Nature Protocols最新文献

Uneven translation rates resulting from mRNA context, tRNA abundance, nascent amino acid sequence or various external factors play a key role in controlling the expression level and folding of the proteome. Inverse toeprinting coupled to next-generation sequencing (iTP-seq) is a scalable in vitro method for characterizing bacterial translation landscapes, complementary to ribosome profiling (Ribo-seq), a widely used method for determining transcriptome-wide protein synthesis rates in vivo. In iTP-seq, ribosome-protected mRNA fragments known as inverse toeprints are generated by using RNase R, a highly processive 3' to 5' RNA exonuclease. Deep sequencing of these fragments reveals the position of the leading ribosome on each mRNA with codon resolution, as well as the full upstream coding regions translated by these ribosomes. Consequently, the method requires no a priori knowledge of the translated sequences, enabling work with fully customizable transcript libraries rather than previously sequenced genomes. As a standardized framework for inverse toeprint generation, amplification and sequencing, iTP-seq can be used in combination with different types of libraries, in vitro translation conditions and data-analysis pipelines tailored to address a range of biological questions. Here, we present a robust protocol for iTP-seq and show how it can be integrated into a broader workflow to enable the study of context-dependent translation inhibitors, such as antibiotics. The time required to complete this workflow is ~10 d, and the workflow can be carried out by an experienced molecular biologist, with data analysis also requiring a working knowledge of command-line tools and Python scripts.

Optical tweezers use focused laser beams to manipulate small particles, primarily for force sensing. Recent advances in nanoscale-trapping approaches have enabled the development of multiplexed sensing applications, such as temperature and viscosity detection. Upconversion particles (UCPs) and, in particular, lanthanide-doped nano-/micro-crystals (~6 nm to 6 μm) exhibit particular anti-Stokes emission properties, which facilitate their visualization when trapped and the detection of changes to their properties based on temperature and orientation. Their ion resonance enhances the trapping force, enabling the manipulation of smaller particles and their use for force sensing. Here we provide step-by-step instructions to build UCP-based holographic optical tweezers systems, including super-resolved photonic force microscopy and fluorescence optical tweezers. We detail the characterization of the setup for subfemtonewton-scale force sensing and include nanoprobe functionalization, force sensitivity validation and comparison with known forces. We further include the procedures for temperature and viscosity sensing, such as calibrating polarized spectra, initiating UCP rotation and analyzing viscosity via spectral fluctuations. Applications, including nanoparticle-DNA-coated gold film interactions and temperature distribution near single cells, are shown as well. The procedure typically requires 6 days to complete and is suitable for users with expertise in photonics.

Nascent proteins begin to fold during their synthesis, while still attached to the ribosome. The dynamic nature of ribosome-nascent chain complexes (RNCs) poses a challenge for conventional structural biology approaches, limiting our understanding of dynamic cotranslational events. Hydrogen-deuterium exchange mass spectrometry (HDX-MS) is a powerful label-free technique for studying the conformational equilibria and refolding of full-length proteins with peptide resolution. However, the large size of the ribosome and the need for stable, highly homogeneous samples have hindered the application of HDX-MS to RNCs. Here we present a strategy for analysing conformational dynamics and interactors of Escherichia coli RNCs using HDX-MS. High-quality RNCs are obtained through the gentle lysis of high-density cultures expressing uniformly stalled ribosomes, followed by ultracentrifugation and tag-based affinity purification. Peptide-resolution information on protein conformational dynamics is obtained by pulse deuterium labeling, quenching with an RNA-compatible low pH buffer and offline digestion with pepsin. Extensive data analysis with use of specific internal controls allows for the confident assignment of mass spectra to specific peptides, ensuring good coverage of the nascent chain and ribosomal proteins. This method provides a valuable complement to existing structural techniques such as cryo-electron microscopy and nuclear magnetic resonance, and enables detailed characterization of large, partially structured nascent chains and their interactions with the ribosomal proteins and molecular chaperones. The protocol takes 1-3 months, from sample preparation and data acquisition to data analysis, and requires standard expertise in cloning and protein purification and intermediate expertise in HDX-MS.

Multimodal profiling of different molecular layers from the same single cell enables more comprehensive characterization of cellular heterogeneity compared with conventional single-modality approaches. A key example is co-detection of chromatin accessibility and gene expression that offers the opportunity to investigate cell type-resolved gene regulatory mechanisms. Here we describe a sensitive and robust protocol for in situ sequencing hetero RNA-DNA-hybrid after assay for transposase-accessible chromatin using sequencing (ISSAAC-seq) for the concurrent measurement of chromatin accessibility and gene expression from the same single nucleus. The method begins with dual Tn5 tagging of open chromatin regions and the RNA-cDNA hybrid produced by reverse transcription that take place in bulk nuclei. Then, various single-nucleus isolation strategies, including plate and droplet barcoding-based approaches, can be used based on the experimental purpose of the user. The protocol is highly modular with a flexible throughput ranging from several hundreds to tens of thousands of nuclei. The generated data are of high quality in both modalities. The entire workflow can be finished within 1 or 2 days, and the procedures work on multiple different single-nucleus isolation and barcoding platforms.

The isolation of small extracellular vesicles (sEVs), viruses and other nanoscale lipid particles from biofluids offers actionable possibilities for advancing disease diagnosis, drug delivery, regenerative medicine, personalized medicine and immunotherapy. Several methods are available to isolate sEVs from biofluids and acoustic techniques provide distinct advantages. Challenges constraining its wider application encompass the absence of adequate procedures for fabrication, implementation and performance validation. These issues impede the development of protocols applicable to nanoscale bioparticles experiencing acoustic isolation effects. Here we present a detailed protocol for acoustic separation of nanoscale bioparticles from biofluids, including plasma and saliva, achieving both high purity and throughput suitable for routine application. This protocol offers a comprehensive, step-by-step guide for the design and fabrication of the acoustic separation device, the establishment of the experimental setup and the isolation of bioparticles. To ensure reliability, rigor and reproducibility, we delineate essential procedures, including acoustic field optimization, channel fabrication and biofluid preparation, subsequently validating the protocol and its performance across different operators. Our protocol further encompasses procedures for data collection and analysis, which are essential for characterizing viruses and sEVs, as well as for evaluating their quality and integrity. This protocol enables researchers to perform high-quality isolation of nanoscale bioparticles, providing access to reliable acoustic separation techniques. Standardizing this technique will pave the way for discoveries in virology and intercellular communication research, with applications in medicine, biology, and materials science.

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.