1
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De Meutter J, Goormaghtigh E. Protein Microarrays for High Throughput Hydrogen/Deuterium Exchange Monitored by FTIR Imaging. Int J Mol Sci 2024; 25:9989. [PMID: 39337477 PMCID: PMC11432650 DOI: 10.3390/ijms25189989] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/13/2024] [Revised: 09/11/2024] [Accepted: 09/12/2024] [Indexed: 09/30/2024] Open
Abstract
Proteins form the fastest-growing therapeutic class. Due to their intrinsic instability, loss of native structure is common. Structure alteration must be carefully evaluated as structural changes may jeopardize the efficiency and safety of the protein-based drugs. Hydrogen deuterium exchange (HDX) has long been used to evaluate protein structure and dynamics. The rate of exchange constitutes a sensitive marker of the conformational state of the protein and of its stability. It is often monitored by mass spectrometry. Fourier transform infrared (FTIR) spectroscopy is another method with very promising capabilities. Combining protein microarrays with FTIR imaging resulted in high throughput HDX FTIR measurements. BaF2 slides bearing the protein microarrays were covered by another slide separated by a spacer, allowing us to flush the cell continuously with a flow of N2 gas saturated with 2H2O. Exchange occurred simultaneously for all proteins and single images covering ca. 96 spots of proteins that could be recorded on-line at selected time points. Each protein spot contained ca. 5 ng protein, and the entire array covered 2.5 × 2.5 mm2. Furthermore, HDX could be monitored in real time, and the experiment was therefore not subject to back-exchange problems. Analysis of HDX curves by inverse Laplace transform and by fitting exponential curves indicated that quantitative comparison of the samples is feasible. The paper also demonstrates how the whole process of analysis can be automatized to yield fast analyses.
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Affiliation(s)
- Joëlle De Meutter
- Center for Structural Biology and Bioinformatics, Laboratory for the Structure and Function of Biological Membranes, Campus Plaine, Université Libre de Bruxelles CP206/2, B1050 Brussels, Belgium
| | - Erik Goormaghtigh
- Center for Structural Biology and Bioinformatics, Laboratory for the Structure and Function of Biological Membranes, Campus Plaine, Université Libre de Bruxelles CP206/2, B1050 Brussels, Belgium
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2
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Eisinger M, Rahn H, Chen Y, Fernandes M, Lin Z, Hentze N, Tavella D, Moussa EM. Elucidation of the Reversible Self-Association Interface of a Diabody-Interleukin Fusion Protein Using Hydrogen-Exchange Mass Spectrometry and In Silico Modeling. Mol Pharm 2024; 21:4285-4296. [PMID: 38922328 DOI: 10.1021/acs.molpharmaceut.4c00169] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/27/2024]
Abstract
Reversible self-association (RSA) of therapeutic proteins presents major challenges in the development of high-concentration formulations, especially those intended for subcutaneous administration. Understanding self-association mechanisms is therefore critical to the design and selection of candidates with acceptable developability to advance to clinical trials. The combination of experiments and in silico modeling presents a powerful tool to elucidate the interface of self-association. RSA of monoclonal antibodies has been studied extensively under different solution conditions and have been shown to involve interactions for both the antigen-binding fragment and the crystallizable fragment. Novel modalities such as bispecific antibodies, antigen-binding fragments, single-chain-variable fragments, and diabodies constitute a fast-growing class of antibody-based therapeutics that have unique physiochemical properties compared to monoclonal antibodies. In this study, the RSA interface of a diabody-interleukin 22 fusion protein (FP-1) was studied using hydrogen-deuterium exchange coupled with mass spectrometry (HDX-MS) in combination with in silico modeling. Taken together, the results show that a complex solution behavior underlies the self-association of FP-1 and that the interface thereof can be attributed to a specific segment in the variable light chain of the diabody. These findings also demonstrate that the combination of HDX-MS with in silico modeling is a powerful tool to guide the design and candidate selection of novel biotherapeutic modalities.
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Affiliation(s)
- Martin Eisinger
- Biologics Analytical Research and Development, AbbVie Deutschland GmbH & Co. KG, Ludwigshafen 67061, Germany
| | - Harri Rahn
- Biologics Analytical Research and Development, AbbVie Deutschland GmbH & Co. KG, Ludwigshafen 67061, Germany
| | - Yong Chen
- Biologics Analytical Research and Development, AbbVie Inc., North Chicago, Illinois 60061, United States
| | - Melissa Fernandes
- Biologics Drug Product Development, AbbVie Inc., North Chicago, Illinois 60061, United States
| | - Zhiyi Lin
- Biologics Drug Product Development, AbbVie Inc., North Chicago, Illinois 60061, United States
| | - Nikolai Hentze
- Biologics Analytical Research and Development, AbbVie Deutschland GmbH & Co. KG, Ludwigshafen 67061, Germany
| | - Davide Tavella
- Biotherapeutics and Genetic Medicine, AbbVie Inc., Worcester, Massachusetts 01604, United States
| | - Ehab M Moussa
- Biologics Drug Product Development, AbbVie Inc., North Chicago, Illinois 60061, United States
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3
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Wood V, Kellerman MA, Groves K, Quaglia M, Topp EM, Matejtschuk P, Dalby PA. Investigation of the Solid-State Interactions in Lyophilized Human G-CSF Using Hydrogen-Deuterium Exchange Mass Spectrometry. Mol Pharm 2024; 21:1965-1976. [PMID: 38516985 PMCID: PMC10988552 DOI: 10.1021/acs.molpharmaceut.3c01211] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/18/2023] [Revised: 03/13/2024] [Accepted: 03/13/2024] [Indexed: 03/23/2024]
Abstract
Hydrogen/deuterium exchange mass spectrometry (HDX-MS) previously elucidated the interactions between excipients and proteins for liquid granulocyte colony stimulating factor (G-CSF) formulations, confirming predictions made using computational structure docking. More recently, solid-state HDX mass spectrometry (ssHDX-MS) was developed for proteins in the lyophilized state. Deuterium uptake in ssHDX-MS has been shown for various proteins, including monoclonal antibodies, to be highly correlated with storage stability, as measured by protein aggregation and chemical degradation. As G-CSF is known to lose activity through aggregation upon lyophilization, we applied the ssHDX-MS method with peptide mapping to four different lyophilized formulations of G-CSF to compare the impact of three excipients on local structure and exchange dynamics. HDX at 22 °C was confirmed to correlate well with the monomer content remaining after lyophilization and storage at -20 °C, with sucrose providing the greatest protection, and then phenylalanine, mannitol, and no excipient leading to progressively less protection. Storage at 45 °C led to little difference in final monomer content among the formulations, and so there was no discernible relationship with total deuterium uptake on ssHDX. Incubation at 45 °C may have led to a structural conformation and/or aggregation mechanism no longer probed by HDX at 22 °C. Such a conformational change was observed previously at 37 °C for liquid-formulated G-CSF using NMR. Peptide mapping revealed that tolerance to lyophilization and -20 °C storage was linked to increased stability in the small helix, loop AB, helix C, and loop CD. LC-MS HDX and NMR had previously linked loop AB and loop CD to the formation of a native-like state (N*) prior to aggregation in liquid formulations, suggesting a similar structural basis for G-CSF aggregation in the liquid and solid states.
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Affiliation(s)
- Victoria
E. Wood
- Department
of Biochemical Engineering, University College
London, London WC1E 6BT, United
Kingdom
| | - Mark-Adam Kellerman
- Department
of Biochemical Engineering, University College
London, London WC1E 6BT, United
Kingdom
| | - Kate Groves
- LGC, Queens Road, Teddington, Middlesex TQ11 0LY, United Kingdom
| | - Milena Quaglia
- LGC, Queens Road, Teddington, Middlesex TQ11 0LY, United Kingdom
| | - Elizabeth M. Topp
- Department
of Industrial and Molecular Pharmaceutics, College of Pharmacy, and
Davidson School of Chemical Engineering, College of Engineering Purdue University, West Lafayette, Indiana 47907, United States
| | - Paul Matejtschuk
- Standardisation
Science, NIBSC, Medicines & Healthcare
Products Regulatory Agency, South Mimms, Hertfordshire EN6 3QG, United
Kingdom
| | - Paul A. Dalby
- Department
of Biochemical Engineering, University College
London, London WC1E 6BT, United
Kingdom
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4
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Haidar Y, Konermann L. Effects of Hydrogen/Deuterium Exchange on Protein Stability in Solution and in the Gas Phase. JOURNAL OF THE AMERICAN SOCIETY FOR MASS SPECTROMETRY 2023. [PMID: 37314114 DOI: 10.1021/jasms.3c00130] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/15/2023]
Abstract
Mass spectrometry (MS)-based techniques are widely used for probing protein structure and dynamics in solution. H/D exchange (HDX)-MS is one of the most common approaches in this context. HDX is often considered to be a "benign" labeling method, in that it does not perturb protein behavior in solution. However, several studies have reported that D2O pushes unfolding equilibria toward the native state. The origin, and even the existence of this protein stabilization remain controversial. Here we conducted thermal unfolding assays in solution to confirm that deuterated proteins in D2O are more stable, with 2-4 K higher melting temperatures than unlabeled proteins in H2O. Previous studies tentatively attributed this phenomenon to strengthened H-bonds after deuteration, an effect that may arise from the lower zero-point vibrational energy of the deuterated species. Specifically, it was proposed that strengthened water-water bonds (W···W) in D2O lower the solubility of nonpolar side chains. The current work takes a broader view by noting that protein stability in solution also depends on water-protein (W···P) and protein-protein (P···P) H-bonds. To help unravel these contributions, we performed collision-induced unfolding (CIU) experiments on gaseous proteins generated by native electrospray ionization. CIU profiles of deuterated and unlabeled proteins were indistinguishable, implying that P···P contacts are insensitive to deuteration. Thus, protein stabilization in D2O is attributable to solvent effects, rather than alterations of intraprotein H-bonds. Strengthening of W···W contacts represents one possible explanation, but the stabilizing effect of D2O can also originate from weakened W···P bonds. Future work will be required to elucidate which of these two scenarios is correct, or if both contribute to protein stabilization in D2O. In any case, the often-repeated adage that "D-bonds are more stable than H-bonds" does not apply to intramolecular contacts in native proteins.
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Affiliation(s)
- Yousef Haidar
- Department of Chemistry, The University of Western Ontario, London, Ontario N6A 5B7, Canada
| | - Lars Konermann
- Department of Chemistry, The University of Western Ontario, London, Ontario N6A 5B7, Canada
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5
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Jethva PN, Gross ML. Hydrogen Deuterium Exchange and other Mass Spectrometry-based Approaches for Epitope Mapping. FRONTIERS IN ANALYTICAL SCIENCE 2023; 3:1118749. [PMID: 37746528 PMCID: PMC10512744 DOI: 10.3389/frans.2023.1118749] [Citation(s) in RCA: 7] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 09/26/2023]
Abstract
Antigen-antibody interactions are a fundamental subset of protein-protein interactions responsible for the "survival of the fittest". Determining the interacting interface of the antigen, called an epitope, and that on the antibody, called a paratope, is crucial to antibody development. Because each antigen presents multiple epitopes (unique footprints), sophisticated approaches are required to determine the target region for a given antibody. Although X-ray crystallography, Cryo-EM, and nuclear magnetic resonance can provide atomic details of an epitope, they are often laborious, poor in throughput, and insensitive. Mass spectrometry-based approaches offer rapid turnaround, intermediate structural resolution, and virtually no size limit for the antigen, making them a vital approach for epitope mapping. In this review, we describe in detail the principles of hydrogen deuterium exchange mass spectrometry in application to epitope mapping. We also show that a combination of MS-based approaches can assist or complement epitope mapping and push the limit of structural resolution to the residue level. We describe in detail the MS methods used in epitope mapping, provide our perspective about the approaches, and focus on elucidating the role that HDX-MS is playing now and in the future by organizing a discussion centered around several improvements in prototype instrument/applications used for epitope mapping. At the end, we provide a tabular summary of the current literature on HDX-MS-based epitope mapping.
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Affiliation(s)
- Prashant N. Jethva
- Department of Chemistry, Washington University in St. Louis, St Louis, MO 63130, USA
| | - Michael L. Gross
- Department of Chemistry, Washington University in St. Louis, St Louis, MO 63130, USA
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6
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Thibodeau MC, Harris NJ, Jenkins ML, Parson MAH, Evans JT, Scott MK, Shaw AL, Pokorný D, Leonard TA, Burke JE. Molecular basis for the recruitment of the Rab effector protein WDR44 by the GTPase Rab11. J Biol Chem 2023; 299:102764. [PMID: 36463963 PMCID: PMC9808001 DOI: 10.1016/j.jbc.2022.102764] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/08/2022] [Revised: 11/22/2022] [Accepted: 11/28/2022] [Indexed: 12/03/2022] Open
Abstract
The formation of complexes between Rab11 and its effectors regulates multiple aspects of membrane trafficking, including recycling and ciliogenesis. WD repeat-containing protein 44 (WDR44) is a structurally uncharacterized Rab11 effector that regulates ciliogenesis by competing with prociliogenesis factors for Rab11 binding. Here, we present a detailed biochemical and biophysical characterization of the WDR44-Rab11 complex and define specific residues mediating binding. Using AlphaFold2 modeling and hydrogen/deuterium exchange mass spectrometry, we generated a molecular model of the Rab11-WDR44 complex. The Rab11-binding domain of WDR44 interacts with switch I, switch II, and the interswitch region of Rab11. Extensive mutagenesis of evolutionarily conserved residues in WDR44 at the interface identified numerous complex-disrupting mutations. Using hydrogen/deuterium exchange mass spectrometry, we found that the dynamics of the WDR44-Rab11 interface are distinct from the Rab11 effector FIP3, with WDR44 forming a more extensive interface with the switch II helix of Rab11 compared with FIP3. The WDR44 interaction was specific to Rab11 over evolutionarily similar Rabs, with mutations defining the molecular basis of Rab11 specificity. Finally, WDR44 can be phosphorylated by Sgk3, with this leading to reorganization of the Rab11-binding surface on WDR44. Overall, our results provide molecular detail on how WDR44 interacts with Rab11 and how Rab11 can form distinct effector complexes that regulate membrane trafficking events.
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Affiliation(s)
- Matthew C Thibodeau
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - Noah J Harris
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - Meredith L Jenkins
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - Matthew A H Parson
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - John T Evans
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - Mackenzie K Scott
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada
| | - Alexandria L Shaw
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada; Department of Biochemistry and Molecular Biology, The University of British Columbia, Vancouver, British Columbia, Canada
| | - Daniel Pokorný
- Max Perutz Labs, Department of Structural and Computational Biology, Vienna, Austria; Department of Medical Biochemistry, Medical University of Vienna, Vienna, Austria
| | - Thomas A Leonard
- Max Perutz Labs, Department of Structural and Computational Biology, Vienna, Austria; Department of Medical Biochemistry, Medical University of Vienna, Vienna, Austria
| | - John E Burke
- Department of Biochemistry and Microbiology, University of Victoria, Victoria, British Columbia, Canada; Department of Biochemistry and Molecular Biology, The University of British Columbia, Vancouver, British Columbia, Canada.
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7
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Investigating how intrinsically disordered regions contribute to protein function using HDX-MS. Biochem Soc Trans 2022; 50:1607-1617. [DOI: 10.1042/bst20220206] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/04/2022] [Revised: 11/11/2022] [Accepted: 11/14/2022] [Indexed: 12/02/2022]
Abstract
A large amount of the human proteome is composed of highly dynamic regions that do not adopt a single static conformation. These regions are defined as intrinsically disordered, and they are found in a third of all eukaryotic proteins. They play instrumental roles in many aspects of protein signaling, but can be challenging to characterize by biophysical methods. Intriguingly, many of these regions can adopt stable secondary structure upon interaction with a variety of binding partners, including proteins, lipids, and ligands. This review will discuss the application of Hydrogen-deuterium exchange mass spectrometry (HDX-MS) as a powerful biophysical tool that is particularly well suited for structural and functional characterization of intrinsically disordered regions in proteins. A focus will be on the theory of hydrogen exchange, and its practical application to identify disordered regions, as well as characterize how they participate in protein–protein and protein–membrane interfaces. A particular emphasis will be on how HDX-MS data can be presented specifically tailored for analysis of intrinsically disordered regions, as well as the technical aspects that are critical to consider when designing HDX-MS experiments for proteins containing intrinsically disordered regions.
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8
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Tajoddin NN, Konermann L. Structural Dynamics of a Thermally Stressed Monoclonal Antibody Characterized by Temperature-Dependent H/D Exchange Mass Spectrometry. Anal Chem 2022; 94:15499-15509. [DOI: 10.1021/acs.analchem.2c03931] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/30/2022]
Affiliation(s)
- Nastaran N. Tajoddin
- Department of Chemistry, The University of Western Ontario, London, Ontario N6A 5B7, Canada
| | - Lars Konermann
- Department of Chemistry, The University of Western Ontario, London, Ontario N6A 5B7, Canada
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9
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ATR-FTIR Biosensors for Antibody Detection and Analysis. Int J Mol Sci 2022; 23:ijms231911895. [PMID: 36233197 PMCID: PMC9570191 DOI: 10.3390/ijms231911895] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/04/2022] [Revised: 09/29/2022] [Accepted: 10/04/2022] [Indexed: 11/17/2022] Open
Abstract
Quality control of drug products is of paramount importance in the pharmaceutical world. It ensures product safety, efficiency, and consistency. In the case of complex biomolecules such as therapeutic proteins, small variations in bioprocess parameters can induce substantial variations in terms of structure, impacting the drug product quality. Conditions for obtaining highly reproducible grafting of 11-mercaptoundecanoic acid were determined. On that basis, we developed an easy-to-use, cost effective, and timesaving biosensor based on ATR-FTIR spectroscopy able to detect immunoglobulins during their production. A germanium crystal, used as an internal reflection element (IRE) for FTIR spectroscopy, was covalently coated with immunoglobulin-binding proteins. This thereby functionalized surface could bind only immunoglobulins present in complex media such as culture media or biopharmaceutical products. The potential subsequent analysis of their structure by ATR-FTIR spectroscopy makes this biosensor a powerful tool to monitor the production of biotherapeutics and assess important critical quality attributes (CQAs) such as high-order structure and aggregation level.
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10
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Bondarchuk TV, Shalak VF, Lozhko DM, Fatalska A, Szczepanowski R, Liudkovska V, Tsuvariev O, Dadlez M, El'skaya A, Negrutskii B. Quaternary organization of the human eEF1B complex reveals unique multi-GEF domain assembly. Nucleic Acids Res 2022; 50:9490-9504. [PMID: 35971611 PMCID: PMC9458455 DOI: 10.1093/nar/gkac685] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/25/2022] [Revised: 07/12/2022] [Accepted: 07/31/2022] [Indexed: 12/24/2022] Open
Abstract
Protein synthesis in eukaryotic cell is spatially and structurally compartmentalized that ensures high efficiency of this process. One of the distinctive features of higher eukaryotes is the existence of stable multi-protein complexes of aminoacyl-tRNA synthetases and translation elongation factors. Here, we report a quaternary organization of the human guanine-nucleotide exchange factor (GEF) complex, eEF1B, comprising α, β and γ subunits that specifically associate into a heterotrimeric form eEF1B(αβγ)3. As both the eEF1Bα and eEF1Bβ proteins have structurally conserved GEF domains, their total number within the complex is equal to six. Such, so far, unique structural assembly of the guanine-nucleotide exchange factors within a stable complex may be considered as a 'GEF hub' that ensures efficient maintenance of the translationally active GTP-bound conformation of eEF1A in higher eukaryotes.
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Affiliation(s)
- Tetiana V Bondarchuk
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
| | - Vyacheslav F Shalak
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
| | - Dmytro M Lozhko
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
| | - Agnieszka Fatalska
- Institute of Biochemistry and Biophysics, PAN, Pawinskiego 5a, 02-109 Warsaw, Poland
- Department of Genetics, University of Cambridge, Cambridge CB2 3EH, UK
| | - Roman H Szczepanowski
- International Institute of Molecular and Cell Biology, Trojdena 4, 02-109 Warsaw, Poland
| | - Vladyslava Liudkovska
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
- International Institute of Molecular and Cell Biology, Trojdena 4, 02-109 Warsaw, Poland
| | - Oleksandr Yu Tsuvariev
- Institute of High Technologies, Taras Shevchenko National University of Kyiv, Akademik Glushkov Ave. 4-g, 03022 Kyiv, Ukraine
| | - Michal Dadlez
- Institute of Biochemistry and Biophysics, PAN, Pawinskiego 5a, 02-109 Warsaw, Poland
| | - Anna V El'skaya
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
| | - Boris S Negrutskii
- Institute of Molecular Biology and Genetics, NAS of Ukraine, 150 Zabolotnogo St., 03143 Kyiv, Ukraine
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11
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Huang J, Chu X, Luo Y, Wang Y, Zhang Y, Zhang Y, Li H. Insights into Phosphorylation-Induced Protein Allostery and Conformational Dynamics of Glycogen Phosphorylase via Integrative Structural Mass Spectrometry and In Silico Modeling. ACS Chem Biol 2022; 17:1951-1962. [PMID: 35675581 DOI: 10.1021/acschembio.2c00393] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/28/2022]
Abstract
Allosteric regulation plays a fundamental role in innumerable biological processes. Understanding its dynamic mechanism and impact at the molecular level is of great importance in disease diagnosis and drug discovery. Glycogen phosphorylase (GP) is a phosphoprotein responding to allosteric regulation and has significant biological importance to glycogen metabolism. Although the atomic structures of GP have been previously solved, the conformational dynamics of GP related to allostery regulation remain largely elusive due to its macromolecular size (∼196 kDa). Here, we integrated native top-down mass spectrometry (nTD-MS), hydrogen-deuterium exchange MS (HDX-MS), protection factor (PF) analysis, molecular dynamics (MD) simulations, and allostery signaling analysis to examine the structural basis and dynamics for the allosteric regulation of GP by phosphorylation. nTD-MS reveals differences in structural stability as well as oligomeric state between the unphosphorylated (GPb) and phosphorylated (GPa) forms. HDX-MS, PF analysis, and MD simulations further pinpoint the structural differences between GPb and GPa involving the binding interfaces (the N-terminal and tower-tower helices), catalytic site, and PLP-binding region. More importantly, it also allowed us to complete the missing link of the long-range communication process from the N-terminal tail to the catalytic site caused by phosphorylation. This integrative MS and in silico-based platform is highly complementary to biophysical approaches and yields valuable insights into protein structures and dynamic regulation.
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Affiliation(s)
- Jing Huang
- School of Pharmaceutical Sciences, Sun Yat-sen University, Guangzhou Higher Education Mega Center, No. 132 Wai Huan Dong Lu, Guangzhou 510006, China
| | - Xiakun Chu
- Advanced Materials Thrust, The Hong Kong University of Science and Technology (Guangzhou), Nansha, Guangzhou, Guangdong 511400, China
| | - Yuxiang Luo
- School of Pharmaceutical Sciences, Sun Yat-sen University, Guangzhou Higher Education Mega Center, No. 132 Wai Huan Dong Lu, Guangzhou 510006, China
| | - Yong Wang
- The Provincial International Science and Technology Cooperation Base on Engineering Biology, International Campus of Zhejiang University, College of Life Sciences, Shanghai Institute for Advanced Study, Institute of Quantitative Biology, Zhejiang University, Haining 314400, China
| | - Ying Zhang
- School of Pharmaceutical Sciences, Sun Yat-sen University, Guangzhou Higher Education Mega Center, No. 132 Wai Huan Dong Lu, Guangzhou 510006, China
| | - Yu Zhang
- State Key Laboratory of Crop Stress Adaptation and Improvement, Key Laboratory of Plant Stress Biology, School of Life Sciences, Henan University, Kaifeng 475004, China
| | - Huilin Li
- School of Pharmaceutical Sciences, Sun Yat-sen University, Guangzhou Higher Education Mega Center, No. 132 Wai Huan Dong Lu, Guangzhou 510006, China.,Guangdong Key Laboratory of Chiral Molecule and Drug Discovery, School of Pharmaceutical Sciences, Sun Yat-sen University, Guangzhou, Guangdong 510006, China
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12
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Correy GJ, Kneller DW, Phillips G, Pant S, Russi S, Cohen AE, Meigs G, Holton JM, Gahbauer S, Thompson MC, Ashworth A, Coates L, Kovalevsky A, Meilleur F, Fraser JS. The mechanisms of catalysis and ligand binding for the SARS-CoV-2 NSP3 macrodomain from neutron and x-ray diffraction at room temperature. SCIENCE ADVANCES 2022; 8:eabo5083. [PMID: 35622909 PMCID: PMC9140965 DOI: 10.1126/sciadv.abo5083] [Citation(s) in RCA: 22] [Impact Index Per Article: 11.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/08/2022] [Accepted: 04/11/2022] [Indexed: 05/04/2023]
Abstract
The nonstructural protein 3 (NSP3) macrodomain of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Mac1) removes adenosine diphosphate (ADP) ribosylation posttranslational modifications, playing a key role in the immune evasion capabilities of the virus responsible for the coronavirus disease 2019 pandemic. Here, we determined neutron and x-ray crystal structures of the SARS-CoV-2 NSP3 macrodomain using multiple crystal forms, temperatures, and pHs, across the apo and ADP-ribose-bound states. We characterize extensive solvation in the Mac1 active site and visualize how water networks reorganize upon binding of ADP-ribose and non-native ligands, inspiring strategies for displacing waters to increase the potency of Mac1 inhibitors. Determining the precise orientations of active site water molecules and the protonation states of key catalytic site residues by neutron crystallography suggests a catalytic mechanism for coronavirus macrodomains distinct from the substrate-assisted mechanism proposed for human MacroD2. These data provoke a reevaluation of macrodomain catalytic mechanisms and will guide the optimization of Mac1 inhibitors.
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Affiliation(s)
- Galen J. Correy
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA 94158, USA
| | - Daniel W. Kneller
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, U.S. Department of Energy, Washington, DC 20585, USA
| | - Gwyndalyn Phillips
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, U.S. Department of Energy, Washington, DC 20585, USA
| | - Swati Pant
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, U.S. Department of Energy, Washington, DC 20585, USA
| | - Silvia Russi
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, CA 94025, USA
| | - Aina E. Cohen
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, CA 94025, USA
| | - George Meigs
- Department of Molecular Biophysics and Integrated Bioimaging, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA 94158, USA
| | - James M. Holton
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, Menlo Park, CA 94025, USA
- Department of Molecular Biophysics and Integrated Bioimaging, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
- Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA 94158, USA
| | - Stefan Gahbauer
- Department of Pharmaceutical Chemistry, University of California, San Francisco, San Francisco, CA 94158, USA
| | - Michael C. Thompson
- Department of Chemistry and Biochemistry, University of California, Merced, Merced, CA 95343, USA
| | - Alan Ashworth
- Helen Diller Family Comprehensive Cancer, University of California, San Francisco, San Francisco, CA 94158, USA
| | - Leighton Coates
- National Virtual Biotechnology Laboratory, U.S. Department of Energy, Washington, DC 20585, USA
- Second Target Station, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
| | - Andrey Kovalevsky
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, U.S. Department of Energy, Washington, DC 20585, USA
| | - Flora Meilleur
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- Department of Molecular and Structural Biochemistry, North Carolina State University, Raleigh, NC 27695, USA
| | - James S. Fraser
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA 94158, USA
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13
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Laurents DV. AlphaFold 2 and NMR Spectroscopy: Partners to Understand Protein Structure, Dynamics and Function. Front Mol Biosci 2022; 9:906437. [PMID: 35655760 PMCID: PMC9152297 DOI: 10.3389/fmolb.2022.906437] [Citation(s) in RCA: 25] [Impact Index Per Article: 12.5] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/28/2022] [Accepted: 04/25/2022] [Indexed: 11/29/2022] Open
Abstract
The artificial intelligence program AlphaFold 2 is revolutionizing the field of protein structure determination as it accurately predicts the 3D structure of two thirds of the human proteome. Its predictions can be used directly as structural models or indirectly as aids for experimental structure determination using X-ray crystallography, CryoEM or NMR spectroscopy. Nevertheless, AlphaFold 2 can neither afford insight into how proteins fold, nor can it determine protein stability or dynamics. Rare folds or minor alternative conformations are also not predicted by AlphaFold 2 and the program does not forecast the impact of post translational modifications, mutations or ligand binding. The remaining third of human proteome which is poorly predicted largely corresponds to intrinsically disordered regions of proteins. Key to regulation and signaling networks, these disordered regions often form biomolecular condensates or amyloids. Fortunately, the limitations of AlphaFold 2 are largely complemented by NMR spectroscopy. This experimental approach provides information on protein folding and dynamics as well as biomolecular condensates and amyloids and their modulation by experimental conditions, small molecules, post translational modifications, mutations, flanking sequence, interactions with other proteins, RNA and virus. Together, NMR spectroscopy and AlphaFold 2 can collaborate to advance our comprehension of proteins.
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14
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The dopamine transporter antiports potassium to increase the uptake of dopamine. Nat Commun 2022; 13:2446. [PMID: 35508541 PMCID: PMC9068915 DOI: 10.1038/s41467-022-30154-5] [Citation(s) in RCA: 10] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/19/2021] [Accepted: 04/18/2022] [Indexed: 12/11/2022] Open
Abstract
The dopamine transporter facilitates dopamine reuptake from the extracellular space to terminate neurotransmission. The transporter belongs to the neurotransmitter:sodium symporter family, which includes transporters for serotonin, norepinephrine, and GABA that utilize the Na+ gradient to drive the uptake of substrate. Decades ago, it was shown that the serotonin transporter also antiports K+, but investigations of K+-coupled transport in other neurotransmitter:sodium symporters have been inconclusive. Here, we show that ligand binding to the Drosophila- and human dopamine transporters are inhibited by K+, and the conformational dynamics of the Drosophila dopamine transporter in K+ are divergent from the apo- and Na+-states. Furthermore, we find that K+ increases dopamine uptake by the Drosophila dopamine transporter in liposomes, and visualize Na+ and K+ fluxes in single proteoliposomes using fluorescent ion indicators. Our results expand on the fundamentals of dopamine transport and prompt a reevaluation of the impact of K+ on other transporters in this pharmacologically important family. The dopamine transporter, DAT, controls dopamine signaling by facilitating its reuptake using the Na+ gradient as driving force. Here, the authors uncover that an antiport of K+ ions also contributes to setting the rate of DAT-mediated dopamine clearance.
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15
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Yılmaz İ, Korkmaz F. Investigations of pH-dependent dynamic properties of OmpG-16SL, an outer membrane protein G mutant by ATR-FTIR spectroscopy. BIOCHIMICA ET BIOPHYSICA ACTA. PROTEINS AND PROTEOMICS 2022; 1870:140780. [PMID: 35405324 DOI: 10.1016/j.bbapap.2022.140780] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 01/18/2022] [Revised: 04/03/2022] [Accepted: 04/05/2022] [Indexed: 06/14/2023]
Abstract
In this paper, the dynamic properties of outer membrane protein G mutant (OmpG-16SL) are investigated with ATR-FTIR spectroscopy. While OmpG-WT has 14 β-strands in its structure, the mutant is designed to have 16 β-strands with the intention of creating an enlarged pore. Loop L6 is elongated by introducing six residues, two of which are negatively charged. The solvent accessibility of the OmpG-16SL mutant is compared with WT and a previously reported mutant OmpG-16S by tracking the 1H/2H exchange kinetics in acidic and neutral buffer conditions. The exchange kinetics and dynamics in the fast and slow exchange phases are separately investigated using the 2DCOS technique, which enables the tracking of the structural changes at each phase of the exchange process. The results suggest that the mutant OmpG-16SL is equally exposed to buffer in both acidic and neutral pH conditions. Additionally, the time range in the fast phase is very short - one-tenth of that for WT - and most of the exchange is completed in this phase. This fast exchange within minutes is also indicative of the presence of highly flexible and/or unstructured regions. In all, the fast exchange rates independent of the buffer pH justify the assumption that there is an altered interaction among the charged residues, which leads to a steadily-open pore. The role of the side-chain interactions within the pore and between the loops involving the loop L6 is also discussed.
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Affiliation(s)
- İrem Yılmaz
- Physics Unit, Biophysics Laboratory, Atilim University, 06836 Ankara, Turkey
| | - Filiz Korkmaz
- Physics Unit, Biophysics Laboratory, Atilim University, 06836 Ankara, Turkey.
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16
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Joyner PM, Tran DP, Zenaidee MA, Loo JA. Characterization of protein-ligand binding interactions of enoyl-ACP reductase (FabI) by native MS reveals allosteric effects of coenzymes and the inhibitor triclosan. Protein Sci 2022; 31:568-579. [PMID: 34882866 PMCID: PMC8862436 DOI: 10.1002/pro.4252] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/24/2021] [Revised: 12/06/2021] [Accepted: 12/07/2021] [Indexed: 01/08/2023]
Abstract
The enzyme enoyl-ACP reductase (also called FabI in bacteria) is an essential member of the fatty acid synthase II pathway in plants and bacteria. This enzyme is the target of the antibacterial drug triclosan and has been the subject of extensive studies for the past 20 years. Despite the large number of reports describing the biochemistry of this enzyme, there have been no studies that provided direct observation of the protein and its various ligands. Here we describe the use of native MS to characterize the protein-ligand interactions of FabI with its coenzymes NAD+ and NADH and with the inhibitor triclosan. Measurements of the gas-phase affinities of the enzyme for these ligands yielded values that are in close agreement with solution-phase affinity measurements. Additionally, FabI is a homotetramer and we were able to measure the affinity of each subunit for each coenzyme, which revealed that both coenzymes exhibit a positive homotropic allosteric effect. An allosteric effect was also observed in association with the inhibitor triclosan. These observations provide new insights into this well-studied enzyme and suggest that there may still be gaps in the existing mechanistic models that explain FabI inhibition.
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Affiliation(s)
- P. Matthew Joyner
- Natural Science DivisionPepperdine UniversityMalibuCaliforniaUSA
- Department of Chemistry & BiochemistryUniversity of California‐Los AngelesLos AngelesCaliforniaUSA
| | - Denise P. Tran
- Department of Chemistry & BiochemistryUniversity of California‐Los AngelesLos AngelesCaliforniaUSA
- Sydney Mass SpectrometryThe University of Sydney, Charles Perkins CentreCamperdownNew South WalesAustralia
| | - Muhammad A. Zenaidee
- Department of Chemistry & BiochemistryUniversity of California‐Los AngelesLos AngelesCaliforniaUSA
- Australian Proteome Analysis FacilityMacquarie UniversityMacquarieNew South WalesAustralia
| | - Joseph A. Loo
- Department of Chemistry & BiochemistryUniversity of California‐Los AngelesLos AngelesCaliforniaUSA
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17
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Correy GJ, Kneller DW, Phillips G, Pant S, Russi S, Cohen AE, Meigs G, Holton JM, Gahbauer S, Thompson MC, Ashworth A, Coates L, Kovalevsky A, Meilleur F, Fraser JS. The mechanisms of catalysis and ligand binding for the SARS-CoV-2 NSP3 macrodomain from neutron and X-ray diffraction at room temperature. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2022:2022.02.07.479477. [PMID: 35169801 PMCID: PMC8845425 DOI: 10.1101/2022.02.07.479477] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Indexed: 11/25/2022]
Abstract
The NSP3 macrodomain of SARS CoV 2 (Mac1) removes ADP-ribosylation post-translational modifications, playing a key role in the immune evasion capabilities of the virus responsible for the COVID-19 pandemic. Here, we determined neutron and X-ray crystal structures of the SARS-CoV-2 NSP3 macrodomain using multiple crystal forms, temperatures, and pHs, across the apo and ADP-ribose-bound states. We characterize extensive solvation in the Mac1 active site, and visualize how water networks reorganize upon binding of ADP-ribose and non-native ligands, inspiring strategies for displacing waters to increase potency of Mac1 inhibitors. Determining the precise orientations of active site water molecules and the protonation states of key catalytic site residues by neutron crystallography suggests a catalytic mechanism for coronavirus macrodomains distinct from the substrate-assisted mechanism proposed for human MacroD2. These data provoke a re-evaluation of macrodomain catalytic mechanisms and will guide the optimization of Mac1 inhibitors.
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Affiliation(s)
- Galen J. Correy
- Department of Bioengineering and Therapeutic Sciences, University of California San Francisco, San Francisco, CA 94158, USA
| | - Daniel W. Kneller
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, US Department of Energy, USA
| | - Gwyndalyn Phillips
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, US Department of Energy, USA
| | - Swati Pant
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, US Department of Energy, USA
| | - Silvia Russi
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Center, Menlo Park, CA 94025, USA
| | - Aina E. Cohen
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Center, Menlo Park, CA 94025, USA
| | - George Meigs
- Department of Molecular Biophysics and Integrated Bioimaging, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
- Department of Biochemistry and Biophysics, University of California San Francisco, CA 94158, USA
| | - James M. Holton
- Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Center, Menlo Park, CA 94025, USA
- Department of Molecular Biophysics and Integrated Bioimaging, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
- Department of Biochemistry and Biophysics, University of California San Francisco, CA 94158, USA
| | - Stefan Gahbauer
- Department of Pharmaceutical Chemistry, University of California San Francisco, San Francisco, CA 94158, USA
| | - Michael C. Thompson
- Department of Chemistry and Chemical Biology, University of California Merced, CA 95343, USA
| | - Alan Ashworth
- Helen Diller Family Comprehensive Cancer, University of California San Francisco, CA 94158, USA
| | - Leighton Coates
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, US Department of Energy, USA
| | - Andrey Kovalevsky
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- National Virtual Biotechnology Laboratory, US Department of Energy, USA
| | - Flora Meilleur
- Neutron Scattering Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA
- Department of Molecular and Structural Biochemistry, North Carolina State University, Raleigh, NC 27695
| | - James S. Fraser
- Department of Bioengineering and Therapeutic Sciences, University of California San Francisco, San Francisco, CA 94158, USA
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18
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Native state fluctuations in a peroxiredoxin active site match motions needed for catalysis. Structure 2022; 30:278-288.e3. [PMID: 34678159 PMCID: PMC8818020 DOI: 10.1016/j.str.2021.10.001] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/22/2021] [Revised: 08/10/2021] [Accepted: 09/29/2021] [Indexed: 02/05/2023]
Abstract
Peroxiredoxins are ubiquitous enzymes that detoxify peroxides and regulate redox signaling. During catalysis, a "peroxidatic" cysteine (CP) in the conserved active site reduces peroxide while being oxidized to a CP-sulfenate, prompting a local unfolding event that enables formation of a disulfide with a second, "resolving" cysteine. Here, we use nuclear magnetic resonance spectroscopy to probe the dynamics of the CP-thiolate and disulfide forms of Xanthomonas campestris peroxiredoxin Q. Chemical exchange saturation transfer behavior of the resting enzyme reveals 26 residues in and around the active site exchanging at a rate of 72 s-1 with a locally unfolded, high-energy (2.5% of the population) state. This unequivocally establishes that a catalytically relevant local unfolding equilibrium exists in the enzyme's CP-thiolate form. Also, faster motions imply an active site instability that could promote local unfolding and, based on other work, be exacerbated by CP-sulfenate formation so as to direct the enzyme along a functional catalytic trajectory.
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19
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Devaurs D, Antunes DA, Borysik AJ. Computational Modeling of Molecular Structures Guided by Hydrogen-Exchange Data. JOURNAL OF THE AMERICAN SOCIETY FOR MASS SPECTROMETRY 2022; 33:215-237. [PMID: 35077179 DOI: 10.1021/jasms.1c00328] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/14/2023]
Abstract
Data produced by hydrogen-exchange monitoring experiments have been used in structural studies of molecules for several decades. Despite uncertainties about the structural determinants of hydrogen exchange itself, such data have successfully helped guide the structural modeling of challenging molecular systems, such as membrane proteins or large macromolecular complexes. As hydrogen-exchange monitoring provides information on the dynamics of molecules in solution, it can complement other experimental techniques in so-called integrative modeling approaches. However, hydrogen-exchange data have often only been used to qualitatively assess molecular structures produced by computational modeling tools. In this paper, we look beyond qualitative approaches and survey the various paradigms under which hydrogen-exchange data have been used to quantitatively guide the computational modeling of molecular structures. Although numerous prediction models have been proposed to link molecular structure and hydrogen exchange, none of them has been widely accepted by the structural biology community. Here, we present as many hydrogen-exchange prediction models as we could find in the literature, with the aim of providing the first exhaustive list of its kind. From purely structure-based models to so-called fractional-population models or knowledge-based models, the field is quite vast. We aspire for this paper to become a resource for practitioners to gain a broader perspective on the field and guide research toward the definition of better prediction models. This will eventually improve synergies between hydrogen-exchange monitoring and molecular modeling.
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Affiliation(s)
- Didier Devaurs
- MRC Institute of Genetics and Cancer, University of Edinburgh, Edinburgh EH4 2XU, U.K
| | - Dinler A Antunes
- Department of Biology and Biochemistry, University of Houston, Houston, Texas 77005, United States
| | - Antoni J Borysik
- Department of Chemistry, King's College London, London SE1 1DB, U.K
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20
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Zhdanova PV, Chernonosov AA, Prokhorova DV, Stepanov GA, Kanazhevskaya LY, Koval VV. Probing the Dynamics of Streptococcus pyogenes Cas9 Endonuclease Bound to the sgRNA Complex Using Hydrogen-Deuterium Exchange Mass Spectrometry. Int J Mol Sci 2022; 23:1129. [PMID: 35163047 PMCID: PMC8834707 DOI: 10.3390/ijms23031129] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/08/2021] [Revised: 01/17/2022] [Accepted: 01/18/2022] [Indexed: 02/04/2023] Open
Abstract
The Cas9 endonuclease is an essential component of the CRISPR-Cas-based genome editing tools. The attainment of high specificity and efficiency of Cas9 during targetted DNA cleavage is the main problem that limits the clinical application of the CRISPR-Cas9 system. A deep understanding of the Cas9 mechanism and its structural-functional relationships is required to develop strategies for precise gene editing. Here, we present the first attempt to describe the solution structure of Cas9 from S. pyogenes using hydrogen-deuterium exchange mass spectrometry (HDX-MS) coupled to molecular dynamics simulations. HDX data revealed multiple protein regions with deuterium uptake levels varying from low to high. By analysing the difference in relative deuterium uptake by apoCas9 and its complex with sgRNA, we identified peptides involved in the complex formation and possible changes in the protein conformation. The REC3 domain was shown to undergo the most prominent conformational change upon enzyme-RNA interactions. Detection of the HDX in two forms of the enzyme provided detailed information about changes in the Cas9 structure induced by sgRNA binding and quantified the extent of the changes. The study demonstrates the practical utility of HDX-MS for the elucidation of mechanistic aspects of Cas9 functioning.
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Affiliation(s)
- Polina V. Zhdanova
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
- Department of Natural Sciences, Novosibirsk State University, 630090 Novosibirsk, Russia
| | - Alexander A. Chernonosov
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
| | - Daria V. Prokhorova
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
| | - Grigory A. Stepanov
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
| | - Lyubov Yu. Kanazhevskaya
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
| | - Vladimir V. Koval
- Institute of Chemical Biology and Fundamental Medicine, Siberian Branch of the Russian Academy of Sciences (SB RAS), 630090 Novosibirsk, Russia; (P.V.Z.); (A.A.C.); (D.V.P.); (G.A.S.); (L.Y.K.)
- Department of Natural Sciences, Novosibirsk State University, 630090 Novosibirsk, Russia
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21
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Trainor K, Doyle CM, Metcalfe-Roach A, Steckner J, Lipovšek D, Malakian H, Langley D, Krystek SR, Meiering EM. Design for Solubility May Reveal Induction of Amide Hydrogen/Deuterium Exchange by Protein Self-Association. J Mol Biol 2021; 434:167398. [PMID: 34902431 DOI: 10.1016/j.jmb.2021.167398] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/29/2021] [Revised: 12/02/2021] [Accepted: 12/03/2021] [Indexed: 10/19/2022]
Abstract
Structural heterogeneity often constrains the characterization of aggregating proteins to indirect or low-resolution methods, obscuring mechanistic details of association. Here, we report progress in understanding the aggregation of Adnectins, engineered binding proteins with an immunoglobulin-like fold. We rationally design Adnectin solubility and measure amide hydrogen/deuterium exchange (HDX) under conditions that permit transient protein self-association. Protein-protein binding commonly slows rates of HDX; in contrast, we find that Adnectin association may induce faster HDX for certain amides, particularly in the C-terminal β-strand. In aggregation-prone proteins, we identify a pattern of very different rates of amide HDX for residues linked by reciprocal hydrogen bonds in the native structure. These results may be explained by local loss of native structure and formation of an inter-protein interface. Amide HDX induced by self-association, detected here by deliberate modulation of propensity for such interactions, may be a general phenomenon with the potential to expose mechanisms of aggregation by diverse proteins.
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Affiliation(s)
- Kyle Trainor
- Department of Chemistry, University of Waterloo, Waterloo N2L 3G1, Ontario, Canada
| | - Colleen M Doyle
- Department of Chemistry, University of Waterloo, Waterloo N2L 3G1, Ontario, Canada.
| | - Avril Metcalfe-Roach
- Department of Chemistry, University of Waterloo, Waterloo N2L 3G1, Ontario, Canada.
| | - Julia Steckner
- Department of Chemistry, University of Waterloo, Waterloo N2L 3G1, Ontario, Canada
| | - Daša Lipovšek
- Bristol Myers Squibb, Cambridge, MA 02140, United States.
| | | | - David Langley
- Bristol Myers Squibb, Cambridge, MA 02140, United States
| | | | - Elizabeth M Meiering
- Department of Chemistry, University of Waterloo, Waterloo N2L 3G1, Ontario, Canada.
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22
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Evangelista W, Knapp J, Zandarashvili L, Esadze A, White MA, Gribenko AV, Lee JC. Signal Transmission in Escherichia coli Cyclic AMP Receptor Protein for Survival in Extreme Acidic Conditions. Biochemistry 2021; 60:2987-3006. [PMID: 34605636 DOI: 10.1021/acs.biochem.1c00388] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/30/2022]
Abstract
During the life cycle of enteric bacterium Escherichia coli, it encounters a wide spectrum of pH changes. The asymmetric dimer of the cAMP receptor protein, CRP, plays a key role in regulating the expressions of genes and the survival of E. coli. To elucidate the pH effects on the mechanism of signal transmission, we present a combination of results derived from ITC, crystallography, and computation. CRP responds to a pH change by inducing a differential effect on the affinity for the binding events to the two cAMP molecules, ensuing in a reversible conversion between positive and negative cooperativity at high and low pH, respectively. The structures of four crystals at pH ranging from 7.8 to 6.5 show that CRP responds by inducing a differential effect on the structures of the two subunits, particularly in the DNA binding domain. Employing the COREX/BEST algorithm, computational analysis shows the change in the stability of residues at each pH. The change in residue stability alters the connectivity between residues including those in cAMP and DNA binding sites. Consequently, the differential impact on the topology of the connectivity surface among residues in adjacent subunits is the main reason for differential change in affinity; that is, the pH-induced differential change in residue stability is the biothermodynamic basis for the change in allosteric behavior. Furthermore, the structural asymmetry of this homodimer amplifies the differential impact of any perturbations. Hence, these results demonstrate that the combination of these approaches can provide insights into the underlying mechanism of an apparent complex allostery signal and transmission in CRP.
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Affiliation(s)
- Wilfredo Evangelista
- Centre for Research and Innovation-Faculty of Health Sciences, Universidad Peruana de Ciencias Aplicadas, Lima 15023, Peru
| | - James Knapp
- Department of Biochemistry and Molecular Biology, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States.,Sealy Center for Structural Biology and Molecular Biophysics, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States
| | - Levani Zandarashvili
- Department of Biochemistry and Molecular Biology, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States.,Sealy Center for Structural Biology and Molecular Biophysics, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States
| | | | - Mark A White
- Sealy Center for Structural Biology and Molecular Biophysics, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States
| | - Alexey V Gribenko
- Department of Biochemistry and Molecular Biology, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States.,Sealy Center for Structural Biology and Molecular Biophysics, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States
| | - J Ching Lee
- Department of Biochemistry and Molecular Biology, The University of Texas Medical Branch at Galveston, Galveston, Texas 77555-1055, United States
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23
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Liu H, Xiang S, Zhu H, Li L. The Structural and Dynamical Properties of the Hydration of SNase Based on a Molecular Dynamics Simulation. Molecules 2021; 26:molecules26175403. [PMID: 34500836 PMCID: PMC8434405 DOI: 10.3390/molecules26175403] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/25/2021] [Revised: 09/02/2021] [Accepted: 09/03/2021] [Indexed: 12/01/2022] Open
Abstract
The dynamics of protein–water fluctuations are of biological significance. Molecular dynamics simulations were performed in order to explore the hydration dynamics of staphylococcal nuclease (SNase) at different temperatures and mutation levels. A dynamical transition in hydration water (at ~210 K) can trigger larger-amplitude fluctuations of protein. The protein–water hydrogen bonds lost about 40% in the total change from 150 K to 210 K, while the Mean Square Displacement increased by little. The protein was activated when the hydration water in local had a comparable trend in making hydrogen bonds with protein– and other waters. The mutations changed the local chemical properties and the hydration exhibited a biphasic distribution, with two time scales. Hydrogen bonding relaxation governed the local protein fluctuations on the picosecond time scale, with the fastest time (24.9 ps) at the hydrophobic site and slowest time (40.4 ps) in the charged environment. The protein dynamic was related to the water’s translational diffusion via the relaxation of the protein–water’s H-bonding. The structural and dynamical properties of protein–water at the molecular level are fundamental to the physiological and functional mechanisms of SNase.
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Affiliation(s)
- Hangxin Liu
- National and Local Joint Engineering Research Center of Biomedical Functional Materials, Nanjing 210023, China; (H.L.); (S.X.)
- Jiangsu Collaborative Innovation Center of Biomedical Functional Materials, Nanjing 210023, China
- Jiangsu Engineering Research Center for Biomedical Function Materials, Nanjing 210023, China
- Jiangsu Key Laboratory of Biofunctional Materials, Nanjing 210023, China
- School of Chemistry and Materials Science, Nanjing Normal University, No. 1 Wenyuan Road, Nanjing 210023, China
| | - Shuqing Xiang
- National and Local Joint Engineering Research Center of Biomedical Functional Materials, Nanjing 210023, China; (H.L.); (S.X.)
- Jiangsu Collaborative Innovation Center of Biomedical Functional Materials, Nanjing 210023, China
- Jiangsu Engineering Research Center for Biomedical Function Materials, Nanjing 210023, China
- Jiangsu Key Laboratory of Biofunctional Materials, Nanjing 210023, China
- School of Chemistry and Materials Science, Nanjing Normal University, No. 1 Wenyuan Road, Nanjing 210023, China
| | - Haomiao Zhu
- National and Local Joint Engineering Research Center of Biomedical Functional Materials, Nanjing 210023, China; (H.L.); (S.X.)
- Jiangsu Collaborative Innovation Center of Biomedical Functional Materials, Nanjing 210023, China
- Jiangsu Engineering Research Center for Biomedical Function Materials, Nanjing 210023, China
- Jiangsu Key Laboratory of Biofunctional Materials, Nanjing 210023, China
- School of Chemistry and Materials Science, Nanjing Normal University, No. 1 Wenyuan Road, Nanjing 210023, China
- Correspondence: (H.Z.); (L.L.)
| | - Li Li
- National and Local Joint Engineering Research Center of Biomedical Functional Materials, Nanjing 210023, China; (H.L.); (S.X.)
- Jiangsu Collaborative Innovation Center of Biomedical Functional Materials, Nanjing 210023, China
- Jiangsu Engineering Research Center for Biomedical Function Materials, Nanjing 210023, China
- Jiangsu Key Laboratory of Biofunctional Materials, Nanjing 210023, China
- School of Chemistry and Materials Science, Nanjing Normal University, No. 1 Wenyuan Road, Nanjing 210023, China
- Correspondence: (H.Z.); (L.L.)
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24
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Ye X, Mayne L, Englander SW. A conserved strategy for structure change and energy transduction in Hsp104 and other AAA+ protein motors. J Biol Chem 2021; 297:101066. [PMID: 34384781 PMCID: PMC8449053 DOI: 10.1016/j.jbc.2021.101066] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/04/2021] [Revised: 08/05/2021] [Accepted: 08/06/2021] [Indexed: 01/09/2023] Open
Abstract
The superfamily of massively large AAA+ protein molecular machines functions to convert the chemical energy of cytosolic ATP into physicomechanical form and use it to perform an extraordinary number of physical operations on proteins, nucleic acids, and membrane systems. Cryo-EM studies now reveal some aspects of substrate handling at high resolution, but the broader interpretation of AAA+ functional properties is still opaque. This paper integrates recent hydrogen exchange results for the typical AAA+ protein Hsp104 with prior information on several near and distantly related others. The analysis points to a widely conserved functional strategy. Hsp104 cycles through a long-lived loosely-structured energy-input "open" state that releases spent ADP and rebinds cytosolic ATP. ATP-binding energy is transduced by allosteric structure change to poise the protein at a high energy level in a more tightly structured "closed" state. The briefly occupied energy-output closed state binds substrate strongly and is catalytically active. ATP hydrolysis permits energetically downhill structural relaxation, which is coupled to drive energy-requiring substrate processing. Other AAA+ proteins appear to cycle through states that are analogous functionally if not in structural detail. These results revise the current model for AAA+ function, explain the structural basis of single-molecule optical tweezer kinetic phases, identify the separate energetic roles of ATP binding and hydrolysis, and specify a sequence of structural and energetic events that carry AAA+ proteins unidirectionally around a functional cycle to propel their diverse physical tasks.
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Affiliation(s)
- Xiang Ye
- Department of Biochemistry and Biophysics, Texas A&M University, College Station, Texas, USA
| | - Leland Mayne
- Department of Biochemistry and Biophysics and Johnson Research Foundation, Perelman School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA
| | - S Walter Englander
- Department of Biochemistry and Biophysics and Johnson Research Foundation, Perelman School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA.
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25
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Grauslund LR, Calvaresi V, Pansegrau W, Norais N, Rand KD. Epitope and Paratope Mapping by HDX-MS Combined with SPR Elucidates the Difference in Bactericidal Activity of Two Anti-NadA Monoclonal Antibodies. JOURNAL OF THE AMERICAN SOCIETY FOR MASS SPECTROMETRY 2021; 32:1575-1582. [PMID: 33683906 DOI: 10.1021/jasms.0c00431] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/12/2023]
Abstract
Characterization of antigen-antibody interactions is crucial for understanding antibody-mediated protection against pathogens, biopharmaceutical development, as well as evaluation of the immune response post vaccination. Bexsero is a multicomponent vaccine against Neisseria meningitidis serogroup B in which one of the key vaccine antigens is Neisserial adhesin A (NadA), a trimeric coiled-coil protein. Two NadA-specific monoclonal antibodies (mAbs) isolated from Bexsero-vaccinated individuals have been shown to have similar binding affinity and appear to recognize a similar antigen region, yet only one of the mAbs is bactericidal. In this study, we use hydrogen/deuterium exchange mass spectrometry (HDX-MS) to perform an in-depth study of the interaction of the two mAbs with NadA antigen using a combined epitope and paratope mapping strategy. In addition, we use surface plasmon resonance (SPR) to investigate the stoichiometry of the binding of the two mAbs to NadA. While epitope mapping only identifies a clear binding impact of one of the mAbs on NadA, the paratope mapping analyses shows that both mAbs are binding to NadA through several complementarity determining regions spanning both heavy and light chains. Our results highlight the advantage of combined epitope and paratope mapping HDX-MS experiments and supporting biochemical experiments to characterize antigen-antibody interactions. Through this combined approach, we provide a rationale for how the binding stoichiometry of the two mAbs to the trimeric NadA antigen can explain the difference in bactericidal activity of the two mAbs.
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Affiliation(s)
- Laura R Grauslund
- Protein Analysis Group, Department of Pharmacy, University of Copenhagen, 2100 Copenhagen O, Denmark
- GSK, Siena 53100, Italy
| | - Valeria Calvaresi
- Protein Analysis Group, Department of Pharmacy, University of Copenhagen, 2100 Copenhagen O, Denmark
- GSK, Siena 53100, Italy
| | | | | | - Kasper D Rand
- Protein Analysis Group, Department of Pharmacy, University of Copenhagen, 2100 Copenhagen O, Denmark
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26
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Abstract
The presence of metamorphism in the protein's native state is not yet fully understood. To shed light on this issue, we present an assessment, in terms of the amide hydrogen exchange protection factor, that aims to determine the possible existence of structural fluctuations in the native-state consistent with both the upper-bound marginal stability of proteins and the presence of metamorphism. The preliminary results enable us to conclude that the native-state metamorphism is, indeed, more probable than previously thought.
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Affiliation(s)
- Jorge A Vila
- IMASL-CONICET, Universidad Nacional de San Luis, Ejército de Los Andes 950, 5700 San Luis, Argentina
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27
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Sivaraman T. A Review on Computational Approaches for Analyzing Hydrogen- Deuterium (H/D) Exchange of Proteins. Protein Pept Lett 2021; 28:372-381. [PMID: 33006533 DOI: 10.2174/0929866527666201002145859] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2020] [Revised: 08/14/2020] [Accepted: 08/17/2020] [Indexed: 11/22/2022]
Abstract
Native state Hydrogen-Deuterium (H/D) exchange method has been used to study the structures and the unfolding pathways for quite a number of proteins. The H/D exchange method is generally monitored using nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry (MS) techniques. NMR-assisted H/D exchange methods primarily monitor the residue level fluctuation of proteins, whereas MS-assisted H/D exchange methods analyze multifold ensemble conformations of proteins. In this connection, quite a large number of computational tools and algorithms have been developed for processing and analyzing huge amount of the H/D exchange data generated from these techniques. In this review, most of the freely available computational tools associated with the H/D exchange of proteins have been comprehensively reviewed and scopes to improve/ develop novel computational approaches for analyzing the H/D exchange data of proteins have also been brought into fore.
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Affiliation(s)
- Thirunavukkarasu Sivaraman
- Drug Design and Discovery Lab, Department of Biotechnology, Karpagam Academy of Higher Education (Deemed to be University), Coimbatore - 641021, Tamil Nadu, India
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28
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Bai Y, Wan W, Huang Y, Jin W, Lyu H, Xia Q, Dong X, Gao Z, Liu Y. Quantitative interrogation of protein co-aggregation using multi-color fluorogenic protein aggregation sensors. Chem Sci 2021; 12:8468-8476. [PMID: 34221329 PMCID: PMC8221170 DOI: 10.1039/d1sc01122g] [Citation(s) in RCA: 21] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/25/2021] [Accepted: 05/19/2021] [Indexed: 12/27/2022] Open
Abstract
Co-aggregation of multiple pathogenic proteins is common in neurodegenerative diseases but deconvolution of such biochemical process is challenging. Herein, we developed a dual-color fluorogenic thermal shift assay to simultaneously report on the aggregation of two different proteins and quantitatively study their thermodynamic stability during co-aggregation. Expansion of spectral coverage was first achieved by developing multi-color fluorogenic protein aggregation sensors. Orthogonal detection was enabled by conjugating sensors of minimal fluorescence crosstalk to two different proteins via sortase-tag technology. Using this assay, we quantified shifts in melting temperatures in a heterozygous model protein system, revealing that the thermodynamic stability of wild-type proteins was significantly compromised by the mutant ones but not vice versa. We also examined how small molecule ligands selectively and differentially interfere with such interplay. Finally, we demonstrated these sensors are suited to visualize how different proteins exert influence on each other upon their co-aggregation in live cells.
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Affiliation(s)
- Yulong Bai
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
- University of Chinese Academy of Sciences Beijing 100049 China
| | - Wang Wan
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
| | - Yanan Huang
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
| | - Wenhan Jin
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
| | - Haochen Lyu
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
| | - Qiuxuan Xia
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
- University of Chinese Academy of Sciences Beijing 100049 China
| | - Xuepeng Dong
- The Second Hospital of Dalian Medical University 467 Zhongshan Road Dalian 116044 China
| | - Zhenming Gao
- The Second Hospital of Dalian Medical University 467 Zhongshan Road Dalian 116044 China
| | - Yu Liu
- CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences 457 Zhongshan Road Dalian 116023 China
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29
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Rosa B, Dickinson ER, Marchetti M, Campanini B, Pioselli B, Bettati S, Rand KD. Revealing the Dynamic Allosteric Changes Required for Formation of the Cysteine Synthase Complex by Hydrogen-Deuterium Exchange MS. Mol Cell Proteomics 2021; 20:100098. [PMID: 34022432 PMCID: PMC8253905 DOI: 10.1016/j.mcpro.2021.100098] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/10/2021] [Accepted: 05/14/2021] [Indexed: 11/30/2022] Open
Abstract
CysE and CysK, the last two enzymes of the cysteine biosynthetic pathway, engage in a bienzyme complex, cysteine synthase, with yet incompletely characterized three-dimensional structure and regulatory function. Being absent in mammals, the two enzymes and their complex are attractive targets for antibacterial drugs. We have used hydrogen/deuterium exchange MS to unveil how complex formation affects the conformational dynamics of CysK and CysE. Our results support a model where CysE is present in solution as a dimer of trimers, and each trimer can bind one CysK homodimer. When CysK binds to one CysE monomer, intratrimer allosteric communication ensures conformational and dynamic symmetry within the trimer. Furthermore, a long-range allosteric signal propagates through CysE to induce stabilization of the interface between the two CysE trimers, preparing the second trimer for binding the second CysK with a nonrandom orientation. These results provide new molecular insights into the allosteric formation of the cysteine synthase complex and could help guide antibacterial drug design. HDX-MS reveals complex formation impact on conformational dynamics of CysK and CysE. CysK binding ensures conformational symmetry within the CysE trimer. Long-range allostery propagates through CysE stabilizing the inter-trimer interface. Insights into the allostery of CS complex could help guide antibacterial drug design.
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Affiliation(s)
- Brenda Rosa
- Biopharmanet-TEC Interdepartmental Center, University di Parma, Parma, Italy
| | - Eleanor R Dickinson
- Protein Analysis Group, Department of Pharmacy, University of Copenhagen, Copenhagen O, Denmark
| | | | | | | | - Stefano Bettati
- Biopharmanet-TEC Interdepartmental Center, University di Parma, Parma, Italy; Department of Medicine and Surgery, University of Parma, Parma, Italy; Institute of Biophysics, CNR, Pisa, Italy.
| | - Kasper Dyrberg Rand
- Protein Analysis Group, Department of Pharmacy, University of Copenhagen, Copenhagen O, Denmark.
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30
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Merkle PS, Trabjerg E, Hongjian S, Ferber M, Cuendet MA, Jørgensen TJD, Luescher I, Irving M, Zoete V, Michielin O, Rand KD. Probing the Conformational Dynamics of Affinity-Enhanced T Cell Receptor Variants upon Binding the Peptide-Bound Major Histocompatibility Complex by Hydrogen/Deuterium Exchange Mass Spectrometry. Biochemistry 2021; 60:859-872. [PMID: 33689297 DOI: 10.1021/acs.biochem.1c00035] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/30/2022]
Abstract
Binding of the T cell receptor (TCR) to its cognate, peptide antigen-loaded major histocompatibility complex (pMHC) is a key interaction for triggering T cell activation and ultimately elimination of the target cell. Despite the importance of this interaction for cellular immunity, a comprehensive molecular understanding of TCR specificity and affinity is lacking. We conducted hydrogen/deuterium exchange mass spectrometry (HDX-MS) analyses of individual affinity-enhanced TCR variants and clinically relevant pMHC class I molecules (HLA-A*0201/NY-ESO-1157-165) to investigate the causality between increased binding affinity and conformational dynamics in TCR-pMHC complexes. Differential HDX-MS analyses of TCR variants revealed that mutations for affinity enhancement in TCR CDRs altered the conformational response of TCR to pMHC ligation. Improved pMHC binding affinity was in general observed to correlate with greater differences in HDX upon pMHC binding in modified TCR CDR loops, thereby providing new insights into the TCR-pMHC interaction. Furthermore, a specific point mutation in the β-CDR3 loop of the NY-ESO-1 TCR associated with a substantial increase in binding affinity resulted in a substantial change in pMHC binding kinetics (i.e., very slow kon, revealed by the detection of EX1 HDX kinetics), thus providing experimental evidence for a slow induced-fit binding mode. We also examined the conformational impact of pMHC binding on an unrelated TRAV12-2 gene-encoded TCR directed against the immunodominant MART-126-35 cancer antigen restricted by HLA-A*0201. Our findings provide a molecular basis for the observed TRAV12-2 gene bias in natural CD8+ T cell-based immune responses against the MART-1 antigen, with potential implications for general ligand discrimination and TCR cross-reactivity processes.
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MESH Headings
- Humans
- Receptors, Antigen, T-Cell/metabolism
- Receptors, Antigen, T-Cell/chemistry
- Receptors, Antigen, T-Cell/immunology
- HLA-A2 Antigen/chemistry
- HLA-A2 Antigen/immunology
- HLA-A2 Antigen/metabolism
- HLA-A2 Antigen/genetics
- Protein Conformation
- Hydrogen Deuterium Exchange-Mass Spectrometry
- Protein Binding
- Peptides/chemistry
- Peptides/metabolism
- Peptides/immunology
- Major Histocompatibility Complex
- Antigens, Neoplasm/chemistry
- Antigens, Neoplasm/metabolism
- Antigens, Neoplasm/immunology
- Antigens, Neoplasm/genetics
- Peptide Fragments/chemistry
- Peptide Fragments/metabolism
- Peptide Fragments/genetics
- Peptide Fragments/immunology
- Deuterium Exchange Measurement
- Mutation
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Affiliation(s)
- Patrick S Merkle
- Department of Pharmacy, University of Copenhagen, 2100 Copenhagen, Denmark
| | - Esben Trabjerg
- Department of Pharmacy, University of Copenhagen, 2100 Copenhagen, Denmark
| | - Song Hongjian
- Department of Pharmacy, University of Copenhagen, 2100 Copenhagen, Denmark
| | - Mathias Ferber
- Swiss Institute of Bioinformatics, Bâtiment Génopode, UNIL Sorge, 1015 Lausanne, Switzerland
| | - Michel A Cuendet
- Swiss Institute of Bioinformatics, Bâtiment Génopode, UNIL Sorge, 1015 Lausanne, Switzerland
- Weill Cornell Medical College, 1300 York Avenue, New York, New York 10065, United States
| | - Thomas J D Jørgensen
- Department of Biochemistry and Molecular Biology, University of Southern Denmark, 5230 Odense, Denmark
| | - Immanuel Luescher
- Ludwig Branch for Cancer Research of the University of Lausanne, 8001 Zurich, Switzerland
| | - Melita Irving
- Swiss Institute of Bioinformatics, Bâtiment Génopode, UNIL Sorge, 1015 Lausanne, Switzerland
- Ludwig Branch for Cancer Research of the University of Lausanne, 8001 Zurich, Switzerland
| | - Vincent Zoete
- Swiss Institute of Bioinformatics, Bâtiment Génopode, UNIL Sorge, 1015 Lausanne, Switzerland
| | - Olivier Michielin
- Swiss Institute of Bioinformatics, Bâtiment Génopode, UNIL Sorge, 1015 Lausanne, Switzerland
| | - Kasper D Rand
- Department of Pharmacy, University of Copenhagen, 2100 Copenhagen, Denmark
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31
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Evaluation of protein secondary structure from FTIR spectra improved after partial deuteration. EUROPEAN BIOPHYSICS JOURNAL: EBJ 2021; 50:613-628. [PMID: 33534058 PMCID: PMC8189984 DOI: 10.1007/s00249-021-01502-y] [Citation(s) in RCA: 17] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 10/12/2020] [Revised: 01/08/2021] [Accepted: 01/13/2021] [Indexed: 11/11/2022]
Abstract
FTIR spectroscopy has become a major tool to determine protein secondary structure. One of the identified obstacle for reaching better predictions is the strong overlap of bands assigned to different secondary structures. Yet, while for instance disordered structures and α-helical structures absorb almost at the same wavenumber, the absorbance bands are differentially shifted upon deuteration, in part because exchange is much faster for disordered structures. We recorded the FTIR spectra of 85 proteins at different stages of hydrogen/deuterium exchange process using protein microarrays and infrared imaging for high throughput measurements. Several methods were used to relate spectral shape to secondary structure content. While in absolute terms, β-sheet is always better predicted than α-helix content, results consistently indicate an improvement of secondary structure predictions essentially for the α-helix and the category called “Others” (grouping random, turns, bends, etc.) after 15 min of exchange. On the contrary, the β-sheet fraction is better predicted in non-deuterated conditions. Using partial least square regression, the error of prediction for the α-helix content is reduced after 15-min deuteration. Further deuteration degrades the prediction. Error on the prediction for the “Others” structures also decreases after 15-min deuteration. Cross-validation or a single 25-protein test set result in the same overall conclusions.
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32
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Sun SC, Huang HW, Lo YT, Chuang MC, Hsu YHH. Unraveling cardiolipin-induced conformational change of cytochrome c through H/D exchange mass spectrometry and quartz crystal microbalance. Sci Rep 2021; 11:1090. [PMID: 33441668 PMCID: PMC7806790 DOI: 10.1038/s41598-020-79905-8] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/16/2020] [Accepted: 12/04/2020] [Indexed: 11/21/2022] Open
Abstract
Cardiolipin (CL), a crucial component in inner mitochondrial membranes, interacts with cytochrome c (cyt c) to form a peroxidase complex for the catalysis of CL oxidation. Such interaction is pivotal to the mitochondrial regulation of apoptosis and is affected by the redox state of cyt c. In the present study, the redox-dependent interaction of cyt c with CL was investigated through amide hydrogen/deuterium exchange coupled with mass spectrometry (HDXMS) and quartz crystal microbalance with dissipation monitoring (QCM-D). Ferrous cyt c exhibited a more compact conformation compared with its ferric form, which was supported by the lower number of deuterons accumulated and the greater amplitude reduction on dissipation. Upon association with CL, ferrous cyt c resulted in a moderate increase in deuteration, whereas the ferric form caused a drastic increase of deuteration, which indicated that CL-bound ferric cyt c formed an extended conformation. These results were consistent with those of the frequency (f) − dissipation (D) experiments, which revealed that ferric cyt c yielded greater values of |ΔD/Δf| within the first minute. Further fragmentation analysis based on HDXMS indicated that the effect of CL binding was considerably different on ferric and ferrous cyt c in the C-helix and the Loop 9–24. In ferric cyt c, CL binding affected Met80 and destabilized His18 interaction with heme, which was not observed with ferrous cyt c. An interaction model was proposed to explain the aforementioned results.
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Affiliation(s)
- Sin-Cih Sun
- Department of Chemistry, Tunghai University, Taichung, Taiwan
| | - Hung-Wei Huang
- Department of Chemistry, Tunghai University, Taichung, Taiwan
| | - Yi-Ting Lo
- Department of Chemistry, Tunghai University, Taichung, Taiwan
| | - Min-Chieh Chuang
- Department of Chemistry, Tunghai University, Taichung, Taiwan. .,Department of Environmental Science and Engineering, Taichung, Taiwan.
| | - Yuan-Hao Howard Hsu
- Department of Chemistry, Tunghai University, Taichung, Taiwan. .,Biological Science Center, Tunghai University, Taichung, Taiwan.
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33
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Lee PS, Bradshaw RT, Marinelli F, Kihn K, Smith A, Wintrode PL, Deredge DJ, Faraldo-Gómez JD, Forrest LR. Interpreting Hydrogen-Deuterium Exchange Experiments with Molecular Simulations: Tutorials and Applications of the HDXer Ensemble Reweighting Software [Article v1.0]. LIVING JOURNAL OF COMPUTATIONAL MOLECULAR SCIENCE 2021; 3:1521. [PMID: 36644498 PMCID: PMC9835200 DOI: 10.33011/livecoms.3.1.1521] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 01/19/2023]
Abstract
Hydrogen-deuterium exchange (HDX) is a comprehensive yet detailed probe of protein structure and dynamics and, coupled to mass spectrometry, has become a powerful tool for investigating an increasingly large array of systems. Computer simulations are often used to help rationalize experimental observations of exchange, but interpretations have frequently been limited to simple, subjective correlations between microscopic dynamical fluctuations and the observed macroscopic exchange behavior. With this in mind, we previously developed the HDX ensemble reweighting approach and associated software, HDXer, to aid the objective interpretation of HDX data using molecular simulations. HDXer has two main functions; first, to compute H-D exchange rates that describe each structure in a candidate ensemble of protein structures, for example from molecular simulations, and second, to objectively reweight the conformational populations present in a candidate ensemble to conform to experimental exchange data. In this article, we first describe the HDXer approach, theory, and implementation. We then guide users through a suite of tutorials that demonstrate the practical aspects of preparing experimental data, computing HDX levels from molecular simulations, and performing ensemble reweighting analyses. Finally we provide a practical discussion of the capabilities and limitations of the HDXer methods including recommendations for a user's own analyses. Overall, this article is intended to provide an up-to-date, pedagogical counterpart to the software, which is freely available at https://github.com/Lucy-Forrest-Lab/HDXer.
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Affiliation(s)
- Paul Suhwan Lee
- Computational Structural Biology Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA
| | - Richard T. Bradshaw
- Computational Structural Biology Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA,For correspondence: (RTB); (LRF)
| | - Fabrizio Marinelli
- Theoretical Molecular Biophysics Laboratory, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, MD, USA
| | - Kyle Kihn
- Department of Pharmaceutical Sciences, School of Pharmacy, University of Maryland, Baltimore, MD, USA
| | - Ally Smith
- Department of Pharmaceutical Sciences, School of Pharmacy, University of Maryland, Baltimore, MD, USA
| | - Patrick L. Wintrode
- Department of Pharmaceutical Sciences, School of Pharmacy, University of Maryland, Baltimore, MD, USA
| | - Daniel J. Deredge
- Department of Pharmaceutical Sciences, School of Pharmacy, University of Maryland, Baltimore, MD, USA
| | - José D. Faraldo-Gómez
- Theoretical Molecular Biophysics Laboratory, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, MD, USA
| | - Lucy R. Forrest
- Computational Structural Biology Section, National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA,For correspondence: (RTB); (LRF)
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34
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Corless EI, Saad Imran SM, Watkins MB, Bacik JP, Mattice JR, Patterson A, Danyal K, Soffe M, Kitelinger R, Seefeldt LC, Origanti S, Bennett B, Bothner B, Ando N, Antony E. The flexible N-terminus of BchL autoinhibits activity through interaction with its [4Fe-4S] cluster and released upon ATP binding. J Biol Chem 2021; 296:100107. [PMID: 33219127 PMCID: PMC7948495 DOI: 10.1074/jbc.ra120.016278] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/05/2020] [Revised: 11/10/2020] [Accepted: 11/20/2020] [Indexed: 11/10/2022] Open
Abstract
A key step in bacteriochlorophyll biosynthesis is the reduction of protochlorophyllide to chlorophyllide, catalyzed by dark-operative protochlorophyllide oxidoreductase. Dark-operative protochlorophyllide oxidoreductase contains two [4Fe-4S]-containing component proteins (BchL and BchNB) that assemble upon ATP binding to BchL to coordinate electron transfer and protochlorophyllide reduction. But the precise nature of the ATP-induced conformational changes is poorly understood. We present a crystal structure of BchL in the nucleotide-free form where a conserved, flexible region in the N-terminus masks the [4Fe-4S] cluster at the docking interface between BchL and BchNB. Amino acid substitutions in this region produce a hyperactive enzyme complex, suggesting a role for the N-terminus in autoinhibition. Hydrogen-deuterium exchange mass spectrometry shows that ATP binding to BchL produces specific conformational changes leading to release of the flexible N-terminus from the docking interface. The release also promotes changes within the local environment surrounding the [4Fe-4S] cluster and promotes BchL-complex formation with BchNB. A key patch of amino acids, Asp-Phe-Asp (the 'DFD patch'), situated at the mouth of the BchL ATP-binding pocket promotes intersubunit cross stabilization of the two subunits. A linked BchL dimer with one defective ATP-binding site does not support protochlorophyllide reduction, illustrating nucleotide binding to both subunits as a prerequisite for the intersubunit cross stabilization. The masking of the [4Fe-4S] cluster by the flexible N-terminal region and the associated inhibition of the activity is a novel mechanism of regulation in metalloproteins. Such mechanisms are possibly an adaptation to the anaerobic nature of eubacterial cells with poor tolerance for oxygen.
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Affiliation(s)
- Elliot I Corless
- Department of Biological Sciences, Marquette University, Milwaukee, Wisconsin, USA; Department of Biochemistry, Saint Louis University School of Medicine, St Louis, Missouri, USA
| | | | - Maxwell B Watkins
- Department of Chemistry, Princeton University, Princeton, New Jersey, USA; Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York, USA
| | - John-Paul Bacik
- Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York, USA
| | - Jenna R Mattice
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana, USA
| | - Angela Patterson
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana, USA
| | - Karamatullah Danyal
- Department of Chemistry and Biochemistry, Utah State University, Logan, Utah, USA
| | - Mark Soffe
- Department of Chemistry and Biochemistry, Utah State University, Logan, Utah, USA
| | - Robert Kitelinger
- Department of Biological Sciences, Marquette University, Milwaukee, Wisconsin, USA
| | - Lance C Seefeldt
- Department of Chemistry and Biochemistry, Utah State University, Logan, Utah, USA
| | - Sofia Origanti
- Department of Biology, Saint Louis University, St Louis, Missouri, USA
| | - Brian Bennett
- Department of Physics, Marquette University, Milwaukee, Wisconsin, USA
| | - Brian Bothner
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana, USA
| | - Nozomi Ando
- Department of Chemistry, Princeton University, Princeton, New Jersey, USA; Department of Chemistry and Chemical Biology, Cornell University, Ithaca, New York, USA.
| | - Edwin Antony
- Department of Biological Sciences, Marquette University, Milwaukee, Wisconsin, USA; Department of Biochemistry, Saint Louis University School of Medicine, St Louis, Missouri, USA.
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35
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Ziegler SJ, Mallinson SJ, St. John PC, Bomble YJ. Advances in integrative structural biology: Towards understanding protein complexes in their cellular context. Comput Struct Biotechnol J 2020; 19:214-225. [PMID: 33425253 PMCID: PMC7772369 DOI: 10.1016/j.csbj.2020.11.052] [Citation(s) in RCA: 17] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/13/2020] [Revised: 11/25/2020] [Accepted: 11/28/2020] [Indexed: 01/26/2023] Open
Abstract
Microorganisms rely on protein interactions to transmit signals, react to stimuli, and grow. One of the best ways to understand these protein interactions is through structural characterization. However, in the past, structural knowledge was limited to stable, high-affinity complexes that could be crystallized. Recent developments in structural biology have revolutionized how protein interactions are characterized. The combination of multiple techniques, known as integrative structural biology, has provided insight into how large protein complexes interact in their native environment. In this mini-review, we describe the past, present, and potential future of integrative structural biology as a tool for characterizing protein interactions in their cellular context.
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Key Words
- CLEM, correlated light and electron microscopy
- Crosslinking mass spectrometry
- Cryo-electron microscopy
- Cryo-electron tomography
- EPR, electron paramagnetic resonance
- FRET, Forster resonance energy transfer
- ISB, Integrative structural biology
- Integrative structural biology
- ML, machine learning
- MR, molecular replacement
- MSAs, multiple sequence alignments
- MX, macromolecular crystallography
- NMR, nuclear magnetic resonance
- PDB, Protein Data Bank
- Protein docking
- Protein structure prediction
- Quinary interactions
- SAD, single-wavelength anomalous dispersion
- SANS, small angle neutron scattering
- SAXS, small angle X-ray scattering
- X-ray crystallography
- XL-MS, cross-linking mass spectrometry
- cryo-EM SPA, cryo-EM single particle analysis
- cryo-EM, cryo-electron microscopy
- cryo-ET, cryo-electron tomography
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Affiliation(s)
- Samantha J. Ziegler
- Biosciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
| | - Sam J.B. Mallinson
- Biosciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
| | - Peter C. St. John
- Biosciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
| | - Yannick J. Bomble
- Biosciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
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Hydrogen-deuterium exchange mass spectrometry captures distinct dynamics upon substrate and inhibitor binding to a transporter. Nat Commun 2020; 11:6162. [PMID: 33268777 PMCID: PMC7710758 DOI: 10.1038/s41467-020-20032-3] [Citation(s) in RCA: 27] [Impact Index Per Article: 6.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/05/2020] [Accepted: 10/15/2020] [Indexed: 11/13/2022] Open
Abstract
Proton-coupled transporters use transmembrane proton gradients to power active transport of nutrients inside the cell. High-resolution structures often fail to capture the coupling between proton and ligand binding, and conformational changes associated with transport. We combine HDX-MS with mutagenesis and MD simulations to dissect the molecular mechanism of the prototypical transporter XylE. We show that protonation of a conserved aspartate triggers conformational transition from outward-facing to inward-facing state. This transition only occurs in the presence of substrate xylose, while the inhibitor glucose locks the transporter in the outward-facing state. MD simulations corroborate the experiments by showing that only the combination of protonation and xylose binding, and not glucose, sets up the transporter for conformational switch. Overall, we demonstrate the unique ability of HDX-MS to distinguish between the conformational dynamics of inhibitor and substrate binding, and show that a specific allosteric coupling between substrate binding and protonation is a key step to initiate transport. XylE is a bacterial xylose transporter and homologue of human glucose transporters GLUTs 1-4. HDX-MS, mutagenesis and MD simulations suggest that protonation of a conserved aspartate triggers conformational transition from outward- to inward facing state only in the presence of substrate xylose. In contrast, inhibitor glucose locks the transporter in the outward facing state.
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37
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Gao S, Thompson EJ, Barrow SL, Zhang W, Iavarone AT, Klinman JP. Hydrogen-Deuterium Exchange within Adenosine Deaminase, a TIM Barrel Hydrolase, Identifies Networks for Thermal Activation of Catalysis. J Am Chem Soc 2020; 142:19936-19949. [PMID: 33181018 DOI: 10.1021/jacs.0c07866] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/27/2022]
Abstract
Proteins are intrinsically flexible macromolecules that undergo internal motions with time scales spanning femtoseconds to milliseconds. These fluctuations are implicated in the optimization of reaction barriers for enzyme catalyzed reactions. Time, temperature, and mutation dependent hydrogen-deuterium exchange coupled to mass spectrometry (HDX-MS) has been previously employed to identify spatially resolved, catalysis-linked dynamical regions of enzymes. We now extend this technique to pursue the correlation of protein flexibility and chemical reactivity within the diverse and widespread TIM barrel proteins, targeting murine adenosine deaminase (mADA) that catalyzes the irreversible deamination of adenosine to inosine and ammonia. Following a structure-function analysis of rate and activation energy for a series of mutations at a second sphere phenylalanine positioned in proximity to the bound substrate, the catalytically impaired Phe61Ala with an elevated activation energy (Ea = 7.5 kcal/mol) and the wild type (WT) mADA (Ea = 5.0 kcal/mol) were selected for HDX-MS experiments. The rate constants and activation energies of HDX for peptide segments are quantified and used to assess mutation-dependent changes in local and distal motions. Analyses reveal that approximately 50% of the protein sequence of Phe61Ala displays significant changes in the temperature dependence of HDX behaviors, with the dominant change being an increase in protein flexibility. Utilizing Phe61Ile, which displays the same activation energy for kcat as WT, as a control, we were able to further refine the HDX analysis, highlighting the regions of mADA that are altered in a functionally relevant manner. A map is constructed that illustrates the regions of protein that are proposed to be essential for the thermal optimization of active site configurations that dominate reaction barrier crossings in the native enzyme.
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Affiliation(s)
| | | | | | - Wenju Zhang
- David R. Cheriton School of Computer Science, University of Waterloo, Waterloo, ON N2L 3G1, Canada
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38
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Patterson A, Zhao Z, Waymire E, Zlotnick A, Bothner B. Dynamics of Hepatitis B Virus Capsid Protein Dimer Regulate Assembly through an Allosteric Network. ACS Chem Biol 2020; 15:2273-2280. [PMID: 32662972 DOI: 10.1021/acschembio.0c00481] [Citation(s) in RCA: 17] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/02/2023]
Abstract
While there is an effective vaccine for Human Hepatitis B Virus (HBV), 257 million people have chronic infections for which there is no cure. The assembly process for the viral capsid is a potential therapeutic target. In order to understand the capsid assembly process, we investigated the dimeric building blocks of the capsid. To understand what blocks assembly, we took advantage of an assembly incompetent mutant dimer, Cp149-Y132A, located in the interdimer interface. This mutation leads to changes in protein dynamics throughout the structure of the dimer as measured by hydrogen-deuterium exchange mass spectrometry (HDX-MS). To further understand how the HBV capsid assembles, the homologue woodchuck HBV (WHV) capsid protein dimer (Cp) was used. WHV is more stable than HBV in HDX-MS and native mass spectrometry experiments. Because the WHV Cp assembles more rapidly into viral capsids than HBV, it was suspected that an increase in stability of the intradimer interface and/or in the contact region leads to increased assembly rates. The differences in dynamics when comparing HBV and human Cp149-Y132A as well as the differences in dynamics when comparing the HBV and WHV Cps allowed us to map an allosteric network within the HBV dimer. Through a careful comparison of structure, stability, and dynamics using four different capsid protein dimers, we conclude that protein subunit dynamics regulate HBV capsid assembly.
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Affiliation(s)
- Angela Patterson
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana 59717, United States
| | - Zhongchao Zhao
- Department of Molecular and Cellular Biochemistry, Indiana University, Bloomington, Indiana 47405, United States
| | - Elizabeth Waymire
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana 59717, United States
| | - Adam Zlotnick
- Department of Molecular and Cellular Biochemistry, Indiana University, Bloomington, Indiana 47405, United States
| | - Brian Bothner
- Department of Chemistry and Biochemistry, Montana State University, Bozeman, Montana 59717, United States
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39
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Huang L, So PK, Chen YW, Leung YC, Yao ZP. Conformational Dynamics of the Helix 10 Region as an Allosteric Site in Class A β-Lactamase Inhibitory Binding. J Am Chem Soc 2020; 142:13756-13767. [PMID: 32686406 DOI: 10.1021/jacs.0c04088] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022]
Abstract
β-Lactamase inhibitory protein (BLIP) can effectively inactivate class A β-lactamases, but with very different degrees of potency. Understanding the different roles of BLIP in class A β-lactamases inhibition can provide insights for inhibitor design. However, this problem was poorly solved on the basis of the static structures obtained by X-ray crystallography. In this work, ion mobility mass spectrometry, hydrogen-deuterium exchange mass spectrometry, and molecular dynamics simulation revealed the conformational dynamics of three class A β-lactamases with varying inhibition efficiencies by BLIP. A more extended conformation of PC1 was shown compared to those of TEM1 and SHV1. Localized dynamics differed in several important loop regions, that is, the protruding loop, H10 loop, Ω loop, and SDN loop. Upon binding with BLIP, these loops cooperatively rearranged to enhance the binding interface and to inactivate the catalytic sites. In particular, unfavorable changes in conformational dynamics were found in the protruding loop of SHV1 and PC1, showing less effective binding. Intriguingly, the single mutation on BLIP could compensate for the unfavored changes in this region, and thus exhibit enhanced inhibition toward SHV1 and PC1. Additionally, the H10 region was revealed as an important allosteric site that could modulate the inhibition of class A β-lactamases. It was suggested that the rigid protruding loop and flexible H10 region might be determinants for the effective inhibition of TEM1. Our findings provided unique and explicit insights into the conformational dynamics of β-lactamases and their bindings with BLIP. This work can be extended to other β-lactamases of interest and inspire the design of novel inhibitors.
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Affiliation(s)
- Liwen Huang
- State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Food Safety and Technology Research Centre, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
| | - Pui-Kin So
- The University Research Facility in Life Sciences, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
| | - Yu Wai Chen
- State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
| | - Yun-Chung Leung
- State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Lo Ka Chung Research Centre for Natural Anti-Cancer Drug Development, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
| | - Zhong-Ping Yao
- State Key Laboratory of Chemical Biology and Drug Discovery, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Food Safety and Technology Research Centre, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China.,State Key Laboratory of Chinese Medicine and Molecular Pharmacology (Incubation) and Shenzhen Key Laboratory of Food Biological Safety Control, Shenzhen Research Institute of The Hong Kong Polytechnic University, Shenzhen 518057, China
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40
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Narang D, Lento C, J. Wilson D. HDX-MS: An Analytical Tool to Capture Protein Motion in Action. Biomedicines 2020; 8:biomedicines8070224. [PMID: 32709043 PMCID: PMC7399943 DOI: 10.3390/biomedicines8070224] [Citation(s) in RCA: 30] [Impact Index Per Article: 7.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/10/2020] [Revised: 07/13/2020] [Accepted: 07/15/2020] [Indexed: 01/07/2023] Open
Abstract
Virtually all protein functions in the cell, including pathogenic processes, require coordinated motion of atoms or domains, i.e., conformational dynamics. Understanding protein dynamics is therefore critical both for drug development and to learn about the underlying molecular causes of many diseases. Hydrogen–Deuterium Exchange Mass Spectrometry (HDX-MS) provides valuable information about protein dynamics, which is highly complementary to the static picture provided by conventional high-resolution structural tools (i.e., X-ray crystallography and structural NMR). The amount of protein required to carry out HDX-MS experiments is a fraction of the amount required by alternative biophysical techniques, which are also usually lower resolution. Use of HDX-MS is growing quickly both in industry and academia, and it has been successfully used in numerous drug and vaccine development efforts, with important roles in understanding allosteric effects and mapping binding sites.
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Affiliation(s)
- Dominic Narang
- Department of Chemistry, York University, Toronto, ON M3J 1P3, Canada; (D.N.); (C.L.)
| | - Cristina Lento
- Department of Chemistry, York University, Toronto, ON M3J 1P3, Canada; (D.N.); (C.L.)
| | - Derek J. Wilson
- Department of Chemistry, York University, Toronto, ON M3J 1P3, Canada; (D.N.); (C.L.)
- Centre for Research of Biomolecular Interactions, York University, Toronto, ON M3J 1P3, Canada
- Centre for Research in Mass Spectrometry, York University, Toronto, ON M3J 1P3, Canada
- Correspondence:
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41
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Xu S, Kennedy MA. Structural dynamics of pentapeptide repeat proteins. Proteins 2020; 88:1493-1512. [PMID: 32548861 DOI: 10.1002/prot.25969] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/22/2020] [Revised: 03/19/2020] [Accepted: 06/06/2020] [Indexed: 12/14/2022]
Abstract
Pentapeptide repeat proteins (PRPs) represent a large superfamily with more than 38 000 sequences in nearly 3500 species, the majority belonging to cyanobacteria but represented among all branches of life. PRPs contain at least eight consecutive pentapeptide repeats with the consensus (A/C/S/V/T/L/I)(D/N/S/K/E/I/R)(L/F)(S/T/R/E/Q/K/V/D)(G/D/E/N/R/Q/K). PRPs fold into right-handed quadrilateral β helices, also known as repeat-five-residue (Rfr)-folds, with four consecutive pentapeptide repeats comprising a single coil, the ~90° change in polypeptide direction in square-shaped coils achieved by type I, II and IV β turns, and hydrogen bonds between coils establishing β ladders on each Rfr-fold face. PRPs are broadly categorized into group 1 and 2 involved in antibiotic resistance and group 3 currently having unknown functions. Motivated by their intriguing structures, we are investigating PRP biophysical characteristics, including Rfr-fold thermal stability, β turn and β ladder hydrogen bond amide exchange rates and backbone dynamics. Here, we present analysis of 20 ns molecular dynamics (MD) simulations and all atom normal mode analysis (aaNMA) calculations for four group 1 and group 2 and four group 3 PRPs whose structures have been determined by X-ray crystallography. The MD cross-correlation matrices and aaNMA indicated strong correlated motion between adjacent coils and weak coupled motion between coils separated by one or more intervening coils. Slow anticorrelated motions were detected between adjacent coils in aaNMA modes that we hypothesize are requisite to access exchange-competent states necessary to permit solvent exchange of amide hydrogens involved in β-ladder and β-turns hydrogen bonds, which can have lifetimes on the order of months.
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Affiliation(s)
- Shenyuan Xu
- Department of Chemistry and Biochemistry, Miami University, Oxford, Ohio, USA
| | - Michael A Kennedy
- Department of Chemistry and Biochemistry, Miami University, Oxford, Ohio, USA
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42
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Min K, Yoon HJ, Park JY, Baidya M, Dwivedi-Agnihotri H, Maharana J, Chaturvedi M, Chung KY, Shukla AK, Lee HH. Crystal Structure of β-Arrestin 2 in Complex with CXCR7 Phosphopeptide. Structure 2020; 28:1014-1023.e4. [PMID: 32579945 DOI: 10.1016/j.str.2020.06.002] [Citation(s) in RCA: 29] [Impact Index Per Article: 7.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/29/2019] [Revised: 04/02/2020] [Accepted: 06/03/2020] [Indexed: 12/21/2022]
Abstract
β-Arrestins (βarrs) critically regulate G-protein-coupled receptor (GPCR) signaling and trafficking. βarrs have two isoforms, βarr1 and βarr2. Receptor phosphorylation is a key determinant for the binding of βarrs, and understanding the intricate details of receptor-βarr interaction is the next frontier in GPCR structural biology. The high-resolution structure of active βarr1 in complex with a phosphopeptide derived from GPCR has been revealed, but that of βarr2 remains elusive. Here, we present a 2.3-Å crystal structure of βarr2 in complex with a phosphopeptide (C7pp) derived from the carboxyl terminus of CXCR7. The structural analysis of C7pp-bound βarr2 reveals key differences from the previously determined active conformation of βarr1. One of the key differences is that C7pp-bound βarr2 shows a relatively small inter-domain rotation. Antibody-fragment-based conformational sensor and hydrogen/deuterium exchange experiments further corroborated the structural features of βarr2 and suggested that βarr2 adopts a range of inter-domain rotations.
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Affiliation(s)
- Kyungjin Min
- Department of Chemistry, College of Natural Sciences, Seoul National University, Seoul 08826, Republic of Korea
| | - Hye-Jin Yoon
- Department of Chemistry, College of Natural Sciences, Seoul National University, Seoul 08826, Republic of Korea
| | - Ji Young Park
- School of Pharmacy, Sungkyunkwan University, Suwon 16419, Republic of Korea
| | - Mithu Baidya
- Biological Sciences and Bioengineering, Indian Institute of Technology, Kanpur 208016, India
| | | | - Jagannath Maharana
- Biological Sciences and Bioengineering, Indian Institute of Technology, Kanpur 208016, India
| | - Madhu Chaturvedi
- Biological Sciences and Bioengineering, Indian Institute of Technology, Kanpur 208016, India
| | - Ka Young Chung
- School of Pharmacy, Sungkyunkwan University, Suwon 16419, Republic of Korea.
| | - Arun K Shukla
- Biological Sciences and Bioengineering, Indian Institute of Technology, Kanpur 208016, India.
| | - Hyung Ho Lee
- Department of Chemistry, College of Natural Sciences, Seoul National University, Seoul 08826, Republic of Korea.
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43
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An intact C-terminal end of albumin is required for its long half-life in humans. Commun Biol 2020; 3:181. [PMID: 32313072 PMCID: PMC7171077 DOI: 10.1038/s42003-020-0903-7] [Citation(s) in RCA: 37] [Impact Index Per Article: 9.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/30/2019] [Accepted: 03/17/2020] [Indexed: 12/19/2022] Open
Abstract
Albumin has an average plasma half-life of three weeks and is thus an attractive carrier to improve the pharmacokinetics of fused therapeutics. The half-life is regulated by FcRn, a cellular receptor that protects against intracellular degradation. To tailor-design the therapeutic use of albumin, it is crucial to understand how structural alterations in albumin affect FcRn binding and transport properties. In the blood, the last C-terminal residue (L585) of albumin may be enzymatically cleaved. Here we demonstrate that removal of the L585 residue causes structural stabilization in regions of the principal FcRn binding domain and reduces receptor binding. In line with this, a short half-life of only 3.5 days was measured for cleaved albumin lacking L585 in a patient with acute pancreatitis. Thus, we reveal the structural requirement of an intact C-terminal end of albumin for a long plasma half-life, which has implications for design of albumin-based therapeutics. Nilsen et al. show that structural alterations in the last C-terminal α-helix of albumin strongly reduce its binding to the neonatal Fc receptor, decreasing the half-life of albumin in humans. This study suggests the structural requirement of the C-terminal of albumin for its long plasma half-life, providing insights into the design of albumin used to carry drugs.
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44
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The Metastable XBP1u Transmembrane Domain Defines Determinants for Intramembrane Proteolysis by Signal Peptide Peptidase. Cell Rep 2020; 26:3087-3099.e11. [PMID: 30865896 DOI: 10.1016/j.celrep.2019.02.057] [Citation(s) in RCA: 22] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/21/2018] [Revised: 12/07/2018] [Accepted: 02/14/2019] [Indexed: 11/23/2022] Open
Abstract
Unspliced XBP1 mRNA encodes XBP1u, the transcriptionally inert variant of the unfolded protein response (UPR) transcription factor XBP1s. XBP1u targets its mRNA-ribosome-nascent-chain-complex to the endoplasmic reticulum (ER) to facilitate UPR activation and prevents overactivation. Yet, its membrane association is controversial. Here, we use cell-free translocation and cellular assays to define a moderately hydrophobic stretch in XBP1u that is sufficient to mediate insertion into the ER membrane. Mutagenesis of this transmembrane (TM) region reveals residues that facilitate XBP1u turnover by an ER-associated degradation route that is dependent on signal peptide peptidase (SPP). Furthermore, the impact of these mutations on TM helix dynamics was assessed by residue-specific amide exchange kinetics, evaluated by a semi-automated algorithm. Based on our results, we suggest that SPP-catalyzed intramembrane proteolysis of TM helices is not only determined by their conformational flexibility, but also by side-chain interactions near the scissile peptide bond with the enzyme's active site.
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45
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Makarov AA, Iacob RE, Pirrone GF, Rodriguez-Granillo A, Joyce L, Mangion I, Moore JC, Sherer EC, Engen JR. Combination of HDX-MS and in silico modeling to study enzymatic reactivity and stereo-selectivity at different solvent conditions. J Pharm Biomed Anal 2020; 182:113141. [PMID: 32036298 DOI: 10.1016/j.jpba.2020.113141] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/09/2019] [Revised: 01/29/2020] [Accepted: 01/31/2020] [Indexed: 12/16/2022]
Abstract
The higher-order structure of a protein defines its function, and protein structural dynamics are often essential for protein binding and enzyme catalysis. Methods for protein characterization in solution are continuously being developed to understand and explore protein conformational changes with regards to function and activity. The goal of this study was to survey the use of combining HDX-MS global conformational screening with in silico modeling and continuous labeling peptide-level HDX-MS as an approach to highlight regions of interest within an enzyme required for biocatalytic processes. We surveyed in silico modeling correlated with peptide level HDX-MS experiments to characterize and localize transaminase enzyme structural dynamics at different conditions. This approach was orthogonally correlated with a global Size-Exclusion-HDX (SEC-HDX) screen for global conformational comparison and global alpha-helical content measurements by circular dichroism. Enzymatic activity and stereo-selectivity of transaminases were compared at different reaction-solution conditions that forced protein conformational changes by increasing acetonitrile concentration. The experimental peptide-level HDX-MS results demonstrated similar trends to the modeling data showing that certain regions remained folded in transaminases ATA-036 and ATA-303 with increasing acetonitrile concentration, which is also associated with shifting stereoselectivity. HDX modeling, SEC-HDX and CD experimental data showed that transaminase ATA-234 had the highest level of global unfolding with increasing acetonitrile concentration compared to the other two enzymes, which correlated with drastically reduced product conversion in transamination reaction. The combined HDX modeling/experimental workflow, based on enzymatic reactions studied at different conditions to induce changes in enzyme conformation, could be used as a tool to guide directed evolution efforts by identifying and focusing on the regions of an enzyme required for reaction product conversion and stereoselectivity.
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Affiliation(s)
- Alexey A Makarov
- Merck & Co., Inc., Process Research & Development, Rahway, NJ 07065, USA.
| | - Roxana E Iacob
- Department of Chemistry and Chemical Biology, Northeastern University, Boston, MA 02115, USA
| | - Gregory F Pirrone
- Merck & Co., Inc., Process Research & Development, Rahway, NJ 07065, USA
| | | | - Leo Joyce
- Merck & Co., Inc., Process Research & Development, Rahway, NJ 07065, USA
| | - Ian Mangion
- Merck & Co., Inc., Process Research & Development, Rahway, NJ 07065, USA
| | - Jeffrey C Moore
- Merck & Co., Inc., Process Research & Development, Rahway, NJ 07065, USA
| | - Edward C Sherer
- Merck & Co., Inc., Computational and Structural Chemistry, Rahway, NJ 07065, USA
| | - John R Engen
- Department of Chemistry and Chemical Biology, Northeastern University, Boston, MA 02115, USA
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46
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Berman HM, Adams PD, Bonvin AA, Burley SK, Carragher B, Chiu W, DiMaio F, Ferrin TE, Gabanyi MJ, Goddard TD, Griffin PR, Haas J, Hanke CA, Hoch JC, Hummer G, Kurisu G, Lawson CL, Leitner A, Markley JL, Meiler J, Montelione GT, Phillips GN, Prisner T, Rappsilber J, Schriemer DC, Schwede T, Seidel CAM, Strutzenberg TS, Svergun DI, Tajkhorshid E, Trewhella J, Vallat B, Velankar S, Vuister GW, Webb B, Westbrook JD, White KL, Sali A. Federating Structural Models and Data: Outcomes from A Workshop on Archiving Integrative Structures. Structure 2019; 27:1745-1759. [PMID: 31780431 DOI: 10.1016/j.str.2019.11.002] [Citation(s) in RCA: 33] [Impact Index Per Article: 6.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/20/2019] [Revised: 10/31/2019] [Accepted: 11/06/2019] [Indexed: 12/23/2022]
Abstract
Structures of biomolecular systems are increasingly computed by integrative modeling. In this approach, a structural model is constructed by combining information from multiple sources, including varied experimental methods and prior models. In 2019, a Workshop was held as a Biophysical Society Satellite Meeting to assess progress and discuss further requirements for archiving integrative structures. The primary goal of the Workshop was to build consensus for addressing the challenges involved in creating common data standards, building methods for federated data exchange, and developing mechanisms for validating integrative structures. The summary of the Workshop and the recommendations that emerged are presented here.
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Affiliation(s)
- Helen M Berman
- Department of Chemistry and Chemical Biology, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA; Department of Biological Sciences, University of Southern California, Los Angeles, CA 90089, USA; Bridge Institute, Michelson Center, University of Southern California, Los Angeles, CA 90089, USA.
| | - Paul D Adams
- Physical Biosciences Division, Lawrence Berkeley Laboratory, Berkeley, CA 94720-8235, USA; Department of Bioengineering, University of California-Berkeley, Berkeley, CA 94720, USA
| | - Alexandre A Bonvin
- Bijvoet Center for Biomolecular Research, Faculty of Science - Chemistry, Utrecht University, Padualaan 8, 3584 CH Utrecht, the Netherlands
| | - Stephen K Burley
- Research Collaboratory for Structural Bioinformatics Protein Data Bank, The State University of New Jersey, Piscataway, NJ 08854, USA; Institute for Quantitative Biomedicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA; Skaggs School of Pharmacy and Pharmaceutical Sciences and San Diego Supercomputer Center, University of California, San Diego, La Jolla, CA 92093, USA; Rutgers Cancer Institute of New Jersey, Rutgers, The State University of New Jersey, New Brunswick, NJ 08903, USA
| | - Bridget Carragher
- Simons Electron Microscopy Center, New York Structural Biology Center, New York, NY 10027, USA; Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY 10032, USA
| | - Wah Chiu
- Department of Bioengineering, Department of Microbiology and Immunology, Stanford University, Stanford, CA 94305-5447, USA; SLAC National Accelerator Laboratory, Menlo Park, CA 94025, USA
| | - Frank DiMaio
- Department of Biochemistry and Institute for Protein Design, University of Washington, Seattle, WA 98195, USA
| | - Thomas E Ferrin
- Department of Pharmaceutical Chemistry, University of California, San Francisco, CA 94158, USA
| | - Margaret J Gabanyi
- Institute for Quantitative Biomedicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA
| | - Thomas D Goddard
- Department of Pharmaceutical Chemistry, University of California, San Francisco, CA 94158, USA
| | | | - Juergen Haas
- Swiss Institute of Bioinformatics and Biozentrum, University of Basel, 4056 Basel, Switzerland
| | - Christian A Hanke
- Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | - Jeffrey C Hoch
- Department of Molecular Biology and Biophysics, UConn Health, Farmington, CT 06030, USA
| | - Gerhard Hummer
- Department of Theoretical Biophysics, Max Planck Institute of Biophysics, 60438 Frankfurt am Main, Germany; Institute for Biophysics, Goethe University Frankfurt, 60438 Frankfurt am Main, Germany
| | - Genji Kurisu
- Protein Data Bank Japan (PDBj), Institute for Protein Research, Osaka University, Osaka 565-0871, Japan
| | - Catherine L Lawson
- Institute for Quantitative Biomedicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA
| | - Alexander Leitner
- Department of Biology, Institute of Molecular Systems Biology, ETH Zurich, 8093 Zurich, Switzerland
| | - John L Markley
- BioMagResBank (BMRB), Biochemistry Department, University of Wisconsin-Madison, Madison, WI 53706, USA
| | - Jens Meiler
- Center for Structural Biology, Vanderbilt University, 465 21st Avenue South, Nashville, TN 37221, USA
| | - Gaetano T Montelione
- Center for Advanced Biotechnology and Medicine, Department of Molecular Biology and Biochemistry, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA; Department of Biochemistry, Robert Wood Johnson Medical School, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA; Center for Biotechnology and Interdisciplinary Studies, Rensselaer Polytech Institute, Troy, NY 12180, USA
| | - George N Phillips
- BioSciences at Rice and Department of Chemistry, Rice University, Houston, TX 77251, USA
| | - Thomas Prisner
- Institute of Physical and Theoretical Chemistry and Center of Biomolecular Magnetic Resonance, Goethe University Frankfurt, 60438 Frankfurt am Main, Germany
| | - Juri Rappsilber
- Wellcome Trust Centre for Cell Biology, Edinburgh EH9 3JR, Scotland
| | - David C Schriemer
- Department of Biochemistry & Molecular Biology, Robson DNA Science Centre, University of Calgary, Calgary, AB T2N 4N1, Canada
| | - Torsten Schwede
- Swiss Institute of Bioinformatics and Biozentrum, University of Basel, 4056 Basel, Switzerland
| | - Claus A M Seidel
- Molecular Physical Chemistry, Heinrich Heine University Düsseldorf, 40225 Düsseldorf, Germany
| | | | - Dmitri I Svergun
- European Molecular Biology Laboratory (EMBL), Hamburg Outstation, Notkestrasse 85, 22607 Hamburg, Germany
| | - Emad Tajkhorshid
- Department of Biochemistry, NIH Center for Macromolecular Modeling and Bioinformatics, Center for Biophysics and Quantitative Biology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA; Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
| | - Jill Trewhella
- School of Life and Environmental Sciences, The University of Sydney, Sydney, NSW 2006, Australia; Department of Chemistry, University of Utah, Salt Lake City, UT 84112, USA
| | - Brinda Vallat
- Institute for Quantitative Biomedicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA
| | - Sameer Velankar
- Protein Data Bank in Europe (PDBe), European Molecular Biology Laboratory, European Bioinformatics Institute (EMBL-EBI), Hinxton, Cambridgeshire CB10 1SD, UK
| | - Geerten W Vuister
- Department of Molecular and Cell Biology, Leicester Institute of Structural and Chemical Biology, University of Leicester, Leicester LE1 9HN, UK
| | - Benjamin Webb
- Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA 94158, USA
| | - John D Westbrook
- Research Collaboratory for Structural Bioinformatics Protein Data Bank, The State University of New Jersey, Piscataway, NJ 08854, USA; Institute for Quantitative Biomedicine, Rutgers, The State University of New Jersey, Piscataway, NJ 08854, USA
| | - Kate L White
- Department of Biological Sciences, University of Southern California, Los Angeles, CA 90089, USA; Bridge Institute, Michelson Center, University of Southern California, Los Angeles, CA 90089, USA
| | - Andrej Sali
- Department of Pharmaceutical Chemistry, University of California, San Francisco, CA 94158, USA; Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, San Francisco, CA 94158, USA; California Institute for Quantitative Biosciences, University of California, San Francisco, San Francisco, CA 94158, USA.
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Abstract
Although many proteins possess a distinct folded structure lying at a minimum in a funneled free energy landscape, thermal energy causes any protein to continuously access lowly populated excited states. The existence of excited states is an integral part of biological function. Although transitions into the excited states may lead to protein misfolding and aggregation, little structural information is currently available for them. Here, we show how NMR spectroscopy, coupled with pressure perturbation, brings these elusive species to light. As pressure acts to favor states with lower partial molar volume, NMR follows the ensuing change in the equilibrium spectroscopically, with residue-specific resolution. For T4 lysozyme L99A, relaxation dispersion NMR was used to follow the increase in population of a previously identified "invisible" folded state with pressure, as this is driven by the reduction in cavity volume by the flipping-in of a surface aromatic group. Furthermore, multiple partly disordered excited states were detected at equilibrium using pressure-dependent H/D exchange NMR spectroscopy. Here, unfolding reduced partial molar volume by the removal of empty internal cavities and packing imperfections through subglobal and global unfolding. A close correspondence was found for the distinct pressure sensitivities of various parts of the protein and the amount of internal cavity volume that was lost in each unfolding event. The free energies and populations of excited states allowed us to determine the energetic penalty of empty internal protein cavities to be 36 cal⋅Å-3.
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48
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Klein SA, Majumdar A, Barrick D. A Second Backbone: The Contribution of a Buried Asparagine Ladder to the Global and Local Stability of a Leucine-Rich Repeat Protein. Biochemistry 2019; 58:3480-3493. [PMID: 31347358 PMCID: PMC7184636 DOI: 10.1021/acs.biochem.9b00355] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/30/2022]
Abstract
Parallel β-sheet-containing repeat proteins often display a structural motif in which conserved asparagines form a continuous ladder buried within the hydrophobic core. In such "asparagine ladders", the asparagine side-chain amides form a repetitive pattern of hydrogen bonds with neighboring main-chain NH and CO groups. Although asparagine ladders have been thought to be important for stability, there is little experimental evidence to support such speculation. Here we test the contribution of a minimal asparagine ladder from the leucine-rich repeat protein pp32 to stability and investigate lattice rigidity and hydrogen bond character using solution nuclear magnetic resonance (NMR) spectroscopy. Point substitutions of the two ladder asparagines of pp32 are strongly destabilizing and decrease the cooperativity of unfolding. The chemical shifts of the ladder side-chain HZ protons are shifted significantly downfield in the NMR spectrum and have low temperature coefficients, indicative of strong hydrogen bonding. In contrast, the HE protons are shifted upfield and have temperature coefficients close to zero, suggesting an asymmetry in hydrogen bond strength along the ladder. Ladder NH2 groups have weak 1H-15N cross-peak intensities; 1H-15N nuclear Overhauser effect and 15N CPMG experiments show this to be the result of high rigidity. Hydrogen exchange measurements demonstrate that the ladder NH2 groups exchange very slowly, with rates approaching the global exchange limit. Overall, these results show that the asparagine side chains are held in a very rigid, nondynamic structure, making a significant contribution to the overall stability. In this regard, buried asparagine ladders can be considered "second backbones" within the cores of their elongated β-sheet host proteins.
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Affiliation(s)
- Sean A. Klein
- T.C. Jenkins Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218 USA
| | - Ananya Majumdar
- The Johns Hopkins University Biomolecular NMR Center, Johns Hopkins University, Baltimore, Maryland, 21218
| | - Doug Barrick
- T.C. Jenkins Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218 USA
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49
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Huang L, So PK, Yao ZP. Protein dynamics revealed by hydrogen/deuterium exchange mass spectrometry: Correlation between experiments and simulation. RAPID COMMUNICATIONS IN MASS SPECTROMETRY : RCM 2019; 33 Suppl 3:83-89. [PMID: 30321473 DOI: 10.1002/rcm.8307] [Citation(s) in RCA: 16] [Impact Index Per Article: 3.2] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 06/12/2018] [Revised: 10/08/2018] [Accepted: 10/08/2018] [Indexed: 05/09/2023]
Abstract
Hydrogen deuterium exchange mass spectrometry (HDX-MS) is a powerful technique for studying protein dynamics, which is an important factor governing protein functions. However, the process of hydrogen/deuterium exchange (HDX) of proteins is highly complex and the underlying mechanism has not yet been fully elucidated. Meanwhile, molecular dynamics (MD) simulation is a computational technique that can be used to elucidate HDX behaviour on proteins and facilitate interpretation of HDX-MS data. This article aims to summarize the current understandings on the mechanism of HDX and its correlation with MD simulation, to discuss the recent developments in the techniques of HDX-MS and MD simulation and to extend the perspectives of these two techniques in protein dynamics study.
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Affiliation(s)
- Liwen Huang
- State Key Laboratory of Chemical Biology and Drug Discovery, Food Safety and Technology Research Centre and Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
- State Key Laboratory of Chinese Medicine and Molecular Pharmacology (Incubation) and Shenzhen Key Laboratory of Food Biological Safety Control, Shenzhen Research Institute of The Hong Kong Polytechnic University, Shenzhen, 518057, China
| | - Pui-Kin So
- State Key Laboratory of Chemical Biology and Drug Discovery, Food Safety and Technology Research Centre and Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
- State Key Laboratory of Chinese Medicine and Molecular Pharmacology (Incubation) and Shenzhen Key Laboratory of Food Biological Safety Control, Shenzhen Research Institute of The Hong Kong Polytechnic University, Shenzhen, 518057, China
| | - Zhong-Ping Yao
- State Key Laboratory of Chemical Biology and Drug Discovery, Food Safety and Technology Research Centre and Department of Applied Biology and Chemical Technology, The Hong Kong Polytechnic University, Hung Hom, Kowloon, Hong Kong Special Administrative Region, China
- State Key Laboratory of Chinese Medicine and Molecular Pharmacology (Incubation) and Shenzhen Key Laboratory of Food Biological Safety Control, Shenzhen Research Institute of The Hong Kong Polytechnic University, Shenzhen, 518057, China
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50
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Yamamoto K. Discovery of Nuclear Receptor Ligands and Elucidation of Their Mechanisms of Action. Chem Pharm Bull (Tokyo) 2019; 67:609-619. [DOI: 10.1248/cpb.c19-00131] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/22/2022]
Affiliation(s)
- Keiko Yamamoto
- Laboratory of Drug Design and Medicinal Chemistry, Showa Pharmaceutical University
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