Renal ischemia-reperfusion injury (IRI), one of the most important causes of acute kidney injury, is a significant health problem that can result in acute kidney failure after surgical procedures in which blood flow to the kidney is temporarily interrupted. Although numerous studies have been conducted to prevent renal IRI damage, this problem remains unresolved due to its complex nature. During IRI, both ischemia and reperfusion contribute to damage, with oxidative stress, inflammation, and the activation of apoptotic mechanisms playing significant roles in exacerbating the damage. This article provides comprehensive protocols for inducing renal IRI in rats, including both bilateral and unilateral models. These protocols can be used in studies designed to understand the cellular and molecular mechanisms of renal IRI and exploring potential therapeutic strategies. When the described procedure is carefully applied in experimental IRI models, the mortality rate in animals will be lower. Furthermore, we outline opportunities for investigating oxidative stress, inflammatory and apoptotic pathways, and highlight how these analyses may contribute to preclinical therapeutic studies. We also provide recommendations on which analyses should be performed in such studies. Specifically, information is provided on which kidney damage biomarkers should be examined in samples obtained from animals. Additionally, we explain which investigations can be performed on apoptotic and inflammatory pathways involved in kidney damage. In summary, this study integrates bilateral and unilateral approaches and provides a detailed methodological framework for mechanistic and therapeutic research on renal ischemia-reperfusion injury. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Bilateral kidney ischemia-reperfusion in rats Alternate Protocol: Unilateral kidney ischemia-reperfusion in rats Basic Protocol 2: Analyses performed on kidney tissues and serum of rats after ischemia-reperfusion.
Apathy and other disorders of motivation represent a significant clinical problem but do not have an agreed treatment approach. The use of translational animal models could facilitate drug development and advance treatment approach. The effort-based forage task provides a readout of motivational state in mouse models based on their intrinsic drive to forage for nesting material. In this task the mouse is placed in an arena composed of an enclosed home area and a foraging area joined via a tube. Throughout the session the mouse can freely choose to traverse the tube to reach the forage area, obtain nesting material, and shuttle it back to the home area. The nesting material requires effort to obtain and is pulled through apertures in a custom designed nesting material box. The amount of nesting material foraged provides a readout of motivational state, where a deficit in foraging indicates a reduction in motivation. The task does not require physiological (food/water) restriction to motivate the animal to perform in the task, and it does not require training beyond initial habituation to the task environment. The task has been used in behavioral phenotyping of disease models and has been used to test the effects of a wide range of pharmacological manipulations on motivational state. The task environment can be altered to test additional behavioral components that contribute to motivational deficit including effort-based modulation of behavior and affective reactivity. Overall, the task provides a rapid, translationally relevant method for understanding changes in motivated behavior independent of physiological restriction at the preclinical level. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Support Protocol: Animal husbandry and arena set up Basic Protocol: Habituation and acute pharmacological manipulation Alternate Protocol 1: The effort curve paradigm Alternate Protocol 2: Affective reactivity test.
Mouse models are essential for understanding gene function, environmental interaction, and brain structure and function. This is reinforced by the ability of mice to perform complex behavioral tasks. Still, their cognitive assessments often rely on aversive paradigms, such as fear conditioning and the Morris water maze. A promising alternative is the automated touchscreen platform, which enables cognitive tests comparable to those used in humans, such as the Cambridge Neuropsychological Test Automated Battery (CANTAB). This approach enhances standardization and reduces stress by employing appetitive reinforcement. Although widely used in non-human primates, touchscreen testing remains underutilized in rodents despite its potential for cross-species cognitive research. Motivation is key to successful touchscreen tasks, often achieved through water restriction, which mice tolerate well. However, water restriction is a stressful condition, combining negative and positive reinforcement. Here, we propose an alternative that uses citric acid (CA) water to avoid classical food privation in the touchscreen paradigm to mitigate mice's stress. By creating a strong contrast with the reward, we increase the reward's positive valence. We used the touchscreen visual discrimination task to assess the effectiveness of CA water in enhancing motivation. Our results show that administering CA water on training days while allowing access to plain water on weekends reduces the learning phase duration without causing significant weight loss in wild-type C57BL/6J mice. In addition, we observed a strong commitment to performing the pattern dissociation task. This approach offers a welfare-friendly alternative for maintaining motivation in touchscreen-based cognitive tasks while minimizing stress. © 2025 Wiley Periodicals LLC. Basic Protocol: Pattern dissociation paradigm using sour water.
Mice carrying patient-associated base changes are powerful tools to define the causality of single-nucleotide variants to disease states. Epitope tags enable immuno-based studies of genes for which no antibodies are available. These alleles enable detailed and precise developmental, mechanistic, and translational research. The first step in generating these alleles is to identify within the target sequence-the orthologous sequence for base changes or the N or C terminus for epitope tags-appropriate Cas9 protospacer sequences. Subsequent steps include design and acquisition of a single-stranded oligonucleotide repair template, synthesis of a single guide RNA (sgRNA), collection of zygotes, and microinjection or electroporation of zygotes with Cas9 mRNA or protein, sgRNA, and repair template followed by screening born mice for the presence of the desired sequence change. Quality control of mouse lines includes screening for random or multicopy insertions of the repair template and, depending on sgRNA sequence, off-target sequence variation introduced by Cas9. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Single guide RNA design and synthesis Alternate Protocol 1: Single guide RNA synthesis by primer extension and in vitro transcription Basic Protocol 2: Design of oligonucleotide repair template Basic Protocol 3: Preparation of RNA mixture for microinjection Support Protocol 1: Preparation of microinjection buffer Alternate Protocol 2: Preparation of RNP complexes for electroporation Basic Protocol 4: Collection and preparation of mouse zygotes for microinjection or electroporation Basic Protocol 5: Electroporation of Cas9 RNP into zygotes using cuvettes Alternate Protocol 3: Electroporation of Cas9 RNP into zygotes using electrode slides Basic Protocol 6: Screening and quality control of derived mice Support Protocol 2: Deconvoluting multiple sequence chromatograms with DECODR.
The mouse is the mammalian model of choice for investigating cardiovascular biology, given our ability to manipulate it by genetic, pharmacologic, mechanical, and environmental means. Imaging is an important approach to phenotyping both function and structure of cardiac and vascular components. This review details commonly used imaging approaches, with a focus on echocardiography and magnetic resonance imaging, with brief overviews of other imaging modalities. In this update, we also emphasize the importance of rigor and reproducibility in imaging approaches, experimental design, and documentation. Finally, we briefly outline emerging imaging approaches but caution that reliability and validity data may be lacking. © 2024 Wiley Periodicals LLC.
Recent advances in cell culture models like air‒liquid interface culture and ex vivo models such as organoids have advanced studies of lung biology; however, gaps exist between these models and tools that represent the complexity of the three-dimensional environment of the lung. Precision-cut lung slices (PCLS) mimic the in vivo environment and bridge the gap between in vitro and in vivo models. We have established the acid injury and repair (AIR) model where a spatially restricted area of tissue is injured using drops of HCl combined with Pluronic gel. Injury and repair are assessed by immunofluorescence using robust markers, including Ki67 for cell proliferation and prosurfactant protein C for alveolar type 2/progenitor cells. Importantly, the AIR model enables the study of injury and repair in mouse lung tissue without the need for an initial in vivo injury, and the results are highly reproducible. Here, we present detailed protocols for the generation of PCLS and the AIR model. We also describe methods to analyze and quantify injury in AIR-PCLS by immunostaining with established early repair markers and fluorescence imaging. This novel ex vivo model is a versatile tool for studying lung cell biology in acute lung injury and for semi-high-throughput screening of potential therapeutics. © 2020 Wiley Periodicals LLC. Basic Protocol 1 : Generation of precision-cut lung slices Basic Protocol 2 : The acid injury and repair model Basic Protocol 3 : Analysis of AIR-PCLS: Immunostaining and imaging
Despite the importance of emotional intelligence, its biological mechanism is still not well understood. For this reason, we have developed a rodent detour task which requires an animal to reach a highly desired object placed directly behind a transparent barrier that blocks the direct route to the target. This apparently simple task is highly dependent on the emotional control that is necessary to inhibit prepotent and counterproductive responses driven by the sight of a desired object. The water escape detour task designed for mice enables testing the ability to solve emotionally challenging problems, as well as identification of an impairment termed perseveration. Such a maladaptive reaction to a challenging situation is characterized by difficulty in terminating an unsuccessful response, leading to persistent repetition of inappropriate behavior. This issue is important because perseveration is associated with schizophrenia, drug abuse, and aging. © 2020 Wiley Periodicals LLC. Basic Protocol : Water escape detour task Support Protocol 1 : Preparation of escape platform Support Protocol 2 : Preparation of the transparent barrier Alternate Protocol : Water escape detour task for testing acute effects
Down syndrome (DS) is the most frequent genetic cause of intellectual disability, characterized by alterations in different behavioral symptom domains: neurodevelopment, motor behavior, and cognition. As mouse models have the potential to generate data regarding the neurological basis for the specific behavioral profile of DS, and may indicate pharmacological treatments with the potential to affect their behavioral phenotype, it is important to be able to assess disease-relevant behavioral traits in animal models in order to provide biological plausibility to the potential findings. The field is at a juncture that requires assessments that may effectively translate the findings acquired in mouse models to humans with DS. In this article, behavioral tests are described that are relevant to the domains affected in DS. A neurodevelopmental behavioral screen, the balance beam test, and the Multivariate Concentric Square Field test to assess multiple behavioral phenotypes and locomotion are described, discussing the ways to merge these findings to more fully understand cognitive strengths and weaknesses in this population. New directions for approaches to cognitive assessment in mice and humans are discussed. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Preweaning neurodevelopmental battery Basic Protocol 2: Balance beam Basic Protocol 3: Multivariate concentric square field test (MCSF).
The 24-hr cycle of activity and sleep provides perhaps the most familiar example of circadian rhythms. In mammals, circadian activity rhythms are generated by a master biological clock located in the hypothalamic suprachiasmatic nuclei (SCN). This clock is synchronized (entrained) to the external light environment via light input from retinal photoreceptors. However, sleep is not a simple circadian output and also is regulated by a homeostatic process whereby extended wakefulness increases the need for subsequent sleep. As such, the amount and distribution of sleep depends upon the interaction between both circadian and homeostatic processes. Moreover, the study of circadian activity and sleep is not confined only to these specialized fields. Sleep and circadian rhythm disruption is common in many conditions, ranging from neurological and metabolic disorders to aging. Such disruption is associated with a range of negative consequences including cognitive impairment and mood disorders, as well as immune and metabolic dysfunction. As circadian activity and sleep are hallmarks of normal healthy physiology, they also provide valuable welfare indicators. However, traditional methods for the monitoring of circadian rhythms and sleep in mice can require separate specialized resources as well as significant expertise. Here, we outline a low-cost, non-invasive, and open-source method for the simultaneous assessment of circadian activity and sleep in mice. This protocol describes both the assembly of the hardware used and the capture and analysis of data without the need for expertise in electronics or data processing. © 2020 Wiley Periodicals LLC. Basic Protocol: Assembly of a PIR system for basic activity and sleep recordings Alternate Protocol: Data collection using Raspberry Pi Support Protocol: Circadian analysis using PIR sensors.
One of the most intriguing functions of neutrophils is the production of neutrophil extracellular traps (NETs), which are formed when neutrophils decondense their internal DNA and extrude it along with cytotoxic proteins in a web-like structure. This process allows neutrophils to trap and kill pathogens, and is also associated with multiple hematological and autoimmune conditions. Due to their rapid degradation, there are many challenges in accurately and specifically detecting and quantifying NETs. Microscopy is the gold standard for NET detection, but is not optimal for large-scale screening. Furthermore, methods relying on detection of free DNA or on flow cytometry-based examination of NET-associated markers can be nonspecific, time-consuming, and expensive. Here, we describe an innovative, quick, specific, and inexpensive conventional flow cytometry method for detecting neutrophils on the verge of forming NETs. These methods utilize pulse-shaped analysis (PulSA) to distinguish resting neutrophils from those with decondensed DNA, a prerequisite for NET formation. An increase in DNA-diffuse neutrophils is found in cell populations after exposure to NET-inducing stimuli, consistent with the DNA decondensation expected during neutrophil NET formation. These populations are only observed in granulocytes, validating the specificity of this method. We describe protocols optimized for neutrophils retrieved from mouse blood, spleen, and bone marrow. The relative speed and simplicity of the method described here makes it a useful tool for detecting NET formation in large-scale experiments. © 2020 Wiley Periodicals LLC. Basic Protocol: Detection of nuclear decondensation in neutrophils from stimulated murine bone marrow Alternate Protocol 1: Detection of nuclear decondensation in neutrophils from splenocytes Alternate Protocol 2: Detection of nuclear decondensation in neutrophils from blood Support Protocol 1: Cryopreservation and defrosting of samples Support Protocol 2: Paraformaldehyde fixation of samples.
Traumatic brain injury (TBI) is a leading cause of morbidity and mortality among trauma patients. Increased intestinal permeability plays an important role in the inflammatory process that accompanies TBI, and therapies that prevent this permeability change may improve outcomes in TBI patients. Different animal models have been developed to test permeability changes, but there has been no agreement on when permeability should be tested after TBI. Here, we describe a method for creating the TBI mouse model and for measuring intestinal permeability. We also detail our permeability measurements at different time points after TBI to help guide future experimental design. The TBI is made using a controlled cortical impact model with the cortical impactor set to speed 6 m/s, depth 3 mm, dwell time 0.2 s, and tip size 3 mm to produce a severe TBI. Permeability is measured at 2, 4, 6, and 24 hr after TBI by removing a piece of terminal ileum, tying the ends, filling the lumen with FITC-labeled dextran, and then measuring how much of the dextran moves into the surrounding solution bath over time using a fluorescent plate reader. Our results show that peak permeability occurs between 4 and 6 hr after TBI. We recommend that future experiments incorporate permeability measurements 4 to 6 hr after TBI in order to take advantage of this peak permeability. © 2020 Wiley Periodicals LLC. Basic Protocol: Mouse CCI traumatic brain injury model and intestinal permeability measurement.
Research into mechanisms underlying lung injury and subsequent repair responses is currently of paramount importance. There is a paucity of models that bridge the gap between in vitro and in vivo research. Such intermediate models are critical for researchers to decipher the mechanisms that drive repair and to test potential new treatments for lung repair and regeneration. Here we report the establishment of a new tool, the Acid Injury and Repair (AIR) model, that will facilitate studies of lung tissue repair. In this model, injury is applied to a restricted area of a precision-cut lung slice using hydrochloric acid, a clinically relevant driver. The surrounding area remains uninjured, thus mimicking the heterogeneous pattern of injury frequently observed in lung diseases. We show that in response to injury, the percentage of progenitor cells (pro surfactant protein C, proSP-C and TM4SF1 positive) significantly increases in the injured region. Whereas in the uninjured area, the percentage of proSP-C/TM4SF1 cells remains unchanged but proliferating cells (Ki67 positive) increase. These effects are modified in the presence of inhibitors of proliferation (Cytochalasin D) and Wnt secretion (C59) demonstrating that the AIR model is an important new tool for research into lung disease pathogenesis and potential regenerative medicine strategies.
The mouse is the most widely used animal model in hearing research. Immunohistochemistry and immunofluorescent staining of murine cochlear sections have, thus, remained a backbone of inner ear research. Since many primary antibodies are raised in mouse, the problem of "mouse-on-mouse" background arises due to the interaction between the anti-mouse secondary antibody and the native mouse immunoglobulins. Here, we describe the pattern of mouse-on-mouse background fluorescence in sections of the postnatal mouse cochlea. Furthermore, we describe a simple double-blocking immunofluorescence protocol to label mouse cochlear cryosections. The protocol contains a conventional blocking step with serum, and an additional blocking step with a commercially available anti-mouse IgG blocking reagent. This blocking technique virtually eliminates the "mouse-on-mouse" background in murine cochlear sections, while adding only a little time to the staining protocol. We provide detailed instructions and practical tips for tissue harvesting, processing, and immunofluorescence-labeling. Further protocol modifications are described, to shorten the duration of the protocol, based on the primary antibody incubation temperature. Finally, we demonstrate examples of immunofluorescence staining performed using different incubation times and various incubation temperatures with a commercially available mouse monoclonal primary antibody. © 2020 The Authors. Basic Protocol: Tackling the Mouse-on-Mouse Problem in Cochlear Immunofluorescence: A Simple Double-Blocking Protocol for Immunofluorescent Labeling of Murine Cochlear Sections with Primary Mouse Antibodies.
Mass spectrometry-based proteomics provides a robust and reliable method for detecting and quantifying changes in protein abundance among samples, including cells, tissues, organs, and supernatants. Physical damage or inflammation can compromise the ocular surface permitting colonization by bacterial pathogens, commonly Pseudomonas aeruginosa, and the formation of biofilms. The interplay between P. aeruginosa and the immune system at the site of infection defines the host's ability to defend against bacterial invasion and promote clearance of infection. Profiling of the ocular tissue following infection describes the nature of the host innate immune response and specifically the presence and abundance of neutrophil-associated proteins to neutralize the bacterial biofilm. Moreover, detection of unique proteins produced by P. aeruginosa enable identification of the bacterial species and may serve as a diagnostic approach in a clinical setting. Given the emergence and prevalence of antimicrobial resistant bacterial strains, the ability to rapidly diagnose a bacterial infection promoting quick and accurate treatment will reduce selective pressure towards resistance. Furthermore, the ability to define differences in the host immune response towards bacterial invasion enhances our understanding of innate immune system regulation at the ocular surface. Here, we describe murine ocular infection and sample collection, as well as outline protocols for protein extraction and mass spectrometry profiling from corneal tissue and extracellular environment (eye wash) samples. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Murine model of ocular infection Basic Protocol 2: Murine model sample collection Basic Protocol 3: Protein extraction from eye wash Basic Protocol 4: Protein extraction from corneal tissue Basic Protocol 5: Mass spectrometry-based proteomics and bioinformatics from eye wash and corneal tissue samples.
Over the last century, the study of mouse behavior has uncovered insights into brain molecular mechanisms while revealing potential causes of many neurological disorders. To this end, researchers have widely exploited the use of mutant strains, including those generated in mutagenesis screens and those produced using increasingly sophisticated genome engineering technologies. It is now relatively easy to access mouse models carrying alleles that faithfully recapitulate changes found in human patients or bearing variants of genes that provide data on those genes' functions. Concurrent with these developments has been an appreciation of the limitations of some current testing platforms, especially those monitoring complex behaviors. Out-of-cage observational testing is useful in describing overt persistent phenotypes but risks missing sporadic or intermittent events. Furthermore, measuring the progression of a phenotype, potentially over many months, can be difficult while relying on assays that may be susceptible to changes in the testing environment. In recent years, there has also been increasing awareness that measurement of behaviors in isolation can be limiting, given that mice attempt to hide behavioral cues of vulnerability. To overcome these limitations, laboratory animal science is capitalizing on progress in data capture and processing expertise. Moreover, as additional recording modes become commonplace, ultrasonic vocalization recording is an appealing focus, as mice use vocalizations in various social contexts. Using video and audio technologies, we record the voluntary, unprovoked behaviors and vocalizations of mice in social groups. Adoption of these approaches is undoubtedly set to increase, as they capture the round-the-clock behavior of mouse strains. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Continuous recording of home cage activity using the Home Cage Analyzer (HCA) system Support Protocol: Subcutaneous insertion of a radio frequency identification microchip in the inguinal area Basic Protocol 2: Continuous recording of mouse ultrasonic vocalizations in the home cage.
In situ hybridization is a powerful technique that allows the visualization of specific RNA species in biological samples in exquisite detail. It has been particularly well explored in the field of developmental genetics. The spatial and temporal patterns of RNA expression provide us with critical information on likely gene function during embryonic development, and often inform the decision on whether to attempt further gene manipulation approaches. Furthermore, once a mouse strain with altered gene function has been created, in situ hybridization is a critical tool for revealing how the development of embryos with the mutation differs from that of wild-type embryos, and thus infer the function of the altered gene. Here, a well-tested protocol used to visualize RNA expression in whole-mount mid-gestation mouse embryos ranging from 8.5 to 14.5 days post-coitum (dpc) is described. © 2020 Wiley Periodicals LLC. Basic Protocol 1: RNA probe synthesis Alternate Protocol: Preparation of DNA template by PCR Basic Protocol 2: Embryo dissection Basic Protocol 3: Whole mount in situ hybridization Support Protocol: Generation of embryo powder.
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 The Authors. 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.
