Practical Aspects of Sample Preparation and Setup of ¹H R_1ρ Relaxation Dispersion Experiments of RNA

Figure 2: Schematic representation of the reported tandem IVT protocol. (A) Tandem transcription from a linearized plasmid template with T7RNAP (left) and successive cleavage by RNAse H of the transcript to achieve target length RNA, directed by a chimeric DNA guide (right). (B) Detailed schematic of the tandem template starting with the viral T7RNAP promoter, an initiation sequence. The target sequence (dark blue, example here is 20 nt long) is repeated “n” times. The repeats flanked by a 5′ and 3′ spacer sequences consisting of the last eight and first four nucleotides, respectively, to allow for removal of the initiation and restriction sequences from the first and last repeat unit. (C) Hybridization of the tandem transcript (red) and the chimeric cleavage guides (green). RNase H cleaves the RNA opposite to the DNA 5′ end. The 2′-OMe RNA flanks increase specificity by enhancing the binding affinity of cleavage guide to the target RNA. This figure has been modified from ²¹. Abbreviations: T7RNAP = T7 RNA polymerase. Please click here to view a larger version of this figure.

1. Preparing work for a new RNA construct

1. Plasmid design and preparation
   1. Write the template sequence in a cloning tool, e.g., Serial Cloner.
   2. Take the T7 promotor sequence and add a high-yield initiation sequence (T7: 5'-TAATACGACTCACTATA ^GGGAGA-3').
      NOTE: Transcription will start at the nucleotide indicated with a caret (^). The initiation sequence GGGAGA is variable, but strongly sequence-dependent; therefore, the use of this sequence is recommended.
   3. Add the last 8 nucleotides (nt) of the target sequence as a 5' spacer (5'S).
   4. Add repeats of the target sequence (TS).
   5. Add the first four nucleotides as a 3' spacer after the repeats (5'S).
   6. Add a BamHI restriction site (RS) or similar unique restriction site.
      NOTE: The total sequence as shown will be cloned or readily ordered in a bacterial high-copy plasmid (e.g., pUC19): 5'-T7-5'S-(TS)_n-3'S-RS-3' (Figure 2B). The number of repeats should be as high as allowed by gene synthesis ( a maximum of 600 nt in this protocol).
   7. Amplify the plasmid in E. coli using a commercial kit.
   8. Linearize the purified plasmid at 20 ng/µL using the appropriate restriction site. Scale restriction digests with BamHI by up to 1 mL.
   9. Purify the digested plasmid, and confirm successful linearization on a 1% agarose gel. Store the linearized plasmid at -20 °C for several months.
2. Cleavage guide design (Figure 2C)
   1. Write the last eight nucleotides of the target RNA sequence in the 5'-3' direction, and add the first four nucleotides of the target RNA sequence on the 3' end also in the 5'-3' direction.
   2. Generate the reverse DNA complement of that sequence
   3. Change the first and last four nucleotides to their 2'-OMe modifications by adding an 'm' before the nucleotide letter.
      NOTE: For synthesis, mU is used instead of mT.
   4. Order the oligo with standard desalting purification.
      NOTE: Check if the generated oligo could bind at another place other than the connection of two RNA sequences. Full complementarity in the central four DNA nucleotides is required, while the flanking regions could permit a mismatch. If necessary, extend the flanks to up to 18 nt to generate a unique binding sequence³⁶.
3. Small-scale IVT
   NOTE: For RNase-free work, prepare all reagents under sterile and RNase-free conditions. Use RNase decontamination reagent (see the Table of Materials) and 95% v/v ethanol to clean work surfaces and pipettes before use. Wash gloves with 95% ethanol and wear lint-free long-sleeved clothes. To minimize RNase contamination, do not breathe over open tubes.
   1. Prepare stock solutions of Tris-Cl (pH 8.0), dithiothreitol, MgCl₂, spermidine, and NTPs/GMP (unbuffered). Mix reagents as shown in Table 1. Prepare a master mix of these reagents in advance, before the addition of enzymes or nucleic acids.
      NOTE: If using frozen reagents, thoroughly mix them after thawing. Reagents might precipitate if mixed at too high concentrations, so it is strongly recommended to follow the order in Table 1.
   2. Add in the following order: plasmid, cleavage guide, inorganic phosphatase (IPPase), RNase H, T7RNAP. As enzyme activity might vary for enzymes produced in-house, test several concentrations before selecting the best one.
      NOTE: Include a negative control for the cleavage reaction, e.g., without RNase H, to attribute a missing target band to flawed RNase H cleavage and not to unsuccessful transcription.
   3. Incubate the reaction at 37 °C for 1 h and confirm reaction on a denaturing polyacrylamide gel electrophoresis (PAGE) (Figure 3A). Dilute the sample 10-fold in loading solution, and load 1 µL onto the gel.
      NOTE: Gel mixture: 8 M urea, 20% acrylamide (19:1 acrylamide:bisacrylamide) in 1x TBE. Loading solution: 5 mM ethylenediamine tetraacetic acid (EDTA), 300 µM bromophenol blue in formamide. RNase H cleavage reactions cannot be expected to be complete after 1 h, as new RNA is produced constantly. At this point, look out for a clear target band and the absence of a species of similar molecular weight (e.g., ±3 nucleotide (nt) products).

Table 1: Reagent table for tandem IVT and simultaneous RNase H cleavage. Stock concentrations can be adapted to the user’s convenience. If RNase H cleavage must be performed after T7 IVT, add cleavage guide and RNase H after heat-inactivation of T7RNAP. Amounts used scale linearly with reaction scale. Abbreviations: T7RNAP = T7 RNA polymerase; IVT = in vitro transcription.

2. NMR sample preparation

1. Scale up the reaction to the desired volume (typically 10 mL), and run the reaction overnight. Test for reaction completion the next day with a denaturing PAGE gel (Figure 3A).
   NOTE: Incomplete cleavage reaction is shown by higher molecular weight species above the target band.
   1. If cleavage was not successful or complete, reanneal RNA and the cleavage guide in the reaction vessel by heating the solution in a conventional microwave at 450 W for 15 s.
   2. Cool the solution slowly to 37 °C for 40 min. Use a heating block for volumes below 1 mL. Note the formation of new precipitate.
   3. Add more IPPase and RNase H, and incubate for another 1-3 h at 37 °C. Confirm completion of the cleavage reaction with denaturing PAGE.
   4. When the RNase H cleavage reaction is completed, quench the reaction by adding EDTA to 50 mM final concentration and vortex thoroughly.
      NOTE: Potential pyrophosphate precipitation will dissolve, and new protein precipitate forms.
   5. Filter the solution through a 0.2 µm syringe filter, and concentrate to a volume injectable into an HPLC system, depending on the injection loop size.
      NOTE: The protocol can be paused here by freezing sample at -20 °C.
2. Large-scale HPLC purification
   1. Prepare ion-exchange buffers A and B within one week of use. Filter and degas the buffers.
      NOTE: Buffer A: 20 mM sodium acetate; 20 mM sodium perchlorate, pH 6.5. Buffer B: 20 mM sodium acetate; 600 mM sodium perchlorate, pH 6.5.
   2. Equilibrate the column with 100% buffer B followed by 100% buffer A for at least 2 column volumes at 75 °C.
   3. Prepare the HPLC sequence (Figure 3B) at a flow rate of 5.5 mL/min. Use the following sequence for purification of an RNA sized between 20 and 30 nt: 0-7 min: 0% B; 7-16 min: gradient 0-20% B; 16-46 min: elution, typically with a gradient of 20-30% B (optimize according to needs); 46-62 min: 100% B; 62-73 min: 0% B.
      NOTE: A change in flow rate from 5.5 to 8 mL/min did not influence the separation in this protocol.
   4. Optimize the elution gradient by the injection of an equivalent to 1 mL of transcription reaction (unlabeled) at a time.
      NOTE: For further details and discussion, refer to Karlsson et al.³¹ and Feyrer et al.²¹.
   5. Test the collected fractions on a denaturing PAGE. If the main elution peak is well isolated and contains the pure target RNA, scale up the purification to an equivalent of 10 mL of transcription reaction.
   6. Collect the fractions of interest, concentrate, and exchange the buffer with NMR buffer. Use an ultracentrifugal filter unit (see the Table of Materials) for volumes above 50 mL.
      NOTE: NMR buffer: 15 mM sodium phosphate; 25 mM sodium chloride; 0.1 mM EDTA, pH 6.5. To minimize loss from RNA adhering to plastic tube walls, wash all collection tubes with 1 mL of water, vortex, and centrifuge to collect all liquid.
   7. Determine the concentration via ultraviolet spectroscopy. Calculate the reaction yield according to Feyrer et al²¹.
      NOTE: The concentration of an NMR sample for RD experiments should not be below 130 nmol, which corresponds to 500 µM in a sample volume of 250 µL using NMR tubes (Table of Materials).
3. Folding of an RNA sample
   1. Dilute and aliquot the sample of a volume of ~10 mL into 1 mL per tube.
   2. Heat the RNA aliquots to 95 °C for 5 min.
   3. Snap-cool the samples by placing them on ice or in a water-ice-salt mixture and incubate for 30 min.
   4. Pool samples and concentrate to ~250 µL in a 2 mL centrifugal filter unit.
4. Filling of an NMR tube
   1. Clean the NMR tube in the NMR tube cleaner by flushing with abundant water, RNase decontamination reagent, water, 95% ethanol (EtOH), and water again. Leave to dry.
   2. Clean the plunger by rinsing with water and wiping with RNase decontamination reagent and 95% EtOH using a lint-free wipe. Leave to dry.
   3. Add 10% (v/v) of D₂O to the NMR sample.
   4. Fill the RNA sample into the NMR tube using a large pipette tip. Let the liquid flow along the side of the tube wall.
   5. Insert the plunger and remove air bubbles by pushing the plunger down together with a fast twisting motion.
   6. Pull the plunger up slowly without creating new air bubbles and fix it with paraffin wax film.
5. Confirm folding by NMR.
   NOTE: At this point, it is necessary to perform at least partial resonance assignment to confirm the secondary structure of the RNA sample and to identify regions of interest for the study of conformational dynamics. An exhaustive description on RNA resonance assignment would exceed this protocol, therefore we refer to well-established literature at this point^19,37,38. An electrophoretic mobility shift assay (EMSA) can be a helpful indicator of RNA folding and serve as complementary data for NMR experiments.
   1. Compare the following spectra of the sample for which ¹H R_1ρ RD experiments are performed with the properly folded reference sample (Figure 4): ¹H 1D, especially the imino region 10–15 ppm; Aromatic ¹H,¹³C-HSQC; ¹H,¹H-SELOPE (optional).
      NOTE: An aromatic fingerprint is also necessary, even in case of agreement between imino signals, because dimer formation often shows the same or similar imino signals as an RNA hairpin. The SELOPE experiment can replace an ¹H,¹³C-HSQC for aromatic fingerprinting, as heteronuclear experiments on unlabeled samples are very time-consuming.
   2. Use the UUCG loop as a fingerprint reference (if present).
   3. Perform this comparison every time before ¹H R_1ρ RD experiments are recorded.

3. ¹H R_1ρ Relaxation dispersion—on-resonance (labeled 1D version)

NOTE: The steps below describe the setup of RD experiments for a labeled sample using the 1D version of the HSQC-based RD pulse sequence. Follow the same steps for the SELOPE-based 1D sequence for unlabeled samples. An overview of parameter names and settings for both cases are shown in Table 2. The focus on 1D versions is because they are more practical for off-resonance measurements, and the setup of the 2D versions of the SELOPE and HSQC-based experiments have been discussed in detail by Schlagnitweit et al.²⁰ and Steiner et al.¹⁰, respectively.

1. Determine ¹H power for a hard 90° pulse (P1).
   1. Option A: Use Bruker pulsecal command.
   2. Option B: In a zg experiment, determine the 360° pulse by measuring a nutation curve at the proton hard-pulse power level on the water peak.
      NOTE: The 90° pulse length is a fourth of the duration where zero signal is observed (if a full nutation curve is measured, then it is the second zero; however, in practice, only the region around the expected value for the 360° is sampled).
2.  Run a ¹H 1D spectrum zgesgp.f2f3dec using the pulse length determined in step 3.1 to confirm the RNA folding before every R_1ρ measurement.
   NOTE: If ¹H SL experiments are run for the first time, check if the calculated SL power corresponds to the power delivered to the sample by calibrating SL power for each desired bandwidth. Detailed calibration steps are described in Steiner et al.¹⁰.
3. Create a ¹H R_1ρ for labeled data set, and set key parameters.
   1. Create a new data set; ideally based on a ¹H-¹³C aromatic HSQC data set as used on fully labeled RNA samples for RNA assignment.
      NOTE: This will ensure that ¹³C as well as ¹⁵N power and decoupling power are already set up.
   2. Set the general parameters according to the first part of Table 2.
   3. Set RD-specific parameters according to the second part of Table 2.
   4. Set ¹H SL power to the lowest value (1.2 kHz) for testing.
   5. Generate a test vd list with only one entry, 0 ms, (to optimize the vd list, as described in step 3.4), set TDF1 to 1, and update D30.
   6. Run a test spectrum with these settings.
4. Optimize the vd list (list of SL lengths to be used).
   1. Run the experiment with a test vd list (e.g., six entries: 0 m, 5 m, 10 m, 20 m, 30 m, 40 m; scramble these values to avoid systematic errors due to heating).
   2. Update D30 and TDF1 accordingly (in this example, D30 = 42m and TDF1 = 6).
   3. Plot intensity of the peak vs. SL length. Identify the SL length at which the intensity of the original peak decreases to 1/3.
   4. Create the final vd list to be used in the experiment, taking into consideration the following: determine the longest SL length as described in the previous step; avoid using a list with decreasing or ascending order; and add some duplicates for statistical studies. Remember to update D30 and TDF1 every time there are any changes in the vd list.
      NOTE: The experiment is run with different SL lengths as given in the vd list in a pseudo-2D way.
   5. Select the number of scans so that the weakest peak of the list has a signal-to-noise ratio (SINO) of at least 10.
      ​NOTE: Although the vd list was optimized for a low SL power (1.2 kHz), this vd list should also be tested at the highest SL power to be used (e.g., 15 kHz). This is because the decay will be much slower at high SL power for peaks with significant k_EX contribution. Therefore, a sufficient decay should also be verified at high SL power.

Table 2: Overview of parameters to set up 1D HCP-based and 1D-SELOPE-based ¹H R_1ρ experiments. Abbreviations: 1D = one-dimensional; HCP = heteronuclear cross-polarization; SELOPE = SELective Optimized Proton Experiment; ppm = parts per million; HSQC= heteronuclear spin quantum correlation; SL = spin lock; RD = relaxation dispersion

5. Set-up and acquisition of on-resonance ¹H R_1ρ experiments
   1. Copy the experiment from section 3.4 into a new folder in Topspin.
   2. In this folder, set up experiments at different SL strengths, each time changing PL25 and CNST12. Determine the correct power level for each SL strength by using the pulse command. Use SL strengths ranging from 1.2 to 15 kHz, with a denser sampling for lower SL strengths (see Figure 5G for selected SL strengths). Add copies of some of the experiments to have duplicates for some of the SL strengths.
   3. Run these experiments.

6. Analysis of on-resonance ¹H R_1ρ experiments
   1. In TopSpin, process each slice of each pseudo-2D data set using the same processing parameters (e.g., line broadening, phase) using the command xf2, and split the dataset into 1Ds using the Bruker AU program split2D.
   2. Obtain signal intensities and volumes for each 1D slice.
      NOTE: In practice, it is better to deconvolve the spectra to get rid of contributions from potentially overlapping peaks and allows the usage of the Bruker AU program multidcon, which conveniently summarizes the intensities or areas of the peaks of all slices in one experiment in the text file decall.txt, which can then be read out easily with other programs (Python scripts written in-house were used here, as described by Steiner et al.¹⁰) in steps 3.6.3 and 3.6.4.
   3. Fit a mono exponential decay for each SL strength to obtain the R_1ρ (or on-resonance, R₂+R_EX) value.
   4. Plot those R₂+R_EX values (y) vs. SL strength (x) (Figure 5F,G).
      NOTE: If the values are significantly higher for low SL strengths and decrease with higher SL power (as shown in Figure 5G), then the investigated peak shows dispersion, and it might be interesting to carry out additional (off-resonance) experiments to obtain information about the population and chemical shift difference of the excited state vs. the ground state.

4. ¹H R_1ρ Relaxation dispersion—off-resonance (labeled 1D version)

1. Set-up and acquisition of off-resonance ¹H R_1ρ experiments
   1. In a new topspin folder, set up experiments at a certain SL strength (usually first at the lowest SL strength as the R_EX contribution is highest there, see Figure 5G for a representative selection of off-resonance SLs), but with different offsets, each time changing CNST30.
   2. Use offsets up to ± (3 or 4)*SL strength, with a denser sampling around 0 offset, as can be seen in Figure 5H,I.
   3. Run these experiments.
2. Analysis of off-resonance ¹H R_1ρ experiments
   1. Use the same processing strategy, as in 3.6.1–3.6.3, to determine an R_1ρ value for each offset.
   2. Plot those values vs. offset (Figure 5G).
      NOTE: An asymmetry in this curve can already indicate that chemical shift information for the excited state can be obtained. Thorough fitting and analysis using Bloch-McConnell or Laguerre equations have to be carried out to obtain information on k_EX, p_ES as well as Δω^10,20 (Figure 5G). Example datasets, pulse programs, and macros for both 1D experiments can be found on the Petzold Lab Github repository (https://github.com/PetzoldLab). An overview of parameters is given in Table 2.