Nature Protocols最新文献

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.

Tissue microenvironments are extremely complex and heterogeneous. It is challenging to study metabolic interaction between the different cell types in a tissue with the techniques that are currently available. Here we describe a multimodal imaging pipeline that allows cell type identification and nanoscale tracing of stable isotope-labeled compounds. This pipeline extends upon the principles of correlative light, electron and ion microscopy, by combining confocal microscopy reporter or probe-based fluorescence, electron microscopy, stable isotope labeling and nanoscale secondary ion mass spectrometry. We apply this method to murine models of hepatocellular and mammary gland carcinomas to study uptake of glucose derived carbon (¹³C) and glutamine derived nitrogen (¹⁵N) by tumor-associated immune cells. In vivo labeling with fluorescent-tagged antibodies (B220, CD3, CD8a, CD68) in tandem with confocal microscopy allows for the identification of specific cell types (B cells, T cells and macrophages) in the tumor microenvironment. Subsequent image correlation with electron microscopy offers the contrast and resolution to image membranes and organelles. Nanoscale secondary ion mass spectrometry tracks the enrichment of stable isotopes within these intracellular compartments. The whole protocol described here would take ~6 weeks to perform from start to finish. Our pipeline caters to a broad spectrum of applications as it can easily be adapted to trace the uptake and utilization of any stable isotope-labeled nutrient, drug or a probe by defined cellular populations in any tissue in situ.

Templates for the acquisition of large datasets such as the Human Connectome Project guide the neuroimaging community to reproducible data acquisition and scientific rigor. By contrast, small animal neuroimaging often relies on laboratory-specific protocols, which limit cross-study comparisons. The establishment of broadly validated protocols may facilitate the acquisition of large datasets, which are essential for uncovering potentially small effects often seen in functional MRI (fMRI) studies. Here, we outline a procedure for the acquisition of fMRI datasets in rats and describe animal handling, MRI sequence parameters, data conversion, preprocessing, quality control and data analysis. The procedure is designed to be generalizable across laboratories, has been validated by using datasets across 20 research centers with different scanners and field strengths ranging from 4.7 to 17.2 T and can be used in studies in which it is useful to compare functional connectivity measures across an extensive range of datasets. The MRI procedure requires 1 h per rat to complete and can be carried out by users with limited expertise in rat handling, MRI and data processing.

Cell-matrix interactions, mediated by cellular force and matrix remodeling, result in dynamic reciprocity that drives numerous biological processes and disease progression. Currently, there is no available method for directly quantifying cell traction force and matrix remodeling in three-dimensional matrices as a function of time. To address this long-standing need, we developed a high-resolution microfabricated device that enables longitudinal measurement of cell force, matrix stiffness and the application of mechanical stimulation (tension or compression) to cells. Here a specimen comprising of cells and matrix self-assembles and self-integrates with the sensor. With primary fibroblasts, cancer cells and neurons we have demonstrated the feasibility of the sensor by measuring single or multiple cell force with a resolution of 1 nN and changes in tissue stiffness due to matrix remodeling by the cells. The sensor can also potentially be translated into a high-throughput system for clinical assays such as patient-specific drug and phenotypic screening. We present the detailed protocol for manufacturing the sensors, preparing experimental setup, developing assays with different tissues and for imaging and analyzing the data. Apart from microfabrication of the molds in a cleanroom (one time operation), this protocol does not require any specialized skillset and can be completed within 4-5 h.

Photoporation with free photothermal nanoparticles (NPs) is a promising technology for gentle delivery of functional biomacromolecules into living cells, offering great flexibility in terms of cell types and payload molecules. However, the translational use of photoporation, such as for transfecting patient-derived cells for cell therapies, is hampered by safety and regulatory concerns as it relies on direct contact between cells and photothermal NPs. A solution is to embed the photothermal NPs in electrospun nanofibers, which form a substrate for cell culture. Here we present a protocol for photothermal electrospun nanofiber (PEN)-mediated photoporation that induces membrane permeabilization by photothermal effects and enables efficient intracellular delivery of payload molecules into various cell types. By incorporating photothermal NPs within biocompatible electrospun nanofibers, direct cellular contact with NPs is avoided, thus largely mitigating safety or regulatory issues. Importantly, PEN photoporation is gentler to cells compared with electroporation, the most commonly used physical transfection method, resulting in higher-quality genetically engineered cells with better therapeutic potential. According to this protocol, it takes 2-3 d to prepare PEN culture wells with the desired cells, 3-4 d to optimize PEN photoporation parameters for intracellular delivery of payload molecules into different cell types in vitro and 4-5 weeks to evaluate the in vivo therapeutic efficacy of PEN-photoporated T cells. The protocol also provides details on how to construct the laser-based setup for performing photoporation experiments.

Implantable multifunctional probes have transformed neuroscience research, offering access to multifaceted brain activity that was previously unattainable. Typically, simultaneous access to both optical and electrical signals requires separate probes, while their integration into a single device can result in the emergence of photogenerated electrical artifacts, affecting the quality of high-frequency neural recordings. Among the nontrivial strategies aimed at the realization of an implantable multifunctional interface, the integration of optical and electrical capabilities on a single, minimally invasive, tapered optical fiber probe has been recently demonstrated using fibertrodes. Fibertrodes require the application of a set of planar microfabrication techniques to a nonplanar system with low and nonconstant curvature radius. Here we develop a process based on multiple conformal depositions, nonplanar two-photon lithography and chemical wet etching steps to obtain metallic patterns on the highly curved surface of the fiber taper. We detail the manufacturing, encapsulation and back end of the fibertrodes. The design of the probe can be adapted for different experimental requirements. Using the optical setup design, it is possible to perform angle selective light coupling with the fibertrodes and their implantation and use in vivo. The fabrication of fibertrodes is estimated to require 5-9 d. Nonetheless, due to the high scalability of a large part of the protocol, the manufacture of multiple fibertrodes simultaneously substantially reduces the required time for each probe. The procedure is suitable for users with expertise in microfabrication of electronics and neural recordings.

Deep and accurate proteome analysis is crucial for understanding cellular processes and disease mechanisms; however, it is challenging to implement in routine settings. In this protocol, we combine a robust chromatographic platform with a high-performance mass spectrometric setup to enable routine yet in-depth proteome coverage for a broad community. This entails tip-based sample preparation and pre-formed gradients (Evosep One) combined with a trapped ion mobility time-of-flight mass spectrometer (timsTOF, Bruker). The timsTOF enables parallel accumulation-serial fragmentation (PASEF), in which ions are accumulated and separated by their ion mobility, maximizing ion usage and simplifying spectra. Combined with data-independent acquisition (DIA), it offers high peak sampling rates and near-complete ion coverage. Here, we explain how to balance quantitative accuracy, specificity, proteome coverage and sensitivity by choosing the best PASEF and DIA method parameters. The protocol describes how to set up the liquid chromatography-mass spectrometry system and enables PASEF method generation and evaluation for varied samples by using the py_diAID tool to optimally position isolation windows in the mass-to-charge and ion mobility space. Biological projects (e.g., triplicate proteome analysis in two conditions) can be performed in 3 d with ~3 h of hands-on time and minimal marginal cost. This results in reproducible quantification of 7,000 proteins in a human cancer cell line in quadruplicate 21-min injections and 29,000 phosphosites for phospho-enriched quadruplicates. Synchro-PASEF, a highly efficient, specific and novel scan mode, can be analyzed by Spectronaut or AlphaDIA, resulting in superior quantitative reproducibility because of its high sampling efficiency.

The clinical potential of current chimeric antigen receptor-engineered T (CAR-T) cell therapy is hampered by its autologous nature that poses considerable challenges in manufacturing, costs and patient selection. This spurs demand for off-the-shelf therapies. Here we introduce an ex vivo feeder-free culture method to differentiate gene-engineered hematopoietic stem and progenitor (HSP) cells into allogeneic invariant natural killer T (^AlloNKT) cells and their CAR-armed derivatives (^AlloCAR-NKT cells). We include detailed information on lentivirus generation and titration, as well as the five stages of ex vivo culture required to generate ^AlloCAR-NKT cells, including HSP cell engineering, HSP cell expansion, NKT cell differentiation, NKT cell deep differentiation and NKT cell expansion. In addition, we describe procedures for evaluating the pharmacology, antitumor efficacy and mechanism of action of ^AlloCAR-NKT cells. It takes ~2 weeks to generate and titrate lentiviruses and ~6 weeks to generate mature ^AlloCAR-NKT cells. Competence with human stem cell and T cell culture, gene engineering and flow cytometry is required for optimal results.