Reagents Setup
Cells
Per experiment, at least 3 cell lines are required: a. The parental cell that does not express APEX2. b. A cell line that expresses a V5-tagged APEX2 localized to a control compartment (often is the cytoplasm). c. A cell line expressing V5-tagged APEX2 localized to the compartment of interest.
For each sample cells should suffice for preparative protein and RNA assays as well as for Western blot and immunofluorescence controls. Thus at least 20x106 cells are required on the day cells are split for the experiment. IMPORTANT – maintain control of all cell lines at similar passage and at similar confluency (<100%) to minimize variability stemming from cell senescence, contact inhibition, variable growth rate etc…
This protocol was developed on Invitrogen HEK293 T-REX FlpIn cells as the parental lines, and stable cell lines were produced according to Invitrogen instructions, however it should be relatively easy to adjust it to any other cell culture systems.
CRITICAL! HEK293 cells are relatively weakly adherent, therefore all steps involving accessing the plates for washing or liquid additions should be done with extreme caution.
Cell media
Standard medium - (DMEM[Gibco, 11995-065], 10% FBS, 1:1000 PenStrep [Gibco, 15140-122]).
Pre-selection: Standard medium supplemented with 100 µg/ml Zeocin (frt site selection) + 15 µg/ml Blasticidin (tetR selection).
Post-selection: Standard medium supplemented with 100 µg/ml Hygromycin (pFRT insert) + 15 µg/ml Blasticidin (tetR selection).
Buffers
RIPA lysis buffer: 50 mM Tris, 150 mM NaCl, 0.1% (wt/vol) SDS, 0.5% (wt/vol) sodium deoxycholate and 1% (vol/vol) Triton X-100 in Millipore water. Adjust the pH to 7.5 with HCl. This solution can be stored at 4 °C for many months.
RNase T1 buffer: 20 mM Tris pH7.4, 150 mM NaCl, 2 mM EDTA, 1% NP40.
PNK buffer without DTT: 50 mM Tris pH7.5, 50 mM NaCl, 10 mM MgCl2.
Procedure
- Defrost cells in standard media.
- Expand each of the control and experimental strains in its own selective media: Parental T-Rex cells in pre-selection medium and APEX2-expressing cells in post selection medium.
- Collect and count cells, and split them into standard media as follows:
Splitting table:

- 16-24 hours after splitting the cells add 4-thiouridine (4SU) to the cell culture media (only to the 6 and 15 cm plates) to a final concentration of 100 μM. NOTE! 4SU stock concentration is 500 mM, so an intermediate x10 dilution in standard media to 50 mM is necessary, from which 2 μl should be added per ml of media. Also add Doxycycline (DOX) to all plates to a final concentration of 2 μg/ml. NOTE! Because of the low volumes of 24-wells an intermediate x100 dilution to 200 μg/ml is necessary from which 10 μl should be added per ml of media.
- Prepare the 500 mM stock of BP: Dissolve BP in DMSO. The 500 mM stock may need to be sonicated (was not needed in our case). Divide the solution into 50-μl aliquots, and store the BP stock at − 80 °C. These aliquots can be stored for several months.
Also prepare the 1M sodium azide stock: Dissolve sodium azide in Millipore water. Aliquots can be stored at – 20 °C or below for several months.
Treatments and fluorescence staining plan:

- 16 hours after 4SU labeling and DOX induction, bring standard media to 37 °C and PBS to room temperature. Have liquid N2 at the bench.
- Weigh required amounts of Sodium ascorbate and Trolox (see step 10b-d).
- Dilute the required amount of 500 mM BP stock 50 times in standard medium to 10 mM, and add 50 μl of the 10 mM BP per 1 ml of media to a final concentration of 500 μM BP. Return cells to incubator for 30 min. CRITICAL! The medium should be prewarmed to 37 °C to facilitate dissolution of the BP. The solution may need to be sonicated to fully dissolve.
- During the BP incubation: a. Make 1500 µl fresh X100 stock of 100 mM H2O2 in DPBS (dilute the 30% (wt/wt) H2O2 reagent (about 10 M H2O2 in water) into Dulbecco’s PBS (DPBS) immediately before using it to label cells. Do not store this solution. b. Make fresh 1M Sodium ascorbate solution by dissolving the pre-weighed powder in Millipore water immediately before making solution. Do not store this stock. c. Make fresh 500 mM Trolox stock by dissolving the pre-weighed powder in DMSO. Sonicate it well. Prepare this stock immediately before making quencher solution. Do not store this stock. d. Make 500 ml of fresh quencher solution: 10 mM sodium ascorbate, 5 mM Trolox and 10 mM sodium azide solution in DPBS. Make this solution immediately before it is to be used to quench the biotinylation reaction. Do not store this solution. e. Make 4% PFA in PBS.
- Dilute the 100 mM H2O2 1:100 directly to the cell media and briefly agitate to achieve a final concentration of 1 mM. Incubate the cells at room temperature for 1 min.
- Quickly aspirate the labeling solution and wash all cells three times in large volumes of quencher solution. CRITICAL! HEK293 cells will be lost at that stage, to reduce cell loss introduce the quencher solution gently and to the same location within the plate. Consider pouring-off the liquid rather than aspirating by vacuum. NOTE! Do not omit washes because they remove excess, unused BP probe left inside the cells. Make sure that the washes are performed using the quencher solution and not merely DPBS, so that BP radicals are quenched. NOTE! Due to the necessity to perform this step rapidly yet with great caution, we recommend limiting Proximity-CLIP experiments to up to 2 compartments of interest on top of the negative and control compartment.
- Pour the last wash, add 750 μl quenching solution, remove plate caps and crosslink the 6 / 15 cm plates with 0.4 J/cm2 of 312 - 365 nm UV light in a UV Crosslinker (Spectronics, Spectrolinker XL-1500 or comparable instrument).
- During UV crosslinking of the 6 / 15 cm plates, fix the 24 wells cells with 4% PFA in PBS for 20 minutes at RT. CAUTION! PFA work should be performed in chemical hood.
- During PFA crosslinking, remove the 6 / 15 cm plates from the UV instrument, place them on ice and collect the cells with the quenching solution to pre-chilled tubes. Pellet cells by 5 min at 300g at 4 °C centrifugation, remove the supernatant and snap freeze cells in liquid N2. Keep the frozen cell pellets at -80 °C, continue working on them as described in sections 20 and 30.
- Back to the 24 well: Wash the PFA-fixed cells 3 times with 1 ml PBS, and fix-perm cells in pre-chilled -20 °C Methanol. Incubate for 20 minutes at -20 °C.
- Block cells with 5% BSA in PBS for 60 minutes and incubate at 4ºC O.N. in the presence of primary antibody (see Table 2 for details). IMPORTANT! Centrifuge the diluted antibody for 10 minutes at max speed to remove aggregates.
- The next day, wash cells 3 times for 5 minutes with PBS, and incubated with secondary antibodies for 1 hour at room temperature (see Table 2 for details). IMPORTANT! Centrifuge the diluted antibody for 10 minutes at max speed to remove aggregates.
- Finally, wash cells 3 times for 5 minutes with PBS and mount over 9 μl vectashield with dapi.
Processing of cells from 6cm control plates:
- Prepare 100 mM PMSF, 7x PI cocktail and fresh sodium ascorbate and Trolox solutions.
- Prepare 4 ml RIPA lysis buffer supplemented with 1× protease inhibitor cocktail, 1 mM PMSF and fresh quenchers (10 mM sodium azide, 10 mM sodium ascorbate [7.9 mg/4 ml] and 5 mM Trolox [5 mg in 40 µl DMSO / 4 ml]).
- Lyse the cell pellets by gentle pipetting in 300 μl of the RIPA lysis buffer. After resuspension, leave the sample on ice for 2 min. Clarify the extract by centrifuging at 15,000g for 10 min at 4 °C, transfer the supernatant to fresh pre-chilled tubes and keep them on ice throughout the procedure. Typically, the protein concentration of the clarified extract is 1.2 μg/μl.
- Quantify the amount of protein in each clarified whole-cell lysate by using the Pierce 660-nm assay. If necessary, dilute the clarified whole-cell lysate first so that the concentrations fall in the linear range of the assay. Prepare triplicate samples for each condition.
- For each sample, take 15 μl aliquots of streptavidin magnetic beads. Note: when you are handling the streptavidin magnetic beads, use 200-μl or larger pipette tips whose tips have been cut off using a clean razor. Wash the beads twice with 1 ml of RIPA lysis buffer.
- Incubate 150 μg of each whole-cell extract with 15 μl of streptavidin magnetic beads at 4 °C overnight on a rotator. Add 150 μl of RIPA buffer to each sample to facilitate rotation. Note: this step can also be done for 1 h at room temperature. Save the remaining whole-cell lysate for gel and western blot analysis.
- Pellet the beads using a magnetic rack and collect the supernatant (‘flow-through’ [FT]). Save the FT on ice for subsequent analysis.
- Wash each bead sample with a series of buffers (1 ml for each wash) to remove nonspecific binders, as follows: Twice with RIPA lysis buffer, once with 1 M KCl, once with 0.1 M Na2CO3, once with 2 M urea in 10 mM Tris-HCl, pH 8.0 (freshly made), and twice with RIPA lysis buffer. Prechill and keep all wash buffers on ice throughout the procedure.
- Elute biotinylated proteins from the beads by heating each sample in 60 μl of 3× protein loading buffer supplemented with 2 mM biotin and 20 mM DTT for 10 min (97 °C, while vortexing). Let the samples cool, spin down and place on a magnetic rack to pellet the beads. Quickly collect the eluate to new tubes, and load 10 μl of them for western blot analysis.
For western blot analysis, prepare and boil the whole-cell lysate and corrected volumes of flow-through samples from the first replicate set in 1× protein loading buffer. Cool the samples on ice and spin them briefly to bring down condensation. Load and run the whole-cell lysate, streptavidin enrichment eluate and flow-through samples on a 4-20% (wt/vol) gradient SDS gel. See excel for amounts and volumes loaded. Perform a streptavidin-HRP western. Check that there is no biotinylated material left in the flow-through.
Processing of cells from 15 cm preparative plates:
- Prepare fresh quenchers.
- Prepare 4 ml RIPA lysis buffer supplemented with 1× protease inhibitor cocktail, 1 mM PMSF and fresh quenchers (10 mM sodium azide, 10 mM sodium ascorbate [7.9 mg/4 ml] and 5 mM Trolox [5 mg in 40 µl DMSO / 4 ml]).
- Lyse the cell pellets derived from the 15 cm plates by gentle pipetting in 800 μl of RIPA lysis buffer. After resuspension leave the sample on ice for 2 min. Clarify the lysates by centrifuging at 15,000g for 10 min at 4 °C. Keep the lysates on ice throughout the procedure. Typically, the protein concentration of the clarified sample is 3 μg/μl.
- Sample 30 μl of cell extracts for total RNA extraction (for RNA-seq) and ~80 μl (the remaining volume) for protein content analysis.
- For each sample, take 60 μl aliquots of streptavidin magnetic beads. Wash each aliquot of beads twice with 1 ml of RIPA lysis buffer. Then, incubate the cell extracts (~500 μl, or 1.5 mg) of each whole-cell lysate sample with 60 μl of streptavidin magnetic beads for 1 h at room temperature on a rotator.
- Pellet the beads using a magnetic rack and collect the FT, which should be saved for subsequent analysis.
- Wash each bead sample with a series of buffers (1 ml for each wash) to remove nonspecific binders: 2x RIPA lysis buffer, 1x with 1 M KCl, 1x with 0.1 M Na2CO3, 1x with 2 M urea in 10 mM Tris-HCl, pH 8.0 (freshly made), and 2x RNase treatment buffer (“NP40 lysis buffer”). Prechill all wash buffers and keep them on ice throughout the procedure.
Use the last wash step to split the beads into 3 aliquots: 1. 300 μl for Mass Spec – remove liquid and keep on ice. 2. 200 μl for RNA seq (No RNase treatment - remove liquid and freeze in -80 °C). 3. 500 μl for RNAse T1 treatment (continue immediately).
RNase T1 treatment and labelling of RNA footprints
- Resuspend the beads aliquots in 100 μl of RNase T1 buffer. Add RNase T1 to a final concentration of 1 U/μl and incubate for 15 min at 22 °C.
- Cool the reaction on ice for 5 min. Wash beads 2 times with RNase T1 buffer and once with dephosphorylation buffer (NEB cutsmart x1).
- Prepare 300 μl of dephosphorylation mix (30 μl of 10x cutSmart, 255 μl ddw, 15 μl CIP [stock conc. is 10 U/ μl]). Resuspend the beads in 60 μl of that mix (can go down to 1 bead volume) and incubate for 10 min at 37 °C with shaking. Note: adjust the shaking speed on the thermomixers so the beads do not settle.
- Wash beads 2 times with 1 ml of dephosphorylation buffer.
- Wash beads 2 times with 1 ml PNK buffer without DTT. Note: Exposure to > 1 mM DTT for a prolonged time may damage magnetic beads and should only be used in the reaction buffer.
- Resuspend the beads in 60 μl (can go down to 1 bead volume) of HOT (radioactive) reaction mix: Hot Mix: 245 μl ddw, 30 μl 10x PNK buffer (with DTT), 30 μl PNK, 5 μl *ATP (0.5 μCi γ-32P-ATP and 1 U/μl of T4 PNK).
- Incubate for 30 min at 37 °C with shaking. Note: adjust the shaking speed on the thermomixers so the beads do not settle.
- Add non-radioactive ATP to a final concentration of 100 μM and incubate for additional 5 min at 37 °C.
- Place tubes on magnet, keep 50 μl of the radioactive waste, which is collected after the first bead wash, to mark the Urea-PAGEs described below.
- Wash beads 5 times with 1 ml of PNK buffer without DTT. Note: Exposure to > 1 mM DTT for a prolonged time may damage magnetic beads and should only be used in the reaction buffer.
- Measure the radioactivity of the beads after washing is done.
Freeze the beads in a radioactive-approved -20 °C freezer and continue with the beads that were kept on ice for Mass Spec analysis.
Preparation of peptides for Mass Spectrometry analysis:
- Defrost the beads in 30 μl of 25 mM Ammonium bicarbonate and 20 mM DTT, shake for 1 hour at increasing temperatures: 25 °C (30 min), 37 °C (20 min), 56 °C (10 min).
- Add 6 μl of 200 mM Iodoacetamide (in 25 mM Ammonium bicarbonate), shake for 1 hour at 25 °C.
- Spin down, place tubes on magnet and collect the fluid (can be kept to verify that no protein was released).
- Wash beads 3 times in 200 μl of 1 mM DTT in 25 mM Ammonium bicarbonate, to wash and quench the remaining of Iodoacetamide, and to fully deplete NP40.
- Dissolve 20 μg of Trypsin in 1 ml of 25 mM Ammonium bicarbonate.
- To the beads tube add 98 μl of 25 mM Ammonium bicarbonate, and 2 μl of Trypsin (40 ng). Shake O.N. at 37 °C with cap to avoid condensation. Add the same components to an empty tube as a control.
- Spin down, place tubes on magnet, and transfer the eluate to fresh tubes.
Boil the remaining of beads in 30 μl of 3× protein loading buffer supplemented with 2 mM biotin and 20 mM DTT for 10 min (97 °C, while vortexing). Spin down and place on a magnetic rack to pellet the beads. Collect the eluate to new tubes and store (in case it will be required to test if any proteins were remained on the beads due to trypsin failure.
Returning to work on beads coupled to intact and RNase T1-treated RNPs:
- Bring the non-labelled intact (for RNA seq) and 32P-labelled (for PAR-CLIP) beads out of freezers, perform cold and hot proteolysis in parallel:
- Release the RNA from the beads by digesting the proteins with proteinase K in three subsequent steps, each time adding to the existing volume to make a final volume of 500 μl:
a. add 1.2 mg/ml proteinase K in 200 μl of 1x Proteinase K buffer (50 mM Tris pH 7.5, 75 mM NaCl, 6.25 mM EDTA, 1% SDS). Incubate the sample at 50 °C in a heat block under vigorous shaking for 30 min (for 9x - weigh 2.16 mg and resuspend in 1.8 ml buffer).
b. add 0.75 mg/ml proteinase K in 150 μl of 1x Proteinase K buffer. Incubate the sample at 50 °C in a heat block under vigorous shaking for 30 min (for 9x - weigh 1.01 mg and resuspend in 1.35 ml buffer).
c. add 0.75 mg/ml proteinase K in 150 μl of 1x Proteinase K buffer. Incubate the sample at 50 °C in a heat block under vigorous shaking for 30 min (for 9x - weigh 1.01 mg and resuspend in 1.35 ml buffer).
Place tubes on magnet and transfer the supernatant to a new 1.5 ml microcentrifuge tube, measure beads radioactivity and discard them.
Extraction of 32P-labelled RNA footprints:
- To the supernatant, add 30 μl of 5 M NaCl and 300 μl acidic phenol-chloroform (pH 4.5), mix by vortexing, and incubate on the bench for 10 minutes.
- Centrifuge at 12000g for 10 min and transfer the top of the aqueous phase (300 μl) to a new 1.5 ml microcentrifuge tube.
- Add 300 μl of chloroform and mix by vortexing.
- Centrifuge at 12000g for 7 min and transfer the aqueous phase to a new 1.5 ml microcentrifuge tube.
- Precipitate the RNA by adding 1 µl of glycol- Blue (10mg/ml), mixing, followed by addition of 3 volumes of ethanol and incubation at -20 °C for at least 1 hr.
- Centrifuge the sample at >12000g for 20 min and remove all ethanol traces.
- Let pellets dry for 5 min at room temperature.
Dissolve the pellet in 20 μl of water.
Extraction of non-labelled bound RNA:
- To the supernatant, add 30 μl of 5 M NaCl and 300 μl acidic phenol-chloroform (pH 4.5), mix by vortexing, and incubate on the bench for 10 minutes.
- Centrifuge at 12000g for 10 min and transfer the top of the aqueous phase (300 μl) to a new 1.5 ml microcentrifuge tube.
- Add 300 μl of chloroform and mix by vortexing.
- Centrifuge at 12000g for 7 min and transfer the aqueous phase to a new 1.5 ml microcentrifuge tube.
- Precipitate the RNA by adding 1 µl of glycol- Blue (10mg/ml), mixing, followed by addition of 3 volumes of ethanol and incubation at -20 °C for at least 1 hr.
Spin at 4 °C for 15 minutes at max speed, wash pellets twice with 75% EtOH, air dry for 5 minutes and resuspend in 20 µl of DEPC ddw.
Extraction of total cell-extract RNA
- To the 30 µl samples of cell extracts, add 370 µl ddw DEPC and immediately after 400 µl biophenol, Vortex, incubate 15 min on bench, spin 10 min at max speed.
- Transfer 200 µl of aqueous phase to new tube, add same volume of ddw-saturated chloroform, vortex and spin for 8 minutes at max speed.
- Transfer 100 µl of aqueous phase to new tube, add 7 µl of 3 M NaAc and 400 µl of cold 100% EtOH, vortex and incubate at -80 °C for at least 3 hours.
- Spin at 4 °C for 15 minutes at max speed, wash pellets twice with 75% EtOH, air dry for 5 minutes and resuspend in 20 µl of DEPC ddw.
Preparation of RNA-seq libraries:
Prepare RNA-seq libraries according to the NEBNext® Ultra™ RNA Library Prep Kit for Illumina manual with the following relevant info: a. Perform rRNA depletion only on total RNA samples. b. Perform RNA fragmentation for 10 minutes.
Preparation of sRNA cDNA libraries from the 32P-labelled RNA footprints:
- Determine the size of RBP-protected RNA fragment by denaturing polyacrylamide electrophoresis as described in5.
- Image the gel after O.N. radiography.
- Excise fragments representing 20 - 40 and 41 - ~80 nt lengths from the gel.
Extract the RNA from the gel by: a. 1 min at Max speed centrifugation in gel breaker tube. b. Addition of 350 μl of 0.3 M NaCl. c. Shaking 1 hour at 60 °C. d. Centrifuge 1 min at 5000g in filter tube. e. Add 1 μl of Glyco-Blue + 1200 μl of EtOH, vortex, incubate >3 hours at -80 °C. f. Centrifugation for 15 min at max speed, air dry and resuspend in 8.7 μl ddw.
3’ Adapters ligation
- Prepare 3’ ligation mix:
- To each 8.7 μl RNA add 8.3 μl mix +2 μl of the adenylated 3’ adapter.
- Incubate 1 min at 90 °C, and re-place in ice.
Add 1 μl T4 Rnl2(1-249)K227Q (1μg/μl), mix gently and incubate O.N. on ice.
3’ ligation gel
- To each tube add 20 μl denaturing PAA gel loading solution.
- Incubate 1 min at 90 °C, and load on 15% denaturing PAA gel.
- Excise ligated footprints from just below the ligated 19 size marker (for 20-40) / 35 size marker (for 40-80) to the top of detected RNA.
Extract the RNA from the gel by: a. 1 min at Max speed centrifugation in gel breaker tube. b. Addition of 350 μl of 0.3 M NaCl. c. Shaking 1 hour at 60 °C. d. Centrifuge 1 min at 5000g in filter tube. e. Add 1 μl of Glyco-Blue + 1200 μl of EtOH, vortex, incubate >3 hours at -80 °C. f. Centrifugation for 15 min at max speed, air dry and resuspend in 9 μl ddw.
5’ ligation
- Prepare the following mix and add 9 μl to each tube (mix):
- Incubate at 90 °C for 1 min to denature the RNA and put back on ice.
Add 2μl Rnl1 (1 mg/ml, ABI), incubate 1 hour at 37 °C.
5’ ligation gel
- To 20 μl RNA add 20 μl denaturing PAA gel loading solution.
- Incubate 1 min at 90 °C, and load on 12% denaturing PAA gel, as described in5.
- Image the gel after O.N. radiography and excise the ligated RNA.
Extract the RNA from the gel by: a. 1 min at Max speed centrifugation in gel breaker tube. b. Addition of 350 μl of 0.3 M NaCl. c. Shaking 1 hour at 60 °C. d. Centrifuge 1 min at 5000g in filter tube. e. Add 1 μl of Glyco-Blue + 1200 μl of EtOH, vortex, incubate >3 hours at -80 °C. f. Centrifugation for 15 min at max speed, air dry and resuspend in 4.6 μl ddw.
Reverse transcription
- Denature the RNA at 90 °C for 1 min.
Chill to 50 °C and add 9.7 μl of the following mix + 0.7 μl Superscript III.
Mix and incubate for 1 hour at 50 °C.
PCR amplification
- Dilute each cDNA by addition of 85 μl of DEPC ddw (reaching a volume of 100 μl).
- Set a 60 μl PCR reaction as follows:
*Not added as a mix
- Split the mixture into 6 tubes and run with the following protocol:
2 min x 94 °C
45 sec x 94 °C
85 sec x 50 °C
60 sec x 72 °C
4 °C…
- Remove a tube from the PCR cycler to ice after 9,11,13,15,17,19 cycles.
- Load the entire tubes content on a 2.5% agarose gel and run for 1 hour. Select the optimal #cycles.
- Setup an identical PCR mixture at a volume of 300 μl, and run the PCR for the optimal number of cycles (split to 3 tubes with 100 μl each).
- Concentrate the PCR product by ZYMO PCR purification kit to 70 μl.
- Size-select 30 μl of the library using PipenPrep to deplete the residual primers and adapter-adapter products based on the following expected sizes: Linker-Linker - 126bp, 20-40 nt footprint libraries: 146-166 bp, 40-80 nt footprint libraries: 166-196 bp.