Cre mRNA V2 (N1-methyl-pseudouridine)

Gene editing modify specific sites within the genome through gene deletions, insertions or conversions to study functionally unknown genes or conduct gene therapy. It is also used to change the biological traits of organisms to establish new varieties. Gene editing techniques include zinc finger nuclease (ZFN), transcription activator-like effector nuclease (TALEN), and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas 9) (CRISPR/Cas9).

By lysing cells, releasing RNA, and removing impurities such as proteins and DNA, high-purity RNA products are finally obtained. The commonly used traditional method is the guanidine isothiocyanate/phenol/chloroform method (Trizol), which is suitable for a variety of animal materials including animal tissues, microorganisms, cultured cells, etc., and most plant materials.

The inducible CreER/CreERT2 system is based on a fusion between Cre recombinase and a modified estrogen receptor ligand-binding domain that retains Cre in the cytoplasm under basal conditions and allows nuclear translocation upon tamoxifen binding, enabling temporal control of site-specific recombination at loxP-flanked genomic loci in vivo or in vitro. Upon tamoxifen administration, CreER translocates to the nucleus and catalyzes recombination between loxP sites, resulting in excision or inversion of floxed DNA segments and enabling temporally defined gene knockout in specific tissues depending on promoter-driven CreER expression. This system has been widely used for inducible gene deletion and lineage tracing in mice, including validation of efficient temporal recombination in developmental and adult tissues using tamoxifen induction paradigms.

CRISPR-Cas9 knockout in cultured mammalian cells uses an sgRNA to direct Cas9 to a complementary genomic sequence adjacent to a compatible PAM; Cas9 creates a targeted DNA double-strand break, and repair by non-homologous end joining can introduce insertions or deletions that disrupt the coding sequence or functional genomic element. The readout of knockout is detection of edited alleles and loss of gene product or phenotype, commonly by PCR/Sanger-sequence trace decomposition, targeted sequencing, immunoblotting, immunostaining, or flow cytometry when the target protein is detectable at the cell surface.

The Cre-lox conditional knockout system is a site-specific recombination technology in which Cre recombinase recognizes loxP sites flanking a genomic sequence, enabling excision, inversion, or rearrangement of the intervening DNA in a spatially and/or temporally controlled manner, allowing tissue-specific gene inactivation in vivo when Cre is expressed under defined promoters. This system is widely used to overcome limitations of conventional germline knockouts, particularly embryonic lethality or systemic effects that obscure tissue-specific gene function, by restricting recombination to selected cell types or developmental stages. Cre-mediated recombination efficiency and specificity depend on factors such as Cre driver expression pattern, loxP configuration, and genomic context, which can lead to mosaic recombination and variable knockout outcomes. The system is typically validated using reporter alleles or floxed target genes to confirm recombination at DNA and protein levels.

CRISPR-Cas9 zebrafish embryo editing introduces targeted double-strand breaks in genomic DNA by delivering Cas9 nuclease with a guide RNA into one-cell-stage embryos; repair by endogenous DNA-repair pathways produces indels or donor-mediated insertions that can be detected by phenotype, PCR-based genotyping, heteroduplex assays, Sanger sequencing, or amplicon sequencing. The readout reflects the frequency and spectrum of edited alleles in mosaic F0 embryos or transmitted F1 animals; because injected embryos can carry multiple alleles, founder screening and sequence confirmation are required before establishing stable mutant lines.

Constitutive germline knockout models are generated by producing a heritable loss-of-function allele in the mouse genome, typically through complete gene disruption in embryonic stem (ES) cells followed by germline transmission or through CRISPR/Cas-mediated editing of zygotes, resulting in offspring that carry a stable null allele in all tissues. Classical approaches rely on homologous recombination in ES cells to introduce targeted gene disruptions, which are then transmitted through chimeric mice to the germline. More recent genome editing strategies use CRISPR/Cas systems to induce double-strand breaks and non-homologous end joining (NHEJ), frequently generating frameshift mutations that abolish gene function, enabling faster generation of knockout alleles directly in embryos. Germline transmission or direct germline editing ensures that the mutation is present in all cells of the resulting animal, allowing systemic functional analysis of gene loss.

CRISPR/Cas9 knockout animal modeling uses guide RNA to direct Cas9 to a genomic target, where Cas9 creates a DNA double-strand break; repair by error-prone non-homologous end joining generates insertions or deletions that can disrupt coding sequence and produce knockout alleles. Classic animal-model workflows deliver Cas9 mRNA or Cas9 protein with sgRNA into fertilized zygotes by microinjection or electroporation, then transfer edited embryos into pseudopregnant recipients and genotype founders for target-site mutations.

Cre-lox lineage tracing labels cells that express Cre recombinase by excising a loxP-flanked STOP cassette in a reporter allele, causing permanent reporter expression in the recombined cell and its descendants. Inducible CreERT2 models add temporal control because tamoxifen activates CreERT2-dependent recombination, allowing cells expressing a selected promoter at a chosen time point to be fate-mapped during development, homeostasis, injury, or disease. Reporter readouts include β-galactosidase, EYFP/ECFP, tdTomato, membrane GFP, and multicolor fluorescent proteins; these readouts detect recombined cells by histology, fluorescence microscopy, flow cytometry, or tissue imaging.