The authors would like to acknowledge Gissell Sanchez, Megan Lee, and Jason Estep for experimental assistance, and Weifeng Gu and Xuemei Chen for sharing reagents.

1. Identification of Homology Regions Around the Desired Deletion
NOTE: Only necessary if using selection-based editing.
<ol>
	<li>Select two regions, initially 1.5-2 kb, on either side of the desired deletion locus, which will serve as homology arms in the HDR template (Figure 1A). Identify regions that lack BsaI recognition sites on either strand (GGTCTC) to facilitate cloning. If BsaI sites are unavoidable, use an alternate type IIs restriction enzyme (BsmBI, SapI, BbsI) and modify the corresponding sites in the recipient plasmid (pUC19-BsaI), resistance marker donor plasmid (pGolden-Neo or pGolden-Hygro), and homology arm PCR product overhangs.</li>
</ol>
2. Generation of Cas9-sgRNA Expression Plasmids
<ol>
	<li>Define the targeting site(s) for mutagenesis. For the single-cut, selection-free method, target 5&#39; proximal coding exons to increase the probability of a non-functional mutation. For the two-cut method, select two sites that span as much of the gene as necessary. 
	NOTE: In our experience, deletions up to 53 kb are efficiently created.</li>
	<li>Input 200 to 300 bp of the targeted region sequence into the crispr.mit.edu design tool. For selection-based cloning, use 200-300 bp of the identified homology regions that are proximal to the deletion locus (Figure 1A). Select 2-3 of the highest-ranked sgRNAs per targeted region to account for differences in their activity, and to isolate independent clones that do not share the same potential off-target effects.</li>
	<li>To design duplexed oligonucleotides with appropriate overhangs for insertion into the expression plasmid, omit the ending PAM sequence (NGG) and append a 5&#39;-CACC overhang followed by a G to the top strand oligo. For the bottom strand oligo, append a 5&#39;-AAAC overhang to the reverse-complemented target sequence, followed by a 3&#39;-C, as illustrated below: 
	<img alt="Sequence 1" src="/files/ftp_upload/55903/55903seq01.jpg" /></li>
	<li>Insert synthetic oligonucleotides corresponding to the sgRNAs into the pSpCas9(BB) plasmid using Golden Gate cloning28 as described in the Zhang lab protocol29.</li>
</ol>
3. Generation of Homology-directed Repair Template Plasmids
NOTE: Only necessary if using selection-based editing. The homology-directed repair template plasmid consists of a drug resistance cassette flanked by two regions that are complementary to the genome just outside the two sgRNA target sites (Figure 1B).
<ol>
	<li>From the broader homology regions identified in step 1.1, select 800-1,000 bp26 no more than 5-10 bp away from the designed Cas9 cut sites, to serve as the homology arms (Figure 1A). Be sure not to include the sgRNA target site and its PAM in the homology arms, as this will cause CRISPR/Cas9 cleavage of the template plasmid itself. If the homology arms do contain internal BsaI sites, use alternate sgRNA sites.</li>
	<li>PCR amplify homology arms from genomic DNA using a high-fidelity DNA polymerase, per manufacturer&#39;s instructions, using forward and reverse primers with additional 5&#39; sequence that introduces BsaI sites (GGTCTC) and unique overhangs on either end of the homology region. The overhangs must contain proper sequences to generate correct assembly order and orientation, as described below (Figure 1B): 
	Left homology arm FP overhang: 5&#39; GGGTCTCAGGCC 
	Left homology arm RP overhang: 5&#39; GGGTCTCTCACG 
	Right homology arm FP overhang: 5&#39; GGGTCTCAGTCC 
	Right homology arm RP overhang: 5&#39; GGGTCTCTCCAC</li>
	<li>Check homology arms for appropriate amplification via gel electrophoresis before proceeding forward.</li>
	<li>Assemble the right and left homology arm regions with the resistance cassette into an HDR template plasmid by Golden Gate cloning28. Use pGolden-Neo or pGolden-Hygro plasmids30 as the source of loxP-flanked resistance cassettes (loxP-PGK-Neo-pA-loxP or loxP-PGK-Hyg-pA-loxP). Use pUC19-BsaI, a modified pUC19 with BsaI sites in the multiple cloning region and eliminated BsaI sites elsewhere, as the recipient vector (available upon request). Use a ratio of 1:1:1:1 for the three inserts and vector.
	<ol>
		<li>Prepare the following reaction mixture30:
		<table>
			<tbody>
				<tr>
					<td>Right homology arm PCR product</td>
					<td>0.06 pmol (30-40ng)</td>
				</tr>
				<tr>
					<td>Left homology arm PCR product</td>
					<td>0.06 pmol (30-40ng)</td>
				</tr>
				<tr>
					<td>pGolden-Neo plasmid (100 ng/&#181;L)</td>
					<td>1 &#181;L</td>
				</tr>
				<tr>
					<td>pUC19-BsaI plasmid (100 ng/&#181;L)</td>
					<td>1 &#181;L</td>
				</tr>
				<tr>
					<td>2x T7 DNA ligase buffer</td>
					<td>5 &#181;L</td>
				</tr>
				<tr>
					<td>BsaI (10 U/&#181;L)</td>
					<td>0.75 &#181;L</td>
				</tr>
				<tr>
					<td>T7 DNA ligase (3000 U/&#181;L)</td>
					<td>0.25 &#181;L</td>
				</tr>
				<tr>
					<td>Water</td>
					<td>up to 10 &#181;L</td>
				</tr>
			</tbody>
		</table>
		</li>
		<li>Use the following thermocycler parameters: 37 &#176;C for 5 min, 20 &#176;C for 5 min. Repeat for 30 cycles.</li>
		<li>Treat the cloning product with an exonuclease per manufacturer&#39;s instructions to digest away any remaining linearized DNA.</li>
		<li>Transform the reaction mixture into a competent E. coli strain according to the protocol supplied with the cells, and plate on ampicillin-containing dishes.</li>
	</ol>
	</li>
	<li>To identify clones with the correct template assembly, perform bacterial colony PCR to amplify the homology arm subregions of the assembled insert (as the whole insert may be too large to amplify reliably). Use one primer annealing to the flanking plasmid sequence and a second primer complementary to the edge of the resistance cassette (Figure 1B, the sequence is common to both Neo and Hygro inserts), generating an assembly-dependent product that is approximately 200 bp longer than the contained homology arm: 
	pUC19-Bsal-Left: 5&#39; GGCTCGTATGTTGTGTGGAATTGTGAG 
	Resistance-Left: 5&#39; AAAAGCGCCTCCCCTACCC 
	pUC19-BsaI-Right: 5&#39; GCTATTACGCCAGCTGGCGAAA 
	Resistance-Right: 5&#39; AAGACAATAGCAGGCATGCTGGG
	<ol>
		<li>Pick bacterial colonies with sterile toothpicks and suspend the bacteria in 20 &#181;L water. Heat at 95 &#176;C for 15 min, and spin down in a tabletop centrifuge at maximal speed for 5 min. Place immediately on ice.</li>
		<li>Prepare the following PCR mixture31:
		<table>
			<tbody>
				<tr>
					<td>Dilute colony template</td>
					<td>1.25 &#181;L</td>
				</tr>
				<tr>
					<td>10x Taq reaction buffer</td>
					<td>1.25 &#181;L</td>
				</tr>
				<tr>
					<td>20mM dNTPs</td>
					<td>0.25 &#181;L</td>
				</tr>
				<tr>
					<td>10 &#181;M Forward primer&#160;</td>
					<td>0.25 &#181;L</td>
				</tr>
				<tr>
					<td>10 &#181;M Reverse primer&#160;</td>
					<td>0.25 &#181;L</td>
				</tr>
				<tr>
					<td>Taq Polymerase</td>
					<td>0.25 &#181;L</td>
				</tr>
				<tr>
					<td>Water&#160;</td>
					<td>9 &#181;L</td>
				</tr>
			</tbody>
		</table>
		</li>
		<li>Use the following thermocycler parameters: 95 &#176;C for 30 s, (95 &#176;C 30 s, 61 &#176;C for 30 s, 72 &#176;C for 1 min/kb), repeat for 25 cycles, 72 &#176;C for 2 min.</li>
		<li>Check PCR product for correct amplicon size by agarose gel electrophoresis.</li>
		<li>Optionally, miniprep plasmid DNA from individual colonies with correct insert size and sequence the homology arm regions by Sanger sequencing with the amplification primers mentioned above.</li>
	</ol>
	</li>
</ol>
4. Transfection of CRISPR Components into Cultured Cells
<ol>
	<li>Prior to transfection: culture T-REx293 cells in DMEM media supplemented with 10% FBS at 37 &#176;C and 5% CO2.</li>
	<li>Plate cells onto 6-well plates and grow to approximately 70% confluency. Include a well for an untransfected control.</li>
	<li>Transfect cells with 2.5 &#181;g total plasmid using a commercial transfection protocol. In parallel, maintain the untransfected control. 
	NOTE: Transfection method and efficiency vary depending on cell type. Determine the appropriate transfection method for the system prior to the experiment.
	<ol>
		<li>For transfection of cells with selection, use 0.75 &#181;g Cas9-sgRNA 1 (left cut), 0.75 &#181;g Cas9-sgRNA 2 (right cut), and 1.0 &#181;g homologous recombination template.</li>
		<li>For transfection of cells without selection, use 2.5 &#181;g Cas9-sgRNA plasmid.</li>
	</ol>
	</li>
</ol>
5. Drug Selection
<ol>
	<li>48 h after transfection, treat the cells with the appropriate drug (Neomycin/Hygromycin). Carry out selection until all cells in the untransfected control die (typically 3-5 days for Neo, 7-14 days for Hygro). 
	NOTE: Use concentrations of 500 &#181;g/mL and 10 &#181;g/mL for Neomycin and Hygromycin, respectively for T-REx293 cells. For other cell types, it may be useful to carry out a prior titration of drug on untransfected cells to determine the effective concentration.</li>
</ol>
6. Isolation of Clonal Populations
<ol>
	<li>Grow cells to 100% confluency in the original well after selection. Seed cells into 96 well plates at a density of 0.33 cells per well. 
	NOTE: Seeding three 96 well plates is a good starting point to ensure isolation of more than one correct clone, but more or less may be needed depending on sgRNA and HDR efficiencies.</li>
	<li>Observe colonies over a 2-4 week period, until colonies are visible to the eye. Pick visible colonies with a sterile pipette tip and reseed in new wells to encourage monolayer growth. When using suspension cells, even lower seeding densities can be used to ensure a greater proportion of single-colony wells.</li>
</ol>
7. Screening Candidates
<ol>
	<li>Screening candidates without the use of selection: dot blot
	<ol>
		<li>Grow individual clones to 50-100% confluency. Dislodge the monolayer of cells by pipetting within the well.</li>
		<li>Aliquot 90 &#181;L of the 100 &#181;L total volume to a clean microcentrifuge tube. Spin down at 6000 rpm for 5 min, remove media, and lyse cell pellet in 10 &#181;L of 1x Lysis Buffer or 1x SDS loading buffer (5x: 250 mM Tris-Cl pH 6.8, 8% SDS, 0.1% bromophenol blue, 40% glycerol, 100 mM DTT). Add 90 &#181;L of new media to the remainder of the cells in the well to continue propagating.</li>
		<li>Pipette 1 &#181;L of cell lysate onto a dry nitrocellulose membrane to form a dot. Blot each sample twice on two separate membranes, creating two identical patterns of samples.</li>
		<li>Block the membranes in 5% milk in TBST (Tris Buffered Saline + 0.01% Tween 20: 8 g NaCl, 0.2g KCl, 3 g Tris base, up to 1 L distilled water, pH 7.4)31 for 1 h at room temperature (with rocking, here and throughout the procedure).</li>
		<li>Blot using the primary antibody against the target protein on one membrane, and the primary antibody for a control protein (tubulin, GAPDH, or any other protein that is not expected to change) on the other. Use the recommended primary antibody dilution (0.2 &#181;g/mL for ELAVL1 and 0.1 &#181;g/mL for Pum2 in this case) in TBST + 5% milk. Incubate 1 h at room temperature.</li>
		<li>Wash 3 times with TBST for 5 min.</li>
		<li>Incubate each membrane with appropriate HRP-conjugated secondary antibody in TBST + 5% milk for 1 h at room temperature.</li>
		<li>Wash 3 times with TBST for 5 min.</li>
		<li>Apply chemiluminescence substrate solution (see Materials Table) following the manufacturer&#39;s instructions and image the blotted membrane on a digital chemiluminescence imager.</li>
		<li>Quantify dot intensities for the target and control proteins using the appropriate software.</li>
		<li>Eliminate candidates with low control protein signal, and calculate background-subtracted ratios of target to control protein intensities for the remaining candidates. Select the candidates with the lowest ratios for passage and further validation by western blot.</li>
	</ol>
	</li>
	<li>Screening candidates with the use of selection: colony PCR
	<ol>
		<li>Collect cell lysate using a DNA prep kit (See Materials Table).
		<ol>
			<li>Duplicate individual colonies in a new 96 well and grow until 100% confluency.</li>
			<li>Remove media from one set of the clones, and resuspend cells in 30 &#181;L of extraction buffer from the kit. Transfer to a clean 1.5 mL microcentrifuge tube.</li>
			<li>Heat the solution to 96 &#176;C for 15 min and let cool to room temperature.</li>
			<li>Add 30 &#181;L of Stabilization buffer from the kit. Mix well.</li>
		</ol>
		</li>
		<li>Identify wildtype and monoallelic/biallelic mutant lines by colony PCR.
		<ol>
			<li>Design two separate sets of PCR primers (for the left and right side of the targeted locus) to amplify regions around the homology arms based on either successful or unsuccessful integration of the resistance cassette (Figure 2C). On each side, use a common primer (red) that anneals outside of the homology arm. Use it with a corresponding paired primer that is complementary to the endogenous sequence, spanning the homology region (blue), to test for the wild-type allele. Use another paired primer, complementary to the inserted resistance cassette (see primers in section 2.3), to test for the desired mutation.</li>
			<li>(Recommended) Validate the primer sets using cell lysate from the original selected bulk cell population, since it will contain a mixture of both WT and mutant alleles (Figure 2D).</li>
			<li>Prepare the following reaction mixture:
			<table>
				<tbody>
					<tr>
						<td>Cell lysate</td>
						<td>0.5 &#181;L</td>
					</tr>
					<tr>
						<td>10x KOD buffer</td>
						<td>1.25 &#181;L</td>
					</tr>
					<tr>
						<td>25mM MgSO4</td>
						<td>0.75 &#181;L</td>
					</tr>
					<tr>
						<td>2 mM dNTPs&#160;</td>
						<td>1.25 &#181;L</td>
					</tr>
					<tr>
						<td>10 &#181;M Forward primer&#160;</td>
						<td>0.375 &#181;L</td>
					</tr>
					<tr>
						<td>10 &#181;M Reverse primer</td>
						<td>0.375 &#181;L</td>
					</tr>
					<tr>
						<td>KOD polymerase</td>
						<td>0.25 &#181;L</td>
					</tr>
					<tr>
						<td>Water</td>
						<td>7.75 &#181;L</td>
					</tr>
				</tbody>
			</table>
			</li>
			<li>Use the following thermocycler conditions: 95 &#176;C for 2 min, (95 &#176;C for 20 s, Primer Tm for 10 s, 70 &#176;C for 20 s/kb) repeat for 25 cycles.</li>
			<li>Visualize the PCR product by agarose gel electrophoresis.</li>
		</ol>
		</li>
		<li>Repeat the colony PCR testing for each side of the predicted integration. Select and expand candidates that show presence of integration and no endogenous sequence product for the deleted region.</li>
		<li>Validate positive candidates by western blot and/or RT-qPCR.</li>
	</ol>
	</li>
</ol>
8. Verify the Genomic Mutation by Sequencing
<ol>
	<li>Isolate genomic DNA from mutant cell lines by phenol chloroform extraction31.</li>
	<li>PCR amplify the sgRNA target region, using primers with 5&#39; overhangs that add BsaI restriction sites to each end. 
	NOTE: For clones edited with HDR, where both alleles may be identical, the PCR product can be sequenced directly.</li>
	<li>Clone the amplified region into pUC19-BsaI by Golden Gate cloning.</li>
	<li>Transform the reaction mixture into a competent E. coli strain.</li>
	<li>Miniprep plasmid DNA from 6-10 individual colonies and sequence the cloned region by Sanger sequencing.</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+cycle+and+genetic+requirements+of+two+pathways+of+nonhomologous+end-joining+repair+of+double-strand+breaks+in+Saccharomyces+cerevisiae.">Cell cycle and genetic requirements of two pathways of nonhomologous end-joining repair of double-strand breaks in Saccharomyces cerevisiae.</a> Mol Cell Biol. 16 (5), 2164-2173 (1996).</li><li>Liang, F., Han, M., Romanienko, P. J., Jasin, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Homology-directed+repair+is+a+major+double-strand+break+repair+pathway+in+mammalian+cells.">Homology-directed repair is a major double-strand break repair pathway in mammalian cells.</a> Proc Natl Acad Sci U S A. 95 (9), 5172-5177 (1998).</li><li>Gratz, S. J., 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=Highly+specific+and+efficient+CRISPR/Cas9-catalyzed+homology-directed+repair+in+Drosophila.">Highly specific and efficient CRISPR/Cas9-catalyzed homology-directed repair in Drosophila.</a> Genetics. 196 (4), 961-971 (2014).</li><li>Cong, L., 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=Multiplex+genome+engineering+using+CRISPR/Cas+systems.">Multiplex genome engineering using CRISPR/Cas systems.</a> Science. 339 (6121), 819-823 (2013).</li><li>Engler, C., Kandzia, R., Marillonnet, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+one+pot,+one+step,+precision+cloning+method+with+high+throughput+capability.">A one pot, one step, precision cloning method with high throughput capability.</a> PLoS One. 3 (11), e3647 (2008).</li><li>Ran, F. 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=Genome+engineering+using+the+CRISPR-Cas9+system.">Genome engineering using the CRISPR-Cas9 system.</a> Nat Protoc. 8 (11), 2281-2308 (2013).</li><li>Luo, Y., Lin, L., Bolund, L., Sorensen, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+construction+of+rAAV-based+gene+targeting+vectors+by+Golden+Gate+cloning.">Efficient construction of rAAV-based gene targeting vectors by Golden Gate cloning.</a> Biotechniques. 56 (5), 263-268 (2014).</li><li>Green, M. R., Sambrook, J., Sambrook, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Molecular cloning: a laboratory manual. , (2012).</li><li>Lin, S., Staahl, B. T., Alla, R. K., Doudna, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+homology-directed+human+genome+engineering+by+controlled+timing+of+CRISPR/Cas9+delivery.">Enhanced homology-directed human genome engineering by controlled timing of CRISPR/Cas9 delivery.</a> Elife. 3, e04766 (2014).</li><li>Kim, S., Kim, D., Cho, S. W., Kim, J., Kim, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+efficient+RNA-guided+genome+editing+in+human+cells+via+delivery+of+purified+Cas9+ribonucleoproteins.">Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins.</a> Genome Res. 24 (6), 1012-1019 (2014).</li></ol>

The authors declare that they have no competing financial interests.

The CRISPR/Cas9 system has allowed for efficient generation of stable genomic modifications, which provide a more consistent alternative to other transient manipulation methods. Here, we have presented two methods for rapid identification of CRISPR/Cas9 gene knockouts in mammalian cell lines. Both methods require little cellular material, so testing can be done in early stages of clonal culture, saving time and reagents. To increase the efficiency of both methods, we recommend testing multiple sgRNAs, as efficiencies var...

Stable genetic alterations provide an advantage over transient methods of cellular perturbation, which can be variable in their efficiency and duration. Genomic editing has become increasingly common in recent years due to the development of target specific nucleases, such as zinc-finger nucleases1,2,3,4,5, transcription activator-like effector nucleases (TALENs)6,7,8,9 and RNA-guided nucleases derived from the clustered, regularly interspaced short palindromic repeats (CRISPR) system10.
The CRISPR/Cas9 editing machinery is adapted from an immune system that bacteria and archaea use to defend against viral infections11,12,13. In this process, short, 20-30 nt fragments of invading viral sequence are incorporated into a genomic locus as &#34;spacers&#34; flanked by repeating units14,15. Subsequent transcription and RNA processing generates small CRISPR-associated RNAs16 (crRNAs) that, together with a trans-activating crRNA17 (tracrRNA), assemble with the effector Cas9 endonuclease. The crRNAs thus provide specificity to Cas9 targeting, guiding the complex to cleave complementary viral DNA sequences and preventing further infections18,19. Any &#34;protospacer&#34; sequence in the targeted DNA can serve as the source of the crRNA, as long as it is directly 5&#39; to a short protospacer adjacent motif (PAM), NGG in the case of S. pyogenes Cas920. The absence of the PAM sequence near the spacer in the host&#39;s CRISPR locus distinguishes between self and non-self, preventing targeting of the host. Because of its universality and flexibility, this biological system has been powerfully adapted for genomic editing, such that nearly any PAM-adjacent DNA site can be targeted. In this version, a further modification fused the crRNA and tracrRNA into a single guide RNA (sgRNA) component that is loaded into the Cas9 protein21.
Upon expression of Cas9 and an sgRNA in eukaryotic cells, the Cas9 protein cleaves both DNA strands at the targeted locus. In the absence of a suitable region of homologous sequence, the cell fixes this break via non-homologous end joining (NHEJ)22,23,24, which typically introduces small deletions or, rarely, insertions. When targeting an open reading frame, the repair likely leads to a translational frameshift that produces a non-functional protein product. In contrast, when provided with an exogenous template with large regions of homology, the cell may fix the double-strand break by homology directed repair25,26. This route allows for larger precise deletions, replacements or insertions in the genome, coupled with the introduction of excisable selection markers27.
Here, we present protocols for generating knockout cell lines by either of these two CRISPR/Cas9 methods (Figure 1A). The NHEJ approach uses a single sgRNA cut site and selection-independent screening, and thus requires little upfront preparation. When using this method, guide RNAs complementary to exons near the 5&#39; end of the transcript, which are most likely to produce a knockout, must be designed. Since the modifications to the genome in this case are small, screening for knockout clones is based on dot blots, where the protein product is assessed in a high-throughput manner. We use the generation of ELAV-like 1 protein (ELAVL1) knockout lines as an example. The second approach relies on homology directed repair (HDR) and uses two sgRNA cut sites that span the gene or region of interest, allowing for deletions of tens of kb. A plasmid with two regions of homology that flank the cleavage sites provides a replacement template (Figure 1B), introducing a selectable resistance marker that increases efficiency of knockout generation. This method can also be adapted to introduce gene modifications with properly designed homology arms. In this case, the integration of a new DNA fragment allows for PCR based screening (Figure 1C). Here, we use the generation of Pumilio RNA binding family member 2 (PUM2) knockout lines as an example.

<table border="0" cellpadding="0" cellspacing="0" width="740">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:227px;">Competent E. coli cells</td>
			<td style="width:95px;"></td>
			<td style="width:120px;"></td>
			<td style="width:299px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">plasmid prep kit</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pSpCas9(BB) plasmid</td>
			<td>Addgene</td>
			<td>42230</td>
			<td>Ran et al 2013, cloning sgRNAs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">BbsI enzyme</td>
			<td>ThermoFisher</td>
			<td>FD1014</td>
			<td>Ran et al 2013, cloning sgRNAs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">T7 DNA ligase</td>
			<td>NEB</td>
			<td>M0318L</td>
			<td>Ran et al 2013, cloning sgRNAs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tango Buffer</td>
			<td>ThermoFisher</td>
			<td>BY5</td>
			<td>Ran et al 2013, cloning sgRNAs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">PlasmidSafe exonuclease</td>
			<td>Epicentre</td>
			<td>E3105K</td>
			<td>Ran et al 2013, cloning sgRNAs</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Q5 hot start high fidelity polymerase</td>
			<td>NEB</td>
			<td>M0494A</td>
			<td>HA amplification</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pUC19-BsaI</td>
			<td></td>
			<td></td>
			<td>Modified pUC19 plasmid, mutated existing BsaI site and inserted two outward facing BsaI sites after BamHI/EcoRI digestion</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pGolden-Neo</td>
			<td>Addgene</td>
			<td>51422</td>
			<td>Resistance cassette</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pGolden-Hygro</td>
			<td>Addgene</td>
			<td>51423</td>
			<td>Resistance cassette</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">BsaI enzyme</td>
			<td>NEB</td>
			<td>R3535S</td>
			<td>Homology arm contruction</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">T7 DNA ligase</td>
			<td>NEB</td>
			<td>M0318L</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">PlasmidSafe exonuclease</td>
			<td>Epicentre</td>
			<td>E3105K</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Toothpicks</td>
			<td></td>
			<td></td>
			<td>bacterial PCR</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Taq polymerase</td>
			<td></td>
			<td></td>
			<td>bacterial colony PCR</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">HEK293 human cells</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMEM</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">FBS</td>
			<td>Corning</td>
			<td>MT35010CV</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Penicillin-Strep (opt.)</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">6 well plates</td>
			<td>BioLite</td>
			<td>12556004</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TransIT-LTI Transfection Reagent</td>
			<td>Mirus</td>
			<td>MIR2300</td>
			<td>for lipofection only</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Opti-MEM</td>
			<td>ThermoFisher</td>
			<td>31985062</td>
			<td>for lipofection only</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">G418 (Neomycin)</td>
			<td>Sigma Aldrich</td>
			<td>A1720-5G</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Hygromycin</td>
			<td>Sigma Aldrich</td>
			<td>H3274-250MG</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">96 well plates</td>
			<td>ThermoFisher</td>
			<td>12556008</td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:18px;">Passive Lysis Buffer, 5x</td>
			<td>Promega</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:18px;">1xSDS loading buffer</td>
			<td></td>
			<td></td>
			<td>recipe decribed in protocol</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Nitrocellulose Membrane</td>
			<td>Bio-Rad</td>
			<td>162-0115</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TBST</td>
			<td></td>
			<td></td>
			<td>recipe decribed in protocol</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dehydrated milk</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">SuperSignal West Dura Extended Duration Substrate</td>
			<td>ThermoFisher</td>
			<td>34075</td>
			<td>for HRP-conjugated secondary antibodies</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Extracta DNA prep for PCR</td>
			<td>Quantabio</td>
			<td>95091-025</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">KOD polymerase</td>
			<td>Novagen</td>
			<td>71316</td>
			<td></td>
		</tr>
	</tbody>
</table>

selection-dependent and independent generation of crispr/cas9-mediated gene knockouts in mammalian cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

For the generation of ELAVL1 knockout lines, a robust antibody was available, so editing using single sgRNAs (Figure 1A, left) was performed, followed by dot immunoblot. Three sgRNAs were transfected independently to compare efficiencies and to rule out off target effects in the resulting clones. After collecting and blotting cell lysates from clonal populations onto two nitrocellulose membranes, the blots were probed for both ELAVL1 and PUM2 (as a normalizat...

Watch this Scientific Journal Video about Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells at JoVE.com

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

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12.4K Views.  University of California, Riverside. Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

Video: Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

Parallel Measurement of Circadian Clock Gene Expression and Hormone Secretion in Human Primary Cell Cultures

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental changes. Considerable progress in our understanding of the tight connection between the circadian clock and most aspects of physiology has been made in the field over the last decade. However, unraveling the molecular basis that underlies the function of the circadian oscillator in humans stays of highest technical challenge. Here, we provide a detailed description of an experimental approach for long-term (2-5 days) bioluminescence recording and outflow medium collection in cultured human primary cells. For this purpose, we have transduced primary cells with a lentiviral luciferase reporter that is under control of a core clock gene promoter, which allows for the parallel assessment of hormone secretion and circadian bioluminescence. Furthermore, we describe the conditions for disrupting the circadian clock in primary human cells by transfecting siRNA targeting CLOCK. Our results on the circadian regulation of insulin secretion by human pancreatic islets, and myokine secretion by human skeletal muscle cells, are presented here to illustrate the application of this methodology. These settings can be used to study the molecular makeup of human peripheral clocks and to analyze their functional impact on primary cells under physiological or pathophysiological conditions.

Here, we describe settings to monitor in parallel circadian bioluminescence and the secretory activity of human islet cells and primary myotubes. For this, we employed lentiviral gene delivery of a luciferase core clock reporter, followed by in vitro synchronization and collection of outflow medium by continuous cell perifusion.

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental ...

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

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 ...

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in which the gene of interest has been disrupted. A gene-disruption DNA construct containing a suitable antibiotic resistance marker gene is useful for the generation of gene-disrupted strains in bacteria. However, conventional construction methods, which require gene cloning steps, involve complex and time-consuming protocols. Here, a relatively facile, rapid, and cost-effective method for targeted gene disruption in Streptococcus mutans is described. The method utilizes a 2-step fusion polymerase chain reaction (PCR) to generate the disruption construct and electroporation for genetic transformation. This method does not require an enzymatic reaction, other than PCR, and additionally offers greater flexibility in terms of the design of the disruption construct. Employment of electroporation facilitates the preparation of competent cells and improves the transformation efficiency. The present method may be adapted for the generation of gene-disrupted strains of various species.

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

We describe a facile, rapid, and relatively inexpensive method for the generation of gene-disrupted Streptococcus mutans strains; this technique may be adapted for the generation of gene-disrupted strains of various species.

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in ...

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 ...

Formaldehyde-assisted Isolation of Regulatory Elements to Measure Chromatin Accessibility in Mammalian Cells

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly defined states of chromatin organization. The dynamic architecture of the nucleus is controlled by multiple mechanisms and shapes the transcriptional output programs. It is, therefore, important to determine locus-specific chromatin accessibility in a reliable fashion that is preferably independent from antibodies, which can be a potentially confounding source of experimental variability. Chromatin accessibility can be measured by various methods, including the Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) assay, that allow the determination of general chromatin accessibility in a relatively low number of cells. Here we describe a FAIRE protocol that allows simple, reliable, and fast identification of genomic regions with a low protein occupancy. In this method, the DNA is covalently bound to the chromatin proteins using formaldehyde as a crosslinking agent and sheared to small pieces. The free DNA is afterwards enriched using phenol:chloroform extraction. The ratio of free DNA is determined by quantitative polymerase chain reaction (qPCR) or DNA sequencing (DNA-seq) compared to a control sample representing total DNA. The regions with a looser chromatin structure are enriched in the free DNA sample, thus allowing the identification of genomic regions with lower chromatin compaction.

The plasticity of nuclear architecture is controlled by dynamic epigenetic mechanisms including non-coding RNAs, DNA methylation, nucleosome repositioning as well as histone composition and modification. Here we describe a Formaldehyde-Assisted Isolation of Regulatory Elements (FAIRE) protocol which allows the determination of chromatin accessibility in a reproducible, inexpensive, and straightforward manner.

Appropriate gene expression in response to extracellular cues, that is, tissue- and lineage-specific gene transcription, critically depends on highly ...

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. ...