Cold Spring Harbor protocols最新文献

Mosquito species vary in their host associations. Although some species are relative generalists, most specialize, to varying extents, on particular types of host animals. Mosquito host associations are among the most important factors that influence the transmission dynamics of mosquito-vectored pathogens, and understanding these associations can provide insight on how such pathogens move within ecosystems. Characterization of the host associations of mosquito species requires applying blood meal analysis to the largest possible sample size of mosquito blood meals. Processing large samples of mosquito blood meals can be time-consuming, especially when chain-termination sequencing is used, necessitating individual processing of each specimen. Various methods and commercially available kits and products are available for extracting DNA from mosquito blood meals. The hot sodium hydroxide and Tris (HotSHOT) method is a rapid and inexpensive method of DNA extraction that is compatible with the recovery of DNA from mosquito blood meals preserved on QIAcard Flinders Technology Associates (FTA) Classic Cards (FTA cards). FTA cards allow nucleic acids found in blood meals to be preserved easily, even in field conditions. DNA prepared using this method is suitable for polymerase chain reaction (PCR)-based blood meal analysis.

Maize genetic transformation is a critical tool for functional genomics and crop improvement. Many laboratories, however, continue to face multiple challenges in attempting to achieve routine genetic transformation of maize inbred genotypes. Here, we describe a rapid and robust maize B104 transformation method using immature embryos as explants. This method uses an Agrobacterium ternary vector system, which includes a conventional T-DNA binary vector (pCBL101-RUBY) and a compatible ternary helper plasmid (pKL2299) that carries extra copies of essential virulence genes. The T-DNA binary vector carries the neomycin phosphotransferase II (NptII) gene for selection and a betalain biosynthesis marker, RUBY, for visual screening. We provide step-by-step instructions for immature embryo explant preparation, Agrobacterium infection, tissue culture procedures, and greenhouse care for acclimatization of regenerated plantlets.

The introduction of maize genetic transformation in the 1990s brought forth a powerful tool for crop improvement and a deeper understanding of plant genetics. Despite decades of genetics research, however, and the promise of CRISPR-mediated gene editing, maize transformation currently faces several challenges, such as genotype dependence and limitations in explant availability. Indeed, although the most commonly used method, immature embryo transformation, has been improved through optimization of tissue culture media composition and selection methods, the approach is only applicable to a limited number of public genotypes, including B104 and Hi II. Recently, genotype-flexible methods have been developed using coexpression cassettes of morphogenic transcription factors (MTFs) Baby boom (Bbm) and Wushel2 (Wus2), which have enabled the successful transformation of many previously recalcitrant maize lines. This MTF-based transformation method has also allowed for the use of alternate explants, such as seedling leaf whorl, whose production is cost-effective and requires only minimum controlled growth space. In this review, we summarize recent advances in Agrobacterium-mediated maize transformation methods that use immature embryos or seedling leaf whorls as starting material.

Conventional maize transformation has largely relied on immature embryos as explants, and is thus often hampered by the limited access to high-quality immature embryos year-round. Here, we present a detailed protocol using seedling leaf whorls as alternative explants for tropical maize inbred transformation. This approach involves the use of a cassette that drives the expression of the morphogenic transcription factors (MTFs) Baby boom (Bbm) and Wuschel2 (Wus2), which have been shown to greatly enhance transformation efficiency. We outline here the steps required for the preparation of seedling leaf whorl explants and subsequent Agrobacterium infection, and describe the tissue culture regimen that results in transgenic plant regeneration. Because constitutive expression of Bbm and Wus2 prevents normal plant regeneration and the production of fertile plants, the cassette containing these genes must be excised. As such, we include the steps for the Cre/loxP-mediated excision of the MTF gene cassette. The protocol outlines a year-round, more affordable, and efficient approach for carrying out maize transformation for crop improvement.

Larvae of the fruit fly Drosophila melanogaster are a popular and tractable model system for studying the development and function of sensorimotor circuits, thanks to the relative numerical simplicity of their nervous system and the wealth of available genetic tools to manipulate the anatomy, activity, and function of specific cell types. Researchers studying the role of a particular gene or cell type in sensorimotor circuit activity or function may wish to observe the effects of an experimental manipulation during behavior in the intact animal. Observing these effects, which may include changes in the intracellular calcium concentration or movement of small numbers of neurons, muscles, etc., typically requires high-spatial-resolution imaging, which poses several difficulties in the freely crawling larva. Freely crawling larvae can move quickly and with changeable heading, making manual or automatic tracking challenging; additionally, they may make three-dimensional movements, such as rearing, that can degrade imaging focus. These challenges are potentially solvable using advanced imaging and algorithmic tracking setups, but cost, space, or development time may be prohibitive. This protocol describes a simple and cost-effective method for placing larvae inside agarose channels, thereby restricting larval crawling to a single dimension and enabling higher-magnification time-series imaging of fluorescently labeled structures during many cycles of locomotion. By using larvae that express fluorescent calcium indicators in cells of interest, researchers can apply this method to study the effects of experimental manipulations on neural or muscular activity during behavior in the intact animal.

In animals, movement is generated by the activity of motor circuits housed in the vertebrate spinal cord or the arthropod nerve cord. How motor circuits form is a fundamental question, with wide-ranging impacts on the fields of development, neurobiology, medicine, evolution, and beyond. Until recently, studying circuit assembly had been experimentally difficult, with a paucity of suitable models. Due to the introduction of novel neuroscience tools (calcium imaging, optogenetics, connectomics), Drosophila embryos and larvae can be used as models to study motor circuit assembly. Here, we briefly review the knowledge relevant to motor circuit assembly in Drosophila larvae. We discuss the larval body and its movements, larval neurons and circuits in the motor system, and how the generation of neural diversity starting from stem cells relates to circuit formation. The long-term goal of Drosophila research in this field is to identify developmental rules, determine when the rules apply, generate an integrated understanding of motor circuit development, and uncover molecular mechanisms driving the assembly process. Motor circuits are an ancient part of the nervous system, and so far, the developmental programs guiding motor circuit assembly appear to be largely conserved across phyla. Thus, as methods improve in other systems, findings in Drosophila will provide foundational concepts that will inspire hypotheses in those systems.

In the Drosophila nerve cord, much is known about the generation of neurons from neuronal stem cells. Over the lifetime of a neuron, the cumulative expression of genes within that neuron determines its fate. Furthermore, gene expression in mature neurons determines their functional characteristics. It is therefore useful to visualize neural gene expression, which is often done via staining with antibodies to a protein of interest. In cases where there is no antibody to a desired gene product, or when it is useful to detect RNA rather than protein products, fluorescent in situ hybridization chain reaction for RNA (HCR RNA-FISH, or HCR for this protocol) can be used to detect and quantify RNA expression. RNA molecules reside predominantly in the cell soma, so HCR can facilitate determining neuron identity because somata position within the nerve cord is stereotyped across animals. HCR provides high-amplitude, high-fidelity signals. In principle, HCR can be broken down into a detection/hybridization stage and an amplification stage. During detection/hybridization, a probe set hybridizes to multiple sequences within a target gene. In the amplification step, concatemerized fluorescent hairpins bind to the hybridized probes. This two-step process increases the specificity of the fluorescent signal and helps reduce the likelihood of background fluorescence compared to traditional in situ hybridization techniques where the hybridizing probe itself contains the fluorescent signal. Here, we describe a protocol for using HCR to study gene expression in the Drosophila embryonic and larval nerve cord. We also describe how to combine HCR with immunofluorescence staining.

Neurons exhibit some of the most striking examples of morphological diversity of any cell type. Thus, when studying neurons, the morphology of each neuron must be considered individually. However, neurons densely populate the central nervous system (CNS), making it difficult to ascertain fine morphological features due to a lack of spatial resolution. In Drosophila, this problem can be partially resolved by using driver lines that express the yeast transcription factor GAL4 in subsets of neurons. GAL4 can activate the expression of other introduced genetic elements such as genes for fluorescent proteins or other markers under the control of the GAL4 upstream activation sequences (UAS effectors). However, even highly specific GAL4 lines often label sets of potentially morphologically heterogeneous neurons. Here, we describe a protocol for using the multicolor flip-out (MCFO) technique in Drosophila melanogaster to stochastically label individual neurons within a GAL4 expression pattern. MCFO relies on the binary GAL4/UAS expression system in Drosophila but adds additional control for how densely the neurons within a GAL4 expression pattern are labeled via user-controlled heat shock. Specifically, three discrete UAS effector elements containing the sequences for unique epitope tags (FLAG, HA, and V5) linked to a gene for nonfluorescent GFP can be independently expressed under the control of GAL4 only when a transcriptional stop sequence in the UAS promoter sequence has been removed by heat shock-induced recombination. This effectively labels multiple individual neurons with either one or a combination of epitope tags that can be spectrally resolved with immunofluorescence. The MCFO technique is ideal for researchers who want to determine morphological features of CNS neurons in wild-type or mutant backgrounds.

The Drosophila larval nerve cord, which is the equivalent of the vertebrate spinal cord, houses the circuits required to process somatosensory stimuli (e.g., tactile, temperature, vibration, and self-movement) and generate the patterned muscle contractions underlying movement and behavior. Within this complex structure reside many cell types and cellular processes, making it difficult to experimentally access, when compared to peripheral parts of the nervous system (i.e., primary sensory neuron dendrites, motor neuron axons and synapses, and muscles). Additionally, the neurons in the larval nerve cord have small cell bodies, precluding traditional electrophysiological approaches. As such, the function of neurons in the nerve cord is less well studied than other parts of the nervous system, severely limiting our understanding of how larvae process sensory information and generate movement. Ca²⁺-sensitive fluorescent proteins enable the study of neuronal activity in live, genetically tractable animals, even those with small neuronal cell bodies. In addition, live imaging of neurons within the nerve cord in whole, intact animals is possible because larvae are translucent, and the use of intact animals allows for the peripheral sensory neuron circuits to remain intact. Ca²⁺-sensitive fluorescent proteins increase their fluorescence when voltage-gated Ca²⁺ channels are opened in depolarized neurons. Here, we describe an assay where a Ca²⁺-sensitive fluorescent protein (GCaMP6m) is expressed under the control of a GAL4 driver in a subset of neurons that reside in a circuit for vibration sensation. External vibration (sound) stimulates sensory neurons that activate the cells expressing the Ca²⁺-sensitive fluorescent protein. Visualization of the calcium-induced fluorescent signal with microscopy allows for quantification of neuronal activity.

The BonnMu resource represents a tagged collection of maize (Zea mays L.) Mutator (Mu) transposon-induced mutants, designed for functional genomics studies. Here, we describe the use of the BonnMu collection for identifying and characterizing mutations. Specifically, we describe workflows for use in both reverse and forward genetics strategies in maize. For reverse genetics, users first acquire a BonnMu F₂ stock of interest based on data accessible at the Maize Genetics and Genomics Database (MaizeGDB). We provide details here for their subsequent propagation and for the confirmation of Mu insertions by genotyping via PCR, with the ultimate goal of establishing genotype-phenotype relationships of interest. For forward genetics studies, we describe a workflow that involves a combined approach of Mutant-Seq (Mu-Seq) and bulked segregant RNA-seq (BSR-Seq), to identify the causal gene underlying a mutant phenotype of interest.