Nature Protocols最新文献

Bioinspired in-sensor computing devices can process information at sensory terminals by leveraging physical principles, thereby reducing latency and energy consumption during computation while simultaneously enhancing the efficiency of data processing and real-time analysis. Optoelectronic devices exhibit in-sensor computing functions, such as feature enhancement and data compression, by tuning the defect states of the semiconductor channels and thereby modulating the photoresponsivity and time constants of the sensors. These functionalities are critically dependent on precise fabrication and testing protocols. Here we present a detailed procedure for fabricating and characterizing in-sensor computing devices based on nanoscale semiconductor thin films. We explain how to test such optoelectronic devices, including the testing of visual adaptation and motion perception responses. When using semiconductor materials obtained from commercial suppliers, this procedure is time efficient and results in highly reproducible device performance. Nevertheless, all device fabrication and testing steps are generalizable and can be extended to other semiconductor thin films grown using different methods. The procedure is intended for researchers experienced in cleanroom operations and microfabrication techniques and can be completed in ~14 d. The use of bioinspired optoelectronic devices enables the development of a framework for advancing in-sensor computing technologies.

Structural variants (SVs) contribute significantly to genomic diversity and disease predisposition as well as development in diverse species. However, their accurate characterization has remained a challenge because of their complexity and size. With the rise of third-generation sequencing technology, analytical strategies to map SVs have been revisited, and software such as NanoVar, a free and open-source package designed for efficient and reliable SV detection in long-read sequencing data, has facilitated their studies. NanoVar has been shown to work effectively in various published genomic studies, including research on genetic disorders, population genomics and genome analysis of non-model organisms. In this article, we describe in detail all the steps of the NanoVar protocol and its interplay with other platforms for SV calling in whole-genome long-read sequencing data such that researchers with minimal experience with command-line interfaces can easily carry out the protocol. It also provides exhaustive instructions for diverse study designs, including single-sample analyses, cohort studies and genome instability analyses. Finally, the protocol covers SV visualization, filtering and annotation details. Overall, users can identify and analyze SVs in a typical human dataset with a conventional computational setup in ~2-5 h after read mapping.

Controlled gene expression programs have a crucial role in shaping cellular functions and activities. At the core of this process lies the RNA life cycle, ensuring protein products are synthesized in the right place at the right time. Here we detail an integrated protocol for imaging-based highly multiplexed in situ profiling of spatial transcriptome using antibody-based protein comapping (STARmap PLUS), spatial translatome mapping (RIBOmap) and spatiotemporal transcriptome mapping (TEMPOmap). These methods selectively convert targeted RNAs, ribosome-bound mRNAs or metabolically labeled RNAs to DNA amplicons with gene-unique barcodes, which are read out through in situ sequencing under a confocal microscope. Compared with other methods, they provide the analytical capacity to track the spatial and temporal dynamics of thousands of RNA species in intact cells and tissues. Our protocol can be readily performed in laboratories experienced in working with RNA and equipped with confocal microscopy instruments. The wet lab experiments in preparing the amplicon library take 2-3 d, followed by variable sequencing times depending on the sample size and target gene number. The spatially resolved single-cell profiles enable downstream analysis, including cell type classification, cell cycle identification and determination of RNA life cycle kinetic parameters through computational analysis guided by the established tutorials. This spatial omics toolkit will help users to better understand spatial and temporal RNA dynamics in heterogeneous cells and tissues.

Photoluminescence imaging is valuable for elucidating biological processes and diagnosing diseases, but its tissue penetration is limited. We developed an imaging technique that utilizes ultrasound to activate the piezoelectric effect of a molecular probe, transforming ultrasound energy into chemical energy. The chemical energy is then converted into light emission through the chemiluminescence effect, improving penetration depth and overcoming traditional photoluminescence imaging constraints. Here we describe how to build two kinds of ultrasound-induced luminescence imaging systems. We introduce a procedure for the synthesis of trianthracene derivative (TD) nanoparticles with ultrasound-induced luminescence properties. The TDs are converted into water-soluble nanoparticles by a simple nanoprecipitation method. Utilizing the constructed ultrasound-induced luminescence imaging systems, TD nanoparticles can be stimulated to exhibit a luminescence spectrum peaking between 625 and 650 nm. Under optimized ultrasound excitation time and excitation power density parameters, the imaging quality and tissue penetration depth are effectively enhanced. Notably, our procedure enables the detection of both subcutaneous tumor models and challenging deep-tissue orthotopic gliomas. This ultrasound-mediated approach represents an important advancement over conventional photoluminescence imaging methods, enabling high-fidelity in vivo tumor imaging with superior signal quality. Establishment of the ultrasound-induced luminescence imaging systems requires ~2 h, the synthesis of TD molecules requires ~4 d, nanoparticle preparation requires ~1 d, ex vivo characterization requires ~1 d, investigation of the ultrasound-induced luminescence of TD nanoparticles requires ~3 d and ultrasound-induced luminescence imaging takes ~1 d. These steps can be performed by operators trained in chemical synthesis, nanomaterial synthesis standards and qualified in relevant animal experiments.

CRISPR-Cas9 technology has transformed the study of gene function, enabling the systematic investigation of host-virus interactions. However, most CRISPR-based screens in the context of viral infections rely on cell survival as a readout, which limits their sensitivity and biases results toward early infection stages. To address these challenges, we developed the virus-encoded CRISPR-based direct readout system (VECOS), a virus-centric approach in which human cytomegalovirus is engineered to express single-guide RNA (sgRNA) libraries directly from its genome. This system allows sgRNA abundance, embedded in the viral genome, to serve as a direct and quantitative readout of gene-perturbation effects on viral propagation. By tracking sgRNA levels at distinct stages of the viral infection cycle, VECOS enables a detailed, multidimensional analysis of virus-host interactions. Here we present a modular detailed Protocol for (1) constructing and reconstituting complex sgRNA libraries in double-stranded DNA viruses using bacterial artificial chromosomes, (2) performing multipassage screens to investigate perturbation effects on various stages of viral infection and (3) analyzing the multipassage and multistage sgRNA abundance measurements utilizing a comprehensive framework for data analysis. Successful implementation of this full Protocol takes 14-22 weeks and requires proficiency in molecular biology, as well as basic familiarity with Unix-based computing and programming in R for data processing. This Protocol offers researchers a robust tool for uncovering the molecular mechanisms that drive viral propagation and host-virus interactions.

A critical component of genetic code expansion applications is an aminoacyl-tRNA synthetase (RS)/tRNA pair that faithfully encodes a noncanonical amino acid (ncAA) in response to a specific codon. Here we detail a procedure to select an ncAA-specific RS from a publicly available 3.2-million-member Methanomethylophilus alvus pyrrolysyl-RS (MaPylRS) active site mutant library. Four main parts of the procedure are: (1) preparing the library for use and creating needed cell lines; (2) life and death selections that, respectively, select for functional RSs and select against RSs that incorporate canonical amino acids; (3) three fluorescence-based status checks that provide information about the efficiency and fidelity of the surviving RSs in incorporating the target ncAA; and (4) characterizing top hits to find the best ones for use in applications. The resulting RS/tRNA pairs can be used in either bacterial or eukaryotic cells to study proteins of interest. Additionally, the stability of the MaPylRSs makes them useful in cell-free ncAA-protein expression and amenable to structural and other in vitro characterizations. This Protocol is usable by those with basic molecular biology expertise and features a reliable positive control scheme for selections, status checks at different stages to interpret the level of success and a robust procedure to characterize newly engineered tRNA-RS pairs. Users of this Protocol can expect to select ncAA-specific RS/tRNA pairs from the library within about 30-50 d depending on preparation needs.

Aspiration-assisted bioprinting (AAB) is a versatile biofabrication technique that enables the precise and selective patterning of biologics, such as tissue spheroids and organoids, addressing limitations of conventional bioprinting techniques. AAB facilitates the fabrication of (1) tissues with physiologically relevant cell densities using spheroids and (2) advanced tissue models that replicate three-dimensional microenvironments essential for studying cellular responses, disease development and drug testing. Here we provide reliable and reproducible guidelines for the precise positioning of abovementioned biologics, incorporating two operational modes: (1) a single-nozzle mode for precise, one-by-one bioprinting and (2) a high-throughput mode using a digitally controllable nozzle array, enabling the rapid and simultaneous placement of multiple spheroids for scalable tissue fabrication. Comprehensive instructions are included for setting up the AAB platform, operating software and key operational procedures, including optimization of bioprinting conditions. This Protocol enables users to build and operate their own AAB platform depending on target applications, achieving fine control over spheroid positioning through successful aspiration and their precise placement under optimized conditions. This Protocol enables the setup of the AAB platform within 1-2 d. Bioprinting time varies depending on the number of spheroids to bioprint: the single-nozzle mode requires ~30 s per spheroid, while the high-throughput mode can print 64 spheroids in 3-4 min. Designed for accessibility and adaptability, this Protocol is suitable for users from a variety of backgrounds, including engineering, biology, pharmacy and medical sciences, who require bioprinting of spheroids for creating microphysiological systems for drug testing and disease modeling and implantable grafts for regenerative medicine.

The majority of the mammalian genome is transcribed into RNAs, most of which are noncapped RNAs (napRNAs) that not only regulate diverse biological processes through their functions as noncoding RNAs but also serve as processing products to delineate specific RNA biogenesis pathways. However, due to their heterogeneous lengths, diverse terminal modifications and complex secondary structures, identifying these napRNAs poses substantial challenges. Recently, we developed a napRNA sequencing technique (NAP-seq) to identify full-length sequences of napRNAs with various terminal modifications at single-nucleotide resolution. Here we describe the experimental design principles and detailed step-by-step procedures for discovering napRNAs across multiple cell types. The procedure includes T4 polynucleotide kinase pretreatment to standardize RNA termini, enabling comprehensive capture of modified napRNAs; size-selection followed by depletion of known high-abundance RNAs via RNase H to enrich long and low-abundance RNAs; and use of custom-designed adapters with random barcodes, permitting identification of full-length napRNAs at single-nucleotide resolution while minimizing PCR biases and adapter ligation inefficiencies. The use of thermally stable reverse transcriptase enzymes and nested reverse transcriptase primers ensures full-length cDNA synthesis across structured or modified RNA regions while minimizing mispriming artifacts. Libraries are sequenced in parallel using Oxford Nanopore (long-read) and Illumina (short-read) platforms, synergizing advantages of third-generation and next-generation sequencing technologies. The entire experimental procedure, from library preparation to deep sequencing and computational analysis, can be completed within 8 d. The NAP-seq approach enables researchers to discover novel classes of noncoding RNAs with regulatory functions and to investigate RNA biogenesis in various tissues and cell lines.

Hydrogels, as 3D cross-linked hydrophilic networks that exhibit favorable flexibility, cargo loading and release abilities and structure and function designability, are desirable for diverse biomedical applications. For in vivo implementation, however, hydrogels often suffer from swelling-weakened mechanical strength, uncontrollable cargo release and complex composition, inevitably hindering further translation. Despite different reported synthetic approaches, the development of a facile yet universal method capable of fabricating hydrogels with dynamically adjustable structure and function remains difficult. Recently, inspired by biological tissues, we have developed a versatile biological membrane hybridization strategy to generate structurally and functionally programmable hydrogels. Specifically, biological membranes are used as a cross-linker to form a cross-linked network through a supramolecular-covalent cascade reaction route. This protocol demonstrates the construction of two biological membrane-hybridized hydrogels, including liposome-hybridized muscle-mimicking hydrogels with swelling-strengthening mechanical behavior and extracellular vesicle-hybridized skin-mimicking hydrogels with enhanced mechanical strength, lubricity, antibacterial activity and immunoactivity. We describe the detailed preparation procedures and characterize the structures and functions of the obtained hydrogels. We also expand the applicability of this biological membrane hybridization strategy to further tune the structure and function of the biomimetic hydrogels by incorporating a second network. This protocol provides a robust preparative platform to develop dual structure- and function-tunable hydrogels for different biomedical applications. Excluding the synthesis of reactive group-functionalized biological membranes, the fabrication of muscle-mimicking hydrogels takes ~3 d, while the construction of skin-mimicking hydrogels takes ~1 d. The implementation of the protocol requires expertise in polymer modification, hydrogel preparation, nanoscale vesicles, surface functionalization and cell culture.

Extracellular vesicles are a heterogeneous group of membrane-bound vesicles involved in cell-cell communication, formed at the plasma membrane (ectosomes) or by endocytosis (exosomes). Most exosome studies so far have focused on in vitro systems or exosomes derived from bodily fluids, while tissue-derived exosomes remain underexplored. Here we present a protocol using cell-type-specific promoter-driven reporter constructs for the targeted labeling and subsequent isolation of exosomes from specific cell types in vivo from mouse tissues. The differentiation between exosomes and ectosomes remains challenging due to limitations of current isolation techniques that are primarily based on size, density or surface markers. To address this issue, our approach leverages genetic engineering to mark exosomes specifically, enabling their precise identification and isolation from a complex biological pool of heterogenous extracellular vesicles. The isolated cell-type-specific exosomes are characterized by electron microscopy, nanoparticle tracking analysis, antibody exosome array assay and other established techniques. The labeling and isolation of exosomes spans 2-3 days and is designed to be accessible to researchers with fundamental laboratory competencies. This protocol facilitates the study of exosome-mediated cellular communication by enabling the isolation of cell-type-specific exosomes from either individual cell types or multiple cell types in combination. Most experiments within the protocol have used murine wound-edge skin tissue, but the protocol can, in principle, also be applied to other tissues to isolate exosomes, with a few modifications as required. This methodology opens new avenues for exploring the functional roles of cell-type-specific exosomes in intercellular communication.