Co-expression of the AsCas12a ultra variant, a T7 RNA Polymerase and a cytosine base editor greatly increases transfection and editing rates in Leishmania species

  1. Nicole Herrmann May
  2. Annika Schmid
  3. Elisabeth Meiser
  4. Tom Beneke

  • Curated by eLife

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    This manuscript describes a method for genetic manipulation of Leishmania species which should be sufficiently efficient to enable genome-wide genetic screens. The authors improved numerous aspects of their previously described method, which is based on sequence-specific genome editing to introduce premature stop codons using a CAS9-cytidine deaminase variant. The work is thoroughly described, with convincing data, and will be very important for Leishmania researchers, as well as perhaps suggesting the use of similar approaches in other organisms in which genetic manipulation is challenging.

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Abstract

The ability to analyse the function of all genes in a genome has obvious appeal. However, this has been challenging in Leishmania due to a repetitive genome architecture, limited DNA repair mechanisms and the absence of RNA interference machinery in most species. While our previous introduction of a cytosine base editor (CBE) tool in Leishmania showcased the potential for bypassing these limits (Engstler and Beneke (2023)), challenges remained in achieving high transfection efficiencies, overcoming species-specific editing rates, minimizing effects on parasite growth and eliminating competition between deleterious and non-deleterious mutations. Here, we present an optimized approach to address these limitations. Firstly, we identified a T7 RNAP promoter variant that ensures high editing rates across Leishmania species without adversely affecting parasite growth. Secondly, we adjusted the scoring of CBE single-guide RNAs (sgRNAs) to prioritize those ensuring STOP codon generation. Thirdly, we developed a triple-expression construct enabling the integration of CBE sgRNA expression cassettes into a Leishmania safe harbor locus via AsCas12a ultra-mediated DNA double-strand breaks. This facilitates the generation of stable CBE sgRNA expression cell lines and increases transfection rates by ∼400-fold, resulting in up to one transfectant per 70 transfected cells. Lastly, we show how the co-expression of AsCas12a ultra, T7 RNAP and CBE can be utilized for hybrid CRISPR gene replacement and base editing approaches in the same cell line. Overall, we believe that these improvements will broaden the range of possible gene editing applications in Leishmania species and will enable a variety of loss-of-function screens in the future.

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  1. Version published to 10.7554/elife.97437.1 on eLife
  2. Version published to 10.7554/elife.97437 on eLife
  3. eLife

    Author response:

    We would like to thank all reviewers and editors for their thorough peer review and valuable suggestions. In these provisional responses, we summarize the main concerns raised by the reviewers and outline our planned revisions to address them in the manuscript.

    Overall, we are pleased to note that the reviewers agree on the potential value of our updated toolbox for gene editing, highlighting its various applications. However, they also raised several valid concerns, which we have summarized and responded to as follows:

    (1) Mutant phenotypes in transfected populations can be occasionally reversed or escaped. This suggests it will not be possible to detect growth-associated phenotypes in pooled screens. An experiment with a pooled loss-of-function screen to test this is missing.

    Escapes or reversals of mutant phenotypes have been observed with other genetic tools used for loss-of-function screening, including lentiviral CRISPR approaches in mammalian systems and RNAi in Trypanosoma brucei. Cells can escape phenotypes through various mechanisms, such as promoter silencing or selection of non-deleterious mutations. Additionally, not every CRISPR guide is efficient in generating a mutant phenotype, and RNAi constructs can also vary in their effectiveness. Despite these challenges, genome-wide loss-of-function screens have been successfully carried out in mammalian cells and Trypanosoma parasites. Therefore, we believe that the observed escape of one mutant phenotype does not preclude the detection of growth-associated or other phenotypes in pooled screens. Moreover, we did not observe a reversal of the mutant phenotype in L. mexicana, L. donovani, and L. major parasites expressing tdTomato from an expression cassette integrated into the 18S rRNA SSU locus (Figure 4). However, the reviewers are rightfully requesting a pooled loss-of-function screen to validate this. Since submitting this manuscript, we have conducted multiple pooled loss-of-function screens, which have confirmed the ability of our here presented method to detect a range of mutant phenotypes in pooled screening formats. We will include these results in our revised manuscript.

    (2) The possibility of mis-integration of the CBE sgRNA expression construct into an entirely different locus is not explored.

    We plan to reanalyze our ONT sequencing data to verify if the CBE sgRNA expression construct was integrated into an unintended loci. If we detect any mis-integration events, we will evaluate their potential negative impacts and discuss these findings in the revised manuscript.

    (3) The achieved increase in editing efficiency compared to the previous base editing method could be more clearly presented.

    We have directly compared our improved method to our previous base editing method in Figures 1E and 4, demonstrating higher editing rates in a much shorter time. In the revised manuscript, we will present and describe the increase in editing rate more clearly.

    (4) The improvements on CBE sgRNA guide design are hypothetical and untested.

    We agree that the improvements to the CBE sgRNA design are currently hypothetical. We plan to systematically test our guide design principles in future studies. Since this will require testing hundreds of guides to draw robust conclusions, we believe that this aspect is beyond the scope of the current study. However, we will discuss our plans for future validation in the revised manuscript.

    Overall, we appreciate the reviewers' insights and are committed to addressing their concerns thoroughly. We believe that the planned revisions and additional experiments will significantly strengthen our manuscript and provide a more comprehensive evaluation of our updated gene editing toolbox.

  4. eLife

    eLife assessment

    This manuscript describes a method for genetic manipulation of Leishmania species which should be sufficiently efficient to enable genome-wide genetic screens. The authors improved numerous aspects of their previously described method, which is based on sequence-specific genome editing to introduce premature stop codons using a CAS9-cytidine deaminase variant. The work is thoroughly described, with convincing data, and will be very important for Leishmania researchers, as well as perhaps suggesting the use of similar approaches in other organisms in which genetic manipulation is challenging.

  5. eLife

    Reviewer #1 (Public Review):

    While CRISPR/Cas technology has greatly facilitated the ability to perform precise genome edits in Leishmania spp., the lack of a non-homologous DNA end-joining (NHEJ) pathway in Leishmania has prevented researchers from performing large-scale Cas-based perturbation screens. With the introduction of base editing technology to the Leishmania field, the Beneke lab has begun to address this challenge (Engstler and Beneke, 2023).

    In this study, the authors build on their previously published protocols and develop a strategy that:

    (1) allows for very high editing efficiency. The cell editing frequency of 1 edit per 70 cells reported in this study represents a 400-fold improvement over the previously published protocol,
    (2) reduces the negative effects of high sgRNA levels on parasite growth by using a weaker T7 promoter to drive sgRNA transcription.

    The combination of these two improvements should open the door to exciting large-scale screens and thus be of great interest to researchers working with Leishmania and beyond.

  6. eLife

    Reviewer #2 (Public Review):

    Summary:

    Previously, the authors published a Leishmania cytosine base editor (CBE) genetic tool that enables the generation of functionally null mutants. This works by utilising a CAS9-cytidine deaminase variant that is targeted to a genetic locus by a small guide RNA (sgRNA) and causes cytosine to thymine conversion. This has the potential to generate a premature stop codon and therefore a loss of function mutant.

    CBE has advantages over existing CAS-based knockout tools because it allows the targeting of multicopy gene families and, potentially, the easier generation of pooled loss of function mutants in complex population experiments. Although successful, the first generation of this genetic tool had several limitations that may have prevented its wider adoption, especially in complex genome-wide screens. These include nonspecific toxicity of the sgRNAs, low transfection efficiencies, low editing efficiencies, a proportion of transfectants that express multiple different sgRNAs, and insufficient effectivity in some Leishmania species.

    Here, the authors set out to systematically solve each of these limitations. By trialling different transfection conditions and different CAS12a cut sites to promote sgRNA expression cassette integration, they increase the transfection efficiency 400-fold and ensure that only a single sgRNA expression cassette integrates that edits with high efficiencies. By trialling different T7 promoters, they significantly reduce the non-specific toxicity of sgRNA expression whilst retaining high editing efficiencies in several Leishmania species (Leishmania major, L. mexicana and L. donovani). By improving the sgRNA design, the authors predict that null mutants will be more efficiently produced after editing.

    This tool will find adoption for producing null mutants of single-copy genes, multicopy gene families, and potentially genome-wide mutational analyses.

    Strengths:

    This is an impressive and thorough study that significantly improves the previous iteration of the CBE. The approach is careful and systematic and reflects the authors' excellent experience developing CRISPR tools. The quality of data and analysis is high and data are clearly presented.

    Weaknesses:

    Figure 4 shows that editing of PF16 is 'reversed' between day 6 and day 16 in L. mexicana WTpTB107 cells. The authors reasonably conclude that in drug-selected cells there is a mixed population of edited and non-edited cells, possibly due to mis-integration of the sgRNA expression construct, and non-edited cells outcompete edited cells due to a growth defect in PF16 loss of function mutants. However, this suggests that the CBE tool will not work well for producing mutants with strong fitness phenotypes without incorporating a limiting dilution cloning step (at least in L. mexicana and quite possibly other Leishmania species). Furthermore, it suggests it will not be possible to incorporate genes associated with a growth defect into a pooled drop-out screen as described in the paper. This issue is not well explored in the paper and the authors have not validated their tool on a gene associated with a severe growth defect, or shown that their tool works in a mixed population setting.

    Although welcome, the improvements to the crRNA CBE design tool are hypothetical and untested.

    The Sanger and Oxford Nanopore Technology analyses on integration sites of the sgRNA expression cassette integration will not detect the mis-integration of the sgRNA expression construct into an entirely different locus.

  7. eLife

    Reviewer #3 (Public Review):

    Genetic manipulation of Leishmania has some challenges, including some limitations in the DNA repair strategies that are present in the organism and the absence of RNA interference in many species. The senior author has contributed significantly to expanding the available routes towards Leishmania genetic manipulation by developing and adapting CRISPR-Cas9 tools to allow gene manipulation via DNA double-strand break repair and, more recently, base modification. This work seeks to improve on some limitations in the tools previously described for the latter approach of base modification leading to base change.

    The work in the paper is meticulously described, with solid evidence for most of the improvements that are claimed: Figure1 clearly describes reduced impairment in the growth of parasites expressing sgRNAs via changes in promoters; Figures 2 and 3 compellingly document the usefulness of using AsCas12a for integration after transformation; and Figures 1 and 4 demonstrate the capacity of the combined modifications to efficiently edit a gene in three different Leishmania species. There is little doubt these new tools will be adopted by the Leishmania community, adding to the growing arsenal of approaches for genetic manipulation.

    There are two weaknesses the authors may wish to address, one smaller and one larger.

    (1) The main advance claimed here is in this section title: 'Integration of CBE sgRNA expression cassettes via AsCas12a ultra-introduced DSBs increase editing rates', with the evidence for this presented in Figure 4. It is hard work in the submission to discern what direct evidence there is for editing rates being improved relative to earlier, Cas9-based approaches. Did they directly compare the editing by the new and old approach? If not, can they more clearly explain how they are able to make this claim, either by adding text or a new figure? A side-by-side comparison would emphasise the advance of the new approach more clearly.

    (2) The ultimate, stated goal of this work is (abstract) to 'enable a variety of loss-of-function screens', as the older approach had some limitations. This goal is not tested for the new tools that have been developed here; the experiment in Figure 5 merely shows that they can, not unexpectedly, make a gene mutant, which was already possible with available tools. Thus, to what extent is this paper describing a step forward? Why have the authors not run an experiment - even the same one that was described previously in Engstler and Beneke (2023) - to show that the new approach improves on previous tools in such a screen, either in scale or accuracy?

  8. Version published to 10.1101/2024.02.28.582587v1 on bioRxiv