Dynamics and Interactions of a 29 kDa Human Enzyme Studied by

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Dynamics and Interactions of a 29-kDa Human Enzyme Studied by Solid-State NMR Suresh Kumar Vasa, Himanshu Singh, Petra Rovó, and Rasmus Linser J. Phys. Chem. Lett., Just Accepted Manuscript • DOI: 10.1021/acs.jpclett.8b00110 • Publication Date (Web): 26 Feb 2018 Downloaded from http://pubs.acs.org on February 27, 2018

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The Journal of Physical Chemistry Letters

Dynamics and Interactions of a 29-kDa Human Enzyme Studied by Solid-State NMR Suresh K. Vasa[1,2], Himanshu Singh[1,2], Petra Rovó[1,2], Rasmus Linser[1,2]* [1]

Department Chemistry and Pharmacy, Ludwig-Maximilians-University Munich, Butenandtstr. 5-13, 81377, Munich, Germany

[2]

Center for Integrated Protein Science (CiPSM)

Corresponding Author Prof. Dr. Rasmus Linser, Department Chemistry and Pharmacy, Ludwig-Maximilians-University Munich, Butenandtstr. 5-13, 81377, Munich, Germany *E-mail: [email protected].

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ABSTRACT: Solid-state NMR has been employed for characterization of a broad range of biomacromolecules and supramolecular assemblies. However, due to limitations in sensitivity and resolution the size of the individual monomeric units has rarely exceeded 15 kDa. As such, enzymes, which are often more complex and comprised of long peptide chains, have not been easily accessible, even though manifold desirable information could potentially be provided by solid-state NMR studies. Here, we demonstrate that more than 1200 backbone and sidechain chemical shifts can be reliably assessed from minimal sample quantities for a 29 kDa human human enzyme of the carbonic anhydrase family, giving access to its backbone dynamics and intermolecular interactions with a small-molecule inhibitor. The possibility of comprehensive assessment of enzymes in this molecular-weight regime without molecular-tumbling-derived limitations enables the study of residue-specific properties important for their mode of action as well as for pharmacological interference in this and many other enzymes.

TOC GRAPHIC KEYWORDS: solid-state NMR • carbonic anhydrase • fast MAS • protein assignment • protein dynamics • protein NMR


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Solid-state NMR overcomes critical molecular-weight limitations of solution NMR and thus it is amenable for characterization of large proteins or protein assemblies. For example, it has been used extensively for structural elucidation of amyloid proteins, which are poorly accessible for more standard structural-biology techniques.1-6 Similarly, detailed insights for supramolecular assemblies like bacterial needle proteins, virus capsid structures, and structural components of bacteriophages are obtained more and more routinely.7-10 Based on partial protein deuteration and/or fast spinning, the detection of protons has recently been used to increase sensitivity and resolution for residue assignment.11-13 Additional advantages include chemical shifts of protons for resonance assignment,14-17 their high gyromagnetic ratio for proton-proton distance restraints,18-20 and assessment of the amide HN pair for characterization of protein backbone dynamics.21-23 Most of solid-state NMR’s target proteins comprise large polymeric networks, however, apart from some larger exceptions in the 30 kDa range24-26 the monomer size of the structures successfully elucidated has rarely been above 15 kDa. This is due to the fact that the difficulty in assigning proteins with increasing size27 scales steeply with the molecular weight due to challenges from sensitivity, spectral overlap and resolution. Just like solution NMR, solid-state NMR enables atomic-resolution characterization of dynamics and interactions,10,23 such as loop motion or ligand binding, which is fundamental for most kinds of protein functionality and complementary to most other structural-biology techniques.28 However, proteins with more complex biological function, in particular enzymes, usually have much larger (monomer) molecular weight, which challenges the assignment approaches currently established in the field.29 The family of carbonic anhydrases (CA) catalyzes the conversion between dissolved CO2 and bicarbonate. They belong to the fastest enzymes known, reaching up to 1×106 turnovers per


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second for the human variant hCAII.30 Despite good structural understanding and various pharmacological studies, in particular the enzyme’s dynamics is poorly understood on the basis of static X-ray structures. Due to the presence of the His64 sidechain in two different conformations, a flip is assumed to be essential for enzymatic activity. It is, however, unclear, whether general distortions of the cavity are possible, which would facilitate the accommodation of the very different binding partners CO2 and HCO3-. Current scientific interest also lies in protein-water interactions present at the active site (where immobilized “deep water” molecules are thought to exist)31 as well as small-molecule inhibitors acting on the cavity surface properties.32


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Figure 1: Assignments and secondary structure of hCAII, obtained from ca. 1 mg deuterated, 13

C, 15N-labeled, micro-crystalline protein. A) Assigned H/N plane and magnified excerpt for the

crowded central region. B) Secondary chemical shifts33 ((δCߙ obs-δCߙ random) - (δCߚ obs-δCߚ random)) as a function of residue34 and secondary structure propensity according to TALOS-N35 prediction. The derived secondary structure predictions are depicted on top. C) Cartoon representation of magnetization transfer pathways for dipolar HNcacoNH and sidechain-tobackbone (S2B) correlations. D) Representative strips from the 3D S2B correlation for sidechain carbon assignment with composite-pulse MOCCA mixing to involve all types of carbons. E) Sequential plane walk from a 4D NUS HNcacoNH amide-to-amide correlation (blue, overlaid with the HN spectrum, gray). We employ dipolar CC transfers for all experiments, both for CO-


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Cα (BSH transfer36) and for Cα-Cβ (using the HORROR condition) as well as non-uniform sampling. See the SI for details. In A) sc and W sc signify sidechain peaks of Asn/Gln and Trp, respectively. Heteronuclear indirect dimensions employed a 90°-shifted squared sine-bell function. 1H dimensions were processed with a Gaussian line broadening of 5 Hz.

Due to their large molecular size of nearly 30 kDa, CAs are prime examples for proteins in the molecular-weight regime on the edge for solution NMR studies. Even though partial hCAII assignments could be obtained early on, unambiguous residue-specific assessment has so far employed single-amino acid labeling and no solution NMR structure has been solved.37,38 Here, based on a combination of tailored proton-detected experiments applied to a deuterated, 13C- and 15

N-labeled sample, we obtain (almost) complete backbone and sidechain assignment of this 29-

kDa protein by solid-state NMR de novo, which allows access to backbone dynamics, water interactions and mapping of protein:inhibitor contacts from 1 mg of protein. Protein assignments via proton-detected solid-state NMR experiments have so far mostly relied on amide H/N correlations providing pairwise carbon chemical shifts in a third dimension.14,15 However, by exclusively using 3D out-and-back experiments (Figure S2 B-D), we could not even assign half of the protein resonances. For de novo assignment of 29-kDa hCAII, only a strategy involving a set of complementary, conceptually new techniques was successful: 15

N shifts bear the most favorable dispersion for unambiguous assignment, due to a wide

spectral width, homogeneous shift distribution, and narrow linewidths.39 Figure S2E shows strips obtained from a backbone walk based on

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N shifts, implemented by a 3D hNcacoNH

experiment. In this experiment, the inter-residual 15N chemical shift is obtained with a dispersion of the H/N 2D plane. The obtained

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N shifts of hCAII are on the order of 20-50 Hz, with a

close-to-normal distribution over 35 ppm (standard deviation of 6.1 ppm, see SI Figure S4). This


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translates into minimal overlap probability among all backbone nuclei and contributes to a high reliability in the sequential walk. Even more importantly, a non-uniformly-sampled 4D HNcacoNH (see Figure 1C bottom and Figure S1A) gives outstanding resolution and facilitates unambiguous sequential linking of amide peaks. Figure 1E shows a plane walk from this experiment extracted at residue positions D32-T35. Despite the crowded H/N plane of the enzyme, most ambiguity could be resolved. Thirdly, while some residue types can be unambiguously identified based on their Cα and Cβ shifts obtained in hCANH and hcaCBcaNH experiments (for example G, S, T, A), most others remain ambiguous (L, V, I, Q, E, N, D, W, F, Y, K, H, C, R, M). In order to resolve this ambiguity, we developed a backbone-to-sidechain (S2B)40,41 correlation (Figure 1C right and Figure S1B) which employs a broadband-MOCCA mixing (See the SI for details.), drastically expanding the hcaCBcaNH information content to a complete set of carbon shifts and yielding fidelity about each residue type. Figure 1D shows the broadband S2B for hCAII, unambiguously identifying the residue types in representative strips. Importantly, this approach enabled partial aromatic sidechain assignments, which tend to escape the established approaches (see Figure S3).


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Figure 2: Use of backbone assignments for assessment of dynamics using 15N relaxation. A) and B) R1ρ and R1 plotted as a function of primary sequence, respectively. Residues in the binding pocket are marked with turquoise shades. C) Representative R1ρ decays. D) and E) R1ρ and R1 rates mapped onto the X-ray structure (PDB 2CBA), respectively. All dynamics data were recorded via 3D

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N R1- and

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N R1ρ-modulated hCANH experiments. (See SI for complete

experimental details.) Using these strategies, overall 234 (90%) out of 260 residues (including 12 prolines) could be assigned unambiguously. Figure 1A shows an assigned H/N correlation spectrum, Figure 1B (also compare Figure S5) shows the derived secondary structure. A full chemical shift assignment (of more than 1200 shifts) is given in Table S3.


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With the availability of comprehensive backbone and sidechain chemical shifts, studies of protein function become possible. Here, we exploit the newly achieved access to residueresolved enzyme characterization in two ways. Figure 2A and 2B shows a characterization of hCAII backbone dynamics as assessed by 15N R1 and R1ρ data. In contrast to previous dynamics studies on smaller proteins, here crowding in the 2D H/N plane made 3D data indispensible for facilitated readout of relaxation. For this purpose, we engineered a 3D hCANH-based pulse sequence for assessment of

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N coherence lifetimes (see the SI for details). This approach now

allows the application of state-of-the-art relaxation techniques developed recently21-23 to a protein as large as hCAII. R1 relaxation rates, which reflect ps-ns motion, are depicted in Figure 3B and E. R1ρ rates, which are sensitive to µs-ms motion, are depicted in Figure 3A, C, and D. (All decay curves are shown in Figures S6 and S7.) Figure 3C shows the R1ρ decay for W208, which is located in the center of the pocket, in comparison with a flexible loop residue. Interestingly, rates for both types of relaxation are generally low (mostly below 10 s-1 and below 0.1 s-1 for R1ρ and R1, respectively. The only slightly elevated values for (µs-timescale) dynamics are found for the short loops interconnecting the secondary structural elements (e.g., between α5 and β9), which are of no functional relevance. In particular, the residues around the active site, marked with turquoise shades in Figure 2A and B, with R1ρ rates of around 4 s-1, are extremely rigid. Biologically, the activity of CAs is constituted of bringing together an adsorbed CO2 and a Lewis-acid-activated water molecule as the nucleophile the most suitable way. As such, any fluctuations of this spatial geometry would be detrimental for an effective turnover, which would agree with the above findings. In line with this, mutation of the conserved Asn62 in an active-site turn has been found to drastically reduce the turnover rate.42 Various crystal structures show that this Asn forms an


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intramolecular hydrogen bond to the backbone amide of catalytically critical H64, a feature that was not further interpreted yet to our knowledge. The benefit of a strong base for its sidechain supports the above interpretation that rigidity of the active-site backbone scaffold is indeed a conserved property of the hCAII structure for a facilitated mechanism of action.

Given the availability of complete residue-specific chemical shift assignment, we are in the position to use solid-state NMR for localizing the binding site of a small molecule. Figure 3A shows chemical shift perturbations (CSPs) upon binding of the CA inhibitor Dorzolamide, a derivative of acetazolamide, used in eye drop solutions against glaucoma.43 Those residues close to the active site in the crystal structure are marked by turquoise shades. Due to the crowding of the H/N plane, CSPs were again deduced by increased dimensionality using 3D hCANH peak positions (Figure 3B) of holo enzyme and complex and quantified from all three obtained shifts in the form [(∆HN)2 + (∆N/10)2+ (∆Cα/4)2]1/2. Perturbations larger than 0.075 ppm are consistently localized to residues close to the active site (Figure 3C), well in line with the positions in which other inhibitors of CAs bind. In particular, residues like F93, H94, T198, T199, and W208 stick out with CSPs larger than 0.1 ppm. Figure S8 shows a correlation between the experimental CSPs and the (inverse) minimum distance between amide protons to any ligand atoms. Figure S9 compares the residues lining the grove (turquoise shades in Figure 3A) with those residues that CSPs larger than 0.075 ppm are obtained for. The possibility to map binding events and ligand dynamics with residue resolution, a routine procedure already for solution NMR, will have tremendous value for structural biology and medicinal chemistry. Whereas in this study exclusively backbone moieties were employed to detect motion and interactions, use of sidechain protons might be beneficial as more widely available reporters in future studies.


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Figure 3: Interaction of hCAII active-site residues with small-molecule inhibitor Dorzolamide, mapped through 3D-hCANH-encoded chemical shift perturbations (CSP). A) CSPs as a function of sequence. B) Representative hCANH strips of perturbed (green) and an unperturbed residue (blue). C) Mapping of CSPs on the enzyme surface (PDB 2CBA). In A), residues in the grove are marked by turquoise shades, matching very well with the experimental CSPs, which are depicted in yellow for cases larger than 0.075 ppm (gray line). In summary, we show here characterization of atom-resolved biophysical features in the human enzyme carbonic anhydrase II, obtained by solid-state NMR. In particular, assessment of backbone dynamics on different timescales can be achieved by higher-dimensionality relaxation experiments. In addition, we demonstrate that solid-state NMR is able to detect binding to and localize the inhibitor dorzolamide by higher-dimensionality chemical shift perturbations. The work shows that comprehensive backbone and sidechain shift assignments for more than 1200 chemical shifts are possible for proteins in the solid state of large monomeric size. Given the access to chemical shift assignments as the major bottleneck for proteins of increasing size,


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which is obtained by potent higher-dimensionality experiments and mixing sequences developed for this purpose, a whole arsenal of NMR parameters including site-resolved dynamics and interactions becomes accessible for complex proteins. The possibility of assessing proteins in the 30 kDa regime critically expands the range of candidates for ssNMR investigations and sets the stage for diverse biological studies on other similarly complex proteins without the tumblingderived limitations in solution. In particular, the possibility to use ssNMR for characterization of enzymes will be of great value for understanding of catalysis mechanisms, substrate handling, reaction intermediates, and thermodynamic features of these processes.

ASSOCIATED CONTENT Supporting Information. Sample preparation, pulse sequences, experimental parameters, chemical shift tables, 15N relaxation data. AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Notes S. K. Vasa and H. Singh contributed equally to the work. The authors declare no competing financial interests. ACKNOWLEDGMENT


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The authors acknowledge the Gerhard Klebe group (Uni Marburg) for the plasmid, protocols, and helpful discussions as well as Karin Giller, Dr. Stefan Becker, and Eszter Najbauer (MPIbpc Göttingen) for initial efforts in this project. Financial support is acknowledged from the Deutsche Forschungsgemeinschaft (SFB 749, TP A13, as well as the Emmy Noether program), the Verband der Chemischen Industrie (VCI) in terms of a Liebig junior group fellowship, the Excellence Cluster CiPS-M, and the Center for NanoScience (CeNS). REFERENCES 1. Jones, E. M.; Wu, B.; Surewicz, K.; Nadaud, P. S.; Helmus, J. J.; Chen, S.; Jaroniec, C. P.; Surewicz, W. K. Structural Polymorphism in Amyloids: New Insights from Studies with Y145Stop Prion Protein Fibrils J. Biol. Chem. 2011, 286, 42777-42784. 2. Lu, J. X.; Qiang, W.; Yau, W. M.; Schwieters, C. D.; Meredith, S. C.; Tycko, R. Molecular Structure of β-Amyloid Fibrils in Alzheimer’s Disease Brain Tissue Cell 2013, 154, 1257–1268. 3. Xiao, Y.; Ma, B.; McElheny, D.; Parthasarathy, S.; Long, F.; Hoshi, M.; Nussinov, R.; Ishii, Y. A[beta](1-42) Fibril Structure Illuminates Self-Recognition and Replication of Amyloid in Alzheimer's Disease Nat. Struct. Mol. Biol. 2015, 22, 499-505. 4. Wälti, M. A.; Ravotti, F.; Arai, H.; Glabe, C. G.; Wall, J. S.; Böckmann, A.; Güntert, P.; Meier, B. H.; Riek, R. Atomic-Resolution Structure of a Disease-Relevant Aβ(1–42) Amyloid Fibril Proc. Natl. Acad. Sci. U.S.A. 2016, 113, E4976-E4984. 5. Colvin, M. T.; Silvers, R.; Ni, Q. Z.; Can, T. V.; Sergeyev, I.; Rosay, M.; Donovan, K. J.; Michael, B.; Wall, J.; Linse, S.; Griffin, R. G. Atomic Resolution Structure of Monomorphic Aβ42 Amyloid Fibrils J. Am. Chem. Soc. 2016, 138, 9663-9674. 6. Tuttle, M. D.; Comellas, G.; Nieuwkoop, A. J.; Covell, D. J.; Berthold, D. A.; Kloepper, K. D.; Courtney, J. M.; Kim, J. K.; Barclay, A. M.; Kendall, A.; Wan, W.; Stubbs, G.; Schwieters, C. D.; Lee, V. M. Y.; George, J. M.; Rienstra, C. M. Solid-State NMR Structure of a Pathogenic Fibril of Full-Length Human α-Synuclein Nat. Struct. Mol. Biol. 2016, 23, 409–415. 7. Loquet, A.; Sgourakis, N. G.; Gupta, R.; Giller, K.; Riedel, D.; Goosmann, C.; Griesinger, C.; Kolbe, M.; Baker, D.; Becker, S.; Lange, A. Atomic Model of the Type III Secretion System Needle Nature 2012, 486, 276-279. 8. Sergeyev, I. V.; Bahri, S.; Day, L. A.; McDermott, A. E. Pf1 Bacteriophage Hydration by Magic Angle Spinning Solid-State NMR J. Chem. Phys. 2014, 141, 22D533. 9. Morag, O.; Sgourakis, N. G.; Baker, D.; Goldbourt, A. The NMR-Rosetta Capsid Model of M13 Bacteriophage Reveals a Quadrupled Hydrophobic Packing Epitope Proc. Natl. Acad. Sci. U.S.A. 2015, 112, 971-976. 10. Zhang, H.; Hou, G.; Lu, M.; Ahn, J.; Byeon, I.-J. L.; Langmead, C. J.; Perilla, J. R.; Hung, I.; Gor’kov, P. L.; Gan, Z.; Brey, W. W.; Case, D. A.; Schulten, K.; Gronenborn, A. M.; Polenova, T. HIV Capsid Function is Regulated by Dynamics: Quantitative Atomic-Resolution Insights by Integrating Magic-Angle-Spinning NMR, QM/MM, and MD. J. Am. Chem. Soc. 2016.


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