Identification of Footprints of RNA:Protein Complexes via RNA Immunoprecipitation in Tandem Followed by Sequencing (RIPiT-Seq)

1. Establishment of Stable HEK293 Cell Lines Expressing Tetracycline-inducible FLAG-tagged Protein of Interest (POI)

1. Seed HEK293 cells with a stably integrated Flp recombination target (FRT) site at a density of 10 x 10⁴ cells/mL in growth medium (Dulbecco's modified Eagle's medium [DMEM] + 10% fetal bovine serum [FBS] + 1% penicillin-streptomycin [penn/strep]) in 6-well plates. Allow cells to grow overnight in a humidified incubator at 37 °C and 5% CO₂ (standard growth conditions for all subsequent steps).
2. The next day, cells should be ~70% confluent. According to transfection reagent protocol, transfect the FRT site-containing HEK293 cells with a 9:1 ratio of pcDNA5-TETO-FLAG-POI:pOG44.
   NOTE: The FLAG tag has the sequence motif DYKDDDDK (D = aspartic acid, Y = tyrosine, and K = lysine).
3. After 24 h, begin antibiotic selection. Remove media and add fresh growth medium supplemented with 100 μg/mL hygromycin. Within 72 h, untransfected cells should begin to die.
4. Every 48-72 h, change growth media and supplement with fresh hygromycin.
5. After ~2 weeks of selection, discrete colonies of stably transfected cells will begin to appear. Once colonies are visible to naked eye, add 1 mL of trypsin to the plate and incubate for 5 min at 37 °C. Resuspend cells in DMEM, transfer cells into a new 10 cm plate, and adjust volume to 10 mL fresh growth medium supplemented with 100 μg/mL hygromycin. Allow the plate to reach ~80% confluency to further expand cells to prepare permanent stocks.
6. Determine the amount of tetracycline (Tet) required to obtain the optimal level of FLAG-POI expression for the experiment. Grow cells in 12-well format and conduct a titration of Tet between 0-1,000 ng/mL range for 16-24 h. Perform western blots on the titration samples with antibody against the POI.
   NOTE: For proteins up to ~60 kDa, the FLAG-tagged and the endogenous copy of the protein can be resolved on the sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) to compare expression levels of FLAG-POI to its endogenous counterpart. For larger proteins, signal intensity in Tet-induced samples can be compared to uninduced sample. The tetracycline stock solution can be prepared at 1 mg/mL in 100% ethanol. Dilutions of the stock for cell culture work should be in sterile water or phosphate-buffered saline (PBS). Tetracycline stock should be prepared fresh once every month.

2. Culturing Cells for Tetracycline Induction and RIPiT Procedure

1. Seed stably integrated FLAG-POI HEK293 at a density of 3 x 10⁵ cells/mL in growth medium in 15 cm plates. Allow cells to grow at 37 °C and 5% CO₂.
   NOTE: In general, three to five 15 cm plates will yield ~2-20 pmol of RNA footprints depending on the abundance of the RNP of interest. If RNPs will be formaldehyde crosslinked and purified under stringent conditions, then twice as much input may be required.
2. Add tetracycline to pre-determined optimal concentration to the media to induce expression of the FLAG-POI (see step 1.6) 16-24 h before cells will be harvested.
   NOTE: It is not necessary to change growth medium. Cells should be ready to harvest about 72 h after seeding or when plates are ~80% confluent. It is important to avoid cells from becoming confluent.

3. Cell harvesting, Formaldehyde Treatment, and Cell Lysis

1. Wash the monolayer cells gently with 15 mL of chilled PBS for each 15 cm plate. Then scrape off cells in 30 mL of PBS. If cells were treated with drugs prior to harvesting, supplement PBS with the drug. Collect all cells into 50 mL conical tube.
   NOTE: For preserving weak interactions, cells can be crosslinked with formaldehyde: add formaldehyde to the cell suspension from step 3.1 to a final concentration of 0.1% and incubate on a rocker at room temperature for 10 min. Add 3 mL of the quenching buffer (Table 1) and rock for 5 additional minutes.
2. Pellet cells at 400 x g for 10 min at 4 °C and discard supernatant.
3. Lyse cells in 4 mL of ice-cold hypotonic lysis buffer (HLB; Table 1). Use a P1000 pipette to resuspend cells. Transfer to a 5 mL tube and incubate lysate on ice for 10 min.
4. Place the lysate in an ice bath and sonicate at 10% amplitude for 30 s with 1 s pulses with 2 s pauses. Then, adjust salt concentration to 150 mM by adding 108 μL of 5 M NaCl.
   NOTE: Formaldehyde crosslinked samples can be subject to more stringent lysis and purification by including 0.1% each of SDS and sodium deoxycholate in the lysis buffer.
5. Clear the lysate by centrifugation at 21,000 x g for 10 min at 4 °C. Collect 20 μL supernatant (cell extract) in a labeled tube for western blot analysis of protein levels in input (see Figure 3A).
   NOTE: While lysate is in centrifuge, FLAG agarose beads can be washed (see step 4.1). FLAG agarose beads should be pre-washed 3x in 4 mL of ice-cold isotonic wash buffer (IsoWB, Table 1).

4. FLAG Immunoprecipitation

1. Apply remaining supernatant from step 3.5 to 750 μL pre-washed FLAG agarose beads in a 5 mL tube (bed volume of washed FLAG agarose beads will be 375 μL). Incubate FLAG agarose beads and cell extract for 1-3 h at 4 °C with gentle mixing.
   NOTE: This volume of FLAG agarose beads should be sufficient for a wide-range of proteins, but may be optimized, if needed.
2. Pellet FLAG agarose beads by centrifugation at 400 x g for 1 min at 4 °C. Collect 20 μL of the supernatant in a labeled tube for Western blot analysis of protein levels in depleted cell extract (see Figure 3A). Discard the remaining supernatant.
3. To wash FLAG agarose beads, add 4 mL of IsoWB and resuspend. Pellet beads at 400 x g for 1 min at 4 °C. Carefully remove supernatant. Repeat 4x.
   NOTE: For stringent washes of the formaldehyde crosslinked IPs, 0.1% each of SDS and sodium deoxycholate can be included in the wash buffer for the first two wash steps.

5. RNase I Digestion

1. Dilute RNase I to 0.002-0.01 units/mL in 750 μL of IsoWB (the appropriate concentration for desired footprint sizes needs to be empirically determined). Add IsoWB-RNase I to washed FLAG agarose beads and incubate with gentle mixing at 4 °C for 10 min.
2. Pellet beads at 400 x g for 1 min at 4 °C. Collect 20 μL supernatant (RNase I elution) in a labeled tube for Western blot analysis. Discard IsoWB-RNase I and wash FLAG agarose beads 4x with IsoWB as described in step 4.3.

6. Affinity Elution

1. Prepare a stock of elution buffer (FLAG peptide at 250 ng/mL in IsoWB). Apply 375 μL of elution buffer to FLAG agarose beads and shake gently at 4 °C for 1-2 h. Pellet FLAG agarose beads and collect a 15 μL aliquot of the elution for Western blot of proteins in FLAG IP (see Figure 3A).
   NOTE: When the affinity elution is underway, section 7 can be performed.

7. Magnetic Bead-antibody Conjugation

1. Wash 50 μL magnetic beads (i.e., Dynabeads; Table of Materials) 3x in 1 mL IsoWB in 1.5 mL tube. Resuspend magnetic beads in 100 μL of conjugation buffer. Add appropriate amount of antibody (exact amount of antibody to use for IP will need to be empirically determined for each antibody).
   NOTE: Protein A magnetic beads are optimal for antibodies produced in rabbit whereas Protein G magnetic beads are more appropriate for mouse antibodies. Magnetic beads compatibility chart is available at supplier website to choose beads appropriate for each antibody.
2. Wash magnetic beads 2x in conjugation buffer (Table 1). Incubate mixture at room temperature for at least 10 min. Resuspend magnetic beads in 375 μL of RIPiT dilution buffer. Store on ice until next step.

8. Second Immunoprecipitation

1. Apply remaining FLAG affinity elution from step 6.1 to magnetic beads coupled to antibodies against the protein of interest. Incubate with gentle mixing at 4 °C for 1-2 h. Capture magnetic beads on a magnet and collect 15 μL of supernatant for analysis of unbound proteins via Western blot. Wash magnetic beads 7x with 1 mL of IsoWB.

9. Denaturing Elution

1. Add 100 μL of clear sample buffer (Table 1) to magnetic beads and resuspend with a P200 pipette. Incubate on ice for 10 min. Flick gently to resuspend beads periodically.
2. Capture magnetic beads on a magnet and collect 15 μL of elution for analysis of proteins in RIPiT elution via western blot (see Figure 3A). Transfer remaining elution into a new labeled 1.5 mL tube.
   NOTE: If samples were formaldehyde crosslinked, then samples must be incubated at 65 °C for 1 h to reverse crosslinking.
3. Perform western blots on samples collected at various steps (input, FLAG IP depletion, FLAG IP, second IP depletion, second IP elution). Blot with antibodies against the two bait proteins, their other interactors if known, and at least one non-interacting RBP as a negative control (Figure 3A).

10. RNA Extraction and End Curing

1. To the RIPiT elution, add 320 μL of RNase-free ddH₂O, 400 μL of phenol-chloroform isoamyl alcohol (PCIAA, pH 4.5), and vortex for 30 s and spin at room temperature at 12,000 x g for 5 min. Collect 350 μL of aqueous phase into a separate tube. Add 35 μL of 3 M sodium acetate, 1 μL of 1 M MgCl₂, 10 μg of glycogen, and 1 mL of 100% ethanol. Incubate overnight at -20 °C.
2. To pellet RNA, centrifuge at 12,000 x g for 30 min at 4 °C. Wash RNA in 70% ethanol.
3. To remove 3' phosphate left on RNA after RNase I cleavage, resuspend RNA pellet in 17 μL of RNase-free ddH₂O, and add 2 μL of 10x T4 polynucleotide kinase (PNK) buffer (Table of Materials) and 1 μL of T4 PNK. Incubate at 37 °C for 30 min.
   NOTE: The 3'phosphatase activity of T4 PNK has optimal activity at pH 6¹⁷. The PNK reaction buffer is optimized for the 5' kinase activity of T4 PNK and has a pH of 7.6. While adjusting the pH of the end-curing reaction has been attempted, this step can be further optimized.
4. Add 380 μL of RNase-free ddH₂O and 400 μL of PCIAA pH 4.5 to the tube. Vortex for 30 s, centrifuge at 12,000 x g for 5 min. Collect aqueous phase and add 35 μL of 3 M sodium acetate, 1 μL of 1 M MgCl₂, 10 μg of glycogen, and 1 mL of 100% ethanol.
5. Incubate overnight at -20 °C. Pellet and wash RNA with 70% ethanol as above. Resuspend RNA in 4.5 μL of RNase-free water.

11. Estimation of RNA Footprint Size and Abundance

1. A successful RIPiT is expected to yield 1 pmol or more of RNA fragments. To quantify actual yield, transfer 0.7 μL of RIPiT RNA (~1/6 of total yield) into a new tube. Add 2 μL of 10x T4 PNK buffer, 1 μL of 1 mM ATP, 40 μCi γ³²P-ATP (0.5−1.0 μL of the stock), and 1 μL of T4 PNK. Adjust volume to 10 μL and incubate at 37 °C for 30 min.
   1. In parallel PNK reactions, label a low range DNA Ladder and 0.1 pmol of a synthetic RNA or DNA oligo (20-40 nt) to use a size and quantity standards.
2. Resolve labeled RNA/DNA on 26% urea-PAGE gel (20 x 27 x 0.45 mm³). Gel must be pre-run for 30 min at 35 W. Flush wells before pre-run and before loading samples and run at 35 W until bromophenol blue dye front has almost reached the end of the gel.
3. Carefully remove gel from glass plates on to a piece of 8 x 11 inch filter paper. With gel on top of paper, place in gel drying apparatus and cover with a piece of plastic wrap. Dry gel at 80 °C for 1 h with vacuum.
4. Expose dried gel to a phosphoscreen overnight or until adequate signal is detected. Image phosphoscreen. Good quality RNA from a RIPiT should appear as a smear in the lane, with minimal prominent bands (Figure 3B). To quantify RNA, compare signal intensity of the desired size RNA fragments in RIPiT lane to the signal from 0.1 pmol of labeled synthetic oligo.
   NOTE: Alternatively, RNA footprint size and amounts can be verified using a high-sensitivity bioanalyzer (Figure 3C).

12. Adapter Ligation

1. Prepare RIPiT RNA such that at least 3 pmol of RNA is dissolved in 3.8 μL of water.
2. In a 0.2 mL polymerase chain reaction (PCR) tube, combine 3.8 μL of RNA, 1 μL of miR-CAT-33 pre-adenylated adapter (7 μM) (Table of Materials). Incubate the mix on a thermal cycler at 65 °C for 10 min, 16 °C for 5 min, then hold at 4 °C.
   NOTE: The pre-adenylated linker can be either ordered from oligo synthesis service, or a custom unadenylated DNA oligo from any oligo synthesis service can be adenylated using Mth RNA ligase (Table of Materials) and gel purified.
3. To the same tube, add 1.5 μL of 10x T4 RNA ligase buffer, 7.5 μL of 50% polyethylene glycol 8000 (PEG-8000), 0.75 μL of 20 mM dithiothreitol (DTT), and 0.45 μL of T4 RNL2 Tr. K227Q (Table of Materials).
   NOTE: 50% PEG-8000 comes with RNA ligase and buffer purchased. PEG-8000 solutions are viscous and should be pipetted slowly.
4. Incubate reaction in the thermal cycler at 30 °C for 6 h, heat inactivate the ligase at 65 °C for 10 min, then hold at 4 °C.

13. Reverse Transcription

1. To the tube with the ligation mix from step 12.4, add 11.25 μL of 4x deoxynucleotide triphosphate (dNTP) mix, which contains a mix of regular and biotinylated dNTPs (see Table 1), 1.0 μL of 10 μM RT primers (Table of Materials), and 6.8 μL of RNase-free water. Incubate at 65 °C for 5 min, then hold at 4 °C.
2. Transfer tubes to ice and add 9.0 μL of 5x first-strand (FS) buffer without MgCl₂ (Table 1), 2.25 μL of 100 mM DTT, 1.2 μL of reverse transcriptase enzyme to a final volume of 45 μL (Table of Materials).
3. Incubate in a thermal cycler at 55 °C for 30-60 min. Heat inactivate reverse transcriptase at 70 °C for 15 min and hold the sample at 4 °C.

14. Purification of RT Product

1. Add 45 μL of 2x urea load buffer (Table 1) to RT reaction. Dilute 1 μg of a low range DNA Ladder in 45 μL and add 45 μL of 2x urea load buffer.
2. Prepare 10% urea-PAGE gel (20 x 28 x 0.15 cm³; Table 1) with 8-well comb. Using syringe or pipet, flush wells with 0.5x Tris/borate/EDTA (TBE) buffer.
   NOTE: The homemade gels above offer better resolution in separating the extended RT product from the unextended RT primer. As an alternative, pre-cast urea-PAGE gels can also be used (Table of Materials). As pre-cast gels allow smaller maximum volumes per well, so samples will need to be divided into multiple wells. Pre-cast gels should be run at 150−200 V.
3. Pre-run the gel at 35 W for 30 min. Flush wells again, load samples and run gel at 35 W until bromophenol blue dye front has migrated to about 1 inch from the end of the gel.
   NOTE: Use metal heat sink during the pre-run and the final run to prevent gel overheating.
4. Stain the gel for 5 min in 1x gold nucleic acid gel stain solution prepared in 0.5x TBE. This dye is light sensitive, so avoid light exposure.
5. Image gel on fluorescent scanner for documentation purposes using 520 nm excitation and 580 nm emission filters. If a fluorescent scanner is unavailable, use a blue light transilluminator. The RT product should appear as a smear starting above the unextended RT primer (Figure 4).
   NOTE: Although the gold nucleic acid gel staining dye is easily visualized on a gel doc with UV-light source, it is vital to not expose the valuable RT product to UV to prevent damage.
6. Visualize the gel on a blue light transilluminator and excise the RT product from the gel. It is recommended to cut DNA with extensions ranging from 30-200 nt (Figure 4). Place the excised gel pieces on a clean surface and mince the slice into small pieces to increase surface area. Carefully transfer to a 1.5 mL tube and add 800 μL of DNA elution buffer (Table 1).
7. Incubate the gel pieces with gentle mixing with DNA elution buffer over night at room temperature.
8. Separate the elution buffer from the gel by passing the slurry through a cellulose acetate filter column (Table of Materials) placed in a 2 mL collection tube.
9. In 1.5 mL tube, wash 10 μL of streptavidin magnetic beads with 500 μL of streptavidin bead wash buffer (Table 1). Repeat for three total washes. Do not let beads dry out. Resuspend beads in 10 μL of DNA elution buffer (Table 1).
10. Transfer the elution buffer separated from the gel pieces in step 14.8 to the tube containing the washed streptavidin magnetic beads.
11. Incubate with gentle mixing for at least 8 h at room temperature. Capture beads on a magnet, remove supernatant and resuspend magnetic beads in 10 μL of RNase-free water and transfer to a 0.2 mL PCR tube.

15. Circularization of RT Product

1. RT products captured on streptavidin beads are circularized while bound on beads. To the magnetic bead slurry, add 2.0 μL of 10x circularization reaction buffer, 1.0 μL of 1 mM ATP, 1.0 μL of 50 mM MnCl₂, 4.0 μL of 5 M betaine, 1.0 μL of ssDNA ligase I (Table of Materials), and 1.0 μL of RNase-free water.
2. Incubate the circularization reaction on a thermal cycler at 60 °C for 4 h. Heat inactivate the ssDNA ligase I by heating at 80 °C for 10 min then hold at 4 °C.

16. Test PCR

1. Before proceeding to a large-scale PCR, use a portion of circularized product to determine the ideal number of amplification cycles for each sample. This step helps prevent over-amplification and limit PCR artifacts as PCR reaction components become limiting at higher PCR cycles.
2. Prepare a 45 μL PCR reaction with 4.0-6.0 μL of the circularized product from step 15.2, 9.0 μL of 5x reaction buffer, 0.9 μL 10 M dNTPs, 2.25 μL of 10 μM PE1.0 primer (Table of Materials), 2.25 μL of 10 μM PE2.0 primer (Table of Materials), and 0.045 μL of high-fidelity DNA polymerase (Table of Materials), and water.
3. Mix reaction well and split into three 15 μL reactions. Each of these three reactions will be subject to a variable number of PCR cycles. The ideal number of cycles is expected to be between 7 and 14. So perform test PCRs for 8, 11, and 14 cycles.
4. Use the following PCR conditions: 98 °C – 30 s; 98 °C – 5 s; 65 °C – 10 s; 72 °C – 15 s; 72 °C – 2 min; 12 °C – hold.
5. Add 3 μL of 6x gel loading dye and resolve on 10% native PAGE gel until blue dye front has migrated 3/4 of the gel. Stain the gel using gold nucleic acid gel stain and image as in steps 14.4 and 14.5 (Figure 5).
6. To choose the ideal number of PCR cycles, compare PCR products from increasing number of cycles. Choose the cycle number that yields the greatest amount of product of the expected size without overamplification artifacts (e.g., DNA smear much larger than expected product, and where no appreciable depletion of the PE1.0 and PE2.0 primers is seen (see red arrow in Figure 5).

17. Large-scale PCR

1. Prepare a 45 μL PCR reaction, as in step 16.2 and repeat the PCR. Resolve PCR on 10% native PAGE at 150 V, stain with 1x gold nucleic acid gel stain and image on a blue light transilluminator.
2. Excise the PCR product from the gel and transfer to a 3 mL syringe. Use the syringe to crush the gel and extrude into a 1.5 mL tube.
   NOTE: The unextended RT product upon circularization yields PCR product of 151 bp. Thus, at this step one should size select products that are larger than 151 bp (Figure 6).
3. Add 900 μL of DNA elution buffer and incubate at room temperature overnight with gentle mixing.
4. Transfer the gel slurry to a cellulose acetate filter column placed in a 2 mL collection tube. Spin at 12,000 x g for 3 min, collecting supernatant into a fresh tube.
5. Add another 400 μL of DNA elution buffer to the crushed gel and transfer to a 1.5 mL tube. Incubate with gentle mixing for an additional 4 hours for a second elution.
6. Pool all elutions and split into 3 tubes with 400 μL each. Precipitate DNA by adding 1 mL of 100% ethanol and 10 μg of glycogen. Vortex and incubate a least 2 h at -20 °C.
7. Pellet DNA at 12,000 x g for 30 min at 4 °C. Wash DNA pellet with 70% ethanol.
8. Carefully remove all ethanol by pipetting and quickly resuspend DNA pellet in 20 μL of water.
   NOTE: At this stage, it is important to not let the DNA pellet become dry, as drying DNA out can denature it.
9. Use a small portion of DNA sample to determine size and concentration of the PCR product via fluorometer and high sensitivity DNA bioanalyzer. Samples can now be submitted for sequencing on one of the platforms.
10. The sequenced reads can be processed (e.g., adapter removal, trimming to keep sequences >30 Phred score), aligned to the reference genome, and visualized on a browser such as UCSC genome browser (Figure 7).