Scott J. Gratz

Genome engineering of Drosophila with the CRISPR RNA-guided Cas9 nuclease

We have adapted a bacterial CRISPR RNA/Cas9 system to precisely engineer the Drosophila genome and report that Cas9-mediated genomic modifications are efficiently transmitted through the germline. This RNA-guided Cas9 system can be rapidly programmed to generate targeted alleles for probing gene function in Drosophila.

Highly Specific and Efficient CRISPR/Cas9-Catalyzed Homology-Directed Repair in Drosophila

We and others recently demonstrated that the readily programmable CRISPR/Cas9 system can be used to edit the Drosophila genome. However, most applications to date have relied on aberrant DNA repair to stochastically generate frameshifting indels and adoption has been limited by a lack of tools for efficient identification of targeted events. Here we report optimized tools and techniques for expanded application of the CRISPR/Cas9 system in Drosophila through homology-directed repair (HDR) with double-stranded DNA (dsDNA) donor templates that facilitate complex genome engineering through the precise incorporation of large DNA sequences, including screenable markers. Using these donors, we demonstrate the replacement of a gene with exogenous sequences and the generation of a conditional allele. To optimize efficiency and specificity, we generated transgenic flies that express Cas9 in the germline and directly compared HDR and off-target cleavage rates of different approaches for delivering CRISPR components. We also investigated HDR efficiency in a mutant background previously demonstrated to bias DNA repair toward HDR. Finally, we developed a web-based tool that identifies CRISPR target sites and evaluates their potential for off-target cleavage using empirically rooted rules. Overall, we have found that injection of a dsDNA donor and guide RNA-encoding plasmids into vasa-Cas9 flies yields the highest efficiency HDR and that target sites can be selected to avoid off-target mutations. Efficient and specific CRISPR/Cas9-mediated HDR opens the door to a broad array of complex genome modifications and greatly expands the utility of CRISPR technology for Drosophila research.

Precise Genome Editing of Drosophila with CRISPR RNA-Guided Cas9

The readily programmable CRISPR-Cas9 system is transforming genome engineering. We and others have adapted the S. pyogenes CRISPR-Cas9 system to precisely engineer the Drosophila genome and demonstrated that these modifications are efficiently transmitted through the germline. Here we provide a detailed protocol for engineering small indels, defined deletions, and targeted insertion of exogenous DNA sequences within one month using a rapid DNA injection-based approach.

Safeguarding gene drive experiments in the laboratory

Multiple stringent confinement strategies should be used whenever possible Gene drive systems promote the spread of genetic elements through populations by assuring they are inherited more often than Mendelian segregation would predict (see the figure). Natural examples of gene drive from Drosophila include sex-ratio meiotic drive, segregation distortion, and replicative transposition. Synthetic drive systems based on selective embryonic lethality or homing endonucleases have been described previously in Drosophila melanogaster (1–3), but they are difficult to build or are limited to transgenic populations. In contrast, RNAguided gene drives based on the CRISPR/Cas9 nuclease can, in principle, be constructed by any laboratory capable of making transgenic organisms (4). They have tremendous potential to address global problems in health, agriculture, and conservation, but their capacity to alter wild populations outside the laboratory demands caution (4–7). Just as researchers working with self-propagating pathogens must ensure that these agents do not escape to the outside world, scientists working in the laboratory with gene drive constructs are responsible for keeping them confined (4, 6, 7).

CRISPR/Cas9-mediated genome engineering and the promise of designer flies on demand.

The CRISPR/Cas9 system has attracted significant attention for its potential to transform genome engineering. We and others have recently shown that the RNA-guided Cas9 nuclease can be employed to engineer the Drosophila genome, and that these modifications are efficiently transmitted through the germline. A single targeting RNA can guide Cas9 to a specific genomic sequence where it induces double-strand breaks that, when imperfectly repaired, yield mutations. We have also demonstrated that 2 targeting RNAs can be used to generate large defined deletions and that Cas9 can catalyze gene replacement by homologous recombination. Zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs) have shown similar promise in Drosophila. However, the ease of producing targeting RNAs over the generation of unique sequence-directed nucleases to guide site-specific modifications makes the CRISPR/Cas9 system an appealingly accessible method for genome editing. From the initial planning stages, engineered flies can be obtained within a month. Here we highlight the variety of genome modifications facilitated by the CRISPR/Cas9 system along with key considerations for starting your own CRISPR genome engineering project.

CRISPR‐Cas9 Genome Editing in Drosophila

The CRISPR‐Cas9 system has transformed genome engineering of model organisms from possible to practical. CRISPR‐Cas9 can be readily programmed to generate sequence‐specific double‐strand breaks that disrupt targeted loci when repaired by error‐prone non‐homologous end joining (NHEJ) or to catalyze precise genome modification through homology‐directed repair (HDR). Here we describe a streamlined approach for rapid and highly efficient engineering of the Drosophila genome via CRISPR‐Cas9‐mediated HDR. In this approach, transgenic flies expressing Cas9 are injected with plasmids to express guide RNAs (gRNAs) and positively marked donor templates. We detail target‐site selection; gRNA plasmid generation; donor template design and construction; and the generation, identification, and molecular confirmation of engineered lines. We also present alternative approaches and highlight key considerations for experimental design. The approach outlined here can be used to rapidly and reliably generate a variety of engineered modifications, including genomic deletions and replacements, precise sequence edits, and incorporation of protein tags.

Fife, a Drosophila Piccolo-RIM homolog, promotes active zone organization and neurotransmitter release.

Neuronal communication depends on the precisely orchestrated release of neurotransmitter at specialized sites called active zones (AZs). A small number of scaffolding and cytoskeletal proteins comprising the cytomatrix of the active zone (CAZ) are thought to organize the architecture and functional properties of AZs. The majority of CAZ proteins are evolutionarily conserved, underscoring the fundamental similarities in neurotransmission at all synapses. However, core CAZ proteins Piccolo and Bassoon have long been believed exclusive to vertebrates, raising intriguing questions about the conservation of the molecular mechanisms that regulate presynaptic properties. Here, we present the identification of a piccolo-rim-related gene in invertebrates, together with molecular phylogenetic analyses that indicate the encoded proteins may represent Piccolo orthologs. In accordance, we find that the Drosophila homolog, Fife, is neuronal and localizes to presynaptic AZs. To investigate the in vivo function of Fife, we generated a deletion of the fife locus. We find that evoked neurotransmitter release is substantially decreased in fife mutants and loss of fife results in motor deficits. Through morphological analysis of fife synapses, we identify underlying AZ abnormalities including pervasive presynaptic membrane detachments and reduced synaptic vesicle clustering. Our data demonstrate the conservation of a Piccolo-related protein in invertebrates and identify critical roles for Fife in regulating AZ structure and function. These findings suggest the CAZ is more conserved than previously thought, and open the door to a more complete understanding of how CAZ proteins regulate presynaptic structure and function through genetic studies in simpler model systems.

BIOSAFETY. Safeguarding gene drive experiments in the laboratory.

Multiple stringent confinement strategies should be used whenever possible Gene drive systems promote the spread of genetic elements through populations by assuring they are inherited more often than Mendelian segregation would predict (see the figure). Natural examples of gene drive from Drosophila include sex-ratio meiotic drive, segregation distortion, and replicative transposition. Synthetic drive systems based on selective embryonic lethality or homing endonucleases have been described previously in Drosophila melanogaster (1–3), but they are difficult to build or are limited to transgenic populations. In contrast, RNAguided gene drives based on the CRISPR/Cas9 nuclease can, in principle, be constructed by any laboratory capable of making transgenic organisms (4). They have tremendous potential to address global problems in health, agriculture, and conservation, but their capacity to alter wild populations outside the laboratory demands caution (4–7). Just as researchers working with self-propagating pathogens must ensure that these agents do not escape to the outside world, scientists working in the laboratory with gene drive constructs are responsible for keeping them confined (4, 6, 7).

Fife organizes synaptic vesicles and calcium channels for high-probability neurotransmitter release

The strength of synaptic connections varies significantly and is a key determinant of communication within neural circuits. Mechanistic insight into presynaptic factors that establish and modulate neurotransmitter release properties is crucial to understanding synapse strength, circuit function, and neural plasticity. We previously identified Drosophila Piccolo-RIM-related Fife, which regulates neurotransmission and motor behavior through an unknown mechanism. Here, we demonstrate that Fife localizes and interacts with RIM at the active zone cytomatrix to promote neurotransmitter release. Loss of Fife results in the severe disruption of active zone cytomatrix architecture and molecular organization. Through electron tomographic and electrophysiological studies, we find a decrease in the accumulation of release-ready synaptic vesicles and their release probability caused by impaired coupling to Ca2+ channels. Finally, we find that Fife is essential for the homeostatic modulation of neurotransmission. We propose that Fife organizes active zones to create synaptic vesicle release sites within nanometer distance of Ca2+ channel clusters for reliable and modifiable neurotransmitter release.

Calcium channel levels at single synapses predict release probability and are upregulated in homeostatic potentiation

Abstract Neurons communicate through Ca2+-dependent neurotransmitter release at presynaptic active zones (AZs). Neurotransmitter release properties play a key role in defining information flow in circuits and are tuned during multiple forms of plasticity. Despite their central role in determining neurotransmitter release properties, little is known about how Ca2+ channel levels are modulated to calibrate synaptic function. We used CRISPR to tag the Drosophila CaV2 Ca2+ channel Cacophony (Cac) and investigated the regulation of endogenous Ca2+ channels during homeostatic plasticity in males in which all endogenous Cac channels are tagged. We found that heterogeneously distributed Cac is highly predictive of neurotransmitter release probability at individual AZs and differentially regulated during opposing forms of presynaptic homeostatic plasticity. Specifically, Cac levels at AZ are increased during chronic and acute presynaptic homeostatic potentiation (PHP), and live imaging during acute expression of PHP reveals proportional Ca2+ channel accumulation across heterogeneous AZs. In contrast, endogenous Cac levels do not change during presynaptic homeostatic depression (PHD), implying that the reported reduction in Ca2+ influx during PHD is achieved through functional adaptions to pre-existing Ca2+ channels. Thus, distinct mechanisms bi-directionally modulate presynaptic Ca2+ levels to maintain stable synaptic strength in response to diverse challenges, with Ca2+ channel abundance providing a rapidly tunable substrate for potentiating neurotransmitter release over both acute and chronic timescales.Communication in neural circuits depends on neurotransmitter release at specialized domains of presynaptic terminals called active zones. Evidence in multiple systems indicates that neurotransmitter release properties vary significantly, even between neighboring active zones of the same neuron. To investigate the role of voltage-gated calcium channels in determining diverse release properties, we combined endogenous tagging of the Cav2 channel Cacophony with functional imaging at Drosophila neuromuscular junctions. We find that calcium channels are differentially localized to active zones and robustly predict release probability at single synapses. Synaptic calcium channel levels, in turn, are highly correlated with ELKS/Bruchpilot levels at the active zone cytomatrix. During acute homeostatic potentiation, active zone cytomatrix remodeling is accompanied by a rapid increase in calcium channel levels. We propose that dynamic reorganization of the active zone cytomatrix during short-term plasticity generates new sites for the incorporation of calcium channels to modulate release properties and circuit function.