This work was supported by NIH/NHGRI R00 HG006922 and NIH/NHGRI R01 HG008974 to J.G., and the Huntsman Cancer Institute. J.B.C. was supported by NIH Training Program in Genetics T32GM007464.

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The authors have nothing to disclose.

This protocol describes a simple and flexible method for dissecting enhancer function at the endogenous genomic locus without physically altering the DNA sequence. While similar in concept to previously published CRISPR interference protocols using dCas9-KRAB27, Enhancer-i differs from these protocols in 3 main ways. First, Enhancer-i utilizes the SIN3A interacting domain of MAD120 to achieve enhancer deactivation. Enhancer deactivation can be rescued using HDAC inhibitors, suggesting that the primary mechanism of deactivation is HDAC dependent. Unlike CRISPR interference with dCas9-KRAB, Enhancer-i does not lead to the deposition of H3K9me3. This is likely due to the fact that Enhancer-i relies on transient introduction of guide RNAs, with the cells being harvested at 3 days post transfection. In CRISPR interference, an increase in H3K9me3 is observed at 7 days post transduction12. Finally, the Enhancer-i protocol provides a strategy to target multiple sites simultaneously and monitor the efficiency of targeting. In Mosaic-seq17, dCas9-KRAB is used to target multiple enhancers simultaneously, but this technique relies on single-cell RNA sequencing to identify expression changes, and many genes (such as estrogen-responsive genes) go undetected due to the low sensitivity of single-cell RNA-seq. Enhancer-i provides a reliable method to study enhancers individually and in combination for any gene.
The most critical step of Enhancer-i is transfection, which should be optimized for the cell line of interest. This protocol relies on puromycin treatment to enrich for transfected cells, but it is possible that co-transfecting guide RNAs with a fluorescent protein and sorting for fluorescent cells using flow cytometry may prove to be a better enrichment method for some cell types. We recommend monitoring the expression level of guide RNAs and SID4x-dCas9-KRAB by qPCR to troubleshoot and confirm transfection. If guide RNA levels are low (cycle threshold &#62;30), users may also consider alternative guide RNA production strategies such as in vitro transcription28. It is also possible that despite high gRNA levels, guide RNA targeting of the SID4x-dCas9-KRAB protein is inefficient, in which case selecting different guide RNA sequences may be necessary. By performing ChIP-seq on the fusion protein with chromatin from Enhancer-i treated cells, the efficiency of targeting can be monitored. If there is high signal of SID4x-dCas9-KRAB at the region of interest, and no expression changes in its putative target gene are detected, then the region likely does not contribute to the expression of that gene under the conditions studied.
One potential limitation of Enhancer-i is that off-target effects may accumulate if too many sites are targeted simultaneously. Nonetheless, CRISPR interference strategies for knockdown have fewer off target effects than RNAi29, particularly when a polyclonal cell line expressing dCas9-KRAB is used. While we have seen off-target genomic binding of SID4X-dCas9-KRAB when targeting 10 sites simultaneously, we have not identified gene expression changes as a result of those binding events. As some enhancers may contact multiple promoters and/or other enhancers, it is possible that many genes may change expression upon targeting of a single enhancer, though it is unclear if this form of gene regulation is common. To confirm that the expression changes observed are due to targeting a specific enhancer, and not off-target effects, users can perform Enhancer-i with two distinct sets of non-overlapping guide RNAs targeting the same region. In addition, the genetic deletion of the region using nuclease-competent Cas9 can further confirm its effects on gene expression.
As Enhancer-i functions through histone deacetylation, it is possible that its deactivation abilities are limited to enhancers that have appreciable levels of histone acetylation. There are a variety of alternative repressive fusions that may be more effective at targeting specific enhancers. DNA methyltransferase fusions to dCas9 can be used to reduce gene expression when targeted to distal enhancers30, but this repression is often not transient. Another repressive fusion uses the Friend of GATA1 (FOG1) domain, which leads to histone H3 lysine 27 trimethylation and represses gene expression at levels similar to dCas9-KRAB across a variety of cell lines and promoters31. Interestingly, adding more copies of FOG1 to dCas9 reduced the repressive potential at promoters, suggesting that a single copy of the SID domain may provide more enhancer deactivation than the 4 copies currently used in Enhancer-i. It is possible that some loci may benefit from dual targeting by different combinations of the above dCas9 fusions. For example, stable long-term repression can be achieved by simultaneous transduction of dCas9-DNMT3a and dCas9-KRAB32. Most of these repressive fusions have only been targeted to a single locus at a time, and it remains unclear which is most effective at manipulating multiple enhancers simultaneously.
Enhancer-i, while a suitable method for studying combinations of enhancers for a handful of genes, is still somewhat limited in throughput if the user wishes to study putative enhancers for hundreds of genes. Future applications of this technique will incorporate imaging-based technologies to quantify multiple genes in multiple samples simultaneously. Importantly, these technologies are compatible with direct detection of RNA molecules from lysate, eliminating the need for time-consuming RNA isolation. These adaptations will facilitate the interrogation of larger sets of enhancers.

Large-scale sequencing projects such as ENCODE1, Roadmap Epigenomics2, and FANTOM3 have identified millions of putative enhancers within the human genome across hundreds of cell types. It is estimated that each promoter associates with an average of 4.9 enhancers and each enhancer contacts an average of 2.4 genes3, suggesting that gene expression is often the result of the integration of multiple distributed regulatory interactions. A significant remaining challenge is to define not only how individual enhancers contribute to gene expression, but how they combine to affect expression. Genetic approaches are commonly used to identify relationships between enhancers in model organisms from Drosophila4 to mice5. However, these experiments are time-consuming and low-throughput for the study of multiple enhancers at multiple genes.
One approach for studying enhancer function on a large scale involves massively parallel reporter assays. These assays allow for the simultaneous screening of thousands of DNA sequences for their ability to drive the expression of a reporter gene6. While these assays have shown that DNA sequence can alone be sufficient to convey gene regulation information7, they come with the caveats of being performed outside of the native chromatin context and with a heterologous promoter. In addition, the size of DNA sequence being analyzed in massively parallel reporter assays is usually less than 200 basepairs, which may exclude relevant surrounding sequence. Importantly, as reporter assays only measure the activity of one sequence at a time, they do not take into account the complex relationships that can exist between enhancers. Thus, while massively parallel reporter assays can be informative about the intrinsic activity of a DNA sequence, they do not necessarily inform us of the function of that DNA sequence in the context of the genome.
Recently developed CRISPR/Cas9 tools8 have facilitated the study of gene regulation as they allow for the deletion of enhancers at the endogenous locus. However, deleting multiple enhancers simultaneously may lead to genomic instability, and it is time consuming to generate successive enhancer deletions in a single cell line. In addition, new genomic sequence is created at the site of the deletion following repair, and this sequence may gain regulatory function. An alternative version of Cas9 has been developed specifically for modulating gene expression, relying on fusions of activating9,10 or repressing11,12 domains to the nuclease-deficient form of Cas9 (dCas9). These fusion proteins are ideal for studying multiple loci simultaneously as they do not physically alter the DNA sequence, and instead modulate epigenetics in order to interrogate a regulatory region. The most widely used repressive fusion is KRAB, which recruits the KAP1 co-repressor complex, promoting the deposition of the repression-associated histone H3 lysine 9 trimethylation (H3K9me3)13. dCas9-KRAB, also known as CRISPR interference14, has been used to target and screen individual enhancers for their contributions to gene expression15,16; however, it has not been optimized for targeting multiple regions simultaneously. One version of multiplex CRISPR interference for enhancers, Mosaic-seq17, uses single cell RNA-seq as a readout, but this technology is expensive and only suitable for the study of highly expressed genes due to the low sensitivity of single cell RNA-seq.
We sought to develop a CRISPR interference-based method for dissecting combinatorial enhancer function within the context of a transcriptional response to estrogen. About half of estrogen-responsive genes contain 2 or more enhancers bound by estrogen receptor alpha (ER) nearby18, suggesting that multiple enhancers may be participating in the estrogen response, and understanding the regulatory logic would require targeting multiple enhancers simultaneously. As initial studies using CRISPR interference at promoters suggested that not all promoters are equally responsive to KRAB-mediated repression19, we reasoned that the addition of a distinct repressive domain to dCas9 may facilitate the deactivation of diverse enhancers. We chose the Sin3a Interacting Domain of Mad1 (SID)20 as it leads to the recruitment of histone deacetylases21, which remove acetyl groups on histones that are associated with transcriptional activity. Importantly, the SID domain was effective at reducing gene expression when fused to dCas922 and TALEs23, and Sin3a has been shown to be a potent repressive co-factor in a variety of enhancer sequence contexts24. We used SID4x-dCas9-KRAB (Enhancer-i) to target 10 different enhancers bound by the ER, and identify ER binding sites (ERBS) that are necessary for the estrogen transcriptional response at 4 genes18. We also targeted the combinations of enhancers to identify the sites that cooperate in the production of the estrogen transcriptional response. We found that up to 50 sites can potentially be targeted simultaneously with detectable gene expression changes. Using ChIP-seq and RNA-seq, we demonstrated that Enhancer-i is a highly specific technique for studying multiple enhancers simultaneously.
In this protocol, we describe the steps involved in performing Enhancer-i, a flexible technique that enables the functional study of multiple enhancers simultaneously in a tissue culture setting. Enhancer-i is highly correlated with genetic deletion but provides transient deactivation that is dependent on histone deacetylases (HDACs). By delivering guide RNAs via transient transfection as opposed to stable integration via viral vectors, this protocol avoids deposition and potential spreading of H3K9me3. This protocol details guide RNA design and cloning via Gibson assembly, the transfection of guide RNAs using lipofection, and the analysis of resulting gene expression changes by qPCR. We also include the methods for evaluating the specificity of Enhancer-i targeting at the level of the genome and transcriptome. While this technique was developed to study gene regulation by ER bound enhancers in human cancer cell lines, it is applicable to the dissection of any mammalian enhancer.

<table><tbody><tr><td>ZR 96-well Quick-RNA Kit</td><td>Zymo Research</td><td>R1053</td><td></td></tr><tr><td>Power SYBR Green RNA-to-CT 1-Step</td><td>Applied Biosystems</td><td>4389986</td><td></td></tr><tr><td>AflII restriction enzyme</td><td>NEB</td><td>R0520S</td><td></td></tr><tr><td>Phusion High-Fidelity PCR Master Mix with HF Buffer</td><td>NEB</td><td>M0531L</td><td></td></tr><tr><td>NEBuilder HiFi DNA Assembly Master Mix</td><td>NEB</td><td>E2621L</td><td></td></tr><tr><td>FuGENE HD</td><td>Promega</td><td>E2312</td><td></td></tr><tr><td>DNA Clean &amp;amp; Concentrator Kit</td><td>Zymo Research</td><td>D4013</td><td></td></tr><tr><td>Buffer RLT Plus</td><td>Qiagen</td><td>1053393</td><td></td></tr><tr><td>b-estradiol</td><td>Sigma-Aldrich</td><td>E2758</td><td></td></tr><tr><td>Human: Ishikawa cells</td><td>ECACC</td><td>99040201</td><td></td></tr><tr><td>H3K27ac rabbit polyclonal</td><td>Active Motif&amp;nbsp;</td><td>39133</td><td></td></tr><tr><td>H3K9me3 rabbit polyclonal</td><td>Abcam&amp;nbsp;</td><td>ab8898</td><td></td></tr><tr><td>FLAG mouse monoclonal</td><td>Sigma-Aldrich&amp;nbsp;</td><td>F1804</td><td></td></tr><tr><td>ER alpha rabbit polyclonal</td><td>Santa Cruz</td><td>sc-544</td><td></td></tr><tr><td>pGL3-U6-PGK-Puro plasmid</td><td>Addgene&amp;nbsp;</td><td>51133</td><td>Shen et al., 2014</td></tr><tr><td>gRNA_cloningVector plasmid</td><td>Addgene&amp;nbsp;</td><td>41824</td><td>Mali et al., 2013</td></tr><tr><td>AflII U6 puromycin plasmid</td><td>Addgene&amp;nbsp;</td><td>106404</td><td>Carleton et al., 2017</td></tr><tr><td>SID4X-dCas9-KRAB plasmid</td><td>Addgene&amp;nbsp;</td><td>106399</td><td>Carleton et al., 2017</td></tr><tr><td>2-Mercaptoethanol</td><td>Sigma-Aldrich</td><td>M6250-10ML</td><td></td></tr><tr><td>UltraPure DNase/RNase-Free Distilled Water</td><td>ThermoFisher Scientific</td><td>10977-023</td><td></td></tr><tr><td>Opti-MEM I Reduced Serum Medium</td><td>Gibco</td><td>31985070</td><td></td></tr><tr><td>KAPA Stranded mRNA-Seq Kit, with KAPA mRNA Capture Beads</td><td>Kapa Biosytems</td><td>KK8420</td><td></td></tr><tr><td>Pierce Protease and Phosphatase Inhibitor Mini Tablets</td><td>ThermoFisher Scientific</td><td>A32959</td><td></td></tr><tr><td>Formaldehyde solution</td><td>Sigma-Aldrich</td><td>252549-25ML</td><td></td></tr><tr><td>Geneticin Selective Antibiotic (G418 Sulfate) (50 mg/mL)</td><td>ThermoFisher Scientific</td><td>10131035</td><td></td></tr><tr><td>LB Broth</td><td>ThermoFisher Scientific</td><td>10855001</td><td></td></tr><tr><td>Quick-DNA Miniprep Kit</td><td>Zymo Research</td><td>D3020</td><td></td></tr><tr><td>Quick-Load Purple 2-Log DNA Ladder</td><td>NEB</td><td>N0050S</td><td></td></tr></tbody></table>

dissection of enhancer function using multiplex crispr-based enhancer interference in cell lines

Multiple enhancers often regulate a given gene, yet for most genes, it remains unclear which enhancers are necessary for gene expression, and how these enhancers combine to produce a transcriptional response. As millions of enhancers have been identified, high-throughput tools are needed to determine enhancer function on a genome-wide scale. Current methods for studying enhancer function include making genetic deletions using nuclease-proficient Cas9, but it is difficult to study the combinatorial effects of multiple enhancers using this technique, as multiple successive clonal cell lines must be generated. Here, we present Enhancer-i, a CRISPR interference-based method that allows for functional interrogation of multiple enhancers simultaneously at their endogenous loci. Enhancer-i makes use of two repressive domains fused to nuclease-deficient Cas9, SID and KRAB, to achieve enhancer deactivation via histone deacetylation at targeted loci. This protocol utilizes transient transfection of guide RNAs to enable transient inactivation of targeted regions and is particularly effective at blocking inducible transcriptional responses to stimuli in tissue culture settings. Enhancer-i is highly specific both in its genomic targeting and its effects on global gene expression. Results obtained from this protocol help to understand whether an enhancer is contributing to gene expression, the magnitude of the contribution, and how the contribution is affected by other nearby enhancers.

Figure 1 shows a schematic of the workflow described in the protocol. To determine the contributions of ER-bound enhancers near the estrogen-regulated gene MMP17, which has 3 binding sites nearby as defined by ChIP-seq (Figure 2A), guide RNAs were designed for each region. To design guide RNAs, a 600 - 900 bp window of sequence surrounding each ER binding site of interest was selected and put into a guide RNA design program. Resulting guide RNA sequences with 0-2 predicted off target sites were aligned to the human genome using BLAT. Four non-overlapping guide RNAs that spanned the region defined by ChIP-seq and DNaseI hypersensitivity were chosen for targeting (Figure 2B). Additional sequence (Table 1) was added to each end to facilitate downstream cloning and the resulting 59 nucleotide fragments were ordered. Upon arrival, guide RNAs were diluted and pooled by site, and a short PCR was performed to add homology regions prior to Gibson assembly. Figure 2C shows the expected guide RNA product after a short PCR using the &#34;U6_internal&#34; primers (Table 1), which will add 20 basepairs of sequence to each end of the 59 basepair guide RNA fragment, resulting in a ~100 basepair sequence. Following Gibson assembly, these guide RNA pools were transformed into bacteria and plasmid minipreps were prepared the following day. Figure 2D shows results from an enhancer dissection experiment, where multiple enhancers nearby MMP17 are targeted alone and in combination using Enhancer-i. Sites targeted by Enhancer-i are indicated with a black hexagon. Guide RNA plasmids targeting the indicated sites were transfected into an estrogen-deprived Ishikawa cell line stably expressing SID4X-dCas9-KRAB. Two days later, the media was changed and puromycin was added to enrich for transfected cells. The following day, the cells were harvested following an 8 h 10 nM estradiol treatment. RNA was isolated, and a one-step qPCR was performed. In this example, sites 1 and 2 are necessary for a complete estrogenic response of MMP17, while site 3 does not contribute under these conditions (Figure 2D, lanes ii-iv). When only sites 2 or 3 are active (vi and vii), the estrogen response is similar to when no sites are active (viii), suggesting that these sites cannot contribute independently. Site 1 can contribute some expression by itself (v), but the greatest activity is seen when sites 1 and 2 are active (iv).
To manipulate 10 enhancers near 4 different genes simultaneously (Figure 3A), complex pools of guide RNAs were generated containing 42 enhancer guides and 16 promoter guides. Guide RNA oligos were pooled before the initial guide RNA extension PCR (Step 3.3), and resulting PCR products were purified and combined with the empty puromycin U6 cloning vector using Gibson assembly. Following the Gibson assembly, multiple independent transformations were performed and plated. The plates were scraped into LB and allowed to grow out for 2 - 4 h prior to maxiprep. Figure 3B shows representative reductions in gene expression by qPCR when these guide RNA pools were transfected into an estrogen-deprived Ishikawa cell line stably expressing SID4X-dCas9-KRAB and treated as described above (Figure 2D). Reductions from Enhancer-i are similar to those obtained by targeting the promoter of the putative target gene. Figure 3C shows the effects of dilution of guide RNAs on reduction of the estrogen response using Enhancer-i. A 1:50 dilution of a guide RNA pool targeting the enhancer near G0S2 still yields significant reduction in gene expression, suggesting that Enhancer-i can be used to target up to 50 sites at once. However, the deactivation can be diluted out, indicating that hundreds of sites cannot be targeted simultaneously unless more sensitive detection methods are employed.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57883/57883fig1.jpg" /> 
Figure 1. Protocol schematic for multiplex enhancer dissection using Enhancer-i. Guide RNAs (red and blue) are designed using e-crisp and selected using the UCSC genome browser. Four guide RNAs are chosen that span the regions of interest (transcription factor binding sites as defined by ChIP-seq). Guide RNA oligonucleotides that have been pooled by the region of interest (red and blue) undergo a PCR to add homology regions (orange) prior to the Gibson assembly and transformation. Resulting plasmid pools are transfected via lipofection into cell lines stably expressing SID4X-dCas9-KRAB or into wild-type cells in conjunction with SID4X-dCas9-KRAB plasmid. Guide RNA plasmid pools can be transfected individually to target one site at a time, or in combination to target multiple sites simultaneously. Transfected cells are treated with antibiotics to enrich for cells containing guide RNAs. At ~72 h post transfection, the cells are harvested. Nucleic acids can be extracted for qPCR, RNA-seq, or ChIP-seq. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57883/57883fig1large.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/57883/57883fig2.jpg" /> 
Figure 2. Guide RNA design and enhancer dissection for MMP17. (A) Genome browser screenshot of the ER alpha-bound enhancers (gray) to be targeted near MMP17. This figure has been modified from Carleton, et al.18. (B) Guide RNA designs for the 3 binding sites18. The binding site for ER as defined by ChIP-seq is the target, and the 4 guide RNAs tile across this region. The DNaseI sensitivity signal, which spans the binding site, can also be used to define target sequence for guide RNA design. Both ChIP-seq and DNaseI HS data were obtained from Ishikawa cells treated with 10 nM estradiol for 1 h. (C) Representative guide RNA sequences that are ready for Gibson assembly, having undergone a short PCR to add homology regions. (D) Relative expression of MMP17 measured via qPCR following targeting of specific regions with Enhancer-i and an 8-h10 nM estradiol treatment. Expression is relative to CTCF and expression level of MMP17 in cells not treated with estradiol. Control guide RNAs target the promoter of IL1RN. All error bars represent SEM, double asterisks indicate p &#60;0.01 and single asterisks indicate p &#60;0.05 in a paired t-test. This figure has been modified from Carleton, et al.18. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57883/57883fig2large.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/57883/57883fig3.jpg" /> 
Figure 3. Targeting multiple enhancers near different genes simultaneously with pooled Enhancer-i. (A) Schematic of the binding sites and promoters to be targeted in pooled Enhancer-i. (B) The effects on expression as measured by qPCR after E2 treatment on Ishikawa cells transfected with Enhancer-i plasmid pool (green), Promoter-i plasmid pool (blue) or control gRNAs (white)18. A significant reduction at all genes is observed with Enhancer-i. This figure was modified from Carleton, et al.18. (C) The effects on G0S2 expression levels after E2 treatment on Ishikawa cells transfected with different amounts of guide RNAs targeting G0S2. A significant reduction can be seen even with small amounts of guide RNA (1:50 dilution), suggesting that up to 50 sites may targeted simultaneously. All error bars represent SEM, double asterisks indicate p &#60;0.01 and single asterisks indicate p &#60;0.05 in a paired t-test. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57883/57883fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Name</td>
			<td>Sequence</td>
		</tr>
		<tr>
			<td>U6_internal_F</td>
			<td>TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCG</td>
		</tr>
		<tr>
			<td>U6_internal_R&#160;</td>
			<td>GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAAC</td>
		</tr>
		<tr>
			<td>U6_PCR_F</td>
			<td>CCAATTCAGTCGACTGGATCCGGTA</td>
		</tr>
		<tr>
			<td>U6_PCR_R</td>
			<td>AAAAAAAGCACCGACTCGGTGCCA</td>
		</tr>
		<tr>
			<td>gRNA_qPCR_F</td>
			<td>GCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCG</td>
		</tr>
		<tr>
			<td>gRNA_qPCR_R</td>
			<td>AAAAAGCACCGACTCGGTGCC</td>
		</tr>
		<tr>
			<td>dCas9_qPCR_F</td>
			<td>GTGACCGAGGGAATGAGAAA</td>
		</tr>
		<tr>
			<td>dCas9_qPCR_R</td>
			<td>AGCTGCTTCACGGTCACTTT</td>
		</tr>
		<tr>
			<td>pAC95_PCR_F</td>
			<td>AGAAGAGAAAGGTGGAGGCC</td>
		</tr>
		<tr>
			<td>pAC95_PCR_R</td>
			<td>CGTCACCGCATGTTAGAAGG</td>
		</tr>
		<tr>
			<td>SID4X_PCR_F</td>
			<td>CAATAGAAACTGGGCTTGTCG</td>
		</tr>
		<tr>
			<td>SID4X_PCR_R</td>
			<td>TCGTGCTTCTTATCCTCTTCC</td>
		</tr>
	</tbody>
</table>
Table 1. Primers used for guide RNA extension and sequencing, qPCR, and detection of the fusion protein.

Watch this Scientific Journal Video about Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines at JoVE.com

Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

This protocol describes the steps needed to design and perform multiplexed targeting of enhancers with the deactivating fusion protein SID4X-dCas9-KRAB, also known as enhancer interference (Enhancer-i). This protocol enables the identification of enhancers that regulate gene expression and facilitates the dissection of relationships between enhancers regulating a common target gene.

Multiple enhancers often regulate a given gene, yet for most genes, it remains unclear which enhancers are necessary for gene expression, and how these ...

dissection-enhancer-function-using-multiplex-crispr-based-enhancer

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Genetics

9.2K Views.  University of Utah. This protocol describes the steps needed to design and perform multiplexed targeting of enhancers with the deactivating fusion protein SID4X-dCas9-KRAB, also known as enhancer interference (Enhancer-i). This protocol enables the identification of enhancers that regulate gene expression and facilitates the dissection of relationships between enhancers regulating a common target gene.

Video: Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

Efficient Generation of hiPSC Neural Lineage Specific Knockin Reporters Using the CRISPR/Cas9 and Cas9 Double Nickase System

Gene targeting is a critical approach for characterizing gene functions in modern biomedical research. However, the efficiency of gene targeting in human cells has been low, which prevents the generation of human cell lines at a desired rate. The past two years have witnessed a rapid progression on improving efficiency of genetic manipulation by genome editing tools such as the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas (CRISPR-associated) system. This manuscript describes a protocol for generating lineage specific human induced pluripotent stem cell (hiPSC) reporters using CRISPR/Cas system assisted homologous recombination. Procedures for obtaining necessary components for making neural lineage reporter lines using the CRISPR/Cas system, focusing on construction of targeting vectors and single guide RNAs, are described. This protocol can be extended to platform establishment and mutation correction in hiPSCs.

Genome editing tools such as the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas (CRISPR-associated) system have greatly improved gene targeting efficiency in human induced pluripotent stem cells (hiPSCs). This manuscript describes a protocol for generating lineage specific hiPSC reporter using CRISPR/Cas system assisted homologous recombination.

Gene targeting is a critical approach for characterizing gene functions in modern biomedical research. However, the efficiency of gene targeting in human ...

Developmental Biology

Genome Editing and Directed Differentiation of hPSCs for Interrogating Lineage Determinants in Human Pancreatic Development

Interrogating gene function in self-renewing or differentiating human pluripotent stem cells (hPSCs) offers a valuable platform towards understanding human development and dissecting disease mechanisms in a dish. To capitalize on this potential application requires efficient genome-editing tools to generate hPSC mutants in disease-associated genes, as well as in vitro hPSC differentiation protocols to produce disease-relevant cell types that closely recapitulate their in vivo counterparts. An efficient genome-editing platform for hPSCs named iCRISPR has been developed through the TALEN-mediated targeting of a Cas9 expression cassette in the AAVS1 locus. Here, the protocols for the generation of inducible Cas9 hPSC lines using cells cultured in a chemically defined medium and a feeder-free condition are described. Detailed procedures for using the iCRISPR system for gene knockout or precise genetic alterations in hPSCs, either through non-homologous end joining (NHEJ) or via precise nucleotide alterations using a homology-directed repair (HDR) template, respectively, are included. These technical procedures include descriptions of the design, production, and transfection of CRISPR guide RNAs (gRNAs); the measurement of the CRISPR mutation rate by T7E1 or RFLP assays; and the establishment and validation of clonal mutant lines. Finally, we chronicle procedures for hPSC differentiation into glucose-responsive pancreatic &#946;-like cells by mimicking in vivo pancreatic embryonic development. Combining iCRISPR technology with directed hPSC differentiation enables the systematic examination of gene function to further our understanding of pancreatic development and diabetes disease mechanisms.

Protocols to generate hPSC mutant lines using the iCRISPR platform and to differentiate hPSCs into glucose-responsive &#946;-like cells are described. Combining genome editing technology with hPSC-directed differentiation provides a powerful platform for the systematic analysis of the role of lineage determinants in human development and disease progression.

Interrogating gene function in self-renewing or differentiating human pluripotent stem cells (hPSCs) offers a valuable platform towards understanding ...

Generating CRISPR/Cas9 Mediated Monoallelic Deletions to Study Enhancer Function in Mouse Embryonic Stem Cells

Enhancers control cell identity by regulating tissue-specific gene expression in a position and orientation independent manner. These enhancers are often located distally from the regulated gene in intergenic regions or even within the body of another gene. The position independent nature of enhancer activity makes it difficult to match enhancers with the genes they regulate. Deletion of an enhancer region provides direct evidence for enhancer activity and is the gold standard to reveal an enhancer's role in endogenous gene transcription. Conventional homologous recombination based deletion methods have been surpassed by recent advances in genome editing technology which enable rapid and precisely located changes to the genomes of numerous model organisms. CRISPR/Cas9 mediated genome editing can be used to manipulate the genome in many cell types and organisms rapidly and cost effectively, due to the ease with which Cas9 can be targeted to the genome by a guide RNA from a bespoke expression plasmid. Homozygous deletion of essential gene regulatory elements might lead to lethality or alter cellular phenotype whereas monoallelic deletion of transcriptional enhancers allows for the study of cis-regulation of gene expression without this confounding issue. Presented here is a protocol for CRISPR/Cas9 mediated deletion in F1 mouse embryonic stem (ES) cells (Mus musculus129 x Mus castaneus). Monoallelic deletion, screening and expression analysis is facilitated by single nucleotide polymorphisms (SNP) between the two alleles which occur on average every 125 bp in these cells.

Experimental validation of enhancer activity is best approached by loss-of-function analysis. Presented here is an efficient protocol that uses CRISPR/Cas9 mediated deletion to study allele-specific regulation of gene transcription in F1 ES cells which contain a hybrid genome (Mus musculus129 x Mus castaneus).

Enhancers control cell identity by regulating tissue-specific gene expression in a position and orientation independent manner. These enhancers are often ...

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 ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...

Enforced Activation of Enhancer RNAs In Situ through the dCas9 Synergistic Activation Mediator System

Enhancers are pivotal genomic elements scattered through the mammalian genome and dictate tissue-specific gene expression programs. Increasing evidence has shown that enhancers not only provide DNA binding motifs for transcription factors (TFs) but also generate non-coding RNAs that are referred to as eRNAs. Studies have demonstrated that eRNA transcripts can play significant roles in gene regulation in both physiology and disease. Commonly used methods to investigate the function of eRNAs are constrained to &#8220;loss-of-function&#8221; approaches by knockdown of eRNAs, or by chemical inhibition of the enhancer transcription. There has not been a robust method to conduct &#8220;gain-of-function&#8221; studies of eRNAs to mimic specific disease conditions such as human cancer, where eRNAs are often overexpressed. Here, we introduce a method for precisely and robustly activating eRNAs for functional interrogation of their roles by applying the dCas9 mediated Synergistic Activation Mediators (SAM) system. We present the entire workflow of eRNA activation, from the selection of eRNAs, the design of gRNAs to the validation of eRNA activation by RT-qPCR. This method represents a unique approach to study the roles of a particular eRNA in gene regulation and disease development. In addition, this system can be employed for unbiased CRISPR screening to identify phenotype-driving eRNA targets in the context of a specific disease.

Enhancer RNAs (eRNAs) are non-coding RNAs produced from active enhancers. An optimal approach to study eRNA functions is to manipulate their levels in the native chromatin regions. Here we introduce a robust system for eRNA studies by using CRISPR-dCas9-fused transcriptional activators to induce the expression of eRNAs of interest.

Enhancers are pivotal genomic elements scattered through the mammalian genome and dictate tissue-specific gene expression programs. Increasing evidence ...

Lentiviral CRISPR/Cas9-Mediated Genome Editing for the Study of Hematopoietic Cells in Disease Models

Manipulating genes in hematopoietic stem cells using conventional transgenesis approaches can be time-consuming, expensive, and challenging. Benefiting from advances in genome editing technology and lentivirus-mediated transgene delivery systems, an efficient and economical method is described here that establishes mice in which genes are manipulated specifically in hematopoietic stem cells. Lentiviruses are used to transduce Cas9-expressing lineage-negative bone marrow cells with a guide RNA (gRNA) targeting specific genes and a red fluorescence reporter gene (RFP), then these cells are transplanted into lethally-irradiated C57BL/6 mice. Mice transplanted with lentivirus expressing non-targeting gRNA are used as controls. Engraftment of transduced hematopoietic stem cells are evaluated by flow cytometric analysis of RFP-positive leukocytes of peripheral blood. Using this method, ~90% transduction of myeloid cells and ~70% of lymphoid cells at 4 weeks after transplantation can be achieved. Genomic DNA is isolated from RFP-positive blood cells, and portions of the targeted site DNA are amplified by PCR to validate the genome editing. This protocol provides a high-throughput evaluation of hematopoiesis-regulatory genes and can be extended to a variety of mouse disease models with hematopoietic cell involvement.

Described are protocols for the highly efficient genome editing of murine hematopoietic stem and progenitor cells (HSPC) by the CRISPR/Cas9 system to rapidly develop mouse model systems with hematopoietic system-specific gene modifications.

Manipulating genes in hematopoietic stem cells using conventional transgenesis approaches can be time-consuming, expensive, and challenging. Benefiting ...

Immunology and Infection

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Gene inactivation is instrumental to study gene function and represents a promising strategy for the treatment of a broad range of diseases. Among traditional technologies, RNA interference suffers from partial target abrogation and the requirement for life-long treatments. In contrast, artificial nucleases can impose stable gene inactivation through induction of a DNA double strand break (DSB), but recent studies are questioning the safety of this approach. Targeted epigenetic editing via engineered transcriptional repressors (ETRs) may represent a solution, as a single administration of specific ETR combinations can lead to durable silencing without inducing DNA breaks.
ETRs are proteins containing a programmable DNA-binding domain (DBD) and effectors from naturally occurring transcriptional repressors. Specifically, a combination of three ETRs equipped with the KRAB domain of human ZNF10, the catalytic domain of human DNMT3A and human DNMT3L, was shown to induce heritable repressive epigenetic states on the ETR-target gene. The hit-and-run nature of this platform, the lack of impact on the DNA sequence of the target, and the possibility to revert to the repressive state by DNA demethylation on demand, make epigenetic silencing a game-changing tool. A critical step is the identification of the proper ETRs&#39; position on the target gene to maximize on-target and minimize off-target silencing. Performing this step in the final ex vivo or in vivo preclinical setting can be cumbersome.
Taking the CRISPR/catalytically dead Cas9 system as a paradigmatic DBD for ETRs, this paper describes a protocol consisting of the in vitro screen of guide RNAs (gRNAs) coupled to the triple-ETR combination for efficient on-target silencing, followed by evaluation of the genome-wide specificity profile of top hits. This allows for reduction of the initial repertoire of candidate gRNAs to a short list of promising ones, whose complexity is suitable for their final&#160;evaluation in the therapeutically relevant setting of interest.

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Here, we present a protocol for the in vitro selection of engineered transcriptional repressors (ETRs) with high, long-term, stable, on-target silencing efficiency and low genome-wide, off-target activity. This workflow allows for reducing an initial, complex repertoire of candidate ETRs to a short list, suitable for further evaluation in therapeutically relevant settings.

Gene inactivation is instrumental to study gene function and represents a promising strategy for the treatment of a broad range of diseases. Among ...

Bioengineering

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 ...

Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

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