Nature Protocols最新文献

Force generation is an integral part of cellular behavior. It plays a crucial role in cell adhesion, migration and division. Mechanical forces are also essential in cell-to-cell interactions, including the widespread interactions involving immune cells. Accurately measuring these forces remains a major challenge, yet it is essential for understanding the mechanobiological mechanisms driving these interactions. Here, we describe a methodology in which deformable and tunable hydrogel microparticles are used to quantify cellular forces. A specific type of acrylamide-based tunable hydrogel microparticles, deformable poly-acrylamide co-acrylic acid microparticles (DAAM-particles), are synthesized in batch using a membrane emulsification approach and conjugated with both biologically active molecules and fluorescent labels through a one-pot functionalization procedure. Cells are then incubated with functionalized DAAM-particles and imaged by confocal microscopy. With a custom image-analysis strategy, local microparticle deformations can be quantified with super-resolution accuracy (<50 nm). Elasticity theory calculations allow for the inference of normal and shear forces, revealing the direction and spatial distribution of cellular forces. The tunability of DAAM-particles enables their adaptation for investigating numerous cellular processes, making them a valuable tool for understanding mechanobiology. The entire protocol takes 2-3 d, requires only basic expertise in mammalian cell culture and fluorescence microscopy and utilizes less specialized equipment and facilities compared with other available techniques. As an example, we demonstrate how this methodology can reveal actin-based force generation during phagocytosis by macrophages.

To understand viral infection and virus-host interactions, real-time, single-cell assays to track viral infection progression are essential. Many conventional assays sample large numbers of cells for single measurements, averaging out the cell-to-cell heterogeneity that is intrinsic to viral infection. Moreover, conventional assays often require cell fixation or lysis, limiting analysis to a single timepoint and masking the temporal and spatial dynamics of infection. We have developed virus infection real-time imaging (VIRIM), a method to visualize the translation of individual RNAs of viruses in real-time. The single-molecule and live-cell nature of VIRIM allows the examination of the earliest events of viral infection, when viral protein and RNA levels are still low, and allows study into the origins and consequences of cell-to-cell heterogeneity during virus infection. Here we provide a step-by-step description of the VIRIM assay, including a detailed procedure for designing, producing and validating the viruses required for VIRIM. In addition, we provide guidelines for generating the reporter cell line, performing the time-lapse imaging and analyzing the fluorescence microscopy data. Once established, a typical VIRIM experiment requires 2-5 days to complete.

Tumor initiation remains one of the least understood events in cancer biology, largely due to the challenge of dissecting the intricacy of the tumorigenic process in laboratory settings. The insufficient biological complexity of conventional in vitro systems makes animal models the primary experimental approach to study tumorigenesis. Despite providing valuable insights, these in vivo models function as experimental black boxes with limited spatiotemporal resolution of cellular dynamics during oncogenesis. In addition, their use raises ethical concerns, further underscoring the need for alternative ex vivo systems. Here we provide a detailed protocol to integrate state-of-the-art microfabrication, tissue engineering and optogenetic approaches to generate topobiologically complex miniature colons ('mini-colons') capable of undergoing tumorigenesis in vitro. We describe the key methodology for the generation of blue light-inducible oncogenic cells, the establishment of hydrogel-based mini-colon scaffolds within microfluidic devices, the development of mini-colons and the induction of spatiotemporally controlled tumorigenesis. This protocol enables the formation and long-term culture of complex cancerous tissues that capture in vivo-like tumoral biology while offering real-time and single-cell resolution analyses. It can be implemented in 4-6 weeks by researchers with prior experience in 3D cell culture techniques. We anticipate that these methodological guidelines will have a broad impact on the cancer research community by opening new avenues for tumorigenesis studies.

The electrocatalytic nitrate reduction reaction (NO₃RR) has emerged as a promising approach for sustainable nitrogen management, enabling the selective conversion of nitrate into targeted nitrogen-containing compounds, such as ammonia and hydroxylamine. However, the efficiency and selectivity of the NO₃RR are highly dependent on the physicochemical properties of the electrocatalysts, necessitating a standardized and comprehensive characterization protocol. Here we provide a detailed methodology for the structural, chemical, electronic and electrochemical characterization of the materials used in the NO₃RR. We outline procedures for evaluating catalyst morphology, composition and redox states, as well as methodologies for quantifying reaction products to determine nitrate conversion efficiency and selectivity. To track catalyst evolution and reaction pathways under reaction conditions, we present real-time monitoring strategies that capture structural changes, key reaction intermediates and electronic transformations associated with chemical bond formation and cleavage. In addition, we incorporate theoretical calculations to comprehensively evaluate the reaction pathways and their interplay with the electronic structures of electrocatalysts, providing deeper mechanistic insights into the reaction kinetics, active site evolution and selectivity-determining factors. This Protocol is designed for researchers in electrocatalysis, environmental chemistry and energy conversion, offering a reproducible workflow for catalyst assessment. The step-by-step methodology ensures reliable data collection and interpretation, enabling direct comparisons across different catalysts and facilitating the development of more efficient NO₃RR catalysts. The entire workflow requires ~8-10 days, depending on sample preparation and measurement duration.

Constraint-based modeling can mechanistically simulate the behavior of a biochemical system, permitting hypothesis generation, experimental design and interpretation of experimental data, with numerous applications, especially the modeling of metabolism. Given a generic model, several methods have been developed to extract a context-specific, genome-scale metabolic model by incorporating information used to identify metabolic processes and gene activities in each context. However, the existing model extraction algorithms are unable to ensure that a context-specific model is thermodynamically flux consistent. Here we introduce XomicsToModel, a semiautomated pipeline that integrates bibliomic, transcriptomic, proteomic and metabolomic data with a generic genome-scale metabolic reconstruction, or model, to extract a context-specific, genome-scale metabolic model that is stoichiometrically, thermodynamically and flux consistent. One of the key advantages of the XomicsToModel pipeline is its ability to seamlessly incorporate omics data into metabolic reconstructions, ensuring not only mechanistic accuracy but also physicochemical consistency. This functionality enables more accurate metabolic simulations and predictions across different biological contexts, enhancing its utility in diverse research fields, including systems biology, drug development and personalized medicine. The XomicsToModel pipeline is exemplified for extraction of a specific metabolic model from a generic metabolic model; it enables omics data integration and extraction of physicochemically consistent mechanistic models from any generic biochemical network. It can be implemented by anyone who has basic MATLAB programming skills and the fundamentals of constraint-based modeling.

The myosin super-relaxed (SRX) state is a biochemical and structural conformation of myosin that modulates contractility and energy expenditure and is in equilibrium with the disordered relaxed (DRX) state of myosin, which can hydrolyze ATP to produce force. The proportion of myosin SRX-DRX states is perturbed in various muscle disorders, and myosin SRX-DRX states have become a promising drug target. There are many approaches that can be used to interrogate myosin conformations, including X-ray diffraction, stopped-flow kinetics and electron microscopy. These techniques are highly informative but necessitate highly skilled researchers and specialist equipment, limiting wider uptake and accessibility. For this reason, we provide a set of protocols detailing established assays to measure biochemically defined myosin SRX-DRX states in skeletal muscle, cardiac muscle, induced pluripotent stem cell-derived cardiomyocytes, myofibrils, reconstituted thick filaments and isolated molecular motors by using a simple chase assay incorporating a fluorescent ATP analogue: methylanthraniloyl (Mant)-ATP. The Mant-ATP assay provides a biochemical measure of myosin states that is distinct from assays that are used to visualize myosin structure directly. These Mant-ATP assays have various protocol lengths, ranging from 1-2 d for preparation and 30 min to run an experiment. With this set of protocols, we make the Mant-ATP assay accessible to those working in biochemistry, muscle physiology and cell biology. At the end of this protocol, users should be able to ascertain a clean fluorescent decay trace that can be fit to define the ratio of SRX/DRX myosin in their sample of choice.

Understanding how cells sense and respond to mechanical forces is crucial for many biological processes, including adhesion, migration, differentiation and immune activation. In this protocol, we describe two advanced DNA-based tension probes, the reversible shearing DNA-based tension probe (RSDTP) and ForceChrono probe, which provide powerful tools for studying mechanotransduction in living cells. RSDTPs enable dynamic quantification of forces ranging from 4 to 60 pN, offering the advantage of reversibility without ligand depletion, making them ideal for ensemble force measurements across populations of cells. ForceChrono probes not only measure the magnitude of force but also capture its duration and loading rate, providing essential insights into the temporal dynamics of single-molecule force transmission. We detail the fundamental principles, design strategies and step-by-step procedures for synthesizing, purifying and applying these probes, including surface preparation, cell experiments, image acquisition and data analysis. In addition, we describe the computational tools for image analysis. Together, these probes enable a detailed analysis of cellular mechanobiology, with applications in integrin mechanobiology and cell adhesion biology. This protocol is suitable for researchers with a background in cell biology, molecular biology, surface chemistry, optical imaging and data analysis and can be completed by a graduate student in 3-4 days.

Here, we describe a practical step-by-step protocol for directional conjugation of monoclonal antibodies to polymersome nanoparticles via the Fc antibody moiety using metal-free click chemistry. This Protocol Extension details procedures for synthesis, quality control and evaluation of nanoparticle-antibody conjugates. The synthesis includes three main stages: (i) a mild oxidation of the glycosyl moiety that is present only on the Fc region of the antibody to produce functional aldehyde groups; (ii) attachment of a heterobifunctional linker, aminooxy-PEG-dibenzocyclooctyne, to the aldehyde groups in the Fc moiety; and (iii) click conjugation of the PEG-linker-modified antibodies to nanoparticles with azide functional groups. This protocol enables covalent surface conjugation of monoclonal antibodies that inherently does not involve the antigen-binding Fab region, thus minimizing the impact of the conjugation on the avidity of antibodies. In contrast, conventional conjugation methods that use amine groups of antibodies for covalent immobilization result in a random antibody orientation that can lead to a loss of the antibody's binding efficiency. The entire conjugation protocol requires ~18 h. The nanoparticle-antibody conjugates synthesized using this method are expected to display a high molecular specificity toward epitopes of the conjugated antibodies while maintaining the physicochemical properties of the core nanoparticles. The protocol described here does not require any special expertise other than general laboratory training on equipment such as a centrifuge, ultraviolet-visible spectrophotometers, dynamic light scattering, ELISA plate readers and cell culturing.

Protein S-nitrosylation (SNO) is a ubiquitous post-translational modification, which regulates a broad range of functional parameters, including protein stability; enzymatic, transcriptional and ion channel activity; and cellular signal transduction. Aberrant protein SNO is associated with diverse pathophysiology, from cardiovascular, metabolic and respiratory disorders to neurodegeneration and cancer. Drugs that enhance or inhibit specific SNO reactions are being developed as potential disease-modifying therapeutics. However, owing to a lack of suitable approaches to monitor SNO proteins, which often exist at low abundance with ephemeral expression, a systematic understanding of their roles in disease remains elusive. Here we report a robust and proteome-wide approach for the exploration of the S-nitrosoproteome in human and mouse tissues, using the brain as an example, with a probe named SNOTRAP (a triphenylphosphine thioester linked to a biotin molecule through a polyethylene glycol spacer group) in conjunction with mass spectrometry (MS)-based detection. In this Protocol, we detail tissue sample preparation, synthesis of SNOTRAP under an argon atmosphere and subsequent MS-based identification and analysis of SNO proteins. In situ labeling of SNO proteins is achieved by the SNOTRAP probe, concomitantly yielding a disulfide-iminophosphorane as a labeling tag. The chemically tagged proteins can be digested, followed by streptavidin capture, release by triscarboxyethylphosphine and relabeling of the liberated free Cys with N-ethylmaleimide. This approach selectively enriches SNO-containing peptides at specific sites for label-free quantification by Orbitrap MS. It requires about 5 d for synthesis of the SNOTRAP probe, 2-2.5 d for sample preparation and about 5 d for nano-liquid chromatography-tandem MS measurement and analysis.

BreakTag is a scalable next-generation sequencing-based method for the unbiased characterization of programmable nucleases and guide RNAs at multiple levels. BreakTag allows off-target nomination, nuclease activity assessment and the characterization of scission profile, that, in Cas9-based gene editing, is mechanistically linked with the indel repair outcome. The method relies on digestion of genomic DNA by Cas9 and guide RNAs in ribonucleoprotein format, followed by enrichment of blunt and staggered DNA double-strand breaks generated by CRISPR nucleases at on- and off-target sequences. Next-generation sequencing and data analysis with BreakInspectoR allows high-throughput characterization of Cas nuclease activity, specificity, protospacer adjacent motif frequency and scission profile. Here we first describe a detailed BreakTag protocol for the nomination of CRISPR off-targets and multilevel characterization of engineered Cas variants and second, we describe a step-by-step protocol for data analysis using BreakInspectoR. Third, we provide a web interface for XGScission, a machine learning model amenable to training with scission-aware BreakTag data to predict the relative frequency of blunt and staggered double-strand breaks at new sequences unseen by the model. XGScission allows a preselection of target sequences predicted to be cut in staggered configuration that are preferably repaired as single-nucleotide templated insertions. Furthermore, XGScisson can be used to assess sequence determinants of blunt and staggered cleavage by SpCas9 and engineered nuclease variants. As a companion strategy, we describe HiPlex for the generation of hundreds to thousands of single guide RNAs in pooled format for the production of robust BreakTag datasets. The BreakTag library preparation takes ~6 h, and the entire protocol can be completed in ~3 d, including sequencing, data analysis with BreakInspectoR and XGScission model training.