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Liu GY, Jouandin P, Bahng RE, Perrimon N, Sabatini DM. An evolutionary mechanism to assimilate new nutrient sensors into the mTORC1 pathway. Nat Commun 2024; 15:2517. [PMID: 38514639 PMCID: PMC10957897 DOI: 10.1038/s41467-024-46680-3] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/15/2023] [Accepted: 03/06/2024] [Indexed: 03/23/2024] Open
Abstract
Animals sense and respond to nutrient availability in their environments, a task coordinated in part by the mTOR complex 1 (mTORC1) pathway. mTORC1 regulates growth in response to nutrients and, in mammals, senses specific amino acids through specialized sensors that bind the GATOR1/2 signaling hub. Given that animals can occupy diverse niches, we hypothesized that the pathway might evolve distinct sensors in different metazoan phyla. Whether such customization occurs, and how the mTORC1 pathway might capture new inputs, is unknown. Here, we identify the Drosophila melanogaster protein Unmet expectations (CG11596) as a species-restricted methionine sensor that directly binds the fly GATOR2 complex in a fashion antagonized by S-adenosylmethionine (SAM). We find that in Dipterans GATOR2 rapidly evolved the capacity to bind Unmet and to thereby repurpose a previously independent methyltransferase as a SAM sensor. Thus, the modular architecture of the mTORC1 pathway allows it to co-opt preexisting enzymes to expand its nutrient sensing capabilities, revealing a mechanism for conferring evolvability on an otherwise conserved system.
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Affiliation(s)
- Grace Y Liu
- Whitehead Institute for Biomedical Research and Massachusetts Institute of Technology, Department of Biology, 455 Main Street, Cambridge, MA, USA.
- Department of Biology, Massachusetts Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA, USA.
- Koch Institute for Integrative Cancer Research and Massachusetts Institute of Technology, Department of Biology, 77 Massachusetts Avenue, Cambridge, MA, USA.
| | - Patrick Jouandin
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA, USA
- Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA
- Institut de Recherche en Cancérologie de Montpellier, Inserm U1194-UM-ICM, Campus Val d'Aurelle, Montpellier, Cedex 5, France
| | - Raymond E Bahng
- Whitehead Institute for Biomedical Research and Massachusetts Institute of Technology, Department of Biology, 455 Main Street, Cambridge, MA, USA
- Department of Biology, Massachusetts Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA, USA
- Koch Institute for Integrative Cancer Research and Massachusetts Institute of Technology, Department of Biology, 77 Massachusetts Avenue, Cambridge, MA, USA
| | - Norbert Perrimon
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA, USA.
- Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA.
| | - David M Sabatini
- Institute of Organic Chemistry and Biochemistry, Flemingovo n. 2, 166 10 Praha 6, Prague, Czech Republic.
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2
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Liu GY, Jouandin P, Bahng RE, Perrimon N, Sabatini DM. An evolutionary mechanism to assimilate new nutrient sensors into the mTORC1 pathway. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.05.25.541239. [PMID: 37292894 PMCID: PMC10245982 DOI: 10.1101/2023.05.25.541239] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/10/2023]
Abstract
Animals must sense and respond to nutrient availability in their local niche. This task is coordinated in part by the mTOR complex 1 (mTORC1) pathway, which regulates growth and metabolism in response to nutrients1-5. In mammals, mTORC1 senses specific amino acids through specialized sensors that act through the upstream GATOR1/2 signaling hub6-8. To reconcile the conserved architecture of the mTORC1 pathway with the diversity of environments that animals can occupy, we hypothesized that the pathway might maintain plasticity by evolving distinct nutrient sensors in different metazoan phyla1,9,10. Whether such customization occurs-and how the mTORC1 pathway might capture new nutrient inputs-is not known. Here, we identify the Drosophila melanogaster protein Unmet expectations (Unmet, formerly CG11596) as a species-restricted nutrient sensor and trace its incorporation into the mTORC1 pathway. Upon methionine starvation, Unmet binds to the fly GATOR2 complex to inhibit dTORC1. S-adenosylmethionine (SAM), a proxy for methionine availability, directly relieves this inhibition. Unmet expression is elevated in the ovary, a methionine-sensitive niche11, and flies lacking Unmet fail to maintain the integrity of the female germline under methionine restriction. By monitoring the evolutionary history of the Unmet-GATOR2 interaction, we show that the GATOR2 complex evolved rapidly in Dipterans to recruit and repurpose an independent methyltransferase as a SAM sensor. Thus, the modular architecture of the mTORC1 pathway allows it to co-opt preexisting enzymes and expand its nutrient sensing capabilities, revealing a mechanism for conferring evolvability on an otherwise highly conserved system.
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Affiliation(s)
- Grace Y. Liu
- Whitehead Institute for Biomedical Research and Massachusetts Institute of Technology, Department of Biology; 455 Main Street, Cambridge, Massachusetts 02142, USA
- Department of Biology, Massachusetts Institute of Technology; 77 Massachusetts Avenue, Cambridge, Massachusetts 02139, USA
- Koch Institute for Integrative Cancer Research and Massachusetts Institute of Technology, Department of Biology; 77 Massachusetts Avenue, Cambridge, Massachusetts 02139, USA
| | - Patrick Jouandin
- Department of Genetics, Blavatnik Institute, Harvard Medical School; Boston, MA 02115, USA
- Howard Hughes Medical Institute, Harvard Medical School; Boston, MA 02115, USA
- Present address: Institut de Recherche en Cancérologie de Montpellier, Inserm U1194-UM-ICM; Campus Val d’Aurelle, F-34298 Montpellier Cedex 5, France
| | - Raymond E. Bahng
- Whitehead Institute for Biomedical Research and Massachusetts Institute of Technology, Department of Biology; 455 Main Street, Cambridge, Massachusetts 02142, USA
- Department of Biology, Massachusetts Institute of Technology; 77 Massachusetts Avenue, Cambridge, Massachusetts 02139, USA
- Koch Institute for Integrative Cancer Research and Massachusetts Institute of Technology, Department of Biology; 77 Massachusetts Avenue, Cambridge, Massachusetts 02139, USA
| | - Norbert Perrimon
- Department of Genetics, Blavatnik Institute, Harvard Medical School; Boston, MA 02115, USA
- Howard Hughes Medical Institute, Harvard Medical School; Boston, MA 02115, USA
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3
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CRISPR-Cas Genome Editing for Insect Pest Stress Management in Crop Plants. STRESSES 2022. [DOI: 10.3390/stresses2040034] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/13/2022]
Abstract
Global crop yield and food security are being threatened by phytophagous insects. Innovative methods are required to increase agricultural output while reducing reliance on hazardous synthetic insecticides. Using the revolutionary CRISPR-Cas technology to develop insect-resistant plants appears to be highly efficient at lowering production costs and increasing farm profitability. The genomes of both a model insect, Drosophila melanogaster, and major phytophagous insect genera, viz. Spodoptera, Helicoverpa, Nilaparvata, Locusta, Tribolium, Agrotis, etc., were successfully edited by the CRISPR-Cas toolkits. This new method, however, has the ability to alter an insect’s DNA in order to either induce a gene drive or overcome an insect’s tolerance to certain insecticides. The rapid progress in the methodologies of CRISPR technology and their diverse applications show a high promise in the development of insect-resistant plant varieties or other strategies for the sustainable management of insect pests to ensure food security. This paper reviewed and critically discussed the use of CRISPR-Cas genome-editing technology in long-term insect pest management. The emphasis of this review was on the prospective uses of the CRISPR-Cas system for insect stress management in crop production through the creation of genome-edited crop plants or insects. The potential and the difficulties of using CRISPR-Cas technology to reduce pest stress in crop plants were critically examined and discussed.
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4
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Katow H, Vasudevan D, Ryoo HD. Drosophila Unfolded Protein Response (UPR) Assays In Vitro and In Vivo. Methods Mol Biol 2022; 2378:261-277. [PMID: 34985706 PMCID: PMC9764256 DOI: 10.1007/978-1-0716-1732-8_17] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
Wildtype or mutant proteins expressed beyond the capacity of a cell's protein folding system could be detrimental to general cellular function and survival. In response to misfolded protein overload in the endoplasmic reticulum (ER), eukaryotic cells activate the Unfolded Protein Response (UPR) that helps cells restore protein homeostasis in the endoplasmic reticulum (ER). As part of the UPR, cells attenuate general mRNA translation and activate transcription factors that induce stress-responsive gene expression.UPR signaling draws research interest in part because conditions that cause chronic protein misfolding in the ER or those that impair UPR signaling underlie several diseases including neurodegeneration, diabetes, and cancers. Model organisms are frequently employed in the field as the UPR pathways are generally well-conserved throughout phyla. Here, we introduce experimental procedures to detect UPR in Drosophila melanogaster.
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Affiliation(s)
- Hidetaka Katow
- Department of Cell Biology, New York University Grossman School of Medicine, New York, NY, USA
| | - Deepika Vasudevan
- Department of Cell Biology, New York University Grossman School of Medicine, New York, NY, USA
| | - Hyung Don Ryoo
- Department of Cell Biology, New York University Grossman School of Medicine, New York, NY, USA.
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5
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Böttcher R, Schmidts I, Nitschko V, Duric P, Förstemann K. RNA polymerase II is recruited to DNA double-strand breaks for dilncRNA transcription in Drosophila. RNA Biol 2021; 19:68-77. [PMID: 34965182 PMCID: PMC8786327 DOI: 10.1080/15476286.2021.2014694] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/25/2022] Open
Abstract
DNA double-strand breaks are among the most toxic lesions that can occur in a genome and their faithful repair is thus of great importance. Recent findings have uncovered local transcription that initiates at the break and forms a non-coding transcript, called damage-induced long non-coding RNA (dilncRNA), which helps to coordinate the DNA transactions necessary for repair. We provide nascent RNA sequencing-based evidence that RNA polymerase II transcribes the dilncRNA in Drosophila and that this is more efficient for DNA breaks in an intron-containing gene, consistent with the higher damage-induced siRNA levels downstream of an intron. The spliceosome thus stimulates recruitment of RNA polymerase II to the break, rather than merely promoting the annealing of sense and antisense RNA to form the siRNA precursor. In contrast, RNA polymerase III nascent RNA libraries did not contain reads corresponding to the cleaved loci and selective inhibition of RNA polymerase III did not reduce the yield of damage-induced siRNAs. Finally, the damage-induced siRNA density was unchanged downstream of a T8 sequence, which terminates RNA polymerase III transcription. We thus found no evidence for a participation of RNA polymerase III in dilncRNA transcription in cultured Drosophila cells.
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Affiliation(s)
- Romy Böttcher
- Department. Of Biochemistry and Gene Center, Ludwig-Maximilians-Universität München, München, Germany
| | - Ines Schmidts
- Department. Of Biochemistry and Gene Center, Ludwig-Maximilians-Universität München, München, Germany
| | - Volker Nitschko
- Department. Of Biochemistry and Gene Center, Ludwig-Maximilians-Universität München, München, Germany
| | - Petar Duric
- Department. Of Biochemistry and Gene Center, Ludwig-Maximilians-Universität München, München, Germany
| | - Klaus Förstemann
- Department. Of Biochemistry and Gene Center, Ludwig-Maximilians-Universität München, München, Germany
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6
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Zirin J, Bosch J, Viswanatha R, Mohr SE, Perrimon N. State-of-the-art CRISPR for in vivo and cell-based studies in Drosophila. Trends Genet 2021; 38:437-453. [PMID: 34933779 PMCID: PMC9007876 DOI: 10.1016/j.tig.2021.11.006] [Citation(s) in RCA: 21] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/15/2021] [Revised: 11/22/2021] [Accepted: 11/23/2021] [Indexed: 12/31/2022]
Abstract
For more than 100 years, the fruit fly, Drosophila melanogaster, has served as a powerful model organism for biological and biomedical research due to its many genetic and physiological similarities to humans and the availability of sophisticated technologies used to manipulate its genome and genes. The Drosophila research community quickly adopted CRISPR technologies and, in the 8 years since the first clustered regularly interspaced short palindromic repeats (CRISPR) publications in flies, has explored and innovated methods for mutagenesis, precise genome engineering, and beyond. Moreover, the short lifespan and ease of genetics have made Drosophila an ideal testing ground for in vivo applications and refinements of the rapidly evolving set of CRISPR-associated (CRISPR-Cas) tools. Here, we review innovations in delivery of CRISPR reagents, increased efficiency of cutting and homology-directed repair (HDR), and alternatives to standard Cas9-based approaches. While the focus is primarily on in vivo systems, we also describe the role of Drosophila cultured cells as both an indispensable first step in the process of assessing new CRISPR technologies and a platform for genome-wide CRISPR pooled screens.
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Affiliation(s)
- Jonathan Zirin
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA 02115, USA.
| | - Justin Bosch
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA 02115, USA
| | - Raghuvir Viswanatha
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA 02115, USA
| | - Stephanie E Mohr
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA 02115, USA
| | - Norbert Perrimon
- Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA 02115, USA; Howard Hughes Medical Institute, Harvard Medical School, Boston, MA 02115, USA.
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7
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Jeronimo C, Angel A, Nguyen VQ, Kim JM, Poitras C, Lambert E, Collin P, Mellor J, Wu C, Robert F. FACT is recruited to the +1 nucleosome of transcribed genes and spreads in a Chd1-dependent manner. Mol Cell 2021; 81:3542-3559.e11. [PMID: 34380014 DOI: 10.1016/j.molcel.2021.07.010] [Citation(s) in RCA: 32] [Impact Index Per Article: 10.7] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/02/2020] [Revised: 05/25/2021] [Accepted: 07/12/2021] [Indexed: 12/29/2022]
Abstract
The histone chaperone FACT occupies transcribed regions where it plays prominent roles in maintaining chromatin integrity and preserving epigenetic information. How it is targeted to transcribed regions, however, remains unclear. Proposed models include docking on the RNA polymerase II (RNAPII) C-terminal domain (CTD), recruitment by elongation factors, recognition of modified histone tails, and binding partially disassembled nucleosomes. Here, we systematically test these and other scenarios in Saccharomyces cerevisiae and find that FACT binds transcribed chromatin, not RNAPII. Through a combination of high-resolution genome-wide mapping, single-molecule tracking, and mathematical modeling, we propose that FACT recognizes the +1 nucleosome, as it is partially unwrapped by the engaging RNAPII, and spreads to downstream nucleosomes aided by the chromatin remodeler Chd1. Our work clarifies how FACT interacts with genes, suggests a processive mechanism for FACT function, and provides a framework to further dissect the molecular mechanisms of transcription-coupled histone chaperoning.
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Affiliation(s)
- Célia Jeronimo
- Institut de recherches cliniques de Montréal, 110 Avenue des Pins Ouest, Montréal, QC H2W 1R7, Canada
| | - Andrew Angel
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU, UK
| | - Vu Q Nguyen
- Department of Biology, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Jee Min Kim
- Department of Biology, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Christian Poitras
- Institut de recherches cliniques de Montréal, 110 Avenue des Pins Ouest, Montréal, QC H2W 1R7, Canada
| | - Elie Lambert
- Institut de recherches cliniques de Montréal, 110 Avenue des Pins Ouest, Montréal, QC H2W 1R7, Canada
| | - Pierre Collin
- Institut de recherches cliniques de Montréal, 110 Avenue des Pins Ouest, Montréal, QC H2W 1R7, Canada
| | - Jane Mellor
- Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU, UK
| | - Carl Wu
- Department of Biology, Johns Hopkins University, Baltimore, MD 21218, USA; Department of Molecular Biology and Genetics, Johns Hopkins School of Medicine, Baltimore, MD 21287, USA
| | - François Robert
- Institut de recherches cliniques de Montréal, 110 Avenue des Pins Ouest, Montréal, QC H2W 1R7, Canada; Département de Médecine, Faculté de Médecine, Université de Montréal, 2900 Boulevard Édouard-Montpetit, Montréal, QC H3T 1J4, Canada.
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8
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Palazzo A, Marsano RM. Transposable elements: a jump toward the future of expression vectors. Crit Rev Biotechnol 2021; 41:792-808. [PMID: 33622117 DOI: 10.1080/07388551.2021.1888067] [Citation(s) in RCA: 16] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/22/2022]
Abstract
Expression vectors (EVs) are artificial nucleic acid molecules with a modular structure that allows for the transcription of DNA sequences of interest in either cellular or cell-free environments. These vectors have emerged as cross-disciplinary tools with multiple applications in an expanding Life Sciences market. The cis-regulatory sequences (CRSs) that control the transcription in EVs are typically sourced from either viruses or from characterized genes. However, the recent advancement in transposable elements (TEs) technology provides attractive alternatives that may enable a significant improvement in the design of EVs. Commonly known as "jumping genes," due to their ability to move between genetic loci, TEs are constitutive components of both eukaryotic and prokaryotic genomes. TEs harbor native CRSs that allow the regulated transcription of transposition-related genes. However, some TE-related CRSs display striking characteristics, which provides the opportunity to reconsider TEs as lead actors in the design of EVs. In this article, we provide a synopsis of the transcriptional control elements commonly found in EVs together with an extensive discussion of their advantages and limitations. We also highlight the latest findings that may allow for the implementation of TE-derived sequences in the EVs feasible, possibly improving existing vectors. By introducing this new concept of TEs as a source of regulatory sequences, we aim to stimulate a profitable discussion of the potential advantages and benefits of developing a new generation of EVs based on the use of TE-derived control sequences.
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Affiliation(s)
- Antonio Palazzo
- Laboratory of Translational Nanotechnology, "Istituto Tumori Giovanni Paolo II" I.R.C.C.S, Bari, Italy
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9
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Trivedi D. Using CRISPR-Cas9-based genome engineering tools in Drosophila melanogaster. PROGRESS IN MOLECULAR BIOLOGY AND TRANSLATIONAL SCIENCE 2021; 180:85-121. [PMID: 33934839 DOI: 10.1016/bs.pmbts.2021.01.006] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
Drosophila melanogaster has been used as a model organism for over a century. Mutant-based analyses have been used extensively to understand the genetic basis of different cellular processes, including development, neuronal function and diseases. Most of the earlier genetic mutants and specific tools were generated by random insertions and deletion strategies and then mapped to specific genomic loci. Since all genomic regions are not equally accessible to random mutations and insertions, many genes still remain uncharacterized. Low efficiency of targeted genomic manipulation approaches that rely on homologous recombination, and difficulty in generating resources for sequence-specific endonucleases, such as ZFNs (Zinc Finger Nucleases) and TALENs (Transcription Activator-Like Effector Nucleases), could not make these gene targeting techniques very popular. However, recently RNA directed DNA endonucleases, such as CRISPR-Cas, have transformed genome engineering owing to their comparative ease, versatility, and low expense. With the added advantage of preexisting genetic tools, CRISPR-Cas-based manipulations are being extensively used in Drosophila melanogaster and simultaneously being fine-tuned for specific experimental requirements. In this chapter, I will discuss various uses of CRISPR-Cas-based genetic engineering and specific design methods in Drosophila melanogaster. I will summarize various already available tools that are being utilized in conjunction with CRISPR-Cas technology to generate specific genetic manipulation and are being optimized to address specific questions. Finally, I will discuss the future directions of Drosophila genetics research and how CRISPR-Cas can be utilized to target specific questions, addressing which has not been possible thus far.
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Affiliation(s)
- Deepti Trivedi
- National Centre for Biological Sciences-TIFR, Bengaluru, India.
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10
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Nitschko V, Kunzelmann S, Fröhlich T, Arnold GJ, Förstemann K. Trafficking of siRNA precursors by the dsRBD protein Blanks in Drosophila. Nucleic Acids Res 2020; 48:3906-3921. [PMID: 32025726 PMCID: PMC7144943 DOI: 10.1093/nar/gkaa072] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/08/2019] [Revised: 01/21/2020] [Accepted: 02/03/2020] [Indexed: 01/03/2023] Open
Abstract
RNA interference targets aberrant transcripts with cognate small interfering RNAs, which derive from double-stranded RNA precursors. Several functional screens have identified Drosophila blanks/lump (CG10630) as a facilitator of RNAi, yet its molecular function has remained unknown. The protein carries two dsRNA binding domains (dsRBD) and blanks mutant males have a spermatogenesis defect. We demonstrate that blanks selectively boosts RNAi triggered by dsRNA of nuclear origin. Blanks binds dsRNA via its second dsRBD in vitro, shuttles between nucleus and cytoplasm and the abundance of siRNAs arising at many sites of convergent transcription is reduced in blanks mutants. Since features of nascent RNAs - such as introns and transcription beyond the polyA site – contribute to the small RNA pool, we propose that Blanks binds dsRNA formed by cognate nascent RNAs in the nucleus and fosters its export to the cytoplasm for dicing. We refer to the resulting small RNAs as blanks exported siRNAs (bepsiRNAs). While bepsiRNAs were fully dependent on RNA binding to the second dsRBD of blanks in transgenic flies, male fertility was not. This is consistent with a previous report that linked fertility to the first dsRBD of Blanks. The role of blanks in spermatogenesis appears thus unrelated to its role in dsRNA export.
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Affiliation(s)
- Volker Nitschko
- Genzentrum & Department Biochemie, Ludwig-Maximilians-Universität, 81377 München, Germany
| | - Stefan Kunzelmann
- Genzentrum & Department Biochemie, Ludwig-Maximilians-Universität, 81377 München, Germany
| | - Thomas Fröhlich
- Laboratory of Functional Genome Analysis, Ludwig-Maximilians-Universität, 81377 München, Germany
| | - Georg J Arnold
- Laboratory of Functional Genome Analysis, Ludwig-Maximilians-Universität, 81377 München, Germany
| | - Klaus Förstemann
- Genzentrum & Department Biochemie, Ludwig-Maximilians-Universität, 81377 München, Germany
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11
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Kleinschnitz K, Vießmann N, Jordan M, Heidmann SK. Condensin I is required for faithful meiosis in Drosophila males. Chromosoma 2020; 129:141-160. [PMID: 32314039 PMCID: PMC7260282 DOI: 10.1007/s00412-020-00733-w] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/11/2019] [Revised: 02/17/2020] [Accepted: 02/20/2020] [Indexed: 01/03/2023]
Abstract
The heteropentameric condensin complexes play vital roles in the formation and faithful segregation of mitotic chromosomes in eukaryotes. While the different contributions of the two common condensin complexes, condensin I and condensin II, to chromosome morphology and behavior in mitosis have been thoroughly investigated, much less is known about the specific roles of the two complexes during meiotic divisions. In Drosophila melanogaster, faithful mitotic divisions depend on functional condensin I, but not on condensin II. However, meiotic divisions in Drosophila males require functional condensin II subunits. The role of condensin I during male meiosis in Drosophila has been unresolved. Here, we show that condensin I-specific subunits localize to meiotic chromatin in both meiosis I and II during Drosophila spermatogenesis. Live cell imaging reveals defects during meiotic divisions after RNAi-mediated knockdown of condensin I-specific mRNAs. This phenotype correlates with reduced male fertility and an increase in nondisjunction events both in meiosis I and meiosis II. Consistently, a reduction in male fertility was also observed after proteasome-mediated degradation of the condensin I subunit Barren. Taken together, our results demonstrate an essential role of condensin I during male meiosis in Drosophila melanogaster.
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Affiliation(s)
| | - Nina Vießmann
- Lehrstuhl für Genetik, University of Bayreuth, Bayreuth, Germany
| | - Mareike Jordan
- Lehrstuhl für Genetik, University of Bayreuth, Bayreuth, Germany
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
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12
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Bosch JA, Knight S, Kanca O, Zirin J, Yang-Zhou D, Hu Y, Rodiger J, Amador G, Bellen HJ, Perrimon N, Mohr SE. Use of the CRISPR-Cas9 System in Drosophila Cultured Cells to Introduce Fluorescent Tags into Endogenous Genes. CURRENT PROTOCOLS IN MOLECULAR BIOLOGY 2020; 130:e112. [PMID: 31869524 PMCID: PMC7213786 DOI: 10.1002/cpmb.112] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 01/08/2023]
Abstract
The CRISPR-Cas9 system makes it possible to cause double-strand breaks in specific regions, inducing repair. In the presence of a donor construct, repair can involve insertion or 'knock-in' of an exogenous cassette. One common application of knock-in technology is to generate cell lines expressing fluorescently tagged endogenous proteins. The standard approach relies on production of a donor plasmid with ∼500 to 1000 bp of homology on either side of an insertion cassette that contains the fluorescent protein open reading frame (ORF). We present two alternative methods for knock-in of fluorescent protein ORFs into Cas9-expressing Drosophila S2R+ cultured cells, the single-stranded DNA (ssDNA) Drop-In method and the CRISPaint universal donor method. Both methods eliminate the need to clone a large plasmid donor for each target. We discuss the advantages and limitations of the standard, ssDNA Drop-In, and CRISPaint methods for fluorescent protein tagging in Drosophila cultured cells. © 2019 by John Wiley & Sons, Inc. Basic Protocol 1: Knock-in into Cas9-positive S2R+ cells using the ssDNA Drop-In approach Basic Protocol 2: Knock-in into Cas9-positive S2R+ cells by homology-independent insertion of universal donor plasmids that provide mNeonGreen (CRISPaint method) Support Protocol 1: sgRNA design and cloning Support Protocol 2: ssDNA donor synthesis Support Protocol 3: Transfection using Effectene Support Protocol 4: Electroporation of S2R+-MT::Cas9 Drosophila cells Support Protocol 5: Single-cell isolation of fluorescent cells using FACS.
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Affiliation(s)
- Justin A Bosch
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
| | - Shannon Knight
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Oguz Kanca
- Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas
- Jan and Dan Duncan Neurological Research Institute, Texas Children's Hospital, Houston, Texas
| | - Jonathan Zirin
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Donghui Yang-Zhou
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Yanhui Hu
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Jonathan Rodiger
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Gabriel Amador
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
| | - Hugo J Bellen
- Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas
- Jan and Dan Duncan Neurological Research Institute, Texas Children's Hospital, Houston, Texas
- Department of Neuroscience, Baylor College of Medicine, Houston, Texas
- Howard Hughes Medical Institute, Baylor College of Medicine, Houston, Texas
| | - Norbert Perrimon
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
- Howard Hughes Medical Institute, Harvard Medical School, Boston, Massachusetts
| | - Stephanie E Mohr
- Department of Genetics, Harvard Medical School, Boston, Massachusetts
- Drosophila RNAi Screening Center, Harvard Medical School, Boston, Massachusetts
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Kanca O, Zirin J, Garcia-Marques J, Knight SM, Yang-Zhou D, Amador G, Chung H, Zuo Z, Ma L, He Y, Lin WW, Fang Y, Ge M, Yamamoto S, Schulze KL, Hu Y, Spradling AC, Mohr SE, Perrimon N, Bellen HJ. An efficient CRISPR-based strategy to insert small and large fragments of DNA using short homology arms. eLife 2019; 8:e51539. [PMID: 31674908 PMCID: PMC6855806 DOI: 10.7554/elife.51539] [Citation(s) in RCA: 82] [Impact Index Per Article: 16.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/09/2019] [Accepted: 10/31/2019] [Indexed: 12/31/2022] Open
Abstract
We previously reported a CRISPR-mediated knock-in strategy into introns of Drosophila genes, generating an attP-FRT-SA-T2A-GAL4-polyA-3XP3-EGFP-FRT-attP transgenic library for multiple uses (Lee et al., 2018a). The method relied on double stranded DNA (dsDNA) homology donors with ~1 kb homology arms. Here, we describe three new simpler ways to edit genes in flies. We create single stranded DNA (ssDNA) donors using PCR and add 100 nt of homology on each side of an integration cassette, followed by enzymatic removal of one strand. Using this method, we generated GFP-tagged proteins that mark organelles in S2 cells. We then describe two dsDNA methods using cheap synthesized donors flanked by 100 nt homology arms and gRNA target sites cloned into a plasmid. Upon injection, donor DNA (1 to 5 kb) is released from the plasmid by Cas9. The cassette integrates efficiently and precisely in vivo. The approach is fast, cheap, and scalable.
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Affiliation(s)
- Oguz Kanca
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
| | - Jonathan Zirin
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | | | - Shannon Marie Knight
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Donghui Yang-Zhou
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Gabriel Amador
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Hyunglok Chung
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
| | - Zhongyuan Zuo
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
| | - Liwen Ma
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
| | - Yuchun He
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Howard Hughes Medical Institute, Baylor College of MedicineHoustonUnited States
| | - Wen-Wen Lin
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
| | - Ying Fang
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
| | - Ming Ge
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
| | - Shinya Yamamoto
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
- Program in Developmental BiologyBaylor College of MedicineHoustonUnited States
- Department of NeuroscienceBaylor College of MedicineHoustonUnited States
| | - Karen L Schulze
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
- Howard Hughes Medical Institute, Baylor College of MedicineHoustonUnited States
| | - Yanhui Hu
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Allan C Spradling
- Department of EmbryologyHoward Hughes Medical Institute, Carnegie Institution for ScienceBaltimoreUnited States
| | - Stephanie E Mohr
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Norbert Perrimon
- Howard Hughes Medical Institute, Harvard Medical SchoolBostonUnited States
- Department of GeneticsHarvard Medical SchoolBostonUnited States
| | - Hugo J Bellen
- Department of Molecular and Human GeneticsBaylor College of MedicineHoustonUnited States
- Jan and Dan Duncan Neurological Research Institute, Texas Children’s HospitalHoustonUnited States
- Howard Hughes Medical Institute, Baylor College of MedicineHoustonUnited States
- Program in Developmental BiologyBaylor College of MedicineHoustonUnited States
- Department of NeuroscienceBaylor College of MedicineHoustonUnited States
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14
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Luhur A, Klueg KM, Roberts J, Zelhof AC. Thawing, Culturing, and Cryopreserving Drosophila Cell Lines. J Vis Exp 2019:10.3791/59459. [PMID: 31058891 PMCID: PMC7032961 DOI: 10.3791/59459] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/14/2022] Open
Abstract
There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.
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Affiliation(s)
- Arthur Luhur
- Drosophila Genomics Resource Center, Department of Biology, Indiana University Bloomington;
| | - Kristin M Klueg
- Drosophila Genomics Resource Center, Department of Biology, Indiana University Bloomington
| | - Johnny Roberts
- Drosophila Genomics Resource Center, Department of Biology, Indiana University Bloomington
| | - Andrew C Zelhof
- Drosophila Genomics Resource Center, Department of Biology, Indiana University Bloomington
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15
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Osswald M, Santos AF, Morais-de-Sá E. Light-Induced Protein Clustering for Optogenetic Interference and Protein Interaction Analysis in Drosophila S2 Cells. Biomolecules 2019; 9:biom9020061. [PMID: 30759894 PMCID: PMC6406598 DOI: 10.3390/biom9020061] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/11/2018] [Revised: 01/30/2019] [Accepted: 01/30/2019] [Indexed: 12/19/2022] Open
Abstract
Drosophila Schneider 2 (S2) cells are a simple and powerful system commonly used in cell biology because they are well suited for high resolution microscopy and RNAi-mediated depletion. However, understanding dynamic processes, such as cell division, also requires methodology to interfere with protein function with high spatiotemporal control. In this research study, we report the adaptation of an optogenetic tool to Drosophila S2 cells. Light-activated reversible inhibition by assembled trap (LARIAT) relies on the rapid light-dependent heterodimerization between cryptochrome 2 (CRY2) and cryptochrome-interacting bHLH 1 (CIB1) to form large protein clusters. An anti-green fluorescent protein (GFP) nanobody fused with CRY2 allows this method to quickly trap any GFP-tagged protein in these light-induced protein clusters. We evaluated clustering kinetics in response to light for different LARIAT modules, and showed the ability of GFP-LARIAT to inactivate the mitotic protein Mps1 and to disrupt the membrane localization of the polarity regulator Lethal Giant Larvae (Lgl). Moreover, we validated light-induced co-clustering assays to assess protein-protein interactions in S2 cells. In conclusion, GFP-based LARIAT is a versatile tool to answer different biological questions, since it enables probing of dynamic processes and protein-protein interactions with high spatiotemporal resolution in Drosophila S2 cells.
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Affiliation(s)
- Mariana Osswald
- Epithelial Polarity and Cell Division, i3S, Instituto de Investigação e Inovação em Saúde, Universidade do Porto, 4200-135 Porto, Portugal.
- IBMC, Instituto de Biologia Molecular e Celular, Universidade do Porto, 4200-135 Porto, Portugal.
| | - A Filipa Santos
- Epithelial Polarity and Cell Division, i3S, Instituto de Investigação e Inovação em Saúde, Universidade do Porto, 4200-135 Porto, Portugal.
- IBMC, Instituto de Biologia Molecular e Celular, Universidade do Porto, 4200-135 Porto, Portugal.
| | - Eurico Morais-de-Sá
- Epithelial Polarity and Cell Division, i3S, Instituto de Investigação e Inovação em Saúde, Universidade do Porto, 4200-135 Porto, Portugal.
- IBMC, Instituto de Biologia Molecular e Celular, Universidade do Porto, 4200-135 Porto, Portugal.
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16
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Supharattanasitthi W, Carlsson E, Sharif U, Paraoan L. CRISPR/Cas9-mediated one step bi-allelic change of genomic DNA in iPSCs and human RPE cells in vitro with dual antibiotic selection. Sci Rep 2019; 9:174. [PMID: 30655567 PMCID: PMC6336765 DOI: 10.1038/s41598-018-36740-2] [Citation(s) in RCA: 22] [Impact Index Per Article: 4.4] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/30/2018] [Accepted: 11/24/2018] [Indexed: 12/27/2022] Open
Abstract
CRISPR/Cas9 causes double-stranded DNA breaks that can undergo DNA repair either via non-homologous end joining (NHEJ) or, in the presence of a template, homology-directed repair (HDR). HDR is typically used to insert a specific genetic modification into the genome but has low efficiency compared to NHEJ, which is lowered even further when trying to create a homozygous change. In this study we devised a novel approach for homozygous single base editing based on utilising simultaneously two donor DNA templates cloned in plasmids with different antibiotic resistant genes. The donor templates were co-transfected alongside the CRISPR/Cas9 machinery into cells and a double antibiotic selection was optimised and allowed the isolation of viable desired clones. We applied the method for obtaining isogenic cells homozygous for variant B cystatin C, a recessive risk factor for age-related macular degeneration and Alzheimer's disease, in both induced Pluripotent Stem Cells (iPSCs) and a human RPE cell line. Bi-allelic gene edited clones were validated by sequencing, demonstrating that the double antibiotic templates approach worked efficiently for both iPSCs and human differentiated cells. We propose that this one step gene editing approach can be used to improve the specificity and frequency of introducing homozygous modifications in mammalian cells.
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Affiliation(s)
- Wasu Supharattanasitthi
- Department of Eye and Vision Science, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, United Kingdom
- Department of Physiology, Faculty of Pharmacy, Mahidol University, Bangkok, Thailand
| | - Emil Carlsson
- Department of Eye and Vision Science, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, United Kingdom
| | - Umar Sharif
- Department of Eye and Vision Science, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, United Kingdom
| | - Luminita Paraoan
- Department of Eye and Vision Science, Institute of Ageing and Chronic Disease, University of Liverpool, Liverpool, United Kingdom.
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17
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Luhur A, Klueg KM, Zelhof AC. Generating and working with Drosophila cell cultures: Current challenges and opportunities. WILEY INTERDISCIPLINARY REVIEWS-DEVELOPMENTAL BIOLOGY 2018; 8:e339. [PMID: 30561900 DOI: 10.1002/wdev.339] [Citation(s) in RCA: 18] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 07/11/2018] [Revised: 10/30/2018] [Accepted: 11/21/2018] [Indexed: 12/26/2022]
Abstract
The use of Drosophila cell cultures has positively impacted both fundamental and biomedical research. The most widely used cell lines: Schneider, Kc, the CNS and imaginal disc lines continue to be the choice for many applications. Drosophila cell lines provide a homogenous source of cells suitable for biochemical experimentations, transcriptomics, functional genomics, and biomedical applications. They are amenable to RNA interference and serve as a platform for high-throughput screens to identify relevant candidate genes or drugs for any biological process. Currently, CRISPR-based functional genomics are also being developed for Drosophila cell lines. Even though many uniquely derived cell lines exist, cell genetic techniques such the transgenic UAS-GAL4-based RasV12 oncogene expression, CRISPR-Cas9 editing and recombination mediated cassette exchange are likely to drive the establishment of many more lines from specific tissues, cells, or genotypes. However, the pace of creating new lines is hindered by several factors inherent to working with Drosophila cell cultures: single cell cloning, optimal media formulations and culture conditions capable of supporting lines from novel tissue sources or genotypes. Moreover, even though many Drosophila cell lines are morphologically and transcriptionally distinct it may be necessary to implement a standard for Drosophila cell line authentication, ensuring the identity and purity of each cell line. Altogether, recent advances and a standardized authentication effort should improve the utility of Drosophila cell cultures as a relevant model for fundamental and biomedical research. This article is categorized under: Technologies > Analysis of Cell, Tissue, and Animal Phenotypes.
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Affiliation(s)
- Arthur Luhur
- Department of Biology, Drosophila Genomics Resource Center, Indiana University Bloomington, Bloomington, Indiana
| | - Kristin M Klueg
- Department of Biology, Drosophila Genomics Resource Center, Indiana University Bloomington, Bloomington, Indiana
| | - Andrew C Zelhof
- Department of Biology, Drosophila Genomics Resource Center, Indiana University Bloomington, Bloomington, Indiana
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18
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Bier E, Harrison MM, O'Connor-Giles KM, Wildonger J. Advances in Engineering the Fly Genome with the CRISPR-Cas System. Genetics 2018; 208:1-18. [PMID: 29301946 PMCID: PMC5753851 DOI: 10.1534/genetics.117.1113] [Citation(s) in RCA: 115] [Impact Index Per Article: 19.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/09/2017] [Accepted: 07/08/2017] [Indexed: 12/26/2022] Open
Abstract
Drosophila has long been a premier model for the development and application of cutting-edge genetic approaches. The CRISPR-Cas system now adds the ability to manipulate the genome with ease and precision, providing a rich toolbox to interrogate relationships between genotype and phenotype, to delineate and visualize how the genome is organized, to illuminate and manipulate RNA, and to pioneer new gene drive technologies. Myriad transformative approaches have already originated from the CRISPR-Cas system, which will likely continue to spark the creation of tools with diverse applications. Here, we provide an overview of how CRISPR-Cas gene editing has revolutionized genetic analysis in Drosophila and highlight key areas for future advances.
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Affiliation(s)
- Ethan Bier
- Cell and Developmental Biology, University of California, San Diego, La Jolla, California 92093-0349
| | - Melissa M Harrison
- Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, Wisconsin 53706
| | - Kate M O'Connor-Giles
- Laboratory of Genetics and Laboratory of Cell and Molecular Biology, Wisconsin 53706
| | - Jill Wildonger
- Biochemistry Department, University of Wisconsin-Madison, Wisconsin 53706
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19
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Korona D, Koestler SA, Russell S. Engineering the Drosophila Genome for Developmental Biology. J Dev Biol 2017; 5:jdb5040016. [PMID: 29615571 PMCID: PMC5831791 DOI: 10.3390/jdb5040016] [Citation(s) in RCA: 16] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/30/2017] [Revised: 12/07/2017] [Accepted: 12/08/2017] [Indexed: 02/07/2023] Open
Abstract
The recent development of transposon and CRISPR-Cas9-based tools for manipulating the fly genome in vivo promises tremendous progress in our ability to study developmental processes. Tools for introducing tags into genes at their endogenous genomic loci facilitate imaging or biochemistry approaches at the cellular or subcellular levels. Similarly, the ability to make specific alterations to the genome sequence allows much more precise genetic control to address questions of gene function.
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Affiliation(s)
- Dagmara Korona
- Department of Genetics, University of Cambridge, Downing Street, Cambridge CB2 3EH, UK.
| | - Stefan A Koestler
- Department of Genetics, University of Cambridge, Downing Street, Cambridge CB2 3EH, UK.
| | - Steven Russell
- Department of Genetics, University of Cambridge, Downing Street, Cambridge CB2 3EH, UK.
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20
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Sun D, Guo Z, Liu Y, Zhang Y. Progress and Prospects of CRISPR/Cas Systems in Insects and Other Arthropods. Front Physiol 2017; 8:608. [PMID: 28932198 PMCID: PMC5592444 DOI: 10.3389/fphys.2017.00608] [Citation(s) in RCA: 90] [Impact Index Per Article: 12.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/30/2017] [Accepted: 08/07/2017] [Indexed: 01/03/2023] Open
Abstract
Clustered regularly interspaced short palindromic repeats (CRISPR) and the CRISPR-associated gene Cas9 represent an invaluable system for the precise editing of genes in diverse species. The CRISPR/Cas9 system is an adaptive mechanism that enables bacteria and archaeal species to resist invading viruses and phages or plasmids. Compared with zinc finger nucleases and transcription activator-like effector nucleases, the CRISPR/Cas9 system has the advantage of requiring less time and effort. This efficient technology has been used in many species, including diverse arthropods that are relevant to agriculture, forestry, fisheries, and public health; however, there is no review that systematically summarizes its successful application in the editing of both insect and non-insect arthropod genomes. Thus, this paper seeks to provide a comprehensive and impartial overview of the progress of the CRISPR/Cas9 system in different arthropods, reviewing not only fundamental studies related to gene function exploration and experimental optimization but also applied studies in areas such as insect modification and pest control. In addition, we also describe the latest research advances regarding two novel CRISPR/Cas systems (CRISPR/Cpf1 and CRISPR/C2c2) and discuss their future prospects for becoming crucial technologies in arthropods.
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Affiliation(s)
- Dan Sun
- Longping Branch, Graduate School of Hunan UniversityChangsha, China.,Department of Plant Protection, Institute of Vegetables and Flowers, Chinese Academy of Agricultural SciencesBeijing, China
| | - Zhaojiang Guo
- Department of Plant Protection, Institute of Vegetables and Flowers, Chinese Academy of Agricultural SciencesBeijing, China
| | - Yong Liu
- Longping Branch, Graduate School of Hunan UniversityChangsha, China
| | - Youjun Zhang
- Department of Plant Protection, Institute of Vegetables and Flowers, Chinese Academy of Agricultural SciencesBeijing, China
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21
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Abstract
The ease of generating genetically modified animals and cell lines has been markedly increased by the recent development of the versatile CRISPR/Cas9 tool. However, while the isolation of isogenic cell populations is usually straightforward for mammalian cell lines, the generation of clonal Drosophila cell lines has remained a longstanding challenge, hampered by the difficulty of getting Drosophila cells to grow at low densities. Here, we describe a highly efficient workflow to generate clonal Cas9-engineered Drosophila cell lines using a combination of cell pools, limiting dilution in conditioned medium and PCR with allele-specific primers, enabling the efficient selection of a clonal cell line with a suitable mutation profile. We validate the protocol by documenting the isolation, selection and verification of eight independently Cas9-edited armadillo mutant Drosophila cell lines. Our method provides a powerful and simple workflow that improves the utility of Drosophila cells for genetic studies with CRISPR/Cas9.
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Affiliation(s)
- Alexandra Franz
- a Institute of Molecular Life Sciences, University of Zurich , Zurich , Switzerland
| | - Erich Brunner
- a Institute of Molecular Life Sciences, University of Zurich , Zurich , Switzerland
| | - Konrad Basler
- a Institute of Molecular Life Sciences, University of Zurich , Zurich , Switzerland
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22
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Kunzelmann S, Förstemann K. Reversible perturbations of gene regulation after genome editing in Drosophila cells. PLoS One 2017; 12:e0180135. [PMID: 28658280 PMCID: PMC5489201 DOI: 10.1371/journal.pone.0180135] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/18/2017] [Accepted: 06/09/2017] [Indexed: 12/04/2022] Open
Abstract
The prokaryotic phage defense CRISPR/cas-system has developed into a versatile toolbox for genome engineering and genetic studies in many organisms. While many efforts were spent on analyzing the consequences of off-target effects, only few studies addressed side-effects that occur due to the targeted manipulation of the genome. Here, we show that the CRISPR/cas9-mediated integration of an epitope tag in combination with a selection cassette can trigger an siRNA-mediated, epigenetic genome surveillance pathway in Drosophila melanogaster cells. After homology-directed insertion of the sequence coding for the epitope tag and the selection marker, a moderate level of siRNAs covering the inserted sequence and extending into the targeted locus was detected. This response affected protein levels less than two-fold and it persisted even after single cell cloning. However, removal of the selection cassette abolished the siRNA generation, demonstrating that this response is reversible. Consistently, marker-free genome engineering did not trigger the same surveillance mechanism. These two observations indicate that the selection cassette we employed induces an aberrant transcriptional arrangement and ultimately sets off the siRNA production. There have been prior concerns about undesirable effects induced by selection markers, but fortunately we were able to show that at least one of the epigenetic changes reverts as the marker gene is excised. Although the effects observed were rather weak (less than twofold de-repression upon ago2 or dcr-2 knock-down), we recommend that when selection markers are used during genome editing, a strategy for their subsequent removal should always be included.
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Affiliation(s)
- Stefan Kunzelmann
- Department of Biochemistry, Gene Center, Ludwig-Maximilians-Universität München, München, Germany
| | - Klaus Förstemann
- Department of Biochemistry, Gene Center, Ludwig-Maximilians-Universität München, München, Germany
- * E-mail:
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23
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Wissel S, Kieser A, Yasugi T, Duchek P, Roitinger E, Gokcezade J, Steinmann V, Gaul U, Mechtler K, Förstemann K, Knoblich JA, Neumüller RA. A Combination of CRISPR/Cas9 and Standardized RNAi as a Versatile Platform for the Characterization of Gene Function. G3 (BETHESDA, MD.) 2016; 6:2467-78. [PMID: 27280787 PMCID: PMC4978900 DOI: 10.1534/g3.116.028571] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 03/01/2016] [Accepted: 05/31/2016] [Indexed: 12/21/2022]
Abstract
Traditional loss-of-function studies in Drosophila suffer from a number of shortcomings, including off-target effects in the case of RNA interference (RNAi) or the stochastic nature of mosaic clonal analysis. Here, we describe minimal in vivo GFP interference (miGFPi) as a versatile strategy to characterize gene function and to conduct highly stringent, cell type-specific loss-of-function experiments in Drosophila miGFPi combines CRISPR/Cas9-mediated tagging of genes at their endogenous locus with an immunotag and an exogenous 21 nucleotide RNAi effector sequence with the use of a single reagent, highly validated RNAi line targeting this sequence. We demonstrate the utility and time effectiveness of this method by characterizing the function of the Polymerase I (Pol I)-associated transcription factor Tif-1a, and the previously uncharacterized gene MESR4, in the Drosophila female germline stem cell lineage. In addition, we show that miGFPi serves as a powerful technique to functionally characterize individual isoforms of a gene. We exemplify this aspect of miGFPi by studying isoform-specific loss-of-function phenotypes of the longitudinals lacking (lola) gene in neural stem cells. Altogether, the miGFPi strategy constitutes a generalized loss-of-function approach that is amenable to the study of the function of all genes in the genome in a stringent and highly time effective manner.
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Affiliation(s)
- Sebastian Wissel
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria
| | - Anja Kieser
- Gene Center, Ludwig-Maximilians-University Munich, 81377, Germany
| | - Tetsuo Yasugi
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria
| | - Peter Duchek
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria
| | - Elisabeth Roitinger
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria Institute of Molecular Pathology, 1030 Vienna, Austria
| | - Joseph Gokcezade
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria
| | | | - Ulrike Gaul
- Gene Center, Ludwig-Maximilians-University Munich, 81377, Germany
| | - Karl Mechtler
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria Institute of Molecular Pathology, 1030 Vienna, Austria
| | - Klaus Förstemann
- Gene Center, Ludwig-Maximilians-University Munich, 81377, Germany
| | | | - Ralph A Neumüller
- Institute of Molecular Biotechnology Austria, 1030 Vienna, Austria Gene Center, Ludwig-Maximilians-University Munich, 81377, Germany
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24
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Schmidts I, Böttcher R, Mirkovic-Hösle M, Förstemann K. Homology directed repair is unaffected by the absence of siRNAs in Drosophila melanogaster. Nucleic Acids Res 2016; 44:8261-71. [PMID: 27353331 PMCID: PMC5041469 DOI: 10.1093/nar/gkw570] [Citation(s) in RCA: 20] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/02/2015] [Accepted: 06/14/2016] [Indexed: 12/28/2022] Open
Abstract
Small interfering RNAs (siRNAs) defend the organism against harmful transcripts from exogenous (e.g. viral) or endogenous (e.g. transposons) sources. Recent publications describe the production of siRNAs induced by DNA double-strand breaks (DSB) in Neurospora crassa, Arabidopsis thaliana, Drosophila melanogaster and human cells, which suggests a conserved function. A current hypothesis is that break-induced small RNAs ensure efficient homologous recombination (HR). However, biogenesis of siRNAs is often intertwined with other small RNA species, such as microRNAs (miRNAs), which complicates interpretation of experimental results. In Drosophila, siRNAs are produced by Dcr-2 while miRNAs are processed by Dcr-1. Thus, it is possible to probe siRNA function without miRNA deregulation. We therefore examined DNA double-strand break repair after perturbation of siRNA biogenesis in cultured Drosophila cells as well as mutant flies. Our assays comprised reporters for the single-strand annealing pathway, homologous recombination and sensitivity to the DSB-inducing drug camptothecin. We could not detect any repair defects caused by the lack of siRNAs derived from the broken DNA locus. Since production of these siRNAs depends on local transcription, they may thus participate in RNA metabolism-an established function of siRNAs-rather than DNA repair.
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Affiliation(s)
- Ines Schmidts
- Gene Center and Department of Biochemistry, Ludwig-Maximilians-Universität München, Feodor-Lynen-Straße 25, D-81377, München, Germany
| | - Romy Böttcher
- Gene Center and Department of Biochemistry, Ludwig-Maximilians-Universität München, Feodor-Lynen-Straße 25, D-81377, München, Germany
| | - Milijana Mirkovic-Hösle
- Gene Center and Department of Biochemistry, Ludwig-Maximilians-Universität München, Feodor-Lynen-Straße 25, D-81377, München, Germany
| | - Klaus Förstemann
- Gene Center and Department of Biochemistry, Ludwig-Maximilians-Universität München, Feodor-Lynen-Straße 25, D-81377, München, Germany
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