Hyperpolarized Water Enhances Two-Dimensional Proton NMR

Subscriber access provided by KEAN UNIV

Communication

Hyperpolarized water enhances two-dimensional proton NMR correlations: a new approach for molecular interactions Aude Sadet, Cristina Stavarache, Mihaela Bacalum, Mihai Radu, Geoffrey Bodenhausen, Dennis Kurzbach, and Paul R. Vasos J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.9b03651 • Publication Date (Web): 01 Aug 2019 Downloaded from pubs.acs.org on August 2, 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 14 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

Journal of the American Chemical Society

Hyperpolarized water enhances two-dimensional proton NMR correlations: a new approach for molecular interactions Aude Sadet1, Cristina Stavarache1,2, Mihaela Bacalum3, Mihai Radu3, Geoffrey Bodenhausen4, Dennis Kurzbach4,5*, and Paul R. Vasos1,6* 1

Research Institute of the University of Bucharest (ICUB), 36-46 B-dul M. Kogalniceanu, RO-050107, Bucharest, Romania ‘‘C.D. Nenitescu” Centre of Organic Chemistry, 202-B Spl. Independentei, RO-060023 Bucharest, Romania

2

”Horia Hulubei” National Institute for Physics and Nuclear Engineering, Department of Life and Environmental Physics, 30 Reactorului Street, RO077125, Bucharest-Magurele, Romania

3

4

Laboratoire des biomolécules, LBM, Département de chimie, École normale supérieure, PSL University, Sorbonne Université, CNRS, 75005 Paris, France,

5

University of Vienna, Faculty of Chemistry, Institute of Biological Chemistry, Währinger Str. 38, 1090 Vienna, Austria

6

ELI-NP, Extreme Light Infrastructure - Nuclear Physics, IFIN-HH, 30 Reactorului Street, RO-077125, Bucharest-Magurele, Romania

Supporting Information material including experimental details and calculations performed on the system is available. ABSTRACT: Proteins and peptides interactions are characterized in the liquid state by multidimensional NMR spectroscopy experiments, which can take hours to record. We show that, starting from hyperpolarized HDO, two-dimensional (2D) proton correlation maps of a peptide, either free in solution or interacting with liposomes, can be acquired in less than 60 s. In standard 2D NMR spectroscopy without hyperpolarization, the acquisition time required for similar spectral correlations is of the order of hours. This hyperpolarized experiment allows to identify amino-acids featuring solvent-interacting hydrogens and provides fast spectroscopic analysis of peptide conformers. These experiments are a useful and straightforward tool for biochemistry and structural biology, as they do not recur to nitrogen-15 or carbon-13 isotope enrichment.

In biomolecular magnetic resonance spectroscopy, long experimental times are regarded as sine qua non for structural accuracy. Multi-dimensional NMR experiments, which yield the most detailed descriptions of molecules in solution, typically last hours, sometimes even days, and curtailing these durations is necessary to obtain relevant information on chemical reactions, biomolecular interactions or biochemical processes. The environment of biomolecules in solution is crucial for their function, in particular at the surface of membranes1, where their conformations are often stabilized through solvent interactions. Fundamental studies of the interactions between water and biomolecules have wide-reaching practical implications in structural biology2 and clinical imaging, as water-proton magnetization transfer to nuclear spins is at the basis of a widely-used method, known as chemical-exchange saturation transfer (CEST)3,4,5. Molecular agents have been developed to use chemical exchange as a contrast mechanism in MRI6,7. Hyperpolarised water protons are a route towards sensitive NMR spectroscopy8,9. Recently, several approaches have been devised. For example, para-hydrogen induced polarization (PHIP) generated via surface-mediated catalysis has been shown to effectively replenish hyperpolarized HDO10. Water-soluble PHIP catalysts have also become available lately11. Furthermore, the perspective of water hyperpolarisation by means of Signal amplification by reversible exchange (SABRE) has recently been proposed12. Here we employ dissolution dynamic nuclear polarization (D-DNP) to obtain hyperpolarized HDO. D-DNP currently yields the strongest

signal enhancements and has the advantage that it is broadly applicable. Enhanced 1H-15N correlations were obtained in this manner in 15N-enriched proteins13,14,15,16,17,18. The short life-time of D-DNP-derived hyperpolarization requires fast 2D acquisition methods for biological NMR studies. Successful approaches speed-up signal acquisition by spatial encoding of 2D spectra19 or reduce magnetization-recovery delays between successive scans20,21. Cross-peaks can be enhanced using the interplay between indirect-dimension evolution and relaxation delays22. However, water-exchanging hydrogens often remain elusive due to the timescales of their exchange. In such instances, hyperpolarization can be used to bring HN signals, which are the cornerstone of spectral correlations, into a detectable range. Transfer occurs from water via exchange and nuclear Overhauser effects, both exchange-relayed and direct, as observed by Hilty and collaborators23. NMR spectroscopy based on protons can be used to study peptide and protein folding and conformational stability in membranes – a field that is much less advanced than protein folding/unfolding in solution, where a wealth of thermodynamic data is available24. For protein-membrane interactions, a few principles have emerged25, but thermodynamic data remain sparse. The energetics of the partitioning of peptides between membranes and solutions yield valuable information in biochemical studies, some of relevancy for cancer treatment26,27. Proton-proton correlation spectroscopy (COSY)28,29 is the most straightforward route to acquiring atomic-resolution structural information for peptides of less than 10 kDa, whenever isotope labelling is not necessary or feasible. Here we describe an experiment that allows high-resolution characterization of peptide conformations in less than 1 minute. The very short duration of the described experiments is obtained using signal enhancement by Dissolution Dynamic Nuclear Polarization (D-DNP). For the proof of concept, we chose the dipeptide alanine-glycine (henceforth, AlaGly) dissolved in hyperpolarized HDO as a model system to study peptide-liposome interactions by hyperpolarized COSY(Fig. 1).

ACS Paragon Plus Environment

Journal of the American Chemical Society 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 14

blue). Both spectra are processed in magnitude mode. Signal assignments are based on a conventional COSY spectrum covering both amide and aliphatic regions (Supporting Information). B) Overlay of projections of proton correlation spectra from Hyperpolarized COSY (red) and conventional COSY (blue, magnified for signal visibility) spectra.

Figure 1. Proton spectrum of AlaGly obtained without hyperpolarization (using thermal-equilibrium magnetization. The inset shows a space-fill representation of AlaGly, with water-exchangeable protons in blue. To obtain hyperpolarized COSY data, we followed a previouslyreported experimental strategy16 and dissolved the AlaGly peptide in a hyperpolarized mixture of 4% H2O in D2O, with enhancements of the HDO protons on the order of ɛHDO =405, which yielded NMR spectra with DNP-enhanced peptide signal intensities, I, stemming from HDO15,17,18 by a factor ɛ = IHyperpolarized/Iconventional ̴ 405. In the hyperpolarized COSY spectrum (Figure 2) we note, in addition to the cross-peaks Gly-HN/Gly-Hα and Ala-HN/Ala-Hα originating from magnetization relayed via intra-residue J-couplings to Hα protons, the appearance of several cross peaks that benefit from enhanced signal intensities. Due to the high sensitivity of the experiment, the Ala-HN/Ala-Hβ correlation stemming from transfer via 4J-couplings is observed and inter-residue correlations of a similar long-range nature30,31 between Gly-HN and Ala-Hα protons can be inferred, despite overlap at these sites. The conventional COSY in thermal equilibrium (i.e. without hyperpolarization) only revealed magnetization transfer from Ala-HN to Ala-Hα and from Gly-HN to Gly-Hα1 and Gly-Hα2. The signal enhancements obtained for different positions of the 2D map, as well as newly-detected signals, are listed in Table 1, column II.

The longitudinal relaxation of magnetization during sample transfer between the polarizer and the detection system is crucial for the obtained enhancement. The key advantage of hyperpolarized HDO is that its relaxation time constant is relatively long, T1(HDO) ̴ 8 s, allowing to observe magnetization up to tens of seconds after hyperpolarization. By comparison, direct polarization of 1H nuclei in AlaGly followed by dissolution, transfer and injection is expected to result in lower signal enhancements, even for the slowestrelaxing methyl protons in Ala, with typical T1 (1H) ̴ 2 s in high magnetic fields, and even shorter during the transfer step in low fields. Therefore, one can obtain multidimensional hyperpolarized spectra within the time window imposed by HDO relaxation rather than being limited by the much faster relaxation of the target peptide. The obtained enhancements (Table 1) allowed us to elevate the signal amplitudes of formerly-invisible correlations above the detection threshold. Proton correlations have been adequately detected in hyperpolarized 2D map. Three points need to be considered in comparison to a conventional COSY experiment: (i) a small spectral window was used in the indirect dimension in order to preserve resolution. Even with reduced spectral windows, resolution in the indirect dimension of the hyperpolarized experiment remains an issue, due to the limited acquisition time, t1max. To improve resolution under these conditions,‘SOFT-COSY’ experiments32 may be useful. (ii) the method is challenged by very intense water resonances, which impose the challenge to avoid radiation damping effects that would distort the acquired spectra ; the HDO hyperpolarization reservoir also has to be maintained. Hence, the direct dimension of the spectra in Fig. 2 is reduced and selective excitation of the peptide resonances was chosen as proposed in earlier work14. (iii) finally, the experiment does not require a conventional recovery delay (typically recovery=4 s) as the proton polarization is replenished by chemical and magnetic exchange with the solvent. We found (Supporting Information) that an overall inter-scan time of is=0.4 s ensures polarisation transfer while optimizing experimental time. Once the protocol of hyperpolarized COSY established, we studied the interaction of AlaGly with liposomes (Figure 3). The liposome consisted of large unilamellar vesicles (LUVs) prepared as detailed in the Supporting Information.

Figure 2. A) Hyperpolarised 2D COSY proton correlations in the amide region of AlaGly and signal assignments. Hyperpolarized COSY recorded in 1 min (in red) is overlaid with a conventional COSY obtained without hyperpolarization in the same time (in

Figure 3. Changes in hyperpolarized COSY maps due to interactions with liposomes. A, B) Hyperpolarized COSY and projections

2 ACS Paragon Plus Environment

Page 3 of 14 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


thereof (integrals over the indirect dimension) for AlaGly without liposomes (shown in red) and in presence of liposomes, shown in blue (the scale is multiplied by a factor 4 in the projection). All COSY spectra were processed in magnitude mode. C) Representation of free AlaGly and a conformation interacting with the liposome. D): Snapshot from an MD trajectory of the peptide forming a hydrogen bound (yellow line) between the Ala residue and DOPC (dioleoyl-phosphocholine) on the bilayer surface. We observed a pronounced difference between spectra recorded in the presence and absence of liposomes. Compared to free AlaGly, an overall reduction of signal intensities of the amide region is immediately detected in the liposome-containing sample. The signal losses in spectra recorded in the presence of liposomes, compared to free AlaGly, can be traced back to two factors: i) hindered exchange between HDO and HN sites and ii) accelerated relaxation of peptide protons in presence of LUVs via slowed dynamics upon binding. While signal losses via factor ii) trigger a mostly uniform scaling of spectral intensities in a small peptide, losses due to effect i) lead to more pronounced decrease in signal amplitudes for sites that are shielded from the solvent. We observed a site-specific effect, i.e., the reduction of the intensity of Ala-NH3-based cross-signals compared to those based on Gly-HN (Table 1, column III). The signal enhancement of the Ala-HN/Ala-Hα correlation is 14-fold reduced, while the Gly-HN/Gly-Hα correlation is reduced only by a factor 6. Hence, signal changes are compatible with effect i), and the impact of liposome binding on the AlaGly signal enhancement indicates stronger interactions of the Ala residue with the liposome surface. Table 1. Signal intensity (S) enhancements ɛ = S(Hyperpolarized COSY)/S(conventional COSY), for selected cross-peaks. “Only in hyperpolarized COSY” indicates that the conventional COSY did not show these correlations. Signal ratios, ρ, between signal intensities of free AlaGly peptide (Sfp) and liposome-interacting AlaGly (Sl_p) observed in Hyperpolarized COSY maps.

and 2D NOESY experiments33 showed that the transfer of polarization is based on a combination of both NOE’s from the solvent and chemical exchange (Supporting Information). Recent advances in observation techniques showed that even small peptides have defined structural propensities34,35,36. These preferences are more marked in the presence of interaction partners. Amphiphilic peptides of a few amino acids bind to lipid vesicles through exothermal reactions with binding energies35, ΔG, that depend on the peptide sequence and vary between -10 and -80 kJ/mol. Energy gaps of this order of magnitude are typical for staunchlyassociated partners. The increase in spectral dispersion observed upon liposome addition (Figure 3B) indicates a spread in peptide conformations. This spread occurs over ca 0.5 ppm, i.e., 400 Hz under our experimental conditions. The major conformations that appear upon liposome introduction feature signals with similar intensities. This implies that the energy gaps that separate these conformations at the temperature of the study, T = 300 K, are of the order of ΔG ̴ -RT = -2.5 kJ/mol (details in the Supporting Information), where R = 8.3 J/(K•mol) is the universal gas constant. To validate the interaction model obtained from the experiments we employed molecular dynamics (MD) simulations. The peptide interacting with the lipid bilayer was sampled for 200 ns using a CHARMM-based simulation platform (details in Supporting Information). The peptide, initially assumed in a random conformation, was seen to spend a significant time in close contact with the bilayer surface. Most of the hydrogen bounds (~ 92 %,) occurred when the peptide is oriented with the Ala residue toward the bilayer surface (Fig. 4), forming hydrogen bonds between the Ala N-terminus and phosphatidylcholine anions. This corroborates our deduction that the positively-charged peptide N-terminus forms the primary interaction site with the membrane surface. The binding energy calculated from MD spans the range -0.98 to -12.89 kJ/mol, reasonably fitting with the experimentally-inferred value mentioned above. Structural information can be obtained within 1 minute by 2D proton correlation spectroscopy using hyperpolarized HDO protons. The residue-dependent enhancement factors yield information about binding sites and can help assign magnetic resonance signals to atoms within molecular and inter-molecular frameworks. The detection of biomolecular interactions without any recourse to isotopic enrichment makes possible fast and straightforward applications, such as drug screening for binding to defined biomolecular sites via hyperpolarized correlation spectroscopy.

Signal

Enhancement ɛ

ρ = Sfp/Sl_p

Gly-HN / Gly-H

114

6

Ala-HN / Ala-H

190

14

Ala-HN / Ala-Hβ

only in hyperpolar-

no signal in lipo-

ized COSY

some sample

only in hyperpolar-

no signal in lipo-

ized COSY

some sample

Corresponding Author

no signal in lipo-

Paul Vasos, University of Bucharest, ELI-NP IFIN-HH [email protected]

Gly-HN / Ala-H

Gly-HN / HDO

some sample Ala-HN / HDO

no signal in liposome sample

Supporting Information

AUTHOR INFORMATION

Dennis Kurzbach, University of Vienna [email protected]

Notes The authors declare no competing financial interests.

The transfer of hyperpolarization from HDO to the detected amides can be based on i) chemical proton exchange, ii) direct magnetic exchange, i.e. nuclear Overhauser effects, from the solvent to the HN protons and iii) on exchange-relayed NOEs. 1D water-selective

ACKNOWLEDGMENT The authors thank G. Necula, F. Teleanu, and P. Ghenuche for assistance with the molecular simulation and useful discussions. We



acknowledge support from the Romanian Ministry of Research, PN 19 06 01 05 / 2019 and UEFISCDI PN-III-P4-ID-PCE-2016-0887. Work has been supported by the Extreme Light Infrastructure Nuclear Physics (ELI-NP) Phase II, a project co-financed by the Romanian Government and the European Union through the European Regional Development Fund and the Competitiveness Operational Programme (1/07.07.2016, ID 1334). This work was supported by the French CNRS and the ERC, contract ‘dilute para water’, grant agreement 339754. The authors thank Bruker BioSpin for the DNP equipment and the DNP-NMR facility at ENS for access supported by the Infrastructure de recherche de resonance magnetique nucleaire a tres hauts champs (IR RMN THC) of the CNRS.

(12)

REFERENCES (1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

(11)

Ball, P. Water Is an Active Matrix of Life for Cell and Molecular Biology. Proc Natl Acad Sci U S A 2017, 114 (51), 13327–13335. https://doi.org/10.1073/pnas.1703781114. Otting, G.; Liepinsh, E.; Wüthrich, K. Protein Hydration in Aqueous Solution. Science 1991, 254 (5034), 974–980. Forsén, S.; Hoffman, R. A. Study of Moderately Rapid Chemical Exchange Reactions by Means of Nuclear Magnetic Double Resonance. J. Chem. 39 (11), 2892–2901. Phys. 1963, https://doi.org/10.1063/1.1734121. Wolff, S. D.; Balaban, R. S. Magnetization Transfer Contrast (MTC) and Tissue Water Proton Relaxation in Vivo. Magnetic Resonance in Medicine 1989, 10 (1), 135–144. https://doi.org/10.1002/mrm.1910100113. Ward, K. M.; Aletras, A. H.; Balaban, R. S. A New Class of Contrast Agents for MRI Based on Proton Chemical Exchange Dependent Saturation Transfer (CEST). Journal of Magnetic Resonance 2000, 143 (1), 79–87. https://doi.org/10.1006/jmre.1999.1956. Zhou, J.; Lal, B.; Wilson, D. A.; Laterra, J.; Zijl, P. C. M. van. Amide Proton Transfer (APT) Contrast for Imaging of Brain Tumors. Magnetic Resonance in Medicine 2003, 50 (6), 1120–1126. https://doi.org/10.1002/mrm.10651. Vallurupalli, P.; Bouvignies, G.; Kay, L. E. Studying “Invisible” Excited Protein States in Slow Exchange with a Major State Conformation. J. Am. Chem. Soc. (19), 8148–8161. 2012, 134 https://doi.org/10.1021/ja3001419. Harris, T.; Szekely, O.; Frydman, L. On the Potential of Hyperpolarized Water in Biomolecular NMR Studies. J. Phys. Chem. B 2014, 118 (12), 3281– 3290. https://doi.org/10.1021/jp4102916. Ardenkjaer-Larsen, J. H.; Fridlund, B.; Gram, A.; Hansson, G.; Hansson, L.; Lerche, M. H.; Servin, R.; Thaning, M.; Golman, K. Increase in Signal-toNoise Ratio of > 10,000 Times in Liquid-State NMR. Proc. Natl. Acad. Sci. U.S.A. 2003, 100 (18), 10158–10163. https://doi.org/10.1073/pnas.1733835100. Zhao, E. W.; Maligal-Ganesh, R.; Du, Y.; Zhao, T. Y.; Collins, J.; Ma, T.; Zhou, L.; Goh, T.-W.; Huang,

(13)

(14)

(15)

(16)

(17)

(18)

(19)

(20)

Page 4 of 14

W.; Bowers, C. R. Surface-Mediated Hyperpolarization of Liquid Water from Parahydrogen. Chem 2018, 4 (6), 1387–1403. https://doi.org/10.1016/j.chempr.2018.03.004. Lehmkuhl, S.; Emondts, M.; Schubert, L.; Spannring, P.; Klankermayer, J.; Blümich, B.; Schleker, P. P. M. Hyperpolarizing Water with Parahydrogen https://onlinelibrary.wiley.com/doi/abs/10.1002/cphc.201700750 (accessed Jun 24, 2019). https://doi.org/10.1002/cphc.201700750. S. Duckett; W. Iali; S. Roy; F. Ahwal; B. Thickner. Achieving the Hyperpolarization of Pyruvate, Glucose and Urea via SABRE; Asilomar Pacific Grove CA, USA., 2019; p 46. Chappuis, Q.; Milani, J.; Vuichoud, B.; Bornet, A.; Gossert, A. D.; Bodenhausen, G.; Jannin, S. Hyperpolarized Water to Study Protein–Ligand Interactions. J. Phys. Chem. Lett. 2015, 6 (9), 1674–1678. https://doi.org/10.1021/acs.jpclett.5b00403. Vuichoud, B.; Bornet, A.; de Nanteuil, F.; Milani, J.; Canet, E.; Ji, X.; Miéville, P.; Weber, E.; Kurzbach, D.; Flamm, A.; et al. Filterable Agents for Hyperpolarization of Water, Metabolites, and Proteins. Chemistry – A European Journal 2016, 22 (41), 14696–14700. https://doi.org/10.1002/chem.201602506. Kurzbach, D.; Canet, E.; Flamm, A. G.; Jhajharia, A.; Weber, E. M. M.; Konrat, R.; Bodenhausen, G. Investigation of Intrinsically Disordered Proteins through Exchange with Hyperpolarized Water. Angew. Chem. Int. Ed. Engl. 2017, 56 (1), 389–392. https://doi.org/10.1002/anie.201608903. Kadeřávek, P.; Ferrage, F.; Bodenhausen, G.; Kurzbach, D. High-Resolution NMR of Folded Proteins in Hyperpolarized Physiological Solvents. Chemistry 2018, 24 (51), 13418–13423. https://doi.org/10.1002/chem.201802885. Szekely, O.; Olsen, G. L.; Felli, I. C.; Frydman, L. High-Resolution 2D NMR of Disordered Proteins Enhanced by Hyperpolarized Water. Anal. Chem. 90 (10), 6169–6177. 2018, https://doi.org/10.1021/acs.analchem.8b00585. Nastasa, V.; Cristina, S.; Hanganu, A.; Coroaba, A.; Nicolescu, A.; Deleanu, C.; Sadet, A.; R Vasos, P. Hyperpolarised NMR to Follow Water Proton Transport through Membrane Channels via Exchange with Biomolecules. Faraday Discussions 2018, 209, 67-82 https://doi.org/10.1039/C8FD00021B. Frydman, L.; Blazina, D. Ultrafast Two-Dimensional Nuclear Magnetic Resonance Spectroscopy of Hyperpolarized Solutions. Nature Physics 2007, 3 (6), 415–419. https://doi.org/10.1038/nphys597. Schanda, P.; Brutscher, B. Very Fast Two-Dimensional NMR Spectroscopy for Real-Time Investigation of Dynamic Events in Proteins on the Time


Page 5 of 14 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

(21)

(22)

(23)

(24)

(25)

(26)

(27)

(28) (29)

(30)

(31)

(32)

Journal of the American Chemical Society Scale of Seconds. J. Am. Chem. Soc. 2005, 127 (22), 8014–8015. https://doi.org/10.1021/ja051306e. Pervushin, K.; Vögeli, B.; Eletsky, A. Longitudinal 1H Relaxation Optimization in TROSY NMR Spectroscopy. J. Am. Chem. Soc. 2002, 124 (43), 12898– 12902. https://doi.org/10.1021/ja027149q. Novakovic, M.; Cousin, S. F.; Jaroszewicz, M. J.; Rosenzweig, R.; Frydman, L. Looped-PROjected SpectroscopY (L-PROSY): A Simple Approach to Enhance Backbone/Sidechain Cross-Peaks in 1H NMR. J Magn Reson 2018, 294, 169–180. https://doi.org/10.1016/j.jmr.2018.07.010. Chen, C.-H.; Wang, Y.; Hilty, C. Intermolecular Interactions Determined by NOE Build-up in Macromolecules from Hyperpolarized Small Molecules. Methods 2018, 138–139, 69–75. https://doi.org/10.1016/j.ymeth.2018.02.015. Wimley, W. C.; Hristova, K.; Ladokhin, A. S.; Silvestro, L.; Axelsen, P. H.; White, S. H. Folding of βSheet Membrane Proteins: A Hydrophobic Hexapeptide Model11Edited by D. C. Rees. Journal of Molecular Biology 1998, 277 (5), 1091–1110. https://doi.org/10.1006/jmbi.1998.1640. White, S. H. Global Statistics of Protein Sequences: Implications for the Origin, Evolution, and Prediction of Structure. Annual Review of Biophysics and Biomolecular Structure 1994, 23 (1), 407–439. https://doi.org/10.1146/annurev.bb.23.060194.002203. White, S. H.; Wimley, W. C.; Ladokhin, A. S.; Hristova, K. Protein Folding in Membranes: Determining Energetics of Peptide-Bilayer Interactions. Meth. Enzymol. 1998, 295, 62–87. Maffei, M.; Arbesú, M.; Le Roux, A.-L.; Amata, I.; Roche, S.; Pons, M. The SH3 Domain Acts as a Scaffold for the N-Terminal Intrinsically Disordered Regions of c-Src. Structure 2015, 23 (5), 893–902. https://doi.org/10.1016/j.str.2015.03.009. Jeener, J. Ampere International Summer School; Basko Polje, Yugoslavia, 1971. Aue, W. P.; Bartholdi, E.; Ernst, R. R. Two‐dimensional Spectroscopy. Application to Nuclear Magnetic Resonance. J. Chem. Phys. 1976, 64 (5), 2229– 2246. https://doi.org/10.1063/1.432450. Vuister, G. W.; Bax, A. Measurement of Four-Bond HN-Hα J-Couplings in Staphylococcal Nuclease. J Biomol NMR 1994, 4 (2), 193–200. https://doi.org/10.1007/BF00175247. Vuister, G. W.; Bax, A. Quantitative J Correlation: A New Approach for Measuring Homonuclear Three-Bond J(HNH.Alpha.) Coupling Constants in 15N-Enriched Proteins. Journal of the American Chemical Society 115 (17), 7772–7777. Emsley, L.; Bodenhausen, G. Selective Two-Dimensional NMR Experiments for Topological Filtration of Fragments of Coupling Networks. Journal of the American Chemical Society 1991, 113 (9), 3309– 3316. https://doi.org/10.1021/ja00009a014.

(33)

(34)

(35)

(36)

Stonehouse, J.; Shaw, G. L.; Keeler, J. Improving Solvent Suppression in Jump-Return NOESY Experiments. J. Biomol. NMR 1994, 4 (6), 799–805. https://doi.org/10.1007/BF00398410. Seelig, J. Thermodynamics of Lipid–Peptide Interactions. Biochimica et Biophysica Acta (BBA) Biomembranes 2004, 1666 (1), 40–50. https://doi.org/10.1016/j.bbamem.2004.08.004. Dyson, H. J.; Wright, P. E. Defining Solution Conformations of Small Linear Peptides. Annual Review of Biophysics and Biophysical Chemistry 1991, 20 (1), 519–538. https://doi.org/10.1146/annurev.bb.20.060191.002511. de Alba, E.; Blanco, F. J.; Jiménez, M. A.; Rico, M.; Nieto, J. L. Interactions Responsible for the PH Dependence of the Beta-Hairpin Conformational Population Formed by a Designed Linear Peptide. Eur. J. Biochem. 1995, 233 (1), 283–292.



Figure 1 261x170mm (96 x 96 DPI)


Page 6 of 14

Page 7 of 14 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 8 of 14

Page 9 of 14 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


Hyperpolarized water enhances two-dimensional proton NMR correlations: a new approach for molecular interactions Aude Sadet1, Cristina Stavarache1,2, Mihaela Bacalum3, Mihai Radu3, Geoffrey Bodenhausen4, Dennis Kurzbach4,5*, and Paul R. Vasos1,6* 1

Research Institute of the University of Bucharest (ICUB), 36-46 B-dul M. Kogalniceanu, RO-050107, Bucharest, Romania ‘‘C.D. Nenitescu” Centre of Organic Chemistry, 202-B Spl. Independentei, RO-060023 Bucharest, Romania

2

”Horia Hulubei” National Institute for Physics and Nuclear Engineering, Department of Life and Environmental Physics, 30 Reactorului Street, RO077125, Bucharest-Magurele, Romania

3

4

Laboratoire des biomolécules, LBM, Département de chimie, École normale supérieure, PSL University, Sorbonne Université, CNRS, 75005 Paris, France,

5

University of Vienna, Faculty of Chemistry, Institute of Biological Chemistry, Währinger Str. 38, 1090 Vienna, Austria

6

ELI-NP, Extreme Light Infrastructure - Nuclear Physics, IFIN-HH, 30 Reactorului Street, RO-077125, Bucharest-Magurele, Romania

Supporting Information material including experimental details and calculations performed on the system is available. ABSTRACT: Proteins and peptides interactions are characterized in the liquid state by multidimensional NMR spectroscopy experiments, which can take hours to record. We show that, starting from hyperpolarized HDO, two-dimensional (2D) proton correlation maps of a peptide, either free in solution or interacting with liposomes, can be acquired in less than 60 s. In standard 2D NMR spectroscopy without hyperpolarization, the acquisition time required for similar spectral correlations is of the order of hours. This hyperpolarized experiment allows to identify amino-acids featuring solvent-interacting hydrogens and provides fast spectroscopic analysis of peptide conformers. These experiments are a useful and straightforward tool for biochemistry and structural biology, as they do not recur to nitrogen-15 or carbon-13 isotope enrichment.

In biomolecular magnetic resonance spectroscopy, long experimental times are regarded as sine qua non for structural accuracy. Multi-dimensional NMR experiments, which yield the most detailed descriptions of molecules in solution, typically last hours, sometimes even days, and curtailing these durations is necessary to obtain relevant information on chemical reactions, biomolecular interactions or biochemical processes. The environment of biomolecules in solution is crucial for their function, in particular at the surface of membranes1, where their conformations are often stabilized through solvent interactions. Fundamental studies of the interactions between water and biomolecules have wide-reaching practical implications in structural biology2 and clinical imaging, as water-proton magnetization transfer to nuclear spins is at the basis of a widely-used method, known as chemical-exchange saturation transfer (CEST)3,4,5. Molecular agents have been developed to use chemical exchange as a contrast mechanism in MRI6,7. Hyperpolarised water protons are a route towards sensitive NMR spectroscopy8,9. Recently, several approaches have been devised. For example, para-hydrogen induced polarization (PHIP) generated via surface-mediated catalysis has been shown to effectively replenish hyperpolarized HDO10. Water-soluble PHIP catalysts have also become available lately11. Furthermore, the perspective of water hyperpolarisation by means of Signal amplification by reversible exchange (SABRE) has recently been proposed12. Here we employ dissolution dynamic nuclear polarization (D-DNP) to obtain hyperpolarized HDO. D-DNP currently yields the strongest signal enhancements and has the advantage that it is broadly applicable. Enhanced 1H-15N correlations were obtained in this manner

in 15N-enriched proteins13,14,15,16,17,18. The short life-time of DDNP-derived hyperpolarization requires fast 2D acquisition methods for biological NMR studies. Successful approaches speed-up signal acquisition by spatial encoding of 2D spectra19 or reduce magnetization-recovery delays between successive scans20,21. Cross-peaks can be enhanced using the interplay between indirectdimension evolution and relaxation delays22. However, water-exchanging hydrogens often remain elusive due to the timescales of their exchange. In such instances, hyperpolarization can be used to bring HN signals, which are the cornerstone of spectral correlations, into a detectable range. Transfer occurs from water via exchange and nuclear Overhauser effects, both exchange-relayed and direct, as observed by Hilty and collaborators23. NMR spectroscopy based on protons can be used to study peptide and protein folding and conformational stability in membranes – a field that is much less advanced than protein folding/unfolding in solution, where a wealth of thermodynamic data is available24. For protein-membrane interactions, a few principles have emerged25, but thermodynamic data remain sparse. The energetics of the partitioning of peptides between membranes and solutions yield valuable information in biochemical studies, some of relevancy for cancer treatment26,27. Proton-proton correlation spectroscopy (COSY)28,29 is the most straightforward route to acquiring atomic-resolution structural information for peptides of less than 10 kDa, whenever isotope labelling is not necessary or feasible. Here we describe an experiment that allows high-resolution characterization of peptide conformations in less than 1 minute. The very short duration of the described experiments is obtained using signal enhancement by Dissolution Dynamic Nuclear Polarization (D-DNP). For the proof of concept, we chose the dipeptide alanine-glycine (henceforth, AlaGly) dissolved in hyperpolarized HDO as a model system to study peptide-liposome interactions by hyperpolarized COSY(Figure 1).



Page 10 of 14

blue). Both spectra are processed in magnitude mode. Signal assignments are based on a conventional COSY spectrum covering both amide and aliphatic regions (Supporting Information). B) Overlay of projections of proton correlation spectra from Hyperpolarized COSY (red) and conventional COSY (blue, magnified for signal visibility) spectra.

Figure 1. Proton spectrum of AlaGly obtained without hyperpolarization (using thermal-equilibrium magnetization. The inset shows a space-fill representation of AlaGly, with water-exchangeable protons in blue. To obtain hyperpolarized COSY data, we followed a previouslyreported experimental strategy16 and dissolved the AlaGly peptide in a hyperpolarized mixture of 4% H2O in D2O, obtaining enhancements of the HDO protons, ɛHDO, in the range 100-200. The first point in the indirect-dimension of the COSY spectrum reveals that peptide signal intensities, I, are enhanced15,17,18 by a factor ɛpeptide=IHyperpolarized/Iconventional=405. In the hyperpolarized COSY spectrum (Figure 2) we note, in addition to the cross-peaks GlyHN/Gly-Hα and Ala-HN/Ala-Hα originating from magnetization relayed via intra-residue J-couplings to Hα protons, the appearance of several cross peaks that benefit from enhanced signal intensities. Due to the high sensitivity of the experiment, the Ala-HN/Ala-Hβ correlation stemming from transfer via 4J-couplings is observed and inter-residue correlations of a similar long-range nature30,31 between Gly-HN and Ala-Hα protons can be inferred, despite overlap at these sites. The conventional COSY in thermal equilibrium (i.e. without hyperpolarization) only revealed magnetization transfer from Ala-HN to Ala-Hα and from Gly-HN to Gly-Hα1 and Gly-Hα2. The signal enhancements obtained for different positions of the 2D map, as well as newly-detected signals, are listed in Table 1, column II.

The longitudinal relaxation of magnetization during sample transfer between the polarizer and the detection system is crucial for the obtained enhancement. The key advantage of hyperpolarized HDO is that its relaxation time constant is relatively long, T1(HDO) ̴ 8 s, allowing to observe magnetization up to tens of seconds after hyperpolarization. By comparison, direct polarization of 1H nuclei in AlaGly followed by dissolution, transfer and injection is expected to result in lower signal enhancements, even for the slowestrelaxing methyl protons in Ala, with typical T1 (1H) ̴ 2 s in high magnetic fields, and even shorter during the transfer step in low fields. Therefore, one can obtain multidimensional hyperpolarized spectra within the time window imposed by HDO relaxation rather than being limited by the much faster relaxation of the target peptide. The obtained enhancements (Table 1) allowed us to elevate the signal amplitudes of formerly-invisible correlations above the detection threshold. Proton correlations have been adequately detected in hyperpolarized 2D map. Three points need to be considered in comparison to a conventional COSY experiment: (i) a small spectral window was used in the indirect dimension in order to preserve resolution. Even with reduced spectral windows, resolution in the indirect dimension of the hyperpolarized experiment remains an issue, due to the limited acquisition time, t1max. To improve resolution under these conditions, ‘SOFT-COSY’ experiments32 may be useful. (ii) the method is challenged by very intense water resonances, which impose to avoid radiation damping effects that would distort the acquired spectra; the HDO hyperpolarization reservoir also has to be maintained. Hence, selective excitation of the peptide resonances was chosen as proposed in earlier work14 (see Supporting Information) and only frequencies beyond 7.5 ppm are acquired in the direct dimension of the spectra in Figure 2. (iii) finally, the experiment does not require a conventional recovery delay (typically recovery=4 s) as the proton polarization is replenished by chemical and magnetic exchange with the solvent. We found (Supporting Information) that an overall inter-scan time of is=0.4 s ensures polarisation transfer while optimizing experimental time. Once the protocol of hyperpolarized COSY established, we studied the interaction of AlaGly with liposomes (Figure 3). The liposome consisted of large unilamellar vesicles (LUVs) prepared as detailed in the Supporting Information.

Figure 2. A) Hyperpolarised 2D COSY proton correlations in the amide region of AlaGly and signal assignments. Hyperpolarized COSY recorded in 1 min (in red) is overlaid with a conventional COSY obtained without hyperpolarization in the same time (in

Figure 3. Changes in hyperpolarized COSY maps due to interactions with liposomes. A, B) Hyperpolarized COSY and projections


Page 11 of 14 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


thereof (integrals over the indirect dimension) for AlaGly without liposomes (shown in red) and in presence of liposomes, shown in blue (the scale is multiplied by a factor 4 in the projection). All COSY spectra were processed in magnitude mode. C) Representation of free AlaGly and a conformation interacting with the liposome. D) Snapshot from an MD trajectory of the peptide forming a hydrogen bound (yellow line) between the Ala residue and DOPC (dioleoyl-phosphocholine) on the bilayer surface. We observed a pronounced difference between spectra recorded in the presence and absence of liposomes. Compared to free AlaGly, an overall reduction of signal intensities of the amide region is immediately detected in the liposome-containing sample. The signal losses in spectra recorded in the presence of liposomes, compared to free AlaGly, can be traced back to two factors: i) hindered exchange between HDO and HN sites and ii) accelerated relaxation of peptide protons in presence of LUVs via slowed dynamics upon binding. While signal losses via factor ii) trigger a mostly uniform scaling of spectral intensities in a small peptide, losses due to effect i) lead to more pronounced decrease in signal amplitudes for sites that are shielded from the solvent. We observed a site-specific effect, i.e., the reduction of the intensity of Ala-NH3-based cross-signals compared to those based on Gly-HN (Table 1, column III). The signal enhancement of the Ala-HN/Ala-Hα correlation is 14-fold reduced, while the Gly-HN/Gly-Hα correlation is reduced only by a factor 6. Hence, signal changes are compatible with effect i), and the impact of liposome binding on the AlaGly signal enhancement indicates stronger interactions of the Ala residue with the liposome surface. Table 1. Signal intensity (S) enhancements ɛ = S(Hyperpolarized COSY)/S(conventional COSY), for selected cross-peaks. “Only in hyperpolarized COSY” indicates that the conventional COSY did not show these correlations. Signal ratios, ρ, between signal intensities of free AlaGly peptide (Sfp) and liposome-interacting AlaGly (Sl_p) observed in Hyperpolarized COSY maps. Signal

Enhancement ɛ

ρ = Sfp/Sl_p

Gly-HN / Gly-H

114

6

Ala-HN / Ala-H

190

14

Ala-HN / Ala-Hβ

only in hyperpolar-

no signal in lipo-

ized COSY

some sample

Gly-HN / Ala-H

Gly-HN / HDO

Recent advances in observation techniques showed that even small peptides have defined structural propensities34,35,36. These preferences are more marked in the presence of interaction partners. Amphiphilic peptides of a few amino acids bind to lipid vesicles through exothermal reactions with binding energies35, ΔG, that depend on the peptide sequence and vary between -10 and -80 kJ/mol. Energy gaps of this order of magnitude are typical for staunchlyassociated partners. The increase in spectral dispersion observed upon liposome addition (Figure 3B) indicates a spread in peptide conformations. This spread occurs over ca 0.5 ppm, i.e., 400 Hz under our experimental conditions. The major conformations that appear upon liposome introduction feature signals with similar intensities. This implies that the energy gaps that separate these conformations at the temperature of the study, T = 300 K, are of the order of ΔG ̴ -RT = -2.5 kJ/mol (details in the Supporting Information), where R = 8.3 J/(K•mol) is the universal gas constant. To validate the interaction model obtained from the experiments we employed molecular dynamics (MD) simulations. The peptide interacting with the lipid bilayer was sampled for 200 ns using a CHARMM-based simulation platform (details in Supporting Information). The peptide, initially assumed in a random conformation, was seen to spend a significant time in close contact with the bilayer surface. Most of the hydrogen bounds (~ 92 %) occurred when the peptide is oriented with the Ala residue toward the bilayer surface (Figure 4), forming hydrogen bonds between the Ala Nterminus and phosphatidylcholine anions. This corroborates our deduction that the positively-charged peptide N-terminus forms the primary interaction site with the membrane surface. The binding energy calculated from MD spans -0.98 to -12.89 kJ/mol, reasonably fitting with the experimentally-inferred value above. Structural information can be obtained within 1 minute by 2D proton correlation spectroscopy using hyperpolarized HDO protons. The residue-dependent enhancement factors yield information about binding sites and can help assign magnetic resonance signals to atoms within molecular and inter-molecular frameworks. The detection of biomolecular interactions without any recourse to isotopic enrichment makes possible fast and straightforward applications, such as drug screening for binding to defined biomolecular sites via hyperpolarized correlation spectroscopy.

Supporting Information

AUTHOR INFORMATION

only in hyperpolar-

no signal in lipo-

Corresponding Author

ized COSY

some sample

Paul Vasos, University of Bucharest, ELI-NP IFIN-HH [email protected]


Ala-HN / HDO

and 2D NOESY experiments33 showed that the transfer of polarization is based on a combination of both NOE’s from the solvent and chemical exchange (Supporting Information).


The transfer of hyperpolarization from HDO to the detected amides can be based on i) chemical proton exchange, ii) direct magnetic exchange, i.e. nuclear Overhauser effects, from the solvent to the HN protons and iii) on exchange-relayed NOEs. 1D water-selective

Dennis Kurzbach, University of Vienna [email protected]

Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT The authors thank G. Necula, F. Teleanu, and P. Ghenuche for assistance with the molecular simulation and useful discussions. We acknowledge support from the Romanian Ministry of Research, PN



19 06 01 05 / 2019 and UEFISCDI PN-III-P4-ID-PCE-2016-0887. Work has been supported by the Extreme Light Infrastructure Nuclear Physics (ELI-NP) Phase II, a project co-financed by the Romanian Government and the European Union through the European Regional Development Fund and the Competitiveness Operational Programme (1/07.07.2016, ID 1334). This work was supported by the French CNRS and the ERC, contract ‘dilute para water’, grant agreement 339754. The authors thank Bruker BioSpin for the DNP equipment and the DNP-NMR facility at ENS for access supported by the Infrastructure de recherche de resonance magnetique nucleaire a tres hauts champs (IR RMN THC) of the CNRS.

REFERENCES (1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

(11)

(12)

Ball, P. Water Is an Active Matrix of Life for Cell and Molecular Biology. Proc Natl Acad Sci U S A (51), 13327–13335. 2017, 114 https://doi.org/10.1073/pnas.1703781114. Otting, G.; Liepinsh, E.; Wüthrich, K. Protein Hydration in Aqueous Solution. Science 1991, 254 (5034), 974–980. Forsén, S.; Hoffman, R. A. Study of Moderately Rapid Chemical Exchange Reactions by Means of Nuclear Magnetic Double Resonance. J. Chem. Phys. 1963, 39 (11), 2892–2901. https://doi.org/10.1063/1.1734121. Wolff, S. D.; Balaban, R. S. Magnetization Transfer Contrast (MTC) and Tissue Water Proton Relaxation in Vivo. Magnetic Resonance in Medicine 1989, 10 (1), 135–144. https://doi.org/10.1002/mrm.1910100113. Ward, K. M.; Aletras, A. H.; Balaban, R. S. A New Class of Contrast Agents for MRI Based on Proton Chemical Exchange Dependent Saturation Transfer (CEST). Journal of Magnetic Resonance 2000, 143 (1), 79–87. https://doi.org/10.1006/jmre.1999.1956. Zhou, J.; Lal, B.; Wilson, D. A.; Laterra, J.; Zijl, P. C. M. van. Amide Proton Transfer (APT) Contrast for Imaging of Brain Tumors. Magnetic Resonance in Medicine 2003, 50 (6), 1120–1126. https://doi.org/10.1002/mrm.10651. Vallurupalli, P.; Bouvignies, G.; Kay, L. E. Studying “Invisible” Excited Protein States in Slow Exchange with a Major State Conformation. J. Am. Chem. Soc. (19), 8148–8161. 2012, 134 https://doi.org/10.1021/ja3001419. Harris, T.; Szekely, O.; Frydman, L. On the Potential of Hyperpolarized Water in Biomolecular NMR Studies. J. Phys. Chem. B 2014, 118 (12), 3281– 3290. https://doi.org/10.1021/jp4102916. Ardenkjaer-Larsen, J. H.; Fridlund, B.; Gram, A.; Hansson, G.; Hansson, L.; Lerche, M. H.; Servin, R.; Thaning, M.; Golman, K. Increase in Signal-toNoise Ratio of > 10,000 Times in Liquid-State NMR. Proc. Natl. Acad. Sci. U.S.A. 2003, 100 (18), 10158–10163. https://doi.org/10.1073/pnas.1733835100. Zhao, E. W.; Maligal-Ganesh, R.; Du, Y.; Zhao, T. Y.; Collins, J.; Ma, T.; Zhou, L.; Goh, T.-W.; Huang,

(13)

(14)

(15)

(16)

(17)

(18)

(19)

(20)

Page 12 of 14

W.; Bowers, C. R. Surface-Mediated Hyperpolarization of Liquid Water from Parahydrogen. Chem 2018, 4 (6), 1387–1403. https://doi.org/10.1016/j.chempr.2018.03.004. Lehmkuhl, S.; Emondts, M.; Schubert, L.; Spannring, P.; Klankermayer, J.; Blümich, B.; Schleker, P. P. M. Hyperpolarizing Water with Parahydrogen https://onlinelibrary.wiley.com/doi/abs/10.1002/cphc.201700750 (accessed Jun 24, 2019). https://doi.org/10.1002/cphc.201700750. S. Duckett; W. Iali; S. Roy; F. Ahwal; B. Thickner. Achieving the Hyperpolarization of Pyruvate, Glucose and Urea via SABRE; Asilomar Pacific Grove CA, USA., 2019; p 46. Chappuis, Q.; Milani, J.; Vuichoud, B.; Bornet, A.; Gossert, A. D.; Bodenhausen, G.; Jannin, S. Hyperpolarized Water to Study Protein–Ligand Interactions. J. Phys. Chem. Lett. 2015, 6 (9), 1674–1678. https://doi.org/10.1021/acs.jpclett.5b00403. Vuichoud, B.; Bornet, A.; de Nanteuil, F.; Milani, J.; Canet, E.; Ji, X.; Miéville, P.; Weber, E.; Kurzbach, D.; Flamm, A.; et al. Filterable Agents for Hyperpolarization of Water, Metabolites, and Proteins. Chemistry – A European Journal 2016, 22 (41), 14696–14700. https://doi.org/10.1002/chem.201602506. Kurzbach, D.; Canet, E.; Flamm, A. G.; Jhajharia, A.; Weber, E. M. M.; Konrat, R.; Bodenhausen, G. Investigation of Intrinsically Disordered Proteins through Exchange with Hyperpolarized Water. Angew. Chem. Int. Ed. Engl. 2017, 56 (1), 389–392. https://doi.org/10.1002/anie.201608903. Kadeřávek, P.; Ferrage, F.; Bodenhausen, G.; Kurzbach, D. High-Resolution NMR of Folded Proteins in Hyperpolarized Physiological Solvents. Chemistry 2018, 24 (51), 13418–13423. https://doi.org/10.1002/chem.201802885. Szekely, O.; Olsen, G. L.; Felli, I. C.; Frydman, L. High-Resolution 2D NMR of Disordered Proteins Enhanced by Hyperpolarized Water. Anal. Chem. 90 (10), 6169–6177. 2018, https://doi.org/10.1021/acs.analchem.8b00585. Nastasa, V.; Cristina, S.; Hanganu, A.; Coroaba, A.; Nicolescu, A.; Deleanu, C.; Sadet, A.; R Vasos, P. Hyperpolarised NMR to Follow Water Proton Transport through Membrane Channels via Exchange with Biomolecules. Faraday Discussions 2018, 209, 67-82 https://doi.org/10.1039/C8FD00021B. Frydman, L.; Blazina, D. Ultrafast Two-Dimensional Nuclear Magnetic Resonance Spectroscopy of Hyperpolarized Solutions. Nature Physics 2007, 3 (6), 415–419. https://doi.org/10.1038/nphys597. Schanda, P.; Brutscher, B. Very Fast Two-Dimensional NMR Spectroscopy for Real-Time Investigation of Dynamic Events in Proteins on the Time


Page 13 of 14 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

(21)

(22)

(23)

(24)

(25)

(26)

(27)

(28) (29)

(30)

(31)

(32)

Journal of the American Chemical Society Scale of Seconds. J. Am. Chem. Soc. 2005, 127 (22), 8014–8015. https://doi.org/10.1021/ja051306e. Pervushin, K.; Vögeli, B.; Eletsky, A. Longitudinal 1H Relaxation Optimization in TROSY NMR Spectroscopy. J. Am. Chem. Soc. 2002, 124 (43), 12898– 12902. https://doi.org/10.1021/ja027149q. Novakovic, M.; Cousin, S. F.; Jaroszewicz, M. J.; Rosenzweig, R.; Frydman, L. Looped-PROjected SpectroscopY (L-PROSY): A Simple Approach to Enhance Backbone/Sidechain Cross-Peaks in 1H NMR. J Magn Reson 2018, 294, 169–180. https://doi.org/10.1016/j.jmr.2018.07.010. Chen, C.-H.; Wang, Y.; Hilty, C. Intermolecular Interactions Determined by NOE Build-up in Macromolecules from Hyperpolarized Small Molecules. Methods 2018, 138–139, 69–75. https://doi.org/10.1016/j.ymeth.2018.02.015. Wimley, W. C.; Hristova, K.; Ladokhin, A. S.; Silvestro, L.; Axelsen, P. H.; White, S. H. Folding of βSheet Membrane Proteins: A Hydrophobic Hexapeptide Model11Edited by D. C. Rees. Journal of Molecular Biology 1998, 277 (5), 1091–1110. https://doi.org/10.1006/jmbi.1998.1640. White, S. H. Global Statistics of Protein Sequences: Implications for the Origin, Evolution, and Prediction of Structure. Annual Review of Biophysics and Biomolecular Structure 1994, 23 (1), 407–439. https://doi.org/10.1146/annurev.bb.23.060194.002203. White, S. H.; Wimley, W. C.; Ladokhin, A. S.; Hristova, K. Protein Folding in Membranes: Determining Energetics of Peptide-Bilayer Interactions. Meth. Enzymol. 1998, 295, 62–87. Maffei, M.; Arbesú, M.; Le Roux, A.-L.; Amata, I.; Roche, S.; Pons, M. The SH3 Domain Acts as a Scaffold for the N-Terminal Intrinsically Disordered Regions of c-Src. Structure 2015, 23 (5), 893–902. https://doi.org/10.1016/j.str.2015.03.009. Jeener, J. Ampere International Summer School; Basko Polje, Yugoslavia, 1971. Aue, W. P.; Bartholdi, E.; Ernst, R. R. Two‐dimensional Spectroscopy. Application to Nuclear Magnetic Resonance. J. Chem. Phys. 1976, 64 (5), 2229– 2246. https://doi.org/10.1063/1.432450. Vuister, G. W.; Bax, A. Measurement of Four-Bond HN-Hα J-Couplings in Staphylococcal Nuclease. J Biomol NMR 1994, 4 (2), 193–200. https://doi.org/10.1007/BF00175247. Vuister, G. W.; Bax, A. Quantitative J Correlation: A New Approach for Measuring Homonuclear Three-Bond J(HNH.Alpha.) Coupling Constants in 15N-Enriched Proteins. Journal of the American Chemical Society 115 (17), 7772–7777. Emsley, L.; Bodenhausen, G. Selective Two-Dimensional NMR Experiments for Topological Filtration of Fragments of Coupling Networks. Journal of the American Chemical Society 1991, 113 (9), 3309– 3316. https://doi.org/10.1021/ja00009a014.

(33)

(34)

(35)

(36)

Stonehouse, J.; Shaw, G. L.; Keeler, J. Improving Solvent Suppression in Jump-Return NOESY Experiments. J. Biomol. NMR 1994, 4 (6), 799–805. https://doi.org/10.1007/BF00398410. Seelig, J. Thermodynamics of Lipid–Peptide Interactions. Biochimica et Biophysica Acta (BBA) Biomembranes 2004, 1666 (1), 40–50. https://doi.org/10.1016/j.bbamem.2004.08.004. Dyson, H. J.; Wright, P. E. Defining Solution Conformations of Small Linear Peptides. Annual Review of Biophysics and Biophysical Chemistry 1991, 20 (1), 519–538. https://doi.org/10.1146/annurev.bb.20.060191.002511. de Alba, E.; Blanco, F. J.; Jiménez, M. A.; Rico, M.; Nieto, J. L. Interactions Responsible for the PH Dependence of the Beta-Hairpin Conformational Population Formed by a Designed Linear Peptide. Eur. J. Biochem. 1995, 233 (1), 283–292.

Table of Contents



228x82mm (96 x 96 DPI)


Page 14 of 14

Hyperpolarized Water Enhances Two-Dimensional Proton NMR

Recommend Documents