This post was contributed by Jordan Ward who is a postdoctoral fellow at UCSF.
Emerging CRISPR/Cas9 editing technologies have transformed the palette of experiments possible in a wide range of organisms and cell lines. In C. elegans, one of the model organisms which I use to study gene regulation during developmental processes, CRISPR/Cas9 allows us to knock out sequences and introduce mutations and epitopes with unprecedented ease. In the last year, several advances in C. elegans genome editing using CRISPR/Cas9 have emerged, which I will describe below. These new C. elegans approaches rapidly enrich for editing events without the need for any selective marker to remain in the edited animal. To my knowledge these approaches have not yet been extended to other organisms/cell lines, though it is likely that many aspects will broadly improve editing efficiency.
Development of CRISPR/Cas9 Editing Strategies in C. elegans
As brilliantly and irreverently illustrated in the parable of the Geneticist vs the Biochemist, we geneticists are a lazy bunch who love to rely on the awesome power of genetic selection. Knock-in events tend to be rare, and the challenge with any experiment is in efficient recovery of these edits. Initial work in C. elegans relied on selective markers such as drug resistance (Chen et al., 2013), fluorescence (GFP; Tzur et al., 2013), or rescue of mutant phenotypes (unc-119, Dickinson et al., 2013). These approaches allowed effective recovery of knock-ins, but did result in 1-2 kilobases of additional sequence being introduced. Cre-mediated excision of the selective cassette minimizes the sequence added, leaving a 34 bp “scar”, but currently requires additional experimental manipulation. Including Cre-mediated marker excision, it takes approximately four weeks to obtain a homozygous, outcrossed knock-in ready for experimentation.
In the last year, several manuscripts published in Genetics detailed approaches in which selection for an editing event that produces a visible phenotype enriches for knock-outs and knock-ins at other genomic loci. The first approach – co-CRISPR from Craig Mello’s lab – used inactivation of the unc-22 gene as their selection marker (Kim et al. 2014). Meanwhile Andy Fire’s lab used oligo-mediated knock-in of a dominant mutation, known as co-conversion (Arribere et al., 2014). In both cases, the selected mutation must be removed, which can be done by isolating animals with particular visible phenotypes. These approaches may become even more powerful following a report that one can use linear repair templates (ie. PCR-derived dsDNA) with 30-60 basepair homology arms to knock in large epitopes, such as GFP (Paix et al., 2014). The co-CRISPR and co-conversion approaches have the advantage of being used in any genetic background, but require variable amounts of experimental manipulation and screening. Additionally, they take from 8-14 days to recover knock-in homozygotes ready for experimentation, depending on the screening strategy.
To Find a Needle, Remove the Haystack
I was fortunate – or unfortunate enough, depending on perspective – to work with inefficient sgRNAs in my initial direct screening efforts, which were similar to the approach detailed by Paix et al. Although I was able to knock-in a 2xFLAG epitope into my gene of interest, I encountered low efficiency (0.13%) and laborious handling (screening 768 F1 animals). Recovering rare editing events in a sea of unedited animals struck me as a “needle in the haystack” type of problem and led to me to explore alternate approaches. Being a “lazy” geneticist, I developed a co-selection approach relying on repair of a conditional-lethal mutation to identify edits with minimal screening effort. Selecting for repair of a lethal mutation should remove much of the unedited “haystack”, facilitating recovery of edited animals.
In my recent Genetics paper (Ward, 2015), I demonstrated that selection for repair of a temperature-sensitive pha-1 mutation significantly enriches for knock-in of 2x and 3xFLAG epitopes into other, non-linked loci; pha-1(e2123) mutant worms are perfectly viable at 15 ºC, yet exhibit complete embryonic lethality at 25 ºC. This method resulted in efficiencies ranging from 11-100% of F1 animals carrying precise knock-ins, and homozygous knock-in animals can be obtained in eight days. The only animals on a plate, other than the parental animal, are rescued progeny, which makes screening extremely rapid. This stringent selection allowed me to optimize a range of editing parameters: oligo repair templates with homology arms of 35-80 bp, and DNA double-strand breaks up to 54 bp from the desired insertion site result in efficient editing. Repair oligos do not need to be PAGE purified, although doing so increases knock-in efficiency. Finally, as shown in Drosophila S2 cells (Böttcher et al., 2014), inactivation of non-homologous end-joining results in a further increase in knock-in efficiency, presumably by channeling DNA breaks into the homologous recombination repair pathway. Reagents required to perform pha-1 co-conversion are available through Addgene.
It is interesting to note that recipients for the 2015 Breakthrough Prize included Jennifer Doudna and Emmanuelle Charpentier for their pioneering CRISPR/Cas9 work, and Victor Ambros and Gary Ruvkun for their seminal work on micro RNAs in C. elegans. The C. elegans work on small RNAs informed and drove work in countless other systems, whereas import of CRISPR technology into C. elegans has been a transformative innovation for our community. Developments in mammalian and yeast cells using modified Cas9 proteins to regulate gene expression (CRISPRi/CRISPRa), visualize specific genomic loci (CRISPR-imaging), or to find the proteins associated with a given genomic locus (CRISPR-ChAP-MS) could further transform the range of experiments imaginable in C. elegans. Equally, some of the “elegans” methods outlined in this post – particularly the use of co-selection methods – could greatly streamline editing in other organisms and systems, allowing rapid progress in a wide range of fields.
Thank you to our guest blogger!
Jordan Ward is a postdoctoral fellow with Keith Yamamoto at UCSF, and is currently studying gene regulation during nematode molting while preparing to transition to his own independent group. He received his PhD in Biochemistry from Cancer Research UK/University of London studying mechanisms of DNA repair in the lab of Professor Simon Boulton. Learn more at his webpage or follow him on Twitter.
Want to learn more about genome editing with CRISPRs?
- Check-out Addgene's CRISPR Resources