Molecular Biotechnology: Principles and Applications - Glick, B., & Pasternak, J. 2002

Molecular Biotechnology of Microbiological Systems
Transgenic Animals
Transgenic Mice: Methodology

Transgenic technologies were originally developed and refined using laboratory mice. Since the early 1980s, hundreds of genes have been introduced into various mouse strains. These studies have significantly contributed to elucidating the mechanisms of Gene regulation and tumor development, The Nature of immunological Specificity, the Molecular Genetics of GROWTH AND DEVELOPMENT, and other fundamental biological processes. Transgenic Mice have played a vital role in exploring the feasibility of large-scale pharmaceutical synthesis, as well as in generating transgenic lines that serve as models for various human genetic diseases. Foreign DNA can be introduced into mice using several approaches: 1) via retroviral vectors that infect embryonic Cells at early developmental stages prior to embryo implantation into a recipient female; 2) by microinjection into the enlarged sperm Nucleus (male pronucleus) of a fertilized oocyte; 3) by introducing genetically modified Embryonic Stem Cells into a preimplantation embryo at an early stage of development. Use of Retroviral Vectors The main advantage of using retroviral vectors (Fig. 19.1) over other transgenesis Methods lies in their high efficiency. However, the insert size in this case is limited to approximately 8 kb, which may result in the transgene lacking the flanking regulatory sequences required for its expression.

The Use of retroviral vectors also presents another major drawback. Although these vectors are engineered to be Replication-defective, The Genome of the retroviral strain (helper virus) required to produce large quantities of vector DNA can occasionally enter the same nucleus as the transgene. Despite all precautions, helper Retroviruses can potentially replicate within the transgenic animal—an entirely unacceptable outcome if these animals are intended for human consumption or as a source of commercial products. Because alternative methods of transgenesis are available, retroviral vectors are rarely employed to generate commercially valuable Transgenic Animals.

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Fig. 19.1. Generation of transgenic mouse lines using retroviral vectors. An embryo, typically at the 8-Cell stage, is infected with a recombinant retrovirus carrying the transgene. Females implanted with the embryo ("surrogate" mothers) give birth to transgenic offspring. A series of test crosses is then performed to identify pups that carry the transgene in their germline cells.

DNA Microinjection Method

Currently, DNA microinjection is the most widely used method for generating transgenic mice. The Procedure involves the following steps (Fig. 19.2).

1. Increasing the number of oocytes available for foreign DNA injection by stimulating superovulation in donor females. Females are first treated with pregnant mare serum gonadotropin, followed approximately 48 hours later by human chorionic gonadotropin. Superovulation yields roughly 35 oocytes instead of the normal 5–10.

2. Mating superovulated females with males and subsequently euthanizing them. Flushing the fertilized oocytes from the oviducts.

3. Microinjection of DNA into the fertilized oocytes—typically performed immediately after isolation. Frequently, the injected transgene construct is in a linear form and lacks prokaryotic vector sequences.

Fig. 19.2. Production of transgenic mouse lines via microinjection. Oocytes are isolated from donor females subjected to induced superovulation and mating. The transgene construct is microinjected into the male pronucleus of a fertilized oocyte. The oocytes are then transferred to a surrogate mother, which gives birth to transgenic pups—the founders of transgenic lines.

In mammals, following sperm penetration into the oocyte, the sperm nucleus (male pronucleus) and the oocyte nucleus exist separately. Once the latter completes mitotic division and becomes the female pronucleus, nuclear fusion (karyogamy) can take place. The male pronucleus is typically much larger than the female one, making it easy to locate under a dissecting Microscope and microinject with foreign DNA. During the microinjection process, the oocyte can be temporarily moved, correctly oriented, and immobilized. An experienced investigator can inoculate several hundred oocytes in a single day.

Following DNA Introduction, 25 to 40 oocytes are microsurgically implanted into a surrogate mother rendered pseudopregnant by mating with a vasectomized male. In mice, mating is the only known physiological trigger that prepares the Uterus for implantation. Because a vasectomized male produces no sperm, none of the surrogate mother's own oocytes are fertilized. Embryos develop exclusively from the transferred oocytes, and pups are born approximately 3 weeks post-implantation.

To identify transgenic animals, DNA is extracted from a small tail biopsy and tested for the presence of the transgene using Southern blot Hybridization or the Polymerase Chain Reaction (PCR). To determine whether the transgene has integrated into the animal's germline, the transgenic mouse is crossed with a wild-type counterpart. Subsequent intercrossing of the progeny allows for the establishment of pure (homozygous) transgenic lines. Although the described approach appears relatively straightforward at first glance, it requires precise coordination across multiple stages. Even highly skilled specialists achieve viable transgenic animals from at best only about 5% of the inoculated oocytes (Fig. 19.3). Because no single experimental step is 100% efficient, A large number of fertilized oocytes must be used for microinjection. For instance, in generating transgenic mice via DNA microinjection, only about 66% of the fertilized oocytes survive; roughly 25% of the implanted oocytes develop into pups, of which only 25% actually carry the transgene. Consequently, out of 1,000 implanted fertilized oocytes, only 30 to 50 transgenic pups typically develop. Furthermore, the introduced DNA can integrate into virtually any genomic locus, frequently resulting in multiple copies inserting at a single site. Finally, not all transgenic pups will exhibit the desired phenotypic characteristics. In some individuals, the transgene may fail to express due to an unfavorable integration site environment, whereas in others, the excessive copy number of the foreign gene can lead to protein overexpression and disrupted normal physiological processes. Nevertheless, despite these challenges, microinjection remains a frequently utilized method for establishing mouse lines harboring functional transgenes.

Fig. 19.3. Overall efficiency of transgenesis following microinjection. All fertilized oocytes (100%) from cattle, pigs, sheep, and mice were inoculated with the transgene; however, successful implantation and live birth were rare events, with transgene-positive offspring resulting from fewer than 5% of the manipulated oocytes.

Use of Modified Embryonic Stem Cells

Cells isolated from mouse embryos at the blastocyst stage can proliferate in culture while maintaining their pluripotency—the capacity to differentiate into any cell type, including germline cells, upon introduction into another blastocyst-stage embryo. These cells are known as pluripotent embryonic stem (ES) cells. In culture, ES cells are readily amenable to Genetic Engineering without loss of pluripotency; for instance, a functional transgene can be targeted to a specific site within a non-essential genomic locus. Genetically altered cells can then be selected, cultured, and used to generate transgenic animals (Fig. 19.4). This strategy avoids the random integration events characteristic of DNA microinjection and retroviral vector systems.

When ES cells in culture are transfected with a vector designed for site-specific chromosomal integration, DNA inserts randomly in some cells, integrates at the targeted site in others, and fails to integrate altogether in the majority of ES cells. To enrich for the first category of targeted events, a protocol known as positive-negative Selection is employed. This strategy combines positive selection for cells that have incorporated the vector DNA via Homologous Recombination at the target site with negative selection against cells containing vector DNA integrated at random chromosomal locations.

The target site must reside within a region of genomic DNA that does not encode vital Proteins, ensuring that foreign DNA insertion does not disrupt developmental processes or essential cellular Functions. Additionally, it is crucial that transgene integration does not block the METABOLISM/31.html">Transcription of neighboring genomic regions. The search for such optimal sites is ongoing. A typical positive-negative selection vector comprises the following elements: 1) two homologous sequences (HB1 and HB2) matching specific Regions of the target site; 2) a transgene (TG) conferring a novel function to the recipient; 3) a sequence encoding resistance to the antibiotic G-418 (Neo^r); and 4) two distinct Herpes simplex virus thymidine kinase genes (tk1 and tk2) derived from types 1 and 2 (HSV-tk1 and HSV-tk2) (Fig. 19.5, A). The spatial arrangement of these elements is critical for successful positive-negative selection. The transgene and the G-418 resistance gene (Neo^r) must be positioned between the two Homology blocks corresponding to the target site, whereas the HSV-tk1 and HSV-tk2 genes flank this entire cassette. If integration occurs at a random site (outside HB1 and HB2), one or both HSV-tk genes will almost certainly integrate alongside the other sequences (Fig. 19.5, A). Conversely, if integration occurs via homologous recombination through a double crossover event at the targeted locus, only the transgene and the Neo^r gene will integrate into the genome, leaving behind the HSV-tk genes (Fig. 19.5, B). When transfected cells are cultured in the presence of G-418, cells lacking the Neo^r gene fail to survive. Only cells in which integration has successfully occurred will live—representing the positive selection phase. If ganciclovir is simultaneously added to the culture medium alongside G-418, the growth of cells expressing thymidine kinase is inhibited, because this enzyme catalyzes The conversion of ganciclovir into a cytotoxic compound lethal to The Cell, thus achieving negative selection. Cells that survive this dual-filter screening process are highly likely to contain the sequence integrated precisely at the desired target site. Although this method is not entirely infallible, it significantly enriches the cell population for clones harboring the transgene at the specific chromosomal Location.

Fig. 19.4. Generation of transgenic mice via genetic modification of embryonic stem (ES) cells. ES cells are derived from the inner cell mass of a mouse blastocyst, transfected with a transgene-bearing vector, cultured, and identified using positive-negative selection or PCR assays. The surviving transfected cell population is expanded in culture and microinjected into blastocysts, which are subsequently transferred into the uteri of surrogate mothers. By intercrossing founder animals carrying the germline transgene, transgenic mouse lines are established.

Fig. 19.5. Positive-negative selection. A. Non-specific (random) integration. Both thymidine kinase genes (tk1 and tk2), the two DNA segments homologous to specific recipient chromosomal sequences (HB1 and HB2), the gene conferring resistance to the cytotoxic agent G-418 (Neo^r), and the transgene (TG) have all integrated into the chromosome. Following transfection, cells are screened for resistance to G-418 and ganciclovir (which becomes cytotoxic specifically to cells expressing thymidine kinase). Alternative random integration patterns may also occur, such as those resulting in the insertion of only the thymidine kinase genes into the chromosome; however, in the presence of both G-418 and ganciclovir, all such cells likewise perish. B. Site-specific integration via homologous recombination. As a result of a double crossover event between the homologous regions (HB1 and HB2) of the vector and chromosomal DNA, a fragment lacking the thymidine kinase genes (tk1 and tk2) is integrated into the host genome. In the presence of G-418 and ganciclovir, only cells that have undergone successful homologous recombination survive.

A simpler approach for identifying ES cells carrying the transgene at the correct locus relies on PCR screening. In this strategy, the DNA vector contains two homology regions matching the target site—one flanking the transgene side, and the other adjacent to a cloned bacterial or synthetic (unique) sequence absent from the mouse genome (Fig. 19.6). Following transfection of ES cells with this vector, the resulting clones are screened by PCR. One PCR primer (P1) is complementary to the cloned bacterial or synthetic unique nucleotide sequence of the integrated vector, whereas the second primer (P2) anneals to a chromosomal DNA sequence immediately adjacent to one of the homology blocks. If the target sequence has integrated randomly, the expected Amplification product will not be generated (Fig. 19.6, A); conversely, site-specific integration yields a DNA fragment of a predetermined, predictable size upon PCR amplification (Fig. 19.6, B). This approach successfully identifies ES cell pools containing the transgene at the correct target site, allowing researchers to isolate and expand cells from these pools to establish stable cell lines with site-specific insertions.

Fig. 19.6. Identification of cells carrying a transgene at a specific site using PCR. A. Due to non-specific Integration of the vector DNA, one of the primers (P2) fails to hybridize with the chromosomal region located at a specific distance from the annealing site of primer P1, resulting in no amplification product of the expected size. P1 hybridizes with a unique sequence (US) of the inserted DNA that is absent in the recipient cell's chromosomal DNA. B. Homologous recombination between the HB1 and HB2 regions of the inserted DNA on one hand, and the complementary chromosomal regions CS1 and CS2 on the other, generates segments that can hybridize with both primers, P1 and P2, separated by a defined distance. During PCR amplification, fragments of a uniform size are synthesized and can be identified via gel Electrophoresis. If the PCR product of the desired length is formed, it indicates that the transgene (TG) located between the homologous regions (HB1 and HB2) has integrated into a specific chromosomal site.

ES cells whose genome contains a site-specific transgene insertion can be cultured and introduced into a blastocyst-stage embryo, which is then implanted into the uterus of a pseudopregnant surrogate mother. Pups in which the genetically modified ES cells contributed to The formation of the germline can give rise to transgenic lines. To achieve this, they must be crossed with mice of the same strain, followed by intercrossing their transgenic offspring. This yields transgenic mice homozygous for the transgene.

A specific chromosomal site in ES cells can not only receive an inserted transgene encoding a novel function, but can also be targeted for disruption by integrating a specific sequence (typically a selectable marker gene) into its coding region (Fig. 19.7). One of the primary objectives of targeted gene disruption, or "knockout," is to investigate its effects on organismal development and underlying physiological processes. Furthermore, it is hoped that transgenic animals with disruptions in specific genes can serve as models for studying human diseases at THE MOLECULAR LEVEL.

For instance, targeted "knockout" of the mouse rhodopsin gene leads to retinal rod inactivation, mimicking human retinitis pigmentosa. Rhodopsin knockout mice can be utilized to study retinal degeneration as well as the therapeutic efficacy of drugs that slow down or halt this genetically driven pathological process altogether. Over 250 knockout mouse lines have already been established and are widely used as models to investigate various human disorders.

Fig. 19.7. Gene knockout via targeted homologous recombination. The vector carries a selectable marker gene (smg) and flanking sequences homologous to the corresponding regions of the target gene. The latter contains five exons (1–5). As a result of homologous recombination (dashed lines), the target gene is disrupted ("knocked out").

In principle, the approaches used to generate transgenic animals with "enhanced functions" and those with "loss of functions" are similar. Unfortunately, pluripotent ES cells analogous to those in mice have not yet been identified in cattle, sheep, pigs, or chickens, though the search for them continues.

Cloning by Nuclear Transfer

Pluripotency can be demonstrated by transferring The Nucleus of a test cell into an enucleated oocyte and subsequently assessing the ability of the reconstructed cell to develop and produce viable offspring. Several laboratories have investigated the pluripotency of embryonic, fetal, and adult cell lines with varying degrees of success. It has been shown that embryonic cell nuclei are capable of supporting development, albeit with low efficiency. For instance, viable offspring have been obtained via nuclear transfer using short-term cultured bovine embryonic cells. The famous sheep named Dolly was cloned via nuclear transfer from an adult mammary gland (udder) cell (Fig. 19.8). This provided the first definitive proof of the pluripotency of a differentiated adult Cell Nucleus. However, the possibility cannot be ruled out that the donor nucleus actually originated from an undifferentiated cell residing within the donor's mammary epithelium.

The successful cloning of Dolly from a differentiated cell nucleus, along with three other sheep from embryonic cell nuclei, was made possible by utilizing nuclei from cells arrested in the resting phase (G₀) and, potentially, Specific features of ovine Embryogenesis. Specifically, During the first three Divisions of the sheep zygote, which span several days, only DNA replication occurs, and no Genes are transcribed. It is hypothesized that during this period, the introduced DNA is stripped of cell-specific regulatory proteins, while the corresponding embryonic development genes become associated with initiator embryonic protein factors originating from the oocyte Cytoplasm.

The primary hurdle that must be overcome to make transgenic animal production via nuclear transfer a practical reality is the maintenance of cellular pluripotency in continuous culture. Once achieved, genetic modification of such cells and the creation of transgenic organisms will become nearly routine Procedures. However, due to species-specific differences in early embryonic Cleavage timing and the onset of transcription, it remains uncertain whether nuclear transfer will be feasible in domestic livestock species other than sheep if the donor Cell Cycle stage matches that of the oocyte.

Fig. 19.8. Cloning a sheep by nuclear transfer. The oocyte nucleus is removed using a micropipette. Adult mammary epithelial cells are cultured and induced to enter the G₀ phase. The G₀ cells and enucleated oocytes are fused, and the reconstructed oocytes are grown in culture or in a ligated oviduct to early embryonic stages before being implanted into the uterus of a surrogate mother for further development. In the experiment described by Wilmut et al. (1997), 277 enucleated oocytes were fused with G₀ mammary cells; out of 29 embryos, only one developed into a viable fetus.

Gene Transfer Using Yeast Artificial Chromosomes

Most transgenes consist of cDNAs, small genes (<20 kb), or gene fragments. Frequently, cDNAs are poorly expressed in mammalian cells, and when genomic DNA serves as the transgene, critical gene-specific regulatory sequences located upstream and downstream of the target gene are typically missing from the insert. Furthermore, full-length genes and multigene complexes (>100 kb) are far too large for incorporation into standard vectors. To address these limitations, yeast artificial chromosomes (YACs), which can accommodate genomic DNA fragments ranging from 100 to >1000 kb in length, have been adopted for transgenesis.

Transgenic mice have been generated by pronuclear microinjection into fertilized oocytes or by transfecting ES cells with YACs harboring multiple related genes or a single large gene. Transgenic mice carrying a cluster of five functional human ß-globin genes spanning a total length of approximately 250 kb expressed all these genes in a tissue-specific and temporally appropriate manner—precisely as they do in humans. This fidelity is ensured by the flanking sequences containing the promoter and other essential regulatory elements.

The generation of mice that synthesize exclusively human Antibodies represents a remarkable feat of YAC-mediated transgenesis. As noted in Chapter 10, Monoclonal Antibodies can be used to treat various human diseases. However, producing human monoclonal antibodies directly is practically unfeasible. Unfortunately, rodent monoclonal antibodies are also immunogenic in humans. To "humanize" existing rodent monoclonal antibodies, sophisticated recombinant DNA strategies have been developed. These laborious procedures have yielded Fv and Fab fragments that often retain some affinity for a specific antigen. A true technological breakthrough in producing full-length human antibodies may ultimately rely on more accessible hybridoma-based methods.

The synthesis of natural antibodies is nothing short of miraculous. An antibody is a highly complex tetrameric Structure composed of two pairs of distinct polypeptide chains: heavy (H) and light (κ or λ). These designations reflect differences in subunit molecular weights. The genetic architecture of each heavy chain is determined by the combinatorial joining of variable (V_H), diversity (D_H), joining (J_H), and constant (C_H) segments of somatic DNA within a B cell. There are two types of light chains, λ and κ, which are generated via the rearrangement of their respective variable (Vλ, Vκ), joining (Jλ, Jκ), and constant (Cλ, Cκ) domains. An individual B cell synthesizes a single type of antibody characterized by a unique combination of H-chain domains and either a rearranged λ- or κ-chain.

The genetic repertoire responsible for generating the vast diversity of human antibody H chains includes approximately 95 V_H domains, 30 D_H domains, 6 J_H domains, and 5 major constant (Cα, Cγ, Cδ, Cε, Cμ) domains. The κ gene locus comprises approximately 76 Vκ domains, 5 Jκ domains, and a single constant (Cκ) region (Fig. 19.9). The size of the H chain loci and κ genes ranges from 1 to 1.5 Mb. To produce transgenic mice capable of synthesizing a diverse repertoire of human antibodies, the endogenous mouse H and L chain genes must first be inactivated, followed by the introduction of YACs containing the human immunoglobulin H and L chain gene repertoires into the mouse chromosomal DNA.

To accomplish this, the mouse H and κ chain genes were replaced ("knocked out") with a compact segment of the human H chain gene cluster (comprising 4 V_H domains, 16 D_H domains, 6 J_H domains, Cγ, and Cμ) and the human κ chain gene cluster (containing 4 Vκ domains, 5 Jκ domains, and Cκ). Transgenic mice carrying this human antibody gene Complement synthesized human antibodies against select Antigens, and hybridomas producing human monoclonal antibodies were successfully derived. However, The Diversity of human antibodies produced by these transgenic mice was relatively limited due to the restricted repertoire of H and κ variable segments. To overcome this limitation, YACs engineered with a much larger array of human heavy and light chain variable region genes were constructed.

By combining four distinct YACs containing human heavy chain genes, a 1000-kb YAC was assembled, carrying 66 V_H domains, approximately 30 D_H segments, 6 J_H domains, Cμ, Cδ, and Cγ. Similarly, utilizing three YACs bearing various Vκ domains, an 800-kb YAC was constructed containing 32 Vκ domains, 5 Jκ domains, and Cκ. ES cells were transfected separately with the H-chain and κ-chain YACs via Cell Fusion, transformants containing the integrated YACs were selected using a selectable marker, and the integrity of each insert was verified by PCR. Cells carrying the inserted H- or κ-chain genes were microinjected into blastocysts to identify founder animals via PCR. Transgenic mice carrying the H- and κ-chain inserts were subsequently crossed individually with mice harboring inactivated loci for the respective chains. The resulting progeny were then intercrossed to yield mice devoid of functional endogenous mouse H and κ chain genes yet carrying both human heavy and light chain transgene insertions.

Transgenic mice possessing an expanded repertoire of human V_H and Vκ domains synthesized human antibodies. Upon immunization with three distinct antigens, hybridomas secreted human monoclonal antibodies exhibiting high affinity for the immunizing antigens in every case. It is highly probable that such transgenic systems will serve as a reliable platform for producing therapeutic human monoclonal antibodies for clinical Applications.

Fig. 19.9. Schematic diagram of the genes for human immunoglobulin κ and H chains. A. STRUCTURE OF THE immunoglobulin κ-chain gene in germline cells. The dashed line indicates intermediate domains not shown here. The functional κ-chain gene, e.g., Vk8-Jk4-Ck, is formed in B cells through several rearrangements of the corresponding DNA domains. The combination shown here is only one of 500 possible variants. B. Structure of the immunoglobulin H-chain gene in germline cells. The dashed line indicates intermediate domains not shown here. The functional H-chain gene, e.g., V_H33-D_H26-J_H4-Ca, is formed in B cells through a series of rearrangements of the corresponding domains. The combination shown here is only one of 140,000 possible variants. The figure shows only one Cy domain, although there are actually four (Cy1, Cy2a, Cy2b, and Cy3).