C.P. is supported by RO1 DK60729 and P30 DK 41301-26. D.P. is supported by a Crohn&#39;s &#38; Colitis Foundation (CCFA) career development award, CURE: Digestive Diseases Research Center (DDRC) DK41301, and UCLA Clinical and Translational Science Institute (CTSI) UL1TR0001881. The UCLA virology core was funded by the Center of AIDS Research (CFAR) grant 5P30 AI028697. The UCLA Integrated Molecular Technologies core was supported by CURE/P30 grant DK041301. This work was also supported by the UCLA AIDS Institute.

NOTE: It is important to note that this protocol uses replication-deficient lentiviruses. Perform viral handling only after appropriate lab safety training. Bleach all items and surfaces that come in contact with live viruses for a minimum of 10 min after handling. Use a disposable lab coat and face/eye protection, as well as double gloves, at all times. Perform virus work in biosafety level 2+ labs with viral certification. Dedicate tissue culture hoods and incubators to viral work.
1. Vector Design and Generation
NOTE: The best way to identify gRNA sequences is to use online design tools (e.g., http://crispr-era.stanford.edu) (Supplemental Figure 1: gRNA design). For the accompanying representative experiment, a company designed and created the gRNA vectors used in the representative experiment. The dCas9 plasmid was also purchased online.
<ol>
	<li>The gRNA sequence is located upstream of the nucleotide sequence &#8220;NGG&#8221;, where &#8220;N&#8221; is any nucleotide, which also is within 100 base-pairs of the transcriptional start site. Search genetic databases such as BLAT (https://genome.ucsc.edu/cgi-bin/hgBlat?command=start) for the gRNA sequence, to make sure there are no other sites in the genome with similar sequences (Supplemental Figure 2: BLAT). This will ensure that the gRNA sequence is unique to the gene of interest (GOI).</li>
	<li>Store the gRNA and dCas9 plasmids, which come as bacterial stubs, at 4 &#176;C. Streak out Escherichia coli on a Luria broth (LB) agar plate with ampicillin (Supplemental Table 1) and grow the bacteria overnight at 37 &#176;C.</li>
	<li>Pick a colony and grow the bacteria in 5 mL of LB with ampicillin (Supplemental Table 1) overnight, vigorously shaking the plate in a 37 &#176;C incubator. The next day, add 2 mL of bacteria to 200 mL of LB with ampicillin grow in a 2 L flask, vigorously shaking the flask overnight in a 37 &#176;C incubator.</li>
	<li>Spin down the bacteria at 3,724 x g for 20 min. Remove the media and place the tube containing the bacteria on ice.</li>
	<li>Purify bacterial DNA using a plasmid purification kit per the manufacturer&#8217;s protocol. 
	NOTE: The plasmid purification kits contain proprietary resuspension buffer, lysis buffer, wash buffer, equilibration buffer, and elution buffer.
	<ol>
		<li>Add 10 mL of resuspension buffer from the kit to the bacteria and pipet the bacteria into the solution. Then, add 10 mL of lysis buffer from the kit to the resuspended bacteria and mix them by inverting the tube. Wait 5 min to lyse the bacteria; then, add 10 mL of ice-cold neutralization buffer from the kit to the lysed bacteria and mix them by inverting the tube.</li>
		<li>Equilibrate the DNA column from the kit with 10 mL of equilibration buffer for 5 min. Pour the samples into the DNA column filter and let the solution pass through the filter.</li>
		<li>Wash the samples 2x with 30 mL of wash buffer and, then, elute the DNA with 30 mL of elution buffer in a 15 mL conical tube. Slowly layer 10.5 mL of isopropanol at room temperature onto the eluted DNA, invert the tube, and immediately spin it at 16,200 x g for 30 min at 4 &#176;C.</li>
		<li>Remove the supernatant and add 1 mL of 70% ethanol to the DNA pellet. Resuspend the pellet by flicking the tube. Spin it at 16,200 x g for 10 min at 4 &#176;C and use a 200 &#956;L pipet tip to remove most of the ethanol. Air-dry the pellet for 5 min and resuspend it in 200 &#956;L of water, and store the DNA at -20 &#176;C. The expected concentration will be &#62;1 &#956;g/&#956;L.</li>
	</ol>
	</li>
</ol>
2. Virus Generation and Particle Count
<ol>
	<li>When ready to create the dCAS9-containing lentivirus, coat 100 mm tissue culture dishes with 5 mL of 0.01% poly-L-lysine (Supplemental Table 1). Remove the excess liquid thoroughly from the plates and let them air-dry for a few minutes in a biosafety cabinet.</li>
	<li>Plate 5 x 106 HEK 293T cells per dish in 10 mL of complete Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) (Supplemental Table 1) and incubate them overnight at 37 &#176;C with 5% CO2. The next day, remove the medium from the HEK 293T dishes and add 10 mL of complete DMEM.</li>
	<li>Mix 6.5 &#956;g of plasmid pMDLg/pRRE containing the gag/pol (components of the virus), 3.5 &#956;g of the plasmid pMDG2.G containing the VSV-G (a component of the virus), 2.5 &#181;g of the plasmid pRSV-Rev containing Rev (a component of the virus), 10 &#956;g of an long terminal repeat (LTR)-containing gRNA vector or an LTR-containing dCas9 vector and add water to a total volume of 450 &#956;L. Filter the mixture through a 0.2 &#181;m filter tip attached to a syringe.</li>
	<li>Add 50 &#181;L of 2.5 M CaCl2 (Supplemental Table 1) to each transfection sample of DNA and mix gently. Filter the calcium/DNA mixture through a 0.2 &#181;m filter tip attached to a syringe. Pipet 500 &#181;L of 2x HBS (Supplemental Table 1) into a 5 mL polystyrene tube. Add the 500 &#181;L DNA/CaCl2 mix dropwise and gently vortex. Incubate at room temperature for 3 min.</li>
	<li>Slowly add 1 mL of the DNA/CaCl2/HBS suspension to each dish. Immediately swirl the dishes to distribute the contents evenly. Incubate each dish overnight in a 5% CO2 incubator at 37 &#176;C.</li>
	<li>On day 3, slowly remove and discard the media from the dishes. Carefully wash the cells 1x with phosphate-buffered saline (PBS). Add 6 mL of complete DMEM supplemented with 20 mM HEPES and 10 mM sodium butyrate. Incubate the cells for 5&#8211;6 h in a 5% CO2 incubator at 37 &#176;C.</li>
	<li>Wash the cells 1x with PBS and add 5 mL of complete DMEM with 20 mM HEPES to the HEK 293T cells. Incubate for 12 h overnight in a 5% CO2 incubator at 37 &#176;C.</li>
	<li>On day 4, collect the HEK 293T cell supernatants and filter the supernatant. Freeze 1 mL aliquots at -80 &#176;C.</li>
	<li>When ready to use the virus, thaw an aliquot of the viral particle containing conditioned media (stored as described in step 2.8) on ice. Bring all reagents to room temperature.</li>
	<li>Use a p24 Enzyme-Linked Immunosorbent Assay (ELISA) kit to quantify the number of viral particles per milliliter.
	<ol>
		<li>Dilute the wash concentrate from the kit by adding 19 parts of distilled deionized water. Dilute the positive control from the kit to 200 ng/mL, using RMPI 1640 as the diluent, and make the dilutions for standard curve in 1.5 mL tubes according to the p24 ELISA dilution table (Supplemental Table 2).</li>
		<li>Add 20 &#956;L of 5% Triton X-100 to all wells except the substrate blank. Add 200 &#956;L of RPMI 1640 to the negative control wells. Add 200 &#956;L of each standard (in duplicate) to the designated wells.</li>
		<li>Using a spectrophotometer, measure the concentration of the virus within the samples, using a standard curve. Start the sample dilution at 1:1,000 and modify the volume as necessary in order to be within the range of the standard curve. Dilute the samples with Triton X-100 to a final concentration of 0.5% and add 200 &#956;L of each sample in RPMI 1640 to designated wells. Seal the plate and incubate it for 2 h at 37 &#176;C.</li>
		<li>Wash the plate 6x with 300 &#956;L of 1x wash buffer per well. Remove any excess fluid by inverting the plate and tapping it on a paper towel. Add 100 &#956;L of detector antibody from the kit to all wells except the substrate blank. Seal the plate and incubate it for 1 h at 37 &#176;C. Wash the plate and, then, remove any excess fluid by inverting the plate and tapping it on a paper towel.</li>
		<li>In order to measure the detector antibody, prepare streptavidin-horseradish peroxidase (SA-HRP) within 15 min of use. Dilute the SA-HRP at 1:100 with SA-HRP diluent. Mix the diluted SA-HRP thoroughly and add 100 &#956;L to all wells except the blank. Seal and incubate the plate for 30 min at room temperature.</li>
		<li>Wash the plate with 1x wash buffer and tap away any excess liquid as in step 2.10.4. Use ortho-phenylenediamine (OPD) substrate solution, which provides the peroxidase the necessary substrates to produce chemiluminescence, within 15 min of preparation. Use one OPD tablet to 11 mL of substrate diluent for each plate.</li>
		<li>Vortex the OPD solution vigorously to dissolve it completely and protect it from light. Add 100 &#956;L of OPD substrate solution to all wells, including the blank. Using a spectrophotometer, read absorbance at 450 nm immediately, repeat this 10x at 1 s intervals, and take the average measurement.</li>
	</ol>
	</li>
</ol>
3. dCas9-VP64 Transduction
<ol>
	<li>Culture Jurkat T cells in complete DMEM in a T75 flask with a density of 2 x 106 cells in 15 mL of media. Culture the cells in a 37 &#176;C incubator with 5% CO2.</li>
	<li>Spin down the cells for 5 min at 233 x g and, then, resuspend them in 10 mL of reduced serum media. Count the cells with a hemocytometer or an automated cell counter.</li>
	<li>Resuspend and plate 1 x 106 Jurkat cells in 5 mL of reduced serum media with polybrene (Supplemental Table 1) in a T75 flask. Add HEK 293T-conditioned media containing 1 x 106 dCas9-containing viral particles. The number of viral particles can be calculated roughly from the p24 ELISA because 1 &#956;g/mL p24 equals 1 x 107 viral particles. Put the flasks in 37 &#176;C incubator with 5% CO2.</li>
</ol>
4. Selection and Clone Creation
<ol>
	<li>Three days post-infection, spin down the cells at 233 x g for 5 min and resuspend them in 10 mL of complete DMEM with puromycin (Supplemental Table 1). Plate the cells in T75 flasks and culture them at 37 &#176;C with 5% CO2. Every third day, for a period of 9 days, spin down the cells at 233 x g for 5 min and replace the media with complete DMEM with puromycin.</li>
	<li>Count the cells using a hemocytometer or an automated cell counter. On a 96-well plate treated with tissue culture, plate 10,000 cells in 100 &#956;L of complete DMEM with puromycin in the first well and serially dilute 1:1 the contents of the following wells with complete DMEM with puromycin. Perform 24 dilutions and repeat this 4x per plate in order to ensure that there is an adequate number of cells per well. Incubate the cells at 37 &#176;C with 5% CO2.</li>
	<li>Expand clonal cells into two 24-well plates, then into a 6-well plate, and eventually into a T75 flask. In order to focus on three of the clones, plate 2 x 106 cells per well of a 6-well plate for three of the clones. Plate the other three wells with nontransduced Jurkat cells as controls. Put the cells in a cell culture incubator overnight.</li>
	<li>For RNA extraction, use a commercially available RNA isolation kit. Spin down the cells at 233 x g for 5 min at room temperature and remove the media. Use a 1 mL pipet tip to resuspend the cells in 350 &#956;L of lysis buffer. Add 350 &#956;L of 70% ethanol to the lysed cells, mix thoroughly with a 1 mL pipet, and transfer the sample to the RNA column.</li>
	<li>Spin the lysates at 10,000 x g for 1 min, remove the flow-through, and add 700 &#956;L of low-stringency wash buffer from the kit to the column. Spin the lysates at 10,000 x g for 1 min, remove the flow-through, and add 80 &#181;L of DNase I solution from the kit to the column; let the samples incubate for 15 min at room temperature.</li>
	<li>Spin the lysates at 10,000 x g for 1 min, remove the flow-through, and add 700 &#181;L of high-stringency wash buffer from the kit to the column. Spin the lysates at 10,000 x g for 1 min, remove the flow-through, and add 700 &#181;L of low-stringency wash buffer to the column.</li>
	<li>Remove the column from the collection tube and transfer it to a new collection tube. Spin the lysates at 10,000 x g for 1 min and, then, transfer the RNA column to a 1.5 mL tube. Add 50 &#181;L of water to the column and spin at 10,000 x g for 1 min.</li>
	<li>Use a spectrophotometer to quantify the concentration of RNA. Expect the concentration to be between 100&#8211;500 ng/&#181;L, with a 260/280 absorbance between 1.9&#8211;2.1. Store the RNA at -80 &#176;C.</li>
	<li>In 250 &#181;L tubes, add 1 &#181;g of RNA, 4 &#181;L of complementary DNA (cDNA) synthesis buffer, 1 &#181;L of reverse transcriptase, and water to a final volume of 20 &#181;L. Include no reverse transcriptase controls. In a thermal cycler, synthesize cDNA at 42 &#176;C for 30 min and at 95 &#176;C for 5 min to inactivate the reverse transcriptase. Dilute the cDNA with 60 &#181;L of water after the synthesis.</li>
	<li>Prepare polymerase chain reactions (PCRs) containing 5 &#181;L of SYBR green, 4 &#181;L of cDNA, 0.5 &#181;L of forward primer (10 &#181;M), and 0.5 &#181;L of reverse primer (10 &#181;M). Run the PCR as follows: step 1 = 95 &#176;C for 3 min, step 2 = 95 &#176;C for 10 s, step 3 = 60 &#176;C for 10 s, and then, repeat steps 2 and 3 for 30x. 
	NOTE: The sequence for the dCas9 forward primer is 5&#39;-TCGCCACAGCATAAAGAAGA and the sequence for the dCas9 reverse primer is 5&#39;-CTTTTCATGGTACGCCACCT. The forward primer for Hypoxanthine Phosphoribosyltransferase 1 (HPRT1) is 5&#39;-GACCAGTCAACAGGGGACAT and the reverse primer is 5&#39;-GCTTGCGACCTTGACCATCT.</li>
</ol>
5. gRNA Transduction, Selection, Clone Creation, and Screening
<ol>
	<li>Retransduce the validated Jurkat-dCas9 cells with gRNA-containing viruses as outlined in section 3. Select cells in DMEM with hygromycin (Supplemental Table 1) for 10 days. Spin down the cells and change the media every 3 days.</li>
	<li>Perform a serial dilution of cells (as described in step 4.2) and expand individual colonies (as described in step 4.3). Purify RNA from the colonies exactly as described in step 4.5 and perform cDNA synthesis as described in step 4.9. Perform real-time PCR against the GOI and HPRT1 or an appropriate housekeeping gene, similarly to the PCR described in step 4.10, except in this case, image the wells each cycle after step 3.</li>
	<li>Use the comparative cycle threshold (Ct) method to determine fold change in the GOI8. Briefly, use the following equation to calculate relative transcript levels (RTLs): 2-(average Ct of GOI &#8211; average Ct of housekeeping gene). Then, calculate the average RTL of the controls and divide all RTL values by the average control RTL to generate a fold change compared to the control samples.</li>
</ol>

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The authors have nothing to disclose.

This manuscript presents a protocol for using activating CRISPR to overexpress lncRNAs in vitro. This is an especially important technique when studying long noncoding RNAs as the transcriptomic product is the functional unit. After overexpression, the researcher can, then, use these cells to increase the signal-to-noise ratio when studying binding partners and even measure the cellular consequences of increased levels of lncRNAs. Additionally, as lncRNAs frequently act on cis-genes, this endogenous overexpression techni...

While most biomedical research has focused on protein-coding transcripts, the majority of transcribed genes actually consists of noncoding RNAs (Ensembl release 93).Current research is beginning to explore this field, with the number of publications on lncRNAs in disease processes rising exponentially between 2007 and 20171. These publications demonstrate that many lncRNAs are associated with disease. However, the molecular mechanisms of these lncRNAs are difficult to study due to their diverse functions as compared to mRNAs. Compounding the problem of understanding the role of lncRNAs in disease, lncRNAs are often expressed at lower levels than coding RNAs2. Additionally, lncRNAs are poorly conserved, which limits the functional studies in human-cell-line-based techniques3. One method to study the mechanism of these novel genes is to endogenously overexpress them in cultured cells. Overexpression studies can provide key information as to the function of specific genes and enable researchers to dissect key molecular pathways.
A new method to activate the transcription of lncRNA genes is based on CRISPR technologies developed initially by the Gersbach laboratory4. This protocol has been adapted for use in lncRNA biology and for the expression of these genes in other model systems. In the CRISPR overexpressing technique, a protein called CRISPR-associated protein 9 (Cas9) can be directed toward a DNA sequence via an antisense gRNA that is recognized by Cas9. Normally, Cas9 will induce DNA cleavage; however, mutations in Cas9, previously developed for the overexpression technique, deactivate this step5. When a transcriptional activator is fused to a &#34;dead&#34; Cas9 (dCas9) and transduced into cell lines, the endogenous overexpression of lncRNAs can be achieved4,6. The addition of additional modifications to the gRNA, enabling additional transcriptional factors to bind to the gRNA, increased the efficacy of the dCas9 gene activation system 10-fold7. Importantly, it was demonstrated that transcriptional activation requires close proximity (&#60;200 bp) to the transcriptional start site genes, enabling a specific upregulation in gene-rich areas7. Unlike CRISPR knockout technologies, dCas9 and gRNA cassettes need to be integrated into the genome to allow for cells to retain an overexpression over multiple generations. One method to achieve this is to use lentiviruses to integrate the dCas9- and gRNA-containing cassettes. After integration, the lncRNA gene expression can be determined.
In this article, a protocol for lncRNA overexpression in a Jurkat T-cell model will be demonstrated. A step-by-step procedure is shown and can also be adapted to adherent cells.

<table><tbody><tr><td>Ampicillin</td><td>Fisher Scientific</td><td>BP1760-5</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>C1016</td><td></td></tr><tr><td>cDNA synthesis kit (iScript)</td><td>BioRad</td><td>1708891</td><td></td></tr><tr><td>dCas9 forward primer</td><td>Integrated DNA Technologies</td><td>n/a</td><td>5&#039;-TCGCCACAGCATAAAGAAGA&amp;nbsp;</td></tr><tr><td>dCas9 reverse primer</td><td>Integrated DNA Technologies</td><td>n/a</td><td>5&#039;-CTTTTCATGGTACGCCACCT</td></tr><tr><td>dCas9 vector</td><td>Addgene</td><td>50918</td><td></td></tr><tr><td>DMEM</td><td>Corning</td><td>10013CM</td><td></td></tr><tr><td>Ethanol</td><td>Acros Organics</td><td>61509-0010</td><td></td></tr><tr><td>FBS</td><td>Sigma Aldrich</td><td>F2442</td><td></td></tr><tr><td>gRNA plasmid</td><td>VectorBuilder</td><td>VB180119-1195qxv</td><td></td></tr><tr><td>HEK293T cells</td><td>ATCC</td><td>&amp;nbsp;CRL-1573</td><td></td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H3375</td><td></td></tr><tr><td>HPRT1 forward primer</td><td>Integrated DNA Technologies</td><td>n/a</td><td>GACCAGTCAACAGGGGACAT&amp;nbsp;</td></tr><tr><td>HPRT1 reverse primer</td><td>Integrated DNA Technologies</td><td>n/a</td><td>GCTTGCGACCTTGACCATCT</td></tr><tr><td>Hygromycin B</td><td>Corning</td><td>MT3024CR</td><td></td></tr><tr><td>Isopropanol</td><td>Fisher Scientific</td><td>BP2618-500</td><td></td></tr><tr><td>Jurkat cells</td><td>ATCC</td><td>&amp;nbsp;TIB-152</td><td>clone E6-1</td></tr><tr><td>L-glutamine</td><td>Corning</td><td>25005Cl</td><td></td></tr><tr><td>LB agar</td><td>Fisher Scientific</td><td>BP1425-500</td><td></td></tr><tr><td>LB broth</td><td>Fisher Scientific</td><td>BP1426-2</td><td></td></tr><tr><td>Maxi-prep kit (Plasmid Purification Kit)</td><td>Qiagen</td><td>12362</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>NIST2186II</td><td></td></tr><tr><td>Optimem I reduced serum media&amp;nbsp;</td><td>Gibco</td><td>31985070</td><td></td></tr><tr><td>p24 elisa</td><td>Perkin Elmer</td><td>NEK050B</td><td></td></tr><tr><td>PBS</td><td>Corning</td><td>21-040-CMR</td><td></td></tr><tr><td>Penicillin and Streptomycin</td><td>Corning</td><td>30-002-Cl</td><td></td></tr><tr><td>pMDG2.G</td><td>Addgene &amp;nbsp;</td><td>#12259</td><td></td></tr><tr><td>pMDLg/pRRE</td><td>Addgene&amp;nbsp;</td><td>#60488</td><td></td></tr><tr><td>Poly-L-Lysine</td><td>Sigma Aldrich</td><td>P-4832</td><td></td></tr><tr><td>Polybrene</td><td>EMD Millipore</td><td>TR-1003</td><td></td></tr><tr><td>pRSV-REV</td><td>Addgene</td><td>#12253</td><td></td></tr><tr><td>Puromycin dihydrochloride</td><td>Sigma Aldrich</td><td>P8833</td><td></td></tr><tr><td>RNA purification kit (Aurum RNA mini)</td><td>BioRad</td><td>7326820</td><td></td></tr><tr><td>Sodium Butyrate</td><td>Sigma Aldrich</td><td>B5887</td><td></td></tr><tr><td>SYBR green (iTaq universal)</td><td>BioRad</td><td>1725122</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>X100</td><td></td></tr></tbody></table>

overexpressing long noncoding rnas using gene-activating crispr

Long noncoding RNA (lncRNA) biology is a new and exciting field of research, with the number of publications from this field growing exponentially since 2007. These studies have confirmed that lncRNAs are altered in almost all diseases. However, studying the functional roles for lncRNAs in the context of disease remains difficult due to the lack of protein products, tissue-specific expression, low expression levels, complexities in splice forms, and lack of conservation among species. Given the species-specific expression, lncRNA studies are often restricted to human research contexts when studying disease processes. Since lncRNAs function at the molecular level, one way to dissect lncRNA biology is to either remove the lncRNA or overexpress the lncRNA and measure cellular effects. In this article, a written and visualized protocol to overexpress lncRNAs in vitro is presented. As a representative experiment, an lncRNA associated with inflammatory bowel disease, Interferon Gamma Antisense 1 (IFNG-AS1), is shown to be overexpressed in a Jurkat T-cell model. To accomplish this, the activating clustered regularly interspaced short palindromic repeats (CRISPR) technique is used to enable overexpression at the endogenous genomic loci. The activating CRISPR technique targets a set of transcription factors to the transcriptional start site of a gene, enabling a robust overexpression of multiple lncRNA splice forms. This procedure will be broken down into three steps, namely (i) guide RNA (gRNA) design and vector construction, (ii) virus generation and transduction, and (iii) colony screening for overexpression. For this representative experiment, a greater than 20-fold enhancement in IFNG-AS1 in Jurkat T cells was observed.

A dual vector system to overexpress the lncRNA IFNG-AS1 
The example experiment in this manuscript is the overexpression of a Jurkat T-cell model system expressing the lncRNA IFNG-AS19. IFNG-AS1 is an lncRNA associated with inflammatory bowel disease, that has been seen to regulate Interferon Gamma10. The IFNG-AS1 gene contains three splice variants that all use the same transcriptional start site (...

Watch this Scientific Journal Video about Overexpressing Long Noncoding RNAs Using Gene-activating CRISPR at JoVE.com

Overexpressing Long Noncoding RNAs Using Gene-activating CRISPR

Traditional cDNA-based overexpression techniques have a limited applicability for the overexpression of long noncoding RNAs due to their multiple splice forms with potential functionality. This review reports a protocol using CRISPR technology to overexpress multiple splice variants of a long noncoding RNA.

Long noncoding RNA (lncRNA) biology is a new and exciting field of research, with the number of publications from this field growing exponentially since ...

overexpressing-long-noncoding-rnas-using-gene-activating-crispr

Research

JoVE Journal

Genetics

8.8K Views.  University of California, Los Angeles. Niet-codering RNAs hebben vaak meerdere isovormen. In in vitro overexpressiestudies is het vaak moeilijk om al deze uitgedrukte isovormen te bestuderen. Deze techniek stelt gebruikers in staat om expressie rechtstreeks uit het genoom af te leiden en de cellen in staat te stellen de endogene isovormen uit te drukken voor een bepaald niet-coderend RNA.Deze krachtige techniek stelt de gebruiker in staat om specifiek de expressie van elk gen in het genoom met trouw en nauwkeurigheid te stu...

Video: Overexpressing Long Noncoding RNAs Using Gene-activating CRISPR

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 ...

Developmental Biology

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.

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 ...

Lentiviral Vector Platform for the Efficient Delivery of Epigenome-editing Tools into Human Induced Pluripotent Stem Cell-derived Disease Models

The use of hiPSC-derived cells represents a valuable approach to study human neurodegenerative diseases. Here, we describe an optimized protocol for the differentiation of hiPSCs derived from a patient with the triplication of the alpha-synuclein gene (SNCA) locus into Parkinson&#8217;s disease (PD)-relevant dopaminergic neuronal populations. Accumulating evidence has shown that high levels of SNCA are causative for the development of PD. Recognizing the unmet need to establish novel therapeutic approaches for PD, especially those targeting the regulation of SNCA expression, we recently developed a CRISPR/dCas9-DNA-methylation-based system to epigenetically modulate SNCA transcription by enriching methylation levels at the SNCA intron 1 regulatory region. To deliver the system, consisting of a dead (deactivated) version of Cas9 (dCas9) fused with the catalytic domain of the DNA methyltransferase enzyme 3A (DNMT3A), a lentiviral vector is used. This system is applied to cells with the triplication of the SNCA locus and reduces the SNCA-mRNA and protein levels by about 30% through the targeted DNA methylation of SNCA intron 1. The fine-tuned downregulation of the SNCA levels rescues disease-related cellular phenotypes. In the current protocol, we aim to describe a step-by-step procedure for differentiating hiPSCs into neural progenitor cells (NPCs) and the establishment and validation of pyrosequencing assays for the evaluation of the methylation profile in the SNCA intron 1. To outline in more detail the lentivirus-CRISPR/dCas9 system used in these experiments, this protocol describes how to produce, purify, and concentrate lentiviral vectors and to highlight their suitability for epigenome- and genome-editing applications using hiPSCs and NPCs. The protocol is easily adaptable and can be used to produce high titer lentiviruses for in vitro and in vivo applications.

Targeted DNA epigenome editing represents a powerful therapeutic approach. This protocol describes the production, purification, and concentration of all-in-one lentiviral vectors harboring the CRISPR-dCas9-DNMT3A transgene for epigenome-editing applications in human induced pluripotent stem cell (hiPSC)-derived neurons.

The use of hiPSC-derived cells represents a valuable approach to study human neurodegenerative diseases. Here, we describe an optimized protocol for the ...

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 ...

A New Toolkit for Evaluating Gene Functions using Conditional Cas9 Stabilization

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

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

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

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 ...

Designing, Packaging, and Delivery of High Titer CRISPR Retro and Lentiviruses via Stereotaxic Injection

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique has been widely used to encode fluorescent proteins, opto- or chemo-genetic controllers of cell activity, or heterologous expression of human genes in model organisms. These viruses have also successfully been used to deliver recombinases to relevant target sites in transgenic animals, or even deliver small hairpin or micro RNAs in order to manipulate gene expression. While these techniques have been fruitful, they rely on transgenic animals (recombinases) or frequently lack high efficacy and specificity (shRNA/miRNA). In contrast, the CRISPR/Cas system uses an exogenous Cas nuclease which targets specific sites in an organism's genome via an exogenous guide RNA in order to induce double stranded breaks in DNA. These breaks are then repaired by non-homologous end joining (NHEJ), producing insertion and deletion (indel) mutations that can result in deleterious missense or nonsense mutations. This manuscript provides detailed methods for the design, production, injection, and validation of single lenti/retro virus particles that can stably transduce neurons to express a fluorescent reporter, Cas9, and sgRNAs to knockout genes in a model organism.

The CRISPR/Cas9 system offers the potential to make targeted genome editing accessible and affordable to the scientific community. This protocol is intended to demonstrate how to create viruses that will knockout a gene of interest using the CRISPR/Cas9 system, and then inject them stereotaxically into the adult mouse brain.

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique ...

Neuroscience

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

Gene Digital Circuits Based on CRISPR-Cas Systems and Anti-CRISPR Proteins

Synthetic gene Boolean gates and digital circuits have a broad range of applications, from medical diagnostics to environmental care. The discovery of the CRISPR-Cas systems and their natural inhibitors-the anti-CRISPR proteins (Acrs)-provides a new tool to design and implement in vivo gene digital circuits. Here, we describe a protocol that follows the idea of the "Design-Build-Test-Learn" biological engineering cycle and makes use of dCas9/dCas12a together with their corresponding Acrs to establish small transcriptional networks, some of which behave like Boolean gates, in Saccharomyces cerevisiae. These results point out the properties of dCas9/dCas12a as transcription factors. In particular, to achieve maximal activation of gene expression, dSpCas9 needs to interact with an engineered scaffold RNA that collects multiple copies of the VP64 activation domain (AD). In contrast, dCas12a shall be fused, at the C terminus, with the strong VP64-p65-Rta (VPR) AD. Furthermore, the activity of both Cas proteins is not enhanced by increasing the amount of sgRNA/crRNA in the cell. This article also explains how to build Boolean gates based on the CRISPR-dCas-Acr interaction. The AcrIIA4 fused hormone-binding domain of the human estrogen receptor is the core of a NOT gate responsive to &#946;-estradiol, whereas AcrVAs synthesized by the inducible GAL1 promoter permits to mimic both YES and NOT gates with galactose as an input. In the latter circuits, AcrVA5, together with dLbCas12a, showed the best logic behavior.

CRISPR-Cas systems and anti-CRISPR proteins were integrated into the scheme of Boolean gates in Saccharomyces cerevisiae. The new small logic circuits showed good performance and deepened the understanding of both dCas9/dCas12a-based transcription factors and the properties of anti-CRISPR proteins.

Synthetic gene Boolean gates and digital circuits have a broad range of applications, from medical diagnostics to environmental care. The discovery of the ...

Bioengineering

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.

Overexpressing Long Noncoding RNAs Using Gene-activating CRISPR

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