Current protocols in mouse biology最新文献

The light (or optical) microscope is the icon of science. The aphorism “seeing is believing” is often quoted in scientific papers involving microscopy. Unlike many scientific instruments, the light microscope will deliver an image however badly it is set up. Fluorescence microscopy is a widely used research tool across all disciplines of biological and biomedical science. Most universities and research institutions have microscopes, including confocal microscopes. This introductory paper in a series detailing advanced light microscopy techniques explains the foundations of both electron and light microscopy for biologists and life scientists working with the mouse. An explanation is given of how an image is formed. A description is given of how to set up a light microscope, whether it be a brightfield light microscope on the laboratory bench, a widefield fluorescence microscope, or a confocal microscope. These explanations are accompanied by operational protocols. A full explanation on how to set up and adjust a microscope according to the principles of Köhler illumination is given. The importance of Nyquist sampling is discussed. Guidelines are given on how to choose the best microscope to image the particular sample or slide preparation that you are working with. These are the basic principles of microscopy that a researcher must have an understanding of when operating core bioimaging facility instruments, in order to collect high-quality images. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Setting up Köhler illumination for a brightfield microscope

Basic Protocol 2: Aligning the fluorescence bulb and setting up Köhler illumination for a widefield fluorescence microscope

Basic Protocol 3: Generic protocol for operating a confocal microscope

Experiments that visualize gene expression in intact tissues or organisms are fundamental to studies of gene function. These experiments, called in situ hybridization, require the production of a riboprobe, which is a labeled antisense RNA corresponding to a particular gene. The most commonly used system for visualizing gene expression via in situ hybridization is the incorporation of a digoxigenin label into an in vitro−transcribed RNA probe. After hybridization of the riboprobe to a target mRNA, its location can be detected via a high-affinity α-digoxigenin antibody conjugated to an alkaline-phosphatase enzyme. The article describes the design and production of digoxigenin-labeled riboprobes transcribed in vitro from template DNA (either plasmid or PCR amplicon). These riboprobes are suitable for use in tissue and whole-mount in situ hybridization protocols. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Plasmid-derived riboprobes

Alternate Protocol: PCR-derived riboprobes

Basic Protocol 2: Riboprobe synthesis with DIG label

Understanding RNA expression in space and time is a key initial step in dissecting gene function. The ability to visualize gene expression in whole-tissue or whole-specimen preparations, called in situ hybridization (ISH), was first developed 50 years ago. Two decades later, these protocols were adapted to establish robust methods for whole-mount ISH to murine embryos. The precise protocols vary somewhat between early-gestation and mid-gestation mouse embryos; the protocol presented here is optimal for use with post-implantation stage mouse embryos (stages 5.5–9.5 dpc). Routine uses of whole-mount ISH include documenting the wild-type expression pattern of individual genes and comparison of the expression pattern of signature genes (i.e., those that identify particular cells and tissues within an embryo) between wild-type and mutant embryos as part of a phenotyping experiment. This technique remains a mainstay of developmental biology studies and complements the massively parallel assessment of gene expression from dissociated tissues and cells via RNA-sequencing techniques. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Dissection of post-implantation (5.5-9.5 dpc) murine embryos

Basic Protocol 2: Whole-mount in situ hybridization in post-implantation embryos

Basic Protocol 3: Visualization of post-WMISH embryos

Support Protocol 1: Creation of siliconized glass pipettes

Support Protocol 2: Creation of embryo powder

The simple protocol described in this article aims to provide all required information, as a comprehensive, easy-to-follow step-by-step method, to ensure the generation of the expected genome-edited mice. Here, we provide protocols for the preparation of CRISPR-Cas9 reagents for microinjection and electroporation into one-cell mouse embryos to create knockout or knock-in mouse models, and for genotyping the resulting offspring with the latest innovative next-generation sequencing methods. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Designing the best RNA guide for your gene disruption/editing strategy

Basic Protocol 2: Preparing and validating CRISPR-Cas9 reagents

Basic Protocol 3: Preparing and injecting CRISPR-Cas9 compounds into fertilized mouse oocytes

Basic Protocol 4: Genotyping genome-edited mice

Support Protocol: Genotyping for CRISPR-generated “indel” mutations

Current preclinical cognitive assessments are highly time intensive, with lengthy assessment procedures. In this regard, a single-day assay that focuses just on assessing learning behavior in a time-effective and relatable manner would be of value. This article describes the box maze as a short-term behavioral procedure to measure learning in mice. The protocol consists of allowing mice to explore an enclosed space that has eight holes. One of these holes leads to a tunnel that connects to an escape cage, and the latency to enter this escape hole is recorded for each mouse. Mice are tested four times within a single day, and the decrease in escape latency over time is used as a measure of learning. Age is a factor that affects escape latency in the box maze. Hence, the box-maze procedure is proposed as an efficient test to probe aging and aging intervention–related research questions. © 2020 by John Wiley & Sons, Inc.

Generating large-fragment knock-ins, such as reporters, conditional alleles, or humanized alleles, directly in mouse embryos is still a challenging feat. We have developed 2C-HR-CRISPR, a technology that allows highly efficient (10-50%) and rapid (generating founders in 2 months) targeting of large DNA fragments. Key to this strategy is the delivery of CRISPR reagents into 2-cell-stage mouse embryos, taking advantage of the high homologous recombination activity during the long G₂ cell cycle phase at this stage. Furthermore, by exploiting a Cas9–monomeric streptavidin (Cas-mSA) and biotinylated PCR template (BioPCR) system to localize the repair template to specific double strand breaks, the efficiency can be further improved to up to 95%. Here we provide a procedure to generate large-fragment knock-in mouse models using 2C-HR-CRISPR. We first describe the principles for designing single guide RNAs and repair templates but refer to published manuscripts and protocols for molecular cloning methods or commercial sources for these reagents. We then describe two unique aspects of 2C-HR-CRISPR that are critical for success: (1) production of the CRISPR reagents for 2C-HR-CRISPR, particularly for applying the Cas9-mSA/BioPCR method, and (2) microinjection of mouse embryos at the 2-cell stage. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Single guide RNA and repair template design

Basic Protocol 2: Preparing reagents for 2C-HR-CRISPR

Basic Protocol 3: Microinjecting 2-cell-stage mouse embryos

Cover: In Jacquot et al. (http://doi.org/10.1002/cpmo.65) image shows Laboratory Information Management System (LIMS). Example of a database controlling the flow of samples, the primer bank, and the operation of the workstation with worklists. The database is connected to the animal management database so that it can receive genotyping requests and transfer animal genotypes at the end of the analyses.

The Collaborative Cross (CC) mouse resource is a next-generation mouse genetic reference population (GRP) designed for high-resolution mapping of quantitative trait loci (QTL) of large effect affecting complex traits during health and disease. The CC resource consists of a set of 72 recombinant inbred lines (RILs) generated by reciprocal crossing of five classical and three wild-derived mouse founder strains. Complex traits are controlled by variations within multiple genes and environmental factors, and their mutual interactions. These traits are observed at multiple levels of the animals’ systems, including metabolism, body weight, immune profile, and susceptibility or resistance to the development and progress of infectious or chronic diseases. Herein, we present general guidelines for design of QTL mapping experiments using the CC resource—along with full step-by-step protocols and methods that were implemented in our lab for the phenotypic and genotypic characterization of the different CC lines—for mapping the genes underlying host response to infectious and chronic diseases. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: CC lines for whole body mass index (BMI)

Basic Protocol 2: A detailed assessment of the power to detect effect sizes based on the number of lines used, and the number of replicates per line

Basic Protocol 3: Obtaining power for QTL with given target effect by interpolating in Table 1 of Keele et al. (2019)

Genotyping consists of searching for a DNA sequence variation localized at a well-defined locus in the genome. It is an essential step in animal research because it allows the identification of animals that will be bred to generate and maintain a colony, euthanized to control the available space in the animal facility, or used in experiment protocols. Here we describe polymerase chain reaction (PCR) genotyping protocols for fast, sensitive, easy, and cost-effective characterization of mouse genotype. We discuss optimization of parameters to improve the reliability of each assay and propose recommendations for enhancing reproducibility and reducing the occurrence of inconclusive genotyping. All steps required for efficient genotyping are presented: tissue collection; sample verification and direct DNA lysis; establishment of a robust genotyping strategy with reliable, rapid, and cost-effective assays; and finally, transition to high-throughput automatized PCR, including mix miniaturization and automation. © 2019 The Authors.

Basic Protocol 1: Tissue sampling methods and procedure

Basic Protocol 2: Sample verification and DNA lysis

Basic Protocol 3: Design of a genotyping strategy

Basic Protocol 4: Moving to high-throughput genotyping

Cover: In Williams et al. (http://doi.org/10.1002/cpmo.64) image shows ENU-mutagenized males are mated to wild-type females. Each G1 offspring carries a unique set of mutations and therefore will exhibit different phenotypes. To segregate the phenotypes with a mutation or mutations, the G1 male mice are mated to wild-type females. The G2 offspring are then either intercrossed to each other or mated back to the original G1. The G3 progeny can be phenotyped for both recessive and dominant mutations. Typically, wild-type females of a different genetic background are used in the breeding scheme to facilitate the genetic mapping of any phenodeviant G3 offspring.