We thank Dr. F. Port, Dr. K. O&#39;Connor-Giles, and Dr. S. Feng for discussions on CRISPR strategy; Dr. T.B. Kornberg, and the Bloomington Stock Center for reagents; UMD imaging core facility; and funding from NIH: R00HL114867 and R35GM124878 to SR.

1. Designing and Constructing the gRNA Expression Vector
<ol>
	<li>To precisely replace a long defined region of an exon, use a dual gRNA approach6, in which each gRNA can specifically target two ends of the selected region of interest. To obtain an accurate gene-specific spatiotemporal expression of the driver, select two gRNA target sites within the first coding exon of the gene.</li>
	<li>For Drosophila melanogaster, select the gRNA target sites using the flyCRISPR Optimal Target Finder tool (http://tools.flycrispr.molbio.wisc.edu/targetFinder/). Other available CRISPR design tools can be used as well, including the Optimized CRISPR Design tool (http://crispr.mit.edu), FlyCas9 (https://shigen.nig.ac.jp/fly/nigfly/cas9), and E-CRISP (www.e-crisp.org/E-CRISP). 
	NOTE: The following protocols of gRNA design and cloning was adopted from methods previously described6,7.
	<ol>
		<li>Copy the actual sequence into the box provided in the online tool. Select the most recently released Drosophila melanogaster genome annotation version, &#8220;r_6,&#8221; in the drop-down menu for &#8220;Select genome.&#8221; In the field &#8220;Select guide length (nt),&#8221; input&#8220;20.&#8221; Select to find &#8220;All CRISPR targets&#8221; and click &#8220;Find CRISPR Targets&#8221;.</li>
		<li>Evaluate all the candidate gRNA targets by setting &#8220;Maximum&#8221; for &#8220;Stringency&#8221; and &#8220;NGG Only&#8221; for &#8220;PAM&#8221;. Try to select the gRNA sites without any potential off-targets or a minimum number of off-targets possible. Use the web-tool described in Doench et al. 2014&#160;8 to select gRNAs with a potential &#8220;good&#8221; activity score.</li>
		<li>Simultaneously, ensure that there is no single nucleotide polymorphism in the gRNA targets in the genome of the parent fly line selected for injection (e.g., nos-Cas9 on X chromosome, BL# 54591). Follow the protocol described in Gratz et al. 20157 to extract the genomic DNA from the parent fly line. PCR amplify, approximately, the 500&#8211;1,000 nt regions using primers that flank the sequence of interest and a high-fidelity DNA polymerase; sequence-verify the PCR amplified products.</li>
	</ol>
	</li>
	<li>For generating a tandem gRNA expression vector, follow a ligation-independent cloning protocol6 to introduce two protospacer sequences into a pCFD4 RNA expression vector. Note that an improved multi-gRNA expression vector (pCFD5) is now available where both the sgRNAs are expressed from the strong U6:3 promoters9 (https://www.crisprflydesign.org). Use a DNA Assembly method to clone the gRNAs into the pCFD4 vector.
	<ol>
		<li>Design and order the forward and reverse primers for introducing two protospacer sequences into the RNA expression vector pCFD4 (Table 1A). As previously described6, the forward primer contains the first protospacer sequence flanked by regions corresponding to the U6-1 promoter and gRNA core in pCFD4; the reverse primer contains the reverse complement sequence of the second protospacer flanked by regions corresponding to the gRNA core and U6-3 promoter in pCDF4.</li>
		<li>Resuspend primers to 100 &#956;M with DNase and RNase-free double distilled water (ddH2O). Make a 10 &#956;M working concentration primer. Use pCFD4 plasmid as a template and set up PCR reaction using a high-fidelity polymerase and recommended settings for PCR reaction6.</li>
		<li>To clone the amplified product into pCFD4, digest pCFD4 plasmid with BbsI enzyme by setting up the following reaction: 2&#8211;5 &#956;g of pCFD4 plasmid, 5 &#956;L of 10x digestion buffer, 1 &#956;L of BbsI enzyme, and ddH2O to bring the final volume to 50 &#956;L. Mix the reaction contents by gently tapping the tube and collecting all the scattered droplets from the wall of the tube to the bottom by a brief spin. Incubate the reaction mix at 37 &#176;C for 2 h.</li>
		<li>Run the PCR product from step 2 and the digested product from step 3 on a 1% agarose gel. Perform the electrophoresis for enough time to completely separate the DNA bands. Cut the correct sized DNA bands under a UV trans-illuminator before purifying the expected products using gel elution column following the manufacturer&#8217;s instruction (see Table of Materials). The PCR product should be 600 bp, and the linearized pCFD4 plasmid should be ~6.4 kb 6.</li>
		<li>Set up the DNA assembly reaction in a PCR tube, per the manufacturer&#8217;s instruction (see Table of Materials).</li>
		<li>Transform 2 &#956;L of the assembly product into competent bacteria (DH5&#945; or similar strains with &#916;recA1, &#916;endA1), plate on Ampicillin (100 &#956;g/mL) containing LB (LB/Amp) agar plates, and incubate at 37 &#176;C overnight. Note that the DNA assembly mix might be toxic to certain bacteria strains, but diluting the assembly mix can reduce the toxicity.</li>
		<li>Pick 3&#8211;4 ampicillin-resistant colonies from the overnight-incubated plate, inoculate the colony in 3 mL LB containing 100 &#956;g/mL ampicillin, and grow inoculated bacteria in a 37 &#176;C shaker overnight. Extract plasmid from the overnight-cultured bacteria using the plasmid mini-prep kit following the manufacturer&#8217;s instruction (see Table of Materials).</li>
		<li>Sequence the plasmids with T3 universal primer to screen for the correct clones containing the tandem gRNA insertion. Establish a glycerol stock of bacteria transformed with a sequence-verified pCFD4-gRNA in 20% glycerol and store in a -80 &#176;C freezer for future use.</li>
	</ol>
	</li>
</ol>
2. Designing and Constructing the HDR Donor
<ol>
	<li>For genomic knock-in of a Gal4 or LexA sequence, design a double-stranded HDR donor containing the transactivator sequence flanked by two homology arms.
	<ol>
		<li>Use &#62;1.5 kb left and right homology arms flanking the gRNA targeting site(s). Extended homology arms increase the efficiency of HDR during the repair process 10.</li>
		<li>PCR amplify the homology arms from the genomic DNA (gDNA) extracted from the parent fly line selected for injection (e.g., nos-Cas9 (on X chromosome), BL# 54591). Use a proof-reading hot-start Taq-DNA polymerase enzyme suitable for long PCR extension (see Table of Materials). Sequence-verify.</li>
	</ol>
	</li>
	<li>To avoid retargeting of the gRNA to the engineered locus, design the replacement donor in such a way that the gRNA recognition sites are disrupted by the exogenous sequence introduced. 
	NOTE: to avoid altering the putative cis-regulatory elements, always select the gRNA recognition sites within the coding exon region.</li>
	<li>For generating a replacement cassette, plan the DNA Assembly strategy to join four segments together: the 5&#8217; and 3&#8217; homology arms, the middle exogenous transactivator DNA sequence, and the linearized cloning vector backbone (e.g., pUC19 or other commonly used cloning vectors). Following the manufacturer&#8217;s guideline for DNA assembly (see Table of Materials), design the suitable primers for PCR amplification and assembly of each segment. Resuspend primers to 100 &#956;M with ddH2O. Make a 10 &#956;M working concentration primer. Use high-fidelity polymerase for all the PCR reactions. Follow the following protocol:
	<ol>
		<li>PCR amplify a Gal4 or LexA expression cassette from an available vector (e.g., nls-LexA:p65; the ideal Gal4/LexA/QF2 expression plasmids can be found and obtained from common resources such as Addgene) using the primers listed in Table 1B.
		<ol>
			<li>To retain all the original transcriptional and post-transcriptional regulations of the replaced exon on the expression of the transactivator DNA sequence, design the cassette in such a way that the edited genomic allele would express a chimeric mRNA of the transactivator and the gene. Preserve the 5&#8217; and 3&#8217; end of the targeted exon to retain any splicing signal.</li>
			<li>Incorporate a T2A self-cleaving peptide sequence between the residual 5&#8217; coding exon and the transactivator sequence to prevent translation into a chimeric protein. Add a translation stop codon after the exogenous transactivator sequence (Figure 5).</li>
		</ol>
		</li>
		<li>PCR amplify the homology arms from the genomic DNA (gDNA) extracted from the parent fly line selected for injection (e.g., nos-Cas9 (on X chromosome), BL# 54591) using the primers listed in Table 1B. Use high-fidelity polymerase for all the PCR reactions using the systems as follows: 5 &#956;L of 5x Reaction Buffer, 0.5 &#956;L of 10 mM dNTPs, 1.25 &#956;L of 10 &#956;M Forward Primer, 1.25 &#956;L of 10 &#956;M Reverse Primer, 0.5 &#956;L of Template DNA, 0.25 &#956;L of High-Fidelity DNA Polymerase, 16.25 &#956;L of ddH2O.</li>
		<li>Use linearized cloning vector such as pUC19. A number of donor vectors are now available. See a comprehensive list at the following website- http://flycrispr.molbio.wisc.edu.</li>
		<li>Run the PCR products on a 1% agarose gel. Perform the electrophoresis for enough time to ensure a clear separation of the desired band from the undesired non-specific bands (if any). Gel-purify the expected DNA fragments. Measure the concentration of each purified DNA fragment using a spectrophotometer.</li>
		<li>Set up the DNA assembly reaction following the manufacturer&#8217;s instruction. Mix the linearized cloning vector and target fragments from step 3 and step 4&#160;in the following reaction: 1 &#956;L of linear cloning vector (50 ng/&#956;L), 2&#8211;3 fold excess of each target fragment, 10 &#956;L of 2x DNA assembly master mix, bring the final reaction volume to 20 &#956;L with ddH2O. Incubate the reaction mix for 1 hour at 50 &#176;C.</li>
		<li>Transform 2 &#956;L of the assembly product into competent bacteria, plate on LB/Amp agar plates containing 100 &#956;g/mL ampicillin. Also, add X-Gal + IPTG on the plate for blue/white screening.</li>
		<li>Pick 3&#8211;4 white colonies from the overnight-incubated plate, inoculate the colony in 3 mL LB containing 100 &#956;g/mL ampicillin, and grow at 37 &#176;C overnight in a shaker. Extract plasmid from the overnight-cultured bacteria using the plasmid mini-prep kit following the manufacturer&#8217;s manual (see Table of Materials).</li>
		<li>Screen the positive colony by PCR or restriction digestion.</li>
		<li>Sequence verify the HDR donor region of the final plasmid. Save a bacterial stock transformed with the correct clones in 20% glycerol at -80 &#176;C for future inoculation.</li>
	</ol>
	</li>
</ol>
3. Embryo Injection, Fly Genetics and Screening for Genome Editing
<ol>
	<li>Prepare the high purity endotoxin-free gRNA expression vector as well as the HDR donor plasmid using a plasmid maxiprep kit (see Table of Materials).</li>
	<li>Co-inject the gRNA expression plasmid (100 ng/&#956;L) and the replacement donor (500 ng/&#956;L) into the germline of nos-Cas9 embryos6. Note: we use a commercial service for injection, but this procedure can be performed in the laboratory as well.</li>
	<li>Fly genetics and screening (Figure 2 and Figure 3):
	<ol>
		<li>When the injected embryos develop into adults, cross each single G0 flies to balancer flies. Select suitable balancer for the chromosome containing the targeted allele.</li>
		<li>Anesthetize the F1 offspring from each G0 cross on a CO2 pad and randomly pick 10-20 males under a stereomicroscope. Cross them individually to the balancer females as shown in Figure 3.</li>
		<li>When the F2 larvae hatch, pick the single F1 father from each cross and extract gDNA using the single fly genomic DNA preparation protocol:
		<ol>
			<li>Prepare gDNA extraction buffer: 10 mM Tris-Cl pH 8.2, 1 mM EDTA, 25 mM NaCl, store at room temperature. Prepare 20 mg/mL Proteinase K stock solution and store in the freezer.</li>
			<li>Put each fly in a 1.5 mL micro-centrifuge tube and label the tube. Keep in the -80 &#176;C freezer overnight.</li>
			<li>Prepare a fresh working volume of gDNA extraction buffer containing 200 &#181;g/mL final concentration of Proteinase K. 
			Note: Do not use an old buffer for this step.</li>
			<li>Squish each fly for 10&#8211;15 s with a pipette tip containing 50 &#956;L of squishing buffer without dispensing the liquid. Dispense the remaining buffer into the tube and mix well. Incubate at 37 &#176;C for 20&#8211;30 min.</li>
			<li>Put tubes in 95 &#176;C heat block for 1&#8211;2 min to inactivate the Proteinase K.</li>
			<li>Spin down for 5 min at 10,000 x g. Store the preparation at 4 &#176;C for further PCR analysis.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Use the same method to prepare gDNA from a nos-Cas9 fly, which serves as a negative control. Perform three-step PCR based screens to identify the correct &#8220;ends out&#8221; HDR (Figure 2, Figure 5) using gDNA of each F1 male as a template5. Use 1 &#956;L of the DNA prep in the following PCR reaction system: 10 &#956;L of 2x PCR Master Mix with Dye, 1 &#956;L of each primer (10 &#956;M), 1 &#956;L of DNA template, and 7 &#956;L of ddH2O.</li>
	<li>As shown in Figure 5A, perform PCR using primers fwd1 and rev1 to screen for the existence of the insertion or replacement; perform PCR using fwd2 and rev2 primers to verify the insertion or replacement from 3&#8217; region; perform PCR using primers M13F and rev3 to check &#8220;ends-in&#8221; HDR (Table 1C).</li>
	<li>Keep the fly lines with the confirmed ends-out HDR and establish balanced stocks from the F2 generation. Outcross to the balancer flies again to remove any unintended mutations on other chromosomes.</li>
	<li>Prepare high-quality gDNA from the established stocks for amplifying long PCR (&#62;800&#8211;1,000 nt) amplicon with high-fidelity Taq polymerase and fully sequence verify the amplicons obtained from the engineered genomic regions. Alternatively, use crude gDNA extract as described earlier to amplify shorter (&#60;800 nt), overlapping PCR products to sequence-validate the edited genome. Use the following protocol to prepare the good quality Drosophila genomic DNA:
	<ol>
		<li>Put about 25 adult flies in a 1.5 mL microcentrifuge tube, and freeze in -20 &#176;C or -80 &#176;C freezer for at least 1 hour.</li>
		<li>Add 250 &#956;L of Solution A (pH 9.0 Tris HCl 0.1 M, EDTA 0.1 M, SDS 1%).</li>
		<li>Homogenize the flies using the homogenizing pestles in microcentrifuge tubes, and put the tube on ice.</li>
		<li>Incubate at 70 &#176;C for 30 min.</li>
		<li>Add 35 &#956;L of KOAc (5 M), shake, and mix well, but do not vortex.</li>
		<li>Incubate on ice for 30 min.</li>
		<li>Spin at 12,000 x g for 15 min.</li>
		<li>With a 1 mL micropipette, carefully transfer only the clear supernatant to a new tube, leaving back any precipitate or interphase.</li>
		<li>Add 150 &#956;L of isopropanol to the supernatant. Gently mix by inversion.</li>
		<li>Spin at 10,000 x g for 5 min.</li>
		<li>Carefully remove the supernatant and leave the pellet in the tube.</li>
		<li>Wash the pellet with 1 mL 70% EtOH.</li>
		<li>Spin at 12,000 x g for 5 min.</li>
		<li>Remove the supernatant. Air dry the pellet for 15 to 20 min. Do not over dry the pellet.</li>
		<li>Dissolve the pellet in 100 &#956;L of ddH2O.</li>
		<li>Use high fidelity DNA polymerase suitable for long amplicon (&#62;800 bp) to amplify the complete edited region. Ideally, take two primers outside the inserted cassette to amplify the genomic region. Gel purify the PCR product, and as described earlier, sequence verify the product with multiple overlapping primers.</li>
	</ol>
	</li>
	<li>Validate the correct spatiotemporal expression patterns from dissected tissues/embryos of the engineered fly lines:
	<ol>
		<li>Perform RT-PCR&#160;from the total RNA to verify the expression of the hybrid mRNA product (Table 1D).</li>
		<li>Perform an in situ hybridization of the mRNA product of the interest in the larval tissues/embryo to validate the expression.</li>
		<li>To screen for the accurate tissue-specific spatiotemporal expression patterns among the sequence-verified ends-out lines, cross each of the edited lines obtained for the binary expression driver with the LexO-GFP or LexO-RFP (for LexA driver) line (available from the Bloomington stock centers) and observe the LexA or Gal4 driven reporter-gene expression in embryo, larval, and adult tissues under a fluorescence microscope.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to disclose.

Traditionally, Drosophila enhancer traps were generated by two different methods. One of the ways includes random insertion of a driver (eg., Gal4) sequence in the genome by transposition (e.g., P-element transposition)1 . Alternatively, the driver sequences can be placed under the transcriptional control of a putative enhancer/promoter region in a plasmid construct, which would then be integrated into an ectopic site of the genome3,<su...

The genetic toolbox for targeted gene expression has been well developed in Drosophila, making it one of the best model systems to investigate the function of genes involved in a wide variety of cellular processes. Binary expression systems, such as yeast Gal4/UAS (upstream activation sequence), was first adopted for tissue-specific enhancer trapping and gene misexpression in the Drosophila genetic model1 (Figure 1). This system facilitated the development of a large number of techniques such as spatiotemporal regulation of gene overexpression, misexpression, knockout in selected groups of cells as well as in cell ablation, cell marking, live tracing of cellular and molecular processes in embryo and tissues, lineage tracing and mosaic analyses during development. A number of binary transcription system, such as the bacterial LexA/LexAop system (Figure 1) and Neurospora Q-system, are powerful genetic tools that are now widely used in Drosophila, in addition to the original Gal4/UAS system for targeted gene expression1,2,3.
Here, we present a method to generate highly reliable tissue-specific binary expression system by employing a genome-editing technique. The recent advancements in CRISPR/Cas9 genome editing technology have allowed unprecedented opportunities to make directed genome changes in a broad range of organisms. Compared to the other available genome editing techniques, the CRISPR/Cas9 system is inexpensive, efficient, and reliable. This technology utilizes components of the bacterial adaptive immune system: a Cas9 endonuclease of Streptococcus pyogenes that creates a double-strand break (DSB), and a chimeric guide RNA (gRNA), which guides the Cas9 to a particular genome site for targeted DSB4. The cells contain the machinery to repair the DSB using different pathways. Non-homologous end joining (NHEJ) leads to small insertions or deletions to disrupt gene function, while homology-directed repair (HDR) introduces a defined directed/desirable genomic knock-in/knock-out by using an exogenous HDR donor as a template. The HDR-based replacement strategy can efficiently be utilized to generate a highly reliable tissue-specific binary expression system, which can overcome all the limitations of the traditional enhancer trap methods. We describe a step-by-step procedure for utilization of CRISPR/Cas9-based HDR repair in generating a binary transcription driver line that is expressed under the control of endogenous transcriptional and post-transcriptional regulation of a Drosophila gene. In this protocol, we demonstrate the generation of a driver line specific for branchless (bnl) gene encoding an FGF family protein that regulates branching morphogenesis of tracheal airway epithelium5. In this example, the first coding exon of the bnl gene was replaced by the sequence of a bacterial LexA transactivator sequence without altering any endogenous cis-regulatory sequences of the bnl gene. We show that the strategy generated a bnl-LexA driver line that spatiotemporally controls the expression of a reporter gene placed downstream of LexAoperator (LexAop or LexO) exclusively in bnl-expressing epithelial/mesenchymal/neuronal cells.

<table><tbody><tr><td>X-Gal/IPTG</td><td>Gentrox (Genesee Scientific)</td><td>18-218</td><td>cloning</td></tr><tr><td>LB-Agar</td><td>BD Difco</td><td>BD&amp;nbsp;244520</td><td>cloning</td></tr><tr><td>Tris-HCl</td><td>Sigma Aldrich</td><td>T3253</td><td>Molecular Biology</td></tr><tr><td>EDTA</td><td>Sigma Aldrich</td><td>E1161</td><td>Molecular Biology</td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S7653</td><td>Molecular Biology</td></tr><tr><td>UltraPure DNase/RNase-Free Water</td><td>ThermoFisher Scientific</td><td>10977-023</td><td>Molecular Biology</td></tr><tr><td>10% SDS</td><td>Sigma Aldrich</td><td>71736</td><td>Molecular Biology</td></tr><tr><td>KOAc</td><td>Fisher-Scientific</td><td>P1190</td><td>Molecular Biology</td></tr><tr><td>EtOH</td><td>Fisher-Scientific</td><td>04-355-451</td><td>Molecular Biology</td></tr><tr><td>GeneJET Miniprep</td><td>ThermoFisher Scientific</td><td>K0503</td><td>Miniprep</td></tr><tr><td>PureLink HiPure Plasmid Maxipep kits</td><td>ThermoFisher Scientific</td><td>K210006</td><td>Maxiprep</td></tr><tr><td>BbsI</td><td>NEB</td><td>R0539S</td><td>Restriction enzyme</td></tr><tr><td>Primers</td><td>IDT-DNA</td><td></td><td>PCR</td></tr><tr><td>pCFD4</td><td>Kornberg Lab</td><td></td><td>DNA template and vector for gRNA</td></tr><tr><td>KAPA HiFi Hot Start- (Kapa Biosystems)</td><td>Kapa biosystems</td><td>KK2601</td><td>PCR</td></tr><tr><td>Q5-high fidelity Taq</td><td>NEB</td><td>NEB #M0491</td><td>PCR</td></tr><tr><td>Gibson Assembly Master Mix</td><td>NEB</td><td>NEB #E2611</td><td>DNA assembly</td></tr><tr><td>pBPnlsLexA:p65Uw</td><td>Addgene</td><td></td><td>DNA template for LexA amplification</td></tr><tr><td>Proteinase K</td><td>ThermoFisher Scientific</td><td>25530049</td><td>Molecular Biology</td></tr><tr><td>2x PCR PreMix, with dye (red)</td><td>Sydlab</td><td>MB067-EQ2R</td><td>Molecular Biology</td></tr><tr><td>Gel elution kit</td><td>Zymo Research (Genesee Scientific)</td><td>11-300</td><td>Molecular Biology</td></tr><tr><td>TRI reagent</td><td>Sigma-Aldrich</td><td></td><td>Molecular Biology</td></tr><tr><td>Direct-zol RNA purification kits</td><td>Zymo Research (Genesee Scientific)</td><td>11-330</td><td>Molecular Biology</td></tr><tr><td>One&lt;em&gt;Taq&lt;/em&gt; One-Step RT-PCR Kit</td><td>NEB</td><td>E5315S</td><td>Molecular Biology</td></tr><tr><td>&lt;em&gt;lexO-&lt;/em&gt;CherryCAAX</td><td>Kornberg Lab</td><td></td><td>Fly line</td></tr><tr><td>UAS-CD8:GFP</td><td>Kornberg lab</td><td></td><td>Fly line</td></tr><tr><td>&lt;em&gt;btl-Gal4&lt;/em&gt;</td><td>Kornberg lab</td><td></td><td>Fly line</td></tr><tr><td>MKRS/TB6B</td><td>Kornberg lab</td><td></td><td>Fly line</td></tr><tr><td>Confocal Microscope SP5X</td><td>Leica</td><td></td><td>Imaging expression pattern</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; station</td><td>Genesee Scientific</td><td>59-122WCU</td><td>fly pushing</td></tr><tr><td>Stereo microscope</td><td>Olympus</td><td>SZ-61</td><td>fly pushing</td></tr><tr><td>Microtube homogenizing pestles</td><td>Fisher-Scientific</td><td>03-421-217</td><td>genomic DNA isolation</td></tr><tr><td>NanoDrop spectrophotometer</td><td>ThermoFisher Scientific</td><td>ND-1000</td><td>DNA quantification</td></tr></tbody></table>

an efficient strategy for generating tissue-specific binary transcription systems in drosophila by genome editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of cells or tissues in model organisms. These systems contain two components as separate transgenic lines. A driver line expresses a transcriptional activator under the control of tissue-specific promoters/enhancers, and a reporter/effector line harbors a target gene placed downstream to the binding site of the transcription activator. Animals harboring both components induce tissue-specific transactivation of a target gene expression. Precise spatiotemporal expression of the gene in targeted tissues is critical for unbiased interpretation of cell/gene activity. Therefore, developing a method for generating exclusive cell/tissue-specific driver lines is essential. Here we present a method to generate highly tissue-specific targeted expression system by employing a &#34;Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR-associated&#34; (CRISPR/Cas)-based genome editing technique. In this method, the endonuclease Cas9 is targeted by two chimeric guide RNAs (gRNA) to specific sites in the first coding exon of a gene in the Drosophila genome to create double-strand breaks (DSB). Subsequently, using an exogenous donor plasmid containing the transactivator sequence, the cell-autonomous repair machinery enables homology-directed repair (HDR) of the DSB, resulting in precise deletion and replacement of the exon with the transactivator sequence. The knocked-in transactivator is expressed exclusively in cells where the cis-regulatory elements of the replaced gene are functional. The detailed step-by-step protocol presented here for generating a binary transcriptional driver expressed in Drosophila fgf/branchless-producing epithelial/neuronal cells can be adopted for any gene- or tissue-specific expression.

This protocol was successfully used to generate a targeted binary expression reporter system specific for bnl expressing cells5. The cis-regulatory elements (CREs) that control complex spatiotemporal bnl expression are not characterized. Therefore, to achieve spatiotemporal expression under the control of the endogenous bnl regulatory sequence, only the first coding exon of bnl was designed to be replaced with the sequen...

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An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Here, we present a method for generating tissue-specific binary transcription systems in Drosophila by replacing the first coding exon of genes with transcription drivers. The CRISPR/Cas9-based method places a transactivator sequence under the endogenous regulation of a replaced gene, and consequently facilitates transctivator expression exclusively in gene-specific spatiotemporal patterns.

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of ...

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Research

JoVE Journal

Developmental Biology

9.0K Views.  University of Maryland. Het algemene doel van deze methode is het genereren van een zeer weefsel-specifieke gerichte expressie systeem. In deze methode demonstreerden we de generatie van een binaire transcriptie driver specifiek voor onze Drosophila FGF homolog, branchless. Dynamische, spatiotemporale expressie van branchless reguleert tak hemogenese van tracheale luchtweg epitheel.Hoewel deze methode inzicht kan geven in de spatiotemporale expressie en migratie van vertakte producerende cellen, kan deze gema...

Video: An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Genetics

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Neuroscience

In Vivo Functional Study of Disease-associated Rare Human Variants Using Drosophila

Advances in sequencing technology have made whole-genome and whole-exome datasets more accessible for both clinical diagnosis and cutting-edge human genetics research. Although a number of in silico algorithms have been developed to predict the pathogenicity of variants identified in these datasets, functional studies are critical to determining how specific genomic variants affect protein function, especially for missense variants. In the Undiagnosed Diseases Network (UDN) and other rare disease research consortia, model organisms (MO) including Drosophila, C. elegans, zebrafish, and mice are actively used to assess the function of putative human disease-causing variants. This protocol describes a method for the functional assessment of rare human variants used in the Model Organisms Screening Center Drosophila Core of the UDN. The workflow begins with gathering human and MO information from multiple public databases, using the MARRVEL web resource to assess whether the variant is likely to contribute to a patient&#39;s condition as well as design effective experiments based on available knowledge and resources. Next, genetic tools (e.g., T2A-GAL4 and UAS-human cDNA lines) are generated to assess the functions of variants of interest in Drosophila. Upon development of these reagents, two-pronged functional assays based on rescue and overexpression experiments can be performed to assess variant function. In the rescue branch, the endogenous fly genes are &#34;humanized&#34;&#160;by replacing the orthologous Drosophila gene with reference or variant human transgenes. In the overexpression branch, the reference and variant human proteins are exogenously driven in a variety of tissues. In both cases, any scorable phenotype (e.g., lethality, eye morphology, electrophysiology) can be used as a read-out, irrespective of the disease of interest. Differences observed between reference and variant alleles suggest a variant-specific effect, and thus likely pathogenicity. This protocol allows rapid, in vivo assessments of putative human disease-causing variants of genes with known and unknown functions.

In Vivo Functional Study of Disease-associated Rare Human Variants Using Drosophila

The goal of this protocol is to outline the design and performance of in vivo experiments in Drosophila melanogaster to assess the functional consequences of rare gene variants associated with human diseases.

Advances in sequencing technology have made whole-genome and whole-exome datasets more accessible for both clinical diagnosis and cutting-edge human ...

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

Quantitative Comparison of cis-Regulatory Element (CRE) Activities in Transgenic Drosophila melanogaster

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE possesses an arrangement of binding sites for several transcription factor proteins that confer a regulatory logic specifying when, where, and at what level the regulated gene(s) is expressed. The full set of CREs within an animal genome encodes the organism&prime;s program for development1, and empirical as well as theoretical studies indicate that mutations in CREs played a prominent role in morphological evolution2-4. Moreover, human genome wide association studies indicate that genetic variation in CREs contribute substantially to phenotypic variation5,6. Thus, understanding regulatory logic and how mutations affect such logic is a central goal of genetics. 
Reporter transgenes provide a powerful method to study the in vivo function of CREs. Here a known or suspected CRE sequence is coupled to heterologous promoter and coding sequences for a reporter gene encoding an easily observable protein product. When a reporter transgene is inserted into a host organism, the CRE&prime;s activity becomes visible in the form of the encoded reporter protein. P-element mediated transgenesis in the fruit fly species Drosophila (D.) melanogaster7 has been used for decades to introduce reporter transgenes into this model organism, though the genomic placement of transgenes is random. Hence, reporter gene activity is strongly influenced by the local chromatin and gene environment, limiting CRE comparisons to being qualitative. In recent years, the phiC31 based integration system was adapted for use in D. melanogaster to insert transgenes into specific genome landing sites8-10. This capability has made the quantitative measurement of gene and, relevant here, CRE activity11-13 feasible. The production of transgenic fruit flies can be outsourced, including phiC31-based integration, eliminating the need to purchase expensive equipment and/or have proficiency at specialized transgene injection protocols. 
Here, we present a general protocol to quantitatively evaluate a CRE&prime;s activity, and show how this approach can be used to measure the effects of an introduced mutation on a CRE&prime;s activity and to compare the activities of orthologous CREs. Although the examples given are for a CRE active during fruit fly metamorphosis, the approach can be applied to other developmental stages, fruit fly species, or model organisms. Ultimately, a more widespread use of this approach to study CREs should advance an understanding of regulatory logic and how logic can vary and evolve.

Quantitative Comparison of cis-Regulatory Element CRE Activities in Transgenic Drosophila melanogaster

Phenotypic variation for traits can result from mutations in cis-regulatory element (CRE) sequences that control gene expression patterns. Methods derived for use in Drosophila melanogaster can quantitatively compare the levels of spatial and temporal patterns of gene expression mediated by modified or naturally occurring CRE variants.

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE ...

Biology

Recombineering Homologous Recombination Constructs in Drosophila

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a genetic model organism for human-disease related research. Recent years have seen technical advancements like homologous recombination and recombineering. However, generating unequivocal null mutations or tagging endogenous proteins remains a substantial effort for most genes. Here, we describe and demonstrate techniques for using recombineering-based cloning methods to generate vectors that can be used to target and manipulate endogenous loci in vivo. Specifically, we have established a combination of three technologies: (1) BAC transgenesis/recombineering, (2) ends-out homologous recombination and (3) Gateway technology to provide a robust, efficient and flexible method for manipulating endogenous genomic loci. In this protocol, we provide step-by-step details about how to (1) design individual vectors, (2) how to clone large fragments of genomic DNA into the homologous recombination vector using gap repair, and (3) how to replace or tag genes of interest within these vectors using a second round of recombineering. Finally, we will also provide a protocol for how to mobilize these cassettes in vivo to generate a knockout, or a tagged gene via knock-in. These methods can easily be adopted for multiple targets in parallel and provide a means for manipulating the Drosophila genome in a timely and efficient manner.

Recombineering Homologous Recombination Constructs in Drosophila

Homologous recombination techniques greatly advance Drosophila genetics by enabling the creation of molecularly precise mutations. The recent adoption of recombineering allows one to manipulate large pieces of DNA and transform them into Drosophila6. The methods presented here combine these techniques to rapidly generate large homologous recombination vectors.

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a ...

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with an attractive off-target profile at both cellular and organismal levels. It is recently reported that the CRISPR/CasRx system can be used to achieve ubiquitous and tissue-specific gene transcript reduction in Drosophila melanogaster. This paper details the methods from the recent work, consisting of three parts: 1) ubiquitous&#160;in vivo endogenous RNA targeting using a two-component CasRx system; 2) ubiquitous in vivo exogenous RNA targeting using a three-component CasRx system; and 3) tissue-specific in vivo RNA targeting using a three-component CasRx system. The effects of RNA targeting observed include targeted gene specific phenotypic changes, targeted RNA transcript reduction, and occasional lethality phenotypes associated with high expression of CasRx protein and collateral activity. Overall, these results showed that the CasRx system is capable of target RNA transcript reduction at the organismal level in a programmable and efficient manner, demonstrating that in vivo transcriptome targeting, and engineering is feasible and lays the foundation for future in vivo CRISPR-based RNA targeting technologies.

Ubiquitous and Tissue-specific RNA Targeting in Drosophila Melanogaster using CRISPR/CasRx

This article outlines a detailed protocol for using the RNA-targeting Cas13D enzyme (RfxCas13D) in flies.

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with ...

Immunology and Infection

One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that allow the tagging of endogenous genes to produce fluorescent fusion proteins. These methods enable researchers to visualize protein behavior in real time, providing valuable insights into their functions and interactions within the cellular environment. Many current gene tagging studies employ a two-step process where visible markers, such as eye color changes, are used to identify genetically modified organisms in the first step, and the visible marker is excised in the second step. Here, we present a one-step protocol to perform precise and rapid endogenous gene tagging in Drosophila melanogaster, which enables screening for engineered lines without the visible eye marker, offering a significant advantage over past methods. To screen for successful gene-tagging events, we employ a PCR-based technique to genotype individual flies by analyzing a small segment from their middle leg. Flies that pass the screening criteria are then used to produce stable stocks. Here, we detail the design and construction of CRISPR editing plasmids and methods for screening and confirmation of engineered lines. Together, this protocol improves the efficiency of endogenous gene tagging in Drosophila significantly and enables studies of cellular processes in vivo.

One-step CRISPR-based Strategy for Endogenous Gene Tagging in Drosophila melanogaster

Here, we present a simplified endogenous gene tagging protocol for Drosophila, which utilizes a PCR-based technique for marker-free identification of successful genetic modifications, facilitating the development of stable knock-in lines.

The study of protein subcellular localization, dynamics, and regulation in live cells has been profoundly transformed by the advent of techniques that ...

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops worldwide. Since adult fruit flies have great flight capacity and females lay their eggs under the skins of fruit, insecticides requiring direct contact with the pest usually perform poorly in the field. With the development of molecular biological tools and high-throughput sequencing technology, many scientists are attempting to develop environmentally friendly pest management strategies. These include RNAi or gene editing-based pesticides that downregulate or silence genes (molecular targets), such as olfactory genes involved in searching behavior, in various insect pests. To adapt these strategies for Oriental fruit fly control, effective methods for functional gene research are needed. Genes with critical functions in the survival and reproduction of B. dorsalis serve as good molecular targets for gene knockdown and/or silencing. The CRISPR/Cas9 system is a reliable technique used for gene editing, especially in insects. This paper presents a systematic method for CRISPR/Cas9 mutagenesis of B. dorsalis, including the design and synthesis of guide RNAs, collecting embryos, embryo injection, insect rearing, and mutant screening. These protocols will serve as a useful guide for generating mutant flies for researchers interested in functional gene studies in B. dorsalis.

Protocols for CRISPR/Cas9 Mutagenesis of the Oriental Fruit Fly Bactrocera dorsalis

This paper presents the step-by-step protocols for CRISPR/Cas9 mutagenesis of the Oriental fruit fly Bactrocera dorsalis. Detailed steps provided by this standardized protocol will serve as a useful guide for generating mutant flies for functional gene studies in B. dorsalis.

The Oriental fruit fly, Bactrocera dorsalis, is a highly invasive and adaptive pest species that causes damage to citrus and over 150 other fruit crops ...

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

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