Nature Protocols最新文献

The use of saturated small-ring bridged hydrocarbons as bioisosteres for aromatic rings has become a popular tactic in drug discovery. Perhaps the best known of such hydrocarbons is bicyclo[1.1.1]pentane, for which the angle between the exit vectors of the bridgehead substituents is identical to that of a para-substituted arene (180°). The development of meta-arene (bio)isosteres is much less explored due to the challenge of identifying an accurate geometric mimic (substituent exit vector angle ~120°, dihedral angle ~0°). To address this, we recently reported straightforward access to bicyclo[3.1.1]heptanes (BCHeps), which exactly meet these geometric properties, via radical ring-opening reactions of [3.1.1]propellane. This required the development of a scalable synthesis of [3.1.1]propellane, as well as the implementation of various ring-opening reactions and derivatizations. Here we describe methodology for a multigram scale synthesis of [3.1.1]propellane in five steps from commercially available ethyl 4-chlorobutanoate, which proceeds in an overall yield of 26-37%. We also describe the functionalization of [3.1.1]propellane to three key classes of BCHep iodides by photocatalyzed-atom transfer radical addition reactions using 456 nm blue light. We further report protocols for the elaboration of these products to other useful derivatives, via iron-catalyzed Kumada coupling with aryl Grignard reagents and conversion of a pivalate ester to a carboxylic acid through hydrolysis/oxidation. The total times required to synthesize [3.1.1]propellane, the BCHep iodides and the BCHep carboxylic acid are ~53, 6-8 and 40 h, respectively, requiring an average level of synthetic chemistry expertise (for example, masters and/or graduate students).

Deep-scale, mass spectrometry-based proteomic studies by the Clinical Proteomic Tumor Analysis Consortium (CPTAC) program involves tissue lysis using urea buffer before data acquisition via mass spectrometry for quantitative global proteomic and phosphoproteomic analysis. This is described in a 2018 protocol¹. Here we report an update to this initial protocol by implementing a sonication step into urea-based tissue lysis. Similar to the initial CPTAC protocol, we identified >12,000 proteins and >25,000 phosphopeptides in a tandem mass tag (TMT) set containing both nonsonicated and sonicated tumor tissues from patient-derived xenograft mouse models. An improvement in the detection of membrane-bound and DNA-binding proteins was observed by including the sonication. We also offer recommendations for optimal sonication conditions such as the buffer composition, timing of sonication cycle, instrumentation settings and a troubleshooting section for potential users. Additionally, the protocol is equally applicable to other biological specimens.

Connectional neuroanatomical maps can be generated in vivo by using diffusion-weighted magnetic resonance imaging (dMRI) data, and their representation as structural connectome (SC) atlases adopts network-based brain analysis methods. We explain the generation of high-quality SCs of brain connectivity by using recent advances for reconstructing long-range white matter connections such as local fiber orientation estimation on multi-shell dMRI data with constrained spherical deconvolution, which yields both increased sensitivity to detecting crossing fibers compared with competing methods and the ability to separate signal contributions from different macroscopic tissues, and improvements to streamline tractography such as anatomically constrained tractography and spherical-deconvolution informed filtering of tractograms, which have increased the biological accuracy of SC creation. Here, we provide step-by-step instructions to creating SCs by using these methods. In addition, intermediate steps of our procedure can be adapted for related analyses, including region of interest-based tractography and quantification of local white matter properties. The associated software MRtrix3 implements the relevant tools for easy application of the protocol, with specific processing tasks deferred to components of the FSL software. The protocol is suitable for users with expertise in dMRI and neuroscience and requires between 2 h and 13 h to complete, depending on the available computational system.

Advances in genomics have identified thousands of risk genes impacting human health and diseases, but the functions of these genes and their mechanistic contribution to disease are often unclear. Moving beyond identification to actionable biological pathways requires dissecting risk gene function and cell type-specific action in intact tissues. This gap can in part be addressed by in vivo Perturb-seq, a method that combines state-of-the-art gene editing tools for programmable perturbation of genes with high-content, high-resolution single-cell genomic assays as phenotypic readouts. Here we describe a detailed protocol to perform massively parallel in vivo Perturb-seq using several versatile adeno-associated virus (AAV) vectors and provide guidance for conducting successful downstream analyses. Expertise in mouse work, AAV production and single-cell genomics is required. We discuss key parameters for designing in vivo Perturb-seq experiments across diverse biological questions and contexts. We further detail the step-by-step procedure, from designing a perturbation library to producing and administering AAV, highlighting where quality control checks can offer critical go-no-go points for this time- and cost-expensive method. Finally, we discuss data analysis options and available software. In vivo Perturb-seq has the potential to greatly accelerate functional genomics studies in mammalian systems, and this protocol will help others adopt it to answer a broad array of biological questions. From guide RNA design to tissue collection and data collection, this protocol is expected to take 9-15 weeks to complete, followed by data analysis.

Bioelectronic medicine, which involves the delivery of electrical stimulation via implantable electrodes, is poised to advance the treatment of neurological conditions. However, current hand-made devices are bulky, invasive and lack specificity. Thin-film neurotechnology devices can overcome these disadvantages. With a typical thickness in the range of micrometers, thin-film devices demonstrate high conformability, stretchability, are minimally invasive and can be fabricated using traditional lithography techniques. Despite their potential, variability and unreliability in fabrication processes hinder their wider utilization. Here, we detail a fabrication method for thin-film poly(ethylene dioxythiophene):poly(styrene sulfonate) (PEDOT:PSS) electrodes and organic electrochemical transistors. The use of organic materials makes these devices particularly well suited for bioelectronic medicine applications as they show superior mechanical and electrical matching of biological tissues compared with devices made of inorganic materials. The procedure details the entire process, including mask design, the fabrication through three photolithography stages, the integration with larger-scale electronics, implantation procedures and the expected electrical characterization metrics. The nanofabrication protocol requires at least 3 d and is suitable for those familiar with lithographic fabrication procedures. The surgery requires up to 10 h and is suitable for those familiar with in vivo implantation procedures.

Sparse, single-cell labeling approaches enable high-resolution, high signal-to-noise recordings from subcellular compartments and intracellular organelles and allow precise manipulations of individual cells and local circuits while minimizing complex changes associated with global network manipulations. However, thus far, only a limited number of approaches have been developed to label single cells with unique combinations of genetically encoded indicators, target deep cortical structures or sustainably use the same chronic preparation for weeks. Here we developed a method to deliver plasmids selectively to single pyramidal neurons in the mouse dorsal hippocampus using two-photon visually guided in vivo single-cell electroporation to address these limitations. This method allows long-term plasmid expression in a controlled number of individual pyramidal neurons, facilitating subcellular resolution imaging, intracellular organelle tracking, monosynaptic input mapping, plasticity induction and targeted whole-cell patch-clamp recordings.

The rise of cellular immunotherapy for cancer treatment has led to the utilization of immune oncology cocultures to simulate T cell interactions with cancer cells for assessing their antitumor response. Previously, we developed BEHAV3D, a three-dimensional live imaging platform of patient-derived tumor organoid (PDO) and engineered T cell cocultures, that analyzes T cells' dynamics to gain crucial insights into their behavior during tumor targeting. However, live imaging alone cannot determine the molecular drivers behind these behaviors. Conversely, single-cell RNA sequencing (scRNA-seq) allows researchers to analyze the transcriptional profiles of individual cells but lacks spatio-temporal resolution. Here we present an extension to the BEHAV3D protocol, called Behavior-Guided Transcriptomics (BGT), for integration of T cell live imaging data with single-cell transcriptomics, enabling analysis of gene programs linked to dynamic T cell behaviors. BGT uses live imaging data processed by BEHAV3D to guide the experimental setup for cell separation based on their PDO engagement levels subsequently followed by fluorescence-activated cell sorting and scRNA-seq. It then integrates in silico simulations of these experiments to computationally infer T cell behavior on scRNA-seq data, uncovering new biomarkers for both highly functional and ineffective T cells, that could be exploited to enhance therapeutic efficacy. The protocol, designed for users with fundamental cell culture, imaging and programming skills, is readily adaptable to diverse coculture settings and takes one month to perform.

Synaptic connections among neurons are critical for information processing and memory storage in the brain, making them hotspots for neuropathologies. Understanding the physiology of synapses, therefore, may facilitate the development of therapeutic approaches. However, synapses are micrometer-sized functional structures involved in many neuronal processes, where the challenge is deciphering differential signaling in presynaptic and postsynaptic compartments of relatively intact microcircuits. Here we developed a method combining two-photon laser microsurgery with compartment-specific electrophysiological activation and readout to improve the specificity with which neuronal signaling is detected. After finding a connection, femtosecond laser pulses are used to sever the presynaptic axon from the cell body with micrometer precision. This microdissection method is effective to a depth of at least 100 µm. The initial segment of the isolated axon is extracellularly stimulated and activated to release neurotransmitters, as detected via a recipient whole-cell neuron, which is being recorded. This methodology is an alternative to axonal patch-clamp recordings, which are short-lasting and difficult. Together with pharmacology and genetic manipulation, our approach allows the interrogation of compartmentalized signaling in intact synapses. The total time of laser exposure is a few seconds and the microsurgery takes 5-10 min, which enables the interrogation of multiple synapses within an experiment. Our protocol provides a tool to investigate compartment-specific signaling in relatively intact brain tissue, enabling a more comprehensive understanding of neuronal synapses.