Nature Protocols最新文献

Lipid nanoparticles (LNPs) have garnered tremendous enthusiasm in preclinical and clinical settings for the delivery of nucleic acids such as mRNA. With applications in protein replacement therapies, vaccines and gene editing, mRNA LNPs have only recently been explored in the context of pregnancy disorders. There is a significant need for the design of novel therapeutic technologies such as mRNA LNPs to treat obstetric disorders like pre-eclampsia that are associated with placental pathology and detrimental effects on maternal and fetal health. Here, we present a step-by-step procedure for the preparation and evaluation of placenta-tropic mRNA LNPs for researchers from varied disciplines to explore their application in treating pregnancy disorders. In this Protocol, we describe steps for synthesizing and purifying the key ionizable lipid excipient of the placenta-tropic LNP formulation (4 d) before preparing mRNA LNPs using microfluidic mixing (1 d). Then, we detail in vitro mechanistic evaluations of the effect of protein adsorption on LNP-mediated mRNA transfection to placental trophoblasts (3 d). Finally, we outline methods for isolating reproductive tissues from time-dated pregnant mice to assess in vivo LNP biodistribution and mRNA transfection to the murine placenta (16 d). Compared to alternative LNP formulation procedures, this Protocol focuses on delivering mRNA LNPs to the placenta with a workflow that can be applied for a range of obstetric disorders. This Protocol seeks to increase interdisciplinary work at the interface of nanomedicine, gene modulation and reproductive health.

Intravenous administration of lipid nanoparticles for the delivery of nucleic acid therapeutics remains constrained by passive uptake mechanisms in the liver, often necessitating high doses to achieve meaningful transfection in specific cells of interest. Targeted LNPs (tLNPs) can overcome these challenges by (i) enabling receptor-mediated endocytosis in difficult-to-transfect cells, thereby reducing passive clearance; (ii) increasing the proportion of LNPs reaching their intended target; and (iii) enabling comparable protein expression at lower doses. Here, we provide a step-by-step guide for formulating tLNPs functionalized with whole antibodies or antibody fragments using traditional laboratory equipment. We outline procedures for antibody preparation and labeling (0.5-1 d), antibody-LNP conjugation (1-2 d), tLNP purification and characterization (1 d) and in vivo and ex vivo targeting evaluation (3-4 d). To demonstrate the versatility of this protocol, we validate in vivo targeting to two mouse tissues: we show that anti-platelet endothelial cell adhesion molecule 1 antibody conjugation to lung-tropic LNPs enhances lung transfection by five times compared to nontargeted LNPs, and anti-epidermal growth factor receptor antibody conjugation to liver-tropic LNPs enhances liver transfection by 20 times. We also demonstrate ex vivo targeting to primary human T cells, where anti-CD5 antibody conjugation to LNPs boosts uptake by 4.5 times and significantly increases mRNA transfection. Importantly, this modular strategy is compatible with any LNP formulation or antibody. In outlining these procedures, we seek to deliver a robust and reproducible workflow for the manufacturing of tLNPs, with the ultimate goal of advancing their therapeutic potential and facilitating clinical translation.

Decentralized, sustainable ammonia production could have an immense global impact. Here we describe an electrolytic approach to synthesizing ammonia directly from air and water under ambient conditions, which could be developed and optimized toward this goal. The system integrates a gliding arc discharge plasma reactor for generating ${\mathrm{NO}}_x$ from air with a membrane electrode assembly reactor for the electrochemical reduction of ${\mathrm{NO}}_x^{-}$ to ammonia, enhancing both the efficiency and scalability of the process. Furthermore, the plasma-generated ${\mathrm{NO}}_x$ feedstock can be substituted with ${\mathrm{NO}}_x$ derived from industrial waste, further extending the potential of this system. In this Protocol, we describe the fundamental principles of this plasma-electrochemical nitrogen reduction reaction (PE-N₂RR) system and provide advice for experimental standardization, operational mechanisms and data analysis methods. The procedure starts with the synthesis of the catalyst-a La_1.5Sr_0.5Ni_0.5Fe_0.5O₄ perovskite oxide-at either laboratory or industrial scale. This catalyst is sufficiently stable to enable the ${\mathrm{NO}}_x^{-}$ RR to continuously work under strongly acidic conditions. We highlight the key operating parameters that are necessary for plasma-based ${\mathrm{NO}}_x$ production and electrochemical ${\mathrm{NO}}_x^{-}$ reduction reaction systems. This information and framework can be used to optimize and streamline the entire PE-N₂RR system. A moderate level of expertise in electrochemistry, plasma systems and catalyst synthesis is recommended to ensure successful execution. The setup of the entire PE-N₂RR system, from catalyst synthesis to the configuration of plasma and electrochemical, is estimated to take 72 h. The full reaction operation test requires 200 h, whereas in situ electrochemical characterizations take 3 h.

In-depth analyses of clinical samples have the potential to provide unparalleled insights into the cellular mechanisms that underlie both health and disease, as well as therapeutic and prophylactic responses. However, these specimens are often paucicellular, necessitating the use of workflows that maximize the amount of information that can be learned. Here we provide a detailed protocol for generating and analyzing single-cell multiomic data from low-input samples with the Seq-Well S³ platform. We further describe a matched pipeline for sample hashing that reduces costs and sources of technical variation in the resulting data while also enhancing throughput. In brief, our streamlined and efficient methodology involves: (1) optionally staining single-cell suspensions with antibody-oligonucleotide conjugates for cell surface protein quantification and/or sample multiplexing; (2) generating Seq-Well S³ sequencing libraries; (3) optionally producing bulk-RNA sequencing libraries via SMART-seq2 to support genetic demultiplexing; and (4) computationally analyzing the resulting data. Each step herein has been designed to leverage readily available reagents and standard laboratory equipment, substantially lowering barriers to entry for researchers. The overall Protocol can yield high-quality multiomic insights from samples in under a week.

Coral reefs are one of the most biodiverse and productive ecosystems on Earth. However, corals are currently under threat from increasing ocean temperatures driven by climate change. Despite the known importance of these fragile ecosystems, our understanding of the molecular mechanisms driving ecologically important traits has been constrained by a lack of genetic tools for functional characterization. To address this limitation, we have developed straightforward and efficient methods to genetically modify corals and study gene function throughout various life history stages using CRISPR-Cas9-based mutagenesis. In this protocol, we first describe how to spawn and collect gametes from the coral Acropora millepora during seasonal spawning events. Next, we describe a method for microinjection of one-cell coral zygotes with CRISPR-Cas9 reagents. We include considerations about effective single-guide RNA design, methods for identifying successfully injected animals, strategies for rearing mutant larvae and juveniles, and methods for the detection and quantification of genomic modifications. This protocol is currently the only way to perform gene editing in corals and takes ~2-4 weeks to complete and has been successfully applied to study genes controlling heat tolerance in coral larvae and skeleton formation in coral juveniles. These technical advances set the foundation for a new field using reverse genetics to study ecologically important traits in corals, such as the establishment of symbiosis and its breakdown upon heat stress.

Brain oscillations are rapidly fluctuating neural activity carried out by large ensembles of neurons. The responsiveness of the brain to external stimuli and the plasticity induced by external stimulation depend strongly on these oscillatory brain states. Here we detail step-by-step instructions for stimulating the brain by tracking human brain activity via the real-time analysis of brain signals recorded by electroencephalography (EEG) and reconstructed in source space, and timing the application of noninvasive transcranial magnetic stimulation (TMS) in synchronization with oscillatory brain states. Real-time EEG-TMS enables the millisecond-precise timing of TMS pulses relative to target oscillatory brain states, such as the phase of an ongoing oscillation. Initial evidence indicates that oscillatory brain state-dependent EEG-TMS is more effective in inducing long-term plasticity than conventional TMS uncoupled to instantaneous oscillatory brain state. This opens the possibility of personalizing therapeutic brain stimulation by coupling it to specific physiological or pathological brain states. In the procedure, we cover brain MRI and image segmentation for anatomical modeling and accurate source reconstruction of the EEG signals, EEG recording without TMS to validate the origin of the oscillation of interest and to determine its phase targeting accuracy, and the main experiment of oscillatory brain state-dependent real-time EEG-TMS to achieve the desired neuroplastic effect. A moderate level of computer science expertise, standard MRI and TMS neuronavigation equipment and TMS-compatible EEG with an accessible online output copy suffice to perform the protocol. The protocol requires ~10 h to complete.

Single-cell epigenome and transcriptome profiling enables the dissection of gene regulatory networks, offering a powerful approach to characterize cellular heterogeneity and regulatory landscapes of cell states. Here we describe a single-cell ultra-high-throughput multiplexed sequencing (SUM-seq) assay for scalable and cost-effective simultaneous profiling of chromatin accessibility and gene expression in single nuclei. SUM-seq combines sample-specific accessible DNA and mRNA in situ barcoding with droplet-based microfluidic barcoding, introducing sample multiplexing and means to resolve multinucleated droplets for multiomic single-cell library preparation. In comparison with existing methods for multimodal profiling of chromatin accessibility and gene expression from the same cell, SUM-seq offers increased throughput and an unmatched multiplexing capability. This permits substantial scaling of the number of samples and nuclei assayed in one experiment, adhering to the needs of large-scale atlas projects, time-course experiments and perturbation screens while considerably reducing costs. We provide guidelines for experimental design and sample handling to accommodate various settings and sample types. Moreover, we discuss potential applications and provide guidelines for data processing. From sample collection to library preparation, the assay can be completed in 2-3 days, followed by sequencing and 1 day of data processing. Although the protocol can be implemented by researchers with general molecular biology skills, prior experience with single-cell assays is recommended.

During tissue regeneration, cells are recruited from surrounding tissue to the defect site. However, when the defect site is large and morphologically complex, cell recruitment often fails to match healthy tissue morphology, resulting in a dysfunctional repair. The integration of bioscaffolds can help to direct the repair process. Here, we present a protocol that integrates electrospinning, weaving, thermal fixation and modified gas-foaming technologies to fabricate 3D hierarchically aligned nanofiber scaffolds. The scaffolds exhibit high porosity, controlled fiber alignment and diverse configurations (uniaxial, bidirectional, radial and gradient alignments), creating effective 'cell highways' for promoting collective cell migration. Applications include hemostatic materials, skin and bone regeneration, hernia repair and biomedical swabs. Both in vitro and in vivo, the highly porous and directionally arranged 3D nanofiber scaffolds markedly enhance cell migration, accelerating the reconstruction of defective tissues. This protocol resolves challenges in production scalability, facilitating the wider adoption of these scaffolds, with a procedure intended for users with expertise in biomaterials and regenerative medicine. The 3D nanofiber scaffolds require 1 d to synthesize and result in improved cell migration during in situ tissue regeneration.

The architecture and function of biological systems are inherently three-dimensional, yet most existing spatial transcriptomic technologies remain restricted to thin tissue sections, limiting their capacity to resolve cellular organization and microenvironments within intact tissue volumes. To address this limitation, we developed volumetric DNA microscopy, a scalable, optics-free approach for spatial transcriptome profiling directly within intact biological specimens. The method encodes spatial information into DNA molecules that form a dense intermolecular network in situ, enabling the reconstruction of three-dimensional spatial relationships through short-read sequencing and computational analysis. Here we detail the complete workflow including in situ cDNA synthesis, spatial encoding through DNA nanoball formation, dual-scale proximity bridging between neighboring nanoballs and spatial reconstruction via geodesic spectral embedding. Sequencing libraries can be generated within 7-8 d by a competent graduate-level molecular biologist, followed by standardized downstream computational analysis. Because the workflow requires only routine molecular biology reagents and a benchtop sequencer, volumetric DNA microscopy provides a versatile platform for exploring genetic and morphological features in intact tissues.