Overexpressing Long Noncoding RNAs Using Gene-activating CRISPR

Noncoding RNAs often have multiple isoforms. In in vitro overexpression studies, it is often difficult to study all these expressed isoforms. This technique allows users to derive expression directly from the genome and enable the cells to express the endogenous isoforms for a particular noncoding RNA.

This powerful technique allows the user to specifically direct the expression of any gene in the genome with fidelity and accuracy. By designing guide RNAs to specific genes, one is able to use the endogenous gene-splicing machinery to express the natural isoforms in a given cell. Demonstrating the procedure will be Robert Rankin, a post-doc from my laboratory.

To design the guide RNA sequence which is located within 100 base pairs of the transcriptional start site, use genetic databases such as BLAT to make sure that the guide RNA is unique to the gene of interest. Store the guide RNA and the deactivated Cas9 plasmids at four degrees Celsius. Streak out E.coli on a Luria broth agar plate with ampicillin and grow the bacteria overnight at 37 degrees Celsius.

The next day, pick a colony and grow the bacteria in five milliliters of LB with ampicillin overnight, vigorously shaking the tube in a 37-degree Celsius incubator. The next day, add two milliliters of bacteria to 200 milliliters of LB with ampicillin in a two-liter flask, vigorously shaking the flask overnight in a 37-degree Celsius incubator. Spin down the bacteria at 3, 724 times g for 20 minutes and proceed to DNA purification.

To create the dCas9-containing lentivirus, first coat 100-milliliter tissue culture dishes with five milliliters of 0.01%Poly-L-Lysine and plate five million HEK293T cells per dish in 10 milliliters of complete Dulbecco's Modified Eagles Medium, then prepare DNA transfection samples by mixing 10 micrograms of an LTR-containing guide RNA vector or an LTR-containing deactivated Cas9 vector with plasmids containing components of viruses. Add water to a total volume of 450 microliters and filter the mixture through a 0.2-micron filter tip attached to a syringe. Next, add 50 microliters of 2.5-molar calcium dichloride to each transfection sample of the DNA and mix gently.

Filter the calcium-DNA mixture through a 0.2-micron filter tip attached to a syringe. Pipette 500 microliters of 2X HBS into a five-milliliter polystyrene tube. Add 500 microliters of the calcium-DNA mixture dropwise and gently vortex.

Incubate at room temperature for three minutes. Add one milliliter of the DNA-calcium HBS suspension to each HEK293T-containing tissue culture dish and incubate them in a 5%carbon dioxide incubator at 37 degrees Celsius overnight. On day three, slowly remove and discard the media from the dishes.

Carefully wash the cells with PBS once. Add six milliliters of complete DMEM supplemented with 20-millimolar HEPES and 10-millimolar sodium butyrate. Then incubate the cells in a 5%carbon dioxide incubator at 37 degrees Celsius for five to six hours.

After the incubation, wash the cells with PBS once and add five milliliters of complete DMEM with 20-millimolar HEPES to the HEK293T cells. Incubate the cells in a 5%carbon dioxide incubator at 37 degrees Celsius for 12 hours. On day four, collect the HEK293T cell supernatants and filter the supernatant containing the virus.

To begin the viral particle counts, first dilute the wash concentrate from the P24 ELISA kit by adding 19 parts of distilled, deionized water. Then, dilute the positive control from this kit to 200 nanograms per milliliter using RPMI 1640 as the diluent and make dilutions for a standard curve in 1.5-milliliter tubes according to the P24 ELISA dilution table. To measure the concentration of the virus within the samples add the sample dilutions to the designated wells of a 96-well plate, starting from 1:1000 dilution and modify the volume as necessary in order to be within the range of the standard curve.

Then dilute the samples with Triton X-100 to a final concentration of 0.5%and add 200 microliters of each sample and RPMI 1640 to the designated wells. Seal the plate and incubate it at 37 degrees Celsius for two hours. Wash the plate with 300 microliters per well of the 1X wash buffer six times.

Remove any excess fluid by inverting the plate and tapping it on a paper towel. Then add 100 microliters of detector antibody from the kit to all wells except the substrate blank. Seal the plate and incubate it at 37 degrees Celsius for one hour.

Wash the plate and then remove any excess fluid by inverting the plate and tapping it on a paper towel. In order to measure the detector antibody, dilute the streptavidin horseradish peroxidase at 1:100 dilution with SA-HRP diluent. Mix the diluted SA-HRP thoroughly and add 100 microliters to all wells except the blank.

Seal and incubate the plate at room temperature for 30 minutes. Next, wash the plate with the 1X wash buffer and tap away any excess liquid. Drop one ortho-phenylenediamine tablet in 11 milliliters of the substrate diluent to make enough substrate solution for one plate.

Vortex the OPD solution vigorously to dissolve it completely and protect it from light. Add 100 microliters of OPD substrate solution to all wells including the blank. Using a spectrophotometer, read absorbance at 450 nanometers immediately.

Repeat 10 times at one-second intervals and take the average measurement. Resuspend and culture Jurkat T-cells in a T75 flask with a density of one million cells in five milliliters of the reduced serum media with polybrene. Then add HEK293T-conditioned media containing one million dCas9-containing viral particles.

Incubate the cells at 37 degrees Celsius with 5%carbon dioxide. Three days post-infection, spin down the cells at 233 time g for five minutes and resuspend them in 10 milliliters of complete DMEM supplemented with puromycin. Culture the cells in T75 flasks and incubate them at 37 degrees Celsius with 5%carbon dioxide.

Every third day for a period of nine days, spin down the cells at 233 times g for five minutes and replace the media with the complete DMEM supplemented with puromycin. Count the cells using a hemocytometer or an automated cell counter. Then, on a 96-well plate with tissue culture-treated tissue, add 10, 000 cells in 100 microliters of the complete DMEM supplemented with puromycin in the first well.

Make 1:1 serial dilutions of the contents of the following wells with the complete DMEM supplemented with puromycin. Incubate the cells at 37 degrees Celsius with 5%carbon dioxide. Expand clonal cells into two 24-well plates, then plate two million cells per well of a six-well plate for three of the clones.

Plate the other three wells with non-transduced Jurkat cells as controls. Finally, plate the cells into a T75 flask and put the cells in a cell culture incubator overnight. To begin quantitative PCR to measure gene expression, first synthesize complementary DNA by adding one microgram of RNA, four microliters of cDNA synthesis buffer, and one microliter of reverse transcriptase to 250 microliter tubes, then add water to a final volume of 20 microliters.

Then, use a thermal cycler at 42 degrees Celsius for 30 minutes and at 95 degrees Celsius for five minutes to inactivate the reverse transcriptase. Dilute the cDNA with 60 microliters of water after the synthesis. Run polymerase chain reactions for tubes containing 0.5 microliters of each forward and reverse primer at final concentration of 10 micromolar, five microliters of SYBR green, and four microliters of cDNA.

Finally select Jurkat-dCas9 cells in DMEM supplemented with Hygromycin B for 10 days. Spin down the cells and change the media every three days. Purify RNAse and proceed to RT-qPCR.

In this study, gRNA sequences that were within 10 to 100 base pairs away from the transcriptional start site were used for directing the Cas9-activating complex to the transcriptional locus of IFNG-AS1, a long noncoding RNA associated with inflammatory bowel disease. To enable the selection of double transduced cells, a two-plasmid system was used to transduce either dCas9 or gRNAse enhancers into cells. Here MS2 proteins enhance the overexpression of IFNG-AS1.

Cas9 expression was confirmed for both clones. Primers against HPRT1 were used to confirm the presence of RNA in the non-transduced cells. mRNA expression was confirmed by omitting the reverse transcriptase from the cDNA reaction.

Three splice variants of IFNG-AS1 could be detected with either transcript-specific primer sets or a primer set against all known IFNG-AS1 transcripts. All fluorescence curves were exponential with the housekeeping gene HPRT1 reaching the exponential phase within a half cycle between control and IFNG-AS1 gRNA-expressing cells. Primers against all known IFNG-AS1 transcripts were the most detectable between experiments, with measurements of 20-fold increases in IFNG-AS1 expression.

However, the third transcript of IFNG-AS1 showed a five-to 10-fold significant increase in IFNG-AS1 levels. To actively overexpress your desired gene, design of the guide RNA in step 2.1 is critical to the success of the protocol. Additionally, production of quality viral particles will ensure a robust expression of your protocol components.

Once the desired gene is overexpressed, functional studies can be performed to investigate the gene mechanisms. In our example, we chose to look at cytokine production following overexpression of our gene of interest. This technique was initially developed by the Griesbach Group In 2013 and has been broadly applied and elaborated on by other groups.

We were excited to apply this technique to long noncoded RNAs in order to explore their specific functions. Replication-defective lentiviruses should be handled in a laboratory that's been approved for viral work. Additionally, human cell lines and E.coli bacteria are considered biohazardous.

Lastly, recombinant DNA plasmids should be handled by trained staff.