How to Make Knockout Mice: Step-by-Step Explanation

Genetically modified mice are essential tools for studying gene function and modeling human diseases. Knockout mice, which have specific genes inactivated, help researchers understand how particular genes contribute to development, physiology, and disease processes.

Creating a knockout mouse requires careful planning and execution to ensure accuracy and efficiency.

Gene Selection And Targeting Strategy

Selecting a gene for knockout experiments involves understanding its function, expression patterns, and potential compensatory mechanisms. Researchers consult literature, genomic databases like Mouse Genome Informatics (MGI) and Ensembl, and use RNA sequencing (RNA-seq) to assess tissue-specific expression.

A common targeting strategy involves deleting critical exons to induce a frameshift mutation, leading to a premature stop codon and nonsense-mediated decay of mRNA. Alternatively, replacing an essential coding region with a reporter gene like β-galactosidase or GFP allows visualization of gene expression while inactivating the gene. The approach depends on factors like gene size, alternative splicing, and redundancy with paralogous genes.

Homology-directed repair (HDR) is used for precise gene targeting in embryonic stem (ES) cells. Long homology arms—typically 5–10 kilobases—flank the mutation site to enhance recombination efficiency. Computational tools such as CRISPOR and Benchling assist in designing homology arms and predicting off-target effects. Selecting a genomic region with high recombination frequency improves success rates.

Constructing The Targeting Vector

The targeting vector facilitates homologous recombination and includes selectable markers to distinguish correctly modified cells. It consists of long homology arms, a selectable resistance gene, and a counterselection marker to eliminate random integrations. Longer homology arms improve targeting efficiency.

Homology arms are amplified from genomic DNA using high-fidelity PCR or retrieved from bacterial artificial chromosome (BAC) libraries. These arms are then cloned into a plasmid backbone, often containing a neomycin resistance cassette for positive selection in ES cells. A counterselection marker like diphtheria toxin A (DTA) or thymidine kinase (TK) helps remove cells with random integrations.

To disrupt gene function, the vector may introduce a frameshift mutation by excising essential exons or inserting a stop codon. A reporter gene like LacZ or GFP can replace the deleted region to track gene expression. Alternative splicing events must be considered to ensure complete gene inactivation.

Once assembled, the vector is validated using restriction enzyme digestion, Sanger sequencing, and pulse-field gel electrophoresis. Southern blot analysis ensures the absence of unwanted rearrangements. The construct is then linearized to enhance recombination efficiency before transfection into ES cells.

Embryonic Stem Cell Transfection

The targeting vector is introduced into ES cells using electroporation, which transiently permeabilizes cell membranes, allowing DNA entry. Electrical parameters—typically 240–300 V with 500–700 µF capacitance—must be optimized to balance efficiency and cell viability.

Following electroporation, ES cells are plated onto mitotically inactivated feeder layers, usually mouse embryonic fibroblasts, which provide essential growth factors. Culture conditions must be maintained with leukemia inhibitory factor (LIF) to prevent differentiation.

Selective pressure is applied 24–48 hours post-transfection using a selective agent like G418 for neomycin resistance. Only cells with homologous recombination survive. A counterselection agent like ganciclovir may be used to eliminate cells with random integration.

Screening For Modified Stem Cells

After selection, individual ES cell colonies are isolated and expanded in culture. PCR is used for initial screening, with primers designed to amplify sequences spanning the junction between the endogenous locus and the inserted vector. Quantitative PCR (qPCR) assesses copy number variation.

Southern blot analysis provides definitive verification. Genomic DNA is digested with restriction enzymes, and hybridization with a radiolabeled probe confirms correct modification. This method distinguishes homologous from random integrations.

Blastocyst Injection And Transfer

Correctly targeted ES cells are introduced into developing embryos to generate chimeric mice. Blastocysts, typically harvested from superovulated female mice, serve as hosts. The donor blastocyst strain is chosen to allow easy identification of chimeric offspring, often differing in coat color from the ES cells.

Microinjection delivers 10–15 targeted ES cells into the blastocoel cavity using a micromanipulator and fine glass capillary needles. Injected blastocysts are transferred into pseudopregnant female mice, hormonally primed for implantation. Timing is critical, with embryos introduced at embryonic day 3.5. Pregnancy progression is monitored via ultrasound or palpation.

Detection Of Chimeric Offspring

Chimeric mice are identified by coat color differences. If ES cells from an agouti strain are injected into albino blastocysts, chimeras exhibit a mottled coat pattern. Higher chimerism increases the likelihood of germline transmission.

Molecular techniques like PCR-based assays detect strain-specific genetic markers, providing a quantitative measure of targeted cell contribution. Fluorescent in situ hybridization (FISH) can visualize chromosomal differences between donor and host cells.

Breeding For Germline Transmission

To establish a stable knockout line, chimeric mice must transmit the modified allele through the germline. High-percentage chimeras are bred with wild-type mice to obtain heterozygous offspring, which are then intercrossed to generate homozygous knockouts.

Genetic confirmation of germline transmission is performed through PCR or Southern blot analysis of offspring DNA. If transmission is unsuccessful, additional chimeras may be bred or alternative strategies like tetraploid complementation may be considered.

Genotyping And Verification

Knockout mice are verified through genotyping. DNA from tail biopsies or ear punches is analyzed using PCR, with primers distinguishing wild-type, heterozygous, and homozygous knockout alleles. Multiplex PCR streamlines screening by amplifying multiple regions simultaneously.

Southern blotting confirms correct integration and absence of unexpected rearrangements. Quantitative RT-PCR assesses gene silencing, ensuring functional knockout.

Characterizing The Knockout

Once genetic modification is confirmed, phenotypic and functional assessments determine the gene’s role. Initial characterization evaluates viability, growth, and reproductive success. Behavioral assessments may be conducted for genes involved in neurological or developmental pathways.

Functional studies include tissue-specific expression analysis via Western blotting or immunohistochemistry. RNA sequencing (RNA-seq) identifies downstream gene expression changes and compensatory pathways. Additional studies, such as metabolic profiling or immune system assessments, further delineate the knockout’s impact.