional data that can be acquired. In all, this protocol shows how to perform calcium imaging experiments in tethered flies, with a focus on acquiring spontaneous and induced sleep data.

The Drosophila embryonic central nervous system has been used for decades as a model for understanding the genetic regulation of axon guidance and other aspects of neural development. Foundational studies using antibody staining to examine the embryonic ventral nerve cord in wild-type and mutant animals led to the discovery of evolutionarily conserved genes that regulate fundamental aspects of axon guidance, including midline crossing of axons. The development of the regular, segmentally repeating structure of axon pathways in the ventral nerve cord can illustrate basic principles of axon guidance to beginning students and can also be used by expert researchers to characterize new mutants, detect genetic interactions between known genes, and precisely quantify variations in gene function in engineered mutant lines. Here, we describe a protocol for collecting and fixing Drosophila embryos and visualizing axon pathways in the embryonic ventral nerve cord using immunofluorescence or immunohistochemical staining methods. As embryogenesis in Drosophila takes ∼24 h to complete, a 1-d collection yields embryos representing all stages of development from newly fertilized through ready-to-hatch larvae, allowing investigation of multiple developmental events within a single batch of collected embryos. The methods described in this protocol should be accessible to introductory laboratory courses as well as seasoned investigators in established research laboratories.

The technique of visualizing axon pathways in the embryonic ventral nerve cord using antibody labeling has been fundamental to our understanding of the genetic and developmental mechanisms underlying nervous system wiring in Drosophila. High-resolution microscopic examination of the ventral nerve cord remains an essential component of many experiments in Drosophila developmental neuroscience. Although it is possible to examine the ventral nerve cord in intact whole-mount embryos, to collect the highest-quality images it is often useful to isolate the nervous system away from the other embryonic tissues through embryo dissection. This protocol describes methods for dissecting ventral nerve cords from Drosophila embryos that have been fixed and stained via immunofluorescence or horseradish peroxidase (HRP) immunohistochemistry. The process of making fine dissection needles for this purpose from electrolytically sharpened tungsten wire is also described. Dissected and mounted ventral nerve cords can be examined and imaged using a variety of microscopy techniques including differential interference contrast (DIC) optics, epifluorescence, or confocal microscopy.

The establishment of neural connectivity is a major part of neural development. The central nervous system (CNS) midline is the most characterized axon guidance choice point, and work in Drosophila has played a pivotal role in understanding the molecular mechanisms responsible. Axons respond to attractive cues such as Netrin via the Frazzled receptor, and repulsive cues such as Slit via Robo receptors. Both signals are expressed at the CNS midline, affect pioneer axons, and have dramatic effects on the axon scaffold as a whole. Here, we focus on previous research analyzing classic mutants in the Slit/Robo pathway, which can readily be detected with a dissecting microscope. We also discuss analyzing these mutants in a teaching lab situation. The combination of sophisticated genetics and reliable axonal markers in Drosophila allows phenotypic analysis to be performed at the single-cell level. The elaborate architecture of neurons is very sensitive to disruption by genetic mutations, allowing the effects of novel mutations to be easily detected and assessed.