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1. The minimal SUF system is not required for Fe–S cluster biogenesis in the methanogenic archaeon Methanosarcina acetivorans

   https://doi.org/10.1038/s41598-023-42400-x  

   Saini, Jasleen ; Deere, Thomas M. ; Lessner, Daniel J. ( September 2023 , Scientific Reports)

   Iron–sulfur (Fe–S) proteins are essential for the ability of methanogens to carry out methanogenesis and biological nitrogen fixation (diazotrophy). Nonetheless, the factors involved in Fe–S cluster biogenesis in methanogens remain largely unknown. The minimal SUF Fe–S cluster biogenesis system (i.e., SufBC) is postulated to serve as the primary system in methanogens. Here, the role of SufBC inMethanosarcina acetivorans, which contains twosufCBgene clusters, was investigated. The CRISPRi-dCas9 and CRISPR-Cas9 systems were utilized to repress or deletesufC1B1andsufC2B2, respectively. Neither the dual repression ofsufC1B1andsufC2B2nor the deletion of bothsufC1B1andsufC2B2affected the growth ofM. acetivoransunder any conditions tested, including diazotrophy. Interestingly, deletion of onlysufC1B1led to a delayed-growth phenotype under all growth conditions, suggesting that the deletion ofsufC2B2acts as a suppressor mutation in the absence ofsufC1B1. In addition, the deletion ofsufC1B1and/orsufC2B2did not affect the total Fe–S cluster content inM. acetivoranscells. Overall, these results reveal that the minimal SUF system is not required for Fe–S cluster biogenesis inM. acetivoransand challenge the universal role of SufBC in Fe–S cluster biogenesis in methanogens.

    

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2. CRISPRi repression of nonhomologous end‐joining for enhanced genome engineering via homologous recombination in Yarrowia lipolytica

   https://doi.org/10.1002/bit.26404  

   Schwartz, Cory ; Frogue, Keith ; Ramesh, Adithya ; Misa, Joshua ; Wheeldon, Ian ( September 2017 , Biotechnology and Bioengineering)

   In many organisms of biotechnological importance precise genome editing is limited by inherently low homologous recombination (HR) efficiencies. A number of strategies exist to increase the effectiveness of this native DNA repair pathway; however, most strategies rely on permanently disabling competing repair pathways, thus reducing an organism's capacity to repair naturally occurring double strand breaks. Here, we describe a CRISPR interference (CRISPRi) system for gene repression in the oleochemical‐producing yeastYarrowia lipolytica. By using a multiplexed sgRNA targeting strategy, we demonstrate efficient repression of eight out of nine targeted genes to enhance HR. Strains with nonhomologous end‐joining repressed were shown to have increased rates of HR when transformed with a linear DNA fragment with homology to a genomic locus. With multiplexed targeting ofKU70andKU80, and enhanced repression with Mxi1 fused to deactivated Cas9 (dCas9), rates of HR as high as 90% were achieved. The developed CRISPRi system enables enhanced HR inY. lipolyticawithout permanent genetic knockouts and promises to be a potent tool for other metabolic engineering, synthetic biology, and functional genomics studies.

    

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3. CRISPR-Cas tools for simultaneous transcription & translation control in bacteria

   https://doi.org/10.1093/nar/gkae275  

   Cardiff, Ryan A. L. ; Faulkner, Ian D. ; Beall, Juliana G. ; Carothers, James M. ; Zalatan, Jesse G. ( April 2024 , Nucleic Acids Research)

   Robust control over gene translation at arbitrary mRNA targets is an outstanding challenge in microbial synthetic biology. The development of tools that can regulate translation will greatly expand our ability to precisely control genes across the genome. In Escherichia coli, most genes are contained in multi-gene operons, which are subject to polar effects where targeting one gene for repression leads to silencing of other genes in the same operon. These effects pose a challenge for independently regulating individual genes in multi-gene operons. Here, we use CRISPR-dCas13 to address this challenge. We find dCas13-mediated repression exhibits up to 6-fold lower polar effects compared to dCas9. We then show that we can selectively activate single genes in a synthetic multi-gene operon by coupling dCas9 transcriptional activation of an operon with dCas13 translational repression of individual genes within the operon. We also show that dCas13 and dCas9 can be multiplexed for improved biosynthesis of a medically-relevant human milk oligosaccharide. Taken together, our findings suggest that combining transcriptional and translational control can access effects that are difficult to achieve with either mode independently. These combined tools for gene regulation will expand our abilities to precisely engineer bacteria for biotechnology and perform systematic genetic screens.

    

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4. Efficient CRISPR/Cas9 gene editing in a tilapia cell line model using endogenous promoters

   Hamar, JC* ; Kültz, D ( January 2021 , SICB Virtual Annual Meeting 2021)

   null (Ed.)

   Cell cultures are effective supplemental models to study specific biochemical pathways used for environmental adaption in animals. They enable isolation from system influence and facilitate control the extracellular environment. For work focusing on fish species many representative cell lines now exist, including a tilapia brain cell line (OmB) developed in our lab. CRISPR/Cas9 gene editing is an additional tool aiding these studies by allowing manipulation of specific genetic loci and evaluating their causal relationship between phenotypes of interest. However, established CRISPR/Cas9 gene targeting tools and methods often have not functioned as efficiently in fish cells as seen in other animal cell models such as mammalian cell lines, consistent with our initial attempts to apply CRISPR/Cas9 in OmB cells that failed to indicate genomic alteration at the targeted sites. Poor expression of heterologous promoters in OmB cells was hypothesized to be a primary cause for this occurrence so we constructed a custom plasmid vector based system utilizing tilapia endogenous promoters (EF1 alpha to express Cas9 and a U6 to express gRNAs). This system demonstrated substantial editing of most target sites attempted with mutational efficiency as high 80%. This work specifically highlighted the importance of phylogenetic proximity in selection of a polymerase III promoter for gRNA expression as commonly used interspecies U6 promoters (human and zebrafish) yielded no detectable gene editing when applied in this system with a common gRNA target sequence. These new tools will allow generation of knockout cell lines for gene targeting studies in tilapia and other phylogenetically close fish species. 

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5. Programmable Gene Knockdown in Diverse Bacteria Using Mobile‐CRISPRi

   https://doi.org/10.1002/cpmc.130  

   Banta, Amy B. ; Ward, Ryan D. ; Tran, Jennifer S. ; Bacon, Emily E. ; Peters, Jason M. ( December 2020 , Current Protocols in Microbiology)

   Facile bacterial genome sequencing has unlocked a veritable treasure trove of novel genes awaiting functional exploration. To make the most of this opportunity requires powerful genetic tools that can target all genes in diverse bacteria. CRISPR interference (CRISPRi) is a programmable gene‐knockdown tool that uses an RNA‐protein complex comprised of a single guide RNA (sgRNA) and a catalytically inactive Cas9 nuclease (dCas9) to sterically block transcription of target genes. We previously developed a suite of modular CRISPRi systems that transfer by conjugation and integrate into the genomes of diverse bacteria, which we call Mobile‐CRISPRi. Here, we provide detailed protocols for the modification and transfer of Mobile‐CRISPRi vectors for the purpose of knocking down target genes in bacteria of interest. We further discuss strategies for optimizing Mobile‐CRISPRi knockdown, transfer, and integration. We cover the following basic protocols: sgRNA design, cloning new sgRNA spacers into Mobile‐CRISPRi vectors, Tn7transfer of Mobile‐CRISPRi to Gram‐negative bacteria, and ICEBs1transfer of Mobile‐CRISPRi to Bacillales. © 2020 The Authors.

   Basic Protocol 1: sgRNA design

   Basic Protocol 2: Cloning of new sgRNA spacers into Mobile‐CRISPRi vectors

   Basic Protocol 3: Tn7transfer of Mobile‐CRISPRi to Gram‐negative bacteria

   Basic Protocol 4: ICEBs1transfer of Mobile‐CRISPRi to Bacillales

   Support Protocol 1: Quantification of CRISPRi repression using fluorescent reporters

   Support Protocol 2: Testing for gene essentiality using CRISPRi spot assays on plates

   Support Protocol 3: Transformation ofE. coliby electroporation

   Support Protocol 4: Transformation of CaCl₂‐competentE. coli

    

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