Nature Protocols最新文献

With the growing demand for accurate yet effortless health monitoring at home, most current approaches to stool analysis rely on self-reported diaries that are prone to recall bias and low adherence. Here we present a fully passive alternative: the Precision Health Integrated Diagnostic (PHIND) system, a smart-toilet-based platform that enables automated defecation monitoring without requiring users to alter their daily routines. By integrating optical and pressure sensors with cloud-based convolutional neural networks, the PHIND system classifies stool form according to the Bristol Stool Form Scale and records key defecatory parameters, including total event time, defecation duration and time to first stool drop. The protocol proceeds in three principal stages: (1) assembling and mounting the hardware onto a conventional toilet; (2) training convolutional neural network models for stool classification and event detection; and (3) image acquisition and deploying cloud infrastructure for real-time analysis, data storage and visualization. Compared with traditional methods that depend on user-reported stool diaries, PHIND provides objective, near real-time data free from recall error, enabling more reliable early detection and long-term management of gastrointestinal conditions. Researchers and clinicians can expect high classification accuracy and robust, longitudinal insights into defecation patterns. The complete protocol-from hardware setup to system validation-can typically be completed within 2 d, excluding printed circuit board manufacturing, which generally requires up to 15 d depending on the manufacturing provider.

In vivo Raman spectroscopy (RS) enables fast, label-free evaluation of tissue biochemistry in situ with high molecular specificity. The Raman spectrum provides a chemical 'fingerprint' of tissue composition, facilitating investigations of dynamic changes in real-time in various physiological and pathophysiological states. This capability makes in vivo RS a promising approach for rapid diagnostics, surgical guidance and biological research. Despite its numerous advantages, the widespread acceptance of RS for in vivo measurements has been hindered by the lack of a standardized stepwise protocol. This protocol serves as a guide for applying RS in vivo and includes steps for proper instrument selection, system alignment, calibration, system parameter setup, in vivo data collection, instrument cleaning, spectral pre-processing, data analysis and interpretation. Troubleshooting information is described for overcoming challenges in acquiring in vivo RS data due to inherently weak Raman signals, variable tissue optical properties, autofluorescence background and interference from ambient lighting and off-target tissues. Specific steps for applying in vivo RS in the skin, cervix, esophagus and colon are described and can be readily adapted to probe other organs. Typical parameters for acquiring and processing in vivo Raman spectra, as well as example spectral output from different organs, are provided for reference. Ultimately, this standardized protocol serves as a guideline to enhance the repeatability of in vivo RS studies and further expand the adoption of this approach as a research and clinical tool.

Studying the molecular mechanisms underlying the assembly of the human nervous system remains a significant challenge. The ability to generate neural cells from pluripotent stem cells, combined with advanced genome-editing techniques, provides unprecedented opportunities to uncover the biology of human neurodevelopment and disease. Organoids and assembloids enable the in vitro modeling of previously inaccessible developmental processes, such as the specification and migration of human neurons, including the integration of cortical interneurons from the ventral into the dorsal forebrain. Here, we present a detailed protocol that combines pooled CRISPR-Cas9 screening with neural organoid and assembloid models and illustrate how it can be applied to map hundreds of disease genes onto cellular pathways and specific aspects of human neural development. Our protocol outlines key steps, from planning and optimizing genetic perturbations to designing readouts for neuronal generation and migration, conducting the screening and validating candidate genes. The screening experiments take ~3 months to complete and require expertise in stem cell culture and neural differentiation, genetic engineering of human induced pluripotent stem cell lines, fluorescence-activated cell sorting and next-generation sequencing and analyses. The integration of genetic screening and human cellular models constitutes a powerful platform for investigating the mechanisms of human brain development and disease, paving the way for the discovery of novel therapeutics.

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.