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Chlamydomonas reinhardtii , a model organism for unicellular green microalgae, is widely used in basic and applied research. Nonetheless, proceeding towards synthetic biology requires a full set of manipulation techniques for inserting, removing, or editing genes. Despite recent advancements in CRISPR/Cas9, still significant limitations in producing gene knock-outs are standing, including (i) unsatisfactory genome editing (GE) efficiency and (ii) uncontrolled DNA random insertion of antibiotic resistance markers. Thus, obtaining efficient gene targeting without using marker genes is instrumental in developing a pipeline for efficient engineering of strains for biotechnological applications. We developed an efficient DNA-free gene disruption strategy, relying on phenotypical identification of mutants, to (i) precisely determine its efficiency compared to marker-relying approaches and (ii) establish a new DNA-free editing tool. This study found that classical CRISPR Cas9-based GE for gene disruption in Chlamydomonas reinhardtii is mainly limited by DNA integration. With respect to previous results achieved on synchronized cell populations, we succeeded in increasing the GE efficiency of single gene targeting by about 200 times and up to 270 times by applying phosphate starvation. Moreover, we determined the efficiency of multiplex simultaneous gene disruption by using an additional gene target whose knock-out did not lead to a visible phenotype, achieving a co-targeting efficiency of 22%. These results expand the toolset of GE techniques and, additionally, lead the way to future strategies to generate complex genotypes or to functionally investigate gene families. Furthermore, the approach provides new perspectives on how GE can be applied to (non-) model microalgae species, targeting groups of candidate genes of high interest for basic research and biotechnological applications.

Background Large-scale cultivation of microalgae provides a carbon–neutral source of biomass for extracting valuable compounds and producing renewable fuels. Owing to their high metabolic activity and rapid reproduction rates, Chlorella species are highly productive when grown in photobioreactors. However, wild-type strains have some biological limitations that make algal bioproducts more expensive than those from more traditional sources. Domestication is thus required for improving strains. Engineering Chlorella species has been made difficult by their chemically complex and highly resistant cell wall, making transformation difficult. Cell wall also restricts diffusion of organic solvents; thus, limiting the extraction of valuable intracellular compounds. Obtaining strains with weakened cell wall is crucial to enhance the extractability of intracellular molecules, reducing the costs of biomass disruption, and to improve genetic transformation efficiency. Results We developed a mutagenesis pipeline combined with single-cell fluorescence scanning on the microalga Chlorella vulgaris to identify mutants with altered cell wall properties. We used the fluorescent dyes erythrosin B and calcofluor white, as markers for cell wall permeability and for binding the structural polysaccharides of the cell wall, respectively. Flow cytometry with fluorescence-activated cell sorting was employed to enrich mutagenized populations with altered emission profiles. After a first round of mutagenesis, we found six mutants with significantly higher cell permeability to erythrosin B than the wild type (CWP lines) and altered cell wall structure and composition. A second round of mutagenesis on a selected CWP strain, followed by selection for lower calcofluor white signal, resulted in the isolation of CFW lines, which exhibited reduced mechanical resistance when the biomass was subjected to cell disruption procedures. This two-steps procedure allowed us to identify new mutant strains with both an increased cell wall permeability and a reduced mechanical resistance, making a novel step towards Chlorella domestication. Conclusions This study demonstrated the feasibility of using mutagenesis and phenotypic selection based on flow cytometry screening to alter the cell wall of C. vulgaris and identify promising strains with improved traits for industrial applications.