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WIREs Dev Biol
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From genes to function: the C. elegans genetic toolbox

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Abstract This review aims to provide an overview of the technologies which make the nematode Caenorhabditis elegans an attractive genetic model system. We describe transgenesis techniques and forward and reverse genetic approaches to isolate mutants and clone genes. In addition, we discuss the new possibilities offered by genome engineering strategies and next‐generation genome analysis tools. WIREs Dev Biol 2012, 1:114–137. doi: 10.1002/wdev.1 This article is categorized under: Technologies > Perturbing Genes and Generating Modified Animals

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Mutant mapping by whole‐genome sequencing (WGS). (a) WGS‐single nucleotide polymorphism (WGS‐SNP) mapping. A homozygous mutant strain (mutation denoted by a red diamond) in a Bristol background is crossed with a SNP‐containing Hawaiian strain. Mutant worms segregating from this cross are reselected in the F2 generation. These mutant lines carry chimeric chromosomes (containing Bristol and Hawaiian SNPs) generated by meiotic recombination. When DNA from multiple F2 recombinant lines is pooled and sequenced, then the relative number of Bristol versus Hawaiian SNP at a given location in the genome will be a reflection of the relative distribution of recombinants in the pool. Therefore, only Bristol SNPs will be found close to the mutation of interest. (Reprinted with permission from Ref 28. Copyright 2010 Public Library of Science) (b) Mapping based on mutagen‐induced DNA variation density across the genome. Mutagenized strains contain not only mutagen‐induced damage (orange asterisks) but also carry background variants (green crosses) present in the mother strain before mutagenesis. By backcrossing a candidate mutant line to the strain used for the original mutagenesis, it is possible to eliminate most mutagen‐induced lesions except for those which are closely linked to the phenotype‐causing mutation (red diamond). In addition, background variation present in the unmutagenized strain can be identified by comparing independent backcrossed mutant lines. After filtering, the genomic region linked to the phenotype‐causing mutation will be highlighted by a high‐density cluster of mutagen‐induced lesions. (Reprinted with permission from Ref 29. Copyright 2010 Genetics Society of America)

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Caenorhabditis elegans fosmid recombineering. Recombineering can be used to efficiently engineer a gene of interest in the context of a fosmid genomic clone. For example, a cassette containing a GFP tag and the FRT‐galK‐FRT selection module (FgF) can be inserted by λ Red recombinase‐mediated homologous recombination into any position of a gene. Recombination is carried out in the SW105 E. coli strain. Recombinant clones are selected on minimal medium with galactose. Excision of the FgF selection module is carried out by Flp recombinase (induced by addition of arabinose) and can be selected for in medium containing deoxy‐galactose. Note that the FRT ‘scar’ remaining after galK excision resides in an intron and therefore has no impact on protein‐coding sequences. Additional applications of fosmid recombineering include: sequence deletion, sequence replacement, and fosmid extension. A comprehensive set of recombineering cassettes including different fluorescent reporters, affinity tags, nuclear localization signals, mutant FRT sites, and gene replacement cassettes is available. (Reprinted with permission from Ref 140. Copyright 2009 Public Library of Science)

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MosTIC, MosDEL and MosSCI strategies. (a) The double‐strand break (DSB) induced by the excision of Mos1 (i) can be repaired by MosTIC using different repair templates to engineer point mutations (ii), fusion proteins (iii), or gene deletions (iv). In the case of MosDEL (v), a unc‐119(+) mini‐gene is used as a positive selection marker. (b) The goal of MosSCI is to engineer single‐copy transgenes at a defined location in the genome. A DSB induced by Mos1 excision is repaired by homologous recombination with an in vitro‐engineered template which contains the desired transgene in combination with the unc‐119(+) mini‐gene.

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Mos1‐based insertional mutagenesis. (a) Mos1 can jump from an extrachromosomal transgene carrying multiple copies of the transposon into the genome, following the expression of the Mos1‐specific transposase via a heat‐shock inducible transgene. Mos1 insertions are generally random in the genome but always take place at a TA dinucleotide. (b) Mos1 insertions can be rapidly mapped with single nucleotide resolution by sequencing the flanking genomic sequences using an inverse PCR strategy. (Reprinted with permission from Ref 44. Copyright 2007 Nature Publishing Group)

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Genome editing with engineered nucleases. (a) Structure of a zinc finger (ZF) DNA‐binding module. Each individual ZF domain is optimized to recognize a 3‐bp sequence. Two cysteines and two histidines coordinate a zinc atom in each module. (b) ZF nucleases (ZFN) are composed of at least four ZF domain repeats, linked to the FokI endonuclease domain. Specificity is increased by using obligatory dimers of two ZFN recognizing two sites which flank the targeted double‐strand break (DSB) site. (c) Structure of TALE nucleases (TALEN). A central array of tandem repeats forms the specific DNA‐binding domain. Two nucleotides (repeat‐variable di‐residues, RVD) in each repeat determine the sequence preference of each module. Examples of four RVDs that recognize all four nucleotides (NT) are indicated. Two TALEN monomers bind to their target sites and allow the dimerization of the FokI endonuclease domain. TALENs also contain a nuclear localization signal (NLS). (d) ZFNs and TALENs allow different types of genome editing strategies. A DSB can be resolved by different pathways. (i) A single DSB can be repaired by non‐homologous end‐joining (NHEJ) and result in a small deletion of a few base pairs. Alternatively, (ii) two simultaneous DSB can generate a larger deletion allele. When a repair template with homology regions is provided, the DSB is resolved by gene conversion to (iii) insert a small sequence (as small as a single nucleotide, i.e., ‘gene correction’) or (iv) a larger transgene (‘gene addition’).

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Generating deletion mutants by library screening. Populations of randomly mutagenized worms can be established using different mutagens and different culture strategies. Mutant strains are either preserved in a frozen library or conserved on NGM plates in a ‘live’ library. To optimize library screening, DNA samples extracted from all worm cultures can be pooled according to different pooling schemes. Deletions with sizes ranging from 500 to 1500 bp are detected by nested PCR screening. Shorter deletions can be detected more easily by including a poison‐primer. Once a candidate pool has been identified, the corresponding well is thawed from the frozen library and all recovered worms are cloned on individual plates for secondary rescreening. Similarly, worms from a ‘live’ library are recultured in small pools and single worms are picked from PCR‐positive pools. Once a single worm harboring a deletion has been isolated, a few steps should be undertaken before the strain can be used for functional studies.

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