There is an increasing appreciation for the heterogeneous nature of extracellular vesicles (EVs). In addition, two nonvesicular extracellular nanoparticles (NVEPs), exomeres and supermeres, have been discovered recently that are enriched in many cargo previously ascribed to EVs. The EV field has largely focused on EV isolation and characterization, while studies on NVEPs are limited. At this juncture, it is critically important to have robust and reliable methods to separate distinct populations of EVs and NVEPs to assign cargo to their correct carrier. Here, we provide a comprehensive step-by-step protocol for sequential isolation of large and small EVs, nonvesicular fractions, exomeres and supermeres from the same starting material. We describe in detail the use of differential ultracentrifugation, filtration, concentration and high-resolution density-gradient fractionation to obtain purified fractions of distinct populations of EVs and NVEPs. This protocol allows assignment and enrichment of a biomolecule of interest to its specific extracellular compartment. Compared to other isolation methods, our protocol has unique advantages, including high purity and reproducibility, with minimal expertise required. The protocol can be applied to purification of EVs and NVEPs from cell culture medium and human plasma and requires ~72 h to complete. Adoption of this protocol will help translational investigators identify potential circulating biomarkers and therapeutic targets for a host of human diseases and allow basic scientists to better understand EV and NVEP biogenesis and function. Overall, this protocol will allow those interested in isolating EVs and extracellular particles to advance scientific inquiry to answer outstanding questions in the field.

Human early development sets the stage for embryonic and adult life but remains difficult to investigate. A solution came from the ability of stem cells to organize into structures resembling preimplantation embryos-blastocysts-that we termed blastoids. This embryo model is available in unlimited numbers and could thus support scientific and medical advances. However, its predictive power depends on how faithfully it recapitulates the blastocyst. Here, we describe how we formed human blastoids that (1) efficiently achieve the morphology of the blastocyst and (2) form lineages according to the pace and sequence of blastocyst development, (3) ultimately forming cells that transcriptionally reflect the blastocyst (preimplantation stage). We employ three different commercially available 96- and 24-well microwell plates with results similar to our custom-made ones, and show that blastoids form in clinical in vitro fertilization medium and can be cryopreserved for shipping. Finally, we explain how blastoids replicate the directional process of implantation into endometrial organoids, specifically when these are hormonally stimulated. It takes 4 d for human blastoids to form and 10 d to prepare the endometrial implantation assay, and we have cultured blastoids up to 6 d (time-equivalent of day 13). On the basis of our experience, we anticipate that a person with ~1 year of human pluripotent stem cell culture experience and of organoid culture should be able to perform the protocol. Altogether, blastoids offer an opportunity to establish scientific and biomedical discovery programs for early pregnancy, and an ethical alternative to the use of embryos.

Skeletal muscle is a complex tissue composed of multinucleated myofibers responsible for force generation that are supported by multiple cell types. Many severe and lethal disorders affect skeletal muscle; therefore, engineering models to reproduce such cellular complexity and function are instrumental for investigating muscle pathophysiology and developing therapies. Here, we detail the modular 3D bioengineering of multilineage skeletal muscles from human induced pluripotent stem cells, which are first differentiated into myogenic, neural and vascular progenitor cells and then combined within 3D hydrogels under tension to generate an aligned myofiber scaffold containing vascular networks and motor neurons. 3D bioengineered muscles recapitulate morphological and functional features of human skeletal muscle, including establishment of a pool of cells expressing muscle stem cell markers. Importantly, bioengineered muscles provide a high-fidelity platform to study muscle pathology, such as emergence of dysmorphic nuclei in muscular dystrophies caused by mutant lamins. The protocol is easy to follow for operators with cell culture experience and takes between 9 and 30 d, depending on the number of cell lineages in the construct. We also provide examples of applications of this advanced platform for testing gene and cell therapies in vitro, as well as for in vivo studies, providing proof of principle of its potential as a tool to develop next-generation neuromuscular or musculoskeletal therapies.

Natural sequence variation within mitochondrial DNA (mtDNA) contributes to human phenotypes and may serve as natural genetic markers in human cells for clonal and lineage tracing. We recently developed a single-cell multi-omic approach, called 'mitochondrial single-cell assay for transposase-accessible chromatin with sequencing' (mtscATAC-seq), enabling concomitant high-throughput mtDNA genotyping and accessible chromatin profiling. Specifically, our technique allows the mitochondrial genome-wide inference of mtDNA variant heteroplasmy along with information on cell state and accessible chromatin variation in individual cells. Leveraging somatic mtDNA mutations, our method further enables inference of clonal relationships among native ex vivo-derived human cells not amenable to genetic engineering-based clonal tracing approaches. Here, we provide a step-by-step protocol for the use of mtscATAC-seq, including various cell-processing and flow cytometry workflows, by using primary hematopoietic cells, subsequent single-cell genomic library preparation and sequencing that collectively take ~3-4 days to complete. We discuss experimental and computational data quality control metrics and considerations for the extension to other mammalian tissues. Overall, mtscATAC-seq provides a broadly applicable platform to map clonal relationships between cells in human tissues, investigate fundamental aspects of mitochondrial genetics and enable additional modes of multi-omic discovery.

Acoustic tweezers provide an effective means for manipulating single cells and particles in a high-throughput, precise, selective and contact-free manner. The adoption of acoustic tweezers in next-generation cellular assays may advance our understanding of biological systems. Here we present a comprehensive set of instructions that guide users through device fabrication, instrumentation setup and data acquisition to study single cells with an experimental throughput that surpasses traditional methods, such as atomic force microscopy and micropipette aspiration, by several orders of magnitude. With acoustic tweezers, users can conduct versatile experiments that require the trapping, patterning, pairing and separation of single cells in a myriad of applications ranging across the biological and biomedical sciences. This procedure is widely generalizable and adaptable for investigations in materials and physical sciences, such as the spinning motion of colloids or the development of acoustic-based quantum simulations. Overall, the device fabrication requires ~12 h, the experimental setup of the acoustic tweezers requires 1-2 h and the cell manipulation experiment requires ~30 min to complete. Our protocol is suitable for use by interdisciplinary researchers in biology, medicine, engineering and physics.

Micro Capture-C (MCC) is a chromatin conformation capture (3C) method for visualizing reproducible three-dimensional contacts of specified regions of the genome at base pair resolution. These methods are an established family of techniques that use proximity ligation to assay the topology of chromatin. MCC can generate data at substantially higher resolution than previous techniques through multiple refinements of the 3C method. Using a sequence agnostic nuclease, the maintenance of cellular integrity and full sequencing of the ligation junctions, MCC achieves subnucleosomal levels of resolution, which can be used to reveal transcription factor binding sites analogous to DNAse I footprinting. Gene dense regions, close-range enhancer-promoter contacts, individual enhancers within super-enhancers and multiple other types of loci or regulatory regions that were previously challenging to assay with conventional 3C techniques, are readily observed using MCC. MCC requires training in common molecular biology techniques and bioinformatics to perform the experiment and analyze the data. The protocol can be expected to be completed in a 3 week timeframe for experienced molecular biologists.

Atropisomers are molecules whose stereogenicity arises from restricted rotation about a single bond. They are of current importance because of their applications in catalysis, medicine and materials science. The defining feature of atropisomeric molecules is that their stereoisomers are related to one another by bond rotation: as a result, evaluating their configurational stability (i.e., the rate at which their stereoisomers interconvert) is central to any work in this area. Important atropisomeric scaffolds include C-C linked biaryls, such as the ligand BINAP and the drug vancomycin, and C-N linked amine derivatives such as the drug telenzepine. This article focuses on the three most widely used experimental methods that are available to measure the rate of racemization in atropisomers, namely: (i) kinetic analysis of the racemization of an enantioenriched sample, (ii) dynamic HPLC and (iii) variable-temperature NMR. For each technique, an explanation of the theory is set out, followed by a detailed experimental procedure. A discussion is also included of which technique to try when confronted with a new molecular structure whose properties are not yet known. None of the three procedures require complex experimental techniques, and all can be performed by using standard analytical equipment (NMR and HPLC). The time taken to determine a racemization rate depends on which experimental method is required, but for a new compound it is generally possible to measure a racemization rate in <1 d.

Spheroid culture systems have allowed in vitro propagation of cells unable to grow in canonical cell culturing conditions, and may capture cellular contexts that model tumor growth better than current model systems. The insights gleaned from genome-wide clustered regularly interspaced short palindromic repeat (CRISPR) screening of thousands of cancer cell lines grown in conventional culture conditions illustrate the value of such CRISPR pooled screens. It is clear that similar genome-wide CRISPR screens of three-dimensional spheroid cultures will be important for future biological discovery. Here, we present a protocol for genome-wide CRISPR screening of three-dimensional neurospheres. While many in-depth protocols and discussions have been published for more typical cell lines, few detailed protocols are currently available in the literature for genome-wide screening in spheroidal cell lines. For those who want to screen such cell lines, and particularly neurospheres, we provide a step-by-step description of assay development tests to be performed before screening, as well as for the screen itself. We highlight considerations of variables that make these screens distinct from, or similar to, typical nonspheroid cell lines throughout. Finally, we illustrate typical outcomes of neurosphere genome-wide screens, and how neurosphere screens typically produce slightly more heterogeneous signal distributions than more canonical cancer cell lines. Completion of this entire protocol will take 8-12 weeks from the initial assay development tests to deconvolution of the sequencing data.

In image-based profiling, software extracts thousands of morphological features of cells from multi-channel fluorescence microscopy images, yielding single-cell profiles that can be used for basic research and drug discovery. Powerful applications have been proven, including clustering chemical and genetic perturbations on the basis of their similar morphological impact, identifying disease phenotypes by observing differences in profiles between healthy and diseased cells and predicting assay outcomes by using machine learning, among many others. Here, we provide an updated protocol for the most popular assay for image-based profiling, Cell Painting. Introduced in 2013, it uses six stains imaged in five channels and labels eight diverse components of the cell: DNA, cytoplasmic RNA, nucleoli, actin, Golgi apparatus, plasma membrane, endoplasmic reticulum and mitochondria. The original protocol was updated in 2016 on the basis of several years' experience running it at two sites, after optimizing it by visual stain quality. Here, we describe the work of the Joint Undertaking for Morphological Profiling Cell Painting Consortium, to improve upon the assay via quantitative optimization by measuring the assay's ability to detect morphological phenotypes and group similar perturbations together. The assay gives very robust outputs despite various changes to the protocol, and two vendors' dyes work equivalently well. We present Cell Painting version 3, in which some steps are simplified and several stain concentrations can be reduced, saving costs. Cell culture and image acquisition take 1-2 weeks for typically sized batches of ≤20 plates; feature extraction and data analysis take an additional 1-2 weeks.This protocol is an update to Nat. Protoc. 11, 1757-1774 (2016): https://doi.org/10.1038/nprot.2016.105.

First envisioned for determining crystalline structures, ptychography has become a useful imaging tool for microscopists. However, ptychography remains underused by biomedical researchers due to its limited resolution and throughput in the visible light regime. Recent developments of spatial- and Fourier-domain ptychography have successfully addressed these issues and now offer the potential for high-resolution, high-throughput optical imaging with minimal hardware modifications to existing microscopy setups, often providing an excellent trade-off between resolution and field of view inherent to conventional imaging systems, giving biomedical researchers the best of both worlds. Here, we provide extensive information to enable the implementation of ptychography by biomedical researchers in the visible light regime. We first discuss the intrinsic connections between spatial-domain coded ptychography and Fourier ptychography. A step-by-step guide then provides the user instructions for developing both systems with practical examples. In the spatial-domain implementation, we explain how a large-scale, high-performance blood-cell lens can be made at negligible expense. In the Fourier-domain implementation, we explain how adding a low-cost light source to a regular microscope can improve the resolution beyond the limit of the objective lens. The turnkey operation of these setups is suitable for use by professional research laboratories, as well as citizen scientists. Users with basic experience in optics and programming can build the setups within a week. The do-it-yourself nature of the setups also allows these procedures to be implemented in laboratory courses related to Fourier optics, biomedical instrumentation, digital image processing, robotics and capstone projects.

Base editing is a powerful CRISPR-based technology for introducing precise substitutions into the genome. This technology greatly advances mutagenesis possibilities in vivo, particularly in zebrafish, for which the generation of precise point mutations is still challenging. Zebrafish have emerged as an important model for genetic studies and in vivo disease modeling. With the development of different base editor variants that recognize protospacer-adjacent motifs (PAMs) other than the classical 5'-NGG-3' PAM, it is now possible to design and test several guide RNAs to find the most efficient way to precisely introduce the desired substitution. Here, we describe the experimental design strategies and protocols for cytosine base editing in zebrafish, from guide RNA design and selection of base editor variants to generation of the zebrafish mutant line carrying the substitution of interest. By using co-selection by introducing a loss-of-function mutation in genes necessary for the formation of pigments, injected embryos with highly efficient base editing can be directly analyzed to determine the phenotypic impact of the targeted substitution. The generation of mutant embryos after base editor injections in zebrafish can be completed within 2 weeks.

The 2022 Nobel Prize in Chemistry was awarded to Professors K. Barry Sharpless, Morten Meldal and Carolyn Bertozzi for their pioneering roles in the advent of click chemistry. Sharpless and Meldal worked to develop the canonical click reaction-the copper-catalyzed azide-alkyne cycloaddition-while Bertozzi opened new frontiers with the creation of the bioorthogonal strain-promoted azide-alkyne cycloaddition. These two reactions have revolutionized chemical and biological science by facilitating selective, high yielding, rapid and clean ligations and by providing unprecedented ways to manipulate living systems. Click chemistry has affected every aspect of chemistry and chemical biology, but few disciplines have been impacted as much as radiopharmaceutical chemistry. The importance of speed and selectivity in radiochemistry make it an almost tailor-made application of click chemistry. In this Perspective, we discuss the ways in which the copper-catalyzed azide-alkyne cycloaddition, the strain-promoted azide-alkyne cycloaddition and a handful of 'next-generation' click reactions have transformed radiopharmaceutical chemistry, both as tools for more efficient radiosyntheses and as linchpins of technologies that have the potential to improve nuclear medicine.

Neural circuits are assembled from an enormous variety of neuronal cell types. Although significant advances have been made in classifying neurons on the basis of morphological, molecular and electrophysiological properties, understanding how this diversity contributes to brain function during behavior has remained a major experimental challenge. Here, we present an extension to our previous protocol, in which we describe the technical procedures for performing juxtacellular opto-tagging of single neurons in freely moving mice by using Channelrhodopsin-2-expressing viral vectors. This method allows one to selectively target molecularly defined cell classes for in vivo single-cell recordings. The targeted cells can be labeled via juxtacellular procedures and further characterized via post-hoc morphological and molecular analysis. In its current form, the protocol allows multiple recording and labeling attempts to be performed within individual animals, by means of a mechanical pipette micropositioning system. We provide proof-of-principle validation of this technique by recording from Calbindin-positive pyramidal neurons in the mouse hippocampus during spatial exploration; however, this approach can easily be extended to other behaviors and cortical or subcortical areas. The procedures described here, from the viral injection to the histological processing of brain sections, can be completed in ~4-5 weeks.This protocol is an extension to: Nat. Protoc. 9, 2369-2381 (2014): https://doi.org/10.1038/nprot.2014.161.

The ability to experimentally measure cell proliferation is the basis for understanding the sources of cells that drive organ development, tissue regeneration and repair. Recently, we generated a genetic approach to detect cell proliferation: we used genetic lineage-tracing technologies to achieve seamless recording of in vivo cell proliferation in a tissue-specific manner. We provide a detailed protocol (generation of mouse lines, characterization of mouse lines, mouse line crossing and cell-proliferation tracing) for using this genetic system to study cell proliferation. This cell-proliferation tracing system, which we term 'ProTracer' (Proliferation Tracer), permits lifelong noninvasive monitoring of cell proliferation of specific cell lineages in live animals. Compared with other short-term strategies that require execution of animals, ProTracer does not require sampling or animal sacrifice for tissue processing. To highlight these features, we used ProTracer to study the proliferation of hepatocytes during liver homeostasis and after tissue injury in mice. We show that the protocol is applicable to study any in vivo cell proliferation, which takes ~9 months to finish from mouse generation to data analysis. This protocol can easily be carried out by researchers skilled in mouse-related experiments.

Enzymes are outstanding natural catalysts with exquisite 3D structures, initiating countless life-sustaining biotransformations in living systems. The flexible structure of an enzyme, however, is highly susceptible to non-physiological environments, which greatly limits its large-scale industrial applications. Seeking suitable supports to immobilize fragile enzymes is one of the most efficient routes to ameliorate the stability problem. This protocol imparts a new bottom-up strategy for enzyme encapsulation using a hydrogen-bonded organic framework (HOF-101). In short, the surface residues of the enzyme can trigger the nucleation of HOF-101 around its surface through the hydrogen-bonded biointerface. As a result, a series of enzymes with different surface chemistries are able to be encapsulated within a highly crystalline HOF-101 scaffold, which has long-range ordered mesochannels. The details of experimental procedures are described in this protocol, which involve the encapsulating method, characterizations of materials and biocatalytic performance tests. Compared with other immobilization methods, this enzyme-triggering HOF-101 encapsulation is easy to operate and affords higher loading efficiency. The formed HOF-101 scaffold has an unambiguous structure and well-arranged mesochannels, favoring mass transfer and understanding of the biocatalytic process. It takes ~13.5 h for successful synthesis of enzyme-encapsulated HOF-101, 3-4 d for characterizations of materials and ~4 h for the biocatalytic performance tests. In addition, no specific expertise is necessary for the preparation of this biocomposite, although the high-resolution imaging requires a low-electron-dose microscope technology. This protocol can provide a useful methodology to efficiently encapsulate enzymes and design biocatalytic HOF materials.

Ultraflexible microelectrode arrays (MEAs) that can stably record from a large number of neurons after their chronic implantation offer opportunities for understanding neural circuit mechanisms and developing next-generation brain-computer interfaces. The implementation of ultraflexible MEAs requires their reliable implantation into deep brain tissues in a minimally invasive manner, as well as their precise integration with optogenetic tools to enable the simultaneous recording of neural activity and neuromodulation. Here, we describe the process for the preparation of elastocapillary self-assembled ultraflexible MEAs, their use in combination with adeno-associated virus vectors carrying opsin genes and promoters to form an optrode probe and their in vivo experimental use in the brains of rodents, enabling electrophysiological recordings and optical modulation of neuronal activity over long periods of time (on the order of weeks to months). The procedures, including device fabrication, probe assembly and implantation, can be completed within 3 weeks. The protocol is intended to facilitate the applications of ultraflexible MEAs for long-term neuronal activity recording and combined electrophysiology and optogenetics. The protocol requires users with expertise in clean room facilities for the fabrication of ultraflexible MEAs.

Biological research and diagnostic applications normally require analysis of trace analytes in biofluids. Although considerable advancements have been made in developing precise molecular assays, the trade-off between sensitivity and ability to resist non-specific adsorption remains a challenge. Here, we describe the implementation of a testing platform based on a molecular-electromechanical system (MolEMS) immobilized on graphene field-effect transistors. A MolEMS is a self-assembled DNA nanostructure, containing a stiff tetrahedral base and a flexible single-stranded DNA cantilever. Electromechanical actuation of the cantilever modulates sensing events close to the transistor channel, improving signal-transduction efficiency, while the stiff base prevents non-specific adsorption of background molecules present in biofluids. A MolEMS realizes unamplified detection of proteins, ions, small molecules and nucleic acids within minutes and has a limit of detection of several copies in 100 μl of testing solution, offering an assay methodology with wide-ranging applications. In this protocol, we provide step-by-step procedures for MolEMS design and assemblage, sensor manufacture and operation of a MolEMS in several applications. We also describe adaptations to construct a portable detection platform. It takes ~18 h to construct the device and ~4 min to finish the testing from sample addition to result.

The zebrafish is a powerful model system for studying animal development, for modeling genetic diseases, and for large-scale in vivo functional genetics. Because of its ease of use and its high efficiency in targeted gene perturbation, CRISPR-Cas9 has recently gained prominence as the tool of choice for genetic manipulation in zebrafish. However, scaling up the technique for high-throughput in vivo functional genetics has been a challenge. We recently developed a method, Multiplexed Intermixed CRISPR Droplets (MIC-Drop), that makes large-scale CRISPR screening in zebrafish possible. Here, we outline the step-by-step protocol for performing functional genetic screens in zebrafish by using MIC-Drop. MIC-Drop uses multiplexed single-guide RNAs to generate biallelic mutations in injected zebrafish embryos, allowing genetic screens to be performed in F0 animals. Combining microfluidics and DNA barcoding enables simultaneous targeting of tens to hundreds of genes from a single injection needle, while also enabling retrospective and rapid identification of the genotype responsible for an observed phenotype. The primary target audiences for MIC-Drop are developmental biologists, zebrafish geneticists, and researchers interested in performing in vivo functional genetic screens in a vertebrate model system. MIC-Drop will also prove useful in the hands of chemical biologists seeking to identify targets of small molecules that cause phenotypic changes in zebrafish. By using MIC-Drop, a typical screen of 100 genes can be conducted within 2-3 weeks by a single user.

The advent of distributed biomanufacturing platforms promises to increase agility in biologic production and expand access by reducing reliance on refrigerated supply chains. However, such platforms are not capable of robustly producing glycoproteins, which represent the majority of biologics approved or in development. To address this limitation, we developed cell-free technologies that enable rapid, modular production of glycoprotein therapeutics and vaccines from freeze-dried Escherichia coli cell lysates. Here, we describe a protocol for generation of cell-free lysates and freeze-dried reactions for on-demand synthesis of desired glycoproteins. The protocol includes construction and culture of the bacterial chassis strain, cell-free lysate production, assembly of freeze-dried reactions, cell-free glycoprotein synthesis, and glycoprotein characterization, all of which can be completed in one week or less. We anticipate that cell-free technologies, along with this comprehensive user manual, will help accelerate development and distribution of glycoprotein therapeutics and vaccines.

CRISPR-Cas nuclease systems, base editors, and CRISPR activation have greatly advanced plant genome engineering. However, the combinatorial approaches for multiplexed orthogonal genome editing and transcriptional regulation were previously unexploited in plants. We have recently established a single Cas9 protein-based CRISPR-Combo platform, enabling efficient multiplexed orthogonal genome editing (double-strand break-mediated genome editing or base editing) and transcriptional activation in plants via engineering the single guide RNA (sgRNA) structure. Here, we provide step-by-step instructions for constructing CRISPR-Combo systems for speed breeding of transgene-free, genome-edited Arabidopsis plants and enhancing rice regeneration with more heritable targeted mutations in a hormone-free manner. We also provide guidance on designing efficient sgRNA, Agrobacterium-mediated transformation of Arabidopsis and rice, rice regeneration without exogenous plant hormones, gene editing evaluation and visual identification of transgene-free Arabidopsis plants with high editing activity. With the use of this protocol, it takes ~2 weeks to establish the CRISPR-Combo systems, 4 months to obtain transgene-free genome-edited Arabidopsis plants and 4 months to obtain rice plants with enrichment of heritable targeted mutations by hormone-free tissue culture.

Luminescent sensing materials are attractive for environmental analysis due to their potential for high selectivity, excellent sensitivity and rapid (even instantaneous) response towards targeted analytes in diverse sample matrices. Many types of analytes have been detected in samples of wastewater for environmental protection, reagents and products in industrial production of drugs and pesticides, and biological markers in blood and urine for early diagnosis. It is still challenging, however, to develop appropriate materials with optimal sensing function for a targeted analyte. Here we synthesize metal-organic frameworks (MOFs) bearing multiple luminescent centers, such as metal cations (for example, Eu3+ and Tb3+), organic ligands and guests, which are chosen for optimal selectivity for the analytes of interest, including industrial synthetic intermediates and chiral drugs. Interaction between the metal node, ligand, guest and analyte results in a complex system with different luminescence properties compared with the porous MOF on its own. The operation time for the synthesis is usually less than 4 h; the quick screening for sensitivity and selectivity takes ~0.5 h and includes steps to optimize the energy levels and spectrum parameters. It can be used to accelerate the discovery of advanced sensing materials for practical applications.

Studying the electrical properties of individual proteins is a prominent research area in the field of bioelectronics. Electron tunnelling or quantum mechanical tunnelling (QMT) probes can act as powerful tools for investigating the electrical properties of proteins. However, current fabrication methods for these probes often have limited reproducibility, unreliable contact or inadequate binding of proteins onto the electrodes, so better solutions are required. Here, we detail a generalizable and straightforward set of instructions for fabricating simple, nanopipette-based, tunnelling probes, suitable for measuring conductance in single proteins. Our QMT probe is based on a high-aspect-ratio dual-channel nanopipette that integrates a pair of gold tunneling electrodes with a gap of less than 5 nm, fabricated via the pyrolytic deposition of carbon followed by the electrochemical deposition of gold. The gold tunneling electrodes can be functionalized using an extensive library of available surface modifications to achieve single-protein-electrode contact. We use a biotinylated thiol modification, in which a biotin-streptavidin-biotin bridge is used to form the single-protein junction. The resulting protein-coupled QMT probes enable the stable electrical measurement of the same single protein in solution for up to several hours. We also describe the analysis method used to interpret time-dependent single-protein conductance measurements, which can provide essential information for understanding electron transport and exploring protein dynamics. The total time required to complete the protocol is ~33 h and it can be carried out by users trained in less than 24 h.

Mass spectrometry imaging (MSI) enables the chemical mapping of molecules and elements in a label-free, high-throughput manner. Because this approach can be accomplished rapidly, it also enables chemical changes to be monitored. Here, we describe a protocol for MSI with subcellular spatial resolution. This is achieved by using a microlensed fiber, which is made by grinding an optical fiber. It is a universal and economic technique that can be adapted to most laser-based mass spectrometry methods. In this protocol, the output of laser radiation from the microlensed fiber causes laser ablation of the sample, and the resulting plume is mass spectrometrically analyzed. The microlensed fiber can be used with matrix-assisted laser desorption ionization, laser desorption ionization, laser ablation electrospray desorption ionization and laser ablation inductively coupled plasma, in each case to achieve submicroscale imaging of single cells and biological tissues. This report provides a detailed introduction of the microlensed fiber design and working principles, sample preparation, microlensed fiber ion source setup and multiple MSI platforms with different kinds of mass spectrometers. A researcher with a little background (such as a trained graduate student) is able to complete all the steps for the experimental setup in ~2 h, including fiber test, laser coupling and ion source modification. The imaging time spent mainly depends on the size of the imaging area. It is suggested that most existing laser-based MSI platforms, especially atmospheric pressure applications, can achieve breakthroughs in spatial resolution by introducing a microlensed fiber module.

Little is known about how social factors contribute to neurobiology or neuropsychiatric disorders. The use of mice allows one to probe the neurobiological bases of social interaction, offering the genetic diversity and versatility to identify cell types and neural circuits of social behavior. However, mice typically show lower social motivation compared with rats, leading to the question of whether mice should be used to model complex social behaviors displayed by humans. Studies on mouse social behavior often rely on measures such as time spent in contact with a social partner or preference for a social-paired context, but fail to assess volitional (subject-controlled) rewarding social interaction. Here, we describe a volitional social self-administration and choice model that is an extension of our previous work on rats. Using mice, we systematically compared female adolescent and adult C57BL/6 mice and outbred CD1 mice, showing that operant social self-administration, social seeking during periods of isolation and choice of social interaction over palatable food is significantly stronger in female CD1 mice than in female C57BL/6J mice, independently of age. We describe the requirements for building the social self-administration and choice apparatus and we provide guidance for studying the role of operant social reward in mice. We also discuss its use to study brain mechanisms of operant social reward, potentially extending its application to mouse models of neuropsychiatric disorders. The training commonly requires ~4 weeks for stable social self-administration and 3-4 additional weeks for tests, including social seeking and choice.

Purification of membrane proteins for biochemical and structural studies is commonly achieved by recombinant overexpression in heterologous cell lines. However, many membrane proteins do not form a functional complex in a heterologous system, and few methods exist to purify sufficient protein from a native source for use in biochemical, biophysical and structural studies. Here, we provide a detailed protocol for the isolation of membrane protein complexes from transgenic Caenorhabditis elegans. We describe how to grow a genetically modified C. elegans line in abundance using standard laboratory equipment, and how to optimize purification conditions on a small scale using fluorescence-detection size-exclusion chromatography. Optimized conditions can then be applied to a large-scale preparation, enabling the purification of adequate quantities of a target protein for structural, biochemical and biophysical studies. Large-scale worm growth can be accomplished in ~9 d, and each optimization experiment can be completed in less than 1 d. We have used these methods to isolate the transmembrane channel-like protein 1 complex, as well as three additional protein complexes (transmembrane-like channel 2, lipid transfer protein and 'Protein S'), from transgenic C. elegans, demonstrating the utility of this approach in purifying challenging, low-abundance membrane protein complexes.

Electrochemical molecular intercalation of layered semiconducting crystals with organic cations followed by ultrasonic exfoliation has proven to be an effective approach to producing a rich family of organic/inorganic hybrid superlattices and high-quality, solution-processable 2D semiconductors. A traditional method for exfoliating 2D crystals relies on the intercalation of inorganic alkali metal cations. The organic cations (e.g., alkyl chain-substituted quaternary ammonium cations) are much larger than their inorganic counterparts, and the bulky molecular structure endows distinct intercalation and exfoliation chemistry, as well as molecular tunability. By using this protocol, many layered 2D crystals (including graphene, black phosphorus and versatile metal chalcogenides) can be electrochemically intercalated with organic quaternary alkylammonium cations. Subsequent solution-phase exfoliation of the intercalated compounds is realized by regular bath sonication for a short period (5-30 min) to produce free-standing, thin 2D nanosheets. It is also possible to graft additional ligands on the nanosheet surface. The thickness of the exfoliated nanosheets can be measured by using atomic force microscopy and Raman spectroscopy. Modifying the chemical structure and geometrical configuration of alkylammonium cations results in different exfoliation behavior and a family of versatile organic/inorganic hybrid superlattices with tunable physical/chemical properties. The whole protocol takes ~6 h for the successful production of stable, ultrathin 2D nanosheet dispersion in solution and another 11 h for depositing thin films and transferring them onto an arbitrary surface. This protocol does not require expertise beyond basic electrochemistry knowledge and conventional colloidal nanocrystal synthesis and processing.

Circular RNAs (circRNAs) have important roles in regulating developmental processes and disease progression. As most circRNA sequences are highly similar to their cognate linear transcripts, the current short-read sequencing-based methods rely on the back-spliced junction signal for distinguishing circular and linear reads, which does not allow circRNAs' full-length structure to be effectively reconstructed. Here we describe a long-read sequencing-based protocol, CIRI-long, for the detection of full-length circular RNAs. The CIRI-long protocol combines rolling circular reverse transcription and nanopore sequencing to capture full-length circRNA sequences. After poly(A) tailing, RNase R treatment, and size selection of polymerase chain reaction products, CIRI-long achieves an increased percentage (6%) of circular reads in the constructed library, which is 20-fold higher compared with previous Illumina-based strategies. This method can be applied in cell lines or tissue samples, enabling accurate detection of full-length circRNAs in the range of 100-3,000 bp. The entire protocol can be completed in 1 d, and can be scaled up for large-scale analysis using the nanopore barcoding kit and PromethION sequencing device. CIRI-long can serve as an effective and user-friendly protocol for characterizing full-length circRNAs, generating direct and convincing evidence for the existence of detected circRNAs. The analytical pipeline offers convenient functions for identification of full-length circRNA isoforms and integration of multiple datasets. The assembled full-length transcripts and their splicing patterns provide indispensable information to explore the biological function of circRNAs.

Despite advances in automated liquid handling and microfluidics, preparing samples for RNA sequencing at scale generally requires expensive equipment, which is beyond the reach of many academic laboratories. Manual sample preparation remains a slow, expensive and error-prone process. Here, we describe a low-cost, semi-automated pipeline to extract cell-free RNA using one of two commercially available, inexpensive and open-source robotic systems: the Opentrons OT1.0 or OT2.0. Like many RNA isolation protocols, ours can be decomposed into three subparts: RNA extraction, DNA digestion and RNA cleaning and concentration. RT-qPCR data using a synthetic spike-in confirms comparable RNA quality to the gold standard, manual sample processing. The semi-automated pipeline also shows improvement in sample throughput (+12×), time spent (-11×), cost (-3×) and biohazardous waste produced (-4×) compared with its manual counterpart. This protocol enables cell-free RNA extraction from 96 samples simultaneously in 4.5 h; in practice, this dramatically improves the time to results, as we recently demonstrated. Importantly, any laboratory already has most of the parts required (manual pipette and corresponding tips and kits for RNA isolation, cleaning and concentration) to build a semi-automated sample processing pipeline of their own and would only need to purchase or three-dimensionally print a few extra parts (US$5.5 K-12 K in total). This pipeline is also generalizable for many nucleic acid extraction applications, thereby increasing the scale of studies, which can be performed in small research laboratories.

Induced pluripotent stem cell-derived brain organoids enable the developmental complexities of the human brain to be deconstructed. During embryogenesis, optic vesicles (OVs), the eye primordium attached to the forebrain, develop from diencephalon. However, most 3D culturing methods generate either brain or retinal organoids individually. Here we describe a protocol to generate organoids with both forebrain entities, which we call OV-containing brain organoids (OVB organoids). In this protocol, we first induce neural differentiation (days 0-5) and collect neurospheres, which we culture in a neurosphere medium to initiate their patterning and further self-assembly (days 5-10). Then, upon transfer to spinner flasks containing OVB medium (days 10-30), neurospheres develop into forebrain organoids with one or two pigmented dots restricted to one pole, displaying forebrain entities of ventral and dorsal cortical progenitors and preoptic areas. Further long-term culture results in photosensitive OVB organoids constituting complementary cell types of OVs, including primitive corneal epithelial and lens-like cells, retinal pigment epithelia, retinal progenitor cells, axon-like projections and electrically active neuronal networks. OVB organoids provide a system to help dissect interorgan interactions between the OVs as sensory organs and the brain as a processing unit, and can help model early eye patterning defects, including congenital retinal dystrophy. To conduct the protocol, experience in sterile cell culture and maintenance of human induced pluripotent stem cells is essential; theoretical knowledge of brain development is advantageous. Furthermore, specialized expertise in 3D organoid culture and imaging for the analysis is needed.

Antibodies play an important role in the immune system by binding to molecules called antigens at their respective epitopes. These interfaces or epitopes are structural entities determined by the interactions between an antibody and an antigen, making them ideal systems to analyze by using docking programs. Since the advent of high-throughput antibody sequencing, the ability to perform epitope mapping using only the sequence of the antibody has become a high priority. ClusPro, a leading protein-protein docking server, together with its template-based modeling version, ClusPro-TBM, have been re-purposed to map epitopes for specific antibody-antigen interactions by using the Antibody Epitope Mapping server (AbEMap). ClusPro-AbEMap offers three different modes for users depending on the information available on the antibody as follows: (i) X-ray structure, (ii) computational/predicted model of the structure or (iii) only the amino acid sequence. The AbEMap server presents a likelihood score for each antigen residue of being part of the epitope. We provide detailed information on the server's capabilities for the three options and discuss how to obtain the best results. In light of the recent introduction of AlphaFold2 (AF2), we also show how one of the modes allows users to use their AF2-generated antibody models as input. The protocol describes the relative advantages of the server compared to other epitope-mapping tools, its limitations and potential areas of improvement. The server may take 45-90 min depending on the size of the proteins.

Strategies for the covalent immobilization of enzymes depend on the type of functional group selected to perform the coupling reaction, and on the relative importance of selectivity, loading capacity, immobilization time and activity/stability of the resulting immobilized preparation. However, no single strategy is applicable for all covalent immobilization methods or can meet all these criteria, exemplifying the challenge of introducing a versatile process broadly compatible with the residues on the surface of proteins and the functional groups of common linkers. Here, we describe the use of isocyanide-based multi-component reactions for the carrier-bound and carrier-free covalent immobilization of enzymes. Isocyanide-based multi-component reactions can accept a wide variety of functional groups such as epoxy, acid, amine and aldehyde, as well as many commercially available bi-functional linkers, and are therefore suitable for either covalent coupling of enzymes on a solid support or intermolecular cross-linking of enzymes. In this strategy, an isocyanide is directly added to the reaction medium, the enzyme supplies either the exposed amine or carboxylic acid groups, and the support (in carrier-bound immobilization) or the bi-functional cross-linking agent (in carrier-free immobilization) provides another reactive functional group. The protocol offers operational simplicity, high efficiency and a notable reduction in time over alternative strategies, and can be performed by users with expertise in chemistry or biology. The immobilization reactions typically require 1-24 h.

RNA-sequencing (RNA-seq) has become an increasingly cost-effective technique for molecular profiling and immune characterization of tumors. In the past decade, many computational tools have been developed to characterize tumor immunity from gene expression data. However, the analysis of large-scale RNA-seq data requires bioinformatics proficiency, large computational resources and cancer genomics and immunology knowledge. In this tutorial, we provide an overview of computational analysis of bulk RNA-seq data for immune characterization of tumors and introduce commonly used computational tools with relevance to cancer immunology and immunotherapy. These tools have diverse functions such as evaluation of expression signatures, estimation of immune infiltration, inference of the immune repertoire, prediction of immunotherapy response, neoantigen detection and microbiome quantification. We describe the RNA-seq IMmune Analysis (RIMA) pipeline integrating many of these tools to streamline RNA-seq analysis. We also developed a comprehensive and user-friendly guide in the form of a GitBook with text and video demos to assist users in analyzing bulk RNA-seq data for immune characterization at both individual sample and cohort levels by using RIMA.

Understanding how proteins and materials interact is useful for evaluating the safety of biomedical micro/nanomaterials, toxicity estimation and design of nano-drugs and catalytic activity improvement of bio-inorganic functional hybrids. However, characterizing the interfacial molecular details of protein-micro/nanomaterial hybrids remains a great challenge. This protocol describes the lysine reactivity profiling-mass spectrometry strategy for determining which parts of a protein are interacting with the micro/nanomaterials. Lysine residues occur frequently on hydrophilic protein surfaces, and their reactivity is dependent on the accessibility of their amine groups. The accessibility of a lysine residue is lower when it is in contact with another object; allosteric effects resulting from this interaction might reduce or increase the reactivity of remote lysine residues. Lysine reactivity is therefore a useful indicator of protein localization orientation, interaction sequence regions, binding sites and modulated protein structures in the protein-material hybrids. We describe the optimized two-step isotope dimethyl labeling strategy for protein-material hybrids under their native and denaturing conditions in sequence. The comparative quantification results of lysine reactivity are only dependent on the native microenvironments of lysine local structures. We also highlight other critical steps including protein digestion, elution from materials, data processing and interfacial structure analysis. The two-step isotope labeling steps need ~5 h, and the whole protocol including digestion, liquid chromatography-tandem mass spectrometry, data processing and structure analysis needs ~3-5 d.

Human skeletal stem cells (hSSCs) hold tremendous therapeutic potential for developing new clinical strategies to effectively combat congenital and age-related musculoskeletal disorders. Unfortunately, refined methodologies for the proper isolation of bona fide hSSCs and the development of functional assays that accurately recapitulate their physiology within the skeleton have been lacking. Bone marrow-derived mesenchymal stromal cells (BMSCs), commonly used to describe the source of precursors for osteoblasts, chondrocytes, adipocytes and stroma, have held great promise as the basis of various approaches for cell therapy. However, the reproducibility and clinical efficacy of these attempts have been obscured by the heterogeneous nature of BMSCs due to their isolation by plastic adherence techniques. To address these limitations, our group has refined the purity of individual progenitor populations that are encompassed by BMSCs by identifying defined populations of bona fide hSSCs and their downstream progenitors that strictly give rise to skeletally restricted cell lineages. Here, we describe an advanced flow cytometric approach that utilizes an extensive panel of eight cell surface markers to define hSSCs; bone, cartilage and stromal progenitors; and more differentiated unipotent subtypes, including an osteogenic subset and three chondroprogenitors. We provide detailed instructions for the FACS-based isolation of hSSCs from various tissue sources, in vitro and in vivo skeletogenic functional assays, human xenograft mouse models and single-cell RNA sequencing analysis. This application of hSSC isolation can be performed by any researcher with basic skills in biology and flow cytometry within 1-2 days. The downstream functional assays can be performed within a range of 1-2 months.

Fast tracking of biological dynamics across multiple murine organs using the currently commercially available whole-body preclinical imaging systems is hindered by their limited contrast, sensitivity and spatial or temporal resolution. Spiral volumetric optoacoustic tomography (SVOT) provides optical contrast, with an unprecedented level of spatial and temporal resolution, by rapidly scanning a mouse using spherical arrays, thus overcoming the current limitations in whole-body imaging. The method enables the visualization of deep-seated structures in living mammalian tissues in the near-infrared spectral window, while further providing unrivalled image quality and rich spectroscopic optical contrast. Here, we describe the detailed procedures for SVOT imaging of mice and provide specific details on how to implement a SVOT system, including component selection, system arrangement and alignment, as well as the image processing methods. The step-by-step guide for the rapid panoramic (360°) head-to-tail whole-body imaging of a mouse includes the rapid visualization of contrast agent perfusion and biodistribution. The isotropic spatial resolution possible with SVOT can reach 90 µm in 3D, while alternative steps enable whole-body scans in less than 2 s, unattainable with other preclinical imaging modalities. The method further allows the real-time (100 frames per second) imaging of biodynamics at the whole-organ level. The multiscale imaging capacity provided by SVOT can be used for visualizing rapid biodynamics, monitoring responses to treatments and stimuli, tracking perfusion, and quantifying total body accumulation and clearance dynamics of molecular agents and drugs. Depending on the imaging procedure, the protocol requires 1-2 h to complete by users trained in animal handling and biomedical imaging.

Mitochondria are key bioenergetic organelles involved in many biosynthetic and signaling pathways. However, their differential contribution to specific functions of cells within complex tissues is difficult to dissect with current methods. The present protocol addresses this need by enabling the ex vivo immunocapture of cell-type-specific mitochondria directly from their tissue context through a MitoTag reporter mouse. While other available methods were developed for bulk mitochondria isolation or more abundant cell-type-specific mitochondria, this protocol was optimized for the selective isolation of functional mitochondria from medium-to-low-abundant cell types in a heterogeneous tissue, such as the central nervous system. The protocol has three major parts: First, mitochondria of a cell type of interest are tagged via an outer mitochondrial membrane eGFP by crossing MitoTag mice to a cell-type-specific Cre-driver line or by delivery of viral vectors for Cre expression. Second, homogenates are prepared from relevant tissues by nitrogen cavitation, from which tagged organelles are immunocaptured using magnetic microbeads. Third, immunocaptured mitochondria are used for downstream assays, e.g., to probe respiratory capacity or calcium handling, revealing cell-type-specific mitochondrial diversity in molecular composition and function. The MitoTag approach enables the identification of marker proteins to label cell-type-specific organelle populations in situ, elucidates cell-type-enriched mitochondrial metabolic and signaling pathways, and reveals functional mitochondrial diversity between adjacent cell types in complex tissues, such as the brain. Apart from establishing the mouse colony (6-8 weeks without import), the immunocapture protocol takes 2 h and functional assays require 1-2 h.

Conducting polymers with conjugated backbones have been widely used in electrochemical energy storage, catalysts, gas sensors and biomedical devices. In particular, two-dimensional (2D) mesoporous conducting polymers combine the advantages of mesoporous structure and 2D nanosheet morphology with the inherent properties of conducting polymers, thus exhibiting improved electrochemical performance. Despite the use of bottom-up self-assembly approaches for the fabrication of a variety of mesoporous materials over the past decades, the synchronous control of the dimensionalities and mesoporous architectures for conducting polymer nanomaterials remains a challenge. Here, we detail a simple, general and robust route for the preparation of a series of 2D mesoporous conducting polymer nanosheets with adjustable pore size (5-20 nm) and thickness (13-45 nm) and controllable morphology and composition via solution-based self-assembly. The synthesis conditions and preparation procedures are detailed to ensure the reproducibility of the experiments. We describe the fabrication of over ten high-quality 2D-ordered mesoporous conducting polymers and sandwich-structured hybrids, with tunable thickness, porosity and large specific surface area, which can serve as potential candidates for high-performance electrode materials used in supercapacitors and alkali metal ion batteries, and so on. The preparation time of the 2D-ordered mesoporous conducting polymer is usually no more than 12 h. The subsequent supercapacitor testing takes ~24 h and the Na ion battery testing takes ~72 h. The procedure is suitable for users with expertise in physics, chemistry, materials and other related disciplines.

Polymeric nanoparticles (NPs) are a promising platform for medical applications in drug delivery. However, their use as drug carriers is limited by biological (e.g., immunological) barriers after intravenous administration. Ionic liquids (ILs), formed from bulky asymmetric cations and anions, have a wide variety of physical internal and external interfacing properties. When assembled on polymeric NPs as biomaterial coatings, these external-interfacing properties can be tuned to extend their circulation half-life when intravenously injected, as well as drive biodistribution to sites of interest for selective organ accumulation. In our work, we are particularly interested in optimizing IL coatings to enable red blood cell hitchhiking in whole blood. In this protocol, we describe the preparation and physicochemical and biological characterization of choline carboxylate IL-coated polymeric NPs. The procedure is divided into five stages: (1) synthesis and characterization of choline-based ILs (1 week); (2) bare poly(lactic-co-glycolic acid) (50:50, acid terminated) Resomer 504H (PLGA) NP assembly, modified from previously established protocols, with dye encapsulation (7 h); (3) modification of the bare particles with IL coating (3 h); (4) physicochemical characterization of both PLGA and IL-PLGA NPs by dynamic light scattering, 1H nuclear magnetic resonance spectroscopy, and transmission electron microscopy (1 week); (5) ex vivo evaluation of intravenous biocompatibility (including serum-protein resistance and hemolysis) and red blood cell hitchhiking in whole BALB/c mouse blood via fluorescence-activated cell sorting (1 week). With practice and technique refinement, this protocol is accessible to late-stage graduate students and early-stage postdoctoral scientists.

Neuroinflammation has either beneficial or detrimental effects, depending on risk factors and neuron-glia interactions in neurological disorders. However, studying neuroinflammation has been challenging due to the complexity of cell-cell interactions and lack of physio-pathologically relevant neuroinflammatory models. Here, we describe our three-dimensional microfluidic multicellular human neural culture model, referred to as a 'brain-on-a-chip' (BoC). This elucidates neuron-glia interactions in a controlled manner and recapitulates pathological signatures of the major neurological disorders: dementia, brain tumor and brain edema. This platform includes a chemotaxis module offering a week-long, stable chemo-gradient compared with the few hours in other chemotaxis models. Additionally, compared with conventional brain models cultured with mixed phenotypes of microglia, our BoC can separate the disease-associated microglia out of heterogeneous population and allow selective neuro-glial engagement in three dimensions. This provides benefits of interpreting the neuro-glia interactions while revealing that the prominent activation of innate immune cells is the risk factor leading to synaptic impairment and neuronal loss, validated in our BoC models of disorders. This protocol describes how to fabricate and implement our human BoC, manipulate in real time and perform end-point analyses. It takes 2 d to set up the device and cell preparations, 1-9 weeks to develop brain models under disease conditions and 2-3 d to carry out analyses. This protocol requires at least 1 month training for researchers with basic molecular biology techniques. Taken together, our human BoCs serve as reliable and valuable platforms to investigate pathological mechanisms involving neuroinflammation and to assess therapeutic strategies modulating neuroinflammation in neurological disorders.

The ability to measure the behavior of a single molecule during a reaction implies the detection of inherent dynamic and static disordered states, which may not be represented when measuring ensemble averages. Here, we describe the building of devices with graphene-molecule-graphene single-molecule junctions integrated into an electrical circuit. These devices are simple to build and are stable, showing tolerance to mechanical changes, solution environment and voltage stimulation. The design of a conductive channel based on a single molecule enables single-molecule detection and is sensitive to variations in physical properties and chemical structures of the detected molecules. The on-chip setup of single-molecule junctions further offers complementary metal-oxide-semiconductor (CMOS) compatibility, enabling logic functions in circuit elements, as well as deciphering of reaction intermediates. We detail the experimental procedure to prepare graphene transistor arrays as a basis for single-molecule junctions and the preparation of nanogapped carboxyl-terminal graphene electrodes by using electron-beam lithography and oxygen plasma etching. We describe the basic design of a molecular bridge with desired functions and terminals to form covalent bonds with electrode arrays, via a chemical reaction, to construct stably integrated single-molecule devices with a yield of 30-50% per chip. The immobilization of the single molecules is then characterized by using inelastic electron tunneling spectra, single-molecule imaging and fluorescent spectra. The whole protocol can be implemented within 2 weeks and requires users trained in using ultra-clean laboratory facilities and the aforementioned instrumentation.

Biological signals occur over time in living cells. Yet most current approaches to interrogate biology, particularly gene expression, use destructive techniques that quantify signals only at a single point in time. A recent technological advance, termed the Retro-Cascorder, overcomes this limitation by molecularly logging a record of gene expression events in a temporally organized genomic ledger. The Retro-Cascorder works by converting a transcriptional event into a DNA barcode using a retron reverse transcriptase and then storing that event in a unidirectionally expanding clustered regularly interspaced short palindromic repeats (CRISPR) array via acquisition by CRISPR-Cas integrases. This CRISPR array-based ledger of gene expression can be retrieved at a later point in time by sequencing. Here we describe an implementation of the Retro-Cascorder in which the relative timing of transcriptional events from multiple promoters of interest is recorded chronologically in Escherichia coli populations over multiple days. We detail the molecular components required for this technology, provide a step-by-step guide to generate the recording and retrieve the data by Illumina sequencing, and give instructions for how to use custom software to infer the relative transcriptional timing from the sequencing data. The example recording is generated in 2 d, preparation of sequencing libraries and sequencing can be accomplished in 2-3 d, and analysis of data takes up to several hours. This protocol can be implemented by someone familiar with basic bacterial culture, molecular biology and bioinformatics. Analysis can be minimally run on a personal computer.

The low number of neural progenitor cells (NPCs) present in the adult and aged primate brains represents a challenge for generating high-yield and viable in vitro cultures of primary brain cells. Here we report a step-by-step approach for the fast and reproducible isolation of high-yield and viable primary brain cells, including mature neurons, immature cells and NPCs, from adult and aged macaques. We describe the anesthesia, transcardial perfusion and brain tissue preparation; the subsequent microdissection of the regions of interest and their enzymatic dissociation, leading to the separation of single cells. The cell isolation steps of our protocol can also be used for routine cell culturing, in particular for NPC expansion and differentiation, suitable for studies of hippocampal neurogenesis in the adult macaque brain. The purified primary brain cells are largely free from myelin debris and erythrocytes, paving the way for multiple downstream applications in vitro and in vivo. When combined with single-cell profiling techniques, this approach allows an unbiased and comprehensive mapping of cell states in the adult and aged macaque brain, which is needed to advance our understanding of human cognitive and neurological diseases. The neural cell isolation protocol requires 4 h and a team of four to six users with expertize in primary brain cell isolation to avoid tissue hypoxia during the time-sensitive steps of the procedure.

The human leukocyte antigen (HLA) locus is associated with more complex diseases than any other locus in the human genome. In many diseases, HLA explains more heritability than all other known loci combined. In silico HLA imputation methods enable rapid and accurate estimation of HLA alleles in the millions of individuals that are already genotyped on microarrays. HLA imputation has been used to define causal variation in autoimmune diseases, such as type I diabetes, and in human immunodeficiency virus infection control. However, there are few guidelines on performing HLA imputation, association testing, and fine mapping. Here, we present a comprehensive tutorial to impute HLA alleles from genotype data. We provide detailed guidance on performing standard quality control measures for input genotyping data and describe options to impute HLA alleles and amino acids either locally or using the web-based Michigan Imputation Server, which hosts a multi-ancestry HLA imputation reference panel. We also offer best practice recommendations to conduct association tests to define the alleles, amino acids, and haplotypes that affect human traits. Along with the pipeline, we provide a step-by-step online guide with scripts and available software ( https://github.com/immunogenomics/HLA_analyses_tutorial ). This tutorial will be broadly applicable to large-scale genotyping data and will contribute to defining the role of HLA in human diseases across global populations.

Organoids have been an exciting advancement in stem cell research. Here we describe a strategy for directed differentiation of human pluripotent stem cells into distal lung organoids. This protocol recapitulates lung development by sequentially specifying human pluripotent stem cells to definitive endoderm, anterior foregut endoderm, ventral anterior foregut endoderm, lung bud organoids and finally lung organoids. The organoids take ~40 d to generate and can be maintained more than 180 d, while progressively maturing up to a stage consistent with the second trimester of human gestation. They are unique because of their branching morphology, the near absence of non-lung endodermal lineages, presence of mesenchyme and capacity to recapitulate interstitial lung diseases. This protocol can be performed by anyone familiar with cell culture techniques, is conducted in serum-free conditions and does not require lineage-specific reporters or enrichment steps. We also provide a protocol for the generation of single-cell suspensions for single-cell RNA sequencing.

The human cerebral cortex undergoes dramatic and critical development during early postnatal stages. Benefiting from advances in neuroimaging, many infant brain magnetic resonance imaging (MRI) datasets have been collected from multiple imaging sites with different scanners and imaging protocols for the investigation of normal and abnormal early brain development. However, it is extremely challenging to precisely process and quantify infant brain development with these multisite imaging data because infant brain MRI scans exhibit (a) extremely low and dynamic tissue contrast caused by ongoing myelination and maturation and (b) inter-site data heterogeneity resulting from the use of diverse imaging protocols/scanners. Consequently, existing computational tools and pipelines typically perform poorly on infant MRI data. To address these challenges, we propose a robust, multisite-applicable, infant-tailored computational pipeline that leverages powerful deep learning techniques. The main functionality of the proposed pipeline includes preprocessing, brain skull stripping, tissue segmentation, topology correction, cortical surface reconstruction and measurement. Our pipeline can handle both T1w and T2w structural infant brain MR images well in a wide age range (from birth to 6 years of age) and is effective for different imaging protocols/scanners, despite being trained only on the data from the Baby Connectome Project. Extensive comparisons with existing methods on multisite, multimodal and multi-age datasets demonstrate superior effectiveness, accuracy and robustness of our pipeline. We have maintained a website, iBEAT Cloud, for users to process their images with our pipeline ( http://www.ibeat.cloud ), which has successfully processed over 16,000 infant MRI scans from more than 100 institutions with various imaging protocols/scanners.

A lack of generic and effective genetic manipulation methods for Pseudomonas has restricted fundamental research and utilization of this genus for biotechnology applications. Phage-encoded homologous recombination (PEHR) is an efficient tool for bacterial genome engineering. This PEHR system is based on a lambda Red-like operon (BAS) from Pseudomonas aeruginosa phage Ab31 and a Rac bacteriophage RecET-like operon (Rec-TEPsy) from P. syringae pv. syringae B728a and also contains exogenous elements, including the RecBCD inhibitor (Redγ or Pluγ) or single-stranded DNA-binding protein (SSB), that were added to enhance the PEHR recombineering efficiency. To solve the problem of false positives in Pseudomonas editing with the PEHR system, the processive enzyme Cas3 with a minimal Type I-C Cascade-based system was combined with PEHR. This protocol describes the utilization of a Pseudomonas-specific PEHR-Cas3 system that was designed to universally and proficiently modify the genomes of Pseudomonas species. The pipeline uses standardized cassettes combined with the concerted use of SacB counterselection and Cre site-specific recombinase for markerless or seamless genome modification, in association with vectors that possess the selectively replicating template R6K to minimize recombineering background. Compared with the traditional allelic exchange editing method, the PEHR-Cas3 system does not need to construct suicide plasmids carrying long homologous arms, thus simplifying the experimental procedure and shortening the traceless editing period. Compared with general editing systems based on phage recombinases, the PEHR-Cas3 system can effectively improve the screening efficiency of mutants using the cutting ability of Cas3 protein. The entire procedure requires ~12 days.

Cell-free DNA (cfDNA) in blood, viewed as a surrogate for tumor biopsy, has many clinical applications, including diagnosing cancer, guiding cancer treatment and monitoring treatment response. All these applications depend on an indispensable, yet underdeveloped task: detecting somatic mutations from cfDNA. The task is challenging because of the low tumor fraction in cfDNA. Recently, we developed the computational method cfSNV, the first method that comprehensively considers the properties of cfDNA for the sensitive detection of mutations from cfDNA. cfSNV vastly outperformed the conventional methods that were developed primarily for calling mutations from solid tumor tissues. cfSNV can accurately detect mutations in cfDNA even with medium-coverage (e.g., ≥200×) sequencing, which makes whole-exome sequencing (WES) of cfDNA a viable option for various clinical utilities. Here, we present a user-friendly cfSNV package that exhibits fast computation and convenient user options. We also built a Docker image of it, which is designed to enable researchers and clinicians with a limited computational background to easily carry out analyses on both high-performance computing platforms and local computers. Mutation calling from a standard preprocessed WES dataset (~250× and ~70 million base pair target size) can be carried out in 3 h on a server with eight virtual CPUs and 32 GB of random access memory.

Developing models of human kidney tissue in vitro is an important challenge in regenerative nephrology research, given the paucity of novel and effective therapies in kidney disease. However, the de novo generation of kidney tissues from human pluripotent stem cells (hPSCs) is challenging owing to the structural and functional complexity of the organ, as well its developmental origin from two distinct embryologic populations: the metanephric mesenchyme and the ureteric bud (UB). Directed differentiation strategies have been developed to generate kidney organoids containing nephron-like structures; we recently reported an efficient and practical method to generate UB tissues. Here, we describe a detailed step-by-step protocol for differentiation of hPSCs into three-dimensonal UB organoids that exhibit complex morphological development and the capacity to differentiate into functional collecting duct tissues. Over 3 d, hPSCs are induced into PAX2+GATA3+ pronephric (anterior) intermediate mesoderm fates in monolayer cultures at high efficiency. The cells are aggregated into three-dimensional spheroids, which then assemble and organize into nephric duct-like tissue over 4 d. When embedded into an extracellular matrix, the spheroids grow into UB organoids that recapitulate fetal branching morphogenesis for 1 week of culture. When switched to permissive conditions, the UB organoids spontaneously differentiate to form collecting duct principal cells. This approach provides robust and reproducible methods that can be readily adopted by users with basic experience in hPSC and organoid differentiation to generate UB tissues, which may be used to investigate human kidney development, model disease processes and catalyze further efforts in engineering functional kidney tissue.

Imaging across different scales is essential for understanding healthy organ morphology and pathophysiological changes. The macro- and microscale three-dimensional morphology of large samples, including intact human organs, is possible with X-ray microtomography (using laboratory or synchrotron sources). Preparation of large samples for high-resolution imaging, however, is challenging due to limitations such as sample shrinkage, insufficient contrast, movement of the sample and bubble formation during mounting or scanning. Here, we describe the preparation, stabilization, dehydration and mounting of large soft-tissue samples for X-ray microtomography. We detail the protocol applied to whole human organs and hierarchical phase-contrast tomography at the European Synchrotron Radiation Facility, yet it is applicable to a range of biological samples, including complete organisms. The protocol enhances the contrast when using X-ray imaging, while preventing sample motion during the scan, even with different sample orientations. Bubbles trapped during mounting and those formed during scanning (in the case of synchrotron X-ray imaging) are mitigated by multiple degassing steps. The sample preparation is also compatible with magnetic resonance imaging, computed tomography and histological observation. The sample preparation and mounting require 24-36 d for a large organ such as a whole human brain or heart. The preparation time varies depending on the composition, size and fragility of the tissue. Use of the protocol enables scanning of intact organs with a diameter of 150 mm with a local voxel size of 1 μm. The protocol requires users with expertise in handling human or animal organs, laboratory operation and X-ray imaging.

Medulloblastoma and high-grade glioma represent the most aggressive and frequent lethal solid tumors affecting individuals during pediatric age. During the past years, several models have been established for studying these types of cancers. Human organoids have recently been shown to be a valid alternative model to study several aspects of brain cancer biology, genetics and test therapies. Notably, brain cancer organoids can be generated using genetically modified cerebral organoids differentiated from human induced pluripotent stem cells (hiPSCs). However, the protocols to generate them and their downstream applications are very rare. Here, we describe the protocols to generate cerebellum and forebrain organoids from hiPSCs, and the workflow to genetically modify them by overexpressing genes found altered in patients to finally produce cancer organoids. We also show detailed protocols to use medulloblastoma and high-grade glioma organoids for orthotopic transplantation and co-culture experiments aimed to study cell biology in vivo and in vitro, for lineage tracing to investigate the cell of origin and for drug screening. The protocol takes 60-65 d for cancer organoids generation and from 1-4 weeks for downstream applications. The protocol requires at least 3-6 months to become proficient in culturing hiPSCs, generating organoids and performing procedures on immunodeficient mice.

The neurovascular unit (NVU), composed of endothelial cells, pericytes, juxtaposed astrocytes and microglia together with neurons, is essential for proper central nervous system functioning. The NVU critically regulates blood-brain barrier (BBB) function, which is impaired in several neurological diseases and is therefore a key therapeutic target. To understand the extent and cellular source of BBB dysfunction, simultaneous isolation and analysis of NVU cells is needed. Here, we describe a protocol for the EPAM-ia method, which is based on flow cytometry for simultaneous isolation and analysis of endothelial cells, pericytes, astrocytes and microglia. This method is based on differential processing of NVU cell types using enzymes, mechanical homogenization and filtration specific for each cell type followed by combining them for immunostaining and fluorescence-activated cell sorting. The gating strategy encompasses cell-type-specific and exclusion markers for contaminating cells to isolate the major NVU cell types. This protocol takes ~6 h for two sets of one or two animals. The isolation part requires experience in animal handling, fresh tissue processing and immunolabeling for flow cytometry. Sorted NVU cells can be used for downstream applications including transcriptomics, proteomics and cell culture. Multiple cell-type analyses using UpSet can then be applied to obtain robust targets from single or multiple NVU cell types in neurological diseases associated with BBB dysfunction. The EPAM-ia method is also amenable to isolation of several other cell types, including cancer cells and immune cells. This protocol is applicable to healthy and pathological tissue from mouse and human sources and to several cell types compared with similar protocols.

Owing to its high sustainable production capacity, cellulose represents a valuable feedstock for the development of more sustainable alternatives to currently used fossil fuel-based materials. Chemical analysis of cellulose remains challenging, and analytical techniques have not advanced as fast as the development of the proposed materials science applications. Crystalline cellulosic materials are insoluble in most solvents, which restricts direct analytical techniques to lower-resolution solid-state spectroscopy, destructive indirect procedures or to 'old-school' derivatization protocols. While investigating their use for biomass valorization, tetralkylphosphonium ionic liquids (ILs) exhibited advantageous properties for direct solution-state nuclear magnetic resonance (NMR) analysis of crystalline cellulose. After screening and optimization, the IL tetra-n-butylphosphonium acetate [P4444][OAc], diluted with dimethyl sulfoxide-d6, was found to be the most promising partly deuterated solvent system for high-resolution solution-state NMR. The solvent system has been used for the measurement of both 1D and 2D experiments for a wide substrate scope, with excellent spectral quality and signal-to-noise, all with modest collection times. The procedure initially describes the scalable syntheses of an IL, in 24-72 h, of sufficient purity, yielding a stock electrolyte solution. The dissolution of cellulosic materials and preparation of NMR samples is presented, with pretreatment, concentration and dissolution time recommendations for different sample types. Also included is a set of recommended 1D and 2D NMR experiments with parameters optimized for an in-depth structural characterization of cellulosic materials. The time required for full characterization varies between a few hours and several days.

Chemical modifications of transcripts with a 5' cap occur in all organisms and function in many aspects of RNA metabolism. To facilitate analysis of RNA caps, we developed a systems-level mass spectrometry-based technique, CapQuant, for accurate and sensitive quantification of the cap epitranscriptome. The protocol includes the addition of stable isotope-labeled cap nucleotides (CNs) to RNA, enzymatic hydrolysis of endogenous RNA to release CNs, and off-line enrichment of CNs by ion-pairing high-pressure liquid chromatography, followed by a 17 min chromatography-coupled tandem quadrupole mass spectrometry run for the identification and quantification of individual CNs. The total time required for the protocol can be up to 7 d. In this approach, 26 CNs can be quantified in eukaryotic poly(A)-tailed RNA, bacterial total RNA and viral RNA. This protocol can be modified to analyze other types of RNA and RNA from in vitro sources. CapQuant stands out from other methods in terms of superior specificity, sensitivity and accuracy, and it is not limited to individual caps nor does it require radiolabeling. Thanks to its unique capability of accurately and sensitively quantifying RNA caps on a systems level, CapQuant can reveal both the RNA cap landscape and the transcription start site distribution of capped RNA in a broad range of settings.

Programmable cytosine base editors show promising approaches for correcting pathogenic mutations; yet, their off-target effects have been of great concern. Detect-seq (dU-detection enabled by C-to-T transition during sequencing) is an unbiased, sensitive method for the off-target evaluation of programmable cytosine base editors. It profiles the editome by tracing the editing intermediate dU, which is introduced inside living cells and edited by programmable cytosine base editors. The genomic DNA is extracted, preprocessed and labeled by successive chemical and enzymatic reactions, followed by biotin pull-down to enrich the dU-containing loci for sequencing. Here, we describe a detailed protocol for performing the Detect-seq experiment, and a customized, open-source, bioinformatic pipeline for analyzing the characteristic Detect-seq data is also provided. Unlike those previous whole-genome sequencing-based methods, Detect-seq uses an enrichment strategy and hence is endowed with great sensitivity, a higher signal-to-noise ratio and no requirement for high sequencing depth. Furthermore, Detect-seq is widely applicable for both mitotic and postmitotic biological systems. The entire protocol typically takes 5 d from the genomic DNA extraction to sequencing and ~1 week for data analysis.

The fluorination of amino acid residues represents a near-isosteric alteration with the potential to report on biological pathways, yet the site-directed editing of carbon-hydrogen (C-H) bonds in complex biomolecules to carbon-fluorine (C-F) bonds is challenging, resulting in its limited exploitation. Here, we describe a protocol for the posttranslational and site-directed alteration of native γCH2 to γCF2 in protein sidechains. This alteration allows the installation of difluorinated sidechain analogs of proteinogenic amino acids, in both native and modified states. This chemical editing is robust, mild, fast and highly efficient, exploiting photochemical- and radical-mediated C-C bonds grafted onto easy-to-access cysteine-derived dehydroalanine-containing proteins as starting materials. The heteroaryl-sulfonyl reagent required for generating the key carbon-centered C• radicals that install the sidechain can be synthesized in two to six steps from commercially available precursors. This workflow allows the nonexpert to create fluorinated proteins within 24 h, starting from a corresponding purified cysteine-containing protein precursor, without the need for bespoke biological systems. As an example, we readily introduce three γCF2-containing methionines in all three progressive oxidation states (sulfide, sulfoxide and sulfone) as D-/L- forms into histone eH3.1 at site 4 (a relevant lysine to methionine oncomutation site), and each can be detected by 19F-nuclear magnetic resonance of the γCF2 group, as well as the two diastereomers of the sulfoxide, even when found in a complex protein mixture of all three. The site-directed editing of C-H→C-F enables the use of γCF2 as a highly sensitive, 'zero-size-zero-background' label in protein sidechains, which may be used to probe biological phenomena, protein structures and/or protein-ligand interactions by 19F-based detection methods.

Targeted metabolite assays that measure tens or hundreds of pre-selected metabolites, typically using liquid chromatography-mass spectrometry, are increasingly being developed and applied to metabolic phenotyping studies. These are used both as standalone phenotyping methods and for the validation of putative metabolic biomarkers obtained from untargeted metabolomics studies. However, there are no widely accepted standards in the scientific community for ensuring reliability of the development and validation of targeted metabolite assays (referred to here as 'targeted metabolomics'). Most current practices attempt to adopt, with modifications, the strict guidance provided by drug regulatory authorities for analytical methods designed largely for measuring drugs and other xenobiotic analytes. Here, the regulatory guidance provided by the European Medicines Agency, US Food and Drug Administration and International Council for Harmonisation of Technical Requirements for Pharmaceuticals for Human Use are summarized. In this Perspective, we have adapted these guidelines and propose a less onerous 'tiered' approach to evaluate the reliability of a wide range of metabolomics analyses, addressing the need for community-accepted, harmonized guidelines for tiers other than full validation. This 'fit-for-purpose' tiered approach comprises four levels-discovery, screening, qualification and validation-and is discussed in the context of a range of targeted and untargeted metabolomics assays. Issues arising with targeted multiplexed metabolomics assays, and how these might be addressed, are considered. Furthermore, guidance is provided to assist the community with selecting the appropriate degree of reliability for a series of well-defined applications of metabolomics.

The interactions between molecules and noble metal nanosurfaces play a central role in many areas of nanotechnology. The surface chemistry of noble metal surfaces under ideal, clean conditions has been extensively studied; however, clean conditions are seldom met in real-world applications. We developed a sensitive and robust characterization technique for probing the surface chemistry of nanomaterials in the complex environments that are directly relevant to their applications. Surface-enhanced Raman spectroscopy (SERS) can be used to probe the interaction of plasmonic nanoparticles with light to enhance the Raman signals of molecules near the surface of nanoparticles. Here, we explain how to couple SERS with surface-accessible plasmonic-enhancing substrates, which are capped with weakly adsorbing capping ligands such as citrate and chloride ions, to allow molecule-metal interactions to be probed in situ and in real time, thus providing information on the surface orientation and the formation and breaking of chemical bonds. The procedure covers the synthesis and characterization of surface-accessible colloids, the preliminary SERS screening with agglomerated colloids, the synthesis and characterization of interfacial nanoparticle assemblies, termed metal liquid-like films, and the in situ biphasic SERS analysis with metal liquid-like films. The applications of the approach are illustrated using two examples: the probing of π-metal interactions and that of target/ligand-particle interactions on hollow bimetallic nanostars. This protocol, from the initial synthesis of the surface-accessible plasmonic nanoparticles to the final in situ biphasic SERS analysis, requires ~14 h and is ideally suited to users with basic knowledge in performing Raman spectroscopy and wet synthesis of metal nanoparticles.