1
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Bondar AN. Interplay between local protein interactions and water bridging of a proton antenna carboxylate cluster. BIOCHIMICA ET BIOPHYSICA ACTA. BIOMEMBRANES 2022; 1864:184052. [PMID: 36116514 DOI: 10.1016/j.bbamem.2022.184052] [Citation(s) in RCA: 5] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 04/16/2022] [Revised: 08/17/2022] [Accepted: 09/11/2022] [Indexed: 06/15/2023]
Abstract
Proteins that bind protons at cell membrane interfaces often expose to the bulk clusters of carboxylate and histidine sidechains that capture protons transiently and, in proton transporters, deliver protons to an internal site. The protonation-coupled dynamics of bulk-exposed carboxylate clusters, also known as proton antennas, is poorly described. An essential open question is how water-mediated bridges between sidechains of the cluster respond to protonation change and facilitate transient proton storage. To address this question, here I studied the protonation-coupled dynamics at the proton-binding antenna of PsbO, a small extrinsinc subunit of the photosystem II complex, with atomistic molecular dynamics simulations and systematic graph-based analyses of dynamic protein and protein-water hydrogen-bond networks. The protonation of specific carboxylate groups is found to impact the dynamics of their local protein-water hydrogen-bond clusters. Regardless of the protonation state considered for PsbO, carboxylate pairs that can sample direct hydrogen bonding, or bridge via short hydrogen-bonded water chains, anchor to nearby basic or polar protein sidechains. As a result, carboxylic sidechains of the hypothesized antenna cluster are part of dynamic hydrogen bond networks that may rearrange rapidly when the protonation changes.
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Affiliation(s)
- Ana-Nicoleta Bondar
- University of Bucharest, Faculty of Physics, Str. Atomiştilor 405, Bucharest-Măgurele 077125, Romania; Forschungszentrum Jülich, Institute for Neuroscience and Medicine and Institute for Advanced Simulations (IAS-5/INM-9), Computational Biomedicine, 52425 Jülich, Germany; Freie Universität Berlin, Department of Physics, Theoretical Molecular Biophysics Group, Arnimallee 14, D-14195 Berlin, Germany.
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2
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Local Attraction of Substrates and Co-Substrates Enhances Weak Acid and Base Transmembrane Transport. Biomolecules 2022; 12:biom12121794. [PMID: 36551222 PMCID: PMC9775063 DOI: 10.3390/biom12121794] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/14/2022] [Revised: 11/25/2022] [Accepted: 11/28/2022] [Indexed: 12/03/2022] Open
Abstract
The transmembrane transport of weak acid and base metabolites depends on the local pH conditions that affect the protonation status of the substrates and the availability of co-substrates, typically protons. Different protein designs ensure the attraction of substrates and co-substrates to the transporter entry sites. These include electrostatic surface charges on the transport proteins and complexation with seemingly transport-unrelated proteins that provide substrate and/or proton antenna, or enzymatically generate substrates in place. Such protein assemblies affect transport rates and directionality. The lipid membrane surface also collects and transfers protons. The complexity in the various systems enables adjustability and regulation in a given physiological or pathophysiological situation. This review describes experimentally shown principles in the attraction and facilitation of weak acid and base transport substrates, including monocarboxylates, ammonium, bicarbonate, and arsenite, plus protons as a co-substrate.
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3
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Du Z, Piguet J, Baryshnikov G, Tornmalm J, Demirbay B, Ågren H, Widengren J. Imaging Fluorescence Blinking of a Mitochondrial Localization Probe: Cellular Localization Probes Turned into Multifunctional Sensors. J Phys Chem B 2022; 126:3048-3058. [PMID: 35417173 PMCID: PMC9059120 DOI: 10.1021/acs.jpcb.2c01271] [Citation(s) in RCA: 5] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/22/2022] [Revised: 03/28/2022] [Indexed: 11/29/2022]
Abstract
Mitochondrial membranes and their microenvironments directly influence and reflect cellular metabolic states but are difficult to probe on site in live cells. Here, we demonstrate a strategy, showing how the widely used mitochondrial membrane localization fluorophore 10-nonyl acridine orange (NAO) can be transformed into a multifunctional probe of membrane microenvironments by monitoring its blinking kinetics. By transient state (TRAST) studies of NAO in small unilamellar vesicles (SUVs), together with computational simulations, we found that NAO exhibits prominent reversible singlet-triplet state transitions and can act as a light-induced Lewis acid forming a red-emissive doublet radical. The resulting blinking kinetics are highly environment-sensitive, specifically reflecting local membrane oxygen concentrations, redox conditions, membrane charge, fluidity, and lipid compositions. Here, not only cardiolipin concentration but also the cardiolipin acyl chain composition was found to strongly influence the NAO blinking kinetics. The blinking kinetics also reflect hydroxyl ion-dependent transitions to and from the fluorophore doublet radical, closely coupled to the proton-transfer events in the membranes, local pH, and two- and three-dimensional buffering properties on and above the membranes. Following the SUV studies, we show by TRAST imaging that the fluorescence blinking properties of NAO can be imaged in live cells in a spatially resolved manner. Generally, the demonstrated blinking imaging strategy can transform existing fluorophore markers into multiparametric sensors reflecting conditions of large biological relevance, which are difficult to retrieve by other means. This opens additional possibilities for fundamental membrane studies in lipid vesicles and live cells.
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Affiliation(s)
- Zhixue Du
- Royal
Institute of Technology (KTH), Experimental Biomolecular Physics,
Department Applied Physics, Albanova Univ
Center, 106 91 Stockholm, Sweden
| | - Joachim Piguet
- Royal
Institute of Technology (KTH), Experimental Biomolecular Physics,
Department Applied Physics, Albanova Univ
Center, 106 91 Stockholm, Sweden
| | - Glib Baryshnikov
- Laboratory
of Organic Electronics, Department of Science and Technology, Linköping University, SE-60174 Norrköping, Sweden
| | - Johan Tornmalm
- Royal
Institute of Technology (KTH), Experimental Biomolecular Physics,
Department Applied Physics, Albanova Univ
Center, 106 91 Stockholm, Sweden
| | - Baris Demirbay
- Royal
Institute of Technology (KTH), Experimental Biomolecular Physics,
Department Applied Physics, Albanova Univ
Center, 106 91 Stockholm, Sweden
| | - Hans Ågren
- Department
of Physics and Astronomy, Uppsala University, P.O. Box 516, SE-751 20 Uppsala, Sweden
| | - Jerker Widengren
- Royal
Institute of Technology (KTH), Experimental Biomolecular Physics,
Department Applied Physics, Albanova Univ
Center, 106 91 Stockholm, Sweden
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4
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Laurence MJ, Carpenter TS, Laurence TA, Coleman MA, Shelby M, Liu C. Biophysical Characterization of Membrane Proteins Embedded in Nanodiscs Using Fluorescence Correlation Spectroscopy. MEMBRANES 2022; 12:membranes12040392. [PMID: 35448362 PMCID: PMC9028781 DOI: 10.3390/membranes12040392] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 03/05/2022] [Revised: 03/29/2022] [Accepted: 03/30/2022] [Indexed: 02/04/2023]
Abstract
Proteins embedded in biological membranes perform essential functions in all organisms, serving as receptors, transporters, channels, cell adhesion molecules, and other supporting cellular roles. These membrane proteins comprise ~30% of all human proteins and are the targets of ~60% of FDA-approved drugs, yet their extensive characterization using established biochemical and biophysical methods has continued to be elusive due to challenges associated with the purification of these insoluble proteins. In response, the development of nanodisc techniques, such as nanolipoprotein particles (NLPs) and styrene maleic acid polymers (SMALPs), allowed membrane proteins to be expressed and isolated in solution as part of lipid bilayer rafts with defined, consistent nanometer sizes and compositions, thus enabling solution-based measurements. Fluorescence correlation spectroscopy (FCS) is a relatively simple yet powerful optical microscopy-based technique that yields quantitative biophysical information, such as diffusion kinetics and concentrations, about individual or interacting species in solution. Here, we first summarize current nanodisc techniques and FCS fundamentals. We then provide a focused review of studies that employed FCS in combination with nanodisc technology to investigate a handful of membrane proteins, including bacteriorhodopsin, bacterial division protein ZipA, bacterial membrane insertases SecYEG and YidC, Yersinia pestis type III secretion protein YopB, yeast cell wall stress sensor Wsc1, epidermal growth factor receptor (EGFR), ABC transporters, and several G protein-coupled receptors (GPCRs).
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Affiliation(s)
- Matthew J. Laurence
- Biosciences and Biotechnology Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA; (M.J.L.); (T.S.C.); (M.A.C.)
| | - Timothy S. Carpenter
- Biosciences and Biotechnology Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA; (M.J.L.); (T.S.C.); (M.A.C.)
| | - Ted A. Laurence
- Materials Science Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA;
| | - Matthew A. Coleman
- Biosciences and Biotechnology Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA; (M.J.L.); (T.S.C.); (M.A.C.)
- Department of Radiation Oncology, University of California Davis, Sacramento, CA 95616, USA
| | - Megan Shelby
- Biosciences and Biotechnology Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA; (M.J.L.); (T.S.C.); (M.A.C.)
- Correspondence: (M.S.); (C.L.)
| | - Chao Liu
- Biosciences and Biotechnology Division, Lawrence Livermore National Laboratory, Livermore, CA 94550, USA; (M.J.L.); (T.S.C.); (M.A.C.)
- Correspondence: (M.S.); (C.L.)
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5
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Kell DB. A protet-based, protonic charge transfer model of energy coupling in oxidative and photosynthetic phosphorylation. Adv Microb Physiol 2021; 78:1-177. [PMID: 34147184 DOI: 10.1016/bs.ampbs.2021.01.001] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/30/2022]
Abstract
Textbooks of biochemistry will explain that the otherwise endergonic reactions of ATP synthesis can be driven by the exergonic reactions of respiratory electron transport, and that these two half-reactions are catalyzed by protein complexes embedded in the same, closed membrane. These views are correct. The textbooks also state that, according to the chemiosmotic coupling hypothesis, a (or the) kinetically and thermodynamically competent intermediate linking the two half-reactions is the electrochemical difference of protons that is in equilibrium with that between the two bulk phases that the coupling membrane serves to separate. This gradient consists of a membrane potential term Δψ and a pH gradient term ΔpH, and is known colloquially as the protonmotive force or pmf. Artificial imposition of a pmf can drive phosphorylation, but only if the pmf exceeds some 150-170mV; to achieve in vivo rates the imposed pmf must reach 200mV. The key question then is 'does the pmf generated by electron transport exceed 200mV, or even 170mV?' The possibly surprising answer, from a great many kinds of experiment and sources of evidence, including direct measurements with microelectrodes, indicates it that it does not. Observable pH changes driven by electron transport are real, and they control various processes; however, compensating ion movements restrict the Δψ component to low values. A protet-based model, that I outline here, can account for all the necessary observations, including all of those inconsistent with chemiosmotic coupling, and provides for a variety of testable hypotheses by which it might be refined.
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Affiliation(s)
- Douglas B Kell
- Department of Biochemistry and Systems Biology, Institute of Systems, Molecular and Integrative, Biology, University of Liverpool, Liverpool, United Kingdom; The Novo Nordisk Foundation Center for Biosustainability, Technical University of Denmark, Lyngby, Denmark.
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6
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Meikle TG, Keizer DW, Babon JJ, Drummond CJ, Separovic F, Conn CE, Yao S. Chemical Exchange of Hydroxyl Groups in Lipidic Cubic Phases Characterized by NMR. J Phys Chem B 2021; 125:571-580. [PMID: 33251799 DOI: 10.1021/acs.jpcb.0c08699] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/04/2023]
Abstract
Proton transportation in proximity to the lipid bilayer membrane surface, where chemical exchange represents a primary pathway, is of significant interest in many applications including cellular energy turnover underlying ATP synthesis, transmembrane mobility, and transport. Lipidic inverse bicontinuous cubic phases (LCPs) are unique membrane structures formed via the spontaneous self-assembly of certain lipids in an aqueous environment. They feature two networks of water channels, separated by a single lipid bilayer which approximates the geometry of a triply periodic minimal surface. When composed of monoolein, the LCP bilayer features two glycerol hydroxyl groups at the lipid-water interface which undergo exchange with water. Depending on the conditions of the aqueous solution used in the formation of LCPs, both resonances of the glycerol hydroxyl groups may be observed by solution 1H NMR. In this study, PFG-NMR and 1D EXSY were employed to gain insight into chemical exchange between the monoolein hydroxyl groups and water in LCPs. Results including the relative population of hydroxyl protons in exchange with water for a number of LCPs at different hydration levels and the exchange rate constants at 35 wt % hydration are reported. Several technical aspects of PFG-NMR and EXSY-NMR for the characterization of chemical exchange in LCPs are discussed, including an alternative way to analyze PFG-NMR data of exchange systems which overcomes the inherent low sensitivity at high diffusion encoding.
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Affiliation(s)
- Thomas G Meikle
- School of Science, College of Science, Engineering and Health, RMIT University, Melbourne, VIC 3000, Australia
| | - David W Keizer
- Bio21 Molecular Science and Biotechnology Institute, The University of Melbourne, Melbourne, VIC 3010, Australia
| | - Jeffrey J Babon
- The Walter and Eliza Hall Institute of Medical Research, Parkville, VIC 3052, Australia.,Department of Medical Biology, The University of Melbourne, Melbourne, VIC 3010, Australia
| | - Calum J Drummond
- School of Science, College of Science, Engineering and Health, RMIT University, Melbourne, VIC 3000, Australia
| | - Frances Separovic
- Bio21 Molecular Science and Biotechnology Institute, The University of Melbourne, Melbourne, VIC 3010, Australia.,School of Chemistry, The University of Melbourne, Melbourne, VIC 3010, Australia
| | - Charlotte E Conn
- School of Science, College of Science, Engineering and Health, RMIT University, Melbourne, VIC 3000, Australia
| | - Shenggen Yao
- Bio21 Molecular Science and Biotechnology Institute, The University of Melbourne, Melbourne, VIC 3010, Australia
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7
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Sligar SG, Denisov IG. Nanodiscs: A toolkit for membrane protein science. Protein Sci 2020; 30:297-315. [PMID: 33165998 DOI: 10.1002/pro.3994] [Citation(s) in RCA: 74] [Impact Index Per Article: 18.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/22/2020] [Revised: 10/30/2020] [Accepted: 10/30/2020] [Indexed: 12/25/2022]
Abstract
Membrane proteins are involved in numerous vital biological processes, including transport, signal transduction and the enzymes in a variety of metabolic pathways. Integral membrane proteins account for up to 30% of the human proteome and they make up more than half of all currently marketed therapeutic targets. Unfortunately, membrane proteins are inherently recalcitrant to study using the normal toolkit available to scientists, and one is most often left with the challenge of finding inhibitors, activators and specific antibodies using a denatured or detergent solubilized aggregate. The Nanodisc platform circumvents these challenges by providing a self-assembled system that renders typically insoluble, yet biologically and pharmacologically significant, targets such as receptors, transporters, enzymes, and viral antigens soluble in aqueous media in a native-like bilayer environment that maintain a target's functional activity. By providing a bilayer surface of defined composition and structure, Nanodiscs have found great utility in the study of cellular signaling complexes that assemble on a membrane surface. Nanodiscs provide a nanometer scale vehicle for the in vivo delivery of amphipathic drugs, therapeutic lipids, tethered nucleic acids, imaging agents and active protein complexes. This means for generating nanoscale lipid bilayers has spawned the successful use of numerous other polymer and peptide amphipathic systems. This review, in celebration of the Anfinsen Award, summarizes some recent results and provides an inroad into the current and historical literature.
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Affiliation(s)
- Stephen G Sligar
- Departments of Biochemistry Chemistry, University of Illinois, Urbana-Champaign, Urbana, Illinois, USA
| | - Ilia G Denisov
- Departments of Biochemistry Chemistry, University of Illinois, Urbana-Champaign, Urbana, Illinois, USA
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8
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Abstract
The interactions between lipids and proteins are one of the most fundamental processes in living organisms, responsible for critical cellular events ranging from replication, cell division, signaling, and movement. Enabling the central coupling responsible for maintaining the functionality of the breadth of proteins, receptors, and enzymes that find their natural home in biological membranes, the fundamental mechanisms of recognition of protein for lipid, and vice versa, have been a focal point of biochemical and biophysical investigations for many decades. Complexes of lipids and proteins, such as the various lipoprotein factions, play central roles in the trafficking of important proteins, small molecules and metabolites and are often implicated in disease states. Recently an engineered lipoprotein particle, termed the nanodisc, a modified form of the human high density lipoprotein fraction, has served as a membrane mimetic for the investigation of membrane proteins and studies of lipid-protein interactions. In this review, we summarize the current knowledge regarding this self-assembling lipid-protein complex and provide examples for its utility in the investigation of a large number of biological systems.
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9
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Yue Z, Li C, Voth GA, Swanson JMJ. Dynamic Protonation Dramatically Affects the Membrane Permeability of Drug-like Molecules. J Am Chem Soc 2019; 141:13421-13433. [PMID: 31382734 DOI: 10.1021/jacs.9b04387] [Citation(s) in RCA: 44] [Impact Index Per Article: 8.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/06/2023]
Abstract
Permeability (Pm) across biological membranes is of fundamental importance and a key factor in drug absorption, distribution, and development. Although the majority of drugs will be charged at some point during oral delivery, our understanding of membrane permeation by charged species is limited. The canonical model assumes that only neutral molecules partition into and passively permeate across membranes, but there is mounting evidence that these processes are also facile for certain charged species. However, it is unknown whether such ionizable permeants dynamically neutralize at the membrane surface or permeate in their charged form. To probe protonation-coupled permeation in atomic detail, we herein apply continuous constant-pH molecular dynamics along with free energy sampling to study the permeation of a weak base propranolol (PPL), and evaluate the impact of including dynamic protonation on Pm. The simulations reveal that PPL dynamically neutralizes at the lipid-tail interface, which dramatically influences the permeation free energy landscape and explains why the conventional model overestimates the assigned intrinsic permeability. We demonstrate how fixed-charge-state simulations can account for this effect, and propose a revised model that better describes pH-coupled partitioning and permeation. Our results demonstrate how dynamic changes in protonation state may play a critical role in the permeation of ionizable molecules, including pharmaceuticals and drug-like molecules, thus requiring a revision of the standard picture.
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Affiliation(s)
- Zhi Yue
- Department of Chemistry, James Frank Institute, and Institute for Biophysical Dynamics , The University of Chicago , Chicago , Illinois 60637 , United States
| | - Chenghan Li
- Department of Chemistry, James Frank Institute, and Institute for Biophysical Dynamics , The University of Chicago , Chicago , Illinois 60637 , United States
| | - Gregory A Voth
- Department of Chemistry, James Frank Institute, and Institute for Biophysical Dynamics , The University of Chicago , Chicago , Illinois 60637 , United States
| | - Jessica M J Swanson
- Department of Chemistry, James Frank Institute, and Institute for Biophysical Dynamics , The University of Chicago , Chicago , Illinois 60637 , United States
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10
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Sharma A, Sarkar A, Goswami D, Bhattacharyya A, Enderlein J, Kumbhakar M. Determining Metal Ion Complexation Kinetics with Fluorescent Ligands by Using Fluorescence Correlation Spectroscopy. Chemphyschem 2019; 20:2093-2102. [PMID: 31240810 DOI: 10.1002/cphc.201900517] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/22/2019] [Revised: 06/25/2019] [Indexed: 11/08/2022]
Abstract
Fluorescence correlation spectroscopy (FCS) has been extensively used to measure equilibrium binding constants (K) or association and dissociation rates in many reversible chemical reactions across chemistry and biology. For the majority of investigated reactions, the binding constant was on the order of ∼100 M-1 , with dissociation constants faster or equal to 103 s-1 , which ensured that enough association/dissociation events occur during the typical diffusion-determined transition time of molecules through the FCS detection volume. However, complexation reactions involving metal ions and chelating ligands exhibit equilibrium constants exceeding 104 M-1 . In the present paper, we explore the applicability of FCS for measuring reaction rates of such complexation reactions, and apply it to binding of iron, europium and uranyl ions to a fluorescent chelating ligand, calcein. For this purpose, we exploit the fact that the ligand fluorescence becomes strongly quenched after binding a metal ion, which results in strong intensity fluctuations that lead to a partial correlation decay in FCS. We also present measurements for the strongly radioactive ions of 241 Am3+ , where the extreme sensitivity of FCS allows us to work with sample concentrations and volumes that exhibit close to negligible radioactivity levels. A general discussion of the applicability of FCS to the investigation of metal-ligand binding reactions concludes our paper.
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Affiliation(s)
- Arjun Sharma
- Radiation & Photochemistry Division, Bhabha Atomic Research Centre, Mumbai, 400085, India.,Chemical Sciences, Homi Bhabha National Institute, Mumbai 400094, India
| | - Aranyak Sarkar
- Radiation & Photochemistry Division, Bhabha Atomic Research Centre, Mumbai, 400085, India.,Chemical Sciences, Homi Bhabha National Institute, Mumbai 400094, India
| | - Dibakar Goswami
- Chemical Sciences, Homi Bhabha National Institute, Mumbai 400094, India.,Bio-organic Division, Bhabha Atomic Research Centre, Mumbai, 400085, India
| | - Arunasis Bhattacharyya
- Chemical Sciences, Homi Bhabha National Institute, Mumbai 400094, India.,Radiochemistry Division, Bhabha Atomic Research Centre, Mumbai, 400085, India
| | - Jörg Enderlein
- III. Institute of Physics - Biophysics, Georg August University, 37077, Göttingen, Germany
| | - Manoj Kumbhakar
- Radiation & Photochemistry Division, Bhabha Atomic Research Centre, Mumbai, 400085, India.,Chemical Sciences, Homi Bhabha National Institute, Mumbai 400094, India
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11
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Gupta A, Sankaran J, Wohland T. Fluorescence correlation spectroscopy: The technique and its applications in soft matter. PHYSICAL SCIENCES REVIEWS 2019. [DOI: 10.1515/psr-2017-0104] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/11/2022]
Abstract
Abstract
Fluorescence correlation spectroscopy (FCS) is a well-established single-molecule method used for the quantitative spatiotemporal analysis of dynamic processes in a wide range of samples. It possesses single-molecule sensitivity but provides ensemble averaged molecular parameters such as mobility, concentration, chemical reaction kinetics, photophysical properties and interaction properties. These parameters have been utilized to characterize a variety of soft matter systems. This review provides an overview of the basic principles of various FCS modalities, their instrumentation, data analysis, and the applications of FCS to soft matter systems.
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12
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Díaz-Vegas AR, Cordova A, Valladares D, Llanos P, Hidalgo C, Gherardi G, De Stefani D, Mammucari C, Rizzuto R, Contreras-Ferrat A, Jaimovich E. Mitochondrial Calcium Increase Induced by RyR1 and IP3R Channel Activation After Membrane Depolarization Regulates Skeletal Muscle Metabolism. Front Physiol 2018; 9:791. [PMID: 29988564 PMCID: PMC6026899 DOI: 10.3389/fphys.2018.00791] [Citation(s) in RCA: 43] [Impact Index Per Article: 7.2] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/25/2018] [Accepted: 06/06/2018] [Indexed: 11/13/2022] Open
Abstract
Aim: We hypothesize that both type-1 ryanodine receptor (RyR1) and IP3-receptor (IP3R) calcium channels are necessary for the mitochondrial Ca2+ increase caused by membrane depolarization induced by potassium (or by electrical stimulation) of single skeletal muscle fibers; this calcium increase would couple muscle fiber excitation to an increase in metabolic output from mitochondria (excitation-metabolism coupling). Methods: Mitochondria matrix and cytoplasmic Ca2+ levels were evaluated in fibers isolated from flexor digitorium brevis muscle using plasmids for the expression of a mitochondrial Ca2+ sensor (CEPIA3mt) or a cytoplasmic Ca2+ sensor (RCaMP). The role of intracellular Ca2+ channels was evaluated using both specific pharmacological inhibitors (xestospongin B for IP3R and Dantrolene for RyR1) and a genetic approach (shIP3R1-RFP). O2 consumption was detected using Seahorse Extracellular Flux Analyzer. Results: In isolated muscle fibers cell membrane depolarization increased both cytoplasmic and mitochondrial Ca2+ levels. Mitochondrial Ca2+ uptake required functional inositol IP3R and RyR1 channels. Inhibition of either channel decreased basal O2 consumption rate but only RyR1 inhibition decreased ATP-linked O2 consumption. Cell membrane depolarization-induced Ca2+ signals in sub-sarcolemmal mitochondria were accompanied by a reduction in mitochondrial membrane potential; Ca2+ signals propagated toward intermyofibrillar mitochondria, which displayed increased membrane potential. These results are compatible with slow, Ca2+-dependent propagation of mitochondrial membrane potential from the surface toward the center of the fiber. Conclusion: Ca2+-dependent changes in mitochondrial membrane potential have different kinetics in the surface vs. the center of the fiber; these differences are likely to play a critical role in the control of mitochondrial metabolism, both at rest and after membrane depolarization as part of an “excitation-metabolism” coupling process in skeletal muscle fibers.
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Affiliation(s)
- Alexis R Díaz-Vegas
- Muscle Physiology Laboratory, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile
| | - Alex Cordova
- Biomedical Neuroscience Institute, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile
| | - Denisse Valladares
- Muscle Physiology Laboratory, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile.,Exercise and Movement Science Laboratory, Universidad Finis Terrae, Santiago, Chile
| | - Paola Llanos
- Muscle Physiology Laboratory, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile.,Institute for Research in Dental Science, Universidad de Chile, Santiago, Chile
| | - Cecilia Hidalgo
- Biomedical Neuroscience Institute, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile
| | - Gaia Gherardi
- Department of Biomedical Sciences, University of Padova, Padova, Italy
| | - Diego De Stefani
- Department of Biomedical Sciences, University of Padova, Padova, Italy
| | | | - Rosario Rizzuto
- Department of Biomedical Sciences, University of Padova, Padova, Italy
| | - Ariel Contreras-Ferrat
- Muscle Physiology Laboratory, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile
| | - Enrique Jaimovich
- Muscle Physiology Laboratory, Center of Studies in Exercise, Metabolism and Cancer, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile
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13
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Rigler R, Widengren J. Fluorescence-based monitoring of electronic state and ion exchange kinetics with FCS and related techniques: from T-jump measurements to fluorescence fluctuations. EUROPEAN BIOPHYSICS JOURNAL : EBJ 2018; 47:479-492. [PMID: 29260269 PMCID: PMC5982452 DOI: 10.1007/s00249-017-1271-1] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 07/05/2017] [Revised: 10/30/2017] [Accepted: 12/03/2017] [Indexed: 11/01/2022]
Abstract
In this review, we give a historical view of how our research in the development and use of fluorescence correlation spectroscopy (FCS) and related techniques has its roots and how it originally evolved from the pioneering work of Manfred Eigen, his colleagues, and coworkers. Work on temperature-jump (T-jump) experiments, conducted almost 50 years ago, led on to the development of the FCS technique. The pioneering work in the 1970s, introducing and demonstrating the concept for FCS, in turn formed the basis for the breakthrough use of FCS more than 15 years later. FCS can be used for monitoring reaction kinetics, based on fluctuations at thermodynamic equilibrium, rather than on relaxation measurements following perturbations. In this review, we more specifically discuss FCS measurements on photodynamic, electronic state transitions in fluorophore molecules, and on proton exchange dynamics in solution and on biomembranes. In the latter case, FCS measurements have proven capable of casting new light on the mechanisms of proton exchange at biological membranes, of central importance to bioenergetics and signal transduction. Finally, we describe the transient-state (TRAST) spectroscopy/imaging technique, sharing features with both relaxation (T-jump) and equilibrium fluctuation (FCS) techniques. TRAST is broadly applicable for cellular and molecular studies, and we briefly outline how TRAST can provide unique information from fluorophore blinking kinetics, reflecting e.g., cellular metabolism, rare molecular encounters, and molecular stoichiometries.
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Affiliation(s)
- Rudolf Rigler
- Department of Medical Biochemistry and Biophysics, Karolinska Institute, Stockholm, Sweden
| | - Jerker Widengren
- Experimental Biomolecular Physics/ Department of Applied Physics, Royal Institute of Technology (KTH), Stockholm, Sweden.
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14
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Liu F, Dong C, Ren J. A study of the diffusion dynamics and concentration distribution of gold nanospheres (GNSs) without fluorescent labeling inside live cells using fluorescence single particle spectroscopy. NANOSCALE 2018; 10:5309-5317. [PMID: 29503992 DOI: 10.1039/c7nr08722e] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/08/2023]
Abstract
Colloidal gold nanospheres (GNSs) have become important nanomaterials in biomedical applications due to their special optical properties, good chemical stability, and biocompatibility. However, measuring the diffusion coefficients or concentration distribution of GNSs within live cells accurately without any extra fluorescent labeling in situ has still not been resolved. In this work, a single particle method is developed to study the concentration distribution of folic acid-modified GNSs (FA-GNSs) internalized via folate receptors, and investigates their diffusion dynamics within live cells using single particle fluorescence correlation spectroscopy (FCS). We optimized the experimental conditions and verified the feasibility of 30 nm GNSs without extra fluorescence labeling being used for single particle detection inside live cells. Then, the FCS characterization strategy was used to measure the concentration and diffusion coefficient distributions of GNSs inside live cells and the obtained results were basically in agreement with those obtained by TEM. The results demonstrate that our strategy is characterized as an in situ, nondestructive, rapid and dynamic method for the assay of live cells, and it may be widely used in the further design of GNP-based drug delivery and therapeutics.
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Affiliation(s)
- Fangchao Liu
- School of Chemistry & Chemical Engineering, State Key Laboratory of Metal Matrix Composites, Shanghai Jiao Tong University, 800 Dongchuan Road, Shanghai, 200240, P. R. China.
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15
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Gennis RB. Proton Dynamics at the Membrane Surface. Biophys J 2017; 110:1909-11. [PMID: 27166799 DOI: 10.1016/j.bpj.2016.04.001] [Citation(s) in RCA: 16] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/29/2016] [Accepted: 04/01/2016] [Indexed: 11/16/2022] Open
Affiliation(s)
- Robert B Gennis
- Department of Biochemistry, University of Illinois, Urbana, Illinois.
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16
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Sato T, Tsukamoto M, Yamamoto S, Mitsuishi M, Miyashita T, Nagano S, Matsui J. Acid-Group-Content-Dependent Proton Conductivity Mechanisms at the Interlayer of Poly(N-dodecylacrylamide-co-acrylic acid) Copolymer Multilayer Nanosheet Films. LANGMUIR : THE ACS JOURNAL OF SURFACES AND COLLOIDS 2017; 33:12897-12902. [PMID: 29058441 DOI: 10.1021/acs.langmuir.7b03160] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 06/07/2023]
Abstract
The effect of the content of acid groups on the proton conductivity at the interlayer of polymer-nanosheet assemblies was investigated. For that purpose, amphiphilic poly(N-dodecylacrylamide-co-acrylic acid) copolymers [p(DDA/AA)] with varying contents of AA were synthesized by free radical polymerization. Surface pressure (π)-area (A) isotherms of these copolymers indicated that stable polymer monolayers are formed at the air/water interface for AA mole fraction (n) ≤ 0.49. In all cases, a uniform dispersion of the AA groups in the polymer monolayer was observed. Subsequently, polymer monolayers were transferred onto solid substrates using the Langmuir-Blodgett (LB) technique. X-ray diffraction (XRD) analyses of the multilayer films showed strong Bragg diffraction peaks, suggesting a highly uniform lamellar structure for the multilayer films. The proton conductivity of the multilayer films parallel to the direction of the layer planes were measured by impedance spectroscopy, which revealed that the conductivity increased with increasing values of n. Activation energies for proton conduction of ∼0.3 and 0.42 eV were observed for n ≥ 0.32 and n = 0.07, respectively. Interestingly, the proton conductivity of a multilayer film with n = 0.19 did not follow the Arrhenius equation. These results were interpreted in terms of the average distance between the AA groups (lAA), and it was concluded that, for n ≥ 0.32, an advanced 2D hydrogen bonding network was formed, while for n = 0.07, lAA is too long to form such hydrogen bonding networks. The lAA for n = 0.19 is intermediate to these extremes, resulting in the formation of hydrogen bonding networks at low temperatures, and disruption of these networks at high temperatures due to thermally induced motion. These results indicate that a high proton conductivity with low activation energy can be achieved, even under weakly acidic conditions, by arranging the acid groups at an optimal distance.
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Affiliation(s)
- Takuma Sato
- Graduate School of Science and Engineering, Yamagata University , 1-4-12 Kojirakawa-machi, Yamagata 990-8560, Japan
| | - Mayu Tsukamoto
- Graduate School of Science and Engineering, Yamagata University , 1-4-12 Kojirakawa-machi, Yamagata 990-8560, Japan
| | - Shunsuke Yamamoto
- Institute for Multidisciplinary Research for Advanced Materials, Tohoku University , 2-1-1 Katahira, Aoba-ku, Sendai 980-8577, Japan
| | - Masaya Mitsuishi
- Institute for Multidisciplinary Research for Advanced Materials, Tohoku University , 2-1-1 Katahira, Aoba-ku, Sendai 980-8577, Japan
| | - Tokuji Miyashita
- Institute for Multidisciplinary Research for Advanced Materials, Tohoku University , 2-1-1 Katahira, Aoba-ku, Sendai 980-8577, Japan
| | - Shusaku Nagano
- Nagoya University Venture Business Laboratory, Nagoya University , Furo-cho, Chikusa, Nagoya 464-8603, Japan
| | - Jun Matsui
- Faculty of Science, Yamagata University , 1-4-12 Kojirakawa-machi, Yamagata 990-8560, Japan
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17
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Weichselbaum E, Österbauer M, Knyazev DG, Batishchev OV, Akimov SA, Hai Nguyen T, Zhang C, Knör G, Agmon N, Carloni P, Pohl P. Origin of proton affinity to membrane/water interfaces. Sci Rep 2017; 7:4553. [PMID: 28674402 PMCID: PMC5495794 DOI: 10.1038/s41598-017-04675-9] [Citation(s) in RCA: 38] [Impact Index Per Article: 5.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/04/2016] [Accepted: 05/18/2017] [Indexed: 11/21/2022] Open
Abstract
Proton diffusion along biological membranes is vitally important for cellular energetics. Here we extended previous time-resolved fluorescence measurements to study the time and temperature dependence of surface proton transport. We determined the Gibbs activation energy barrier ΔG ‡r that opposes proton surface-to-bulk release from Arrhenius plots of (i) protons' surface diffusion constant and (ii) the rate coefficient for proton surface-to-bulk release. The large size of ΔG ‡r disproves that quasi-equilibrium exists in our experiments between protons in the near-membrane layers and in the aqueous bulk. Instead, non-equilibrium kinetics describes the proton travel between the site of its photo-release and its arrival at a distant membrane patch at different temperatures. ΔG ‡r contains only a minor enthalpic contribution that roughly corresponds to the breakage of a single hydrogen bond. Thus, our experiments reveal an entropic trap that ensures channeling of highly mobile protons along the membrane interface in the absence of potent acceptors.
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Affiliation(s)
- Ewald Weichselbaum
- 0000 0001 1941 5140grid.9970.7Institute of Biophysics, Johannes Kepler University Linz, 4040 Linz, Austria
| | - Maria Österbauer
- 0000 0001 1941 5140grid.9970.7Institute of Inorganic Chemistry, Johannes Kepler University Linz, 4040 Linz, Austria
| | - Denis G. Knyazev
- 0000 0001 1941 5140grid.9970.7Institute of Biophysics, Johannes Kepler University Linz, 4040 Linz, Austria
| | - Oleg V. Batishchev
- 0000 0001 2192 9124grid.4886.2A.N. Frumkin Institute of Physical Chemistry and Electrochemistry, Russian Academy of Sciences, Leninskiy pr. 31/4, Moscow, 119071 Russian Federation ,0000000092721542grid.18763.3bMoscow Institute of Physics and Technology, Institutsky lane, 9, 141700 Dolgoprudniy, Russian Federation
| | - Sergey A. Akimov
- 0000 0001 2192 9124grid.4886.2A.N. Frumkin Institute of Physical Chemistry and Electrochemistry, Russian Academy of Sciences, Leninskiy pr. 31/4, Moscow, 119071 Russian Federation ,0000 0001 0010 3972grid.35043.31National University of Science and Technology “MISiS”, Leninskiy pr. 4, Moscow, 119991 Russian Federation
| | - Trung Hai Nguyen
- 0000 0001 0728 696Xgrid.1957.aComputational Biomedicine (IAS-5 / INM-9) Forschungszentrum Jülich, 52425 Jülich, Germany, RWTH Aachen University, 52056 Aachen, Germany
| | - Chao Zhang
- 0000 0001 0728 696Xgrid.1957.aComputational Biomedicine (IAS-5 / INM-9) Forschungszentrum Jülich, 52425 Jülich, Germany, RWTH Aachen University, 52056 Aachen, Germany ,0000000121885934grid.5335.0Department of Chemistry, University of Cambridge, Lensfield Rd, Cambridge, CB2 1EW United Kingdom
| | - Günther Knör
- 0000 0001 1941 5140grid.9970.7Institute of Inorganic Chemistry, Johannes Kepler University Linz, 4040 Linz, Austria
| | - Noam Agmon
- 0000 0004 1937 0538grid.9619.7Institute of Chemistry, The Hebrew University of Jerusalem, Jerusalem, 9190401 Israel
| | - Paolo Carloni
- 0000 0001 0728 696Xgrid.1957.aComputational Biomedicine (IAS-5 / INM-9) Forschungszentrum Jülich, 52425 Jülich, Germany, RWTH Aachen University, 52056 Aachen, Germany
| | - Peter Pohl
- Institute of Biophysics, Johannes Kepler University Linz, 4040, Linz, Austria.
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18
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Ter Beek J, Kahle M, Ädelroth P. Modulation of protein function in membrane mimetics: Characterization of P. denitrificans cNOR in nanodiscs or liposomes. BIOCHIMICA ET BIOPHYSICA ACTA-BIOMEMBRANES 2017; 1859:1951-1961. [PMID: 28668220 DOI: 10.1016/j.bbamem.2017.06.017] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Subscribe] [Scholar Register] [Received: 02/21/2017] [Revised: 06/03/2017] [Accepted: 06/27/2017] [Indexed: 10/19/2022]
Abstract
For detailed functional characterization, membrane proteins are usually studied in detergent. However, it is becoming clear that detergent micelles are often poor mimics of the lipid environment in which these proteins function. In this work we compared the catalytic properties of the membrane-embedded cytochrome c-dependent nitric oxide reductase (cNOR) from Paracoccus (P.) denitrificans in detergent, lipid/protein nanodiscs, and proteoliposomes. We used two different lipid mixtures, an extract of soybean lipids and a defined mix of synthetic lipids mimicking the original P. denitrificans membrane. We show that the catalytic activity of detergent-solubilized cNOR increased threefold upon reconstitution from detergent into proteoliposomes with the P. denitrificans lipid mixture, and above two-fold when soybean lipids were used. In contrast, there was only a small activity increase in nanodiscs. We further show that binding of the gaseous ligands CO and O2 are affected differently by reconstitution. In proteoliposomes the turnover rates are affected much more than in nanodiscs, but CO-binding is more significantly accelerated in liposomes with soybean lipids, while O2-binding is faster with the P. denitrificans lipid mix. We also investigated proton-coupled electron transfer during the reaction between fully reduced cNOR and O2, and found that the pKa of the internal proton donor was increased in proteoliposomes but not in nanodiscs. Taking our results together, the liposome-reconstituted enzyme shows significant differences to detergent-solubilized protein. Nanodiscs show much more subtle effects, presumably because of their much lower lipid to protein ratio. Which of these two membrane-mimetic systems best mimics the native membrane is discussed.
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Affiliation(s)
- Josy Ter Beek
- Department of Biochemistry and Biophysics, Stockholm University, Svante Arrhenius väg 16C, SE-106 91 Stockholm, Sweden.
| | - Maximilian Kahle
- Department of Biochemistry and Biophysics, Stockholm University, Svante Arrhenius väg 16C, SE-106 91 Stockholm, Sweden.
| | - Pia Ädelroth
- Department of Biochemistry and Biophysics, Stockholm University, Svante Arrhenius väg 16C, SE-106 91 Stockholm, Sweden.
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19
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Sjöholm J, Bergstrand J, Nilsson T, Šachl R, Ballmoos CV, Widengren J, Brzezinski P. The lateral distance between a proton pump and ATP synthase determines the ATP-synthesis rate. Sci Rep 2017; 7:2926. [PMID: 28592883 PMCID: PMC5462737 DOI: 10.1038/s41598-017-02836-4] [Citation(s) in RCA: 33] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/17/2017] [Accepted: 04/19/2017] [Indexed: 11/09/2022] Open
Abstract
We have investigated the effect of lipid composition on interactions between cytochrome bo 3 and ATP-synthase, and the ATP-synthesis activity driven by proton pumping. The two proteins were labeled by fluorescent probes and co-reconstituted in large (d ≅ 100 nm) or giant (d ≅ 10 µm) unilamellar lipid vesicles. Interactions were investigated using fluorescence correlation/cross-correlation spectroscopy and the activity was determined by measuring ATP production, driven by electron-proton transfer, as a function of time. We found that conditions that promoted direct interactions between the two proteins in the membrane (higher fraction DOPC lipids or labeling by hydrophobic molecules) correlated with an increased activity. These data indicate that the ATP-synthesis rate increases with decreasing distance between cytochrome bo 3 and the ATP-synthase, and involves proton transfer along the membrane surface. The maximum distance for lateral proton transfer along the surface was found to be ~80 nm.
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Affiliation(s)
- Johannes Sjöholm
- Department of Biochemistry and Biophysics, The Arrhenius Laboratories for Natural Sciences, Stockholm University, SE-106 91, Stockholm, Sweden
| | - Jan Bergstrand
- Experimental Biomolecular Physics, Department of Applied Physics, Royal Institute of Technology (KTH), SE-106 91, Stockholm, Sweden
| | - Tobias Nilsson
- Department of Biochemistry and Biophysics, The Arrhenius Laboratories for Natural Sciences, Stockholm University, SE-106 91, Stockholm, Sweden
| | - Radek Šachl
- Department of Biophysical Chemistry, J. Heyrovský Institute of Physical Chemistry of the A.S.C.R. v.v.i., Prague, Czech Republic
| | | | - Jerker Widengren
- Experimental Biomolecular Physics, Department of Applied Physics, Royal Institute of Technology (KTH), SE-106 91, Stockholm, Sweden
| | - Peter Brzezinski
- Department of Biochemistry and Biophysics, The Arrhenius Laboratories for Natural Sciences, Stockholm University, SE-106 91, Stockholm, Sweden.
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20
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van der Loop TH, Ottosson N, Vad T, Sager WFC, Bakker HJ, Woutersen S. Communication: Slow proton-charge diffusion in nanoconfined water. J Chem Phys 2017; 146:131101. [DOI: 10.1063/1.4979714] [Citation(s) in RCA: 18] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/14/2022] Open
Affiliation(s)
- Tibert H. van der Loop
- Van ’t Hoff Institute for Molecular Sciences (HIMS), University of Amsterdam, Science Park 904, 1098 XH Amsterdam, The Netherlands
| | - Niklas Ottosson
- FOM Institute for Atomic and Molecular Physics, Science Park 104, 1098 XG Amsterdam, The Netherlands
| | - Thomas Vad
- Institut für Textiltechnik, RWTH Aachen University, Otto-Blumenthal-Strasse 1, 52074 Aachen, Germany
| | - Wiebke F. C. Sager
- Peter Grünberg Institute, Forschungszentrum Jülich GmbH, 52425 Jülich, Germany
| | - Huib J. Bakker
- FOM Institute for Atomic and Molecular Physics, Science Park 104, 1098 XG Amsterdam, The Netherlands
| | - Sander Woutersen
- Van ’t Hoff Institute for Molecular Sciences (HIMS), University of Amsterdam, Science Park 904, 1098 XH Amsterdam, The Netherlands
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21
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Abstract
Membrane proteins play a most important part in metabolism, signaling, cell motility, transport, development, and many other biochemical and biophysical processes which constitute fundamentals of life on the molecular level. Detailed understanding of these processes is necessary for the progress of life sciences and biomedical applications. Nanodiscs provide a new and powerful tool for a broad spectrum of biochemical and biophysical studies of membrane proteins and are commonly acknowledged as an optimal membrane mimetic system that provides control over size, composition, and specific functional modifications on the nanometer scale. In this review we attempted to combine a comprehensive list of various applications of nanodisc technology with systematic analysis of the most attractive features of this system and advantages provided by nanodiscs for structural and mechanistic studies of membrane proteins.
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Affiliation(s)
- Ilia G Denisov
- Department of Biochemistry and Department of Chemistry, University of Illinois , Urbana, Illinois 61801, United States
| | - Stephen G Sligar
- Department of Biochemistry and Department of Chemistry, University of Illinois , Urbana, Illinois 61801, United States
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22
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Lancaster CRD, Betz YM, Heit S, Lafontaine MA. Transmembrane Electron and Proton Transfer in Diheme-Containing Succinate : Quinone Oxidoreductases. Isr J Chem 2017. [DOI: 10.1002/ijch.201600139] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/12/2022]
Affiliation(s)
- C. Roy D. Lancaster
- Department of Structural Biology; Center of Human and Molecular Biology (ZHMB); Saarland University; Faculty of Medicine Building 60 D-66421 Homburg (Saar) Germany
| | - Yamila M. Betz
- Department of Structural Biology; Center of Human and Molecular Biology (ZHMB); Saarland University; Faculty of Medicine Building 60 D-66421 Homburg (Saar) Germany
| | - Sabine Heit
- Department of Structural Biology; Center of Human and Molecular Biology (ZHMB); Saarland University; Faculty of Medicine Building 60 D-66421 Homburg (Saar) Germany
| | - Michael A. Lafontaine
- Department of Structural Biology; Center of Human and Molecular Biology (ZHMB); Saarland University; Faculty of Medicine Building 60 D-66421 Homburg (Saar) Germany
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