We thank previous members of our laboratory and scientist Serif Senturk for previous work. We thank Danilo Segovia for critically reading this manuscript. This study was possible and supported by Swim Across America and the National Cancer Institute Cancer Target Discovery and Development Center program.

1.The DD-Cas9 vector
<ol>
	<li>Obtain DD-Cas9 vector from Addgene (DD-Cas9 with filler sequence and Venus (EDCPV), Plasmid 90085). 
	&#8203;NOTE: This is a lentiviral DD-Cas9 plasmid with a U6 promoter that drives the single guide RNA (sgRNA) transcription while the EFS promoter drives the DD-Cas9 transcription. The DYKDDDDK sequence (flag-tag) is present at the C-terminal of Cas9 followed by 2A self-cleaving peptide (P2A) that separates DD-Cas9 and modified fluorescent protein Venus (mVenus).</li>
</ol>
2. Small guide RNA (sgRNA) design
<ol>
	<li>Design small guide RNA (sgRNA) that will be cloned into the DD-Cas9 vector by using one of several algorithms, targeting a specific genomic region. 
	NOTE: To reduce the off-target effects, select sgRNA sequences with the highest score by using the website: http://crispr.mit.edu.</li>
	<li>Design and order two oligonucleotides per sgRNA sequence to make a complete sgRNA. 
	&#8203;NOTE: Since the plasmid will be digested by the BsmBI enzyme, design the forward and the reverse oligonucleotide by adding BsmBI digestion overhangs to the sgRNA sequence. Use the template from Table 1.</li>
</ol>
3. Cloning of sgRNA into the lentiviral DD-Cas9 vector
<ol>
	<li>Dephosphorylate the 5&#8242;-ends of DD-Cas9 plasmid by using alkaline phosphatase (FastAP) in a restriction enzyme digestion reaction. 
	NOTE: Vectors can re-circularize during the ligation especially when they are linearized by a single restriction enzyme. To ensure the vector will not re-circularize, de-phosphorylate the plasmid by adding phosphatase directly into the digestion reaction.
	<ol>
		<li>Dephosphorylate and digest the plasmid with BsmBI enzyme by preparing a mix of 5 &#181;g of DD-Cas9 plasmid, 6 &#181;L of digest buffer (10x), 0.6 &#181;L of DTT (100 mM), 3 &#181;L of BsmBI, 3 &#181;L of FastAP, add up to 60 &#181;L nuclease-free water to make 60 &#181;L the total reaction volume. 
		NOTE: Add the BsmBI enzyme and FastAP to the mixture in the end.</li>
		<li>Incubate the mixture for 30 min at 37&#160;&#176;C heat block or thermocycler.</li>
	</ol>
	</li>
	<li>Purify the digested DD-Cas9 plasmid.
	<ol>
		<li>Prepare an 0.8% DNA agarose gel to remove incompletely digested plasmid and traces of enzymes. Use a wider gel-comb for better separation and run the gel at 90 V.</li>
		<li>Visualize bands under 360-365 nm UV light and cut the larger plasmid band out of the gel. Discard the 2 kb fragment which is corresponding to the filler.</li>
		<li>Purify the large cut gel-band by using a commercial gel purification kit and follow the manufacturer&#39;s instructions.</li>
	</ol>
	</li>
	<li>Ligate annealed oligos with digested DD-Cas9 plasmid.
	<ol>
		<li>Phosphorylate and anneal the sgRNA oligos.
		<ol>
			<li>Prepare the mix of 1 &#181;L of T4 Ligation Buffer (10x), 1 &#181;L of Oligo 1 (100 &#181;M), 1 &#181;L of Oligo 2 (100 &#181;M), 6.5 &#181;L of nuclease-free water. Lastly, add 0.5 &#181;L of T4 PNK. The total volume of this reaction is 10 &#181;L.</li>
			<li>Add the reaction-mix to the thermocycler with the following conditions: 37&#160;&#176;C for 30 min, 95 &#176;C for 5 min, and then ramp down to 25 &#176;C with the speed of 5 &#176;C/min. 
			&#8203;NOTE: The PNK must be heat-inactivated before the oligonucleotides are put into the ligation otherwise the PNK will phosphorylate the vector. The phosphorylation step and annealing step are performed together in a thermocycler. The phosphorylation step where 5&#39; phosphate is added to the sgRNA oligonucleotides is required for the ligation to occur.</li>
		</ol>
		</li>
		<li>Ligate DD-Cas9 digested plasmid and sgRNA oligos.
		<ol>
			<li>Dilute the annealed oligos to 1:200 in nuclease-free water and use 50 ng of plasmid DNA in the ligation reaction. 
			&#8203;NOTE: Use the molar ratio of the 6 insert : 1 vector, to promote insert integration or use a ligation calculator to calculate the molar ratio: http://nebiocalculator.neb.com/#!/ligation.</li>
			<li>Prepare the ligation reaction mix: 50 ng of digested and purified DD-Cas9 vector, appropriate volume of diluted annealed oligos, 1 &#181;L of T4 Ligation Buffer (10x), 1 &#181;L of T4 DNA Ligase, up to 10 &#181;L nuclease-free water to obtain the total volume of the reaction 10 &#181;L.</li>
			<li>If using &#34;high concentration&#34; ligase, incubate the reaction at room temperature for 5 min. Otherwise, incubate at room temperature for 2 h, or to achieve a higher yield of ligation, incubate at 16 &#176;C overnight. 
			&#8203;NOTE: Heat-inactivate if using the standard T4 DNA Ligase at 65 &#176;C for 20 minutes. Do not heat-inactivate if you are using ligase master mixes.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Bacterial transformation
<ol>
	<li>Pre-warm 10 cm LB-ampicillin agar plates at room temperature.</li>
	<li>Thaw 50 &#181;L of &#34;One shot Stbl3&#34; competent bacterial cells on ice.</li>
	<li>Add 2-3 &#181;L of ligation mixture to 50 &#181;L of competent bacterial cells and gently mix by flicking the bottom of the tube with a finger 4 times. Incubate on ice for 30 min.</li>
	<li>Heat-shock the cells at exactly 42 &#176;C for 40 seconds, using a timer. Cool down the bacterial cells on ice for 5 min.</li>
	<li>Add 500 &#181;L of SOC media without antibiotic to the bacterial cells and grow them in the shaking incubator at 250 rpm for 45 min, at 37 &#176;C.</li>
	<li>Spread 100 &#181;L of transformed bacteria on pre-warm LB-ampicillin plates and incubate overnight at 37 &#176;C.</li>
</ol>
5. Mini/maxi-prep of ligated plasmid
<ol>
	<li>Prepare a maxi/mini-prep starter culture.
	<ol>
		<li>The next day pick a single bacterial clone and inoculate a starter culture with 8 mL of LB media containing ampicillin.</li>
		<li>Grow the bacteria in the shaking incubator at 250 rpm, 37 &#176;C for 10-12 h.</li>
		<li>Use 3-5 mL of bacteria for mini-prep or use it as a starter culture for maxi-prep. Use the rest of the bacteria as a bacterial glycerol-stock by adding 100% glycerol in the ratio of bacterial culture:glycerol is 1:1. 
		&#8203;NOTE: Here, we will describe the mini-prep procedure.</li>
	</ol>
	</li>
	<li>Mini-prep procedure
	<ol>
		<li>Harvest 3-5 mL of bacterial cells and centrifuge at 12,000 x g for 1 min.</li>
		<li>Resuspend the pellet of bacterial cells with 200 &#181;L of P1 buffer, containing RNase A and stored at 4 &#176;C.</li>
		<li>Add 200 &#181;L of buffer P2 and mix gently by inverting the tube 10 times until the liquid clarifies.</li>
		<li>Add 300 &#181;L of buffer P3 and mix by inverting the tube 10 times. The liquid will become cloudy. Proceed immediately with centrifuging the samples at 12,000 x g for 10 min.</li>
		<li>Transfer clear supernatant to the spin column and centrifuge at 12,000 x g for 1 min. 
		Discard the flow-through. 
		&#8203;NOTE: Make sure the supernatant is clarified. Particles can clog the spin column and reduce the column efficacy.</li>
		<li>Add 400 &#181;L of PD buffer and centrifuge at 12,000 x g for 1 min. Discard the flow-through.</li>
		<li>Add 600 &#181;L of PW buffer to wash the spin column and centrifuge at 12,000 x g for 1 min. Discard the flow-through.</li>
		<li>Centrifuge again the spin column at top speed for 3 min to remove the buffer residues. Discard the flow-through and let it sit for 2 min.</li>
		<li>Place the spin column to a fresh tube and add 50 &#181;L of elution buffer. Incubate for 5 min and centrifuge at top speed for 2 min.</li>
		<li>Use the nanodrop device to measure the concentration of purified DD-Cas9 plasmid with the sgRNA insert (ng/&#181;L).</li>
	</ol>
	</li>
	<li>Validate the sgRNA cloning by DNA sequencing of the DD-Cas9 plasmid. Use the U6 primer sequence in Table 2.</li>
</ol>
6. Lentiviral preparation
<ol>
	<li>Prepare HEK293T packaging cells for transfection.
	<ol>
		<li>Use healthy HEK293T up to passage 10 and seed them evenly at density 1-2 x 106 cells per 10 cm tissue culture plate. Use 10&#160;mL of Glucose Dulbecco&#39;s Modified Eagle Medium (DMEM) with 10% filtered Fetal Bovine Serum (FBS). Incubate cells in the tissue culture incubator, at 5% CO2 and 37 &#176;C for 20 h.</li>
		<li>Next day, confirm that cells reached 70% confluency by brightfield microscopy and that they are evenly dispersed across the plate. Change media to cells 1 h before the transfection with the final volume of 10 mL. 
		&#8203;NOTE: The media should be absent of antibiotics and antimycotics.</li>
		<li>Prepare the mixture of 2 tubes with 500 &#181;L of warm media (e.g., OptiMEM).
		<ol>
			<li>Add 25 &#181;L of transfection reagent to tube 1 containing warm 500 &#181;L of media, and incubate at room temperature for 5 min.</li>
			<li>Meanwhile prepare&#160;a mix of plasmids to tube 2 containing 3.5 &#181;g of DD-Cas9 containing cloned sgRNA, 6 &#181;g of the packaging plasmid (psPAX2), and 3 &#181;g of the envelope plasmid (pMD2.G) into warm 500 &#181;L of media.</li>
			<li>Mix tube 1 with tube 2 to form a transfection mixture and incubate at room temperature for 20 min.</li>
			<li>Dropwise the transfection mixture to HEK293T cells and incubate plate in the tissue culture incubator, at 5% CO2 and 37 &#176;C overnight.</li>
			<li>After 18 h, carefully change the media to 10 mL of fresh DMEM with 10% FBS to remove the transfection reagent. Incubate for the next 48 h.</li>
			<li>After 48 h, collect the supernatant with a 10 mL syringe and pass it through a 0.45 &#181;m filter.</li>
			<li>Aliquot and store the virus supernatant at -80 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Determining virus titer and transduction efficacy with flow cytometry
<ol>
	<li>Seed the HEK293T cells with 5 x 105 cells/well into a 6-well plate. Seed cells into two 6-well plates. Use one plate for counting the next day.</li>
	<li>Incubate the plates in the incubator at 37 &#176;C, 95% humidity, 5% CO2 overnight to reach 50-60% confluency the next day.</li>
	<li>The next day, use one of the 6-well plates to count the cells in 6 wells.</li>
	<li>Thaw the virus aliquot that will be used to perform the serial dilution and add 8 &#181;g/mL of polybrene reagent.</li>
	<li>Prepare DMEM with 10% FBS for serial dilution of the virus and add 8 &#181;g/mL of polybrene reagent.</li>
	<li>Prepare 2 mL of ten-fold serial dilutions of the lentivirus from 1 x 10-1 to 1 x 10-4 in polybrene-containing media.</li>
	<li>Remove media from 6-well plate and add 1 mL of viral dilutions to wells. Leave one well with media alone as negative control and one well with 100% virus.</li>
	<li>Incubate cells with the virus in the incubator for 24 h.</li>
	<li>The next day, remove the media with virus from 6-well plates and change it for 2 mL of fresh DMEM with 10% FBS. Incubate cells for 48-72 h. Observe GFP by using a fluorescent microscope every day.</li>
	<li>After 48-72 h, detach the cells and resuspend them in MACS buffer.</li>
	<li>Use a flow cytometer to determine the percentage of GFP expression.</li>
	<li>Calculate virus titer by using the following formula: TU/mL = (Number of cells transduced x Percent fluorescent x Dilution Factor)/(Transduction Volume in mL)</li>
</ol>
8. Lentiviral transduction of target cells
<ol>
	<li>Plate the cell line of interest to a 10 cm plate and incubate overnight to reach confluency 50-60% the next day. 
	NOTE: We used the A549 cell line expressing DD-Cas9 and two independent RPA3-gene sgRNAs. We also used the A549 cell line expressing vector with Renilla control. We seeded those cells at the density of 2 x 103cells/cm2 and incubated them at 37 &#176;C, 95% humidity, 5% CO2 overnight to reach 50-60% confluency the next day. We used RPMI media with 10% filtered FBS. We proceeded with the next steps as follow:</li>
	<li>The next day, infect the cells with 500-2000 &#181;L of virus particles in 10 mL total volume of culture medium. Incubate viral media for 24 h.</li>
	<li>The next day, change the viral media to culture media and determine the percentage of GFP positive cells by using flow cytometry.</li>
	<li>Select GFP positive cells by FACS cell sorting or by using bleomycin selection.</li>
	<li>Expand positively selected cells and freeze a stock.</li>
</ol>
9. Conditional induction of Cas9 mediated gene editing
<ol>
	<li>Plate positively selected cells 24 h after infection as well as untransduced cells to 12-well plate separately. Wait until the cells attach and change the media to cell culture media containing 200 nM of Shield-1. Replace the media in the plates with transduced and untransduced-cells, leaving 2 wells per plate with regular media as a negative control.</li>
	<li>Incubate and extract proteins from each well at different time points: time 0, 2 h, 6 h, 12 h, 24 h, 48 h, and 72 h after adding the Shield-1 to the wells.</li>
	<li>Then remove the media with Shield-1 from the rest of the wells and change it for regular cell media.</li>
	<li>Collect the proteins from those wells 2 h, 6 h, and 12 h after changing the media.</li>
	<li>Visualize by western blot analysis the reversibility and rapidity of destabilized DD-Cas9 protein regulation after the addition and the withdrawal of its ligand, Shield-1. Use antibody directed towards DYKDDDDK Tag to visualize proteins and a control antibody targeting for example beta-tubulin.</li>
	<li>Determine the optimal dose of Shield-1 by dose-response curve observed by Western Blot analysis.</li>
</ol>
10. Validation of gene editing
NOTE: The GFP expression assays, such as flow cytometry analysis and bleomycin selection marker only confirm successful CRISPR reagent delivery but they do not determine if the desired sequence was successfully targeted. The most common assays to confirm successful gene targeting by the CRISPR experiment are Sanger DNA sequencing, Next-generation sequencing, the Surveyor Nuclease Assay, the Tracking of Indels by Decomposition (TIDE) Assay, or western blot analysis16,17,18.
<ol>
	<li>Plate positively selected cells and change media to 200 nM of Shield-1 containing media. Incubate cells for 5 days, changing media with Shield-1 every 3 days.</li>
	<li>Use the above-mentioned validation techniques 5 days after DD-Cas9 induction by Shield-1.</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+context+affects+levels+of+a+chemically+dependent+destabilizing+domain.">Intracellular context affects levels of a chemically dependent destabilizing domain.</a> PLOS One. 7 (9), 43297 (2012).</li><li>Iwamoto, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+general+chemical+method+to+regulate+protein+stability+in+the+mammalian+central+nervous+system.">A general chemical method to regulate protein stability in the mammalian central nervous system.</a> Chemistry &amp; Biology. 17 (9), 981-988 (2010).</li><li>Tai, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Destabilizing+domains+mediate+reversible+transgene+expression+in+the+brain.">Destabilizing domains mediate reversible transgene expression in the brain.</a> PLOS One. 7 (9), 46269 (2012).</li><li>Auffenberg, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Remote+and+reversible+inhibition+of+neurons+and+circuits+by+small+molecule+induced+potassium+channel+stabilization.">Remote and reversible inhibition of neurons and circuits by small molecule induced potassium channel stabilization.</a> Scientific Reports. 6, 19293 (2016).</li></ol>

The authors declare that they have no conflict of interest.

The CRISPR/Cas9 technology has revolutionized the capability of functionally interrogate genomes2. However, the inactivation of genes often results in cell lethality, functional deficits, and developmental defects, limiting the utility of such approaches for studying gene functions7. Additionally, constitutive expression of Cas9 may result in toxicity and the generation of off-target effects6. Different approaches have been developed to temporally co...

CRISPR-Cas9 which stands for &#34;clustered regularly interspaced short palindromic repeats-associated protein 9&#34; was first discovered as part of studies on bacterial adaptive immunity1,2. Today, CRISPR/Cas9 has become the most recognized tool for programmable gene editing and different iterations of the system have been developed to allow transcriptional and epigenetic modulations3. This technology enables the highly precise genetic manipulation of almost any sequence of DNA4.
The essential components of any CRISPR gene editing are a customizable guide RNA sequence and the Cas9 nuclease5. The RNA guide binds to the target-complementary sequence in the DNA, directing the Cas9 nuclease to perform a double-strand break at a specific point in the genome3,4. The resulting cleavage site is then repaired by non-homologous end-joining (NHEJ) or homology-directed repair (HDR), with the consequent introduction of changes in the targeted DNA sequence5.
CRISPR/Cas9 based gene editing is easy to use, and relatively inexpensive compared to previous gene-editing techniques and it has been proven to be both efficient and robust in a multiplicity of systems2,4,5. Yet, the system presents some limitations. The constitutive expression of Cas9 has often been shown to result in an increased number of off-targets and high cell toxicity4,6,7,8. Additionally, the constitutive targeting of essential and cell survival genes by Cas9 takes away from its ability to perform certain types of functional studies such as kinetic studies of cell death7.
Different inducible or conditionally controlled CRISPR-Cas9 tools have been developed to address those issues6, such as Tet-ON and Tet-Off9; site-specific recombination10; chemically-induced proximity11; intein dependent splicing3; and 4-Hydroxytamoxifen Estrogen Receptor (ER) based nuclear localization systems12. In general, most of these procedures (intein splicing and chemically induced proximity split systems) do not offer reversible control, present a very slow kinetic response to drug treatment (Tet-On/Off system), or are not amenable to high-throughput manipulation6.
To address these limitations we developed a novel toolkit that not only provides fast and robust temporal-controlled gene editing but also ensures traceability, tunability, and amenability to high throughput gene manipulation. This novel technology can be used in cell lines, organoids, and animal models. Our system is based on an engineered domain, when fused to Cas9, it induces its rapid degradation. However, it can be rapidly stabilized with a highly selective, non-toxic, cell-permeable small molecule. More specifically, we engineered the human FKBP12 mutant &#34;destabilizing domain&#34; (DD) to Cas9, marking Cas9 for rapid and constitutive degradation via the ubiquitin-proteasome system when expressed in mammalian cells13. The DD synthetic ligand, Shield-1 can stabilize DD conformation, thereby preventing the degradation of proteins fused to DD (such as Cas9) in a very efficient manner, and with a fast kinetic response14,15. Of note, Shield-1 binds with three orders of magnitude tighter to the mutant FKBP12 than to its wild-type counterpart14.
The DD-Cas9/Shield-1 pair can be used to study the systematic identification and characterization of essential genes in cultured cells and animal models as we previously showed by conditionally targeting the CypD gene, which plays an important role in the metabolism of mitochondria; EGFR, a key player in oncogenic transformation; and Tp53, a central gene in DNA damage response. In addition to temporally and conditionally controlled gene editing, another advantage of the method is that the stabilization&#160;of DD-Cas9 is independent of its transcription. This feature enables co-expression, under the same promoter, of traceable markers as well as recombinases, such as the estrogen receptor-dependent recombinase, CREER. In this work, we show how our method can be successfully used in vitro, to conditionally target for example, DNA replication gene, RPA3.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100mM DTT</td>
			<td>Thermosfisher</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10X FastDigest buffer</td>
			<td>Thermosfisher</td>
			<td>B64</td>
			<td></td>
		</tr>
		<tr>
			<td>10X T4 Ligation Buffer</td>
			<td>NEB</td>
			<td>M0202S</td>
			<td></td>
		</tr>
		<tr>
			<td>colorimetric BCA kit</td>
			<td>Pierce</td>
			<td>23225</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM, high glucose, glutaMax</td>
			<td>Thermo Fisher</td>
			<td>10566024</td>
			<td></td>
		</tr>
		<tr>
			<td>FastAP</td>
			<td>Thermosfisher</td>
			<td>EF0654</td>
			<td></td>
		</tr>
		<tr>
			<td>FastDigest BsmBI</td>
			<td>Thermosfisher</td>
			<td>FD0454</td>
			<td></td>
		</tr>
		<tr>
			<td>Flag [M2] mouse mAb</td>
			<td>Sigma</td>
			<td>F1804-50UG</td>
			<td></td>
		</tr>
		<tr>
			<td>Genomic DNA extraction kit</td>
			<td>Macherey Nagel</td>
			<td>740952.1</td>
			<td></td>
		</tr>
		<tr>
			<td>lipofectamine 2000</td>
			<td>Invitrogen</td>
			<td>11668019</td>
			<td></td>
		</tr>
		<tr>
			<td>Phusion High-Fidelity DNA Polymerase</td>
			<td>NEB</td>
			<td>M0530S</td>
			<td></td>
		</tr>
		<tr>
			<td>oligonucleotides</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pMD2.G</td>
			<td>Addgene</td>
			<td>12259</td>
			<td></td>
		</tr>
		<tr>
			<td>polybrene</td>
			<td>Sigma Aldrich</td>
			<td>TR-1003-G</td>
			<td></td>
		</tr>
		<tr>
			<td>psPAX2</td>
			<td>Addgene</td>
			<td>12260</td>
			<td></td>
		</tr>
		<tr>
			<td>QIAquick PCR &#38; Gel Cleanup Kit</td>
			<td>Qiagen</td>
			<td>28506</td>
			<td></td>
		</tr>
		<tr>
			<td>secondary antibodies</td>
			<td>LICOR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Shield-1</td>
			<td>Cheminpharma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stbl3 competent bacterial cells</td>
			<td>Thermofisher</td>
			<td>C737303</td>
			<td></td>
		</tr>
		<tr>
			<td>SURVEYOR Mutation Detection Kit</td>
			<td>Transgenomic/IDT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T4 PNK</td>
			<td>NEB</td>
			<td>M0201S</td>
			<td></td>
		</tr>
		<tr>
			<td>Taq DNA Polymerase</td>
			<td>NEB</td>
			<td>M0273S</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-tubulin [DM1A] mouse mAb</td>
			<td>Millipore</td>
			<td>CP06-100UG</td>
			<td></td>
		</tr>
	</tbody>
</table>

a new toolkit for evaluating gene functions using conditional cas9 stabilization

The clustered regularly interspaced short palindromic repeat- (CRISPR-) associated protein 9 (CRISPR/Cas9) technology has become a prevalent laboratory tool to introduce accurate and targeted modifications in the genome. Its enormous popularity and rapid spread are attributed to its easy use and accuracy compared to its predecessors. Yet, the constitutive activation of the system has limited applications. In this paper, we describe a new method that allows temporal control of CRISPR/Cas9 activity based on conditional stabilization of the Cas9 protein. Fusing an engineered mutant of the rapamycin-binding protein FKBP12 to Cas9 (DD-Cas9) enables the rapid degradation of Cas9 that in turn can be stabilized by the presence of an FKBP12 synthetic ligand (Shield-1). Unlike other inducible methods, this system can be adapted easily to generate bi-cistronic systems to co-express DD-Cas9 with another gene of interest, without conditional regulation of the second gene. This method enables the generation of traceable systems as well as the parallel, independent manipulation of alleles targeted by Cas9 nuclease. The platform of this method can be used for the systematic identification and characterization of essential genes and the interrogation of the functional interactions of genes in in vitro and in vivo settings.

To enable the conditional expression of Cas9, we developed a dual lentiviral vector construct consisting of a U6-driven promoter to constitutively express sgRNA, and an EF-1&#945; core promoter to drive the expression of the DD-Cas9 fusion protein (Figure 1A)19. As a paradigm to illustrate the robustness and efficiency of the system, we transduced the lung carcinomatous A549 cell line with the lentiviral construct. The levels of Cas9 i...

Watch this Scientific Journal Video about A New Toolkit for Evaluating Gene Functions using Conditional Cas9 Stabilization at JoVE.com

A New Toolkit for Evaluating Gene Functions using Conditional Cas9 Stabilization

Here, we describe a genome-editing tool based on the temporal and conditional stabilization of clustered regularly interspaced short palindromic repeat- (CRISPR-) associated protein 9 (Cas9) under the small molecule, Shield-1. The method can be used for cultured cells and animal models.

The clustered regularly interspaced short palindromic repeat- (CRISPR-) associated protein 9 (CRISPR/Cas9) technology has become a prevalent laboratory ...

a-new-toolkit-for-evaluating-gene-functions-using-conditional-cas9

Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

4.0K Views.  Cold Spring Harbor Laboratory. Here, we describe a genome-editing tool based on the temporal and conditional stabilization of clustered regularly interspaced short palindromic repeat- (CRISPR-) associated protein 9 (Cas9) under the small molecule, Shield-1. The method can be used for cultured cells and animal models.

Video: A New Toolkit for Evaluating Gene Functions using Conditional Cas9 Stabilization

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

A Protocol for the Production of Integrase-deficient Lentiviral Vectors for CRISPR/Cas9-mediated Gene Knockout in Dividing Cells

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells and mediating high levels of gene expression. However, their ability to integrate into the host cell genome enhances the risk of insertional mutagenicity and thus raises safety concerns and limits their usage in clinical settings. Further, the persistent expression of gene-editing components delivered by these integration-competent lentiviral vectors (ICLVs) increases the probability of promiscuous gene targeting. As an alternative, a new generation of integrase-deficient lentiviral vectors (IDLVs) has been developed that addresses many of these concerns. Here the production protocol of a new and improved IDLV platform for CRISPR-mediated gene editing and list the steps involved in the purification and concentration of such vectors is described and their transduction and gene-editing efficiency using HEK-293T cells was demonstrated. This protocol is easily scalable and can be used to generate high titer IDLVs that are capable of transducing cells in vitro and in vivo. Moreover, this protocol can be easily adapted for the production of ICLVs.

We describe the production strategy of integrase-deficient lentiviral vectors (IDLVs) as vehicles for delivering CRISPR/Cas9 to cells. With an ability to mediate quick and robust gene editing in cells, IDLVs present a safer and equally effective vector platform for gene delivery compared to integrase-competent vectors.

Lentiviral vectors are an ideal choice for delivering gene-editing components to cells due to their capacity for stably transducing a broad range of cells ...

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 ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively correlated with maternal age, age alone is not always predictive of the risk of generating an aneuploid embryo. Therefore, gene variants might account for incorrect chromosome segregation during oogenesis. Given that access to human oocytes is limited for research purposes, a series of assays were developed to study human gene function during meiosis I using mouse oocytes. First, messenger RNA (mRNA) of the gene and gene variant of interest are microinjected into prophase I-arrested mouse oocytes. After allowing time for expression, oocytes are synchronously released into meiotic maturation to complete meiosis I. By tagging the mRNA with a sequence of a fluorescent reporter, such as green fluorescent protein (Gfp), the localization of the human protein can be assessed in addition to the phenotypic alterations. For example, gain or loss of function can be investigated by establishing experimental conditions that challenge the gene product to fix meiotic errors. Although this system is advantageous in investigating human protein function during oogenesis, adequate interpretation of results should be undertaken given that protein expression is not at endogenous levels and, unless controlled for (i.e. knocked out or down), murine homologs are also present in the system.

As the genetic variants associated with human disease begin to become uncovered, it is becoming increasingly important to develop systems with which to rapidly evaluate the biological significance of those identified variants. This protocol describes methods for evaluating human gene function during female meiosis I using mouse oocytes.

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A New Toolkit for Evaluating Gene Functions using Conditional Cas9 Stabilization

Summary

Explore More Videos

Chapters in this video

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

A Protocol for the Production of Integrase-deficient Lentiviral Vectors for CRISPR/Cas9-mediated Gene Knockout in Dividing Cells

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

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9