Nature Protocols最新文献

Advances in genomics have identified thousands of risk genes impacting human health and diseases, but the functions of these genes and their mechanistic contribution to disease are often unclear. Moving beyond identification to actionable biological pathways requires dissecting risk gene function and cell type-specific action in intact tissues. This gap can in part be addressed by in vivo Perturb-seq, a method that combines state-of-the-art gene editing tools for programmable perturbation of genes with high-content, high-resolution single-cell genomic assays as phenotypic readouts. Here we describe a detailed protocol to perform massively parallel in vivo Perturb-seq using several versatile adeno-associated virus (AAV) vectors and provide guidance for conducting successful downstream analyses. Expertise in mouse work, AAV production and single-cell genomics is required. We discuss key parameters for designing in vivo Perturb-seq experiments across diverse biological questions and contexts. We further detail the step-by-step procedure, from designing a perturbation library to producing and administering AAV, highlighting where quality control checks can offer critical go-no-go points for this time- and cost-expensive method. Finally, we discuss data analysis options and available software. In vivo Perturb-seq has the potential to greatly accelerate functional genomics studies in mammalian systems, and this protocol will help others adopt it to answer a broad array of biological questions. From guide RNA design to tissue collection and data collection, this protocol is expected to take 9-15 weeks to complete, followed by data analysis.

Bioelectronic medicine, which involves the delivery of electrical stimulation via implantable electrodes, is poised to advance the treatment of neurological conditions. However, current hand-made devices are bulky, invasive and lack specificity. Thin-film neurotechnology devices can overcome these disadvantages. With a typical thickness in the range of micrometers, thin-film devices demonstrate high conformability, stretchability, are minimally invasive and can be fabricated using traditional lithography techniques. Despite their potential, variability and unreliability in fabrication processes hinder their wider utilization. Here, we detail a fabrication method for thin-film poly(ethylene dioxythiophene):poly(styrene sulfonate) (PEDOT:PSS) electrodes and organic electrochemical transistors. The use of organic materials makes these devices particularly well suited for bioelectronic medicine applications as they show superior mechanical and electrical matching of biological tissues compared with devices made of inorganic materials. The procedure details the entire process, including mask design, the fabrication through three photolithography stages, the integration with larger-scale electronics, implantation procedures and the expected electrical characterization metrics. The nanofabrication protocol requires at least 3 d and is suitable for those familiar with lithographic fabrication procedures. The surgery requires up to 10 h and is suitable for those familiar with in vivo implantation procedures.

Sparse, single-cell labeling approaches enable high-resolution, high signal-to-noise recordings from subcellular compartments and intracellular organelles and allow precise manipulations of individual cells and local circuits while minimizing complex changes associated with global network manipulations. However, thus far, only a limited number of approaches have been developed to label single cells with unique combinations of genetically encoded indicators, target deep cortical structures or sustainably use the same chronic preparation for weeks. Here we developed a method to deliver plasmids selectively to single pyramidal neurons in the mouse dorsal hippocampus using two-photon visually guided in vivo single-cell electroporation to address these limitations. This method allows long-term plasmid expression in a controlled number of individual pyramidal neurons, facilitating subcellular resolution imaging, intracellular organelle tracking, monosynaptic input mapping, plasticity induction and targeted whole-cell patch-clamp recordings.

The rise of cellular immunotherapy for cancer treatment has led to the utilization of immune oncology cocultures to simulate T cell interactions with cancer cells for assessing their antitumor response. Previously, we developed BEHAV3D, a three-dimensional live imaging platform of patient-derived tumor organoid (PDO) and engineered T cell cocultures, that analyzes T cells' dynamics to gain crucial insights into their behavior during tumor targeting. However, live imaging alone cannot determine the molecular drivers behind these behaviors. Conversely, single-cell RNA sequencing (scRNA-seq) allows researchers to analyze the transcriptional profiles of individual cells but lacks spatio-temporal resolution. Here we present an extension to the BEHAV3D protocol, called Behavior-Guided Transcriptomics (BGT), for integration of T cell live imaging data with single-cell transcriptomics, enabling analysis of gene programs linked to dynamic T cell behaviors. BGT uses live imaging data processed by BEHAV3D to guide the experimental setup for cell separation based on their PDO engagement levels subsequently followed by fluorescence-activated cell sorting and scRNA-seq. It then integrates in silico simulations of these experiments to computationally infer T cell behavior on scRNA-seq data, uncovering new biomarkers for both highly functional and ineffective T cells, that could be exploited to enhance therapeutic efficacy. The protocol, designed for users with fundamental cell culture, imaging and programming skills, is readily adaptable to diverse coculture settings and takes one month to perform.

Synaptic connections among neurons are critical for information processing and memory storage in the brain, making them hotspots for neuropathologies. Understanding the physiology of synapses, therefore, may facilitate the development of therapeutic approaches. However, synapses are micrometer-sized functional structures involved in many neuronal processes, where the challenge is deciphering differential signaling in presynaptic and postsynaptic compartments of relatively intact microcircuits. Here we developed a method combining two-photon laser microsurgery with compartment-specific electrophysiological activation and readout to improve the specificity with which neuronal signaling is detected. After finding a connection, femtosecond laser pulses are used to sever the presynaptic axon from the cell body with micrometer precision. This microdissection method is effective to a depth of at least 100 µm. The initial segment of the isolated axon is extracellularly stimulated and activated to release neurotransmitters, as detected via a recipient whole-cell neuron, which is being recorded. This methodology is an alternative to axonal patch-clamp recordings, which are short-lasting and difficult. Together with pharmacology and genetic manipulation, our approach allows the interrogation of compartmentalized signaling in intact synapses. The total time of laser exposure is a few seconds and the microsurgery takes 5-10 min, which enables the interrogation of multiple synapses within an experiment. Our protocol provides a tool to investigate compartment-specific signaling in relatively intact brain tissue, enabling a more comprehensive understanding of neuronal synapses.

The developing human brain displays unique features that are difficult to study in animal models. Current in vitro models based on human brain tissue face several challenges, including the limited cellular heterogeneity in two- or three-dimensional neural stem cell cultures, while tissue slice cultures suffer from short survival. We recently established culture conditions to derive organoid cultures directly from human fetal brain tissue by preserving tissue integrity, which can be long-term expanded and display cellular heterogeneity and complex organization. In this Protocol, we describe detailed procedures to establish human fetal brain organoids (FeBOs) that broadly retain regional characteristics, along with procedures for their passaging and characterization. In addition, we describe genome engineering approaches applied to FeBOs to generate mutant FeBO lines that serve as versatile bottom-up brain cancer models. Lastly, we exemplify various downstream applications applicable to both healthy and mutant FeBOs. Scientists with experience in tissue culture can expect the establishment of human FeBO cultures to take 2-3 weeks, while genome engineering of FeBOs takes 2-4 months.

Pair correlation microscopy is a unique approach to fluorescence correlation spectroscopy that can track the long-range diffusive route of a population of fluorescent molecules in live cells with respect to intracellular architecture. This method is based on the use of a pair correlation function (pCF) that, through spatiotemporal comparison of fluctuations in fluorescence intensity recorded throughout a microscope data acquisition, enables changes in a molecule's arrival time to be spatially mapped and statistically quantified. In this protocol, we present guidelines for the measurement and analysis of line scan pair correlation microscopy data acquired on a confocal laser scanning microscope (CLSM), which will enable users to extract a fluorescent molecule's transport pattern throughout a living cell, and then quantify the molecular accessibility of intracellular barriers encountered or the mode of diffusion governing a molecular trafficking event. Finally, we demonstrate how this protocol can be extended to a two-channel line scan acquisition that, when coupled with a cross pCF calculation, enables a fluorescent molecule's transport pattern to be selectively tracked as a function of complex formation with a spectrally distinct fluorescent ligand. For a skilled user of a CLSM, the line scan data acquisition and analysis described in this protocol will take ~1-2 d, depending on the sample and the number of experiments to be processed.

Causal control of short- and long-range projections between networks is necessary to study complex cognitive processes and cortical computations. Neural circuits can be studied via optogenetic approaches, which provide excellent genetic and temporal control and electrophysiological recordings. However, in nonhuman primates (NHPs), these approaches are commonly performed at a single location, missing out on the potential to test connections between separate networks. We have recently developed an approach for optogenetic manipulation in NHPs which targets intra- and interareal cortical projections. Here we describe the combination of optogenetic stimulation with standard chamber-based electrophysiological recordings in awake NHPs to monitor and manipulate both short- and long-range feedforward and feedback circuits. We describe the injection of viral constructs, the simultaneous electrophysiological recordings with the optical stimulation of neurons at various cortical distances and the evaluation of gene expression using a focal biopsy technique. We focus on details that are specific to NHP preparations, such as the precise targeting of injection sites, choosing appropriate viral constructs and considerations for behavioral measures. When combined with laminar electrode configurations (to functionally identify cortical layers) and complex cognitive behavioral tasks, our approach can be used to investigate an array of systems neuroscience questions, such as the role of feedback circuits in attention and the role of lateral connections in contrast normalization. The procedure requires 2-3 active days and 45 waiting days to transduce selected neural circuits and several weeks to complete experiments. The procedure is appropriate for users with expertise in in vivo, awake electrophysiology with NHPs.

Nanostructured devices have proven useful in a broad range of applications, from diagnosing diseases to discovering and screening new drug molecules. We developed vertical silicon nanopillar (SiNP) arrays for on-chip multiplex capture of selected biomolecules using a light-induced release of the array's selectively captured biomarkers. This platform allows the rapid, reusable and quantitative capture and release of a selection of biomarkers, followed by their downstream analysis. Here we outline a standardized protocol for producing the SiNP-based capture-and-release device, which involves the detailed fabrication steps for single-zone nanopillar arrays, their morphological characterization and the chemical modification procedures applied for the anchoring of selective bioreceptors together with the light-controlled on-demand release of the chemical agent. In addition, we provide a detailed approach for the fabrication of a multizone-SiNP array, allowing the simultaneous capture and release of multiple biomarkers of interest. Finally, we demonstrate the entire process of selective and quantitative capture and release of biomolecules from biosamples by means of a commercial low-volume microplate reader system, using green fluorescent protein as a biomarker example. The entire protocol can be conducted within 45 h and requires knowledge in nanoscience, surface chemistry, device micro- and nanofabrication procedures, microfluidics and protein quantification techniques. These SiNP array devices have already demonstrated applications for highly selective and quantitative analysis of a wide range of biological and chemical species, including proteins, nucleic acids, small molecules and ionic species.