Nature Protocols最新文献

Covalent organic frameworks (COFs) are versatile materials platforms for precise function integration owing to their high crystallinity, large surface areas, tunable characteristics and diverse and predictable structures. However, the dominant solvothermal method for COF synthesis requires harsh conditions, including high temperatures, toxic organic solvents, sealed and pressurized reactors, and extended reaction times that often exceed several days. Here we present a probe-based sonochemical method for synthesizing COFs in an aqueous environment under ambient atmosphere, offering a safer and faster alternative. By employing sonication in an aqueous acetic acid solution, the protocol avoids harmful organic solvents and high-pressure systems and reduces the reaction times to under 1 h. COFs synthesized through this method have large surface areas and high crystallinity, making them ideal for applications in photocatalysis, gas sorption, food contaminant removal and chemical sensing. The broad applicability of this synthesis method has been demonstrated by successfully preparing 62 COFs with various linkage types, including imine-linked, β-ketoenamine-linked, azine-linked and hydrazone-linked COFs, as well as frameworks with one-dimensional, two-dimensional and three-dimensional topologies. The process (performed at the 50-100 mg scale) involves steps such as preparing monomer solutions, using sonication to induce COF formation and postsynthesis purification. The surface area is characterized using nitrogen sorption, while the crystallinity is assessed by powder X-ray diffraction and transmission electron microscopy. The entire protocol can be completed within 24 h and requires moderate expertise in materials chemistry and access to standard laboratory equipment.

High-throughput chemical synthesis plays a critical role in generating compound libraries and optimizing reaction conditions. Increasing adoption of robots and multiwell formats means that hundreds of reactions can be accessed with ease. However, conventional methods for quantitative detection of the product (for example, by liquid chromatography-mass spectrometry) create a bottleneck in the workflow because analyses need to be customized separately for each sample. Here we describe a tandem mass spectrometry approach where the samples are analyzed directly using acoustic ejection mass spectrometry. This approach features simple method development enabling accurate quantification for reactions where a characteristic neutral lost fragment is common to both the chemical starting material and the expected reaction products. Combining this precise fragmentation signature with acoustic ejection mass spectrometry allows whole 384-well plates of chemical reaction data to be collected at the same pace as two liquid chromatography-mass spectrometry samples, while maintaining the same levels of accuracy. To explain the principles involved in designing these experiments, we show four examples of common medicinal chemistry transformations for C-N and C-C bond formation that have been used in the synthesis of analogs of cereblon binding molecular glues, antifungals, antibiotics, and building blocks for automated small molecule synthesis. The whole procedure requires ~2 days of work to complete, including the 384-well plate reaction setup, analytical sample preparation, mass spectrometry data collection and analysis.

Macrophages are crucial in immune responses, tissue repair and homeostasis, making them prime candidates for translational applications. Induced pluripotent stem cell (iPS cell)-derived macrophages hold considerable promise for regenerative medicine, cancer therapy, inflammatory disease treatment and in vitro bioassays. However, cost-effective, standardized intermediate-scale bioreactor systems tailored for early-stage research and drug discovery in academia remain limited. Here, we present an extension of our previously published protocol that is feeder free, semi-defined and user friendly, enabling the standardized production of iPS cell-derived macrophages in an intermediate (10-50 mL)-scale benchtop bioreactor. This Protocol can be implemented by users with basic iPS cell culture experience without requiring advanced bioprocessing expertise. This method consists of two primary endpoints: the generation of mesoderm-primed aggregates with hematopoietic potential, termed hemanoids, and the standardized production of iPS cell-derived macrophages that are ready for downstream applications. This Protocol enables continuous macrophage generation in long-term cultures, with a minimum of five consecutive collections, yielding an average of 2-3 × 10⁷ cells per collection per vessel. Four vessels operate independently, each with a maximum culture volume of up to 50 mL, while critical process parameters (CO₂, temperature and pH) are monitored. This semi-automated platform and in-process monitoring improve process control, leading to higher yields, reproducibility and cell quality compared with other systems. The simplified process spans 24 d, starting from single-cell iPS cells to ready-to-use macrophages. By bridging the gap between small- and large-scale systems, this approach provides scalable, standardized manufacturing of iPS cell-derived macrophages, making it a valuable tool for academics focused on human immune cells such as macrophages.

Microplastics (MPs) are persistent and widely distributed pollutants that have become an emerging environmental problem. When present in the environment, they are increasingly being incorporated into the carbon cycle and thus participate in the Earth's biochemical cycle alongside other pollutants. However, upon entering the environment (as primary or secondary MPs), various environmental factors collectively influence the physical and chemical characteristics of MPs, leading to changes in surface morphology, particle size and microstructure. These alterations can change their physical behavior and the effect that they have on living organisms. The aging processes of MPs are complex and influenced by multiple factors relating to both the size and composition of the MPs themselves and the environmental conditions. We developed a protocol for studying MP aging across different environmental settings (soil, water, air and organisms) over timescales from several weeks to months, applicable to common types of MPs. This protocol integrates modular aging experiments with an array of analytical techniques (for example, scanning electron microscopy, Fourier transform IR spectroscopy and pyrolysis-gas chromatography-mass spectrometry) to characterize MP aging according to micromorphology, chemical characteristics, dissolution characteristics and mass fraction. We have established a quantitative method for evaluating MP aging along with graded property assessment criteria for measuring the degree of aging-the composite aging index (CAI). This protocol takes approximately 4 days to 6 months, depending on the specific environmental aging conditions selected, the sample matrix and the suite of analytical techniques used. By implementing this protocol, we can gain mechanistic insights into MP aging processes, ultimately contributing to sustainable plastics development through strategic design that guides their fate and behavior.

Drosophila is an important model organism for studying epithelial barrier pathogenesis by Pseudomonas aeruginosa because it allows clinically important infectivity to be mimicked. Here, we present a Protocol Extension to introduce P. aeruginosa and other bacteria into the adult Drosophila intestine by feeding, further to our previously published protocol that used the needle-pricking method to impose a wound infection and the injector-pumping method of bacterial delivery directly into the fly hemocoel. We start with Drosophila preparation and priming for feeding, in parallel with bacterial growth and infection mix preparation. We continue with the feeding setup followed by the output measurements, which include, but are not limited to, fly survival, bacterial load and systemic spread, intestinal regeneration and gene expression. The protocol can be used to infect Drosophila with tens of different clinically important and insect-related bacterial species, one at a time or in combination; modify the inoculum consistency to control and even eliminate P. aeruginosa virulence; measure bacterial load in the hemolymph; assess intestinal histopathology of tumor-prone flies; and assess the host transcriptome. Preparation of adult flies for feeding lasts ≤1 week, and infection mixes can be prepared in 2 d, which require a minimum level of expertise in fly handling and microbiological techniques, respectively. All output measurements usually take ≤10 d or until all flies die from a virulent infection. Fly survival and bacterial load assessment require a short training, while intestinal histopathology and host gene expression assessment need up to a month of systematic training. Assessing bacterial load and gut measurements require 2 d of work per infection time point considered.

Biomolecular condensates formed through liquid-liquid phase separation regulate cellular processes, and their dysregulation causes disease. Current methods for identifying endogenous phase-separating proteins have low throughput and cannot capture dynamic responses to stimuli. Here we present a protocol combining osmotic compression or transforming growth factor-β (TGF-β) treatment to induce condensation with sucrose density gradient centrifugation and quantitative mass spectrometry to enable systematic, high-throughput identification of endogenous condensates and phase-separating proteins. The method exploits the density changes that occur when phase-separating proteins undergo oligomerization during condensate formation. In H1975 cells, we identified over 1,500 phase-separating proteins under osmotic compression or TGF-β treatment; 538 of these candidates were not present in PhaSepDB, a database that compiles in vivo, in vitro and omics-derived proteins. The approach detects constitutive condensates and proteins that dynamically phase-separate in response to osmotic stress or TGF-β signaling. This protocol provides proteome-wide analysis of fractions of proteins having different densities and enables temporal resolution of phase-separation events. The procedure takes ~9 d and requires expertise in cell culture, biochemistry and mass spectrometry. This method enables systematic study of biomolecular condensates and disease-associated phase-separation mechanisms.

Molecular recording is an emerging paradigm for measuring biology over time. Enhancer-mediated genomic recording of activity in multiplex (ENGRAM) is a recently described synthetic biology circuit architecture that converts the transient activity of cis-regulatory elements (CREs) into stable genomic records that can be retrospectively recovered via DNA sequencing. Here we provide a step-by-step protocol for conducting ENGRAM experiments and analyzing the resulting data. We also describe key design considerations for ENGRAM recorders, summarize the strengths and limitations of ENGRAM, and highlight applications, including multiplex signal recording and high-throughput CRE screening. In contrast to other systems for DNA-based recording in mammalian systems, ENGRAM relies on prime editing-mediated insertions to record the activity of a given CRE, such that it is inherently multiplexable-for example, four-base-pair insertions can represent the activities of up to 256 distinct CREs. A further contrast lies with ENGRAM's compatibility with DNA Typewriter, which facilitates the capture of signal order. For users with basic skills in molecular biology, mammalian cell culture and DNA sequencing analysis, ENGRAM experiments can typically be completed within 5-6 weeks.

The persistent disparity between organ donation rates and clinical demand has driven the increasing use of extended-criteria donor livers. However, conventional static cold storage inadequately preserves extended-criteria donor with severe ischemia-reperfusion injury (IRI), contributing to high rates of mortality and morbidity. Although different machine perfusion technologies have been used to reduce IRI in clinical practice, organ ischemia remains unavoidable throughout the entire transplantation procedure. To minimize IRI to the greatest extent possible, we developed a novel ischemia-free liver transplantation (IFLT) method based on surgical innovation and continuous normothermic machine perfusion. IFLT not only effectively preserves graft quality but also expands the donor pool, making it possible to utilize high-risk livers. Classic IFLT increases the complexity of donor liver procurement and prolongs the anhepatic phase during implantation. Here we develop a simplified IFLT (SIFLT) technique. By streamlining the donor liver retrieval procedure and optimizing the sequence of vascular anastomosis during implantation, the efficacy and safety data for SIFLT are comparable to those of classic IFLT, with similar rates of postoperative complications, graft survival and patient survival. Thus, SIFLT represents a more efficient, safer and widely applicable approach to minimize organ ischemia, offering a robust strategy to improve outcomes and maximize organ utilization.

Synthetic gene circuits are powerful tools for precisely programming gene expression and introducing novel cellular functions. However, their development and application in plants has lagged behind other systems, due mainly to the limited availability of modular genetic parts. We recently developed a CRISPR interference (CRISPRi)-based synthetic gene circuit system for programming gene expression in plants. Using a robust and high-throughput protoplast-based dual luciferase assay, we demonstrated the development, testing and functionality of these circuits in various plant species. Here we detail the key design principles and considerations for building and testing programmable and reversible CRISPRi-based gene circuits in plants. We also provide detailed procedures for isolating protoplasts from multiple plant species, including Arabidopsis thaliana, Brassica napus, Triticum aestivum and Physcomitrium patens. Furthermore, we provide step-by-step instructions for the 96-well plate-based protoplast transfection assay for testing genetic parts and synthetic circuits, using a dual luciferase assay. The detailed descriptions of these developed systems will enhance the efficiency and reproducibility of the construction, testing, and implementation of synthetic gene circuits in a variety of plant species. This protocol enables the design and testing of CRISPRi-based gene circuits in plants within ~4 weeks.

This protocol describes a standardized in vitro method to determine the digestibility and digestible indispensable amino acid score (DIAAS) of dietary proteins. This 'INFOGEST Quant' method is an extension of our previous INFOGEST static digestion model (INFOGEST 2.0) and adds a workflow for the quantification of total protein digestibility, individual amino acid digestibility and DIAAS. The protocol was validated using in vivo data obtained by digesting the same food samples, and the results showed a high degree of agreement, confirming its relevance for nutritional assessments. To establish the DIAAS of a protein source, nonabsorbable peptides and proteins are precipitated after in vitro digestion and the resulting absorbable fraction is analyzed using ultrahigh-performance liquid chromatography with ultraviolet detection (to assess total and individual amino acids). Two alternative quantification strategies are also described: protein titration using the Kjeldahl method (to assess total nitrogen) and spectrophotometric analysis with o-phthalaldehyde (to assess total amino groups). Both alternative methods are valid only for the calculation of total digestibility and the proxy-digestible indispensable amino acid ratio, which gives an approximation of the DIAAS of the tested protein sources. Compared to existing approaches, this protocol is suitable for routine application in nutrition and food science laboratories. The preparatory steps take ~6 d, whereas the full workflow can be completed in triplicate in ~8 d. Analysis of the digesta takes an additional 3-5 d, depending on the method. The procedure requires only standard laboratory equipment and reagents and can be performed by anyone with basic training in biochemistry or a related discipline.