Since its introduction by Streisinger, the zebrafish has become a popular genetic model for studying development and disease because of its many advantages (Jagadeeswaran et al., 2005; Streisinger et al., 1981). One advantage is that zebrafish embryos are transparent, so a developing embryo and its morphology could be easily observed. Breeding zebrafish under laboratory conditions is extremely easy; the embryos grow for 72 hours until they hatch. Other advantages include high fecundity (the female fish can lay up to 200 eggs), short generation times, ease of in vitro fertilization, and amenability for large-scale mutagenesis. In addition, features such as transparency of embryos and generation of diploids from haploid eggs make the model even more attractive for identifying recessive mutations. Described below are the principles of genetics in zebrafish and how mutants are isolated and mapped to identify the genes that are affected. Mutagenesis is performed by dipping zebrafish males in ethylnitrosourea (ENU) to cause multiple, random point mutations throughout the genome of the germ cells (spermatogonia). Mating of an ENU-treated male with a wild-type female zebrafish will yield the F1 progeny that will carry as many as 1000 random mutations. Thus, saturating the genome with mutations is possible because theoretically, in 100 fish, 105 mutations can be generated. The fish from the F1 generation is then bred to a wild-type fish to generate the heterozygous F2 progeny. Homo-zygous fish are generated by performing brother-sister matings of the F2 generation. A screen on this F3 progeny for mutant phenotypes constitutes a classical three-generation screen. More than 1000 developmental mutants have been isolated using this strategy. Recently, several imaginative screens have been performed to isolate mutants related to thrombosis, lipid absorption, bone defects, and so forth. Adult screens are more difficult when compared to embryonic and larval screens, because rearing the F3 generation fish to adulthood requires more time and space. In identifying embryonic mutant pheno-types, many laboratories have also used two-generation screens as an alternative method. In this method, haploid or homozygous mutant progeny can be generated directly from the F1 females. Haploid embryos are generated by the in vitro fertilization of eggs obtained from a heterozygous F1 female with male sperm whose DNA was destroyed by treatment with ultraviolet light. Although these sperm do not contribute DNA, they can still initiate fertilization and allow the development of haploid embryos, which are viable for up to 3 days postfertilization. If these eggs are subjected to early high-pressure treatment (EP) immediately after fertilization, these haploids will become diploid embryos, which are fully viable. In the EP treatment, hydrostatic pressure causes disruption of spindle formation, which is required for the separation of sister chromatids and polar body extrusion following fertilization. The resulting gynogene-tic diploid larvae possess only maternally derived genes that are homozygous for most loci (heterozygosity may arise from recombination events in meiosis I). Thus, this EP treatment allows for a two-generation mutagenesis screen that will identify mutant-carrier females one generation ahead of classical approaches. Once a mutant of interest is identified, the mutant loci can be identified by a positional cloning strategy as described below.
An F1 female mutant carrier is crossed with a male zebrafish that is polymorphic for many loci when compared with the strain used to generate mutations. From this progeny, homozygous mutants are generated by brother-sister mating. To map the mutant locus, two pools of genomic DNA are created from normal and mutant zebrafish (20 zebrafish per genomic DNA preparation) in the above progeny of brother-sister mating. Multiple sets of primers are then used to amplify known microsatellite markers that span the zebrafish genome at an average resolution of 10 cM. Due to the fact that recombination will shuffle both genes and markers in meiosis, markers not linked to the mutant loci, or distal to the loci, will be present in both normal and mutant pools, while linked markers will be present in only one of the two pools. Once a marker is identified showing linkage to a mutant gene, further analysis using additional flanking markers and a larger number of genomes will establish close linkage. Bacterial artificial chromosomes that contain the linked markers could then be sequenced to identify the mutant gene. However, with the availability of genome information, the painstaking efforts of positional cloning will be replaced by the candidate gene approach once a closer linkage is identified. By rescuing the mutant through introducing the functional cDNA or by inhibiting the gene function by antisense approaches, the mutant gene could be confirmed.
In addition to the above approach, there has recently been a major success in the retroviral insertion mutagen-esis approach, which identified several hundred genes involved in development (Amsterdam et al., 2004). It has been shown that mouse retroviral vectors pseudotyped with a VSV-G envelope infect the fish germ line after injecting the virus into blastula-stage embryos at the 1000-2000-cell stage. Retroviruses are attractive candidates for insertional mutagenesis, because they had been shown to integrate into many different sites in mammalian and avian chromosomes. Importantly, they integrate without rearrangement of their own sequences or significant alterations to host DNA sequences at the site of insertion—essential features for easily cloning genes disrupted by insertions. Once the infectious retrovirus is injected into the zebrafish embryos and the founder lines are generated, they are mated pairwise and the resulting F1 fish are tested for multiple insertions of the retroviral genome. Those with multiple insertions are then pair-mated and F2 families are raised. Again mating six pairs of siblings will generate homozygous insertional mutants, and one could score for the phenotypes similar to the screening done in ENU mutagenesis. However, the relatively low efficiency of insertional mutagenesis in contrast to the ENU mutagenesis limits the utility of this method for small laboratories. Also, this method will result in the inactivation of the gene rather than producing hyperactive and hypoactive alleles that are possible by the ENU mutagenesis.
Because of the above advantages of the zebrafish model, we tested its utility as a genetic model for studying aging by initially establishing the zebrafish lifespan (Gerhard et al., 2002; Herrera and Jagadeeswaran, 2002; Herrera and Jagadeeswaran, 2004). Another group independently performed such studies (Gerhard et al., 2002). Both of these studies revealed that zebrafish have a lifespan of approximately 4 years. Furthermore, Kishi and his colleagues have identified several potential aging biomarkers such as senescent associated /-galactosidase activity in skin and oxidized protein accumulation in muscle (Kishi, 2004). In contrast, markers such as accumulation of lipofuschin granules (which accumulate in muscle cells with advancing age) were not found. Another study revealed the decline of muscle function with advancing age by studying the bone curvature (Gerhard et al., 2002). Even though some differences exist in the expression of biomarkers when compared to mammals, zebrafish nevertheless showed gradual senescence as observed in other fish species. In spite of this information on biomarkers, classical genetic studies will take long periods of study since zebrafish have long lifespans. Therefore, we suggested that zebrafish do not qualify as an ideal model because of their long lifespan. However, Gerhard and Cheng have recently argued that studying more than one fish may be useful for comparative aging (Gerhard et al., 2004). While it is a good argument, the utility of such comparative studies in identifying longevity genes is questionable.
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