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Review
The Wellcome Trust Sanger Institute, Hinxton, Cambs CB10 1SA, United Kingdom
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ABSTRACT |
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 TOP ABSTRACT 1) BACKGROUND2) GENE TARGETING3) MUTAGENESIS TECHNOLOGIES4) CONCLUSIONS AND FUTURE...References |
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gene targeting; homologous recombination; embryonic stem cell; mutagenesis; gene trap; mouse genome project
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1) BACKGROUND |
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TOPABSTRACT1) BACKGROUND2) GENE TARGETING3) MUTAGENESIS TECHNOLOGIES4) CONCLUSIONS AND FUTURE...References |
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In the last few years, a number of significant technological advances have dramatically increased our ability to create mouse models of human disease. These technological advances have been greatly aided by the sequencing of the mouse genome and the subsequent mouse genomic resources that have been developed.
1.2) The Mouse Genome Sequencing Project
The C57BL/6J strain has been selected as the reference strain for the production of a finished genome sequence (17). The mouse genome sequencing project has two main aims; the first is to generate a finished sequence of the mouse genome that is freely accessible to the public, and the second is to create a richly annotated information resource consisting of a database containing information about the underlying genomic sequence (reviewed in Ref. 129). Mouse genomic resources include mapping resources [such as dense genetic maps (51, 168), sequence tagged site (STS)-based physical maps (168), radiation hybrid (gene-based) maps (11, 88, 250), and simple sequence length polymorphism (SSLP) marker-anchored bacterial artificial chromosome (BAC) framework maps (33)]; DNA resources [such as the generation of expressed sequence tags (ESTs) (144), cDNAs (109, 233) and BAC libraries (175, 283)]; and database resources, which help to provide community access to this information (see below for URLs).
1.2.1) How is the mouse genome being sequenced?
The sequencing of the mouse genome involves two parts: a genome-wide program, and a targeted program (summarized in Ref. 129). The genome-wide sequencing program is being undertaken by the Mouse Genome Sequencing Consortium (MGSC), which is an international collaboration between four centers [the Wellcome Trust Sanger Institute; The Whitehead Institute/MIT Center for Genome Research; The Washington University Genome Sequencing Center; and Ensembl (a joint project between the Sanger Institute and the European Bioinformatics Institute)]. Incorporated into the genome-wide sequence will be the sequence of regions targeted by specific projects. These include the Trans-NIH BAC sequencing program (funding sequencing of specific BACs/regions of biological interest by sequencing facilities at Cold Spring Harbor Laboratory, University of Oklahoma, and the Albert Einstein College of Medicine), the UK-MRC Mouse Sequencing Programme (adopting a regional and functional approach to target four "core regions" on chromosomes 4, 13, 2, and X), and the Joint Genome Institute (performing comparative sequencing of mouse chromosomal regions syntenic with human chromosome 19). The annotation of the mouse genome will be facilitated by cDNA sequencing programs, including that carried out by the RIKEN Institute (who collect data on most, if not all, full-length cDNAs, their primary structures, and expression sites) and the Mammalian Gene Collection [MGC; who provide a complete set of full-length (open reading frame) sequences and cDNA clones of human and mouse genes (233)]. In addition, the MGSC will do clone-by-clone BAC sequencing to a high standard.
There are several components to the genome-wide sequencing effort. First, there is the development of a BAC map, consisting of overlapping BAC clones covering the vast majority of the mouse genome. For this, two BAC libraries have been selected: RPCI-23 (female) and RPCI-24 (male) (175). Although previously characterized by sequencing both ends of the cloned inserts ("BAC-end sequencing") (283), as part of the MGSC they are being characterized by restriction-digest fingerprint of each clone ("fingerprinting"), which can be used to assemble the BACs into overlapping clone "contigs" (continuous stretches of assembled sequences) covering the mouse genome. The BAC map and BAC end-sequences provide a crucial scaffold for assembling the sequence of the mouse genome. The second component uses paired-end whole-genome shotgun reads to generate light (
2.5-fold) and deeper (5- to 6-fold) coverage of the mouse genome; the sequence is obtained by preparing plasmid libraries (with inserts of 2–10 kb) and sequencing both ends of the inserts ("paired-end sequencing"). By combining the BAC map and the shotgun coverage, a "hybrid" genome assembly can be constructed, consisting of sequence contigs, linked into "supercontigs" (neighboring contigs that have been properly organized and orientated), which are further linked into "ultracontigs" (neighboring supercontigs that have been properly organized and orientated). In addition, comparative sequence information from other strains of mice, such as 129S1/SvImJ, BALB/cByJ, and C3H/HeJ will also be obtained, as such analysis is important for identifying sequence variants, such as single nucleotide polymorphisms (SNPs).
As of April 2002, two assemblies of the complete whole genome data set for the mouse genome have been generated: one by the Whitehead group using the ARACHNE program (18), and the other by the Sanger Institute using the PHUSION program. Both assemblies used the February 2002 freeze of data, and were done using the same set of files (each assembly included about 33 million reads, corresponding to x7 coverage of the genome). Although both assemblies were evenly matched by many criteria, the ARACHNE assembly was chosen by the MGSC as the one with which to proceed for further analysis. This assembly provides 96% coverage of the euchromatic mouse genome and predicts with high confidence 22,444 genes across the genome, of which 75% have a firm human genome counterpart. This "MGSC Version 3" assembly can be downloaded (ftp://ftp.ensembl.org/pub/assembly/mouse/mgsc_assembly_3) or searched using SSAHA (http://www.ensembl.org/Mus_musculus/ssahaview) or BLAST (http://www.ncbi.nlm.nih.gov/genome/seq/MmBlast.html). In addition, the BAC map resource has been aligned to the sequence, and an initial functional annotation of these genes has been added, as well as a comparison to the human genome (at the DNA level). The final phase of the MGSC’s work is to now fill in the "gaps" and correct any errors or misassemblies (for a completed genome sequence to be available within the next 3 yr).
All information generated by the MGSC is rapidly released to the scientific community; a constantly updated and comprehensive view of the project’s data is provided at a central MGSC server (http://mouse.ensembl.org/). In addition, MGSC data is incorporated into other key genome servers, including mouse genome resources at the National Center for Biotechnology Information (NCBI; http://www.ncbi.nlm.nih.gov/genome/guide/mouse/index.html), the MRC UK mouse sequencing program (http://mrcseq.har.mrc.ac.uk), and the mouse genomic sequence reads aligned against the human draft sequence at the University of California at Santa Cruz (http://genome.cse.ucsc.edu).
1.3) Basic Genetic Technologies for Mining the Mouse Genome
With the exponential increase in the number of genes identified by the genome sequencing projects, it has become imperative that efficient methods be developed for determining gene function. It has now become possible to make essentially any mutation in the germ line of mice by utilizing homologous recombination (the recombining of an exogenous piece of DNA with its endogenous homologous sequence in vivo) and embryonic stem (ES) cells (the pluripotent derivatives of the inner cell mass of the blastocyst) (34). Gene targeting (using homologous recombination to alter specific endogenous genes) provides the highest level of control over producing mutations. The combination of gene targeting techniques in mouse ES cells and the Cre-loxP recombination system has resulted in the emergence of chromosomal engineering technology in mice. This advance has opened up new opportunities for modeling new diseases that are associated with chromosomal rearrangements [in humans, chromosomal abnormalities are a principle cause of fetal loss and developmental disorders (214), and chromosomal translocations are involved in the genesis of many types of human tumor (189)]. Mutagenesis studies in the mouse have emphasized modeling human diseases through phenotype-driven assays. The chromosome engineering strategy for generating mice with precise chromosomal rearrangements, in combination with mutagenesis approaches, provides an invaluable tool for deciphering gene function using phenotype-based methods. In addition, gene trap approaches in ES cells also offer a route to gene discovery by providing information on gene sequence, expression, and mutant phenotype.
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2) GENE TARGETING |
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TOPABSTRACT1) BACKGROUND2) GENE TARGETING3) MUTAGENESIS TECHNOLOGIES4) CONCLUSIONS AND FUTURE...References |
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The procedure for the generation of mice that have been genetically modified using gene targeting strategies is essentially the same regardless of the specific targeting strategy used and is outlined in Fig. 1. Briefly, the targeting construct (containing sequence homologous to the targeted gene and a selectable marker) is electroporated into ES cells. These cells are then cultured in the presence of a selection agent to remove any cells that have not stably integrated the construct into their genome. The surviving ES cell colonies are then isolated and examined for the presence of the targeted allele (to ensure the desired recombination event has occurred) by PCR amplification or Southern blot analysis. Those containing the targeted allele are then injected into blastocysts (embryonic day 3.5) and transferred to the uterus of a foster mother. The resulting pups are then examined for their degree of chimerism (percentage of the genetic makeup of the mouse that was contributed by the ES cell). Male mice showing a high percentage of chimerism are then mated with wild-type mice to check for germ-line transmission of the targeted allele in the F1 offspring. The F1 heterozygotes may then be intercrossed to breed to homozygosity.
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locus (202), and the cystic fibrosis transmembrane conductance regulator (Cftr) locus (49) (with germ-line transmission of the mutated allele being demonstrated in the latter two examples). Although hit-and-run targeting procedures are typically performed in ES cells in vitro, with the modified ES cells subsequently being injected into blastocysts to generate mice carrying the mutation, one group has modified this technique to generate mice carrying a subtle mutation in the K-ras gene, whereby the targeted insertion allele was created in cultured ES cells, and the excision was performed in mice in vivo (95).
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2 Na,K-ATPase gene (7), and the Huntington’s disease homolog (Hdh) gene (36) (with germ-line transmission of the mutated allele demonstrated in the latter example). A modified approach, termed "stable tag exchange," incorporates an additional positive selection cassette to circumvent the problem of high levels of nonhomologous recombinants, which survive selection due to loss of tag gene expression (266). The advantage of this technique is that any number of different vectors can be used in the second step to generate an array of mutations at the same locus, as has been used in ES cells to generate a series of different targeted alterations to the prion protein gene (153). However, as with the hit-and-run method, a limitation of this procedure is the spontaneous loss of sensitivity to the negative selection agent due to events other than homologous recombination, such as mutation of the negative selectable marker (82).
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2.1.3) Generating conditional mice.
Conditional activation ("gain-of-function") or inactivation ("loss-of-function") of gene expression in vivo can be used to control gene expression in a temporal-, spatial-, or tissue-specific manner. The tight regulation of gene expression offered by conditional gene targeting means avoidance of any potential embryonic lethality associated with the mutation and allows for a more accurate mouse model of human disease and sporadic cancer initiation and progression (reviewed in Ref. 97).
Such control can be achieved using binary transgenic systems, in which gene expression is regulated by the interaction of an effector protein product on a target transgene. These interactions can be controlled by crossing mouse lines or by adding/removing an exogenous inducer. Binary transgenic systems fall into two categories: transcriptional transactivation (used to activate transgenes in gain-of-function experiments) and site-specific DNA recombination (used to activate transgenes or to generate tissue-specific gene knockouts). Transcriptional transactivation is more widely used than DNA recombination to bring about transgene activation in mice, because the latter is irreversible, whereas the former occurs only when the transactivator is present (thus if an inducible system is used, then transgene activation can be reversed at a given time, providing greater control over transgene expression) (reviewed in Ref. 123).
2.1.3.1) transcriptional transactivation. The most widely used binary transcription transactivation systems are the tetracycline-dependent regulatory systems ("Tet-on" and "Tet-off"; see http://www.clontech.com/tet/index.shtml or http://www.zmbh.uni-heidelberg.de/bujard/Homepage.html) (70). These systems use a chimeric transactivator to control transcription of the gene of interest from a silent promoter and are based on two regulatory elements derived from the E. coli tetracycline-resistance operon: the Tet repressor protein (TetR) and the Tet operator sequence (tetO). The Tet-off system uses a plasmid that expresses a fusion protein known as the tetracycline-controlled transactivator (tTA), composed of TetR and the VP16 activation domain of herpes simplex virus (Fig. 5A). tTA binds to the tetracycline response element (TRE) and activates transcription of the target gene in the absence of the inducer, doxycyclin (Dox).2 The Tet-on system uses a form of TetR containing four amino acid changes that result in altered binding characteristics and create the reverse TetR (rTetR). As shown in Fig. 5B, rTetR binds the TRE and activates transcription of the target gene in the presence of Dox. Recent examples of use of the Tet system in transgenic mice include the regulation of delta-FosB expression to control bone formation and bone mass (221), regulation of the SV40 T-antigen oncogene to control ß-cell autoimmunity and hyperplasia (21), and regulation of the endothelin receptor B (ednrb) to determine when ednrb signaling is required during embryogenesis (218). In addition, there are many variations on the basic Tet system, such as allowing two target transgenes to be coregulated (14), thereby enabling monitoring of transactivation in transgenic mice when one promoter drives a reporter gene [such as luciferase or ß-galactosidase (ß-gal)] (120). Other variations include modification of the DNA-binding specificity of the transactivator (15) and the generation of rtTA variants with increased Dox sensitivity, allowing a greater Dox responsiveness in mouse tissues to which Dox has limited access (such as the brain) (246).
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2.1.3.2) site-specific dna recombination. The most widely used site-specific DNA recombination system uses the Cre recombinase from bacteriophage P1, and more recently the Flp recombinase from S. cerevisiae has begun to show utility (discussed in more detail in Section 2.2). By using gene targeting techniques to produce mice with modified endogenous genes that can be acted on by Cre or Flp recombinases expressed under the control of tissue-specific promoters, site-specific recombination can be used to inactivate endogenous genes in a spatially controlled manner. Cre/Flp activity can also be controlled temporally by 1) delivering cre/FLP-encoding transgenes in viral vectors, 2) administering exogenous steroids to mice that carry a chimeric transgene consisting of the cre gene fused to a mutated ligand-binding domain, 3) using transcriptional transactivation to control cre/FLP expression, and 4) using soluble Cre. There are numerous examples of conditional mice that have been generated using the loxP/Cre recombinase system (19b).
2.1.4) Generating "knock-in" mice.
Knock-in experiments are used to place a transgene, such as a cDNA or a reporter construct contained in a targeting vector, under the transcriptional control of an endogenous gene. For example, the endogenous gene may be replaced with a homolog (to assess whether members of the same gene family have similar functions when expressed in the same spatial/temporal pattern); human cyclin E has been knocked into the murine cyclin D1 gene (which rescues all phenotypic manifestations of cyclin D1 deficiency and restores normal development in cyclin D1-dependent tissues) (67), and murine Otx1 has been knocked into the murine Otx2 locus (to demonstrate the functional equivalency between Otx2 and Otx1 in development of the rostral head) (236). The most widely used knock-in strategy is the replacement of a gene by a reporter gene such as LacZ or GFP, to monitor expression pattern of the gene during development and in the adult mouse, both in a spatial and temporal manner. For example, LacZ has been knocked into the Pax3 locus (to examine the role of Pax3 in the neuroepithelium and somites) (143). A knock-in allele usually results in the loss-of-function of the endogenous gene.
Although the early knock-in studies in mouse ES cells utilized the double replacement technique (Section 2.1.2) (69, 228), more recent knock-in experiments have been performed using conventional gene targeting approaches (Fig. 6A), often in combination with the Cre-loxP system to remove the selection cassette (detailed in Section 2.2; Fig. 6B) (76). As shown in Fig. 6, such knock-in vectors are essentially replacement targeting vectors containing the transgene and a positive selectable marker and are designed such that after homologous recombination, the transgene is transcriptionally regulated by the endogenous promoter of the locus. This method is based on the production of a fusion protein between the endogenous and knocked in products. However, if a fusion product is undesirable (such as in the case of secreted proteins), then 1) the transgene can be knocked into the untranslated region upstream of the endogenous translational start site (although splicing may occur around the inserted transgene) or 2) an internal ribosome entry site (IRES) element (181) can be used to generate a bicistronic mRNA [the encephalomyocarditis virus IRES is most commonly used in mammalian cells (110)] (113).
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Genetic mosaics can be generated when mitotic recombination between homologous chromosomes occurs during the G2 phase of the cell cycle and the recombinant chromatids segregate to different daughter cells (X segregation). Recombinant chromatids produced in G2 can also segregate to the same daughter cell, and the nonrecombinant chromatids to the other daughter cell (Z segregation). It is X segregation that is useful for genetic mosaic analysis, because it produces clones of homozygous mutant daughter cells from heterozygous mothers (whereas Z segregation produces daughter cells that are phenotypically indistinguishable from the parent cell or from cells produced by G1 recombination).
There are numerous approaches to generate mosaics in mice (see Fig. 7). The most common is the use of site-specific Cre/loxP technology (described in Section 2.2) in a conditional knockout strategy, in which mosaics are generated in specific tissues by the activity of Cre on a loxP-flanked allele (Fig. 7A) (74). Alternatively, homozygous mutant ES cells can be produced in culture, and these can be used to generate chimeras. For example, direct selection of loss of heterozygosity (LOH) events in neo cassette-containing ES cells can be achieved using high concentrations of G418 (which causes chromosomal nondisjunction followed by reduplication of the remaining chromosome; Fig. 7B) (154). The mechanism that duplicates the targeted neo cassette (thereby providing enhanced G418 resistance) also allows mutations on the same chromosome to become homozygous. Homozygous clones can also be generated by repetitive neo targeting, followed by puro targeting of the second allele. Another strategy (Fig. 7C), based on the principles of the Flp-FRT site-specific recombination system in Drosophila (22), uses the spatial and temporal regulation of Cre expression, which can be used to trigger mitotic recombination (with recombination rates of 10-4–10-2 in ES cells transiently transfected with Cre) (134). Alternatively, Bloom’s mice can be used, as they possess a genetic background that displays increased rates of mitotic recombination (Fig. 7D). Bloom’s mice (Blm-/-) are deficient in the RecQ DNA helicase/Blm protein (140), analogous to the human condition, Bloom’s syndrome [a rare cancer-prone disorder in which the cells of affected persons have a high frequency of somatic mutation and genomic instability due to high frequency of sister chromatid exchange (68)]. Bloom’s mice and ES cells exhibit greatly elevated rates of mitotic recombination, allowing homozygous mutant daughter cells to be generated from heterozygous mothers both in vitro and in vivo. The rate of mitotic recombination in Blm-deficient cells is sufficient to enable homozygous mutant clones to be recovered for any gene, regardless of its location in the genome (140).
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-integrase superfamily) include the Cre-loxP system from the bacteriophage P1 and the Flp-FRT system from the budding yeast S. cerevisiae. Both Cre and Flp cleave DNA at a distinct target sequence (Fig. 8A) and ligate it to the cleaved DNA of a second identical site, to generate a contiguous strand. The orientation of these target sites relative to each other directs the type of modification catalyzed by the recombinase (detailed in Fig. 8B). When compared with one another, Cre appears more effective in recombining a transgene array in vitro and in vivo than Flp (198). However, mutated versions of Flp, such as the temperature-sensitive FlpL (30) and the enhanced Flp (Flpe) (31), show improved activity over wild-type Flp. Indeed, the generation of high-efficiency Flpe deleter mice [the ROSA26 locus was targeted to create a mouse strain with generalized expression of FLPe, termed the FLPeR ("flipper") strain (56)] show that Flpe is an alternative to Cre-loxP (199). In general, Cre and Flpe are used in situations requiring high-efficiency recombination (30–31, 56, 245); Flp and FlpL are used when tight regulation is needed (255). Currently, the most widely used site-specific DNA recombinase system in both ES cells and mice is the Cre-loxP system.
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2.2.3) Systems for delivery of Cre in vitro and in vivo.
The methods used to introduce Cre into ES cells are numerous. The usual in vitro introduction method for transient expression or stable integration is calcium precipitation or electroporation [which can reach as high as 70% efficiency (163)]. Alternative methods of delivery include infection of ES cells with a recombinant Cre adenovirus (2, 5, 259). Adenoviral transfection of Cre results in transient Cre expression, but mosaic mice can be produced, which transmit the targeted allele to their offspring with high frequency (106). The method of introduction of Cre in vivo depends heavily upon the specific application. The simplest solution to achieve overall excision in the developing embryo is crossing of the floxed mice with transgenic mice carrying the Cre recombinase gene (a general Cre expresser transgenic line) (119, 206). Another variation of this approach is pronuclear injection of a Cre expression vector [with the resultant transient Cre expression inducing recombination during preimplantation development (8)] or injecting Cre RNA or protein.
2.2.4) Temporal- and tissue-specific expression of Cre.
As discussed above (Section 2.1.3), the Cre recombinase system can be used to conditionally activate ("gain-of-function") or inactivate ("loss-of-function") gene expression in a temporal-, spatial-, or tissue-specific manner, to allow for a more accurate mouse model of human disease and cancer initiation and progression. To create spatiotemporally controlled somatic mutations in the mouse, chemically inducible forms of Cre have also been used, such as Cre-ERT and Cre-ERT2, in which the ligand-binding domain of a mutated human estrogen receptor (ERT), which recognizes tamoxifen or its derivative 4-hydroxytamoxifen (4-OHT), has been added to Cre (29, 59, 65, 90). Accordingly, Cre-mediated recombination is 4-OHT dependent in mice bearing a CreT transgene. This approach has been used to selectively ablate expression of the retinoid X receptor- (RXR) in adult mouse keratinocytes, by putting expression of these recombinases under control of the bovine keratin-5 promoter (126). A mouse strain expressing Cre-ERT from the ubiquitously expressed ROSA26 locus has also been generated (256).
The main methodology for tissue-specific Cre-mediated excision is the use of established transgenic lines expressing Cre under the control of a promoter with the required specificity. For example, to create a mouse model for BRCA2-associated breast cancer [as inheritance of one defective BRCA2 allele predisposes humans to breast cancer (reviewed in Ref. 213)], mice conditional for the tumor suppressor genes (TSGs) Brca2 (floxed at exon 11) and/or p53 (floxed at exons 2–10) were mated with mice expressing Cre under control of the epithelial-specific K14 promoter; although no tumors arose in mice carrying conditional Brca2 alleles alone, mammary and skin tumors developed in females carrying conditional Brca2 and p53 alleles, showing that inactivation of both Brca2 and p53 combine to mediate mammary tumorigenesis (96). There are numerous more examples of the use of Cre-expressing mice (19a). In addition, a voluntary database of Cre-expressing mice has been established (see http://www.mshri.on.ca/nagy).
However, a stumbling block is simply the limited number of existing transgenic mouse lines that express recombinase in the appropriate cell type. To address this issue, recombinant Cre fusion proteins bearing hydrophobic peptides from the Kaposi fibroblast growth factor (FGF-4) (94) or basic peptides derived from HIV-TAT (98, 180) have been produced to promote cellular uptake of recombinant Cre. Recombination has been observed in a variety of cultured cell types and in specific tissues examined in mice following intraperitoneal administration. This new cell-permeable form of Cre will likely open up new opportunities for genetically manipulating cells both in vitro and in vivo (40).
In certain applications, the retroviral or adenoviral delivery systems have been used. For example, after intranasally administered adenoviral Cre, mice expressing a mutated K-ras oncogene gene placed downstream of a floxed transcriptional termination stop element developed lung tumors by 2–16 wk of age (due to Cre-mediated removal of the floxed stop element, and hence expression of the mutant K-ras) (95). An alternative is the combination of avian retrovirus with the TVA (EnvA receptor) delivery system, which provides the possibility of viral delivery of Cre in a tissue- or cell-specific manner, as there is a requirement for the expression of a special avian receptor (tv-a) in the mouse cells from a cell-type-specific transgene to make these cells susceptible to infection (85). More recently, an adeno-associated virus [used to transfer foreign genes into the adult and neonatal central nervous system in animals (23, 136, 146)] expressing Cre recombinase has been shown to mediate extensive in vivo recombination in neural cells of defined brain regions in the mouse (108).
2.3) Chromosome Engineering
2.3.1) Chromosomal rearrangements.
A large array of mice containing chromosomal rearrangements (deletions, inversion, duplications, and translocations) have been generated by exposure to chemical mutagens [cyclophosphamide, ethylene oxide, chlorambucil, and N-ethyl-N-nitrosourea (ENU)] or radiation (X-rays) (47, 124, 192, 205, 234, 237). However, although these mutagens have generated some valuable mouse models for human diseases, such as the mouse model of trisomy 21 (47, 192), their usefulness for inducing rearrangements is limited by the fact that the endpoints of the induced rearrangements cannot be predetermined (see SECTION 3, below, for details on mutagenesis strategies). To this end, gene targeting-based strategies have been developed to introduce defined chromosomal rearrangements into the mouse genome by engineering them in ES cells using the Cre/loxP site-specific recombination system. This technology, known as "chromosome engineering," has successfully generated numerous mouse models that accurately recapitulate human chromosomal rearrangements, such as the chromosomal deletion within band 22q11 (del22q11) causing DiGeorge syndrome (130–131), the paternal deficiency of chromosome 15q11-q13 causing Prader-Willi syndrome (244), the translocation between chromosomes 8 and 21 [t(8;21)] found in acute myeloid leukemia (AML) (32), and the reciprocal translocation between chromosomes 9 and 11 [t(9;11)(p22;q23)] associated with acute leukemia (42).
2.3.2) Chromosome engineering technology.
2.3.2.1) selection of endpoints.
Generating chromosomal rearrangements in the mouse involves the sequential insertion of two targeting vectors into two separate loci in the ES cell genome. Thus an important decision is to define the two endpoints of the rearrangement. Endpoints can be chosen at any genomic region. For example, SSLP microsatellite markers make useful endpoints as they have been genetically mapped (see the Whitehead Institute STS Physical Map of the Mouse at http://www-genome.wi.mit.edu/cgi-bin/mouse/index). Numerous SSLP markers have been successfully used as the endpoints for engineering chromosomal deletions on mouse chromosome 11 (286). In addition, the high-resolution mapping information available for the mouse genome (see the MGSC Ensembl Mouse Genome Server at http://www.ensembl.org/Mus_musculus/) means that genes of known chromosomal location may also be used as endpoints. Genes used as endpoints have included the epidermal growth factor receptor (Egfr) gene (242), the amyloid precursor protein gene (125), the myeloperoxidase (MPO) gene (10), Wnt3, p53, and Hoxb9 (190, 286), p63 (149), and the HoxB cluster (148), to name a few.
2.3.2.2) generating the chromosomal rearrangement. Once the endpoints have been selected, the first step involves the targeting of an insertion vector containing a loxP site, a positive selection cassette (e.g., neomycin), and one of two complementary, but nonfunctional, fragments of the Hprt gene into the desired locus (selected endpoint) of the ES cell genome (see Fig. 9A) (285). ES cell clones with a loxP site targeted to a first endpoint can be identified by positive selection and Southern blot analysis. The second step involves the targeting of a second insertion vector containing a loxP site, a different positive selection cassette (e.g., puromycin), and the complementary fragment of the Hprt gene into the second endpoint (see Fig. 9B). The generation of these targeting vectors is detailed below. The doubly targeted ES cell is then transiently transfected (electroporated) with a vector expressing Cre, which facilitates recombination between loxP sites such that the intervening DNA is deleted. Commonly used Cre-expression vectors include pOG231 (171), pTurboCre (GenBank accession no. AF334827), pCrePAC (238), and pBS185 (125). The type of chromosomal rearrangement derived from the double-targeted ES cells is determined by the loxP configuration (reviewed in Fig. 8B). As shown in Fig. 10A, loxP sites in the same orientation generate a chromosomal deletion (or duplication event; not shown). Alternatively, loxP sites in the opposite orientation generate a chromosomal inversion (Fig. 10B). The generation of chromosomal translocations is discussed below (in Section 2.3.4).
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2.3.2.3) generation of gene targeting vectors for chromosome engineering. Gene targeting vectors, such as those shown in Fig. 9, can be generated in the conventional way by sequentially inserting various genetic components into a plasmid construct (133, 190). However, the "two library" system (285) greatly reduces the number of cloning steps required for generating gene targeting vectors. This system is composed of two complementary libraries of pre-made gene targeting insertion vectors. The 5'-Hprt library was generated by cloning a genomic library into a vector backbone that contained the 5'-Hprt cassette, a loxP site, and neomycin (PGKneobpA) as a positive selectable marker. The complementary 3'-Hprt library was generated by cloning a genomic library into a vector backbone that contained the 3'-Hprt cassette, a loxP site, and puromycin (PGKpurobpA) as a positive selectable marker. In addition, both of these libraries are equipped with genes encoding visible coat color markers. The tyrosinase minigene (Ty) has been used to "tag" transgenes with a visible pigment marker in albino mice (176, 275), and the K14-Agouti transgene (Ag) uses the keratin-14 promoter to constitutively express agouti (114). Mice carrying the Ty transgene have a grayish coat on an otherwise albino background, and the Ag confers a "butterscotch" coat color in black agouti or non-agouti mice (285).
To make a defined rearrangement between any two desired loci, clones for each endpoint are first isolated from the appropriate library. Linearization of the construct within the genomic insert generates a gene targeting insertion vector that integrates at the target locus. Targeting can be assessed by Southern blot analysis using a DNA fragment that was removed from the genomic insert before gene targeting, by an external probe, or by PCR amplification using primers specific for the gap and vector. An additional feature of this gene targeting library system is that clones isolated from these libraries can also be used for analyzing single gene function via the knockout (null allele) approach, as demonstrated by targeted disruption of the p63 locus (149).
2.3.3) Chromosomal deletions.
2.3.3.1) uses for the engineering of chromosomal deletions.
The chromosomal engineering of deletions can be used to identify TSGs without prior knowledge of the gene function. For example, mice possessing a deletion encompassing a putative TSG should exhibit increased tumorigenesis, as they only possess a single copy of a TSG, and tumor-specific loss or inactivation of the remaining allele can be used to clone the causative gene. Chromosome engineering can also be used to generate mouse models of human microdeletion syndromes. For example, mice heterozygous for a 1.2-Mb deletion between Es2 and Ufd1l on mouse chromosome 16 show cardiovascular abnormalities resembling those found in DiGeorge syndrome patients (130). When these mice were crossed to a strain harboring a duplication of the same region, the mutant phenotype was functionally rescued, indicating that the defects seen in DiGeorge patients are due to haploinsufficiency (loss of one functional copy of a gene). Thus the potential embryonic lethality of a particular deletion can be functionally determined by assessing the ability of the duplicated allele to rescue the embryonic lethality of the deletion (as mice containing both the deleted and duplicated alleles are genetically balanced).
The major factor limiting the generation of deletions in ES cells is the size of the rearranged interval, with deletions >22 cM leading to ES cell lethality or severe growth disadvantage of the cells in culture (286). Although Cre/loxP recombination has been shown to occur over such large distances, clones can emerge from these experiments that have undergone a compensatory genetic change, such as a chromosomal duplication (286). X-ray- and UV-induced mutagenesis have also been successfully performed to generate deletion complexes in ES cells. Radiation-induced deletions can be localized and made selectable by targeting a vector that carries a negative selection cassette to a predetermined locus; for example, homologous recombination can be used to insert a herpes simplex virus thymidine kinase (HSV-tk) gene into a specific locus, followed by irradiation and then selection for clones that have deleted the tk cassette (240, 276). In addition, deletion alleles in ES cells of several centimorgans have been successfully transmitted through the germ line (190, 276).
2.3.3.2) generation of nested chromosomal deletions. An extension of the chromosome engineering strategy is the generation of nested chromosomal deletions, a series of variably sized, overlapping deletions surrounding a predetermined genomic locus. This strategy has been used to identify that haploinsufficiency of the Tbx1 gene contained within the 1.2-Mb deletion interval of DiGeorge mice was responsible for the aortic arch defects seen in these patients (131). Furthermore, if the genomic locations of the endpoints are known, then nested deletions can be extremely useful for mapping novel recessive mutations (278). However, to get around the task of having to generate targeting vectors for the nested endpoints, retroviral integration of a second loxP site and selection cassette can be used (235). More specifically, deletion complexes can be anchored to a predetermined location in the genome by targeting the 5'-Hprt-loxP cassette. The 3'-Hprt-loxP cassette is then randomly inserted into the ES cell genome by retrovirus-mediated integration, generating a library of ES cell clones with the same targeted endpoint and a collection of random endpoints (Fig. 11). This method has been used to generate nested deletions extending from a few kilobases to several megabases at the Hprt locus, also on chromosome 11 (235). Electroporation has also been used to randomly insert the second loxP site and selection cassette into the ES cell genome (122). However, insertion by electroporation increases the risk of genomic rearrangements occurring at the insertion site, and tandem repeats of a vector may be introduced into the insertion site (although these should be reduced to a single locus by the activity of Cre on a head-to-tail concentrate) (278). Endpoints generated by random insertion can be defined by cloning the genomic DNA that flanks the deletion endpoints and by mapping these junction fragments onto a physical map of the region. Nested deletions can also be efficiently generated by irradiation (116, 240, 276), although they require additional extensive characterization to define each deletion interval (278).
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Using the Cre-loxP system, chromosomal translocations are generated using the same techniques as for deletions, duplications and inversions (see Section 2.3.2), except the loxP sites, orientated in the same direction relative to their respective centromeres, are targeted to nonhomologous (different) chromosomes. If the loxP sites are in opposite directions, then recombination will result in acentric chromosomes (without a centromere) and dicentric chromosomes (containing two centromeres), which will be unable to survive. Thus, to prevent the generation of acentric and dicentric chromosomes, only pairs of genes with the same transcriptional orientations relative to their centromeres can be engineered to produce fusion proteins (278). To generate a fusion protein from a chromosomal translocation, the targeting vectors need to be designed to allow the two genes to be linked through their introns (with the loxP site embedded in the breakpoint), so RNA splicing will generate an in-frame fusion mRNA and protein (278). To induce the translocation, mice carrying both targeted endpoints can be crossed with transgenic mice expressing a tissue- and/or temporal-specific Cre. In addition, this strategy circumvents the problem of transmitting the translocation through the male germ line, as the presence of chromosomal translocations in male germ cells can cause infertility (141).
2.3.5) Balancer chromosomes.
A balancer chromosome is one containing an inverted region(s). Balancer chromosomes can be generated using chromosomal engineering technology, as chromosomal inversions can be generated in mouse ES cells by successive gene targeting of a loxP site to the two endpoints, followed by Cre-mediated recombination between the two loxP sites (Fig. 10B) (284). In addition, the inversion can be marked with a dominant marker (such as the dominant K14-Agouti coat-color gene) so that progeny carrying the balancer can be readily identified.
As inversions suppress crossing over during mitotic recombination (the genetic exchange of information between sister chromatids), balancer chromosomes are genetic reagents that can be used for stock maintenance (to maintain the integrity of mutagenized chromosomes). Balancer chromosomes can also be used for large-scale mutagenesis screens (see Section 3.1.2). For example, in intercrosses between siblings that have inherited the balancer chromosome and a mutagenized chromosome, absence of non-balancer-carrying progeny indicates the presence of one or more recessive lethal mutations on the mutagenized chromosome (284, 278). Indeed, the first mouse balancer chromosome was constructed on chromosome 11, to facilitate the isolation of ENU-induced recessive mutations on mouse chromosome 11 (http://www.mouse-genome.bcm.tmc.edu/ENU/ENUHome.asp) (284). This balancer chromosome is based on a 24-cM inversion between the Trp53 gene and the Wnt3 gene (in addition, a coat color marker, K14-Agouti, has been inserted into the mutated Wnt3 locus). Balancers will also be useful to facilitate analysis and maintenance of other types of knockouts.
2.3.6) Homologous recombination in E. coli.
To simplify the generation of knockout constructs, recombineering technologies have been developed. This form of DNA engineering utilizes methods based on homologous recombination in E. coli that enable large segments of genomic DNA in BACs or P1 artificial chromosomes (PACs) to be modified and subcloned, without the need for restriction enzymes or DNA ligase.
2.3.6.1) reca homologous recombination. Several recombination pathways have been identified in E. coli, including the RecA pathway. A homologous recombination-based system allowing modification of BACs in recombination-deficient E. coli is the temperature-sensitive shuttle-vector-based system (75, 170). This temperature-sensitive plasmid replicates in cells growing at the permissive temperature (30°C) but is lost in cells growing at the restrictive temperature (42–44°C) because its origin of replication cannot function (79). Introduction of the E. coli RecA gene into the temperature-sensitive shuttle vector allows the RecA- host E. coli strain containing the BAC to become competent to perform homologous recombination of the resident BAC in vivo (150, 272). Using this method, transgenic mice have been generated by pronuclear injection of the modified BAC, and germ-line transmission of the intact BAC obtained (272).
2.3.6.2) et recombination.
Another recombination pathway in E. coli is the RecBCD pathway involved in repairing double-strand breaks (225). RecBCD unwinds and degrades DNA to generate 3' single-stranded DNA (ssDNA) tails, which are used by RecA to initiate recombination. However, E. coli possessing wild-type RecBCD degrade the introduced linear DNA molecules before recombination has proceeded; therefore, to restore recombination activity, a suppressor mutation that activates expression of a nuclease that produces 3' overhangs may be used (117). RecBCD can be inactivated by 1) the sbcA mutation, which removes a repressor for the endogenous lamboid prophage, Rac, which in turn induces the expression of recE and recT [two Rac genes that encode homologous recombination functions (135)], or 2) the gam protein of phage [in the presence of gam, phage-encoded recombination function stimulates homologous recombination by the red genes (156, 188)].
Engineering DNA by homologous recombination mechanisms involving the RecE/RecT and Red/Redß proteins in E. coli is termed "ET recombination" (also known as Red recombination and lambda-mediated cloning). ET recombination was pioneered by Stewart and colleagues (280), who showed that a PCR-amplified fragment of linear dsDNA, flanked by short regions of homology (42 bp) to a plasmid or BAC can be efficiently targeted to a plasmid or BAC by electroporating the dsDNA into recBC sbcA strains (see http://www-db.embl-heidelberg.de/jss/servlet/de.embl.bk.wwwTools.GroupLeftEMBL/ExternalInfo/stewart/ETcloning-textonly.html) (280). As shown in Fig. 12, ET recombination involves two steps: first, the amplification of a fragment of DNA by PCR with flanking regions of homology, plus the introduction of phage recombination functions into a BAC-containing bacterial strain; and second, the transformation of the cassette into the bacterial cells that contain a BAC and recombinase functions (the bacterial cells generate a recombinant in vivo, and detection of the recombinant is done by selection, counter-selection, or by direct screening). ET recombination allows alterations such as point mutations, sequence insertions and/or deletions to be carried out at any position on a target DNA molecule and has greatly simplified the generation of transgenic and knockout constructs (reviewed in Refs. 43, 160).
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