Transgenic Technology

The generation and characterization of genetically altered mice is intensive, time consuming, and technically demanding, and it includes preparation of the construct, selection of the mouse strain for embryo donation or ES (embryonic stem) clone, microinjection of DNA into pronuclei or targeted ES cells into blastocysts, identification of founder animals or chimeras, breeding and testing of transgenic progeny, and maintenance of colony records and health. It is essential to have the necessary expertise and resources to be successful in the development of new genetically altered animal models.

Popular mouse background strains used to generate mouse models include (C57Bl/6XC3H/He)F1 and (C57Bl/6XDBA/2)F1. However, the problems of genetic variation between F2 animals, genetic drift in subsequent generations, and the extensive backcrossing onto an inbred background required to regain genetic definition, are greatly minimized by using an inbred embryo strain. The disadvantages of inbred strains are decreased breeding efficiency, poor ovulators, eggs with small pronuclei (thus more challenge for microinjection), and increased susceptibility to lysis. The inefficiencies of C57BL/6 in superovulation, microinjection, and reproduction can be minimized by monitoring specific biological end points such as pronuclear egg formation and response to gonadotropin as well as diet, age, and light exposure. FVB and SWR female mice are highly suitable for the propagation of transgenes, as they have high ovulation rates, and oocytes are resistant to lysis.4 For production of mutant mice via the microinjection of gene-targeted embryonic stem (ES) cells, inbred strains are routinely used as blastocyst donors to provide a constant genetic background for the production of chimeras. Generally, targeted ES cells (most commonly 129/SvJ and C57Bl/6 derived) are injected into BALB/c or C57BL/6 blastocysts, respectively. Pseudopregnant recipients can be any strain with high breeding efficiency. A popular outbred strain is Swiss Webster, which is an excellent breeder and exceptional foster mother (personal observations).

Procedures for generating transgenic mice have been described in detail.5 In general, the transgene constructs for generating transgenic mice consist of a functional promoter, initiation codon, polyadeny-lation site, and full length cDNA or genomic DNA for a specific gene fused to the enhancer and promoter sequence and cloned into an appropriate vector. Because many vectors used in cloning can interfere with expression of the transgene, it is important that unique restriction sites at the 5' and 3' ends of the transgene are available to remove plasmid sequences prior to injection. Thermocycling with probes specific to the transgene is the most frequently used method for analyzing transgene integration, although Southern and dot blot analyses are also used. The choice of promoter depends on the target tissue of interest or the desire to direct the ubiquitous expression of a transgene. Promoters such as chick P-actin and cytomegalovirus are frequently used as broad-spectrum promoters by focusing the expression of genes to a broad array of tissues.6 However, these promoters have limitations, because expression in liver is low or undetectable, and it is variable in other organs. Organ-specific promoters such as the rat albumin enhancer and promoter, which targets gene expression to the liver, and the lck promoter, which targets immature T cells, have been used successfully in various studies. Inducible promoters such as the tetracycline-inducible system can also be employed, because they are preferable in a number of situations in order to obtain maximum quantity of the gene product and to regulate its expression.

The gene of interest is excised from its vector sequences, purified by gel electrophoresis, and linearized. Linearized DNA of less than 10 kb in length is commonly used, because DNA with higher molecular weight becomes too viscous, making it difficult to load and inject through a 1 to 2 |im opening of a microinjection needle. The usual concentration of DNA for injection is 2 to 5 ng/uL. The foreign DNA is introduced into the murine germ line by microinjection of the pronuclei of one-cell fertilized eggs. Routinely, between 100 and 200 embryos are injected, which are then implanted into pseudopregnant females. Approximately 10 to 30% of the transferred eggs result in live births with litter sizes of five to eight per recipient. The pregnant females are monitored for phenotypic abnormalities during gestation, such as embryo reabsorption. Pups in the initial perinatal period are monitored for lack of suckling reflex, as well other abnormalities. At 2 to 4 weeks of age, the founder pups are screened for the presence of the transgene integration using tail or ear punch biopsies and PCR analysis. Southern blot analysis, while technically more cumbersome, provides an estimate of transgene copy number and is used to confirm PCR results. Once transgene positive pups are identified, they are raised to sexual maturity and mated to wild-type mice of the desired background strain. Transgene positive offspring will confirm that line as a permanent founder.

Embryonic stem cell procedures have been described in detail.5 The ES cell lines used are of 129 derivation or C57Bl/6 origin. Homologous recombination vectors are transfected into ES cells via electroporation. The vectors are linearized and used at a concentration of about 1 | g/mL free of toxic contaminants, such as ethanol, ethidium bromide, and excess salt. Electroporation is performed on 2 x 107 ES cells in PBS (without Ca++ and Mg++) containing 25 |g vector. A short high-voltage electrical pulse is applied to the ES-vector mixture, which allows for pore development in the ES cell membranes and entry of the vector. A postelectroporation killing rate of 50%, as assessed by tryptan blue exclusion, is an early indication of a successful electroporation. Following electroporation, the cells are plated with selection media to isolate the targeted clones. When colonies of resistant cells are detected, they are tested by PCR or Southern blot for proper genetic manipulations. Because genetic drift and maintenance of totipotency are exquisitely sensitive to culture conditions, generating large pools of low-passage frozen stocks are required for successful thaw and chimera production.

Microinjection of the targeted ES cells into blastocyst (E3.5) stages is used to produce targeted genetic mutant mice. ES cells are injected into the space between the uncompacted eight-cell embryo and the zona pellucida, into the center of a compacted eight-cell/morula or into the blastocoel of a blastocyst, whether unhatched or hatched. Five to 15 ES cells are injected into each blastocyst, and nine to 15 of the microinjected blastocysts are transferred to the uterus of a pseudopregnant recipient. Pups are born approximately 18 days from transfer. Detection of chimerism is possible as early as 3 days later but is not confirmed and quantified until hair grows in, 5 to 7 days after birth. Only male chimeras, where >50% of the coat color is from the ES contribution, are selected for testing germ line transmission of the induced mutation. The male chimeras are test mated with C57BL/6 females. The resulting black offspring are derived only from a host embryo background, whereas black agouti offspring are derived from the ES cell contribution, due to coat color genetics of all the ES 129 substrains. These black agouti pups are further tested, using tail or ear biopsies, to determine whether the wild type or the targeted allele is transmitted. Pups determined to be heterozygous (+/-) for the mutations are retained for breeding.

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