Jaryn Perkins et al. (2005) International Worm Meeting "A Caenorhabditis elegans fosmid library"

Rene Warren, Kim Wong, Jeff Stott, Rob Holt, Jacquie Schein, Jaryn Perkins, Steve Jones, Marco Marra, Don Moerman

[International Worm Meeting, 2005]

The availability of genomic clones with 35 to 40kb inserts has been a critical resource to the C. elegans community. Fingerprinted and ordered cosmid clones harboring such inserts were the backbone of the sequencing project. Further utility was found when individual research labs were trying to map mutants. Complementing a mutant by injecting a strain with overlapping cosmid clones eventually led to mutation identity. This approach is still widely used and is an important tool for mutational analysis. Many of the cosmid libraries are now 20 years old and there are reports of some of them losing viability. This persuaded us to consider building a new DNA library with total coverage of the genome. We also felt it was best to make a 40kb clone insert library rather than rely on YAC or BAC clone libraries. The large insert clones are not trivial to manipulate and do not in many cases give you the resolution that is required for many experiments. With these considerations in mind we have chosen to make a fosmid rather than a cosmid library. The cosmid libraries have been excellent workhorses but many clones within the libraries have been prone to rearrangements. By using a fosmid vector (pCC1FOS from Epicentre) that is maintained at low copy number until induced we hope that the occurrence of such rearrangements will be reduced. The use of fosmids as backbones allow for the maintenance of large pieces of DNA (around 40kB) in limited number (1-5) per bacterial host. Up to this point, such a resource was unavailable for C. elegans We have created an N2 fosmid library and are currently mapping clones to the genome. We are mapping all fosmid clones to the C. elegans genome by pair-wise alignment of fosmid end-reads. Initial analysis of the library quality shows that the average insert size is ~35 kb and 86% of the clones have paired end-reads with correct relative orientation. Currently, with over 6,100 clones sequenced we have obtained approximately 2X clone coverage of the genome. From this, we estimate that we will require ~16,000 clones to obtain our goal of 5X coverage of the genome. This resource was created for public domain use and is available to interested researchers.

Tenenhaus T et al. (1997) International C. elegans Meeting "GENETIC REQUIREMENTS FOR PIE-1 EXPRESSION AND RNA POLYMERASE II INHIBITION IN THE EMBRYONIC GERM LINEAGE OF C. ELEGANS"

Tenenhaus T, Seydoux GC

[International C. elegans Meeting, 1997]

The germline blastomeres (P1-P4) fail to transcribe many messenger RNAs expressed in somatic cells(1). This deficiency in mRNA transcription requires the germline factor PIE-1(1,2), and correlates with the absence of a particular phosphoepitope on RNA polymerase II (RNAP II-H5) (see abstract by Dunn and Seydoux). To begin to understand the genetic requirements for the repression of RNA polymerase II activity in the germ lineage, we have analysed pre-existing embryonic lethal mutations for the expression of PIE-1 and RNAP II-H5 by immunohistochemistry. We find that many mutations (par-1, par-2, par-3, par-4, par-6, cib-1, mes-1, emb-1, emb-6, emb-8, emb-16, emb-20, emb-21, emb-25, emb-27, emb-30, zyg-1, zyg-2, zyg-9) result in the complete or progressive loss of PIE-1 expression in the germ lineage. In all cases, these mutations cause inappropriate expression of RNAP II-H5 in the germ lineage, suggesting that PIE-1 is required in each germline blastomere to repress RNAP II activity. We have also found two mutations (mex-1, mex-3) that cause ectopic expression of PIE-1 in somatic blastomeres; preliminary results suggests that these mutations also affect RNAP II-H5 expression, suggesting that RNAP II-H5 expression is sensitive to variations in PIE-1 levels. Further analysis of these and other mutations will test the hypothesis that PIE-1 is necessary and sufficient to repress RNA polymerase II phosphorylation, and will identify loci required to restrict PIE-1-dependent repression of transcription to the germ lineage. We thank the CGC for strains and Craig Mello, Charlotte Schubert, Jim Priess and Steve Warren for antibodies. 1. Seydoux et al. (1996). Nature 382, 713-716. 2. Mello et al. (1996). Nature 382, 710-712.

Mihail Sarov et al. (2007) International Worm Meeting "Recombineering based high throughput protein tagging in C. elegans."

Mihail Sarov, Francis Stewart, Anthony Hyman

[International Worm Meeting, 2007]

The methods for protein tagging in C. elegans so far have been inefficient and largely dependent on artificial cDNA based constructs, which can lack important regulatory elements. The recent generation of a genomic fosmid library[1] opened the possibility for direct modification of any gene of interest in its native genomic environment by recombineering in E. coli. We recently published a method for precise in vivo engineering of large genomic clones into GFP tagged transgenes for ballistic transformation[2]. Using such transgenes, we reproduce known and document new expression patterns. We further demonstrate that the tagged protein can complement the function of its endogenous counterpart. Applying the latest developments of the recombineering field and introducing several innovations of our own we have now streamlined the protocol to allow rapid 96 well format liquid culture processing in a continuous recombineering pipeline that takes just four days. This unprecedented throughput and the achieved economy of scale allow us to rapidly produce thousands of tagged transgenes. Our method has become the basis for a large-scale project aimed at comprehensive description of the expression pattern (through fluorescent reporters) and the DNA binding sites (through affinity tag based chromatin immunoprecipitation) of more than 400 C. elegans transcription factor genes (as part of the NIH funded ModENCODE program). At the meeting we will discus the advantages of a distributed, community based collaboration model that will bring this technology to any worm lab and will make the genome wide protein tagging a feasible goal. 1. Perkins, J., K. Wong, R. Warren, J.E. Schein, J. Stott, R. Holt, S. Jones, M.A. Marra, and D. Moerman. 2005. A Caenorhabditis elegans fosmid library. In 15th International C. elegans Conference, University of California, Los Angeles. 2. Sarov, M., S. Schneider, A. Pozniakovski, A. Roguev, S. Ernst, Y. Zhang, A.A. Hyman, and A.F. Stewart. 2006. A recombineering pipeline for functional genomics applied to Caenorhabditis elegans. Nature Methods 3: 839-844.

van Luenen HGAM et al. (1996) European Worm Meeting "The use of Tc1 and Tc3 transposons in Caenorhabditis elegans and other species"

Vos JC, van der Linden KM, Hazendonk E, De Baere I, van Luenen HGAM, Plasterk RHA

[European Worm Meeting, 1996]

Tc1 and Tc3 transposable elements are the most active transposons in the nematode C. elegans. They are responsible for most of the spontaneous mutations in certain strains. Tc1 and Tc3 belong to a large family of transposons found in many different organisms, ranging from fungi to vertebrates, including man. We have investigated the mechanism of excision and integration of Tc1 and Tc3 (1). The requirements for this reaction are limited: the transposase protein encoded by the element (2), and the sequences at both ends of the transposon (see the abstract of Rene Rezsohazy). These limited requirements for transposition might explain the successful spread of the Tc1-like elements throughout the animal kingdom. The limited requirements for transposition might allow the use of Tc1 or Tc3 as a gene delivery system in C. elegans and other species. We are currently collaboration with several groups to investigate whether Tc1/Tc3 transposes in mice, zebrafish and insects. The C. elegans genome sequencing project will be making the complete genomic sequence available within two years. Efficient reverse genetics will be needed to study the functions of the newly identified genes. One method to determine the function of a gene is to disrupt the coding sequence by the insertion of a transposon. We have developed an efficient method for the isolation of transposon insertion mutants in a specific sequence (3). A large collection of different cultures of strain MT3126 has been generated and frozen away. The strain MT3126 contains many Tc1 transposons which are mobile in the germ line. DNA samples of all these cultures have been pooled in an organized way. With only a few PCR's we can identify a transposon insertion in a specific sequence and the frozen culture can be thawed. A transposon insertion in a gene is not a guaranteed null allele of a gene. Therefore, isolation of a transposon insertion is followed by the isolation of a transposon excision induced deletion of the genomic region. We screen our collection on a routine basis for laboratories all over the world and we have obtained thus far transposon insertions in over a 100 different genes. Transposons can also be used for forward genetic analysis. We have developed a quick method to display all transposon insertions in a genome in a single lane of a sequencing gel. Screening a panel of different cultures, it is possible to determine which transposon insertion co-segregates with a phenotype caused by a transposon integration. This band can be excised from gel and the transposon flank can be sequenced. This sequence is used to screen the data base and the mutated gene can identified. With this method it is possible to identify the gene that causes the phenotype within 2 or 3 days, whereas with the classical method of transposon tagging and cloning it would at least take a couple of months. Secondly, we have determined most Tc1 insertion sites in three strains with a high Tc1 copy number. These insertions can be visualized by PCR and they can be used for mapping mutant genes (see abstract of Rik Korswagen). 1. Van Luenen, H. G. A. M., Colloms, S. D. & Plasterk, R. H. A. (1994) Cell 79, 293-301. 2. Vos, J. C., de Baere, I. & Plasterk, R. H. A. (1996) Genes & Dev. 10, 755-761. 3. Zwaal, R. R., Broeks, A., Van Meurs, J., Groenen, J. T. M. & Plasterk, R. H. A. (1993) Proc. Natl. Acad. Sci. USA 90, 7431-7435.