Observation of Fast 2D NMR Spectra During Protein Folding Using

Subscriber access provided by UNIV OF GEORGIA

Biophysical Chemistry, Biomolecules, and Biomaterials; Surfactants and Membranes

Observation of Fast 2D NMR Spectra During Protein Folding Using Polarization Transfer from Hyperpolarized Water Jihyun Kim, Ratnamala Mandal, and Christian Hilty J. Phys. Chem. Lett., Just Accepted Manuscript • DOI: 10.1021/acs.jpclett.9b02197 • Publication Date (Web): 23 Aug 2019 Downloaded from pubs.acs.org on August 26, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

The Journal of Physical Chemistry Letters

Observation of Fast 2D NMR Spectra During Protein Folding Using Polarization Transfer from Hyperpolarized Water Jihyun Kim, Ratnamala Mandal, and Christian Hilty* Chemistry Department, Texas A&M University, 3255 TAMU, College Station, TX 77843, USA Corresponding Author *E-mail: [email protected]

ABSTRACT

Nuclear spin hyperpolarized water is utilized to obtain protein spectra not only in the folded state but also during the refolding process. Polarization transfer to Ribonuclease Sa through proton exchange and the nuclear Overhauser effect (NOE) results in NMR signal enhancements of amide protons by up to 24-fold. These enhancements enable the measurement of fast 2D NMR spectra on the same time scale as the folding. Resolved amide proton signals corresponding to the folded protein are observed both under folded and refolding conditions, whereby the refolding protein shows smaller transferred signals. Residue-specific evaluation of contributions to the polarization transfer indicates that signals attributed to a relayed intramolecular NOE are not observable in the refolding experiment. These differences are explained by the absence of long-range contacts and

ACS Paragon Plus Environment

1

The Journal of Physical Chemistry Letters 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 23

faster molecular motions in the unfolded protein. Applications of this method include accessing residue-specific information on structure and dynamics during multi-state protein folding.

TOC GRAPHICS

KEYWORDS Protein folding, Hyperpolarization, Dissolution dynamic nuclear polarization, NMR spectroscopy


2

Page 3 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Structural changes occurring during the folding process of a protein are not directly accessible with most methods of structural biology. Insight into this process can be indirectly obtained through nuclear magnetic resonance (NMR), for example, quenched-flow hydrogen-exchange labeling experiments which provide structural information on folding intermediates.1,2 Backbone dynamics in folded and unfolded equilibrium also give insights into the nature of the conformational structure at the start and the end points of protein folding, allowing to identify potential folding pathways.3 NMR spectroscopy combined with a stopped-flow device can be used to measure the spectra of folding intermediates in real-time that allows obtaining the information of folding directly. Balbach et al. have demonstrated the measurement of transient spectra of protein using a rapid injection of unfolded protein into refolding buffer, followed by a series of 1D 1H

NMR acquisition.4 Instead of 1H NMR, other NMR active nuclei can be used as labels to avoid

difficulties of assignment and analysis of spectra caused by overlapped 1H signals. Using

19F-

tryptophan labeled proteins, the changes in side-chain environment during the folding have been examined.5,6 Pulse sequences with rapid data acquisition through band-selective optimized flipangle excitations7,8 or spatial encoding9,10 were developed for the real-time measurement of multidimensional spectra. With this sequence, Schanda et al. have studied the folding of ɑ-lactalbumin and ubiquitin by measuring real-time 2D spectra and H/D exchange kinetics, respectively.11 The sensitivity of real-time signal acquisitions can be substantially enhanced by hyperpolarization techniques, which increase the population difference of Zeeman energy levels prior to the NMR experiment. Using chemically induced dynamic nuclear polarization (CIDNP) coupled with 1D experiments, Dobson and Hore have proposed the use of hyperpolarized signals of tryptophan residues on the protein for folding kinetics.12 Overhauser dynamic nuclear


3


Page 4 of 23

polarization (Overhauser DNP) with spin-labeled proteins has recently been introduced to study protein aggregation13 and protein folding14 by monitoring surface protein hydration dynamics. Our group has recently demonstrated the use of dissolution dynamic nuclear polarization (DDNP) for improving signal sensitivity of proteins and monitoring the folding process in real-time.15 Dissolution dynamic nuclear polarization is a hyperpolarization technique, generating polarization in solid-state but the NMR measurements are performed in liquid-state by dissolving the sample with dissolution solvent.16 Utilizing the dissolution step of D-DNP, a wide variety of conditions for protein folding can easily be accessible, for example, changing the concentration of denaturant or pH jump before and after the dissolution. Although this dissolution step causes dilution of the spin magnetization, even after the dilution, the hyperpolarized signal can show a large enhancement. Our group has demonstrated the use of D-DNP for polarizing polypeptides directly.17 Folding of a protein can be followed in real-time by measuring a series of 13C 1D NMR spectra of hyperpolarized protein after pH jump15, and information on molecular dynamics can be obtained from spin relaxation properties.18 Enhancements of protein signals can also be achieved by proton exchange or NOE from hyperpolarized water.19,20 This method has been applied to identify residue-specific enhancements using fast 2D measurements for intrinsically disordered proteins (IDPs)21 and the folded protein.22 However, it has not been applied to measure the protein signals during the folding process. Here, we demonstrate the application of this approach to obtain the protein spectra not only in the static state but also during folding after the dilution of denaturant by a fast 2D NMR. Ribonuclease Sa (RNase Sa) from E. coli was chosen as a model protein.23,24 This protein is a small enzyme comprising a two-stranded β-sheet, a four-stranded antiparallel β-sheet, and a turn of 3/10 helix followed by an α-helix in a total of 96 amino acid residues. Two cysteine residues near the


4

Page 5 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


N and C termini are linked through a disulfide bond. Structures determined by NMR spectroscopy25 and X-ray crystallography26 are available. Stopped-flow fluorescence measurements indicated that the protein folds in a two-step process with rate constants of 1.7 and 0.36 s-1 in the absence of denaturant.27 The two folding rates potentially indicate the existence of an intermediate conformation, which, however, has not been identified yet. These rates decrease at higher denaturant concentrations. Based on the folding kinetics of various mutants of this protein, the contributions of specific interactions of hydrophobic28 and electrostatic27 nature to protein stability were further determined. In the context of the present work, the corresponding time constants are on the same order as the observable window of a stopped-flow DNP-NMR experiment.15 With the ability to enhance amide proton NMR signals through hyperpolarized water, we measure high-resolution spectra of the folded protein, as well as during the refolding of the protein. We evaluate the mechanism of polarization transfer on each residue and discuss the difference between the spectra of the folded and refolding proteins.


5


Page 6 of 23

Figure 1. Comparisons between SOFAST-HMQC spectra of a) folded and b) refolding RNase Sa. Spectra shown in blue were recorded after addition of hyperpolarized water, resulting in a final 1H fraction of 3 % in D2O buffer. The protein concentrations were 0.23 mM for (a) and 0.32 mM for (b). Non-hyperpolarized reference spectra from a sample with 1 mM protein in 90 % protonated solvent are shown in gray. Residue specific peak assignments are indicated for the signals visible in the spectra with hyperpolarized water. 1D slices at δ(15N) = 108 ppm are shown in c) for folded and in d) for refolding protein.

Spectra of RNase Sa observed after admixing of hyperpolarized water are shown in Figure 1. The experiments were conducted under two different conditions. For the spectrum on the left side (Figure 1a), the protein was maintained in its native form in a phosphate buffer at pH 6.5 (see Materials and Methods section). The same buffer was used for the dissolution of hyperpolarized water. The spectrum on the right (Figure 1b) was acquired for a protein sample that was initially


6

Page 7 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


denatured in 8 M urea-d4 and subjected to dilution of the denaturant upon admixing the hyperpolarized water, resulting in the initiation of refolding. The measurement of the spectrum was complete within 5 s, before decay of the hyperpolarized water signal governed by the spinlattice relaxation of water protons (T1 = 4.7 ± 0.3 s in a mixture containing 3 % H2O in D2O buffer). Both of the resulting spectra show peak patterns corresponding to folded protein, as seen by comparison with the underlaid reference spectra shown in gray. This observation is consistent with the published faster folding rate of 1.1 s-1,27 the time of 700 ms elapsed in-between sample mixing and the start of data acquisition, and the total experimental duration of 5 s.29 Although the strongest signals are substantially enhanced by the transfer of polarization from hyperpolarized water, not all peaks of the protein are visible. Since polarization transfer from water occurs mainly to exposed residues, selective enhancements of solvent-exposed amide protons are expected. Through modeling of polarization transfer observed by one-dimensional NMR, we had previously found that signal transfers primarily through proton exchange, followed by intramolecular NOE between protein-bound protons.20 Therefore, some signals may result from NOE contributions, which depend on the local structure and dynamics of the protein. The signal enhancements can be estimated by comparing the peak integrals obtained from 1D slices of DNP and reference spectra, shown in Figure 1c and d. Despite a lower concentration of the protein and lower protonation level, the DNP experiment results in ×1, ×4, and ×4 larger signal integrals than those of the reference sample for the residues 73, 66 in Figure 1c and the residue 66 in Figure 1d. The strongest peak, not shown in these slices, was observed for the residue 61 when the protein is in the folded form, showing 6 times larger signal than that of the reference (Figure S5). After considering the difference of protein concentrations between DNP and reference spectra, the maximum enhancement of 24 can be calculated, which is in good agreement with the


7


Page 8 of 23

previously reported value of 20 from our group,20 despite the fact that this value was determined for a different protein. Comparing the spectrum of folded with the spectrum of refolding protein in Figure 1a and b, it is evident that the pattern of enhanced peaks is different between the two experiments. This difference is also clearly visible in 1D slices (Figure 1c and d) where the signals for both of the residues 73 and 66 were enhanced in the spectrum of the folded protein but only for residue 66 in that of refolding protein. Contrary to what would be expected based on solvent exposure alone, fewer peaks are observed for the refolding protein. This observation suggests that molecular dynamics and the magnitude of exchange rates may play an important role in the transfer and retention of spin polarization.

Figure 2. Averaged peak intensities of each of the resolved residues from three repetitions of DNP SOFAST-HMQC experiments for folded a), refolding b) of the protein, and the residue-specific intensities from non-hyperpolarized measurements of water-selective NOESY c) and CLEANEX-


8

Page 9 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


PM d) at mixing time of 100 ms with the reference sample. In the DNP data, the intensities were normalized by the initial signal enhancements of water and protein concentrations and then averaged. The error bars represent the standard deviation of the three repetitions. The peak intensities were obtained by peak integration using the CARA software package.30

Given the multiple potential contributions to the differences in the signals from the two sample conditions, a comparison to conventional NMR spectroscopy was attempted. Figure 2 shows the signal intensities of the spectra with hyperpolarized water (Figure 2a and b) compared to nonhyperpolarized measurements of NOE and exchange from water (Figure 2c and d). The nonhyperpolarized water-selective NOESY spectrum31 contains contributions from exchange and NOE, whereas the latter is suppressed in the CLEANEX-PM spectrum.32 The majority of the residues identified in the CLEANEX-PM experiments are visible in the DNP experiment with the folded protein, showing a similar pattern of intensities. This suggests that the exchangeable amide protons on the protein in the DNP experiment mainly obtain the polarization through the exchange process. On the other hand, some signals of residues enhanced in the DNP spectrum are observable in the NOESY spectrum but not in the CLEANEX-PM, for example, the residues 8, 9, 28, 30, 34, 56 – 59, 68, 69, 72, 73, and 88. This difference may be caused by indirect polarization transfer from the water via exchangeable protons on the protein, known as exchange𝑘𝑒𝑥

relayed NOEs (𝐻𝑤 𝐻𝑒𝑥

𝑁𝑂𝐸

𝐻𝑁𝐻). Similar observations were reported by our group20 and by

Kadeřávek et al.22, where the polarization transferred to the protein through proton exchange can further spread to neighboring protons within the protein. A direct NOE from water to protein can also occur, which would produce negative enhancements. However, the degree of these


9


Page 10 of 23

contributions is small compared to that of exchange or exchange-relayed NOEs.20 The exchange𝑘𝑒𝑥

relayed NOEs can be present along with the direct polarization transfer via exchange (𝐻𝑤 𝐻𝑁𝐻) depending on the structure, resulting in further contributions in signals. Comparing Figures 2a and c, more signals are observed in the NOESY spectrum compared to that in the DNP experiment. A larger intensity contrast is seen in the DNP data when comparing signals of residues in loop structures to those in the regions of α-helical and β-sheet secondary structure elements, where amide protons are protected from exchange through hydrogen bonding. These differences were also observed when comparing the DNP data to a 3D NOESY-HSQC spectrum, confirming that the difference is not caused by a selection artifact of the water selective pulse (Figure S4). The DNP and conventional NMR experiments differ in the NOE mixing time, which was 100 ms for the water-selective NOESY experiment and 5 s for the DNP experiment, in the protonation level, as well as in the initial spin polarization of water. Therefore, similar, but not necessarily identical signal patterns are expected. This observation also applies to the comparison between the DNP and CLEANEX-PM spectra (Figures 2a and d). In the DNP data measured of refolding protein (Figure 2b), several residues show signal enhancements; the pattern of signal intensities, however, is different when compared to that of the folded protein (Figure 2a). In the refolding protein, the intensities for the residues with fast exchange rates are still observable but show weaker signals. On the other hand, most of the signals identified above as having obtained polarization through intramolecular NOE in the folded form are not observable in the refolding protein. The absence of these signals is consistent with the absence of long-range contacts and faster molecular motions in the unfolded protein. Rate constants were calculated from the initial slopes of the non-hyperpolarized NOESY and EXSY spectra measured with mixing times ranging from 5 to 20 ms (Figure S5). The resulting


10

Page 11 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


rates correlate with the DNP intensities of the folded protein, after normalization of these intensities with the non-hyperpolarized reference spectrum. The comparison between the normalized DNP intensities and the rate constants result in an identical conclusion as from Figure 2. For this reason, and because the rate constants are not obtained from the DNP experiment, the comparison between intensities from the DNP and NOESY or EXSY measurements is preferred.

Figure 3. A color map of the residue-specific peak intensities from the DNP experiment, shown in Figure 2a and b, a) of folded, and b) of refolding forms of the protein. c) The intensities from the CLEANEX-PM, shown in Figure 2d, mapped onto the structure of the protein. In (a)-(c), the backbone amide protons are shown as spheres. d) Representation of the protein showing the backbone amide protons identified to obtain polarization through NOE (residues 8, 9, 28, 30, 34, 56 – 59, 68, 69, 72, 73, and 88) as red spheres. Other backbone amide protons are shown as gray spheres, and the hydroxyl protons and amide protons on side-chains on the protein are shown as


11


Page 12 of 23

green and yellow spheres, respectively. All figures are plotted based on the solution structure of RNase Sa (PDB code 1C54).33 Images were created using UCSF Chimera software.34

The difference of observed signals in folded and refolding cases are visible on the protein structures, where the signal intensities of the respective peaks are mapped (Figure 3a and b). The signals of residues that are exposed to the surface and not located in secondary structures are observed in both folded and refolding proteins, even though they are weaker in the refolding experiment. Comparing Figures 3a and c, the locations of the residues showing signals in the DNP experiment but not in the CLEANEX-PM, are seen. These residues are mainly located near the βsheet and part of the loops at the center of the protein, as well as near the turn of 3/10 helix, where slow exchange is expected. For example, residue 9 on the 3/10 helix and the residue 73 on the βsheet are visible in Figure 3a but not in Figure 3c. The difference suggests that these residues obtain the polarization through exchange-relayed NOE. The signals for these residues are not observable under the refolding conditions (Figure 3b), for the reasons discussed above. The residues thus identified as receiving polarization through NOE are colored red in Figure 3d. The origin of exchange-relayed NOEs can be estimated by identifying closely located exchangeable protons. From comparing to Figure 3a, amide proton signals enhanced through NOE are not directly explained by proximity to other backbone amide protons that are enhanced through proton exchange. For example, no other amide protons with observable signals in the DNP spectra are located within 6 Å from the amide protons of residues 9 and 73. Polarization can also be transferred through multiple steps, or from other protons including hydroxyl protons (OH) and amide protons on side-chains. The locations of OH and amide protons on side-chains are shown as green and yellow spheres, respectively, in Figure 3d. Many of the


12

Page 13 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


amide protons identified as enhanced by exchange-relayed polarization transfer (red spheres) are in proximity to OH protons (green spheres). For example, the distances from the amide protons of residues 9 and 73 to nearby OH protons are 4.5 Å and 4.7 Å, respectively. On the other hand, distances to amide protons on side-chains (yellow spheres) are longer, suggesting a contribution of OH protons to the transferred NOE. Polarization transfer through aliphatic protons in multiple steps is further possible. This polarization would also originate at the locations of the exchangeable protons indicated. For comparison, the solvent-accessible surface (areaSAS) was calculated for each amide proton using UCSF chimera software34 (Figure S6). Comparing to Figure 3c, most of the amide protons showing signals in the CLEANEX-PM experiment are identified in the areaSAS calculation. The signal intensities are not related quantitatively to the areaSAS, since the proton exchange process also depends on the existence of hydrogen bonding. Most of the amide protons identified to obtain polarization through NOE (red spheres in Figure 3d) are not predicted to be accessible to the solvent by the areaSAS, further supporting that these protons obtain the polarization through NOE rather than through proton exchange. In summary, we have measured enhanced 2D NMR spectra of RNase Sa under folded and refolding conditions with polarization transfer from water hyperpolarized by dissolution DNP. The residue-specific contributions to the polarization transfer mechanism, including proton exchange and NOE, were evaluated. In both cases, the spectrum of folded protein was ultimately observed. However, in the dynamic sample undergoing refolding after the dilution of denaturant, a smaller number of residues obtained polarization through NOE compared to the folded protein. The combined use of hyperpolarized water and rapid 2D NMR measurement allows detecting residue-


13


Page 14 of 23

specific signals within the folding timescale of many proteins and opens the possibility to characterize relevant structural and dynamic features.

ASSOCIATED CONTENT Supporting Information. The supporting Information is available free of charge. 1. Materials and Methods (Protein Expression, Purification, and Sample Preparation/Protein Characterization and Chemical Shift Assignments/Hyperpolarized NMR/Non-hyperpolarized NOE and Exchange Measurements), 2. Comparison between Water-selective 2D NOESY-HSQC and 3D NOESY-HSQC, 3. Comparison DNP with Rates determined Non-hyperpolarized Measurements, 4. Summary of DNP Experiments, 5. Calculation of Solvent Accessible Surface Area of Amide Protons (PDF)

AUTHOR INFORMATION Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT Financial support from the National Science Foundation (Grant CHE-1362691) and the Welch Foundation (A-1658) is gratefully acknowledged.


14

Page 15 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


REFERENCES (1) Roder, H.; Elöve, G. A.; Englander, S. W. Structural Characterization of Folding Intermediates in Cytochrome c by H-Exchange Labelling and Proton NMR. Nature 1988, 335, 700–704. (2) Udgaonkar, J. B.; Baldwin, R. L. NMR Evidence for an Early Framework Intermediate on the Folding Pathway of Ribonuclease A. Nature 1988, 335, 694–699. (3) Farrow, N. A.; Zhang, O.; Forman-Kay, J. D.; Kay, L. E. Comparison of the Backbone Dynamics of a Folded and an Unfolded SH3 Domain Existing in Equilibrium in Aqueous Buffer. Biochemistry 1995, 34, 868–878. (4) Balbach, J.; Forge, V.; van Nuland, N. A. J.; Winder, S. L.; Hore, P. J.; Dobson, C. M. Following Protein Folding in Real Time Using NMR Spectroscopy. Nat. Struct. Mol. Biol. 1995, 2, 865–870. (5) Ropson, I. J.; Frieden, C. Dynamic NMR Spectral Analysis and Protein Folding: Identification of a Highly Populated Folding Intermediate of Rat Intestinal Fatty Acid-Binding Protein by 19F NMR. Proc. Natl. Acad. Sci. 1992, 89, 7222–7226. (6) Hoeltzli, S. D.; Frieden, C. Real-Time Refolding Studies of 6-19F-Tryptophan Labeled Escherichia Coli Dihydrofolate Reductase Using Stopped-Flow NMR Spectroscopy. Biochemistry 1996, 35, 16843–16851.


15


Page 16 of 23

(7) Schanda, P.; Kupče, Ē.; Brutscher, B. SOFAST-HMQC Experiments for Recording TwoDimensional Heteronuclear Correlation Spectra of Proteins within a Few Seconds. J. Biomol. NMR 2005, 33, 199–211. (8) Schanda, P.; Brutscher, B. Very Fast Two-Dimensional NMR Spectroscopy for Real-Time Investigation of Dynamic Events in Proteins on the Time Scale of Seconds. J. Am. Chem. Soc. 2005, 127, 8014–8015. (9) Gal, M.; Schanda, P.; Brutscher, B.; Frydman, L. UltraSOFAST HMQC NMR and the Repetitive Acquisition of 2D Protein Spectra at Hz Rates. J. Am. Chem. Soc. 2007, 129, 1372– 1377. (10) Seginer, A.; Olsen, G. L.; Frydman, L. Acquiring and Processing Ultrafast Biomolecular 2D NMR Experiments Using a Referenced-Based Correction. J. Biomol. NMR 2016, 66, 141–157. (11) Schanda, P.; Forge, V.; Brutscher, B. Protein Folding and Unfolding Studied at Atomic Resolution by Fast Two-Dimensional NMR Spectroscopy. Proc. Natl. Acad. Sci. 2007, 104, 11257–11262. (12) Dobson, C. M.; Hore, P. J. Kinetic Studies of Protein Folding Using NMR Spectroscopy. Nat. Struct. Mol. Biol. 1998, 5, 504–507. (13) Pavlova, A.; R. McCarney, E.; W. Peterson, D.; W. Dahlquist, F.; Lew, J.; Han, S. SiteSpecific Dynamic Nuclear Polarization of Hydration Water as a Generally Applicable Approach to Monitor Protein Aggregation. Phys. Chem. Chem. Phys. 2009, 11, 6833–6839.


16

Page 17 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(14) Armstrong, B. D.; Choi, J.; López, C.; Wesener, D. A.; Hubbell, W.; Cavagnero, S.; Han, S. Site-Specific Hydration Dynamics in the Nonpolar Core of a Molten Globule by Dynamic Nuclear Polarization of Water. J. Am. Chem. Soc. 2011, 133, 5987–5995. (15) Chen, H.-Y.; Ragavan, M.; Hilty, C. Protein Folding Studied by Dissolution Dynamic Nuclear Polarization. Angew. Chem. Int. Ed. 2013, 52, 9192–9195. (16) Ardenkjær-Larsen, J. H.; Fridlund, B.; Gram, A.; Hansson, G.; Hansson, L.; Lerche, M. H.; Servin, R.; Thaning, M.; Golman, K. Increase in Signal-to-Noise Ratio of > 10,000 Times in Liquid-State NMR. Proc. Natl. Acad. Sci. 2003, 100, 10158–10163. (17) Ragavan, M.; Chen, H.-Y.; Sekar, G.; Hilty, C. Solution NMR of Polypeptides Hyperpolarized by Dynamic Nuclear Polarization. Anal. Chem. 2011, 83, 6054–6059. (18) Ragavan, M.; Iconaru, L. I.; Park, C.-G.; Kriwacki, R. W.; Hilty, C. Real-Time Analysis of Folding upon Binding of a Disordered Protein by Using Dissolution DNP NMR Spectroscopy. Angew. Chem. Int. Ed. 2017, 56, 7070–7073. (19) Harris, T.; Szekely, O.; Frydman, L. On the Potential of Hyperpolarized Water in Biomolecular NMR Studies. J. Phys. Chem. B 2014, 118, 3281–3290. (20) Kim, J.; Liu, M.; Hilty, C. Modeling of Polarization Transfer Kinetics in Protein Hydration Using Hyperpolarized Water. J. Phys. Chem. B 2017, 121, 6492–6498. (21) Szekely, O.; Olsen, G. L.; Felli, I. C.; Frydman, L. High-Resolution 2D NMR of Disordered Proteins Enhanced by Hyperpolarized Water. Anal. Chem. 2018, 90, 6169–6177. (22) Kadeřávek, P.; Ferrage, F.; Bodenhausen, G.; Kurzbach, D. High-Resolution NMR of Folded Proteins in Hyperpolarized Physiological Solvents. Chem. – Eur. J. 2018, 24, 13418–13423.


17


Page 18 of 23

(23) Hebert, E. J.; Grimsley, G. R.; Hartley, R. W.; Horn, G.; Schell, D.; Garcia, S.; Both, V.; Sevcik, J.; Pace, C. N. Purification of Ribonucleases Sa, Sa2, and Sa3 after Expression in Escherichia Coli. Protein Expr. Purif. 1997, 11, 162–168. (24) Marley, J.; Lu, M.; Bracken, C. A Method for Efficient Isotopic Labeling of Recombinant Proteins. J. Biomol. NMR 2001, 20, 71–75. (25) Laurents, D. V.; Pérez-cañadillas, J. M.; Santoro, J.; Rico, M.; Schell, D.; Hebert, E. J.; Pace, C. N.; Bruix, M. Letter to the Editor: Sequential Assignment and Solution Secondary Structure of Doubly Labelled Ribonuclease Sa. J. Biomol. NMR 1999, 14, 89–90. (26) Sevcík, J.; Lamzin, V. S.; Dauter, Z.; Wilson, K. S. Atomic Resolution Data Reveal Flexibility in the Structure of RNase Sa. Acta Crystallogr. D Biol. Crystallogr. 2002, 58, 1307– 1313. (27) Trefethen, J. M.; Pace, C. N.; Scholtz, J. M.; Brems, D. N. Charge–Charge Interactions in the Denatured State Influence the Folding Kinetics of Ribonuclease Sa. Protein Sci. Publ. Protein Soc. 2005, 14, 1934–1938. (28) Pace, C. N.; Fu, H.; Fryar, K. L.; Landua, J.; Trevino, S. R.; Shirley, B. A.; Hendricks, M. M.; Iimura, S.; Gajiwala, K.; Scholtz, J. M.; et al. Contribution of Hydrophobic Interactions to Protein Stability. J. Mol. Biol. 2011, 408, 514–528. (29) Bowen, S.; Hilty, C. Rapid Sample Injection for Hyperpolarized NMR Spectroscopy. Phys. Chem. Chem. Phys. 2010, 12, 5766–5770. (30) Keller, R. L. J. Optimizing the Process of Nuclear Magnetic Resonance Spectrum Analysis and Computer Aided Resonance Assignment. Doctoral Thesis, ETH Zurich, 2005.


18

Page 19 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(31) Böckmann, A.; Penin, F.; Guittet, E. Rapid Estimation of Relative Amide Proton Exchange Rates of 15N-Labelled Proteins by a Straightforward Water Selective NOESY-HSQC Experiment. FEBS Lett. 1996, 383, 191–195. (32) Hwang, T.-L.; Zijl, P. C. M. van; Mori, S. Accurate Quantitation of Water-Amide Proton Exchange Rates Using the Phase-Modulated CLEAN Chemical EXchange (CLEANEX-PM) Approach with a Fast-HSQC (FHSQC) Detection Scheme. J. Biomol. NMR 1998, 11, 221–226. (33) Laurents, D.; Pérez-Cañadillas, J. M.; Santoro, J.; Rico, M.; Schell, D.; Pace, C. N.; Bruix, M. Solution Structure and Dynamics of Ribonuclease Sa. Proteins Struct. Funct. Bioinforma. 2001, 44, 200–211. (34) Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. UCSF Chimera-a Visualization System for Exploratory Research and Analysis. J. Comput. Chem. 2004, 25, 1605–1612.


19


Figure 1. Comparisons between SOFAST-HMQC spectra of a) folded and b) refolding RNase Sa. Spectra shown in blue were recorded after addition of hyperpolarized water, resulting in a final 1H fraction of 3 % in D2O buffer. The protein concentrations were 0.23 mM for (a) and 0.32 mM for (b). Non-hyperpolarized reference spectra from a sample with 1 mM protein in 90 % protonated solvent are shown in gray. Residue specific peak assignments are indicated for the signals visible in the spectra with hyperpolarized water. 1D slices at δ(15N) = 108 ppm are shown in c) for folded and in d) for refolding protein.


Page 20 of 23

Page 21 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 2. Averaged peak intensities of each of the resolved residues from three repetitions of DNP SOFASTHMQC experiments for folded a), refolding b) of the protein, and the residue-specific intensities from nonhyperpolarized measurements of water-selective NOESY c) and CLEANEX-PM d) at mixing time of 100 ms with the reference sample. In the DNP data, the intensities were normalized by the initial signal enhancements of water and protein concentrations and then averaged. The error bars represent the standard deviation of the three repetitions. The peak intensities were obtained by peak integration using the CARA software package.30



Figure 3. A color map of the residue-specific peak intensities from the DNP experiment, shown in Figure 2a and b, a) of folded, and b) of refolding forms of the protein. c) The intensities from the CLEANEX-PM, shown in Figure 2d, mapped onto the structure of the protein. In (a)-(c), the backbone amide protons are shown as spheres. d) Representation of the protein showing the backbone amide protons identified to obtain polarization through NOE (residues 8, 9, 28, 30, 34, 56 – 59, 68, 69, 72, 73, and 88) as red spheres. Other backbone amide protons are shown as gray spheres, and the hydroxyl protons and amide protons on sidechains on the protein are shown as green and yellow spheres, respectively. All figures are plotted based on the solution structure of RNase Sa (PDB code 1C54).33 Images were created using UCSF Chimera software.34


Page 22 of 23

Page 23 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


TOC GRAPHICS


Observation of Fast 2D NMR Spectra During Protein Folding Using

Recommend Documents