Name: selection-dependent and independent generation of crispr/cas9-mediated gene knockouts in mammalian cells
Uploaded: 2017-06-16T15:00:00.000+00:00
Duration: 11 min 35 s
Description: Watch this Scientific Journal Video about Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells at JoVE.com

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 vary depending on sequence and genomic location. In addition, using multiple sgRNAs is useful in identifying off-target effects by detecting outlier phenotypes as a result of mutations in non-targeted genes. Effective transfection and Cas9 cleavage will greatly increase chances of generating knockouts. Alternatively, transfecting the cells directly with commercially sourced sgRNA/Cas9 ribonucleoprotein complexes32,33 may expedite the protocol and lower off-target effects.
Dot blots are well-suited to assess protein knockouts caused by the small genomic alterations due to NHEJ. Since only a single sgRNA/Cas9 construct is necessary, this approach may be the quickest in obtaining null lines, and the blot procedure circumvents additional steps, expediting the screening. The normalized immunoblot protein signal serves as a reliable predictor of successful validation (Figure 2B). This protocol is ideal for generation of knockouts with minimal genomic perturbation. Alternatively, this method of screening can be adapted to probe for protein addition or modification, such as transgenic proteins or protein tags. However, this screening method requires an antibody with high specificity to the protein of interest, as cross-reactivity leads to high background in dot blots. This method is naturally limited to mutations targeting protein coding regions, as the final output depends on the presence/absence of a protein product. It is important to also note that mutations caused by NHEJ can lead to a variety of protein outcomes, including a protein with constitutively active, partial or dominant negative function, or a functional protein lacking the detection antibody epitope. To minimize these issues, it is helpful to target the Cas9 cut as early into the coding region as possible, in order to increase the chances of generating a fully non-functional protein. Additionally, targeting genomic regions that are less accessible to the Cas9 machinery can produce low deletion yields in the absence of selection. Finally, knockouts that confer a selective disadvantage may be difficult to obtain by this method for similar reasons.
Using homology-directed repair mechanisms together with CRISPR/Cas9 cleavage allows for much larger and more specific genomic manipulations, which can include both large scale deletions (delineated by two sgRNA cleavages) and insertions. Since two cleavage events and a precise repair process need to take place, integration of a drug resistance marker substantially increases knockout yields using this method. Although dot blot detection can also be used, screening by colony PCR is more generally applicable to accurately detect correct integration/knockout candidates with such large genomic perturbations. Further, testing by PCR allows for identification of clones with imperfect and incomplete integrations, which can provide insight into the extent and nature of the HDR process itself. Additionally, the introduced marker can be subsequently removed (by transient Cre expression or addition in membrane-permeable form), leaving a minimal, 34 bp deletion scar (loxP site) and permitting future re-use of the selection marker. The flexibility of this approach increases with the use of multiple resistance markers, allowing for the generation of multiple cumulative protein knockouts. It is important when applying this method that the absence of product is further validated, for example by RT-qPCR of the expected transcript. Since this screen can be antibody independent, this method is easily applied to protein knockouts without a robust antibody, as well as to detect changes that do not fall in protein coding genes.

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><tbody><tr><td>Competent &lt;em&gt;E. coli&lt;/em&gt; cells</td><td></td><td></td><td></td></tr><tr><td>plasmid prep kit</td><td></td><td></td><td></td></tr><tr><td>pSpCas9(BB) plasmid</td><td>Addgene</td><td>42230</td><td>Ran &lt;em&gt;et al.&lt;/em&gt; 2013, cloning sgRNAs</td></tr><tr><td>BbsI enzyme</td><td>ThermoFisher</td><td>FD1014</td><td>Ran &lt;em&gt;et al.&lt;/em&gt; 2013, cloning sgRNAs</td></tr><tr><td>T7 DNA ligase</td><td>NEB</td><td>M0318L</td><td>Ran &lt;em&gt;et al.&lt;/em&gt; 2013, cloning sgRNAs</td></tr><tr><td>Tango Buffer</td><td>ThermoFisher</td><td>BY5</td><td>Ran &lt;em&gt;et al.&lt;/em&gt; 2013, cloning sgRNAs</td></tr><tr><td>PlasmidSafe exonuclease</td><td>Epicentre</td><td>E3105K</td><td>Ran &lt;em&gt;et al.&lt;/em&gt; 2013, cloning sgRNAs</td></tr><tr><td>Q5 hot start high fidelity polymerase</td><td>NEB</td><td>M0494A</td><td>HA amplification</td></tr><tr><td>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><td>pGolden-Neo</td><td>Addgene</td><td>51422</td><td>Resistance cassette</td></tr><tr><td>pGolden-Hygro</td><td>Addgene</td><td>51423</td><td>Resistance cassette</td></tr><tr><td>BsaI enzyme</td><td>NEB</td><td>R3535S</td><td>Homology arm contruction</td></tr><tr><td>T7 DNA ligase</td><td>NEB</td><td>M0318L</td><td></td></tr><tr><td>PlasmidSafe exonuclease</td><td>Epicentre</td><td>E3105K</td><td></td></tr><tr><td>Toothpicks</td><td></td><td></td><td>bacterial PCR</td></tr><tr><td>Taq polymerase</td><td></td><td></td><td>bacterial colony PCR</td></tr><tr><td>HEK293 human cells</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td>Corning</td><td>10-013-CV</td><td></td></tr><tr><td>FBS</td><td>Corning</td><td>MT35010CV</td><td></td></tr><tr><td>Penicillin-Strep (opt.)</td><td>Gibco</td><td>15140-122</td><td></td></tr><tr><td>6 well plates</td><td>BioLite</td><td>12556004</td><td></td></tr><tr><td>TransIT-LTI Transfection Reagent</td><td>Mirus</td><td>MIR2300</td><td>for lipofection only</td></tr><tr><td>Opti-MEM</td><td>ThermoFisher</td><td>31985062</td><td>for lipofection only</td></tr><tr><td>G418 (Neomycin)</td><td>Sigma Aldrich</td><td>A1720-5G</td><td></td></tr><tr><td>Hygromycin</td><td>Sigma Aldrich</td><td>H3274-250MG</td><td></td></tr><tr><td>96 well plates</td><td>ThermoFisher</td><td>12556008</td><td></td></tr><tr><td>Passive Lysis Buffer, 5x</td><td>Promega</td><td></td><td></td></tr><tr><td>1x SDS loading buffer</td><td></td><td></td><td>recipe decribed in protocol</td></tr><tr><td>Nitrocellulose Membrane</td><td>Bio-Rad</td><td>162-0115</td><td></td></tr><tr><td>TBST</td><td></td><td></td><td>recipe decribed in protocol</td></tr><tr><td>Dehydrated milk</td><td></td><td></td><td></td></tr><tr><td>SuperSignal West Dura Extended Duration Substrate</td><td>ThermoFisher</td><td>34075</td><td>for HRP-conjugated secondary antibodies</td></tr><tr><td>Extracta DNA prep for PCR</td><td>Quantabio</td><td>95091-025</td><td></td></tr><tr><td>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 normalization control) (Figure 2A). The quantified ratio of ELAVL1 to PUM2 signal demonstrated a very broad, &#62;104-fold range among the isolated lines (Figure 2B). Using this ratio as a selection criterion, we validated a number of lines using ELAVL1 western blots (Figure 3A). In most cases, clonal populations with the lowest relative ELAVL1 signal were correctly identified as knockouts (Figure 2B), confirming the predictive power of this quantification (despite considerable variability in the control signal, which likely arose from differing numbers of collected cells). Conversely, most of the clones with intermediate ratios exhibited ELAVL1 signal upon validation, potentially representing monoallelic deletion events. However, no general tiering of the ratios into WT, monoallelic and full (biallelic) mutant pools was observed, and even the ELAVL1-positive clones displayed a wide and continuous range of levels, likely due to clonal variability of expression. The overall effectiveness of ELAVL1 knockout generation using this method ranged from 6 to 36% among the sgRNAs.
For other gene targets (such as PUM2), the quality of the available antibodies prevents accurate screening of candidate clones by dot blot due to high background from cross-reactivity. Additionally, the efficiency of targeting some genomic loci may be substantially lower due to chromatin accessibility. In such cases, a selection-dependent method is appropriate to increase efficiency. Two sgRNAs targeting early and late coding regions of PUM2 were cotransfected with a homology-directed repair template in order to excise 45 kb of genomic DNA and replace it with a neomycin resistance cassette (Figure 1A, right). Assembly of the homology template is illustrated in Figure 1B. PCR primers were designed to amplify regions around the homology arms based on either successful or unsuccessful integration of the resistance cassette (Figure 2C), and were tested on untransfected and the bulk selected cells (left panel). As expected, in untransfected cells, only an amplicon corresponding to the endogenous state (top) was observed, while the bulk selected cells showed amplicons from both wild-type and mutant alleles. Cell lysates from generated clonal candidates were collected and screened for the presence of both the endogenous sequence and the resistance cassette. Candidate knockouts were identified by the presence of the resistance cassette amplicon alone, indicating biallelic deletion/integration (Figure 2D, stars). Clones with monoallelic insertions were also observed, and were fairly common, since such populations can also survive the selection process. These clones were indicated by the presence of both the endogenous sequence and the resistance cassette. In a few cases, clonal wildtypes at the tested locus were also observed, suggesting that random, non-sgRNA-assisted genomic integration of the homology-directed repair template occurs at a low frequency. Biallelic mutant lines were further validated by western blot (Figure 3B). The overall efficiency of knockout generation using the HDR procedure was 33%.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55903/55903fig1.jpg" /> 
Figure 1: A workflow for generating CRISPR-mediated knockouts in mammalian cell lines. (A) Genome editing events produced by CRISPR/Cas9 cleavage and NHEJ (left) or HDR (right). The site of Cas9 cleavage is denoted by a white vertical line. (B) Components and assembly of the HDR template plasmid. The type IIs restriction enzymes used in Golden Gate cloning, such as BsaI, recognize an asymmetric DNA sequence and leave a staggered cut outside of the recognition sequence, allowing for the design of arbitrary overlapping segments for assembly (pictured). Arrows indicate the primer positions to check for correct plasmid assembly. (C) Workflow: after transfection (and drug selection if applicable), cells are plated at limiting dilutions in 96-well plates. Clonal populations are screened either by dot immunoblot (selection-independent, NHEJ) or colony PCR (selection-dependent, HDR). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55903/55903fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55903/55903fig2.jpg" /> 
Figure 2: Screening candidate knockout lines by dot blots or colony PCR. (A) Dot immunoblot of isolated ELAVL1-targeted clones with ELAVL1 antibody (left), and separately with an antibody for a control protein, Pum2 (right). Clones that displayed little control signal, likely due to low input protein amounts, are marked with an X and excluded from further analysis. Clones with low ELAVL1 signal (circles) were further tested by western blots that confirmed (green) or invalidated (grey) the candidates. (B) Quantification of KO candidates from dot blot. Ratios of ELAVL1 to control protein signal in the dot blot shown in A, in increasing order. (C) Primer design for knockout/insertion testing via colony PCR. A primer complementary to the genomic locus outside of the homology arm region (red) is paired with a primer complementary to the endogenous sequence (blue) or to the resistance cassette (green), probing for the unedited state, or a deletion/insertion event, respectively. Separate primer pairs are designed to test both sides of the integration. (D) Example agarose gel of colony PCR results using primers spanning the upstream integration junction using endogenous inside primers (top) and knockout inside primers (bottom). Primers are first validated in both parental T-REx293 cells and bulk selected cells (left). Individual candidate clones (middle) and the expected band patterns for WT, KO and monoallelic deletion (WT/KO) clones (right). Biallelic knockout clones are indicated by a star. The correct amplification products are indicated by black arrows. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55903/55903fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55903/55903fig3.jpg" /> 
Figure 3: Validation of knockout candidates by Western blot. (A) Western blot for ELAVL1 (top) of a subset of clones identified by dot blot. PUM2 was blotted on the same membrane as a loading control (bottom). (B) Western blot for PUM2 (top) of a subset of clones identified by colony PCR. GAPDH was blotted on the same membrane as a loading control (bottom). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55903/55903fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

selection-dependent-independent-generation-crisprcas9-mediated-gene

Nanyang Technological University

Research

JoVE Journal

Genetics

12.4K Views.  University of California, Riverside.  Recente vooruitgang in het vermogen om somatische cellijnen genetisch te manipuleren, biedt een groot potentieel voor basis en toegepast onderzoek. Hier presenteren wij twee benaderingen voor CRISPR / Cas9 gegenereerde knockout productie en screening in zoogdiercellijnen, met en zonder het gebruik van selecteerbare markers. 

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

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 ...

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. ...

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been successfully repurposed for genome engineering in many different living organisms. Most commonly, the wildtype CRISPR associated 9 (Cas9) or Cas12a endonuclease is used to cleave specific sites in the genome, after which the DNA double-stranded break is repaired via the non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway depending on whether a donor template is absent or present respectively. To date, CRISPR systems from different bacterial species have been shown to be capable of performing genome editing in mammalian cells. However, despite the apparent simplicity of the technology, multiple design parameters need to be considered, which often leave users perplexed about how best to carry out their genome editing experiments. Here, we describe a complete workflow from experimental design to identification of cell clones that carry desired DNA modifications, with the goal of facilitating successful execution of genome editing experiments in mammalian cell lines. We highlight key considerations for users to take note of, including the choice of CRISPR system, the spacer length, and the design of a single-stranded oligodeoxynucleotide (ssODN) donor template. We envision that this workflow will be useful for gene knockout studies, disease modeling efforts, or the generation of reporter cell lines.

CRISPR-Cas is a powerful technology to engineer the complex genomes of plants and animals. Here, we detail a protocol to efficiently edit the human genome using different Cas endonucleases. We highlight important considerations and design parameters to optimize editing efficiency.

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been ...

Pooled CRISPR-Based Genetic Screens in Mammalian Cells

Genome editing using the CRISPR-Cas system has vastly advanced the ability to precisely edit the genomes of various organisms. In the context of mammalian cells, this technology represents a novel means to perform genome-wide genetic screens for functional genomics studies. Libraries of guide RNAs (sgRNA) targeting all open reading frames permit the facile generation of thousands of genetic perturbations in a single pool of cells that can be screened for specific phenotypes to implicate gene function and cellular processes in an unbiased and systematic way. CRISPR-Cas screens provide researchers with a simple, efficient, and inexpensive method to uncover the genetic blueprints for cellular phenotypes. Furthermore, differential analysis of screens performed in various cell lines and from different cancer types can identify genes that are contextually essential in tumor cells, revealing potential targets for specific anticancer therapies. Performing genome-wide screens in human cells can be daunting, as this involves the handling of tens of millions of cells and requires analysis of large sets of data. The details of these screens, such as cell line characterization, CRISPR library considerations, and understanding the limitations and capabilities of CRISPR technology during analysis, are often overlooked. Provided here is a detailed protocol for the successful performance of pooled genome-wide CRISPR-Cas9 based screens.

CRISPR-Cas9 technology provides an efficient method to precisely edit the mammalian genome in any cell type and represents a novel means to perform genome-wide genetic screens. A detailed protocol discussing the steps required for the successful performance of pooled genome-wide CRISPR-Cas9 screens is provided here.

Genome editing using the CRISPR-Cas system has vastly advanced the ability to precisely edit the genomes of various organisms. In the context of mammalian ...

A Protocol for Multiple Gene Knockout in Mouse Small Intestinal Organoids Using a CRISPR-concatemer

CRISPR/Cas9 technology has greatly improved the feasibility and speed of loss-of-function studies that are essential in understanding gene function. In higher eukaryotes, paralogous genes can mask a potential phenotype by compensating the loss of a gene, thus limiting the information that can be obtained from genetic studies relying on single gene knockouts. We have developed a novel, rapid cloning method for guide RNA (gRNA) concatemers in order to create multi-gene knockouts following a single round of transfection in mouse small intestinal organoids. Our strategy allows for the concatemerization of up to four individual gRNAs into a single vector by performing a single Golden Gate shuffling reaction with annealed gRNA oligos and a pre-designed retroviral vector. This allows either the simultaneous knockout of up to four different genes, or increased knockout efficiency following the targeting of one gene by multiple gRNAs. In this protocol, we show in detail how to efficiently clone multiple gRNAs into the retroviral CRISPR-concatemer vector and how to achieve highly efficient electroporation in intestinal organoids. As an example, we show that simultaneous knockout of two pairs of genes encoding negative regulators of the Wnt signaling pathway (Axin1/2 and Rnf43/Znrf3) renders intestinal organoids resistant to the withdrawal of key growth factors.

This protocol describes the steps for cloning multiple single guide RNAs into one guide RNA concatemer vector, which is of particular use in creating multi-gene knockouts using CRISPR/Cas9 technology. The generation of double knockouts in intestinal organoids is shown as a possible application of this method.

CRISPR/Cas9 technology has greatly improved the feasibility and speed of loss-of-function studies that are essential in understanding gene function. In ...

Bioengineering

Construction of CRISPR Plasmids and Detection of Knockout Efficiency in Mammalian Cells through a Dual Luciferase Reporter System

Although highly efficient, modification of a genomic site by the CRISPR enzyme requires the generation of a sgRNA unique to the target site(s) beforehand. This work describes the key steps leading to the construction of efficient sgRNA vectors using a strategy that allows the efficient detection of the positive colonies by PCR prior to DNA sequencing. Since efficient genome editing using the CRISPR system requires a highly efficient sgRNA, a preselection of candidate sgRNA targets is necessary to save time and effort. A dual luciferase reporter system has been developed to evaluate knockout efficiency by examining double-strand break repair via single strand annealing. Here, we use this reporter system to pick up the preferred xCas9/sgRNA target from candidate sgRNA vectors for specific gene editing. The protocol outlined will provide a preferred sgRNA/CRISPR enzyme vector in 10 days (starting with appropriately designed oligonucleotides).

Here, we present a protocol describing a streamlined method for the efficient generation of plasmids expressing both the CRISPR enzyme and associated single guide RNA (sgRNAs). Co-transfection of mammalian cells with this sgRNA/CRISPR vector and a dual luciferase reporter vector that examines double-strand break repair allows evaluation of knockout efficiency.

Although highly efficient, modification of a genomic site by the CRISPR enzyme requires the generation of a sgRNA unique to the target site(s) beforehand. ...

Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific eukaryotic genome engineering. CRISPR/Cas9 is an inexpensive, facile, and efficient genome editing tool that allows genetic perturbation of genes and genetic elements. Here we present a simple methodology for CRISPR design, cloning, and delivery for the production of genomic deletions. In addition, we describe techniques for deletion, identification, and characterization. This strategy relies on cellular delivery of a pair of chimeric single guide RNAs (sgRNAs) to create two double strand breaks (DSBs) at a locus in order to delete the intervening DNA segment by non-homologous end joining (NHEJ) repair. Deletions have potential advantages as compared to single-site small indels given the efficiency of biallelic modification, ease of rapid identification by PCR, predictability of loss-of-function, and utility for the study of non-coding elements. This approach can be used for efficient loss-of-function studies of genes and genetic elements in mammalian cell lines.

CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific ...

Biology

Electroporation-Based CRISPR-Cas9-Mediated Gene Knockout in THP-1 Cells and Single-Cell Clone Isolation

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including their interplay with significant human pathogens such as the human immunodeficiency virus (HIV). Compared to other immortalized cell lines of myeloid origin, THP-1 cells retain many intact inflammatory signaling pathways and display phenotypic characteristics that more closely resemble those of primary monocytes, including the ability to differentiate into macrophages when treated with phorbol-12-myristate 13-acetate (PMA). The use of CRISPR-Cas9 technology to engineer THP-1 cells through targeted gene knockout (KO) provides a powerful approach to better characterize immune-related mechanisms, including virus-host interactions. This article describes a protocol for efficient CRISPR-Cas9-based engineering using electroporation to deliver pre-assembled Cas9:sgRNA ribonucleoproteins into the cell nucleus. Using multiple sgRNAs targeting the same locus at slightly different positions results in the deletion of large DNA fragments, thereby increasing editing efficiency, as assessed by the T7 endonuclease I assay. CRISPR-Cas9-mediated editing at the genetic level was validated by Sanger sequencing followed by Inference of CRISPR Edits (ICE) analysis. Protein depletion was confirmed by immunoblotting coupled with a functional assay. Using this protocol, up to 100% indels in the targeted locus and a decrease of over 95% in protein expression were achieved. The high editing efficiency makes it convenient to isolate single-cell clones by limiting dilution.

The THP-1 cell line is widely used as a model to investigate the functions of human monocytes/macrophages across various biology-related research areas. This article describes a protocol for efficient CRISPR-Cas9-based engineering and single-cell clone isolation, enabling the production of robust and reproducible phenotypic data.

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including ...

Immunology and Infection

CRISPR-Cas9 Mediated Gene Deletion in Human Pluripotent Stem Cells Cultured Under Feeder-Free Conditions

The CRISPR-Cas9 system for genome editing has revolutionized gene function studies in mammalian cells, including stem cells. However, the practical application of this technique, particularly in pluripotent stem cells, presents certain challenges, such as being time- and labor-intensive and having low editing efficiency. Here, we describe the generation of a CRISPR-mediated gene knockout in a human embryonic stem cell (hESC) line stably expressing sgRNAs for the L2HGDH gene, using a highly efficient and stable lentiviral-mediated gene delivery system. The sgRNAs targeting exon 1 of the L2HGDH gene were chemically synthesized and cloned into the lentiCRISPR v2-puro vector, which combines the constitutive expression of sgRNAs with Cas9 in a highly efficient single-vector system to achieve higher lentiviral titers for hESC infection and stable selection using puromycin. Puromycin-selected cells were further expanded, and single-cell clones were obtained using the limited dilution method. The single clones were expanded, and several homozygous knockout clones for the L2HGDH gene were obtained, as confirmed by a 100% reduction in L2HGDH expression using Western blot analysis. Furthermore, using MSBSP-PCR, the CRISPR mutation site was mapped upstream of the PAM recognition sequence of Cas9 in the selected homozygous clones. Sanger sequencing was performed to analyze the exact insertions/deletions, and functional characterization of the clones was conducted. This method produced a significantly higher percentage of homozygous deletions compared to previously reported non-viral gene delivery methods. Although this report focuses on the L2HGDH gene, this robust and cost-effective approach can be used to create homozygous knockouts for other genes in pluripotent stem cells for gene function studies.

The presented method describes the generation of a CRISPR-mediated gene knockout in the human embryonic stem cell (hESC) line H9, which stably expresses sgRNAs targeting the L2HGDH gene using a highly efficient lentiviral-mediated gene delivery system.

The CRISPR-Cas9 system for genome editing has revolutionized gene function studies in mammalian cells, including stem cells. However, the practical ...

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

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