Nature Protocols最新文献

Extracellular vesicles (EVs), present in blood as well as other biological fluids, encapsulate nucleic acid biomarkers used for diagnosis, prognosis and treatment monitoring of disease via minimally invasive liquid biopsy. EVs are a reliable source of biomarkers because their contents reflect the cells from which they are derived, and their lipid bilayer membranes protect nucleic acids from degradation. Previously, analyzing EVs in blood was difficult because of time-consuming, labor-intensive EV isolation methods. Here, we provide a protocol for an EV detection approach in which reagent-loaded liposomes fuse with EVs directly in patient blood to sensitively detect RNA within the EVs. In this 'liposome-EV fusion assay', antibodies capture EVs in blood, and reagent-loaded liposomes initiate liposome-EV fusion and CRISPR-based nucleic acid detection. We originally used this assay to detect EV-encapsulated viral RNA and accurately diagnose infectious diseases from patient plasma. It has since been adopted by many other research groups to detect mRNA, microRNA, DNA, DNA mutations and EV surface proteins in a variety of patient-derived tumor samples, incorporating enzymatic and nonenzymatic detection reagents and different diagnostic readouts. As a clinical and research tool, this approach has great potential for the diagnosis, treatment and study of cancer, infectious diseases and neurological dysfunction.

Cryogenic-electron tomography (cryo-ET) permits the in situ visualization of biological macromolecules at the molecular level. Owing to the variable thickness of cells, tissues and organisms, frozen specimens may need to be thinned by cryo-focused ion beam (FIB) milling to produce thin (<500 nm) cryo-lamellae suitable for cryo-ET. Locating regions of interest remains a challenge because untargeted milling can lead to inadvertent ablation and removal of regions of interest. Correlative light and electron microscopy, combined with cryo-FIB milling, can guide the identification of labeled targets in the cellular milieu. Multiple transfers between cryo-imaging instruments, cumbersome correlation algorithms, limited accuracy and low throughput have hindered the routine adoption of cryo-FIB milling within a multimodal correlative workflow for in situ structural biology. Here we present a workflow for 3D correlative cryo-fluorescence light microscopy-FIB-ET that streamlines fluorescence light microscopy-guided FIB milling, improving throughput while preserving both structural and contextual information. The complete integration of hardware and software described here minimizes sample contamination from cross-platform exchanges and greatly enhances the efficiency of 3D targeting in cryo-milling. We then describe procedures for implementing montage parallel array cryo-ET (MPACT), which can be easily adapted to any modern life-science transmission electron microscope. MPACT supports high-throughput cryo-ET acquisitions (10 tilt series in 1.5 h) for structure determination and comprehensive contextual understanding of macromolecules within their native surroundings. A complete session from sample preparation to MPACT data processing takes 5-7 d for an individual experienced in both cryo-EM and cryo-FIB milling.

Single-cell RNA sequencing quantifies biological samples at an unprecedented scale, allowing us to decipher biological differentiation dynamics such as normal development or disease progression. As conventional single-cell RNA sequencing experiments are destructive by nature, reconstructing cellular trajectories computationally is an essential aspect of analysis pipelines. To infer trajectories in a consistent and scalable manner, we have developed CellRank. In its first iteration, CellRank quantitatively recovered trajectories from RNA velocity estimates and transcriptomic similarity. Given these data views, CellRank constructed a cell-cell transition matrix, inducing a Markov chain to automatically infer terminal states and describe their lineage formation. However, CellRank did not enable incorporating complementary data views such as experimental time points, pseudotime or stemness potential. To facilitate these and future views, CellRank 2 generalizes CellRank's trajectory inference framework to multiview single-cell data, leading to a general and scalable framework for cellular fate mapping. Overall, the CellRank framework enables the consistent quantification of cellular fate, combining complementary views and analyzing lineage priming consistently. Here we provide detailed protocols on how to run exemplary CellRank analyses at scale and across different data views. Using CellRank requires basic apprehension and knowledge of single-cell omics data and the Python programming language.

As DNA sequencing technologies improve, it is becoming easier to sequence and assemble new genomes from non-model organisms. However, before a newly assembled genome sequence can be used as a reference, it must be annotated with genes and other features. This can be conducted by individual laboratories using publicly available software. Modern genome annotations integrate gene predictions from the assembled DNA sequence with gene homology information from other high-quality reference genomes and take into account functional evidence (e.g., protein sequences and RNA sequencing information). Many genome annotation pipelines exist but have varying accuracies, resource requirements and ease of use. This genome annotation Tutorial describes a streamlined genome annotation pipeline that can create high-quality genome annotations for animals in the laboratory. Our workflow integrates existing state-of-the-art genome annotation tools capable of annotating protein-coding and non-coding RNA genes. This Tutorial also guides the user on assigning gene symbols and annotating repeat regions. Finally, we describe additional tools to assess annotation quality and combine and format the results.

Mammalian development starts at fertilization and continually progresses until birth, except in cases in which an interruption is favorable to the embryo and the mother. Many mammals have the ability to pause development in case of suboptimal resources or routinely as part of their reproductive cycle-a phenomenon called 'embryonic diapause'. Diapause can be mimicked in vivo in mice via surgical removal of the ovaries or hormone injections. This procedure is laborious and invasive, ruling out its use across species. We have developed in vitro protocols through which mouse blastocysts, human blastoids and pluripotent stem cells from both species can be induced to enter a diapause-like dormant state via pharmacological inhibition of mTOR. Here, we describe in detail how embryos, blastoids and stem cells can be transitioned into and out of dormancy under different culture conditions. We further explain critical parameters to ensure success and propose experimental readouts. These in vitro embryonic dormancy setups can be used to uncover molecular mechanisms of dormancy, to test environmental or pharmacological effectors and to further innovate culture systems for species in which in vitro reproductive technologies are limited. We anticipate that researchers with ~1 year of embryo- and stem cell-handling experience should be able to achieve consistent results and evaluate outcomes. Altogether, inducing dormancy in vitro offers the possibility to slow down embryonic development for exploratory investigations of molecular mechanisms and eventually to expand the time window before implantation for clinical assays.

Elucidating the mechanical regulation of angiogenesis remains a challenge owing to the complexities of measuring cellular forces in this dynamic, multicellular and three-dimensional (3D) process. Current methods for force measurements typically involve traction force microscopy (TFM) applied to single cells or monolayers on 2D substrates or to individual cells within 3D extracellular matrix (ECM)-like gels. Here we present a protocol for mimicking and imaging dynamic early angiogenic sprouting into biomimetic matrices compatible with 3D TFM and for visualizing matrix degradation. Given that reliably acquiring sufficiently large 3D TFM datasets in multicellular systems is challenging, our protocol emphasizes best practices for higher-throughput data acquisition and for accurately imaging, analyzing and interpreting cell-ECM forces using our open-source TFMLAB software. As such, this assay provides a defined and reproducible system to study cellular forces and matrix degradation during angiogenesis in response to perturbations of cell-intrinsic signaling and mixed cell populations, as well as ECM cues. We further provide protocols for immunofluorescence analysis of angiogenic sprouts formed within the matrices and their retrieval from the hydrogel for downstream sequencing. Depending on the number of samples, sample preparation can take between 2 h and 4 h followed by a 15-17 h overnight wait time for angiogenic invasion. The 3D TFM data acquisition can take 2-6 h, while downstream processing of samples can take either 1 h (endothelial cell isolation) or up to 5 d (immunofluorescence). Notably, this workflow demands minimal prior expertise in programming, biophysics or molecular biology.

Practical lignin valorization strategies have the potential to enhance the profitability of lignocellulosic biorefineries. Leveraging the phenolic structure of lignin, commercially available lignins (for example, kraft, soda or biorefinery lignin) have been extensively studied as renewable substitutes for phenol in the synthesis of lignin-phenol-formaldehyde resin adhesives. However, the large-scale production and adoption of lignin-phenol-formaldehyde resin adhesives remain limited owing to challenges related to appearance, performance and cost. To address these limitations, this protocol outlines a comprehensive process for producing lignin adhesives with light colors, high adhesion properties and superior water and weather resistance. The procedure encompasses: (1) isolation of lignin from various types of biomass pretreatment liquors; (2) screening of high-quality lignins from the isolated lignins; and (3) performance assessment of lignin adhesives prepared directly from high-quality lignins without the need for chemical modification or additional processing. Additionally, this protocol provides a rapid and quantitative method for determining the condensation degree of lignin using a small amount of the isolated lignin sample (~50 mg) within a relatively short experimental time (5 h 20 min). This enables efficient quality evaluation and screening of high-quality lignins derived from different extraction methods and biomass sources. The entire process, from biomass to lignin adhesive fabrication, requires a total of 8 h 10 min and is designed for users with prior experience in biomass fractionation, chromatographic analysis and wood-based panel production.

Glioblastomas (GBMs) functionally integrate into diverse neuronal circuits within the central nervous system, which can promote tumor progression and affect neurons via neuron-to-glioma synapses. It remains challenging to identify and manipulate tumor-innervating neurons, which may remain localized or widely distributed throughout the brain. Building on GBM organoids (GBOs) derived from patient-resected surgical tissue, we present here detailed procedures for assessing interactions between tumors and neurons. We first discuss retrograde trans-monosynaptic tracing approaches to study the neuron-tumor connectome by using a rabies viral system in ex vivo human tissue and in xenogenic animal models. As a complementary approach, we then describe the use of anterograde transsynaptic tracing using herpes simplex virus in vivo and ex vivo to assess brain region-specific connectivity in GBMs. In addition, to facilitate the adaptability of these tracing methodologies in diverse systems, we provide procedures for the viral transduction into GBOs, the generation of assembloids comprising GBOs and human induced pluripotent stem cell-derived cortical organoids and the establishment of air-liquid interface cultures from surgical human brain tissue. Together, these techniques permit the flexible characterization and manipulation of tumor-neural circuits and can be easily adapted to other cancers with nervous system involvement. After the generation of GBOs and/or cortical organoids, transsynaptic tracing requires 12-35 d to complete ex vivo or in vivo. The procedure is suitable for users with expertise in human cell and organoid culture, viral production and transduction, rodent surgery and microscopy.

The construction of carbon‒nitrogen (C‒N) bonds is an essential transformation for synthesizing value-added organonitrogen compounds, including the raw materials for fertilizers, synthetic materials and pharmaceuticals. Electrocatalytic C‒N bond construction has emerged as an alternative strategy to traditional thermochemical routes, avoiding harsh and energy-intensive processes. This protocol describes an electrocatalytic C‒N bond construction strategy to synthesize organonitrogen from nitrogen oxides in water under ambient conditions, with a focus on several vital chemicals, such as urea, formamide, cyclohexanone oxime, amino acids and ¹⁵N-labeled amino acids. Here, we provide detailed procedures for catalyst and reaction device design, electrosynthesis, product quantification and measurements for investigating the reaction mechanisms. Four catalysts, namely, vacancy-rich ZnO, core-shell Cu@Zn, AgRu alloy and low-coordination Ag, are synthesized as cathode catalysts. Two types of electrolyzers, an H-type cell and a flow cell, are used for the electrocatalytic reaction. Characterization techniques such as electrochemical in situ Raman spectroscopy, in situ attenuated total reflectance-Fourier transform infrared spectroscopy, ex situ electron paramagnetic resonance and scanning flow cell-differential electrochemical mass spectrometry have been adopted to study the reaction mechanism. The synthesis amount of urea is at the micromole level, while the synthesis amounts of formamide, cyclohexanone oxime and amino acids are at the millimole level. The catalyst synthesis protocol requires 0.5-1.5 d, the electrosynthesis requires ≤11 h and the in situ characterization requires 0.5-1.5 h.

Cartilage is an essential component of the vertebrate skeleton, providing biomechanical support via its extracellular matrix composition. However, in many mammals, including humans and mice, numerous head, neck and chest cartilages produce little extracellular matrix and, instead, contain many large intracellular lipid vacuoles, which determine tissue size, shape and biomechanics. Such cartilages, termed lipocartilages, are made of individual cells called lipochondrocytes with distinct gene expression, lipid composition and metabolism. Lipochondrocytes significantly influence tissue-level physiology, regenerative potential and aging of skeletal elements. Here we provide a step-by-step protocol for the isolation of lipocartilage from mouse ear and the purification of its lipochondrocytes. We include instructions on how to microdissect ear lipocartilage for the purposes of lipid staining, wholemount imaging, morphometric analyses and biomechanical assays. Furthermore, we include a guide for the efficient dissociation of lipocartilages and the purification of individual lipochondrocytes by means of lipid-based buoyancy or cell sorting following fluorescent staining with neutral lipid dyes. With adequate dissection tools and sufficient practice, a researcher can cleanly isolate mouse ear lipocartilage within 20 min and purify lipochondrocytes within 4 h. Tissue biomechanics can be assayed by tensile testing within 30 min per sample. Although the protocol has only been validated in mice, it might be possible to adapt it for larger mammals, but modifications would probably be necessary, as lipocartilage is thicker. These guidelines will serve as a standard for future experiments on lipocartilage and have applications in the fields of developmental biology, bioengineering and metabolism.