Nature Protocols最新文献

As microRNAs (miRNA) regulate almost all physiopathological activities in the human body, miRNA therapeutics that deliver miRNA regulators have attracted considerable attention in the field of nucleic acid drug development. The use of tetrahedral DNA nanostructures to deliver miRNA regulators is promising because of their simple fabrication, enhanced cell entry, effective tissue penetration, biocompatibility and functional editability. This protocol extension builds on our previous protocol for the use of tetrahedral DNA nanostructures and was designed to establish an updated bioswitchable delivery system (BDS) for achieving controlled cargo loading and release. A ribonuclease H-sensitive sequence is designed as a bioswitchable apparatus for the targeted release of the miRNA regulator. The functional sequence of the miRNA regulator and minimal secondary structure formation tendency during annealing are two key points in cargo design. We provide two BDS design strategies; BDS-A comprises an intact DNA tetrahedron with the RNA cargo hanging outside, offering the merits of lower cost, simplicity, and more direct structural design. In the BDS-B design, the RNA regulators are embedded into the DNA tetrahedron, which is beneficial for dermal tissue permeation applications. Following sequence design in Oligo 7 and Tiamat, the BDS assembly is completed and then ribonuclease H achieves controlled release of the miRNA regulator by triggering the bioswitchable apparatus. This is verified via polyacrylamide and agarose gel electrophoresis or fluorophore modifications. Both BDSs show promising cellular membrane permeability, tissue permeability and target inhibition in vitro and in vivo. The assembly and characterization of the BDS can be completed in 4 d, and the validation time for biostability and biological applications will depend on the specific use.

Measuring forces within living cells remains a technical challenge. In this Tutorial, we cover the development of hydrophobic mechanosensing fluorescent probes called Flippers, whose fluorescence lifetime depends on lipid packing. Flipper probes can therefore be used as reporters for membrane tension via the measurement of changes in their fluorescence lifetime. We describe the technical optimization of the probe for imaging and provide working examples for their characterizations in a variety of biological and in vitro systems. We further provide a guideline to measure biophysical parameters of cellular membranes by fluorescence lifetime imaging microscopy using Flipper probes, providing evidence that flippers can report long range forces in cells, tissues and organs.

To understand the dynamic nature of the genome, the localization and rearrangement of DNA and DNA-binding proteins must be analyzed across the entire nucleus of single living cells. Recently, we developed a computational light microscopy technique, called high-resolution diffusion (Hi-D) mapping, which can accurately detect, classify and map diffusion dynamics and biophysical parameters such as the diffusion constant, the anomalous exponent, drift velocity and model physical diffusion from the data at a high spatial resolution across the genome in living cells. Hi-D combines dense optical flow to detect and track local chromatin and nuclear protein motion genome-wide and Bayesian inference to characterize this local movement at nanoscale resolution. Here we present the Python implementation of Hi-D, with an option for parallelizing the calculations to run on multicore central processing units (CPUs). The functionality of Hi-D is presented to the users via user-friendly documented Python notebooks. Hi-D reduces the analysis time to less than 1 h using a multicore CPU with a single compute node. We also present different applications of Hi-D for live-imaging of DNA, histone H2B and RNA polymerase II sequences acquired with spinning disk confocal and super-resolution structured illumination microscopy.

Targeted integration of large DNA cargoes (>10 kb) or genomic replacements in mammalian cells, such as human pluripotent stem cells (hPS cells), remains challenging. Here we describe a platform termed serine and tyrosine recombinase-assisted integration of genes for high-throughput investigation (STRAIGHT-IN) to circumvent this. First, a landing pad cassette is precisely inserted or used to replace specific genomic regions. The site-specific integrase Bxb1 then enables DNA constructs, including those >50 kb, to be integrated into the genome, while Cre recombinase excises auxiliary DNA sequences to prevent postintegrative silencing. Using a strategy whereby the positive selection marker is only expressed if the donor plasmid carrying the payload is correctly targeted, we can obtain 100% enrichment for cells containing the DNA payload. Procedures for expressing Cre efficiently also mean that a clonal isolation step is no longer essential to derive the required genetically modified hPS cells containing the integrated DNA, potentially reducing clonal variability. Furthermore, STRAIGHT-IN facilitates rapid and multiplexed generation of genetically matched hPS cells when multiple donor plasmids are delivered simultaneously. STRAIGHT-IN has various applications, which include integrating complex genetic circuits for synthetic biology, as well as creating panels of hPS cells lines containing, as necessary, hundreds of disease-linked variants for disease modeling and drug discovery. After establishing the hPS cell line containing the landing pad, the entire procedure, including donor plasmid synthesis, takes 1.5-3 months, depending on whether single or multiple DNA payloads are integrated. This protocol only requires the researcher to be skilled in molecular biology and standard cell culture techniques.

The production of living therapeutics, cell-based delivery of drugs and gene-editing tools and the manufacturing of bio-commodities all share a common concept: they use either a synthetic or a living cell chassis to achieve their primary engineering or therapeutic goal. Live-cell chassis face limitations inherent to their auto-replicative nature and the complexity of the cellular context. This limitation highlights the need for a new chassis combining the engineering simplicity of synthetic materials and the functionalities of natural cells. Here, we describe a protocol to assemble a synthetic polymeric network inside bacterial cells, rendering them incapable of cell division and allowing them to resist environmental stressors such as high pH, hydrogen peroxide and cell-wall-targeting antibiotics that would otherwise kill unmodified bacteria. This cellular bioengineering protocol details how bacteria can be transformed into single-lifespan devices that are resistant to environmental stressors and possess programable functionality. We designate the modified bacteria as cyborg bacterial cells. This protocol expands the synthetic biology toolset, conferring precise control over living cells and creating a versatile cell chassis for biotechnology, biomedical engineering and living therapeutics. The protocol, including the preparation of gelation reagents and chassis strain, can be completed in 4 d. The implementation of the protocol requires expertise in microbiology techniques, hydrogel chemistry, fluorescence microscopy and flow cytometry. Further functionalization of the cyborg bacterial cells and adaptation of the protocol requires skills ranging from synthetic genetic circuit engineering to hydrogel polymerization chemistries.

Nitrogen heterocycles are found in the structures of many biologically important compounds, as well as materials used in the synthesis of fine chemicals. Notably, ~59% of US Food and Drug Administration-approved small-molecule drugs contain nitrogen heterocycles. It is therefore meaningful to explore greener or more sustainable methods for their synthesis. The use of alcohols as reagents is attractive as they can be readily obtained from biomass derived natural resources. In the last two decades, alcohol dehydrogenative coupling reaction to synthesize various heterocycles were extensively explored which furnished hydrogen (H₂) and water (H₂O) as the two greener byproducts. In this protocol, we describe several efficient catalytic transformations to synthesize quinolines, 1,8-naphthyridines, quinoxalines, quinazolines, pyrimidines, benzimidazoles, pyrroles and pyridines, using alcohol as starting materials. We also describe the synthesis of several homogeneous iridium/ruthenium catalysts and heterogeneous cobalt/copper catalysts that can be used in these transformations. The reaction setup is simple; in a Schlenk/reaction tube with magnetic stir-bar, alcohol, corresponding coupling reagents (nucleophiles), catalyst, base and solvent (water or organic solvent such as toluene, dioxane or p-xylene) are added. The reaction mixture is refluxed at the specified temperature (110-150 °C)-either in air or under argon-to furnish these heterocycles. Synthesis of the catalysts takes 3-5 h and the coupling reactions take 4-5 h depending on the target product. The cobalt- and copper-based heterogeneous catalytic systems displayed an good catalyst recyclability.

The mammalian membrane is composed of various eukaryotic lipids interacting with extensively post-translationally modified proteins. Probing interactions between these mammalian membrane proteins and their diverse and heterogeneous lipid cohort remains challenging. Recently, native mass spectrometry (MS) combined with bottom-up 'omics' approaches has provided valuable information to relate structural and functional lipids to membrane protein assemblies in eukaryotic membranes. Here we provide a step-by-step protocol to identify and provide relative quantification for endogenous lipids bound to mammalian membrane proteins and their complexes. Using native MS to guide our lipidomics strategies, we describe the necessary sample preparation steps, followed by native MS data acquisition, tailored lipidomics and data interpretation. We also highlight considerations for the integration of different levels of information from native MS and lipidomics and how to deal with the various challenges that arise during the experiments. This protocol begins with the preparation of membrane proteins from mammalian cells and tissues for native MS. The results enable not only direct assessment of copurified endogenous lipids but also determination of the apparent affinities of specific lipids. Detailed sample preparation for lipidomics analysis is also covered, along with comprehensive settings for liquid chromatography-MS analysis. This protocol is suitable for the identification and quantification of endogenous lipids, including fatty acids, sterols, glycerolipids, phospholipids and glycolipids and can be used to interrogate proteins from recombinant sources to native membranes.

Complex cellular functions occur via the coordinated formation and dissociation of protein complexes. Functions such as the response to a signaling ligand can incorporate dozens of proteins and hundreds of complexes. Until recently, it has been difficult to measure multiple protein complexes at the single-cell level. Here, we present a step-by-step procedure for proximity sequencing, which enables the simultaneous measurement of proteins, mRNA and hundreds of protein complexes located on the outer membrane of cells. We guide the user through probe creation, sample preparation, staining, sequencing and computational quantification of protein complexes. This protocol empowers researchers to study, for example, the interplay between transcriptional states and cellular functions by coupling measurements of transcription to measurements of linked effector molecules, yet could be generalizable to other paired events. The protocol requires roughly 16 h spread over several days to complete by users with expertise in basic molecular biology and single-cell sequencing.

Nanozymes are nanomaterials with enzyme-like catalytic properties. They are attractive reagents because they do not have the same limitations of natural enzymes (e.g., high cost, low stability and difficult storage). To test, optimize and compare nanozymes, it is important to establish fundamental principles and systematic standards to fully characterize their catalytic performance. Our 2018 protocol describes how to characterize the catalytic activity and kinetics of peroxidase nanozymes, the most widely used type of nanozyme. This approach was based on Michaelis-Menten enzyme kinetics and is now updated to take into account the unique physicochemical properties of nanomaterials that determine the catalytic kinetics of nanozymes. The updated procedure describes how to determine the number of active sites as well as other physicochemical properties such as surface area, shape and size. It also outlines how to calculate the hydroxyl adsorption energy from the crystal structure using the density functional theory method. The calculations now incorporate these measurements and computations to better characterize the catalytic kinetics of peroxidase nanozymes that have different shapes, sizes and compositions. This updated protocol better describes the catalytic performance of nanozymes and benefits the development of nanozyme research since further nanozyme development requires precise control of activity by engineering the electronic, geometric structure and atomic configuration of the catalytic sites of nanozymes. The characterization of the catalytic activity of peroxidase nanozymes and the evaluation of their kinetics can be performed in 4 h. The procedure is suitable for users with expertise in nano- and materials technology.