Current protocols最新文献

Advances in sequencing technology have led to the discovery of diverse types of regulatory RNAs. Differential transcript levels regulate cellular processes and influence disease severity. Identifying these variations through reliable methods is crucial for understanding the regulatory roles and disease mechanisms of regulatory RNAs. Northern blotting, which is considered the gold standard for differential expression analysis, poses challenges due to various limitations associated with RNA quality and integrity, radioactivity exposure, and associated reagents and expenses. In this protocol, we employ a biotin-based northern blotting (BiNoB) approach that is both convenient and inexpensive, eliminating the need for specialized settings as required with radioactivity-based northern blotting. We comprehensively target various RNA types, making this technique a versatile tool for RNA detection. Additionally, we conduct a comparison between 3′-end labeled probes that were labeled in-house and 5′-end labeled probes that were obtained commercially. Remarkably, our results reveal relatively higher sensitivity with 3′-end labeled probes. Furthermore, we demonstrated that the use of an in-house buffer offered comparable sensitivity to a commercially available buffer, providing another cost-effective alternative. We also aimed to determine the minimum quantity of total RNA required to detect small non-coding RNAs such as tRNA fragments. Whereas previous studies reported the use of 5-10 µg total RNA for tRNA fragment detection, our findings revealed that as little as 1 µg total RNA is sufficient to detect small RNAs like tRNAs and their fragments. This concentration may vary depending on the expression levels of the specific RNAs being detected. © 2024 Wiley Periodicals LLC.

Basic Protocol: Biotin-based northern blotting

Cultivated meat represents a transformative solution to environmental and ethical concerns of traditional meat industries, replicating livestock meat's texture and sensory attributes in vitro with a focus on cost, safety, and nutritional quality. Central to this process are biomaterial scaffolds that support tissue development from isolated animal cells grown in or on these matrices. Understanding scaffold interactions with cells, including scaffold degradation and biomass production, is crucial for process design and for scaling-up goals. In this article, we outline comprehensive methods to quantify scaffold-cell interactions for such scenarios, focusing on biomaterial scaffold degradation and changes in cell biomass [measured by cell weight, extracellular matrix (ECM) deposition, and cell coverage] during cell culture. We introduce two methodologies for assessing cell coverage: fixation and staining for detailed imaging analysis, and non-invasive, real-time evaluation across scaffolds. Here we focus on fiber-based scaffolds, while the assessments can be extrapolated to 2-dimensional (2D; films), and in part to 3-dimensional (3D; sponge) systems. Utilizing the C2C12 mouse myoblast cell line as a gold standard, the protocols deliver precise, step-by-step instructions for preparing fiber scaffolds (using silk proteins here), seeding cells, and monitoring key parameters for cultivated meat production, providing a framework for advancing cellular agriculture techniques. © 2024 Wiley Periodicals LLC.

Basic Protocol 1: Fabrication and preparation of silk fiber scaffolds for cell seeding

Support Protocol 1: Cultivation of C2C12 cells and seeding onto fibrous scaffolds

Basic Protocol 2: Quantification of decellularized yarn scaffold degradation during cell culture

Basic Protocol 3: Quantification of biomass variation and ECM deposition on yarn scaffolds during C2C12 cell culture

Basic Protocol 4: Visualization of cell-laden yarn scaffolds and determination of cell coverage ratio using confocal microscopy

Support Protocol 2: Real-time imaging of cell-laden yarn scaffolds using Celigo system

Support Protocol 3: Applying green CellTracker fluorescent probes to C2C12 cells seeded on yarn scaffolds

Current Protocols is issuing corrections for the following protocol article.

Reuven, O., Mikula, I., Ashkenazi-Preiser, H., Twaik, N., Ben-Meir, K., Meirow, Y., Daniel, L., Kariv, G., Kurd, M., & Baniyash, M. (2022). Functional assays evaluating immunosuppression mediated by myeloid-derived suppressor cells. Current Protocols, 2, e557. doi: 10.1002/cpz1.557

In the above-referenced article:

In step 11 of Support Protocol 2, “0.03125 µg/ml” has been changed to “0.03125 mg/ml”.

The current version online now includes this correction and may be considered the authoritative version of record.

Studying adipogenesis and adipocyte biology requires the isolation of primary preadipocytes from adipose tissues. However, primary preadipocytes have a limited lifespan, can only undergo a finite number of divisions, and often lose their original biological characteristics before becoming senescent. The repeated isolation of fresh preadipocytes, particularly from young pups or aged animals, is costly and time consuming. Immortalization of these cells offers a solution by overcoming cellular senescence and maintaining proliferative capacity, allowing for long-term studies without the continuous need to isolate new cells from animals. Immortalized cell lines thus provide a consistent and reproducible experimental model, significantly reducing variability across different animals. However, successfully establishing immortalized preadipocyte cell lines presents challenges, including selecting appropriate adipose tissue depots, isolating primary preadipocytes, and choosing an effective immortalization strategy. In this article, we present optimized protocols and share first-hand experiences establishing immortalized brown and white preadipocyte cell lines from young and aging mice. These protocols offer a valuable resource for researchers studying adipogenesis and metabolism. © 2024 Wiley Periodicals LLC.

Support Protocol 1: Retrovirus production

Basic Protocol 1: Isolation and culture of primary brown and white preadipocytes from mouse interscapular brown adipose tissue (iBAT) and subcutaneous white adipose tissue (sWAT) in the same region

Basic Protocol 2: Immortalization of mouse brown and white preadipocytes

Basic Protocol 3: Selection of immortalized preadipocytes

Basic Protocol 4: Selection of single-cell clones of immortalized mouse preadipocytes

Basic Protocol 5: Single-cell sorting in a 96-well plate using a flow cytometer for the selection of single-cell clones of immortalized preadipocytes

Support Protocol 2: Cryopreservation of immortalized mouse preadipocytes

Support Protocol 3: Thawing and culture of cryopreserved immortalized mouse preadipocytes

Support Protocol 4: Subculture and expansion of immortalized mouse preadipocytes

Basic Protocol 6: Differentiation of immortalized mouse brown and white preadipocytes

Support Protocol 5: Identification of differentiated white and brown adipocytes

Metastasis remains a leading cause of cancer-related mortality, yet its study has been constrained by the lack of reliable animal models that faithfully replicate this complex process. Syngeneic models for studying lung cancer metastasis are limited, with the Lewis lung carcinoma (LLC) model being the most commonly employed. The conventional LLC orthotopic model involves injecting LLC cells intravenously (i.v.) via the tail vein into syngeneic C57BL/6 mice. However, this model has significant drawbacks, such as tumor development in multiple anatomical sites, incomplete lung tumor penetrance, and challenges in monitoring lung tumor growth. This article highlights the advantages of using luciferase-expressing LLC cells combined with bioluminescence imaging (BLI) to quantify tumor progression in live animals. We demonstrate that both white- and black-furred C57BL/6 mice can be used for BLI. Finally, we propose that intranasal (i.n.) instillation of LLC cells offers a valuable alternative to the traditional i.v. tail vein injection method, particularly for its simplicity and improved reproducibility. Although the LLC i.n. model does not recapitulate the metastasis process via the blood vascular route, it is an effective model for studying tumor seeding within the lungs and is particularly useful for analyzing the impact of the lung microenvironment on tumor initiation and progression. © 2024 Wiley Periodicals LLC.

Basic Protocol 1: Lewis lung carcinoma intravenous injection method

Support Protocol: In vivo bioluminescence imaging

Basic Protocol 2: Lewis lung carcinoma intranasal instillation method

Gene therapies are being developed for several central nervous system (CNS) disorders. These therapies are primarily administered to the CNS via the cerebrospinal fluid (CSF), as the blood–brain barrier prevents the transport of large molecules to the brain. Currently, intrathecal injection is the most commonly used route of administration over cisterna magna injections in the clinic for gaining access to the CSF. However, studies in nonhuman primates (NHPs) have shown that administering gene therapies via suboccipital cisterna magna injection results in superior distribution and more cells being transduced in the brain compared to lumbar injection. It has also been reported that comparable CNS size is important when translating therapeutic dosages from animal studies to human trials. Therefore, we chose to develop a computed tomography (CT)-guided cisterna magna injection protocol in pigs as they are anatomically closer in size to humans than nonhuman primates and rodents. Pigs are also a readily available and cost-effective large animal model for preclinical studies compared to nonhuman NHPs. In this paper, we describe a method for CT-guided suboccipital cisterna magna injections in pigs. We developed this protocol utilizing CT to confirm needle placement with three-dimensional visualization. A CT-guided injection minimizes procedural risk and can be performed without a contrast agent, which is required in magnetic resonance and fluoroscopy imaging. © 2024 Wiley Periodicals LLC.

Basic Protocol: Computed tomography–guided suboccipital cisterna magna injection in pigs to confirm needle placement prior to the administration of a test article or vehicle

Ever-increasing availability of experimental volumetric data (e.g., in .ccp4, .mrc, .map, .rec, .zarr, .ome.tif formats) and advances in segmentation software (e.g., Amira, Segger, IMOD) and formats (e.g., .am, .seg, .mod, etc.) have led to a demand for efficient web-based visualization tools. Despite this, current solutions remain scarce, hindering data interpretation and dissemination. Previously, we introduced Mol* Volumes & Segmentations (Mol* VS), a web application for the visualization of volumetric, segmentation, and annotation data (e.g., semantically relevant information on biological entities corresponding to individual segmentations such as Gene Ontology terms or PDB IDs). However, this lacked important features such as the ability to edit annotations (e.g., assigning user-defined descriptions of a segment) and seamlessly share visualizations. Additionally, setting up Mol* VS required a substantial programming background. This article presents an updated version, Mol* VS 2.0, that addresses these limitations. As part of Mol* VS 2.0, we introduce the Annotation Editor, a user-friendly graphical interface for editing annotations, and the Volumes & Segmentations Toolkit (VSToolkit) for generating shareable files with visualization data. The outlined protocols illustrate the utilization of Mol* VS 2.0 for visualization of volumetric and segmentation data across various scales, showcasing the progress in the field of molecular complex visualization. © 2024 The Author(s). Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1: VSToolkit—setting up and visualizing a user-constructed Mol* VS 2.0 database entry

Basic Protocol 2: VSToolkit—visualizing multiple time frames and volume channels

Support Protocol 1: Example: Adding database entry idr-13457537

Alternate Protocol 1: Local-server-and-viewer—visualizing multiple time frames and volume channels

Support Protocol 2: Addition of database entry custom-tubhiswt

Basic Protocol 3: VSToolkit—visualizing a specific channel and time frame

Basic Protocol 4: VSToolkit—visualizing geometric segmentation

Basic Protocol 5: VSToolkit—visualizing lattice segmentations

Alternate Protocol 2: “Local-server-and-viewer”—visualizing lattice segmentations

Basic Protocol 6: “Local-server-and-viewer”—visualizing multiple volume channels

Support Protocol 3: Deploying a server API

Support Protocol 4: Hosting Mol* viewer with VS extension 2.0

Support Protocol 5: Example: Addition of database entry empiar-11756

Support Protocol 6: Example: Addition of database entry emd-1273

Support Protocol 7: Editing annotations

Support Protocol 8: Addition of database entry idr-5025553

Most pathological conditions of the central nervous system do not affect all cell types to the same extent. Delineation of molecular events underlying disease symptoms, including genetic, epigenetic, and transcriptional changes, thus relies on the ability to characterize a specific cell type separately from others. We have developed a methodology for the collection of nuclear RNA and genomic DNA of specific cell types from frozen post-mortem striatum and cerebral cortex. This allows deep transcriptomic profiling of specific cell populations and characterization of their genomes and epigenetic properties. The method is based on the purification of cell nuclei, followed by fluorescence-activated sorting of nuclei (FANS) labeled with nucleic acid probes or antibodies binding to targets present in specific cell types. The protocol describes in detail the procedure for isolating and labeling neuronal and glial nuclei from human brain tissue, the steps that can be taken to extract RNA and genomic DNA, a way to combine the procedure with ATAC-seq to yield information about chromatin accessibility, as well as computational measures for assessing the quality of cell type-specific RNA-seq and ATAC-seq datasets. © 2024 The Author(s). Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1: Tissue homogenization, isolation of cell nuclei by ultracentrifugation and formaldehyde-fixation

Basic Protocol 2: Antibody-based labeling and isolation of nuclei of specific neocortical neuron types

Support Protocol 1: Generation of ATAC-seq libraries from the nuclei of specific neuron types of the cerebral cortex

Basic Protocol 3: Nucleic acid hybridization-based labeling and isolation of nuclei of specific striatal projection neuron types

Alternate Protocol 1: Labeling and isolation of nuclei of specific striatal interneuron types

Support Protocol 2: Generation of ATAC-seq libraries from the nuclei of specific striatal neuron types

Basic Protocol 4: Extraction of genomic DNA and nuclear RNA and preparation of sequencing libraries

Basic Protocol 5: Processing and quality control of FANS-seq and ATAC-seq data

Dendritic spine morphology is associated with the current state of the synapse and neuron, and changes during synaptic plasticity in response to stimulus. At the same time, dendritic spine alterations are reported during various neurodegenerative and neurodevelopmental disorders and other brain states. Accurate and informative analysis of spine shape has an urgent need for studying the synaptic processes and molecular pathways in normal and pathological conditions, and for testing synapto-protective strategies during preclinical studies. Primary neuronal cultures enable high quality imaging of dendritic spines and offer a wide spectrum of accessible experimental manipulations. This article outlines the protocol for isolating, culturing, fluorescent labeling, and imaging of mouse primary hippocampal neurons by three-dimensional (3D) confocal microscopy in a normal state and in conditions of low amyloid toxicity—an in vitro model of Alzheimer's disease. An alternate protocol describes the neuronal morphology analysis using the EGFP expressing neurons in line-M transgenic mouse brain slices. Since the dendritic spines are relatively small structures lying close to the confocal microscope resolution limit, their proper segmentation on the images is challenging. This protocol highlights the image-preprocessing steps, including generation of theoretical point spread function and deconvolution, which enhances resolution and removes noise, thereby enhancing the 3D spine reconstruction results. SpineTool, an open source Python–based script, enables 3D segmentation of dendrites and spines and numerical metric calculation, including key measures, such as spine length, volume, and surface area, with a new feature, the chord length distribution histogram, improving clustering results. SpineTool supports both manual and machine learning spine classification (i.e., mushroom, thin, stubby, filopodia) and automated clustering using k-means and DBSCAN methods. This protocol provides detailed instructions for using SpineTool to analyze and classify dendritic spines in control and experimental groups, enhancing our understanding of spine morphology across different experimental conditions. © 2024 Wiley Periodicals LLC.

Basic Protocol 1: Obtaining 3D confocal dendritic spine images of hippocampal neuronal culture in normal state and conditions of low amyloid toxicity

Alternate Protocol: Obtaining confocal dendritic spine images of mice hippocampal neurons from fixed brain slices

Support Protocol: Post-processing deconvolution of confocal images

Basic Protocol 2: Segmentation of dendritic spines with SpineTool

Basic Protocol 3: Spine dataset preparation using SpineTool

Basic Protocol 4: Clustering of dendritic spines with SpineTool

Basic Protocol 5: Machine classification of dendritic spines with SpineTool