Summary

Paramagnetic Relaxation Enhancement for Detecting and Characterizing Self-Associations of Intrinsically Disordered Proteins

Published: September 23, 2021
doi:

Summary

A protocol for the application of paramagnetic relaxation enhancement NMR spectroscopy to detect weak and transient inter- and intra-molecular interactions in intrinsically disordered proteins is presented.

Abstract

Intrinsically disordered proteins and intrinsically disordered regions within proteins make up a large and functionally significant part of the human proteome. The highly flexible nature of these sequences allows them to form weak, long-range, and transient interactions with diverse biomolecular partners. Specific yet low-affinity interactions promote promiscuous binding and enable a single intrinsically disordered segment to interact with a multitude of target sites. Because of the transient nature of these interactions, they can be difficult to characterize by structural biology methods that rely on proteins to form a single, predominant conformation. Paramagnetic relaxation enhancement NMR is a useful tool for identifying and defining the structural underpinning of weak and transient interactions. A detailed protocol for using paramagnetic relaxation enhancement to characterize the lowly-populated encounter complexes that form between intrinsically disordered proteins and their protein, nucleic acid, or other biomolecular partners is described.

Introduction

Intrinsic disorder (ID) describes proteins (IDPs) or regions within proteins (IDRs) that do not spontaneously fold into stable secondary or tertiary structures but are biologically active. Generally, the function of IDP/IDRs is to facilitate specific yet reversible interactions with biomolecules at physiological conditions1. Thus, IDPs and IDRs are involved in a range of cellular functions, including recruitment, organization, and stabilization of multi-protein complexes, for example, the assembly and activity of the spliceosome2, recruitment and organization of components at sites of DNA damage3, organization, and stabilization of the recruitment of transcription complexes4, or of the chromatin remodeler BAF5. Additionally, IDPs are found at signaling nexuses where their promiscuity for different binding partners enables them to mediate information transfer through cellular protein networks6. Recent work has also revealed a proclivity for IDR regions to self-associate forming biomolecular condensates through the process of liquid-liquid phase separation7. Many of the aforementioned functions involving ID are also now thought to involve some aspect of condensate formation8. Despite the importance of ID for biomolecular complex assembly, stabilization, scaffolding, and signal transduction, the atomic details of their specific interactions are difficult to identify since IDPs and IDRs are typically not amenable to structural investigations using x-ray crystallography or cryogenic electron microscopy.

Nuclear magnetic resonance (NMR) is an ideal technique for investigating ID as it is not dependent on the presence of rigid or homogenous structural ensembles but reports on the immediate local environment of individual nuclei. The resonant frequency, or chemical shift, of a nucleus in a given molecule is influenced by weak magnetic fields induced by the local electronic distribution, which in turn is dependent on bond lengths, angles, nearness of other nuclei, interactions with binding partners, and other factors9. Thus, each nucleus acts as a unique, site-specific structural probe sensitive to changes in its local chemical environment. Despite these advantages, NMR is a bulk technique, and the observed chemical shift is the average of all the environments sampled by a particular nucleus. A range of NMR techniques, many of which are described in this issue, have been developed to recover structural, dynamic, and kinetic information about high energy, lowly-populated biomolecular conformations contained in the averaged chemical shift10,11. Although transiently populated, identification and quantification of these states are important for determining the details of functional mechanisms12. For example, in the case of IDPs and IDRs, the conformational ensemble may be biased to preferentially sample conformations that are productive for the formation of encounter complexes with physiological binding partners. Detection of these states, as well as identification of the residue-specific inter- and intra-molecular interactions and dynamics, are important to determine the underlying structural mechanisms of protein function and complex formation.

A protocol for using paramagnetic relaxation enhancement (PRE) NMR to investigate transient, lowly-populated states important for the formation of IDP/IDR-mediated biomolecular complexes is described13. This approach is useful for studying the transient protein-protein interactions such as those that promote the assembly of amyloid fibrils from α-synuclein14,15 or the self-association of FUS16, as well as to characterize specific protein-protein interactions such as between signaling proteins17. An example of a self-associating IDP is presented, where specific inter- and intra-molecular interactions result in preferentially compacted states as well as site-specific interactions that drive self-association.

The PRE arises from the magnetic dipolar interaction of a nucleus to a paramagnetic center with an isotropic g-tensor, commonly supplied in the form of an unpaired electron on a nitroxide group or as a paramagnetic metal atom18 (Figure 1). While atoms with anisotropic g-tensors also produce a PRE effect, analysis of these systems is more difficult due to confounding effects contributed by the pseudo-contact shifts (PCS) or residual dipolar coupling (RDC)13,19. The strength of the interaction between a nucleus and the paramagnetic center is dependent on the <r-6> distance between the two. This interaction results in an increase in nuclear relaxation rates, which causes detectable line broadening even for long-range interactions (~10-35 Å), because the magnetic moment of the unpaired electron is so strong20,21. Detection of transient states with the PRE is possible if the following two conditions are met; (1) the transient interaction is in fast exchange on the NMR timescale (observed chemical shift is a population-weighted average of the exchanging states); and (2) the nuclei to paramagnetic center distance is shorter in the transiently populated state than in the major state11. The transverse PRE is denoted Γ2 and, for practical purposes, is calculated from the difference in 1H transverse relaxation rates between a sample containing a paramagnetic center and a diamagnetic control. For an in-depth treatment of the theory of the PRE and related pseudocontact shifts in fast and slow exchange regimes, the reader is referred to the comprehensive reviews by Clore and co-workers13,22. Here, only the situation where 1HNΓ2 is in the fast exchange regime is considered, where because of the r-6-dependence of the PRE, the observed relaxation rate is related to both the distance to which the paramagnetic center approaches the nucleus as well as the amount of time it spends in that conformation. Therefore, transient conformations that do not involve a close approach produce a small PRE while closer interactions, even if short-lived, will produce a larger PRE.

For IDPs, the PRE is used to measure and differentiate the interactions occurring within a single molecule (intramolecular) and between separate molecules (intermolecular). By attaching a paramagnetic center to an NMR visible (e.g., 15N-labeled) or NMR invisible (e.g., natural abundance 14N) protein, the source (inter- or intra-molecular) of the PRE may be determined (Figure 2). Site-directed mutagenesis that introduces a cysteine residue is a convenient approach to attach a paramagnetic center (spin-label) to a protein23. Several types of molecules have been proposed for use as spin labels, including metal chelating (EDTA-based) and free-radical (nitroxide-based)24. Various nitroxide spin labels have been described and are available with different cysteine-reactive chemistries such as methanethiosulphonate, maleimide, and iodoacetamide25,26 (Figure 1). Inherent flexibility of the tag or of the linker may be problematic for certain analyses, and in these situations, different strategies have been proposed to limit the motion of the tag, such as by adding bulky chemical groups or the use of a second linker to anchor the tag to the protein (two site attachment)27,28. Additionally, commercially available tags may contain diastereomeric proteins but generally this will not contribute to the observed PRE29. The use of the 3-Maleimido-PROXYL attached to a free cysteine via maleimide chemistry is described since it is readily available, cost-effective, non-reversible, and the reducing agent tris(2-carboxyethyl)phosphine (TCEP) can be maintained in the solution throughout the labeling reaction. Since 3-Maleimido-PROXYL has an isotropic g-tensor, no PCS or RDCs are induced, and the same chemical shift assignments can be used for both the paramagnetic and diamagnetic samples13.

The 1HNT2 is measured using a two time-point strategy (Ta, Tb) that has previously been shown to be as accurate as collecting a full evolution series consisting of 8 to 12 time points30. The first time point (Ta) is set as close to zero as practical, and the optimal length of the second time point is dependent on the magnitude of the largest expected PRE for a given sample and can be estimated from: Tb ~ 1.15/(R2,dia + Γ2) where R2,dia represents the R2 of the diamagnetic sample13. If the magnitude of the largest PREs is unknown, setting Tb to ~ one times the 1H T2 of the protein is a good initial estimate and further optimized by adjusting T2 to improve the signal to noise. This two-point measurement strategy significantly reduces the experimental time required to measure PREs and allows time for more signal averaging, particularly since relatively dilute samples are used to minimize the effects of non-specific contacts between molecules. An HSQC-based pulse sequence is used to measure 1HNT2 and has been described in detail elsewhere30. For improved sensitivity, the hard pulses of the forward and back INEPT transfers may be replaced with shaped pulses; alternatively, the sequence is readily converted to a TROSY-based readout31. Since IDPs typically have much longer transverse relaxation rates resulting in narrower line widths (due to the inherent disorder) than similarly sized globular proteins, long acquisition times in the indirect dimension may be used to improve spectral resolution and alleviate the chemical shift dispersion limitation inherent for IDPs.

PRE is a useful tool for studying protein-protein and protein-nucleic acid interactions, particularly interactions that are transient or lowly populated. A detailed protocol for the preparation of an NMR sample suitable for measuring PREs, including steps for protein purification, site-directed spin labeling, setting up and calibrating the pulse program, processing, and interpreting the NMR data, is provided. Important experimental considerations are noted throughout that may impact data quality and experimental outcome, including sample concentration, selection of the spin-label, and removal of paramagnetic components.

Protocol

General requirements for the protocol: protein purification facilities, UV-Vis spectrometer, high-field NMR spectrometer and operating software, post-processing analysis software including; NMRPipe32, Sparky33, (or CCPN Analysis34, or NMRViewJ35).

1. Recombinant expression and purification of a protein for PRE measurements

  1. Design an expression construct for the protein of interest so that there is a single cysteine residue present. Multiple mutations will be required to introduce a free cysteine at different positions in the protein of interest36.
  2. Express and purify a natural abundance (14N) or 15N-labeled sample of the protein of interest using an established protocol37.
    NOTE: E. coli expression systems provide cost effective and robust method for recombinant protein expression since isotopic enrichment of 15N is a minimal requirement for biomolecular heteronuclear NMR spectroscopy. Typical steps are expression in minimal media, chromatographic purification, and removal of affinity purification tag. This protocol assumes a robust expression and purification protocol has been established that can produce sufficient protein of suitable quality for NMR investigations.
    1. Maintain 1 mM reducing agent (DTT or TCEP) in buffers at all purification steps to prevent reaction of the free cysteine and formation of intermolecular disulfide bonds for IDPs.
      NOTE: Some systems may be more tolerant and less aggregation prone to non-reducing conditions depending on the specific characteristics of the protein, as well as the temperature, pH, and buffer system chosen for purification38.
    2. Remove affinity tags used for purification before proceeding since they may non-specifically interact with the protein in unpredictable ways or possibly contain reactive cysteine residues that could inadvertently serve as an unintended attachment site.
    3. Prepare a 15N labeled reference sample without cysteine mutation(s) mixed with a soluble version of the spin-label to assess the contribution of solvent PREs.

2. Conjugating the 3-Maleimido-PROXYL nitroxide spin label

  1. Store or exchange the purified protein into a degassed buffer containing 50 mM Tris pH 7 and 1 mM TCEP; the buffer may also contain up to 8 M urea if needed to aid protein solubility.
    ​Alternatively, rapidly dilute a protein stock solution into at least 10 volume equivalents of degassed 50 mM Tris pH 7 and 1 mM TCEP buffer. Ensure that the protein concentration prior to adding spin-label is at least 100 µM.
  2. Add 3-Maleimido proxyl from a stock solution to 20x molar excess of the protein of interest. Protect the sample from light and oxygen and incubate overnight at room temperature or 4 °C; gentle rocking or nutation may improve labeling efficiency.
  3. Prepare stock solutions of the spin-label by dissolving 3-Maleimido proxyl powder in 95% ethanol. Aliquots of the stock can be stored at -80 °C for less than 6 months.
  4. Critical step: Remove the non-reacted free spin-label to prevent non-specific solvent PREs. Achieve this by gel filtration or (preferably) extensive dialysis of the protein sample. This step will also introduce the protein into a buffer suitable for NMR.
    NOTE: Reducing agents should be prepared fresh, and compatibility between buffer components should be considered; for example, TCEP degrades quickly in phosphate-based buffers, and this combination should be avoided39.
  5. Treat all buffers used from this step forward with a chelating resin selective for divalent and transition metals to remove paramagnetic ions or spin-label quenchers. If the protein cannot be stored in an NMR buffer, concentrate the protein to be rapidly diluted into a buffer suitable for NMR.
  6. Monitoring the efficiency of spin-label incorporation.
    1. Use Ellman's reagent (5,5-dithio-bis-(2-nitrobenzoic acid) for quantifying free sulfhydryl groups in solution40.
      NOTE: Detailed protocols are available from the manufacturer. For the purposes here, it is important to determine incorporation of the spin-label, the concentration of free sulfhydryl groups is compared with the total protein concentration. The percent of free sulfhydryl groups is the percent of molecules that do not have a nitroxide spin label attached.
    2. Monitor the intensity of the peak corresponding to the tagged cysteine residue to judge spin-label incorporation into the protein of interest.
      ​NOTE: This is a rapid and effective approach to determine the degree of spin labeling of the protein. Complete incorporation of the spin-label will result in the disappearance of the peak from the spectrum. With the poor dispersion characteristic of IDPs the peak corresponding to the mutant cysteine residue may not always be readily identified, and thus the use of Ellman's reagent (step 2.6.1) is recommended.

3. Prepare NMR sample for measuring intra- or inter-molecular PRE

  1. Prepare sample for measurement of intramolecular PRE
    1. Prepare 15N isotopically enriched, spin-labeled protein to a concentration of at least 100 µM but not more than 300 µM in a buffer suitable for NMR. Total sample volume (including D2O) is 500 – 550 µL.
      NOTE: Common NMR buffers include phosphate, acetate, (bi)carbonate, and TRIS. Good's buffers such as MES, HEPES may also be appropriate. Exercise caution when selecting buffers to ensure no cross-reactivity with other solution components.
    2. Ensure that the pH is ~7.2 or lower to minimize the effects of amide proton exchange with water. Keep the concentration of salt as low as possible (typically less than 150 mM), although the primary consideration is to maintain protein stability.
      NOTE: Approaches for conducting NMR experiments in high-salt conditions have been described elsewhere41.
  2. Prepare sample for measurement of intermolecular PRE
    1. Follow this step or step 3.1; they are not performed simultaneously. Prepare 14N natural abundance, spin-labeled protein in the chosen NMR buffer.
    2. Prepare the protein sample by mixing 15N isotopically enriched non-spin-labeled protein with 1%-50% 14N natural abundance spin-labeled protein so that the final concentration is identical to the sample prepared in 3.1.1. The total sample volume (including D2O) is 500 – 550 µL.
    3. Empirically optimize the ratio of the 15N and 14N proteins for each protein studied. The ratios of 1%, 5%, and 20% of 14N-spin-labeled protein are good starting points.
      NOTE: A buildup of the PRE as a function of added 14N-spin-labeled protein indicates a specific effect; the observed PRE is sample-specific since it depends on distance and population (as discussed above), and therefore higher ratios of 14N-spin-labeled protein will be required if the interaction is particularly transient17.
  3. Transfer the NMR (either intra- or inter-molecular) sample to a 5 mm NMR tube that is appropriate for use in high-field magnets using a long-stem (9") glass pipette or micropipette. Ensure that all NMR samples include 5%-10% of D2O to facilitate field locking.
    ​NOTE: NMR tubes that utilize polymer plugs to reduce the necessary sample volume are not recommended for PRE measurements due to difficulties related to effective sample shimming.

4. Set up NMR spectrometer and experiment specific parameters

  1. Exercise extreme caution when working around superconducting, high-field NMR spectrometers.
    NOTE: Hazards include injuries due to the sudden acceleration of metallic objects toward the magnet, interference with implanted medical devices, and asphyxiation due to a sudden release of N2 and He2 gas in the event of a magnet quench. The following steps assume that the reader has undergone the required training, is aware of these and other local hazards, and has received approval from the facility manager to operate the NMR spectrometer. When in doubt of a step or instruction, consult with the facility manager or experienced user to prevent potential personal injury or damage to the spectrometer.
  2. The following steps assume a commercial NMR spectrometer running a modern version of the acquisition control software. Download the pulse program and parameter files and place them in the appropriate directories.
    NOTE: A pulse program and parameter set suitable for use with a Bruker spectrometer and TopSpin (3.2 or later) are available upon request from the authors.
    1. Critical step: Familiarity with installing non-native NMR pulse programs is assumed; consult with the facility manager or an experienced user if necessary.
  3. Place the sample in the magnet, lock on the 2H signal using the Lock command, tune and match the 1H channel according to facility protocols (the exact procedure will depend on if the probe is equipped with a remote tune and match module).
  4. Adjust the shims using the topshim subroutine to optimize solvent signal suppression.
  5. Calibrate the 1H and 15N 90° pulses using standard methods.
    1. Calibrate the 1H pulse using the popt program (use pulsecal first to estimate pulse length).
    2. Calibrate the 15N pulse against a standard sample; make sure this value has been calibrated recently by discussing with a technical director or experienced user.
    3. Alternatively, calibrate the 15N pulse on the sample by varying one of the 90° pulses of an HMQC experiment until a null signal is achieved.
    4. Determine the correct attenuation for shaped pulses using the shape tool (stdisp) subroutine.
    5. Open the appropriate pulse shape file by clicking on the folder icon. The shaped pulses are found in the pulse parameters section of ACQUPARS.
    6. Load the pulse definition file and click on Analyze Waveform > Integrate Shape. Input the calibrated 1H 90° hard pulse, desired shaped pulse length, and rotation (90° or 180°).
    7. Calculate the power level of the shaped pulse by adding the change of power level to the attenuation for the calibrated 90° pulse.
  6. Record a standard 1H, 15N HSQC (hsqcetfpf3gpsi) to optimize sweep width, carrier frequency and check water suppression25.
  7. Adjust the sweep width and the number of indirect dimension increments using the sw and td commands or directly in the appropriate dialog boxes. Typically, for collecting PREs, spectral widths are chosen so that the spectrum is not folded.

5. Setup the 1HNT2 experiment

  1. Calibrate the shaped pulses as described above (4.4.5-4.5.7). The shaped pulse parameter files for the PRE experiment are Eburp2.1000 (90° pulse), Reburp.1000, and Iburp2.1000. Enter the calibrated pulse lengths in the pulse parameters section on the ACQUPARS tab.
  2. This experiment measures the 1HNT2 using the two time-delay point approach30.
    1. Set the time delays by editing the vdlist file, the first delay (Ta) is set to 0.01 ms.
    2. Choose the second delay, (Tb) using the relationship to the expected maximum PRE (Tb ~ 1.15/(R2,dia + Γ2) where R2,dia represents the R2 of the diamagnetic sample13. Without prior knowledge of the magnitude of the PRE contribution to the observed relaxation, a good starting point is setting Tb to ~1x 1H T2.
    3. Then determine a suitable value by comparing the first increments (processed with the efp command) of the Ta and Tb spectra and adjusting Tb such that the signal decays to between 40%-50% of its initial value.
      NOTE: This approach optimizes the spectral signal-to-noise, a necessary consideration for samples that cannot be highly concentrated (< 50 µM). Suitable values of Tb are sample dependent but typically range from 8 – 40 ms for an average sized protein.
  3. Determine the number of complex points to record and number of scans for sufficient signal averaging. Since IDPs have longer 15N T2 than folded proteins of comparable size, longer acquisition times in the indirect dimension may be used.
    NOTE: This value is dependent on the specific characteristics of the protein but can be roughly estimated from the 15N T2 and optimized by monitoring signal decay in the FID. For the direct dimension, 1024* complex points (13 ppm sweep width, 112.6 ms acquisition time) is sufficient for most samples.
  4. Use the command expt to calculate the experiment time and then start the experiment with the command zg.

6. Make a diamagnetic sample by reducing spin-label with ascorbic acid

  1. Dissolve sodium ascorbate in the NMR buffer and adjust the pH to match the original NMR buffer.
  2. Calculate the concentration of sodium ascorbate stock so that a 10x molar excess of ascorbate over the concentration of the spin-label can be added with the least change of sample volume. For example, for a 100 µM protein sample, a 100 mM stock of ascorbate is appropriate. Reducing the spin-label will require adding 5.5 µL of ascorbic acid stock solution, which is only 1 % of the total sample volume.
  3. Add the required amount of ascorbic acid to the NMR tube by placing a droplet below the rim of the tube, cap the tube, carefully invert the tube to mix, and then spin at 200-400 x g for 10-20 s in a hand-cranked centrifuge to settle the sample at the bottom of the tube.
  4. Wrap the NMR tube in foil to protect from light and allow the reaction to proceed for at least 3 h.
  5. Record 1HNT2 on the diamagnetic sample using the same parameters used for the paramagnetic sample.
  6. Recalibrate the pulses. However, they should not have changed from the paramagnetic measurements; if they are significantly different (> 0.5 µs difference), consider sample quality (e.g., degradation, precipitation).
  7. Ensure that all acquisition parameters, including the specified relaxation delays (vdlist), number of dummy scans, number of scans collected, number of complex points collected, acquisition time, sweep widths, and carrier frequencies remain the same for the diamagnetic and paramagnetic samples.

7. Process paramagnetic and diamagnetic spectra

  1. Copy the data to local computer or workstation that has NMRPipe and Sparky installed and configured. Make a folder named proc in the experiment data directory that contains the ser file.
  2. Copy the NMRPipe scripts fid.com, p3d.com, and nmrproc.com into proc (processing scripts are available upon request from the authors).
    1. Use the fid.com script to convert the Bruker data format (ser) into NMRPipe format.
    2. Use the p3D.com script to split the pseudo3D planes into individual spectra.
    3. Use the nmrproc.com script to read the output of the fid.com script, apply solvent suppression, a window function, append zeros to the raw data (zero fill), apply phase correction, execute a Fourier transformation, trim the data for display and write the processed data to disk. The script will output one file for each relaxation delay recorded (Ta and Tb).
      ​NOTE: Each of these scripts is customizable to optimize the processing for the specific details of each experiment. Tutorials and example data sets are included in the NMRPipe distribution available from the NMRPipe website32. NMRDraw may be used for spectral viewing during processing (e.g., setting proper phase angles etc.). Options available for NMRPipe commands can be viewed using the command nmrPipe -help.

8. Transfer resonance assignments and extract peak heights

  1. Change the file header information for each spectrum file (Ta, Tb for both paramagnetic and diamagnetic samples) using the command sethdr [filename] -ndim 2.
  2. Use Sparky to extract peak heights33 following steps 8.3-8.5. Other software packages, including NMRPipe (NMRDraw)32, CCPN Analysis34, and NMRViewJ35 are also appropriate.
  3. Read the spectral files into Sparky. At this step the data set will consist of one spectrum for each time point spectra (Ta, Tb), for both the paramagnetic and diamagnetic samples, measured for each position of the spin-label in the protein.
  4. Use Sparky to pick peaks (command: F8, then click and drag) and transfer assignments using the transfer peak list tool from a reference peaklist.
    NOTE: Resonance assignments of the protein of interest are necessary for sequence-specific interpretation of the observed PREs36.
    1. Set contours in both paramagnetic and diamagnetic spectra to the same level. Ensure to set the contours so that the spectra collected after the time delay do not purposely exclude peaks but are high enough so that the Ta spectra are not overly noisy.
  5. Save the new peak lists for each spectrum and include the measured peak intensity and Sparky calculated signal to noise ratio (command: lt to open peaklist, click options to include intensity and SNR columns, command: save).

9. Extract 1HNT2 rates for each residue and calculate PRE

  1. Import the peak lists into spreadsheet software or a preferred programming language such as Python.
    NOTE: For each spin-label position on the protein, the dataset will consist of four peak lists with associated peak intensities, one each of Ta and Tb for both the paramagnetic and diamagnetic experiments.
  2. Calculate 1HN R2 for both the paramagnetic and diamagnetic samples using the equation:
    Equation 1
    Equation 2
  3. Use the above equation to determine the relaxation rate for each residue for the paramagnetic and diamagnetic samples.
  4. Determine the 1HNΓ2 rate for each residue using the equation:
    Equation 3
  5. Use the Sparky calculated signal to noise ratio (SNR) to compute the uncertainty of the peak height for each residue.
  6. Propagate the error using the equation:
    Equation 4
  7. Plot 1HNΓ2 as a function of residue number using a scatter plot including the error calculated in 9.6.

Representative Results

Intramolecular 1HNΓ2 PREs were recorded on a self-associating, intrinsically disordered fragment (residues 171-264) derived from the low-complexity domain of the RNA-binding protein EWSR142 (Figure 3). Residues in close sequential proximity to the spin-label attachment point (e.g., residue 178 or 260 in Figure 3) are expected to be significantly broadened and are not detectable in the spectrum. Residues sequentially spaced from the attachment point yet show enhanced Γ2 were spatially close (10-35 Å) to the spin-label. In the case of EWSR1 171-264, attributing the source of the PRE effect is complicated since it may arise from a combination of inter- and intra- residue contacts and is dependent on the distance from the nucleus to the paramagnetic center, the population of that conformation, and the dynamics of the vector connecting the electron and nuclear spins. Further, the magnitude of PREs arising from intramolecular contacts is not concentration-dependent, while PREs arising from intermolecular contacts depend on concentration as well as the kinetics and dynamics of the association between protein molecules.

A possible interpretation of these data is that the IDP ensemble samples conformations that are more compact than an extended chain. Alternatively, the PREs could arise from intermolecular contacts responsible for the self-association of EWSR1, or the PREs could be from a combination of both intra- and intermolecular contacts. In the case presented here, what remains unknown is how close the residues approach the spin-label or for how long they remain in close proximity. With highly flexible molecules such as EWSR1 171-264, it can be difficult to qualitatively disentangle these parameters. By placing the spin-label at different residue positions, contacts between different parts of the chain may be identified, providing a more accurate interpretation of specific interactions that may be functionally relevant for self-association (Figure 3). Measuring intermolecular PREs (14N spin-labeled protein mixed with 15N non-spin labeled protein), employing a mutational strategy of residues with larger than average PREs (e.g., residues 196 or 215, Figure 3), and utilizing other biophysical methods such as dynamic light scattering, size exclusion chromatography, and analytical ultracentrifugation, are useful for characterizing the conformational ensemble of an IDP.

Figure 1
Figure 1: Molecules containing an unpaired electron and various functional groups to facilitate attachment to free cysteine residues that are typically used as paramagnetic relaxation agents. Diamagnetic molecules may be used as controls. (A) 3-Maleimido-2,2,5,5-tetramethyl-1-pyrrolidinyloxy, free radical (3-Maleimido-PROXYL) (B) 3-Carboxy-2,2,5,5-tetramethyl-1-pyrrolidinyloxy, free radical (3-Carboxy-PROXYL) (C) 3-(2-Iodoacetamido)-2,2,5,5-tetramethyl-1-pyrrolidinyloxy, free radical (3-(2 – Iodoacetamido-PROXYL) (D) 1-Oxyl-2,2,5,5-tetramethylpyrrolidin-3-yl) Methyl Methanethiosulfonate (MTSL) (E) (1-Acetoxy-2,2,5,5-tetramethyl-δ-3-pyrroline-3-methyl) Methanethiosulfonate (Acetoxy-MTSL) Please click here to view a larger version of this figure.

Figure 2
Figure 2: Depiction of intra- and intermolecular PRE. (A) Intramolecular PRE, the red circle represents the effective radius of a paramagnetic center attached to a 15N-labeled protein. The PRE effect decreases with an <r-6> dependence on distance from the paramagnetic molecule. (B) Intermolecular PRE, the paramagnetic group (red circle), is located on a 14N (natural abundance) protein (blue) that is invisible to NMR. The effects of the paramagnetic group on the non-NMR active protein are observed as increased relaxation rates when it comes into close contact with the 15N protein (black). Please click here to view a larger version of this figure.

Figure 3
Figure 3: 1HNΓ2 rates for residues 171-264 of the intrinsically disordered domain of EWSR1. A serine residue at position (A) 178 or (B) 260 that has been mutated to a cysteine serves as the attachment point for a 3-Maleimido-PROXYL spin-label (red *). Increased relaxation rates occur at the location of the tag, other sites of increased relaxation are indicative of intramolecular interactions. Please click here to view a larger version of this figure.

Discussion

A method for characterizing transient interactions that exist at low populations between intrinsically disordered proteins and various binding partners using PRE has been presented. In the example shown, the protein is self-associating, and thus the PRE may arise from a combination of inter and intramolecular interactions. This method is readily extended to heterogeneous samples where the interactions between two different proteins may be characterized. Complementary information about how different regions of the protein interact is available by placing the spin-label at different positions within the protein. Additionally, by alternating the spin-label between NMR active (15N) and NMR inactive (14N) species, the intra- and inter-molecular sources of observed PRE may be differentiated from one another, providing information about encounter complexes. The experiment outlined here can report on encounter complex interactions even if they occur on a microsecond timescale13.

Central to this method is the incorporation of a spin-label tag into the protein of interest by attachment to a cysteine residue. Some proteins may contain a native cysteine that is suitable (does not participate in disulfide bonds, is surface exposed) for attaching a spin-label. For IDPs, solvent exposure of cysteine is usually not an issue. In the majority of cases, it is desirable to introduce cysteines as conservative mutations (serine to cysteine or other uncharged polar amino acids to cysteine) using site-directed mutagenesis43. In the example presented, the fragment of EWSR1 does not contain native cysteines and is enriched in serines; thus, devising a mutational strategy was straightforward. Proteins that contain native cysteine(s) present a more complicated case, and care needs to be taken to not disrupt the native function (e.g., break a structurally important disulfide bond)44. Further, to incorporate a single cysteine for spin-labeling, the native cysteines must be mutated to a residue that does not react with the spin-label (no mercapto group) and based on its size and other properties, serine is a good substitute for cysteine. If native cysteines need to be mutated, careful characterization of the mutants is required to ensure they maintain native structure and function is essential. Simple 1H,15N HSQCs of mutants in comparison to the wildtype protein are powerful indicators of perturbations (even minor) to protein structure, and this approach is also useful for IDPs45. Other methods to consider are circular dichroism, analytical ultracentrifugation, or biochemical approaches such as activity assays46.

Technical considerations for obtaining reproducible, rigorous, high-quality data include the removal of ionic impurities during the preparation of the NMR sample. This is achieved by passing all solutions over chelating resin prior to use. Further, using a properly degassed buffer is important during the attachment of the nitroxide spin label as the presence of oxygen can reduce the efficiency of labeling. Diamagnetic contamination will contribute to a decrease in the observed PRE; however, the effect is less pronounced on intramolecular PREs and can be reduced by decreasing ΔT13. Therefore, it is not necessary to obtain 100% label incorporation to proceed with the experiment, particularly for the qualitative interpretation presented here. If free cysteines from incomplete spin-label attachment are problematic, some mercapto-reactive chemistries (e.g., maleimide) are amenable to maintaining a reducing agent in the sample throughout the experiment26. It is important that the paramagnetic and diamagnetic samples match each other as closely as possible. When reducing the spin-label with ascorbic acid to create a diamagnetic control, consider the dilution factor introduced from titrating in an ascorbic acid stock solution. This dilution can be minimized by maintaining the ascorbic acid stock at least 10x the expected working concentration in the NMR buffer.

There are many software packages available for analyzing NMR data, including NMRPipe32, Sparky33, CCPN Analysis34, NMRViewJ35, among others. The combination of NMRPipe for spectral processing and Sparky for spectral analysis (peak picking and quantification) was described here due to the ease of use of this combination. NMRPipe is commonly used by many NMR groups for spectral processing, but the NMRPipe suite contains the tools necessary for completing all steps of the analysis, albeit with a significant learning curve. Data may also be processed using the NMR spectrometer control software. Sparky was chosen for spectral analysis because of its ease of use and rapid uptake by novice users. There are several options available for spectral analysis (peak picking and measuring peak heights) that can easily substitute for the functionality of Sparky including CCPN Analysis, or NMRViewJ. Notably, many of these programs have overlapping functionalities and the user is advised to select the combination of programs with which they are most comfortable.

Poor chemical shift dispersion is an inherent problem with IDPs leading to significant resonance overlap and introduction of error into the measurement of peak height. Different strategies have been proposed to alleviate this problem. One of the most straightforward, and the one employed here, is to take advantage of the long transverse relaxation characteristic of IDPs and simply extend the acquisition time in the 15N (indirect) dimension. Alternatively, the triple resonance HNCO experiment is useful for resolving resonance overlap in IDPs due to the superior dispersion of C' resonances. Both TROSY and HSQC versions of the HNCO for measuring PREs have been proposed and are described elsewhere47. However, the improved resolution is not always significant enough to warrant the increased complexity of the experiment, longer time for data collection, and added cost for preparing a suitable sample (enrichment of 13C). This is indeed the case for EWSR1 171-264 presented here, where no significant improvement in the number of non-overlapped residues was observed between a TROSY-HNCO and an 1H, 15N HSQC collected with long acquisition time in the indirect dimension.

This procedure outlined above focuses on the utility of PRE experiments for characterizing weak interactions that exist within and between intrinsically disordered proteins. The PRE has a much broader utility in biomolecular NMR, including determining long-range structural restraints and quantitative determination of sparsely populated conformational states. For example, Clore and co-workers have pioneered the use of the PRE to detect and quantify transient interactions arising from interactions between individual domains of a single protein48 or between the subunits of assembled protein complexes17. There are many examples of the PRE used to derive long-range distance restraints, including for large proteins49, or with novel PRE tags50, to help determine the overall fold of a protein51, as well as in highly paramagnetic systems52. Finally, while PCS is beyond the scope of this discussion, they have been applied to important biomolecular problems which have been described elsewhere53. The method presented above is suitable for probing the conformation and interactions of IDPs using PREs and was designed to be accessible for novice users. For more quantitative approaches to the analysis of the PRE, the user is referred to the many excellent articles referenced within11,24,30,31.

Disclosures

The authors have nothing to disclose.

Acknowledgements

We thank Drs. Jinfa Ying and Kristin Cano for helpful discussions and technical assistance. DSL is a St. Baldrick's Scholar and acknowledges the support of the St. Baldrick's Foundation (634706). This work was supported in part by the Welch Foundation (AQ-2001-20190330) to DSL, the Max and Minnie Tomerlin Voelcker Fund (Voelcker Foundation Young Investigator Grant to DSL), UTHSA Start-Up Funds to DSL, and a Greehey Graduate Fellowship in Children's Health to CNJ. This work is based upon research conducted in the Structural Biology Core Facilities, part of the Institutional Research Cores at the University of Texas Health Science Center at San Antonio supported by the Office of the Vice President for Research and the Mays Cancer Center Drug Discovery and Structural Biology Shared Resource (NIH P30 CA054174).

Materials

0.45 µm and 0.22 µm syringe filters Millipore Sigma SLHVM33RS
SLGVR33RS
Filter lysate before first purification step and before size exclusion chromatography.
100 mm Petri Dish Fisher FB0875713 Agar plates for bacterial transformation.
14N Ammonium chloride Sigma Aldrich 576794 Use of 15N in M9 medium will produce an NMR visible protein, 14N will produces an NMR invisible protein
15N Ammonium chloride Sigma Aldrich 299251 Use of 15N in M9 medium will produce an NMR visible protein, 14N will produces an NMR invisible protein
3 L Fernbach baffled flask Corning 431523 Bacterial expression culture
3-Maleimido-Proxyl Sigma Aldrich 253375 Nitroxide spin label
50 mL conical centrifuge tubes Thermo Fisher 14-432-22 Solution/protein storage
Amicon centrifugal filter Millipore Sigma UFC900308 Protein concentration
Ampicillan Sigma Aldrich A5354 Antibiotic for a selective marker, exact choice depends on the expression construct plasmid
Analytical balance Oahus 30061978 Explorer Pro, for weighing reagents
Ascorbic acid Sigma Aldrich AX1775 Reduces nitroxide spin label
Autoclave Sterilize glassware and culture media
Calcium chloride Sigma Aldrich C4901 M9 media component
Centrifuge bottles Thermo Fisher 010-1459 Harvest E. coli cells after recombinant protein expression
Centrifuge, hand-crank Thomas Scientific 0241C68 Boekel hand-driven, low-speed centrifuge with 15 mL buckets that can accommodate NMR tubes
Chelex 100 Sigma Aldrich C7901 Remove contaminating paramagnetic compounds from buffer solutions
Computer workstation Linux or Mac OS compatable with NMR data processing and analysis software packages such as NMRPipe and Sparky
Deuterium oxide Sigma Aldrich 151882 Needed for NMR lock signal
Dextrose Sigma Aldrich D9434 M9 media component
Dibasic Sodium Phosphate Sigma Aldrich S5136 M9 media component
Ellman's reagent (5,5-dithio-bis-(2-nitrobenzoic acid) Thermo Fisher 22582 Quantification of free cystiene residues
High speed centrifuge tubes Thermo Fisher 3114-0050 Used to clear bacterial lysate.
High-field NMR instrument (600 – 800 MHz) Bruker Equiped with a multichannel cryogenic probe and temperature control
IMAC column, HisTrap FF Cytvia 17528601 Initial fractionation of crude bacterial lysate
Isopropyl B-D-thiogalactoside (IPTG) Sigma Aldrich I6758 Induces protein expression for genes under control of lac operator
LB agar Thermo Fisher 22700025 Items are used for transforming E. coli to express protein of interest, substitions for any of these items with like products is acceptable.
LB broth Thermo Fisher 12780052
Low-pressure chromatography system Bio-Rad 7318300 BioRad BioLogic is used for low-pressure chomatograph such as running IMAC columns
Magnesium sulfate Sigma Aldrich M7506 M9 media component
Medium pressure chromatography system Bio-Rad 7880007 BioRad NGC equipped with a multi-wavelength detector, pH and conductivity monitors, and automatic fraction collector
MEM vitamin solution Sigma Aldrich M6895 M9 media component
Microfluidizer Avestin EmulsiFlex-C3 Provides rapid and efficient bacterial cell lysis
Micropipettes Thermo Fisher Calibrated set of micropippetters with properly fitting disposable tips (available from multiple manufacturers e.g. Eppendorf)
Monobasic potassium phosphate Sigma Aldrich 1551139 M9 media component
NMR pipettes Sigma Aldrich 255688 To remove sample from NMR tube
NMR sample tube NewEra NE-SL5 Suitable for high-field NMR spectrometers
Preparative Centrifuge Beckman Coulter Avanti J-HC Harvest E. coli cells after recombinant protein expression
Round bottom polystyrene centrifuge tubes Corning 352057 Clear bacterial lysate
Shaking incubator Eppendorf S44I200005 Temperature controlled growth of E. coli starter and expression cultures
Sodium chloride Sigma Aldrich S5886 M9 media component
Sonicating water bath and vacuum source Thomas Scientific Used to degas buffer solutions
Sonicator Thermo Fisher FB505110 Used for bacterial cell lysis or shearing bacterial DNA
Spectrophotometer Implen OD600 Diluphotometer Monitor growth of E.coli protein expression cultures
Superdex 200 16/600 size exculsion colum Cytvia 28989333 Final protein purification step
Topspin software, version 3.2 or later Bruker Operating software for the NMR instrument
Transformation competent E. coli cells Thermo Fisher C600003 One Shot BL21 Star (DE3) chemically competent E. coli, other strains may be compatable
Tris(2-carboxyethyl)phosphine (TCEP) ThermoFisher 20490 Reducing agent compatable with some sulfhydryl-reactive conjugations
UV-Vis spectrophotometer Implen NP80 Measure protein concentration.
Water bath, temperature controlled ThermoFisher FSGPD25 For heat shock step of bacterial transformation
Yeast extract Sigma Aldrich Y1625 For supplementing M9 media if required

References

  1. Dyson, H. J., Wright, P. E. Intrinsically unstructured proteins and their functions. Nature Reviews: Molecular Cell Biology. 6 (3), 197-208 (2005).
  2. Korneta, I., Bujnicki, J. M. Intrinsic disorder in the human spliceosomal proteome. PLoS Computational Biology. 8 (8), 1002641 (2012).
  3. Frege, T., Uversky, V. N. Intrinsically disordered proteins in the nucleus of human cells. Biochemistry and Biophysics Reports. 1, 33-51 (2015).
  4. Liu, J., et al. Intrinsic disorder in transcription factors. 생화학. 45 (22), 6873-6888 (2006).
  5. El Hadidy, N., Uversky, V. N. Intrinsic disorder of the BAF complex: Roles in chromatin remodeling and disease development. International Journal of Molecular Sciences. 20 (21), (2019).
  6. Wright, P. E., Dyson, H. J. Intrinsically disordered proteins in cellular signalling and regulation. Nature Reviews: Molecular Cell Biology. 16 (1), 18-29 (2015).
  7. Brangwynne, C. P. Phase transitions and size scaling of membrane-less organelles. Journal of Cell Biology. 203 (6), 875-881 (2013).
  8. Shin, Y., Brangwynne, C. P. Liquid phase condensation in cell physiology and disease. Science. 357 (6357), (2017).
  9. Cavanagh, J. . Protein NMR spectroscopy : principles and practice. 1st edition. , (2018).
  10. Sekhar, A., Kay, L. E. NMR paves the way for atomic level descriptions of sparsely populated, transiently formed biomolecular conformers. Proceedings of the National Academy of Sciences of the United States of America. 110 (32), 12867-12874 (2013).
  11. Anthis, N. J., Clore, G. M. Visualizing transient dark states by NMR spectroscopy. Quarterly Reviews of Biophysics. 48 (1), 35-116 (2015).
  12. Alderson, T. R., Kay, L. E. NMR spectroscopy captures the essential role of dynamics in regulating biomolecular function. Cell. 184 (3), 577-595 (2021).
  13. Clore, G. M., Iwahara, J. Theory, practice, and applications of paramagnetic relaxation enhancement for the characterization of transient low-population states of biological macromolecules and their complexes. Chemical Reviews. 109 (9), 4108-4139 (2009).
  14. Wu, K. P., Baum, J. Detection of transient interchain interactions in the intrinsically disordered protein alpha-synuclein by NMR paramagnetic relaxation enhancement. Journal of the American Chemical Society. 132 (16), 5546-5547 (2010).
  15. Janowska, M. K., Wu, K. P., Baum, J. Unveiling transient protein-protein interactions that modulate inhibition of alpha-synuclein aggregation by beta-synuclein, a pre-synaptic protein that co-localizes with alpha-synuclein. Scientific Reports. 5, 15164 (2015).
  16. Murthy, A. C., et al. Molecular interactions underlying liquid-liquid phase separation of the FUS low-complexity domain. Nature Structural & Molecular Biology. 26 (7), 637-648 (2019).
  17. Fawzi, N. L., Doucleff, M., Suh, J. Y., Clore, G. M. Mechanistic details of a protein-protein association pathway revealed by paramagnetic relaxation enhancement titration measurements. Proceedings of the National Academy of Sciences of the United States of America. 107 (4), 1379-1384 (2010).
  18. Griffith, O. H., Waggoner, A. S. Nitroxide free radicals: spin labels for probing biomolecular structure. Accounts of Chemical Research. 2 (2), 17-24 (1969).
  19. Bertini, I., Luchinat, C., Parigi, G., Ravera, E. . NMR of Paramagnetic Macromolecules, Applications to Metallobiomolecules and Models. 2 edn. , (2016).
  20. Bloembergen, N., Purcell, E. M., Pound, R. V. Relaxation effects in nuclear magnetic resonance absorption. Physical Review. 73 (7), 679-712 (1948).
  21. Solomon, I. Relaxation processes in a system of two spins. Physical Review. 99 (2), 559 (1955).
  22. Clore, G. M. Practical aspects of paramagnetic relaxation enhancement in biological macromolecules. Methods in Enzymology. 564, 485-497 (2015).
  23. Klare, J. P. Site-directed spin labeling EPR spectroscopy in protein research. Biological Chemistry. 394 (10), 1281-1300 (2013).
  24. Clore, G. M., Tang, C., Iwahara, J. Elucidating transient macromolecular interactions using paramagnetic relaxation enhancement. Current Opinion in Structural Biology. 17 (5), 603-616 (2007).
  25. Melanson, M., Sood, A., Torok, F., Torok, M. Introduction to spin label electron paramagnetic resonance spectroscopy of proteins. Biochemistry and Molecular Biology Education. 41 (3), 156-162 (2013).
  26. Czogalla, A., Pieciul, A., Jezierski, A., Sikorski, A. F. Attaching a spin to a protein — site-directed spin labeling in structural biology. Acta Biochimica Polonica. 54 (2), 235-244 (2007).
  27. Lindfors, H. E., de Koning, P. E., Drijfhout, J. W., Venezia, B., Ubbink, M. Mobility of TOAC spin-labelled peptides binding to the Src SH3 domain studied by paramagnetic NMR. Journal of Biomolecular NMR. 41 (3), 157-167 (2008).
  28. Fawzi, N. L., et al. A rigid disulfide-linked nitroxide side chain simplifies the quantitative analysis of PRE data. Journal of Biomolecular NMR. 51 (1-2), 105-114 (2011).
  29. Bleicken, S., et al. gem-Diethyl pyrroline nitroxide spin labels: Synthesis, EPR characterization, rotamer libraries and biocompatibility. ChemistryOpen. 8 (8), 1035 (2019).
  30. Iwahara, J., Tang, C., Clore, G. M. Practical aspects of 1H transverse paramagnetic relaxation enhancement measurements on macromolecules. Journal of Magnetic Resonance. 184, 185-195 (2007).
  31. Venditti, V., Fawzi, N. L. Probing the atomic structure of transient protein contacts by paramagnetic relaxation enhancement solution NMR. Methods in Molecular Biology. 1688, 243-255 (2018).
  32. Delaglio, F., et al. NMRPipe: a multidimensional spectral processing system based on UNIX pipes. Journal of Biomolecular NMR. 6 (3), 277-293 (1995).
  33. Lee, W., Tonelli, M., Markley, J. L. NMRFAM-SPARKY: enhanced software for biomolecular NMR spectroscopy. Bioinformatics. 31 (8), 1325-1327 (2015).
  34. Vranken, W. F., et al. The CCPN data model for NMR spectroscopy: development of a software pipeline. Proteins. 59 (4), 687-696 (2005).
  35. Johnson, B. A. Using NMRView to visualize and analyze the NMR spectra of macromolecules. Methods in Molecular Biology. 278, 313-352 (2004).
  36. Sjodt, M., Clubb, R. T. Nitroxide labeling of proteins and the determination of paramagnetic relaxation derived distance restraints for NMR studies. Bio-Protocol. 7 (7), (2017).
  37. Zhang, H., van Ingen, H. Isotope-labeling strategies for solution NMR studies of macromolecular assemblies. Current Opinion in Structural Biology. 38, 75-82 (2016).
  38. Rabdano, S. O., et al. Onset of disorder and protein aggregation due to oxidation-induced intermolecular disulfide bonds: case study of RRM2 domain from TDP-43. Scientific Reports. 7 (1), 11161 (2017).
  39. Burns, J. A., Butler, J. C., Moran, J., Whitesides, G. M. Selective reduction of disulfides by tris(2-carboxyethyl)phosphine. Journal of Organic Chemistry. 56 (8), 2648-2650 (1991).
  40. Ellman, G. L. Tissue sulfhydryl groups. Archives of Biochemistry and Biophysics. 82 (1), 70-77 (1959).
  41. Binbuga, B., Boroujerdi, A. F., Young, J. K. Structure in an extreme environment: NMR at high salt. Protein Science. 16 (8), 1783-1787 (2007).
  42. Schwartz, J. C., Cech, T. R., Parker, R. R. Biochemical properties and biological functions of FET proteins. Annual Review of Biochemistry. 84, 355-379 (2015).
  43. Nabuurs, S. M., de Kort, B. J., Westphal, A. H., van Mierlo, C. P. Non-native hydrophobic interactions detected in unfolded apoflavodoxin by paramagnetic relaxation enhancement. European Biophysics Journal. 39 (4), 689-698 (2010).
  44. Wiedemann, C., Kumar, A., Lang, A., Ohlenschlager, O. Cysteines and disulfide bonds as structure-forming units: Insights from different domains of life and the potential for characterization by NMR. Frontiers in Chemistry. 8, 280 (2020).
  45. Wommack, A. J., et al. NMR solution structure and condition-dependent oligomerization of the antimicrobial peptide human defensin 5. 생화학. 51 (48), 9624-9637 (2012).
  46. Taylor, A. M., et al. Detailed characterization of cysteine-less P-glycoprotein reveals subtle pharmacological differences in function from wild-type protein. British Journal of Pharmacology. 134 (8), 1609-1618 (2001).
  47. Hu, K., Doucleff, M., Clore, G. M. Using multiple quantum coherence to increase the 15N resolution in a three-dimensional TROSY HNCO experiment for accurate PRE and RDC measurements. Journal of Magnetic Resonance. 200 (2), 173-177 (2009).
  48. Anthis, N. J., Doucleff, M., Clore, G. M. Transient, sparsely populated compact states of apo and calcium-loaded calmodulin probed by paramagnetic relaxation enhancement: interplay of conformational selection and induced fit. Journal of the American Chemical Society. 133 (46), 18966-18974 (2011).
  49. Battiste, J. L., Wagner, G. Utilization of site-directed spin labeling and high-resolution heteronuclear nuclear magnetic resonance for global fold determination of large proteins with limited nuclear overhauser effect data. 생화학. 39 (18), 5355-5365 (2000).
  50. Donaldson, L. W., et al. Structural characterization of proteins with an attached ATCUN motif by paramagnetic relaxation enhancement NMR spectroscopy. Journal of the American Chemical Society. 123 (40), 9843-9847 (2001).
  51. Gaponenko, V., et al. Protein global fold determination using site-directed spin and isotope labeling. Protein Science. 9 (2), 302-309 (2000).
  52. Trindade, I. B., Invernici, M., Cantini, F., Louro, R. O., Piccioli, M. PRE-driven protein NMR structures: an alternative approach in highly paramagnetic systems. FEBS Journal. 288 (9), 3010-3023 (2021).
  53. Nitsche, C., Otting, G. Pseudocontact shifts in biomolecular NMR using paramagnetic metal tags. Progress in Nuclear Magnetic Resonance Spectroscopy. 98-99, 20-49 (2017).

Play Video

Cite This Article
Johnson, C. N., Libich, D. S. Paramagnetic Relaxation Enhancement for Detecting and Characterizing Self-Associations of Intrinsically Disordered Proteins. J. Vis. Exp. (175), e63057, doi:10.3791/63057 (2021).

View Video