Real-time Analysis of Transcription Factor Binding, Transcription, Translation, and Turnover to Display Global Events During Cellular Activation

All methods are in accordance and compliance with the Helmholtz Zentrum München institutional, state, and federal guidelines.

1. Preparation

1. Make a detailed plan of the experimental setup including a time schedule, when to add 4sU to the cell culture medium, and when to harvest the cells for each method. Depending on the biological question, carefully consider time points for sample drawing, 4sU-labeling time, and concentration (Table 1).
   NOTE: Verify the impact of 4sU on cell viability and stress response in advance (see "Verify optimal 4sU-labeling conditions" in Representative Results and Figure 1). It is recommended to perform a preliminary test of each method with at least one sample. Verify if the quality and amount of RNA/DNA is sufficient for deep sequencing (see dedicated parts of the protocol), and practice fast but gentle handling of the cells during the experiment.
2. Calculate the number of cells needed for every time point of each method (see Table 2 for a rough estimation when using primary T cells). Also, consider sequencing fewer samples of total RNA than just that of 4sU RNA (e.g., just for time point translation rates or RNA turnover rates should be calculated). To confirm and verify the significance of the results (recommended), create at least one biological replicate.
   NOTE: Importantly, for all methods and consecutive time points, samples must be from the same starting pool of cells. At least one dedicated researcher for each method is recommended.
3. Prepare everything necessary in advance (e.g., aliquoted 4sU, cycloheximide, Mammalian Lysis Buffer, and 1% formaldehyde). After addition of 4sU, avoid exposing the cells to bright light, as this may lead to crosslinking of 4sU-labeled RNA to cellular proteins¹⁸.
4. Pool all cells of interest in one flask just prior to treatment (Figure 2). Count the cells (e.g., with a hemocytometer) and use the required amount for the untreated control of each method (do not forget to also label the untreated control for 4sU-seq with 4sU). Treat remaining cells and immediately split the required number of cells for each time point and method. Handle cells as quickly as possible to minimize stress due to changes in temperature or CO₂ levels.
   NOTE: For example, samples are taken 1 h, 2 h, and 3 h after treatment, then one quarter of the cells is used for untreated control, and three quarters are used for treated control.

2. 4sU-Labeling

NOTE: This protocol is modified from Rädle, et al.¹⁹ Refer to their protocol for further information regarding metabolic labeling with 4sU. For all methods and consecutive time points, samples must originate from the same starting pool of cells.

1. Start of Labeling
   1. Thaw aliquoted 4sU just before use. Add 4sU at each time point directly to the medium containing cells of interest (refer to Table 2 for recommended T cell numbers, least 60 µg RNA per time point), mix gently, and place back into the incubator. Dispose remaining 4sU (do not refreeze).
   2. At the end of labeling, collect cells (e.g., cell scraper) and centrifuge at 330 x g for 5 min at 4 °C in polypropylene tubes (which resist high g forces). Aspirate medium and add reagent for RNA isolation (≥1 mL per 3 x 10⁶ cells, see materials for recommendation which to use) to each tube. Fully resuspend the pellet (≥1 mL per 3 x 10⁶ cells), incubate for 5 min at room temperature (RT), and freeze samples at -20 °C. Samples can be stored at -20 °C for at least 1 month.
      Caution: The reagents used for RNA isolation are extremely hazardous when getting in contact with skin or eyes. Handle these with care and consider the safety instructions.
2. RNA preparation using modified RNA isolation protocol
   1. Add 0.2 mL chloroform per 1 mL reagent for RNA isolation, and mix thoroughly by shaking for 15 s. Proceed as mentioned in the metabolic labeling protocol (step 1 – 12, 2. RNA preparation using modified trizol protocol) from Rädle et al.¹⁹
   2. Measure RNA concentration (see Table of Materials), according to the manufacturer's instructions. Use this RNA for total RNA-seq as well (see step 3. Total RNA-seq) or store at -80 °C for at least 1 month.
3. Thiol-Specific Biotinylation of Newly Transcribed RNA
   1. Start with 30 – 80 µg of total cellular RNA. 60 µg RNA should yield sufficient amounts of newly transcribed RNA.
   2. Prepare labeling reaction. Pipette in the following order (per µg RNA): 1 µL 10x Biotinylation Buffer, 7 µL RNA (containing 1 µg RNA diluted in nuclease-free H₂O), and 2 µL biotin-HPDP (1 mg/mL). Add biotin-HPDP last and mix immediately by pipetting. Wrap tubes with aluminum foil to avoid exposure to bright light. See the discussion for an alternative to biotin-HPDP. Incubate at RT for 1.5 h with rotation.
   3. Centrifuge appropriate 2 mL tubes (see Table of Materials) at 15,000 x g for 2 min. Pipette all of the biotinylated RNA into the pre-spun 2 mL tube, add an equal volume of chloroform, and mix vigorously. Incubate for 2 – 3 min until the phases begin to separate and bubbles start to disappear.
   4. Centrifuge at 15,000 x g for 15 min at 4 °C. Carefully transfer the upper aqueous phase into a new tube.
   5. Repeat steps 2.3.3. and 2.3.4. once. Add 10% of the volume of NaCl (5 M) and an equal volume of isopropanol to the aqueous phase. Centrifuge at 20,000 x g for 20 min at 4 °C. Discard supernatant.
   6. Add an equal volume freshly prepared 75% ethanol. Centrifuge at 20,000 x g. Discard the supernatant, spin briefly, and remove the remaining ethanol. Fully resuspend in 30 – 100 µL H₂O (use 1 µL H₂O per 1 µg input RNA from step 2.3.1) by pipette mixing.
   7. Verify RNA integrity by electrophoretic analysis, or take an aliquot and verify later.
4. Separation of Newly Transcribed (Labeled) and Preexisting (Unlabeled) RNA
   1. Remove paramagnetic beads (see Table of Materials) from 4 °C storage and let it stand for at least 30 min to bring them to room temperature. Heat 4sU washing buffer (3 mL per sample) to 65 °C.
   2. Prepare 100 mM dithiothreitol (DTT) solution. Weigh DTT on an ultra-fine scale and add the required amount of nuclease-free water. Always prepare fresh. Use 200 µL per sample.
   3. Heat biotinylated RNA samples (1 µg/µL) to 65 °C for 10 min to denature and immediately place on ice. Add 100 µL streptavidin beads to biotinylated RNA and incubate at RT for 15 min with rotation.
   4. Place an appropriate column (see Table of Materials for recommendations) for each sample into the magnetic stand, and pre-equilibrate each column with 1 mL room temperature 4sU washing buffer.
      NOTE: This will take approximately 5 – 10 min. If any of the columns do not initiate draining after 5 min, press gently on top of the column with a gloved finger.
   5. Apply an RNA/beads mix to the middle of each column. Discard the flow-through, unless unlabeled RNA needs to be recovered. If so, collect the flow-through and at least the first wash. Perform the recovery of RNA as described by Rädle et al. (step 1 – 7, 7. Recovery of Unlabeled, Unbound RNA)¹⁹.
   6. Wash three times with 0.9 mL 4sU washing buffer (preheated to 65 °C from step 2.4.2.) and 0.9 mL RT 4sU washing buffer, respectively.
   7. Use paramagnetic beads to recover newly transcribed RNA. Pipette 400 µL of well-dispersed RT paramagnetic beads in a tube per sample and place underneath each column. Elute newly transcribed RNA with 100 µL 100 mM DTT. Wait for 3 min and perform a second elution with 100 µL 100 mM DTT. (Optional: Perform elution and recovery as described by Rädle et al.)¹⁹
   8. Mix newly transcribed RNA/beads thoroughly by pipette mixing 10 times and proceed according to the manufacturer's guideline. Elute RNA in 11 µL nuclease-free H₂O.Quantify RNA using a suitable fluorometer (see Table of Materials). RNA can be stored at -80 °C for at least 1 month.
      NOTE: Newly transcribed RNA can be used to prepare cDNA libraries for next-generation sequencing (see Table of Materials for a suggestion on which kit to use) or further downstream analyses. 100 – 500 ng RNA are sufficient for most library preparation kits (see Discussion).

3. Total RNA-Seq

1. Directly take RNA from 4sU labeled RNA after RNA preparation using the modified reagent for RNA isolation protocol for total RNA-seq (see step 2.2.2.)
2. For library preparation, dilute an RNA aliquot to a final concentration of 50 – 100 ng/µL. Use the same kit as for library preparation of newly transcribed RNA. 100 – 500 ng RNA are sufficient for most library preparation kits.

4. Ribosome Profiling

NOTE: For all methods and consecutive time points, samples must be from the same starting pool of cells . For recommendations on which kit to use, refer to the Table of Materials.

1. Preparation and Isolation of Ribosome Protected Fragments (RPFs):
   1. Use appropriate amounts of cells for each time point (refer to Table 2 for recommended T cell numbers). Treat adherent cells with cycloheximide as described in the manufacturer's protocol.
      Caution: Cycloheximide is highly toxic and may cause mutations. Avoid skin contact and inhalation.
   2. Collect and pool non- or semi-adherent cells from each time point into one polypropylene tube and adjust the final concentration to 1 x 10⁶ cells per mL of cell-specific medium (e.g., supplemented RPMI for T cells). Add cycloheximide with a final concentration of 0.1 mg/mL, mix by inverting the polypropylene tube, and incubate for 1 min. Centrifuge cells for 5 min at 330 x g at 4 °C. Aspirate medium and wash cells with at least 10 mL PBS supplemented with cycloheximide (final concentration of 0.1 mg/mL).
   3. Centrifuge cells for 5 min at 330 x g at 4 °C. Aspirate medium and add 100 µL mammalian cell lysis buffer per 10 x 10⁶ cells. Mix by pipetting and expel through a sterile 22 25-gauge needle to lyse the cells completely.
   4. Transfer the cell lysate to a precooled 1.5 mL tube. Incubate for 10 min on ice with periodic inversions. Centrifuge for 10 min at 20,000 x g at 4 °C to clarify the lysate. Transfer the supernatant to a precooled 1.5 mL tube.
   5. Prepare a 1:10 dilution of the lysate with nuclease-free water and record an A₂₆₀reading using a spectrophotometer. Use nuclease-free water as a blank and a 1:10 dilution of mammalian cell lysis buffer as the standard. Calculate the A₂₆₀/mL concentration of the lysate according to the following equation:
      (A₂₆₀ cell lysate – A₂₆₀ Mammalian Lysis Buffer) x 10 dilution factor = A₂₆₀/mL
   6. Create 200 µL aliquots of the lysate on ice and proceed with nuclease treatment.
      NOTE: Optionally, prepare a 100 µL aliquot for total RNA, add 10 µL of 10% SDS and mix. Store at 4 °C and proceed with 4.3.2. Using total RNA from 4sU-labeled RNA is recommended (see step 3. Total RNA-seq).
2. Ribosome footprinting
   1. Perform nuclease treatment immediately without freezing the lysate. Add 7.5 units of nuclease (included in the recommended kit) for each A₂₆₀ of lysate. For example: 80 A₂₆₀/mL lysate x 0.2 mL lysate x 7.5 U/A₂₆₀nuclease = 120 U nuclease.
      NOTE: Optionally, titrate the nuclease for digestion as described by the manufacturer.
   2. Incubate the nuclease reaction for 45 min at RT with gentle mixing. Freeze 200 µL aliquots of the lysate with liquid nitrogen and store at -80 °C, or stop the nuclease reaction by adding 15 µL RNase inhibitor to each 200 µL aliquot, and continue with step 4.3.2.
3. Purification of RPFs
   1. Unfreeze one sample of nuclease digested RPFs and add 15 µL RNase inhibitor. Keep the samples on ice.
   2. Purify the RPFs according to the manufacturer's protocol (column purification is recommended) and measure RNA concentration on a spectrophotometer.
4. rRNA Depletion
   1. Use 5 µg of purified RPFs for rRNA depletion.
   2. Follow the manufacturer's protocol (step 1 – 2, Primary rRNA depletion) for rRNA depletion. Measure RNA concentration of rRNA depleted RPFs on a spectrophotometer.
5. PAGE Purification of RPFs
   1. Use 500 ng of rRNA depleted RPFs for PAGE purification.
   2. Prepare RNA Control, samples, and ladder for PAGE purification. Mix 5 µL RNA Control and 5 µL denaturing gel loading dye in a 0.5 mL microcentrifuge tube. Mix 10 µL of each RPF with 10 µL of denaturing gel loading, respectively. Prepare a ladder aliquot (4 µL 20/100 ladder, 1 µL nuclease-free water, and 5 µL denaturing gel loading dye). Load it between each sample and control to prevent crosscontamination.
   3. Denature samples and ladder by incubating at 95 °C for 5 min and immediately place on ice. Load 20 µL of each sample (optionally, load 10 µL and freeze remaining samples at 20 °C) separated by 10 µL of prepared ladder onto a 12% or 15% urea-polyacrylamide gel. Load 10 µL of RNA Control. Run the gel until the bromophenol blue band reaches the bottom of the gel (180 V, ~70 min) (Figure 3).
   4. Stain the gel according to the manufacturer's protocol at 4 °C. Use a dark-field transilluminator that emits blue light to visualize the RNA. Excise gel slices for each sample corresponding to ~28 and 30 nt in length. Take RNA Control as reference and excise it.
      NOTE: RPFs are hardly visible. Excise slices at the size indicated by the RNA Control – containing two oligos of 28 and 30 nt in length – even if samples are not visible.
   5. Puncture a hole in the bottom of 0.5 mL microcentrifuge tubes with a sterile 20-gauge needle. Transfer each gel slice into a separate tube and place capped tubes in a 1.5 mL tube. Centrifuge for 2 min at 12,000 x g. Repeat centrifugation if gel slices are not completely shred into the 1.5 mL tube.
   6. Elute the RNA from disrupted gel slices with 400 µL nuclease-free water, 40 µL ammonium acetate (5 M), and 2 µL SDS (10%) each overnight at 4 °C.
   7. Transfer the slurry to 1.5 mL filter tubes (provided with the recommended kit) with a 1 mL pipette tip (wide-bore tip or self-made 1 mL tip with cut end). Centrifuge for 3 min at 2,000 x g to separate eluted RNA from gel slices. Gently pipette aqueous solution into a 1.5 mL tube. Add 2 µL glycogen (provided with the recommended kit) and 700 µL 100% isopropanol and store at -20 °C for at least 1 h.
   8. Centrifuge at 4 °C for 20 min at 13,000 x g. Discard supernatant. Wash the pellet with pre-chilled freshly prepared 80% ethanol at 4 °C for 10 min at 13,000 x g. Discard supernatant and air-dry. Resuspend each sample in 20 µL and the RNA Control in 8 µL nuclease-free water. Store at -20 °C if needed.
6. Fragmentation, End Repair, 3' Adaptor Ligation, Reverse Transcription
   1. Perform the procedure as described by the manufacturer's protocol (Fragmentation and end repair, 3' adapter ligation, and reverse transcription).
7. PAGE Purification of cDNA
   1. Prepare samples, RNA Control and ladder for PAGE purification: Mix 10 µL of each sample and RNA Control with 10 µL denaturing gel loading dye, respectively. Prepare a ladder aliquot (4 µL 20/100 ladder, 1 µL nuclease-free water, 5 µL denaturing gel loading dye). Load it between each sample and control to prevent crosscontamination.
   2. Denature samples and ladder by incubating at 95 °C for 5 min and immediately place on ice. Load 20 µL of each sample (Optionally, load 10 µL and freeze remaining samples at 20 °C) separated by 10 µL of prepared ladder onto a 10% polyacrylamide/7 – 8 M urea/TBE gel. Load 10 µL of RNA Control. Run the gel until the bromophenol blue completely migrates out of the gel (180 V, ~60 min).
   3. Stain the gel according to the manufacturer's protocol at 4 °C. Use a dark-field transilluminator that emits blue light to visualize the RNA and excise the gel slices for each sample corresponding to ~70 – 80 nt.
   4. Proceed as described in step 4.5.5 – 4.5.8 and resuspend each sample in 10 µL nuclease-free water.
8. cDNA Circularization
   1. Prepare enough circularization master mix for all reactions by combining the following reagents for each sample on ice: 4.0 µL Circularization Reaction Mix, 2.0 µL ATP, 2.0 µL MnCl₂, and 2.0 µL Ligase.
   2. Add 10 µL of the master mix to each sample. Mix gently and centrifuge briefly. Incubate samples at 60 °C for 2 h. Immediately place samples on ice.
9. PCR Amplification
   1. Follow the manufacturer's protocol (step 1 – 3, PCR amplification) for PCR amplification. Use 4 µL of circularized cDNA for amplification with 9 PCR cycles for primary T cells, to achieve best results.
   2. Purify libraries and check their size distribution, according to the manufacturer's protocol (step 4 – 8, PCR amplification). The expected size of the amplified library is 140 – 160 bp (see Figure 4).
   3. For sequencing libraries, refer to the manufacturer's protocol and the sequencing facility for further guidance.

5. ChIP-Seq

NOTE: This protocol is modified from Blecher-Gonen, et al.¹⁴ Refer to their protocol for further information regarding ChIP-seq. For all methods and consecutive time points, samples must be from the same starting pool of cells.

1. Crosslinking and Harvesting the Cells
   1. Crosslink appropriate number of cells (refer to Table 2 for recommended T cell numbers) for each time point with a final concentration of 1% formaldehyde in a cellspecific medium (e.g., supplemented RPMI for T cells) for 10 min at room temperature with gentle rocking. Stop the crosslinking reaction by the addition of glycine to a final concentration of 0.125 M.
   2. Centrifuge cells at 330 x g for 5 min at 4 °C. Discard the supernatant and wash the cells in ice-cold PBS. Repeat step 5.1.2 twice and freeze cell pellets at -80 °C. Frozen pellets can be stored for at least 6 months.
2. Cell Lysis and Sonication
   NOTE: During all cell lysis and sonication steps, samples must be kept on ice or at 4 °C to minimize crosslink reversal and protein degradation.
   1. Resuspend cell pellets in 1 mL ice-cold cell lysis buffer with freshly added protease inhibitors to isolate the nuclei (add phosphatase inhibitors optionally). Incubate for 10 min on ice and centrifuge at 2,600 x g for 5 min at 4 °C.
   2. Aspirate supernatant and resuspend nuclei pellet in 1 mL ice-cold nuclei lysis buffer with freshly added protease inhibitors (optionally: add phosphatase inhibitors). Incubate for 10 min on ice. Sonicate the cells to generate a mean DNA size fraction of 0.2 – 1.0 kb (see Figure 5).
      NOTE: Sonication conditions need to be optimized according to the cell type and further conditions (e.g., cell number, volume, and buffer). For primary T cells, sonication for 20 – 25 cycles is recommended (see materials for detailed description).
   3. Take a 20 – 50 µL aliquot of the sheared chromatin and heat for 10 min at 95 °C and 1,000 rpm shaking to perform a fast reverse crosslink and verify chromatin size. Add 2 – 5 µL Proteinase K and incubate for 20 min at 56 °C and 1,000 rpm shaking. Perform heat inactivation for 10 min at 95 °C and 1,000 rpm shaking. Purify chromatin with an appropriate kit (see Table of Materials). Check chromatin size on a 1% agarose gel and use 100 bp Plus Marker.
   4. Centrifuge sheared chromatin with a mean DNA size fraction of 0.2 – 1.0 kb for 10 min at 20,000 x g and 4 °C to pellet insoluble chromatin and debris. Transfer the supernatant to a new tube and keep on ice.
   5. Keep 5 – 10% of sonicated chromatin as input. Freeze at -20 °C (used in step 5.5.2).
3. Couple Antibody to Beads
   1. Couple 10 µg antibody (e.g., anti-RNA Pol II; anti-Histon H3K36me3) in 220 µL PBS (with 0.5% BSA and 0.5% Tween 20) to 80 µL superparamagnetic beads coupled to protein G (see Table of Materials) for at least 1 h at room temperature with rotation.
   2. Place the tubes on a magnet. Wait until all beads are bound to the magnet and remove the supernatant. Further block with 6 µL sonicated salmon sperm DNA in PBS (with 0.5% BSA and 0.5% Tween 20) for 30 min at room temperature with rotation.
   3. Place the tubes on a magnet. Wait until all beads are bound to the magnet and remove the clear supernatant. Wash beads with ChIP IP buffer three times.
4. Chromatin Immunoprecipitation
   1. Dilute chromatin to 1 mL total volume in nuclei lysis buffer with freshly added protease inhibitors (optionally, add phosphatase inhibitors). Add ChIP IP buffer with freshly added protease inhibitors (optionally, add phosphatase inhibitors) to a final volume of 3 mL. Keep on ice or at 4 °C while antibody is coupled to the beads.
   2. Add diluted chromatin to the antibody coupled beads from step 5.3.3 and incubate over night at 4 °C with gentle rotation.
   3. Wash with the following buffers (1 mL each, see Table of Materials) at room temperature for 5 min with rotation, place tubes back on the magnet and remove supernatant: Wash Buffer I, Wash Buffer II, Wash Buffer III, and 2x TE pH 8.0 respectively.
   4. Discard supernatant and air-dry for ~5 min.
5. Reverse crosslinking
   1. Remove samples from the magnet. Add 50 µL elution buffer and mix by pipetting to elute protein-DNA complexes from the beads.
   2. Include input sample(s) from this step forward. Add elution buffer to input sample(s) for a final volume of 50 µL (to keep the buffer composition similar to the ChIP samples) and process together with the ChIP samples.
   3. Mix 3 µL of elution buffer and 2 µL of RNase (DNase free). Add 5 µL of the mix to each sample and incubate for 30 min at 37 °C.
   4. Mix 2.5 µL proteinase K, 1 µL glycogen, and 1.5 µL elution buffer per sample. Add 5 µL of the mix to each sample (1 U proteinase K and 20 µg glycogen per sample) and incubate for 2 h at 37 °C.
   5. Incubate the samples at 65 °C overnight (at least 4 h) with shaking to perform reverse crosslinking.
   6. Place the tubes on the magnet for at least 30 s and transfer the supernatant to a new tube. Samples can be frozen at -20 °C for up to 12 months.
6. DNA purification
   1. Add 140 µL of well-dispersed paramagnetic beads to 60 µL of sample (2.3:1 ratio). Carefully pipette up and down 25 times to mix thoroughly. Ensure that the liquid in each tube is homogenous. Incubate at room temperature for 2 min. Place the tubes on the magnet for 4 min, or until all beads are bound to the magnet, and discard the supernatant.
   2. Leave the tubes on the magnet and add 200 µL of freshly prepared 70% ethanol. Incubate the tubes for 30 s without disturbing the beads. Discard the supernatant and repeat this step once more. Aspirate ethanol completely and let the paramagnetic beads air-dry for 4 min.
      NOTE: Incomplete ethanol removal may seriously reduce DNA recovery and yield. Dry the pellet only until it is dry. Over-drying the pellet may reduce DNA recovery and yield.
   3. Remove the tubes from the magnet and add 20 µL 10 mM Tris-HCl (pH 8.0). Gently pipette the entire volume up and down 25 times to mix thoroughly. Incubate for 2 min at room temperature. Place the tubes back on the magnet for 4 min and transfer the supernatant to another tube.
   4. Measure the amount of DNA with a suitable fluorometer (see Table of Materials).
   5. Verify the ChIP was successful by qPCR (dilute 1 µL in 100 µl H₂O and use 2 – 5 µL for qPCR). Use specific primers for a positive (known binding site of protein of interest) and negative control (e.g., a gene that is silent and/or not a target of protein of interest).
      NOTE: Library preparation can be performed with 2 ng of ChIP DNA depending on the kit (see Table of Materials for a suggestion on which kit to use).