Specific aim #4:
To identify and characterize genes involved in axial extension, morphogenetic movements, axial patterning, and eye development via
A. chemical mutagenesis
B. insertional mutagenesis; and
C. To assess the feasibility of gene targeting in S. tropicalis by evaluating nuclear transfer from cultured somatic cells into enucleated eggs.
Rationale:
This proposal up to this point is already broad. While it is not unambitious to include mutagenesis, there are several reasons to conclude that the labor required for limited screens is outweighed by the likelihood of obtaining useful research tools and insights. First, mutagenisis by transgenesis (insertional mutagenesis) needs to be quantitated if only to optimize the transgenic procedure for generating stable lines, and offers a direct path to molecular characterization of mutant loci. Second, combining chemical mutagenesis with transgenesis- especially tissue-specific reporter lines- offers ways for us to focus our screens on the pathways that interest us, and may also make discerning subtly aberrant phenotypes easier. Third, the riskiest proposals, such as testing gene targeting in frogs, have very large potential payoffs relative to the investment of labor, since preliminary assays of feasibility can be conducted with available reagents (e.g. nuclear transfer from extant X. laevis euploid cell lines). Fourth, some of the same techniques that made early zebrafish screens feasible, including the ready production of gynogenetic haploids and diploids, likewise make it possible for us avoid carrying large numbers of animals for the full duration of the screen. Should the need arise, we have the facilities to maintain up to 20,000 S. tropicalis in a new facility recently constructed at the University of Virginia (see Grainger/Keller facilities).
Specific aim #4.A: Chemical Mutagenesis
Rationale:
Conventional chemical mutagenesis is likely to be more efficient and easier to titrate than insertional mutagenesis, with better prospects for generating a varied set of single-gene mutants for analysis. Molecular characterization of chemically-induced point mutants remains a stumbling block. However, our screens include design elements to help focus on particular developmental processes; we are not planning to evaluate the very large number of lines required for a saturation mutagenesis. Instead, we hope to use streamlined assays for genes expressed in recognizable patterns indicative of normal early development (see 'focused screens' below) to help isolate mutants in relevant pathways. Gene expression assays may also create a more readily-recognizable, patterned "search image" to help workers spot subtle aberrations in development, much as the regular patterns of larval denticle belts facilitated the identification of axis formation mutants in Drosophila.
Methods
Titration of mutagen:
It is useful to calibrate dosage prior to initiating large-scale mutagenesis: too low a mutagen dose and much effort will be wasted examining genotypically wild-type embryos, too high a dose may be toxic and analysis will be complicated by multigene phenotypes in unpredictable ratios. We will determine the specific locus mutation rate (the rate at which a given locus is mutated) at different doses of mutagen; multiplied by the number of genes with a visible embryonic phenotype, estimated to be 2400 in zebrafish30, this gives the average number of mutations per haploid genome.
Ideally, one would calculate this rate by mating a mutagenized animal to animals that were each homozygous for one of several different recessive marker alleles, to quantitate the mutagenic "hit" frequency at a variety of loci by counting the frequency with which the homozygous recessive phenotypes appear. Identification of viable S. tropicalis marker genes by gynogenetic screens of wild-caught individual animals is described in Specific Aim 1.C. Mutagenized wild-type sperm are then used to fertilize S. tropicalis eggs mutant for viable recessive marker loci, and the specific locus mutation rate will be calculated by observing the frequency of embryos displaying the homozygous marker phenotype, indicating a mutagenic "hit" at the marker locus in the paternal genome.
For chemical mutagenesis, N-nitroso-N-ethylurea (ENU) "isopacs" (Sigma) will be dissolved in water and diluted to 0.5, 1.5, 3, and 5mM in 'adult frog water' (0.1% rock salt, 1mM NaPO4). Two wild-type male S. tropicalis per dose will be immersed for two hours three times weekly for two weeks to mutagenize prespermatogonial stem cells using Environmental Health and Safety-approved protocols. After a one to two month sperm maturation period, mutagenized males will be mated to two pairs of tester females homozygous for different marker loci for calculation of the specific locus mutation rate as described above. Female S. tropicalis typically lay 1,000-3,000 eggs, so approximately 8,000 to 24,000 haploid genomes per mutagen dose will be screened. If we have not identified useful S. tropicalis mutants prior to initiating the chemical mutagenesis, mutagen dosage will be optimized first on X. laevis, in which several mutations are available74, then normalized to the lower body mass of S. tropicalis.
One- and two-generation Screens
Following determination of the specific locus mutation rate at a range of ENU doses, an optimal dose for delivering one mutagenic 'hit' causing a visible embryonic phenotype per haploid sperm genome will be delivered to five males by immersion as above (F0) as above, and following a sperm maturation period, the males will allowed to mate with five hormonally-induced females. Our plan is to initially process the progeny of one mutagenized male at a time in our screen, and total four males in an eight-month screen; we may adjust our throughput as we go along.
From the eggs of five females crossed to mutagenized F0 males, we plan to raise 1000 F1 frogs to maturity, of which half are expected to be female. The 500 F1 "candidate heterozygous" females will be individually marked with a coded brand, and then simultaneously outbred and assayed by gynogenesis at a rate of 25 females per day. ~300 eggs per female will be fertilized with UV-irradiated sperm to make F2 haploids, and with unirradiated sperm to generate F2 outbred progeny. Half of the F2 haploids will be subjected to late pressure treatment to inhibit first cleavage and restore diploidy (F2 gynogenetic diploids). Haploids and homozygous diploids will be scored for mutant phenotypes at gastrula, neurula, and tailbud stages. Assuming normal survival rates at early stages, this will require sorting and visual screening of ~10,000 embryos per day for a team of five or six people, and perhaps two months to score the progeny of 500 F1 females.
Formally, half of the haploid and gynogenetic diploid progeny of parents carrying a single scorable recessive mutation should be phenotypically mutant. We are proposing to score both treatments. At later tailbud stages haploid syndrome abnormalities may obscure some phenotypes, and pressure treatment rarely rescues all of the embryos, so it is difficult to unambiguously identify mendelian ratios of subtle phenotypes. A haploid screen allows us to discard lines that are phenotypically wildtype or carry "uninteresting" phenotypes. Lines that carry "interesting" tissue-specific phenotypes will be maintained and characterized further by conventional inbreeding. We estimate that we will discard >90% of the lines at this point, and will be keeping <50 F2 families per F0 mutagenized male. 20 to 25 F2 outbred siblings of putative mutant haploid and gynogenetic diploid F2 embryos will be cultured to maturity, and inbred in natural matings. At one mutagenic "hit" per sperm, on average every other F2 sibling natural mating will result in 25% mutant phenotype, and will be free of the haploid syndrome background. The presence of additional mutations will result in higher and more variable phenotypic ratios, and will require additional segregation in order to characterize the individual mutations.
Focused screens:
How can we hope to characterize genes in pathways in which we are interested without the enormous investment involved in screening and maintaining large numbers of random mutations? We propose two strategies to focus our screens and make it easier to spot the rare mutation that affects a specific process in a background of wildtypes, less-relevant mutants, and epigenetic variations:
1. By mutagenizing and screening isogenic lines bearing tissue-specific GFP reporter activity. For example, lines homozygous for neural tubulin-GFP, which is expressed in the neural plate in a striped pattern during neural tube closure119 should provide a distinct search image on which to discern mutations affecting morphogenesis of the neural tube.
2. By screening haploid and gynogenetic diploid F2 embryos by in situ hybridization with various probe cocktails which reflect patterned gene expression in the wild-type embryo. Embryos screened in this fashion obviously cannot be rescreened later in development, and the labor involved is considerably greater than unaided visual scoring, but the available spectrum of molecular probes is much larger. Using mixed probes facilitates screening for phenotypes in multiple pathways simultaneously.
Target phenotypes:
1) mutations affecting movements of axial convergent extension, which define and elongate the anterior-posterior axis of the vertebrate body; screened on a neural tubulin-GFP background to enhance detection of axial defects:
a. blastopores that stall in closure at the midgastrula stage, when convergent extension normally begins;
b. short axes and widened notochordal and somitic tissues, indicating decreased rate or amount of convergent extension64, 65,
c.arched backs, indicating retarded mesodermal convergent extension;
d. blastopores that re-open, indicating locally fractured circumblastoporal arrays of converging cells, forming the classical ring embryo phenotype120,63.
2) mutations affecting eye formation:
a. small or missing lenses, assayed by diminished or missing GFP expression in crystallin promoter-GFP-bearing transgenic lines of S. tropicalis.
b. altered, reduced or missing retinal tissue, assayed by monitoring distribution of GFP expression in Six-3 promoter-GFP bearing transgenic lines of S. tropicalis.
We feel that it is important to be fairly stringent in restricting the phenotypes that we choose for extensive characterization. Our screen is designed so that visible morphological defects can be readily correlated with aberrations in relevant gene expression. As other developmental phenotypes which do not meet these criteria emerge in screens, they will be briefly evaluated and distributed to other Xenopus laboratories with cognate research interests.
Further characterization of morphogenic mutants:
First, axial defects will be analyzed for altered patterning of tissues, using whole mount in situ hybridization and immunohistochemistry for notochordal and somitic tissue, as well as a battery of region-specific neural markers. These include gsc (head mesoderm and notochord), tor 70 (notochord), 12-101 (somitic mesoderm), brachyury (pre- and post-involution notochord, preinvolution somitic mesoderm) for the mesoderm, and Otx2 (forebrain), engrailed (midbrain-hindbrain), hoxb1 (rhombomere 4), krox 20 (rhombomeres 3 and 5), hoxb9 (posterior neural), Xslu and Xsna (neural folds), and f-spondin (floorplate), among others. Abnormal gene expression may also reveal morphogenic defects; for example, tor 70 staining of the notochord may reveal abnormally wide and short notochords, indicating a failure of convergent extension, rather than mis-patterning. Morphogenesis can be analyzed by time lapse video, low light video fluorescence video, and time lapse recording of labeled cells, epi-illuminated embryos or explants to reveal defects in gross tissue movements4 or cell motility64. If defective mechanical properties of tissues are suspected, the relevant tissues can be measured for the forces they produce as well as the mechanical properties necessary for such movements as convergent extension67.
Further characterization of eye mutants:
A number of retinal121 and lens (Mullins, pers. comm.) mutations emerged from recent fish genetic screens. Because of the robust assays for discerning eye phenotypes described above, we also expect to uncover such mutants. Since a large number of candidate regulators of retinal or lens determination100,122,108 have already been cloned in Xenopus, a panel of genes may be tested for their ability to rescue mutations, and ordered into a hierarchy of epistasis, as mice mutated in the Pax-6 gene have been used to test whether other eye regulatory genes are upstream or downstream94. The embryology of amphibians should be invaluable in characterizing the mutants as well. For example, if a mutant were to fail to form a lens, extensive information about the inductive interactions and progression through the determination process100 should allow us to determine by transplantation and recombinant assays which step in the process is affected.
Cloning mutant loci:
The conventional method to isolate chemically-induced point mutations is positional cloning: isolating closely linked markers and mapping by recombination followed by chromosome walking. Even with a fine-structure genetic map, this is laborious. We will test the following alternatives:
1. candidate gene approaches:
Many genes with activity in pathways of interest have been identified. Assays used to evaluate candidates for a given mutation will include rescue of the mutant phenotype by injection of candidate gene mRNA and transgenic rescue of mutants with candidate wild-type gene driven by a suitable characterized promoter, and correlation of RAPD map position of the mutation and the candidate gene (see Specific Aim 1.D).
2. expression cloning approaches:
Specific biological activities have been identified in X. laevis mRNA pool injection and sib-selection assays by their ability to rescue the 'phenotype' of eggs exposed to ultraviolet light8 9. It may be possible to similarly rescue certain zygotic mutant phenotypes, by injecting pools of synthetic mRNA to deliver the missing gene product. Pools that rescue can then be split and re-assayed until the activity is cloned. X. laevis cDNA libraries in expression vectors designed for this purpose have already been constructed9, 123. Correlation of RAPD map position of mutations and sib-selected genes will then be used to confirm identity (see Specific Aim 1.D).
potential pitfalls:
Due to ENU toxicity, it could be difficult to implement a dose giving one mutagenic event per haploid genome. In this case, larger numbers of F1 frogs will be screened. The haploid syndrome background may preclude scoring mendelian ratios of late-developing phenotypes on F2 haploids and incompletely-rescued gynogenetic diploids; non-mendelian phenotypic ratios can also ensue from epigenetic processes. As a result, some of the outbred F2 families cultured for further analysis may not carry heritable abnormalities. However, mutants in the pathways we are interested in are likely to offer phenotypes prior to the appearance of the haploid syndrome (at tailbud stages). Multigene F2 phenotypes will likewise become apparent in F3 inbred progeny, with complex mendelian ratios of different phenotypes; interesting synthetic phenotypes may be less compelling once segregated.
timeframe:
titrate mutagenesis 1/98; screen first F2 haploids/gynogenetic diploids 6/98; screen first inbred F3 11/98
implementation:
Grainger, Keller, and DeSimone laboratories