Book of Abstracts - fret.uni-duesseldorf.de · Book of Abstracts 0,01 1 100 0 5 10 15 20 25 t R2 t diff sCCF amplitude correlation time t c [ms] 0.0 0.2 0.8 10 t R1 MgCl 2 [mM] Welcome

International Discussion Meeting

Förster Resonance Energy Transferin life sciences: FRET 2

Molecular imaging – structure – dynamics ‐ hybrid methods

April 3 – 6, 2016Max Planck Institute for Biophysical Chemistry

Göttingen, Germany

Under the auspices of:

Max Planck Institute for Biophysical Chemistry

CMST COST Action CM1306 Study Group Biophysical Chemistry

Book of Abstracts

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Welcome to Göttingen! On the occasion of the 106th anniversary of his birth and 70 years after his first publication on resonance energy transfer we are again honoring the achievements of Theodor Förster by a discussion meeting about FRET, a key topic of his rich scientific legacy which began in Göttingen. During the years 1946 to 1948, Theodor Förster published papers in Naturwissenschaften and Annalen der Physik outlining the quantum-mechanical behaviour of the transfer of electronic excitation energy between two molecules in a solution. Förster's groundbreaking work in spectroscopy was built upon the earlier theories of J. and F. Perrin, and explained the non-radiative transfer of energy between two molecules. FRET, is an acronym for Förster resonance energy transfer or (less precisely) fluorescence resonance energy transfer. Equations determined by Förster were the basis for a quantitative interpretation of FRET and feature parameters that can be derived experimentally. FRET is a common and fundamental photophysical technique in the life and materials sciences. After absorption of light, intrachromophore processes, such as radiative decay (e.g., fluorescence, phosphorescence) and radiationless transitions (e.g., internal conversion, intersystem crossing), dissipate the absorbed energy. FRET is an interchromophoric relaxation process that transfers the electronic excitation nonradiatively from an initially excited donor to a ground-state acceptor. While resonance energy transfer was first observed in fluorescence polarization studies in the 1920s, interest in FRET was limited to interpreting the concentration dependence of fluorescence depolarization. The intention of Förster's papers was to describe in a quantitative fashion the energy migration in molecular crystals and during photosynthesis in plants. Innumerable manifestations and uses of FRET have since been described over the ensuing years. Some notable examples include light harvesting in photosynthesis, design of high performance sensors, structural determination in macromolecules, and detection of biomolecular interactions in the life sciences. The number of citations using FRET measurements has increased almost 300-fold over the past 20 years. We call this a “Discussion” Meeting because we will not have long plenary lectures but rather lively discussions after short presentations and around posters about the latest experimental results, ideas, or theories related to FRET. We warmly welcome you to our discussion meeting! The organizing committee: Donna Arndt-Jovin, Stefan Hell, Thomas Jovin, Niko Hildebrandt, Igor Medintz, Claus Seidel and Jürgen Troe

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We thank the scientific committee for helping setting up the program:

– Philippe Bastiaens - MPI of Molecular Physiology, Dortmund, DE – Helmut Grubmüller - MPIBPC, Göttingen, DE – Thorsten Hugel - Albert-Ludwigs-University, Freiburg, DE – Dagmar Klostermeier - Westfälische Wilhelms-University, Münster, DE – Marcia Levitus - Arizona State University, Tempe, AZ, USA – Diane Lidke - University of New Mexico, Albuquerque, NM, USA – Fraser MacMillan - University of East Anglia, Norwich, UK – Arwen R. Pearson, University Hamburg, Hamburg, DE – Steve Vogel - NIH, Bethesda, MD, USA

This conference is organized under the auspices of:

Max-Planck-Institut für Biophysikalische Chemie Göttingen, Germany

German Society for Biochemistry and Molecular Biology (Study Group Biophysical Chemistry)

CMST COST Action CM1306 Understanding Movement and Mechanism in Molecular Machines

DFG funded Collaborative Research Center SFB 755 “Nanoscale Photonic Imaging” Georg-August-Universität Göttingen DFG funded Collaborative Research Center SFB 803 “Functionality controlled by organization in and between membranes” Georg-August-Universität Göttingen


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Theodor Förster: A giant of modern photochemistry May 15th, 1910 - May 20th 1974

1933 Ph.D. (with Erwin Madelung) University of Frankfurt am Main 1942 o. Professor, the State University of Poznan 1946 Arrival in Niedernjesa, Kr. Göttingen (old school) 27.9.1946 First Publication on FRET (now 70 years)

Energy transport and fluorescence [in German] Naturwissenschaften 33:166-175.

1.12.1947-30.4.1951

Max-Planck-Institute for physical chemistry, Göttingen, leader of the department for "structure research".

1948 Most cited work on FRET (5777 citations until 9.3.2016) Zwischenmolekulare Energiewanderung und Fluoreszenz [in German] Annalen der Physik 437(2): 55-75.

1.5.1951 Full Professor, Technical University Stuttgart 17.7.1952 External scientific member of the Max Planck Institute for Physical Chemistry Three main research fields (77 Publications from 1933 to 1975)

1. Förster Resonance Energy Transfer (FRET), 2. the Förster cycle, linking protolysis and reprotonation of molecules in the excited

state, 3. excimer formation by association of an excited with an electronic-ground-state

molecule. Literature (see our webpage: http://fret.uni-duesseldorf.de/cms/Literature.html)

1. Albert Weller, Nachruf auf Theodor Förster, Berichte der Bunsengesellschaft für Physikalische Chemie 1974, 78(10): 969-971 [German].

2. Albert Weller, In Memoriam Theodor Förster, EPA Newsletter 1980, 9(April): 6-19 [English].

3. Special Issue: Förster Resonance Energy Transfer, ChemPhysChem 2011, 12(3) 421-719.

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The Max Planck Society thanks the following organizations for support for research in conjunction with this forum:

Acal BFi Germany GmbH Oppelner Straße 5 82194 Gröbenzell, Germany

AHF analysentechnik AG Kohlplattenweg 18 72074 Tübingen, Germany

ATTO-TEC GmbH Am Eichenhang 50 57076 Siegen, Germany

chemosensors MDPI AG Klybeckstrasse 64, 4057 Basel, Switzerland

Fond der Chemischen Industrie Verband der Chemischen Industrie e.V. (VCI) Mainzer Landstraße 55 60329 Frankfurt am Main, Germany http://fonds.vci.de/

Gesellschaft für Biochemie und Molekularbiologie GBM-Geschäftsstelle Mörfelder Landstr. 125 60598 Frankfurt am Main, Germany http://www.gbm-online.de

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Hidex vertreten durch FCI - Laborgeräte & Consulting Zedernweg 28 D-55128 Mainz, Germany

HORIBA Jobin Yvon GmbH Neuhofstr. 9 64625 Bensheim, Germany

IBA GmbH IBA Headquarters Rudolf-Wissell-Str. 28 37079 Göttingen, Germany

Methods and Applications in Fluorescence IOP Publishing Temple Circus, Temple Way Bristol BS1 6HG, UK

NKT Photonics GmbH Schanzenstraße 39, Bldg. D9-D13 51063 Köln, Germany

Max-Planck-Institut für Biophysikalische Chemie Am Faßberg 11 37077 Göttingen, Germany http://www.mpibpc.mpg.de/en

Office of Naval Research One Liberty Center 875 N. Randolph Street, Suite 1425 Arlington, VA 22203-1995, USA


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Picoquant GmbH Rudower Chaussee 29 (IGZ) 12489 Berlin, Germany

Princeton Instruments vertreten durch Roper Scientific GmbH Einsteinstraße 39a 82152 Planegg/Martinsried, Germany

PURIMEX Dr. Gerd Kotzorek Auf dem Wildhagen 8 34393 Grebenstein, Germany

RoentDek Handels GmbH Im Vogelshaag 8 65779 Kelkheim, Germany

DFG funded Collaborative Research Center SFB 755 “Nanoscale Photonic Imaging” Georg-August-Universität Göttingen https://www.uni-goettingen.de/de/318955.html

DFG funded Collaborative Research Center SFB 803 “Functionality controlled by organization in and between membranes” Georg-August-Universität Göttingen http://www.uni-goettingen.de/en/213080.html

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Program of the International Discussion Meeting “Förster Resonance Energy transfer in Life Sciences 2" (FRET 2)

Göttingen, April 3-6, 2016

The organizers thank the following scientific organizations for support of organization and speakers: Max Planck Institute for Biophysical Chemistry; GBM Study Group Biophysical Chemistry; Fonds der Chemischen Industrie; Office of Naval Research Grant; CMST Cost Action CM1306 [1]; SFB 755 [2]; SFB 803 [3].

Sun, April 3: Opening session

18:00-19:00 Registration, poster setup

19:00-19:10 Jürgen Troe Welcome

19:10-19:40 Niko Hildebrandt History of FRET

19:40-20:10 Ulrich Steiner The scientific work of Theodor Förster: A personal view

20:15-22:00 Reception

Mon, April 4: Day 1

Session 1: Molecular machinery [1] Chair: Fraser MacMillan

9:00-9:30 Dagmar Klostermeier T1: Unwind and relax: Using single-molecule FRET to dissect the mechanism of ATP-driven molecular machines

9:30-9:50 Mateusz Dyla T2: Single-molecule dynamics of a SERCA homologue

9:50-10:10 Sarah Rauscher T3: Structural ensembles of intrinsically disordered proteins from simulation and experiment

10:10-10:30 Katherina Hemmen T4: Multiparameter fluorescence spectroscopic toolkit resolves the heterogeneity of unfolded states

10:30-11:00 Coffee

Session 2: Single molecule theory and application Chair: William Eaton

11:00-11:30 Irina Gopich T5: Maximum likelihood analysis in single-molecule FRET

11.30-11:50 Bettina Keller T6: Accounting for experimental errors in HMM analyses of single-molecule FRET experiments

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11:50‐12:20 Hoi Sung Chung T7: Multi‐color single molecule FRET study of intrinsically disordered protein binding

12:20‐12:50 Ben Schuler T8: Protein folding dynamics from single‐molecule FRET – from in vitro to in vivo

13:00‐14:00 Lunch

14:00‐17:00 Poster session & coffee/tea

Session 3: Imaging, multimode applications, acceptors Chair: Steven Vogel

17:00‐17:20 Alexy Chizhik T9: Metal‐induced energy transfer for live cell nanoscopy

17:20‐17:50 Philippe Bastiaens T10: Quantitative FRET imaging reveals the interdependence of vesicular membrane dynamics and growth factor signal processing

17:50‐18:10 Enrico Gratton T11: 3D fluorescence anisotropy imaging using selective plane illumination microscopy

18:10‐18:30 Marisa Martin‐Fernandez

T12: Combining FRET with single particle localisation microscopy to ascertain in cellulo EGFR oligomer structure

18:30‐18:50 Alessandro Esposito T13: Frontiers in biochemical imaging: multiplexed FRET sensors and Optogenetics

18:50‐19:10 Kees Jalink T14: FRET sensing of metabolites: dedicated FLIM sensors, dedicated FLIM readout

19:10‐19:30 Theodorus Gadella, Jr.

T15: mScarlet, a novel high quantum yield monomeric red fluorescent protein with strongly enhanced properties for sensitized emission‐based FRET

19:30‐19:45 Meeting of the GBM Study Group Biophysical Chemistry

20:00‐22:00 Conference Dinner

Tues, April 5: Day 2

Session 4: Structural analysis Chair: Helmut Grubmüller

9:00‐9:20 Marcia Levitus [2] T16: Cyanine dyes in biophysical research: The photophysics of Cy‐dyes on DNA

9:20‐9:50 Victoria Birkedal T17: Single‐molecule FRET studies of DNA structures and devices

9:50‐10:10 Wolf Holtkamp T18: Co‐translational protein folding on the ribosome monitored in real time

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10:10-10:30 Diane Lidke T19: Enhanced dimerization of oncogenic EGFR mutants revealed by single molecule imaging and FRET microscopy

10:30-11:00 Coffee

Session 5: New instrumentation and analysis Chair: Philippe Bastiaens

11:00-11:20 Johannes Hohlbein T20: Camera-based single-molecule FRET detection with improved time resolution

11.20-11:40 Wieb van der Meer T21: Estimating the distribution of FRETefficiencies from fluorescence Lifetime-FRET data

11:40-12:00 Olaf Schulz T22: Recent advances in lifetime FRET

12:00-12:20 Martin Masip T23: Reversible cryo-arrest for imaging molecules in living cells at high spatial resolution

12:20-12:40 Thomas Jovin T24: Extended excitation FLIM (eeFLIM)

13:00-14:00 Lunch

14:00-16:00 Poster session & coffee/tea

Session 6: Hybrid approaches: FRET combined with modelling and other spectroscopies Chair: Attila Szabo

16:00-16:30 Gerhard Hummer T25: Assembling the pieces of protein puzzles

16:30-16:50 Timothy Craggs

T26: Combining coarse-grained and all-atom simulations with smFRET to obtain high-precision DNA-protein structures and conformational dynamics

16:50-17:10 Jens Michaelis T27: Fast NPS for quantitative structural information from single molecule FRET

17:10-17:30 Björn Hellenkamp T28: Distance distribution analysis reveals the dynamic structure of Hsp90

17:30-17:50 Thomas Peulen T29: Mapping motions to a state necessary for oligomerization of a large GTPase: a joint SAXS, NSE, EPR and FRET study

18:00-19:00 Dinner

19:00 - 20:00

Concert : Hyperion Trio (Hagen Schwarzrock, Klavier, Oliver Kipp, Violine und Katharina Troe, Violoncello)

– Franz Schubert, Klaviertrio B-Dur op.99 D898– Rubin Goldmark, Klaviertrio d-Moll op.1

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Parallel evening discussion groups

20.00 Standards and measurement protocols

Discussion leaders: Thorsten Hugel, Claus Seidel

Narain Karedla A‐P17: Single-Molecule Metal Induced EnergyTransfer (smMIET): Resolving nanometerdistances at single molecule level

Peter Nagy

B‐P46: Understanding and reducing the error in the evaluation of intensity‐based microscopic FRET experiments in the presence of low signal‐to‐noise ratio

– Report on a recent FRET comparison study (18 groups participated)– Report on the activities of the wwPDB Hybrid/Integrative Methods

Task Force– Establishing a taskforce for quantitative FRET measurements

(Structural modelling, kinetic analyses and imaging):Defining standards for measurements, analyses and interpretationwith protocols for: (i) FRET restrained structural modelling anddepositing of the obtained in data banks; (ii) Analysis of kineticnetworks.

20.00 Probe and Biosensor Development

Discussion leaders: Dorus Gadella Jr., Alexander Savitsky

Jasper van der Velde A‐P35: Enhancing single‐molecule FRET studies with photostabilizer‐dye conjugates

Asko Uri B‐P48: Organic photoluminescent probes possessing triplet –singlet energy transfer by Förster mechanism

Zongwen Jin E‐P80: Multiplexed micro‐RNA assays using time‐resolved FRET with biospectral correction

20.00 Nanoparticles

Discussion leaders: Niko Hildebrandt, Igor Medintz

Kateryna Trofymchuk E‐P85: Exploiting fast exciton diffusion in dye‐doped polymer nanoparticles to engineer efficient photoswitching through FRET

Igor Medintz E‐P81: Energy transfer‐based sensitization of luminescent gold nanoclusters

Jurriaan Zwier E‐P88: Time gated FRET microscopy‐ and plate based assays to study G‐Protein Coupled Receptors using bright Eu3+ and Tb3+ donors.

Lauren Field E‐P74: Semiconductor quantum dots as Förster Resonance Energy Transfer donors for intracellularly‐based biosensors

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Wed, April 6: Day 3

Session 7: N-way FRET Chair: Alexander Demchenko

8:30-9:00 Adam Hoppe [3] T30: 3D N-way FRET Microscopy Seeing more molecular interactions throughout the living cell

9:00-9:20 Sebastián Díaz T31: Homogenous FRET in molecular photonic wires

9:20-9:40 Michael Schlierf T32: farFRET: Extending the range in single-molecule FRET experiments beyond 10 nm

9:40-10:10 Don Lamb T33: Testing biomolecular coordination: quantitative analysis of single-triad FRET data

10:10-10:30 Tuan Nguyen T34: Deciphering CaMKII multimerization using holoenzyme assembly mutants, FCS, and concurrent homo- and hetero-FRET analysis

10:30-10:50 Coffee

Session 8: New imaging strategies Chair: Donna Arndt-Jovin

10:50-11:20 Gerard Marriott T35: On new classes of genetically-encoded fluorescent proteins optimized for fluorescence polarisation and FRET

11:20-11:40 Marcel Bruchez T36: Genetically targeted and activated fluorogenic FRET-based indicators

11:40-12:00 Alexander Savitsky T37: Peptide molecular dynamic beacon for FRET-sensor

12:00-12:30 Edward Lemke T38: Decoding molecular plasticity in the dark proteome

12:30-13:00 Russ Algar T39: Concentric FRET with quantum dots: energy transfer pathways and application to multiplexed biological sensing

13:00-13:15 Claus Seidel Closing Remarks

End of the meeting

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List of Posters

Topic A: Single‐molecule and ensemble in vitro FRET studies of Proteins, Nucleic Acids and Membranes; FRET spectroscopy.

A‐P1 Aznauryan, M. Probing the folding dynamics of human telomeric G‐quadruplex with single‐molecule FRET

A‐P2 Boening, D. 3D Light microscopy of protein structure with Angstrom resolution

A‐P3 Castellanos, M. Conformational mechanisms of homing‐to‐target in protein‐DNA binding and characterization of binding partners with different affinities

A‐P4 Cerminara, M. Unravelling the domain contributions to the folding mechanism of a multi‐domain protein

A‐P5 de Boer, M. Asymmetric conformational states of the ribosome recycling factor ABCE1 probed by single‐molecule FRET

A‐P6 Doroshenko, O. Accurate determination of the RNA junction via single molecule high‐precision FRET measurements

A‐P7 Fiorini, E. Point mutations reveal specific intra‐domain interactions essential for group II intron ribozyme folding

A‐P8 Gracia, P. Folding Thermodynamics and Kinetics of the Outer Membrane Phospholipase A investigated by Single‐Molecule FRET Spectroscopy

A‐P9 Gosse, C. Engineering FRET pairs to study the dynamics of protein conformational changes with a frequency‐domain perturbative technique involving temperature oscillations

A‐P10 Hartmann, A. Two step millisecond kinetics of the fourU RNA thermometer revealed by single‐molecule FRET

A‐P11 Hartmann, S. Tools in protein chemistry for multi‐subunit and multidomain enzymes in smFRET

A‐P12 Heil, C. S. Investigating Structural Dynamics of Mega‐Enzymes by smFRET

A‐P13 Hugel, T. Mechanistic insights into the multi‐component Hsp90 machinery from smFRET

A‐P14 Inan, D. Investigation of Energy Transfer in Light Harvesting Antenna Systems

A‐P15 Isbaner, S. Simultaneous Measurements of Thickness and Diffusion of Lipid Bilayers using MIET and 2f‐FLCS

A‐P16 Kallis, E. Single‐molecule FRET experiments for investigating the binding mechanism of the enzyme PARP‐1 to DNA single‐strand breaks

A‐P17 Karedla, N. Single‐Molecule Metal Induced Energy Transfer (smMIET): Resolving nanometer distances at single molecule level

A‐P18 Kramm, K. Selective bioorthogonal labeling of protein complexes in cell extracts for fast single‐molecule pulldown assays

A‐P19 Kubiak, J. Fluorescence spectroscopy reveals large‐amplitude conformational dynamics of the multifunctional protein GABARAP

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A-P20 Kühnemuth, R. Effect of dye-linker dynamics on the interpretation of FRET experiments

A-P21 Lerner, E. An off-pathway route in Escherichia Coli transcription initiation

A-P22 Ljubetič, A. Folding pathways of designed protein polyhedral A-P23 Margeat, E. Structural dynamics of metabotropic glutamate receptors

by single-molecule FRET A-P24 Melinger, J. S. FRET in Fluorophore-Labeled DNA Crystals A-P25 Molle, J. Probing the same inter-dye-distance of 6 nm by super-

resolution DNA-PAINT and FRET on DNA Origami A-P26 Moparthi, S. B. A Comparison of MreB Conformations upon Interactions

with GroEL/ES and Tail-less Complex Polypeptide 1 Ring Complex (TRiC) Chaperonins

A-P27 Nettels, D. Excited-state annihilation reduces power dependence of single molecule FRET experiments

A-P28 Prakash, A. Position dependent fluorescent properties of coupled fluorescent dyes in large doublestranded RNA

A-P29 Prieto, M. Amyloid-like fibers and the role of lipids: Multibilayer structure and protein oligomerization from FLIM-FRET and homo-FRET methodologies

A-P30 Salsi, E. Following the Structural Dynamics of Elongation Factor G during Ribosomal Translocation

A-P31 Sarangamath, S. qAN4: a second generation adenine analog as fluorescent probe to understand conformation and dynamics of nucleic acids.

A-P32 Sohail, A. The environment shapes the inner vestibule of LeuT A-P33 Schneider, S. Pressure unfolding of a model folding protein followed by

smFRET A-P34 Steffen, F. D. Dissecting carbocyanine photophysics in the context of

RNA A-P35 van der Velde,

J. H. M. Enhancing single-molecule FRET studies with photostabilizer-dye conjugates

A-P36 Wengler, D. Studying the function of BAP in the nucleotide cycle of BiP by spFRET using MFD-PIE

A-P37 Wilhelmsson, L. M.

Fluorescent nucleobase analogue development and their use in FRET-investigations

A-P38 Yushchenko, D. Probes to study AS-membrane interactions A-P39 Zhao, M. Site-specific labeling of large RNA with fluorophores for

the application in single molecule FRET studies

Topic B: FRET Theory, Analysis and Instrumentation

B-P40 Barth, A. PIE analysis in MATLAB (PAM) – A software package for the analysis of fluorescence experiments using pulsed interleaved excitation

B-P41 Cook, N. P. Extended excitation FLIM for rapid FLIM-FRET determinations of EGFR conformational dynamics and association states

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B-P42 Demchenko, A. P. Multiple FRET in molecular ensembles: site-photoselection effects

B-P43 Eilert, T. A specialized Sampling Engine for the Bayesian Inference Software Fast-NPS

B-P44 Hanley, Q. S. N-FRET in Assemblies and Clusters by Steady State Anisotropy: Theory and Practice

B-P45 Langhals, H. Förster Resonant Energy Transfer (FRET) in Orthogonally Arranged Chromophores. Limitations of the Theory

B-P46 Nagy, P. Understanding and Reducing the Error in the Evaluation of Intensity-based Microscopic FRET Experiments in the Presence of Low Signal-to-noise Ratio

B-P47 Schmid, S. Quantitative Protein Kinetics From smFRET Time Traces B-P48 Uri, A. Organic photoluminiscent probes possessing triplet-singlet

energy transfer by Förster mechanism B-P49 Zeug, A. Advanced spectral FRET approaches for quantitative

Imaging reveal importance of receptor oligomerization in serotonergic signaling

Topic C: FRET microscopy

C-P50 Abdollahi, E. Establishment of Sensitive Probes for Radiation-Induced Chromatin Decondensation Using Fluorescence Lifetime Imaging Microscopy

C-P51 Agam, G. Quantitative three-color FRET for the study of coordinated intramolecular motion

C-P52 Alexiev, U. FRET-FLIM as a tool to investigate interactions of hyaluronic acid with the skin and implications for the dermal delivery of biomacromolecules

C-P53 Diekmann, S. Next to CENP-A also the H3 variants H3.1 and H3.3 have defined cell-cycle dependent locations in centromeric chromatin

C-P54 Doll, F. Using FLIM-FRET microscopy to study protein-specific glycosylation in living cells

C-P55 Ebrecht, R. Imaging Gephyrin network formation at GABAergic postsynapsis by FLIM-FRET

C-P56 Haas, K. Mapping biochemical networks in single living cells by FLIM and multiplexed FRET: a Systems Microscopy approach

C-P57 Majoul, I. Bacterial toxin induced cAMP gradients shape Gap Junction responses

C-P58 Schuermann, K. C.

Simultaneous imaging of three FRET sensors by fluorescence anisotropy microscopy

C-P59 Seidel, C. A. M. Revealing structural features and affinities of protein complexes in living cells by MFIS-FRET analysis

C-P60 Weber, P. FRET-FLIM-Imaging: From 2D to 3D Cell Culture Systems

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Topic D: Hybrid Approaches: FRET combined with Modelling and other Spectroscopies

D-P61 Dimura, M. Toolkit for multi-conformation biomolecular structure determination by high-precision FRET and molecular simulations

D-P62 Lenger, K. Homeopathy – Applied quantum physics D-P63 Ploetz, E. PIFE meets ALEX: observing both binding and

conformational changes in unlabelled proteins D-P64 Šachl, R. Determination of nanodomain sizes by FRET D-P65 Schmid, J. A. Studying dynamics of protein interactions by FRET-FRAP D-P66 Stockner, T. Determination of the Conformation of LeuT: Using an

Integrated Computational and Experimental Approach D-P67 Vanderberk, N. The E. coli Sec reaction pathway for cellular protein

sorting under a single molecule loupe Topic E: Biosensors, Probes and Nanoparticles

E-P68 Bhuckory, S. Multiplexed Detection of Three Different Tumor Biomarkers in a Rapid Time-Gated Terbium-to-Quantum Dot Homogeneous FRET Immunoassay

E-P69 Bouchaala, R. Near-Infrared FRET imaging reveals the fate and integrity of lipid nanocarriers in healthy and tumor-bearing mice

E-P70 Bremer, D. Longitudinal investigation of the calcium concentration in retinal neurons during chronic inflammation in vivo- a FRET-based approach

E-P71 Cardoso Dos Santos, M.

Intra and Extracellular Biosensing Using Time-Gated Terbium to Quantum Dot FRET

E-P72 Diaz, S. A. Utilizing Photochromic FRET to Develop Color Switching Quantum Dots

E-P73 Drees, C. Resolving Interactions Inside Living Cells With Engineered Upconversion Nanoparticles

E-P74 Field, L. D. Semiconductor Quantum Dots as Förster Resonance Energy Transfer Donors for Intracellularly-Based Biosensors

E-P75 Goryashchenko, A. S.

Genetically encoded caspase-3 FRET-sensor based on terbium chelate and red fluorescent protein

E-P76 Guo, J. Single Terbium-Quantum Dot FRET Pair for Time-Gated Lifetime Multiplexing of microRNA

E-P77 Hendrix, J. Using FRET to follow protein oligomerization throughout a viruses’ life cycle

E-P78 Hildebrandt, N. Multiplexed MicroRNA Detection Using Terbium to Quantum Dot FRET

E-P79 Höfig, H. Time-resolved FRET measurements of CFP-YFP-based biosensors

E-P80 Jin, Z. Multiplexed micro-RNA assays using time-resolved FRET with biospectral correction

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E-P81 Medintz, I. L. Energy Transfer-Based Sensitization of Luminescent Gold Nanoclusters

E-P82 Parimi, H. Homogeneous Proximity-Ligation FRET Immunoassays for VEGF Detection

E-P83 Petreto, A. Multiplexed and Homogenous FRET Diagnostics of MicroRNA

E-P84 Phillips, G. K. Developing fluorogen activating protein-fluorescent protein FRET pairs to investigate signalling of the high affinity IgE receptor (FcεR1)

E-P85 Trofymchuk, K. Exploiting fast exciton diffusion in dye-doped polymer nanoparticles to engineer efficient photoswitching through FRET

E-P86 Ulbricht, C. Towards functional imaging in vivo A FRET-based Genetically Encoded Calcium Indicator in B Cells

E-P87 Wiens, M. D. Development and characterization of tandem heterodimeric fluorescent proteins

E-P88 Zwier, J. Time gated FRET microscopy- and plate based assays to study G-Protein Coupled Receptors using bright Eu3+ and Tb3+ donors

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International Discussion Meeting “Förster Resonance Energy transfer in Life Sciences 2" (FRET 2)


Abstracts of oral presentations (in order of the schedule)

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Unwind and relax: Using single-molecule FRET

to dissect the mechanism of ATP-driven molecular machines

Alexandra Z. Andreou, Ulf Harms, Airat Gubaev, Dagmar Klostermeier

* University of Muenster, Institute for Physical Chemistry, Corrensstraße 30, 48149

Muenster, Muenster, Germany

[email protected]

Single molecule fluorescence techniques have been instrumental in understanding the mechanism of ATP-driven molecular machines. RNA helicases use the energy of ATP hydrolysis to separate RNA duplexes and to remodel RNA structures. They are involved in virtually all processes in RNA metabolism, from transcription, mRNA splicing and editing, transport and translation to RNA degradation. DEAD-box helicases are the largest family of RNA helicases, and share a common helicase core of two flexibly linked RecA-like domains. The activity of the core is modulated by additional domains or other protein factors. DNA topoisomerases inter-convert different DNA topoisomers and impact on key cellular processes such as replication, transcription, recombination, and the storage of the genome as chromatin. DNA gyrase introduces negative supercoils into DNA at the expense of ATP hydrolysis. DNA supercoiling is believed to follow a strand passage mechanism, in which a double-stranded DNA segment is cleaved, and a second double-stranded segment is passed through the gap, leading to the conversion of a positive DNA node into a negative node. In a combined approach of biochemical and biophysical methods, including single molecule FRET, we have dissected the conformational cycle of DEAD-box RNA helicase eIF4A [1]-[3], involved in translation initiation, and of DNA gyrase [4]-[8]. Recent insights into the mechanism of these enzymes will be presented, and the possibilities and limitations of single molecule FRET approaches will be discussed.

[1] Andreou, A.Z. & Klostermeier, D. (2014) eIF4B and eIF4G jointly stimulate eIF4A ATPase and unwinding activities by modulation of the eIF4A conformational cycle, J. Mol. Biol. 426(1): 51-61. [2] Harms, U., Andreou, A.Z., Gubaev, A., Klostermeier, D. (2014) eIF4B, eIF4G and RNA regulate eIF4A activity in translation initiation by modulating the eIF4A conformational cycle, Nucleic Acids Res. 42(12):7911-7922. [3] Andreou, A.Z. & Klostermeier, D.: „Fluorescence methods to investigate DEAD-box protein mechanisms“, pp. 161-192 in “Fluorescent methods applied to molecular motors: from single molecules to whole cells” (292 pages), edited by C. Toseland & N. Fili, 2013, Springer, Basel 2014. [4] Gubaev, A. & Klostermeier, D. (2014) Single molecule FRET reveals DNA- and nucleotide-induced conformational changes in DNA gyrase preceding the strand passage reaction, for the DNA repair Special Issue “Single molecule approaches: watching DNA repair one molecule at a time”, DNA repair 16: 23-34. [4] Lanz, M.A., Farhat, M., Klostermeier, D. (2014) The acidic C-terminal tail of GyrA down-regulates the DNA supercoiling activity of B. subtilis gyrase, J. Biol. Chem. 289(18):12275-12285. [5] Lanz, M.A. & Klostermeier, D. (2012) The GyrA-box determines the geometry of DNA bound to gyrase and couples DNA binding to the nucleotide cycle, Nucleic Acids Res. 40(21):10893-903. [6] Gubaev, A. & Klostermeier, D. (2012) Potassium ions are required for nucleotide-induced closure of the gyrase N-gate, J. Biol. Chem. 287(14):10916-21. [7] Lanz, M.A., Klostermeier, D. (2011) Guiding strand passage: DNA-induced movement of the gyrase C-terminal domains defines an early step in the supercoiling cycle, Nucleic Acids Res. 39(22):9681-94. [8] Gubaev, A., Klostermeier, D. (2011) DNA-induced narrowing of the gyrase N-gate coordinates T-segment capture and strand passage, Proc. Natl. Acad. Sci. 108(34):14085-90.

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Single-molecule dynamics of a SERCA homologue Mateusz Dyla*, Daniel Terry**, Magnus Kjaergaard*, Jacob Lauwring Andersen*, Charlotte

Rohde Knudsen*, Victoria Birkedal***, Scott C. Blanchard**, Poul Nissen*

* Centre for Membrane Pumps in Cells and Disease – PUMPKIN. Danish National ResearchFoundation & Danish Research Institute of Translational Neuroscience – DANDRITE,

Nordic-EMBL Partnership for Molecular Medicine. Aarhus University, Denmark, Department of Molecular Biology and Genetics, Gustav Wieds Vej 10C, DK – 8000 Aarhus C.

** Department of Physiology and Biophysics, Weill Cornell Medical College, New York, NY 10065, USA

*** Interdisciplinary Nanoscience Center (iNANO), Aarhus University, Gustav Wieds Vej 14, DK – 8000 Aarhus C.

[email protected]

Approximately 30% of the ATP generated in the living cell is utilized by the P-type ATPase primary active transporters to generate and maintain electrochemical gradients across biological membranes. P-type ATPases undergo large conformational changes alternating between the E1 and E2 states during their functional cycle to couple ATP hydrolysis (via phosphoenzyme formation and breakdown) in the cytoplasmic domains to ion transport across the membrane region, ~50 Å away. A well-characterized and representative P-type ATPase, the Listeria monocytogenes Ca2+-ATPase LMCA1, was engineered and characterized tofacilitate single-molecule FRET studies of transport-related structural changes of a P-type ATPase. LMCA1 was found to reside in the high-FRET E1 conformational state through most of its functional cycle, even in the absence of Ca2+. Binding of Ca2+ brought the cytoplasmicdomains of LMCA1 closer together, whereas transitions to the E2 state triggered by ATP binding and phosphorylation were very brief, and could only be characterized in a dephosphorylation-deficient LMCA1 mutant. Owing to a spontaneous dephosphorylation of this mutant, full transport cycles at a single-molecule resolution were observed for the first time.

Fig. 1. LMCA1 dynamics during its functional cycle. Homology models of LMCA1 are shown in six functional states, labeled with the corresponding Post-Albers state on top of the picture. Amino acids constituting the labeling sites are shown as spheres, and are labeled with amino acid name,

number and the domain it belongs to in the leftmost homology model. The distances between labeling sites are shown as black lines with distances in Å shown above. Two intrinsic cysteines located in a

transmembrane region are shown as black spheres.

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Structural Ensembles of Intrinsically Disordered Proteins From Simulation and Experiment

Sarah Rauscher*, Reinhard Klement*, Timo Graen*, Vytautas Gapsys*, Man Zhou**, Qui Van**, Michal Gajda***, Markus Zweckstetter***, Joerg Enderlein**, Bert L. de Groot*,

Helmut Grubmüller*

*Dept. of Theoretical and Computational Biophysics, Max Planck Institute for BiophysicalChemistry, Göttingen, Germany, **III Institute of Physics, University of Goettingen, Göttingen, Germany, ***Department of NMR-based Structural Biology, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany. [[email protected]]

Intrinsically disordered proteins (IDPs) are notoriously challenging to study both experimentally and computationally. The structure of IDPs cannot be described by a single conformation, but must instead be described as an ensemble of interconverting conformations. Atomistic simulations are increasingly used to obtain such IDP conformational ensembles. Here, we focus on obtaining accurate structure ensembles by combining FRET, FCS, NMR, and SAXS data with all atom molecular dynamics simulations for three IDPs: α-synuclein, an RS peptide and FG-nucleoporin peptides. In a first step, to assess the quality of available force fields, we have compared the IDP ensembles generated by nine all-atom empirical force fields against primary small angle x-ray scattering (SAXS), NMR, and fluorescence correlation spectroscopy (FCS) data for an RS peptide and FG-nucleoporin peptides. Ensembles obtained with different force fields exhibit marked differences in chain dimensions, hydrogen-bonding, and secondary structure content. These differences are unexpectedly large: changing the force field is found to have a stronger effect on secondary structure content than changing the entire peptide sequence. The CHARMM 22* ensemble performs best in this force field comparison: it has the lowest error in chemical shifts and J-couplings, and agrees well with the SAXS and FCS data. To eliminate inadequate sampling as a reason for differences between force fields, extensive simulations were carried out (1.351 ms in total); the remaining small sampling uncertainty is shown to be much smaller than the observed differences. These findings highlight how IDPs, with their rugged energy landscapes, are highly-sensitive test systems that are capable of revealing force field deficiencies and, therefore, contributing to force field development. In a second step, we have simulated ensemble Forster Resonance Energy Transfer (FRET) measurements with a tryptophan/coumarin FRET dye pair at eight different labeling positions in α-synuclein. Each labeled peptide was sampled from eight different starting structures for >50 µs, using different force field / water model combinations. Taking the three competing decay channels of tryptophan as well as their environment dependence into account, FRET fluorescence decay curves were calculated and compared to experiment. We found that the AMBER03Ws//TIP4P2005 and CHARMM22*//TIP4P-D ensembles agree best with the trFRET measurements.

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Multiparameter Fluorescence spectroscopic toolkit resolves the heterogeneity of unfolded states

Hemmen, Katherina*; Rodnin, Dmitro*; Markovic, Igor**; Felekyan, Suren*; Kühnemuth, Ralf*; Sanabria, Hugo**; Seidel, Claus*

* Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie,Heinrich-Heine-Universität, 40255 Düsseldorf, Germany.

** Department of Physics and Astronomy, Clemson University, Clemson, SC, U.S.A.

[email protected]

Intense debate exists on the possibility that unfolded proteins show, to certain extent, residual secondary structure. Transient structure formation might facilitate folding and/ or enhance binding to ligands. Some proteins such as those categorized as intrinsically disordered proteins take advantage of these features. Here, we used the model enzyme lysozyme from the phage T4 (T4L), a simple two subdomain protein, which already exists in an equilibrium of at least three conformations under native conditions. We created a set of 24 double mutants of the cysteine-free pseudo-wild type by inserting an unnatural amino acid and a cysteine mutation and site-specifically labeled them using orthogonal chemistry with a Förster resonance energy transfer (FRET) dye pair. Our set of variants allows us to build up a network of distances spanning the enzyme in order to monitor the ensemble of conformations. The behavior of the protein under highly denaturing conditions was observed by a combination of ensemble (ensemble time-resolved fluorescence lifetime and anisotropy) and single-molecule spectroscopic (multiparameter fluorescence detection, photon distribution analysis, (filtered) fluorescence correlation spectroscopy) methods. Our network covered all possible directions over the whole protein allowing us to map specifically the local motions (dye mobility) and global changes. We identified regions with residual structure which exist even under highly denaturing conditions. Additionally, the combination of ensemble and single-molecule methods allows us to resolve the full heterogeneity of the proteins’ denatured conformations.

1E-8 1E-5 0.01 10

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Fig. 1. Our combinational approach probes states distribution and dynamics over wide range of time scales. Fluorescence lifetime probes the static distribution of states, whereas fluorescence anisotropy experiments describe the local, backbone dynamics of the sample. Single-molecule experiments in terms of multiparameter fluorescence detection (MFD) and photon distribution

analysis (PDA) show the population distribution and its dynamics on timescales similar to burst duration (~ 1 ms). Fast dynamics can be resolved with fluorescence correlation methods (fullFCS and

filteredFCS) from both single and double labeled molecules.

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Maximum Likelihood Analysis in Single-Molecule FRET

Irina V. Gopich

Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892, USA

[email protected]

Single-molecule fluorescence spectroscopy is widely used to study macromolecular dynamics. The output of such measurements is a sequence of photons of different colors separated by random time intervals. In the experiments with pulsed lasers, fluorescence lifetimes can also be monitored. To improve the range of the measured dynamics at a given photon count rate, we consider each and every photon and use a maximum likelihood method to get the information about fast conformational dynamics. For a photon trajectory with recorded photon colors, interphoton times, and delay times (relative to laser pulses), the parameters of a model describing molecular dynamics are obtained by maximizing the appropriate likelihood function [1-3]. We discuss various aspects of the maximum likelihood analysis, such as likelihood functions and their applicability, the accuracy of the extracted parameters [4], their sensitivity to the model assumptions [3], and the influence of fast blinking [5]. The method has been applied to study fast folding proteins [3, 5].

[1] I.V. Gopich, A. Szabo. Decoding the pattern of photon colors in single-molecule FRET. J. Phys. Chem. B.; v. 113, 10965–10973 (2009).[2] I.V. Gopich, A. Szabo. Theory of the energy transfer efficiency and fluorescence lifetime distribution in single-molecule FRET. Proc. Natl. Acad. Sci. U.S.A.; v. 109, 7747 (2012). [3] H. S. Chung and I. V. Gopich. Fast single-molecule FRET spectroscopy: theory and experiment. Phys. Chem. Chem. Phys.; v. 16, 18644-18657 (2014). [4] I. V. Gopich. Accuracy of Maximum Likelihood Estimates of a Two-State Model in Single-Molecule FRET. J. Chem. Phys.; v. 142, 034110 (2015). [5] H. S. Chung, J. M. Louis, and I. V. Gopich. Analysis of Fluorescence Lifetime and Energy Transfer Efficiency in Single-Molecule Photon Trajectories of Fast-folding Proteins. J. Phys. Chem. B; (2016), in press.

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Accounting for experimental errors in HMM analyses of single-molecule FRET experiments

Oliver Lemke*, Frank Noé**, and Bettina G. Keller *

* Department of Biology, Chemistry, and Pharmacy, Freie Universität Berlin ** Department of Mathematics and Computer Science, Freie Universität Berlin

[email protected]

In recent years hidden Markov models (HMM) have been successfully applied for the analysis of single-molecule FRET experiments. In HMMs, a model of the conformational dynamics and of the FRET process is varied such that the likelihood with which this model could have generated the experimental data set is maximized. The experimental FRET trace is however not only determined by the underlying time-series of conformations and their associated FRET efficiencies, but it is modified and skewed by a whole range of other photo-physical processes, such as spectral cross-talk, background noise, or differences in the quantum yields of the chromophores. To obtain meaningful models of the conformational dynamics, it is therefore of crucial importance that the likelihood function models the experiment as accurately as possible. I will discuss how different experimental errors can be incorporated into the likelihood function of the HMM and thereby improve the quality of the model. Related to this is the question of how to compare and rate different models derived from the same data set. I will demonstrate tests to check the self-consistency of the HMM and its consistency with the experimental data.

Fig. 1. The two components of a hidden Markov model: hidden conformational dynamics x(t) and emission process o(t).

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Multi-color single molecule FRET study of intrinsically disordered protein binding Hoi Sung Chung*

* Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and KidneyDiseases, National Institutes of Health (NIH), Bethesda, MD, 20892-0520

[email protected]

Intrinsically disordered proteins (IDPs) are unstructured at the native condition and fold when attaching to their binding partners. Understanding the mechanism of this process requires probing conformational changes of IDPs during binding processes. Since multiple binding pathways should exist as protein folding, single-molecule spectroscopy is expected to provide unique information such as the distribution of binding pathways. In order to probe conformational changes of IDPs and their interactions with binding targets simultaneously, it is necessary to obtain the distance information between more than two fluorophores. As a first step toward this direction, we performed two- and three-color FRET spectroscopy to study the oligomerization of the tetramerization domain (TD) of the tumor suppressor protein p53. Two monomers of TD form a dimer at low nM concentration and subsequently two dimers form a tetramer at higher concentration. In the three-color FRET experiment of the dimerization, one monomer TD construct was labeled with Alexa 488 and Alexa 647 and immobilized on a PEG-coated glass coverslip via a biotin-streptavidin linkage. Another TD construct was labeled with Alexa 750 as a binding partner in solution. Using the alternating excitation of two picosecond-pulsed lasers (485 and 640 nm) at 40 MHz, it was possible to detect all three FRET efficiencies between three fluorophores. In addition, from the average delay times between photon arrivals and laser excitation, the fluorescence lifetime of each state was measured. Interestingly, we observed multiple conformations in the dimer state that are different from the conformation in the tetramer. The faster formation of the tetramer than the dissociation of the dimer at high TD concentration suggests that these dimers with different conformations should be on pathway intermediates during the tetramerization process.

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Protein folding dynamics from single-molecule FRET – from in vitro to in vivo Schuler B*

* University of Zurich, Department of Biochemistry, 8057 Zurich, Switzerland

[email protected]

Single-molecule spectroscopy provides new opportunities for investigating the structure, folding and dynamics of proteins. The combination of single-molecule Förster resonance energy transfer (FRET) with nanosecond correlation spectroscopy, microfluidic mixing, and related methods can be used to probe intramolecular distance distributions, reconfiguration dynamics, folding, and interactions on a wide range of timescales [1], and even in heterogeneous environments [2]. Recent developments now allow us to compare these processes directly in vitro and in live cells [3].

[1] Schuler, B. & Hofmann, H. (2013) Single-molecule spectroscopy of protein folding dynamics - expanding scope and timescales. Curr. Opin. Struct. Biol. 23, 36-47. [2] Soranno, A., Koenig, I., Borgia, M.B., Hofmann, H., Zosel, F., Nettels, D., & Schuler, B. (2014) Single-molecule spectroscopy reveals polymer effects of disordered proteins in crowded environments. Proc. Natl. Acad. Sci. USA 111, 4874-4879. [3] König, I., Zarrine-Afsar, A., Aznauryan, M., Soranno, A., Wunderlich, B., Dingfelder, F., Stüber, J.C., Plückthun, A., Nettels, D. & Schuler, B. (2015) Single-molecule spectroscopy of protein conformational dynamics in live eukaryotic cells. Nat. Methods 12, 773-779.

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Metal-induced energy transfer for live cell nanoscopy Chizhik A. I. *, Ruhlandt D.*, Baronsky, T.**, Chizhik A. M.*, Karedla, N.*, Rother, J.**,

Gregor, I.*, Janshoff, A.**, Enderlein, J.*

*III. Institute of Physics, Georg August University, 37077 Göttingen, Germany.

** Institute of Physical Chemistry, University of Göttingen, Tammannstrasse 6, 37077 Göttingen, Germany.

[email protected]

The discovery of Förster resonance energy transfer (FRET) [1] has revolutionized our ability to measure inter- and intramolecular distances on the nanometre scale using fluorescence imaging. The phenomenon is based on electromagnetic-field-mediated energy transfer from an optically excited donor to an acceptor. We replace the acceptor molecule with a metallic film and use the measured energy transfer efficiency from donor molecules to metal surface plasmons [2] to accurately deduce the distance between the molecules and metal [3]. Like FRET, this makes it possible to localize emitters with nanometre accuracy, but the distance range over which efficient energy transfer takes place is an order of magnitude larger than for conventional FRET. This creates a new way to localize fluorescent entities on a molecular scale, over a distance range of nearly 200 nm. We demonstrate the power of this method by profiling the basal lipid membrane of living cells.

Fig. 1. 3D reconstruction of the basal cell membrane allows for observing its motion with 2 nanometer accuracy. See ref. 3 for further details.

[1] Förster, Th. “Zwischenmolekulare energiewanderung und fluoreszenz”, Ann. Phys. 437, 55-75 (1948). [2] Drexhage, K. H. “Interaction of light with monomolecular dye layers”, Prog. Opt. 12, 163-232 (1974). [3] Chizhik, A. I., Rother, J. Gregor, I., Janshoff, A., Enderlein, J. “Metal-induced energy transfer for live cell nanoscopy”, Nature Photon. 8, 124-127 (2014).

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Quantitative FRET imaging reveals the interdependence of vesicular membrane dynamics and growth factor signal processing

Philippe Bastiaens

Department of Systemic Cell Biology, Max Planck Institute for Molecular Physiology, Otto Hahn Str. 11, 44227, Dortmund.

Autocatalytic phosphorylation of receptor tyrosine kinases (RTKs) enables diverse, context-dependent responses to extracellular signals but comes at the price of autonomous, ligand-independent activation. In order to illuminate the relevance of the spatial dimension in the response properties of growth factor receptors, I will describe how quantitative FRET imaging approaches enabled us to elucidate where and when protein tyrosine phosphatases interact with RTKs. For example, by comparing epidermal growth factor receptor (EGFR) spatial phosphorylation patterns upon knock-down or overexpression of PTPs using cell-array fluorescence lifetime imaging microscopy (CA-FLIM), we could identify which PTPs regulate EGFR phosphorylation where and when in the cell. The spatially exerted control of PTPs on vesicular RTK activity preserves ligand responsiveness but also shapes the finite response to growth factors. Vesicular membrane dynamics thereby control the autocatalytic activation and response of receptor tyrosine kinases by regulating when receptors encounter spatially partitioned PTP activities in the cell.

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3D fluorescence anisotropy imaging using selective plane illumination microscopy

Per Niklas Hedde, Suman Ranjit, and Enrico Gratton

Laboratory of Fluorescence Dynamics, Department of Biomedical Engineering, University of California, Irvine, CA, USA.

[email protected]

Fluorescence anisotropy imaging is a popular method to visualize changes in organization and conformation of biomolecules within cells and tissues. In such an experiment, depolarization effects resulting from differences in orientation, FRET and rotational mobility of fluorescently labeled molecules are probed with high spatial resolution. Fluorescence anisotropy is typically imaged using laser scanning and epifluorescence-based approaches. Unfortunately, those techniques are limited in either axial resolution, image acquisition speed, or by photobleaching. In the last decade, however, selective plane illumination microscopy has emerged as the preferred choice for three-dimensional time lapse imaging combining axial sectioning capability with fast, camera-based image acquisition, and minimal light exposure. We demonstrate how selective plane illumination microscopy can be utilized for three-dimensional fluorescence anisotropy imaging of live cells. We further examined the formation of focal adhesions by three-dimensional time lapse anisotropy imaging of CHO-K1 cells expressing an EGFP-paxillin fusion protein.

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Combining FRET with single particle localisation microscopy to ascertain in cellulo EGFR oligomer structure

Chris Tynan, Laura Zanetti-Domingues, Sarah Needham, Daniel Rolfe, Michael Hirsch, Teodor Boyadzhiev, Selene Roberts, David Clarke and Marisa Martin-Fernandez

[email protected]

STFC Central Laser Facility, Research Complex at Harwell, Rutherford Appleton Laboratory, OX11 0QX, UK

The epidermal growth factor receptor (EGFR) initiates signals for cell proliferation and transformation. This receptor has an extracellular growth factor-binding domain (ECD), a single-pass transmembrane region, and an intracellular domain that has tyrosine kinase activity. The activation of the EGFR is triggered by the binding of small growth factor polypeptide ligands and involves the formation of dimers of these receptors.

FRET-microscopy is a useful tool to investigate receptor complexes on the cell surface, with the potential of providing crucial details on the activation mechanism of EGFR in the physiological context of the cell. The ubiquitous presence of FRET in EGFR dimers, where crystal structures suggested donor/acceptor probe separations >11 nm1,2, led to the hypothesis that EGFR also formshigher order oligomers, subsequently detected on the cell surface using image correlation methods3.Yet, little is known about their structures and their functional role in EGFR signalling. This is largely attributable to the lack of methods with sufficient resolution that can cover the range between 8-60 nm, where higher order oligomers are expected to fall.

To address this issue, we developed a super-resolution method based on fluorophore localisation imaging with photobleaching (FLImP)4 to investigate the geometry and size of oligomers of theEGFR family on the cell surface with ~ 6 nm resolution. By using non-activating peptide markers and combining the FLImP super-resolution method with FRET to determine intra-receptor conformation, we are beginning to determine conformational changes, conformational coupling, and interactions in higher order oligomers that regulate the basal state and EGFR signal transduction across the plasma membrane.

[1] R. Ishitani, O. Nureki, S. Fukai, M. Yamanaka, J. H. Kim, K. Saito, A. Sakamoto, M. Inoue, M. Shirouzu, S. Yokoyama, Crystal structure of the complex of human epidermal growth factor and receptor extracellular domains. Cell 110, 775-787 (2002). [2] T. P. Garrett, N. M. McKern, M. Lou, T. C. Elleman, T. E. Adams, G. O. Lovrecz, H. J. Zhu, F. Walker, M. J. Frenkel, P. A. Hoyne, R. N. Jorissen, E. C. Nice, A. W. Burgess, C. W. Ward, Crystal structure of a truncated epidermal

growth factor receptor extracellular domain bound to transforming growth factor alpha. Cell 110 (2002) 763-773.3 [3] Saffarian, S., Li, Y., Elson, E. L. & Pike, L. J. Oligomerization of the EGF receptor investigated by live cell fluorescence intensity distribution analysis. Biophys J 93, 1021-1031, doi:10.1529/biophysj.107.105494 (2007).[4] S. R. Needham, M. Hirsch, D. J. Rolfe, D. T. Clarke, L. C. Zanetti-Domingues, R. Wareham, M. L. Martin-Fernandez, Measuring EGFR Separations on Cells with similar to 10 nm Resolution via Fluorophore Localization Imaging with Photobleaching. PLoS One 8 (2013) e62331.

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Frontiers in biochemical imaging: multiplexed FRET sensors and OptogeneticsEsposito A, Haas K, Fries MW, Campbell CJ, De S and Venkitaraman AR

The Medical Research Council Cancer Unit at the University of Cambridge,

Hutchison/MRC Research Centre, Box 197, Biomedical Campus

Cambridge, CB2 0XZ, United Kingdom

[email protected]

A large number of molecules cooperate in an intricate network of interactions for the maintenance of the structural integrity, the metabolism and the function of the living cell. A challenge for engineering and physics in optical microscopy is to provide tools that could offer the highest spatio-temporal resolution with the capability to decode complex networks of molecular interactions [1] by the development of technologies and methods that, at the same time, may provide cost-effective and user-friendly instruments [2].

We have developed a number of technology platforms based on multi- or hyper- spectral FLIM and novel FRET pairs dedicated to biochemical multiplexing. We are now integrating these sensing platforms with Optogenetics [3], in order to enable the quantitative spatio-temporal control of biochemical reactions in the living cells. With these techniques, we aim to perform perturbation analysis of biochemical networks in living cells and to correlate network topologies and their heterogeneity with cellular decisions.

We will describe the challenges we are confronting and our solutions, with particular emphasis on the development of novel efficient and fast FLIM technologies and how they can be integrated with Optogenetics in order to deploy a Systems Microscopy approach to the study of cell biology.

FFig. 1. A vision for Systems Microscopy. a) perturbational analysis of biochemical networks may

allow to quantify how biochemical pathways are disrupted in cancer in order to optimize our therapeutic approach. b) Integrating of biochemical multiplexing technologies with optogenetics.

[1] S. D. M. Santos et al., “Growth factor-induced MAPK network topology shapes Erk response determining PC-12 cell fate” Nat. Cell Biol., 9, 324–330 (2007) [2] A. Esposito et al., “Maximizing the Biochemical Resolving Power of Fluorescence Microscopy” PLOS ONE DOI: 10.1371/journal.pone.0077392 (2013) [3] A. Levskaya et al., “Spatiotemporal Control of Cell Signalling Using A Light-Switchable Protein Interaction” Nature, 461(7266) 997–1001 (2009)

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FRET sensing of metabolites: dedicated FLIM sensors, dedicated FLIM readout

Marcel Raspe1, Katarzyna M. Kedziora1, Bram van den Broek1, Qiaole Zhao2, Sander de Jong3, Johan Herz3, Marieke Mastop4, Joachim Goedhart4,5, Theodorus W.J. Gadella4,5, Ian Ted Young2, Kees Jalink1,5

1 The Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX, Amsterdam, The Netherlands 2 Delft University of Technology, Lorentzweg 1, 2628 CJ, Delft, The Netherlands 3 Lambert Instruments B.V., Leonard Springerlaan 19, 9727 KB, Groningen, The Netherlands 4 University of Amsterdam, Science Park 904, 1098 XH Amsterdam, The Netherlands. 5 Van Leeuwenhoek Centre for Advanced Microscopy, Amsterdam, The Netherlands

Corresponding author: KJ ([email protected])

Förster(Fluorescence) Resonance Energy Transfer (FRET) has become a powerful tool to study the inner workings of the cell. FRET may occur when an excited donor fluorophore is in very close proximity to a suitable acceptor: the excited state energy is transferred from the donor to the acceptor, with a consequent increase in emission from the acceptor and decrease in donor emission. Typical applications include the readout, both in living cells and entire organisms, of protein-protein interactions and the analysis of metabolic states of cells.

FRET is commonly read out either by detecting the ratio of the donor and acceptor intensities (sensitized emission) or by detecting the excited state lifetime of the donor, which decreases with increasing FRET (Fluorescence Lifetime IMaging or FLIM). Sensitized emission is fast and simple, but due to channel leak-through and ongoing bleaching, it requires quite some calibration in order to arrive at quantitative results. FLIM, on the other hand, is more immune to bleaching and inherently quantitative, but typically much slower and less photon-efficient. Moreover, FLIM detection typically requires several images to be collected from the cells, leading to potential artifacts in lifetime when FRET changes rapidly, for example, during fast transients in metabolite concentrations, and when vesicles move within the cell.

Using a new generation of cameras capable of collecting two phase images simultaneously, we developed a fast and artifact-immune technique to obtain lifetime images in just a single image. This approach, which we termed single-image FLIM (siFLIM), dramatically increases speed and photon efficiency of fluorescence lifetime detection[1]. We show that despite of ultimate photon efficiency. siFLIM lifetimes closely follow those of conventional (12-frame) FLIM experiments.

To match the power of siFLIM, we also developed a series of dedicated FRET sensors tailored for FLIM detection. These sensors employ mTurquoise2[2] , regarded as currently the best cyan fluorescent protein with excellent quantum yield, very high photostability and single-exponential decay, as a donor, and they have dark variants of YFP as acceptors. We sandwiched the cAMP-binding protein EPAC between the mTurquoise2 donor and a tandem of two monomeric dark Venus acceptors to generate a superior cAMP sensor[3], and also generated a novel dark-acceptor sensor to report activity of Gαq in living cells.

[1] Raspe et al, siFLIM: Single Image Frequency-Domain FLIM provides fast and photon-efficient lifetime data; submitted. [2] Goedhart et al, Nat Commun. 2012; [3] Klarenbeek et al, 4th-generation cAMP FRET sensors, PLoS One, 2015.

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mScarlet, a novel high quantum yield monomeric red fluorescent protein with strongly enhanced properties for sensitized emission-based FRET

Lindsay Haarbosch*, Daphne Bindels*, Laura van Weeren*, Marieke Mastop*, Marten Postma*, Antoine Royant** and Dorus Gadella*

*Section of Molecular Cytology & van Leeuwenhoek Centre for Advanced Microscopy,Swammerdam Institute for Life Sciences, University of Amsterdam, Science Park 904, NL-

1098 XH, The Netherlands.

**Structural Biology Group, European Synchrotron Radiation Facility, 38043 Grenoble, France

[email protected]

mScarlet, a novel red fluorescent protein was generated from a synthetic template based on a consensus amino acid sequence derived from naturally occurring red fluorescent proteins and purple chromoproteins and on consensus monomerization mutations. The encoded synthetic red fluorescent protein was optimized by molecular evolution through site directed and random mutagenesis. Improved variants were selected by quantitative multimode screening for increased fluorescence lifetime, increased photo stability, increased quantum yield and for increased chromophore maturation.

Very bright variants were obtained with high fluorescence lifetimes up to 3.8 ns, quantum yields >65% and complete maturation. The monomeric status of the variants was confirmed by OSER analysis and with a-tubulin fusions. The brightness of mScarlet is >2-fold increased as compared to bright red fluorescent proteins such as mCherry, mRuby2 and tagRFP-T as was analyzed with quantitative (single plasmid with viral 2A sequence) co-expression with mTurquoise2 in mammalian cells.

During evolution mScarlet variants with substantially altered spectroscopic properties were generated including fluorescence lifetime variants, photo labile variants and strongly spectrally shifted variants.

Because of their efficient maturation and high quantum yield, mScarlet vastly outperforms existing monomeric red fluorescent proteins in ratiometric FRET applications due to seriously enhanced sensitized emission.

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Cyanine Dyes in Biophysical Research: The Photophysics of Cy-dyes on DNAMarcia Levitus *

* School of Molecular Sciences and The Biodesign Institute at Arizona State University.Tempe, AZ, USA

[email protected]

Cyanine dyes are very commonly used in biophysical applications of single molecule fluorescence and fluorescence spectroscopy in general. For instance, the FRET pair Cy3-Cy5 is a popular choice in biophysical studies of DNA structure and dynamics, DNA-protein interactions, etc. As single molecule FRET measurements become more quantitative, it is becoming increasingly important to understand how the properties of the probes influence the signals measured in these experiments, and how the presence of the probes may affect the structure of the biopolymers under investigation. We have done extensive work characterizing the photophysical properties of Cy3 covalently attached to DNA. In this talk I will summarize our recent work on the photophysical properties of Cy-dyes, emphasizing the significance of the results for the interpretation of FRET data. The fluorescence quantum yield and lifetime of Cy3 are particularly sensitive to the physical properties of its microenvironment, and we have demonstrated that this is due to the existence of a non-fluorescent photoisomer that is formed with an efficiency that decreases when dye-DNA interactions hinder the rotation of a double bond. Using experiments and computer simulations we demonstrated that Cy3-DNA interactions are dynamic in nature, and depend strongly on DNA sequence. These results are significant for the analysis of FRET data because Cy3-DNA interactions impact the well-known kappa-square factor, and also affect the photophysical properties of the dye that are relevant in the calculation of Förster's distance.

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Single molecule FRET studies of DNA structures and devicesVictoria Birkedal *,**

* Interdisciplinary Nanoscience center (iNANO), Aarhus University, Aarhus, Denmark

** Centre for DNA nanotechnology (cDNA), Aarhus University, Aarhus, Denmark

[email protected]

FRET spectroscopy and single molecule FRET microscopy are powerful tools to investigate structural dynamics properties of DNA-based devices and structures. DNA has been used as a building block in the design and assembly of a number of DNA devices, whose operation is often based on conformational changes between several states [1]. Sizes range from a few nanometers to several hundreds of nanometers. Using single molecule methods, individual molecules can be followed in action and thus a direct view can be obtained of how they work. We focus here on two DNA devices: a DNA actuator consisting of two piston arms that can move with respect to each other with sub-nanometre steps and a total movement of 7 nm [2] and on a DNA origami box, a nanocontainer whose lid can be opened and closed with DNA triggers [3] (Figure 1). Our results give a direct insight into the movement, conformation changes and performance of these DNA devices. I will also discuss our data analysis procedures using the single molecule FRET iSMS platform [4].

Fig. 1. Schematic representation of A) the surface immobilized DNA actuator in two different conformations giving a total movement of 7 nm [2]. The two piston arms are shown in blue and red

respectively. B) the DNA origami box with two locks shown in blue and orange. Fluorophores, attached for FRET studies, are shown with stars [3].

[1] Y. Krishnan and F. C. Simmel, “Nucleic acid based molecular devices”, Angew. Chem. Int. Ed. 50, 3124 (2011). [2] L. L. Hildebrandt, S. Preus, Z. Zhang, N. V. Voigt, K. V. Gothelf, V. Birkedal, “Single Molecule FRET Analysis of the 11 Discrete Steps of a DNA Actuator”, J. Am. Chem. Soc. 136, 8957 (2014). [3] R. M. Zadegan, M. D. E. Jepsen, L. L. Hildebrandt, V.Birkedal, J. Kjems, “Construction of a fuzzy and all Boolean logic gates based on DNA”, Small 11, 1811 (2015) and E. S. Andersen, Mingdong Dong, M. M. Nielsen, K. Jahn, R. Subramani, W. Mamdouh, M. M. Golas, B. Sander, H. Stark, C. L. P. Oliveira, J. S. Pedersen, V. Birkedal, F. Besenbacher, K. V. Gothelf, and J. Kjems, “Self-assembly of a nano-scale DNA box with a controllable lid”, Nature 459, 73 (2009). [4] S. Preus, S. L. Noer, L. L. Hildebrandt, D. Gudnason, V. Birkedal, “iSMS: singlemolecule FRET microscopy software”, Nat. Methods, 12, 593 (2015).

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Co-translational Protein Folding on the Ribosome Monitored in Real Time Wolf Holtkamp1, Goran Kokic1, Marcus Jäger1, Joerg Mittelstaet1, Anton A. Komar3, Marina V.Rodnina1

1Department of Physical Biochemistry, Max Planck Institute for Biophysical Chemistry, 37077 Goettingen, Germany. 3Center for Gene Regulation in Health and Disease and the Department of Biological, Geological and Environmental Sciences, Cleveland State University, Cleveland, Ohio 44115, USA

Correspondence to: [email protected].

One of the major questions in the field of protein folding is how proteins attain their native three-dimensional structure during their synthesis on the ribosome when the polypeptide chain emerges from the peptide exit tunnel. Studies utilizing stalled ribosome-nascent chain complexes (RNC) suggested that protein domains can fold co-translationally into stable tertiary structures early during translation before the C-terminal part of the polypeptide chain is synthesized. However, the timing of these events in relation to translation is largely unknown. To investigate co-translational protein folding in real time we used a reconstituted in vitro translation system from E. coli and followed folding of the N-terminal domain (NTD) of PrmC (a N5-glutaminemethyltransferase) during ongoing translation in a time-resolved manner. We employed Förster resonance energy transfer (FRET) and photoinduced-electron transfer (PET) between two fluorophores introduced at different positions in the polypeptide chain, which serve as sensitive probes for ternary structure formation of the polypeptide during ongoing translation. The results suggest that compaction of the nascent polypeptide chain begins early during protein synthesis, whereas the native-like structure is attained after the whole domain emerged from the exit tunnel of the ribosome. Compaction and native state formation during elongation of the polypeptide are rapid and limited by the rate of translation. These finding support the idea of domain-wise folding of multi-domain proteins on the ribosome as an evolutionary mechanism to prevent kinetic trapping and unproductive protein folding.

Fig. 1. Co-translational HemK NTD folding monitored by FRET. Top panel, secondary structure elements of HemK NTD with a linker, helices H1-H5 are indicated by different colors. HemK112 denotes

the length (in amino acids) of the nascent peptide. PTC, peptidyl transferase center. Fluorescence labels BOF (BodipyFL) and BOP (Bodipy576/589) were introduced at the N-terminal Met and Lys34 residues, respectively. Kinetics of peptide synthesis was determined by analysis of fluorescence-labeled peptide

products on SDS-PAGE (bottom left panel). The two reporters, BOF and BOP, provide a donor-acceptor pair to FRET. When the two dyes come close due to folding, donor fluorescence decreases (bottom right

panel; BOF, green), while acceptor fluorescence increases (BOP, red) due to FRET.

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Enhanced dimerization of oncogenic EGFR mutants revealed by single molecule imaging and FRET microscopy

Christopher C. Valley*, Donna J. Arndt-Jovin**, Narain Karedla***, Mara P. Steinkamp*, Alexey I. Chizhik***, William S. Hlavacek****, Bridget S. Wilson*, Keith A. Lidke***** and

*Diane S. Lidke

*Department of Pathology and Cancer Research and Treatment Center, University of NewMexico, Albuquerque, NM, USA

**Laboratory of Cellular Dynamics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

***III. Institute of Physics, Georg-August University of Göttingen, Göttingen, Germany****Theoretical Biology and Biophysics Group, Theoretical Division, Los Alamos National

Laboratory, Los Alamos, NM, USA*****Department of Physics and Astronomy, University of New Mexico, Albuquerque, NM,

USA

[email protected]

Mutations within the epidermal growth factor receptor (EGFR/erbB1/Her1) are often associated with tumorigenesis. In particular, a number of EGFR mutants that demonstrate ligand-independent signaling are common in non-small cell lung cancer (NSCLC), including kinase domain mutations L858R and exon 19 deletions (e.g. ΔL747-P753insS). The molecular mechanisms by which these mutations confer constitutive activity remain unresolved. Using multiple sub-diffraction-limit imaging modalities, we reveal the altered receptor structure and interaction kinetics of NSCLC-associated EGFR mutants. Live cell FRET-FLIM measurements revealed that the L858R mutation alters the ectodomain structure such that unliganded mutant EGFR adopts an extended conformation (Fig. 1). This suggested that the mutants would be more available for dimerization. To test this this, we used two-color single particle tracking to quantify receptor dimerization kinetics on live cells. We found that the mutants are capable of forming stable, ligand-independent dimers. These data support a model where NSCLC-associated EGFR mutations induce a structural change in the

receptor that enhances dimerization and facilitates oncogenic signaling.

Fig. 1. FRET-FLIM measurements of EGFR structure in live cells [1]. (A) Schematic of the EGFR ectodomain in the tethered, autoinhibited conformation (left) and the extended conformation (right). The donor fluorophore (Oregon Green 488, green) is covalently linked at the EGFR N-terminus via a small acyl carrier protein (ACP) tag, and the acceptor fluorophore (NR12S, a derivate of Nile Red, red) is embedded in the outer leaflet of the plasma membrane. Intensity and lifetime images of EGFR-WT (B) or EGFR-L858R (C) cells.

[1] Valley et. al. “Enhanced dimerization drives ligand-independent activity of mutant EGFR in lung cancer title”, Molecular Biology of the Cell, vol. 26, page 4087, (2015).

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Camera-based Single-molecule FRET Detection With Improved Time ResolutionShazia Farooq* and Johannes Hohlbein*

*Laboratory of Biophysics, Wageningen University, Dreijenlaan 3,6703 HA Wageningen, The Netherlands

[email protected]

The achievable time resolution of camera-based single-molecule detection is often limited by the frame rate of the camera. Especially in experiments utilizing single-molecule Förster resonance energy transfer (smFRET) to probe conformational dynamics of biomolecules, increasing the frame rate by either pixel-binning or cropping the field of view decreases the number of molecules that can be monitored simultaneously. Here, we present a generalised excitation scheme termed stroboscopic alternating-laser excitation (sALEX) that significantly improves the time resolution without sacrificing highly parallelised detection in total internal reflection fluorescence (TIRF) microscopy [1,2]. In addition, we adapt a technique known from diffusion-based confocal microscopy to analyse the complex shape of FRET efficiency histograms. We apply both sALEX and dynamic probability distribution analysis (dPDA) to resolve conformational dynamics of interconverting DNA hairpins in the millisecond time range.

[1] S. Farooq and J. Hohlbein, “Camera-based single-molecule FRET detection with improved time resolution”, Phys. Chem. Chem. Phys., 17, 27862, (2015)

[2] J. Hohlbein, T.D. Craggs, and T. Cordes, “Alternating-laser excitation: single-molecule FRET and beyond”, Chem. Soc. Rev., 43, 1156-1171, (2014)

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Estimating the Distribution of FRET efficiencies from Fluorescence Lifetime-FRET Data

B.Wieb VanDerMeer *, Paul S. Blank** and Steven S.Vogel***

* Department of Physics and Astronomy, Western Kentucky University, Bowling Green,Kentucky, United States of America

**Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland, United States of America

*** Laboratory of Molecular Physiology, National Institute on Alcohol Abuse and Alcoholism, National Institutes of Health, Bethesda, Maryland, United States of America

[email protected]

Populations of FRET donor and accepter are typically characterized by an ensemble average FRET efficiency; the underlying distribution of individual FRET efficiencies remains unknown. Here we describe an approach to calculate this distribution from Time-resolved FRET data. We assume that the relevant transition moment motion rates of the Donor and Acceptor are either much smaller or much larger than the average lifetime of the excited donor state. The initial time-resolved-efficiency (TRE) [1] reflects the average rate or lifetime. At later times the contributions of smaller rates become increasingly more important. This suggests that the FRET rate distribution is contained in the TRE-versus-time-curve. It is possible to estimate this unknown distribution by expanding the distribution in a complete set of functions, so that the TRE becomes a specific time-dependent expression with calculable coefficients representing the contributions of the individual functions to the complete set. The coefficients are determined by fitting this function expression to the experimental TRE curve. In theory, any set of functions approximating the unknown distribution can be used. In practice, the set must be carefully chosen to ensure rapid convergence. Förster theory for a unique transfer rate constant, k , and a Donor lifetime in the absence of transfer, τ , gives the well-known relationship between the steady state efficiency, E , and k . This can be written

as kτ = R0 rDA( )6 = E 1− E( ) , with R0 representing the Förster distance at kappa-squared =23 , and rDA the distance between Donor and Acceptor. For the case of a distribution of rates

(i.e. static regime) this equation must be replaced by k τ = E 1− E( )N , where the brackets

denote averages in the distribution and N depends on E . N can be calculated from the distribution estimated from the TRE. For some distributions N does not strongly depend on E . For example, when the Acceptor-Donor distance is unique and kappa-squared varies in the standard fashion between 0 and 4, N is virtually independent of E and approximately equal to 2 [1]. However, it is possible and likely that, in other cases, N depends strongly on E and differs significantly from 1 or 2. Monte-Carlo simulations are useful in exploring how N varies with E for possible lifetime distributions. [1] B.W. VanDerMeer, D.M. VanDerMeer, and S.S. Vogel. “Optimizing the Orientations Factor Kappa-Squared for More Accurate FRET Measurements, Chapter 4, in FRET - Förster Resonance Energy Transfer, Igor Medintz and Niko Hildebrandt (editors), 2014 Wiley-VCH Verlag GmbH & Co. KGaH].

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Recent Advances in Lifetime FRET

Benedikt Krämer, Marcelle König, Paja Reisch, Rhys Dowler, Sandra Orthaus, MatthiasPatting, Tino Roehlicke, Felix Koberling and Rainer Erdmann

PicoQuant GmbH, Rudower Chausse 29, 12489 Berlin, Germany

[email protected]

Confocal laser scanning microscopes play an essential role in biological and biomedicalresearch. Their application can be further extended by incorporating Time-Correlated SinglePhoton Counting (TCSPC), giving access to time-resolved applications such as FluorescenceLifetime Imaging (FLIM).By fully exploiting the information contained in TCSPC decays, one can quantitativelyextract two types of information by investigating samples exhibiting Förster ResonanceEnergy Transfer (FRET). The first being the FRET efficiency and the second representing thepercentage of binding, i.e. the ratio of complete FRET molecular pairs compared to thenumber of molecules featuring only the donor part. A prerequisite for carrying out thisanalysis method is a mono exponential fluorescence decay of the donor fluorophore as foundin, e.g., T-Sapphire. We present here a pattern-matching analysis technique that allows identifying selectedfluorescence decay patterns at different spatial locations of the sample. The technique is easyto apply and allows for excellent discrimination of fluorophores even when they featuresimilar emission properties, e.g., dye molecules quenched due to FRET from theirunquenched counterparts. The speed of TCSPC FLIM measurements is generally restricted by the pile up limit. In orderto overcome this challenge, we have developed a TCSPC unit featuring a very short dead-time (below 1ns) and multi-stop capability. Combining this unit with a fast hybridphotomultiplier detector (PMA Hybrid) allows performing FLIM imaging in much shortertime spans than with conventional TCSPC systems. We call this novel approach RapidFLIM.These shorter acquisition times allow imaging with several FLIM images per second formonitoring e.g., transient molecular interactions as well as fast moving species. Even at veryhigh count rates, the mean fluorescence lifetime can be described accurately by a mono-exponential fit. However, due to residual dead-time effects, the fit quality of the TCSPC curveis compromised.

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Reversible cryo-arrest for imaging molecules in living cells at high spatial resolution

Martin E. Masip*, Jan Huebinger*, Jens Christmann*,**, Frank Wehner*, Günther R. Fuhr*** and Philippe I. H. Bastiaens*,**

*Department of Systemic Cell Biology, Max Planck Institute of Molecular Physiology, Otto-Hahn-Str.11, 44227 Dortmund, Germany

**Faculty of Chemistry and Chemical Biology, TU Dortmund, Otto-Hahn-Str. 6, 44227 Dortmund, Germany

***Fraunhofer Institute for Biomedical Engineering, Ensheimer Str. 48, 66386 St. Ingbert, Germany

Correspondence to: [email protected]

Dynamic movements of molecules that maintain structures within living matter hamper the observation of molecular patterns in cells. Therefore, the precise detection of molecular arrangements by single molecule localization microscopy (SMLM) or Förster Resonance Energy Transfer imaging (anisotropy-FRET, FLIM) usually requires irreversible and lethal chemical fixation. To circumvent this problem, an on-stage cryo-fixation approach was developed that allows for reversible arrest and therefore for the precise determination of molecular patterns at different points in time within the same living cell. We demonstrate the broad applicability of this reversible cryo-arrest by mapping the active states of epidermal growth factor receptor as well as its clustering in the plasma membrane by FLIM and SMLM, respectively.

Fig. 1. Fluorescence anisotropy of EGFR-QG-mCitrine before and 5 min after stimulation with EGF (right) and EGFR phosphorylation monitored by FLIM imaging during cryo-fixation (left).

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Figure 2. FLIM of 3T3 cells stained with Acridine Orange, low and high concentrations. Metachromatic stains: DNA, green; RNA, red. The lifetime images of designated cells on the left are shown on the right, with their respective color tables for the range of values.

Extended Excitation FLIM (eeFLIM) Thomas M. Jovin * and Nathan P. Cook*

* Laboratory of Cellular Dynamics, Max Planck Institute for Biophysical Chemistry,37077 Göttingen, Germany

[email protected]

The usual dogma in the field of fluorescence lifetime determination is that “the shorter the excitation pulse the better”. WRONG! We demonstrate that by recording the integrated emission of an emitting species excited with a rectangular light pulse with a duration ≥ ∼10·lifetime (reciprocal decay constant), sensitive and accurate determinations of the intensitymean lifetime are feasible. A series of successive determinations (2 suffice) are taken in the region corresponding to constant excitation intensity and at integration times > 6·the longest lifetime in the sample population. These points correspond to a straight line, the slope and position of which are referenced to a companion measurement of a sample with 0 lifetime (e.g. scattered excitation light) or known lifetime so as to yield the absolute mean lifetime. That is, the displacement on the integration time (gate width) axis is given by the lifetime (Fig. 1). The mixtures can be of arbitrary heterogeneity. For a two-component system (e.g. a binding reaction), the mean lifetime can be expressed analytically as a function of the fraction of species engaged in FRET. The mean value is very useful in numerous other applications (for FRET-based studies of the EGFR using eeFLIM see poster Cook et al.). eeFLIM ha s been implemented in an imaging system based on the gated intensified camera PI-MAX4-1024EMB of Princeton Instruments using laser diode for excitation (Fig. 2). This camera features excellent spatial resolution and linearity (emCCD detector), and powerful software + electronics for control of multimode acquisition and external synchronization. The system is very sensitive and allows real-time full-field (1K×1K) FLIM at rates that can exceed 1 Hz. Some important advantages of eeFLIM can be emphasized: (1) the rectangular excitation pulses (e.g. 10-50 ns) are easy to generate and provide very high pulse energies and thus intense response signals; (2) virtually all the light emitted per pulse (discounting detection efficiencies) is utilized; (3) the temporal resolution is tens of ps; (4) lifetime image calculations are very fast, involving only simple, linear, non-iterative calculations. It is anticipated that light sources based on pulsed LEDs will be more versatile (wide spectral range, no speckle) and cost effective. eeFLIM is also applicable to single or array detectors and TCSPC. Figure 1. Validation of eeFLIM. Mean

normalized integrated signals from images of IRF (scattering from focal plane) and 3 fluorescence dye solutions. The inset highlights the horizontal (temporal) displacements (equal to the lifetimes) of the 4 measured dyes: Rhodamine B (1.6 ns), Coumarin 6 (2.5 ns), Rhodamine 110 (3.8 ns), and dianionic Fluorescein (4.1 ns).

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Assembling the Pieces of Protein Puzzles

*Hummer G *DepartmentofTheoreticalBiophysics,MaxPlanckInstituteofBiophysics,Frankfurtam

Main,Germany

[email protected]

Dynamic supramolecular assemblies are central to many biological functions, from the processing of genetic information to the formation of cellular structures. Structural and functional studies face enormous challenges associated with large size, transient interactions, extensive motions, and partial disorder. To overcome these problems in the characterization of assembly structure and dynamics, we developed a hybrid modeling approach. The combination of ensemble refinement techniques with coarse-grained molecular simulations allows us to integrate data from diverse experiments, including X-ray crystallography, single-molecule fluorescence energy transfer (FRET), small-angle X-ray scattering (SAXS), spin-label distance measurements (EPR/DEER), paramagnetic relaxation enhancement (PRE), nuclear magnetic resonance (NMR), as well as single-particle cryo electron microscopy (cryo-EM). In a Bayesian formulation, we account for uncertainties in the different experiments in a quantitative and transparent manner. The resulting ensemble representations of the dynamic structures of protein supercomplexes balance inputs from a wide range of experiments and from simulation. Applications to the ESCRT machinery involved in membrane protein trafficking and the Atg1 complex in the formation of autophagosomes shed light on the functional principles of the molecular machinery carrying out these complex biological processes.

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Combining Coarse-Grained and All-Atom simulations with smFRET to obtain High-Precision DNA-Protein Structures and Conformational Dynamics

Timothy D. Craggs*.**, Marko Sustarsic*, Majid Mosayebi*, Hendrik Kaju*, Johannes Hohlbein***, Phillip C. Biggin*, Jonathan P.K. Doye*, Achilles N. Kapanidis*

*University of Oxford, UK, **University of Bristol, UK

*** University of Wageningen, The Netherlands [email protected]

We have used single-molecule FRET in combination with all-atom and coarse-grained molecular dynamics simulations to study the structure and conformational dynamics of DNA polymerase I (Pol) substrates both alone and in Pol-DNA complexes.

Using a rigid-body docking approach (with 73 FRET restraints) we determined the structure of the single-nucleotide-gapped DNA-Pol binary complex, which showed a 120° bend in the DNA substrate. This novel structure formed the starting point for all-atom molecular dynamics simulations, revealing 4-5 nt of downstream DNA proximal to the gap are unwound. This melting was verified experimentally using a single-molecule FRET quenching assay.

Coarse-grained simulations on the gapped substrate alone reproduced experimentally determined FRET values with surprising accuracy (<FRET> = -0.0025, across 34 independent distances). The gapped DNA frequently adopted bent conformations as observed in the Pol-bound state (free energy < 4 kT). These conformations were less accessible to nicked (> 7 kT) or duplex (>> 10 kT) DNA, consistent with weaker Pol binding to these substrates. Bent conformations were also more often frayed around the gap, consistent with a mechanism in which Pol recognises a pre-bent, partially-melted conformation of the gapped-DNA. Taken together, these results suggest a mechanism for substrate recognition by Pol in which specificity is encoded through the conformational dynamics of the DNA substrates.

This work exemplifies the power of an iterative approach combining different single-molecule FRET techniques with molecular modelling to yield both high-precision structural information and also detailed molecular dynamics.

Fig. 1 (A) Rigid-body docking of Pol and two duplex DNAs (B) smFRET and coarse-grained modelling of the gapped DNA substrate

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Fast NPS for quantitative structural information from single molecule FRETMichaelis J , Eilert T, Beckers M, Nagy J, and Drechsler F

Ulm University, Institute of Biophysics, Albert-Einstein Allee 11, 89081 Ulm, Germany www.uni-ulm.de/biophys

[email protected]

Single-molecule studies can be used to study biological processes directly and in real- time. In particular, the fluorescence energy transfer between reporter dye molecules attached to specific sites on macromolecular complexes can be used to infer distance information [1,2]. When several measurements are combined, the information can be used to determine the position and conformation of certain domains with respect to the complex. However, data analysis schemes that include all experimental uncertainties are highly complex, and the outcome depends on assumptions about the state of the dye molecules. Here, we present a new analysis algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering termed Fast-NPS [3]. Fast NPS can analyse large smFRET networks in a relatively short time and yields the position of the dye molecules together with their respective uncertainties. Moreover, we show what effects different assumptions about the dye molecules have on the outcome and how to test different models using statistical means. We discuss the possibilities and pitfalls in structure determination based on smFRET using experimental data for an archaeal transcription pre-initiation complex, whose architecture has recently been unravelled by smFRET measurements [4].

Fig. 1. Fast NPS analysis of an archaeal transcription initian complex. Credible volumes are shown at 68% credibility together with x-ray model for polymerase and NPS derived models for the DNA, TFE,

TBP and TFB. Left, analysis was done using the classic NPS model making conservative estimates about the dye molecule prior (static position, fixed cone angle). Right, analysis was done using the

meanposition-iso model, i.e. dynamic averaging over the accessible volume and κ2=2/3, leading to a drastic reduction in the size of the credible volumes. However, statistical tests showed that the

assumptions of the meanposition-iso model are not valid for this smFRET scenario.

[1] A. Muschielok, J.Andrecka, F. Brückner, A. Jawhari, P. Cramer and J. Michaelis “A Nanopositioning system for macromolecular structural analysis”, Nature Methods, 5, 965 (2008). [2] A. Muschielok, and J. Michaelis “Application of the Nano-Positioning System to the Analysis of FRET Networks”, Journal of Physical Chemistry B, 115, 11927 (2011). [3] M. Beckers, F. Drechsler, T. Eilert, J. Nagy and J. Michaelis “Quantitative structural information from single-molecule FRET”, Faraday Discussions, 184, 117 (2015). [4] J. Nagy, D. Grohmann, A.C.M. Cheung, S. Schulz, K. Smollett, F. Werner and J. Michaelis “Complete architecture of the archaeal RNA polymerase open complex from single-molecule FRET and NPS”, Nature Communications, 6, doi: 10.1038/ncomms7161 (2015).

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DISTANCE DISTRIBUTION ANALYIS REVEALS THE DYNAMIC STRUCTURE

OF HSP90

Bjoern Hellenkamp*, Philipp Wortmann*, Florian Kandzia**, Martin Zacharias**,

Thorsten Hugel*

* Institute of physical chemistry, University of Freiburg, Freiburg, Germany** Physics department, Technische Universitaet Muenchen, Garching, Germany

[email protected]

In solution, the protein exists as a dynamic equilibrium of local and global conformations. A comprehensive understanding of protein function requires knowledge on the interrelation between structure and dynamics. We developed a hybrid approach based on single molecule Förster resonance energy transfer (FRET) that integrates intramolecular distance fluctuations together with x-ray structure information and MD simulations. Self-consistent networks of more than 100 distance distributions and time-resolved anisotropies enable a new level of accuracy and confidence. We demonstrate the power of this approach by determining the dynamic structure of the heat shock protein Hsp90. Our resulting mean structure for the closed state resembles the corresponding x-ray structure of yeast Hsp90 with an RMSD of 2.5 Å. Beyond that, we resolved the previously unknown dynamic open structure of this multi-domain protein. Finally, we show how this method can be applied to quantify inter-domain dynamics, dynamics of small elements and protein-protein interactions.

Fig. 1. The highly flexible multi-domain protein Hsp90 (red surface) interacts dynamically with a model client protein (yellow surface).

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MAPPING MOTIONS TO A STATE NECESSARY FOR OLIGOMERIZATION OF A LARGE GTPASE: A JOINT SAXS, NSE, EPR AND FRET STUDY

Thomas-Otavio Peulen1, Carola S. Hengstenberg2, Ralf Biehl3, Mykola Dimura1,5, AlessandroValeri1, Semra Ince2, Tobias Vöpel2, Bela Farago4, Holger Gohlke5, Christian Herrmann2,

Johann Klare6, Andreas Stadler3, Claus A.M. Seidel1.1Chair for Molecular Physical Chemistry, Heinrich Heine University, Düsseldorf, Germany,

2Physical Chemistry I, Ruhr-University Bochum, Bochum, Germany 3Institute of Complex System, Forschungszentrum Jülich, Jülich, Germany

4Institut Laue-Langevin, Grenoble, France 5Institut für Pharmazeutische und Medizinische Chemie, Heinrich Heine University,

Düsseldorf, Germany 6Macromolecular Structure Group, University of Osnabrück, Osnabrück, Germany

[email protected]

By combining double electron-electron resonance (EPR), small angle x-ray scattering (SAXS) and time-resolved fluorescence-measurements (TCSPC) we resolved two distinct conformational states in hGBP1 a large GTPase composed out of three domains, a GTPase-, a middle- and a helical-domain. We find that the C-terminal helix of the helical domain, which is important for oligomerization, has two distinct binding modes on the GTPase domain. By neutron spin echo spectroscopy we show that the correlation time of the conformational change is slower than nanoseconds. By FRET lifetime filtered species cross-correlation (fFCS) we fully characterize the relaxation time distribution of the conformational changes and find that it happens mainly in the microsecond-regime. By multiple FRET-fFCS measurements we determined the internal flexibility of the protein and find that the C-terminal helices a12/a13 are highly flexible relatively to the middle domain. This finding is consistent with the two conformational changes that involve a domain-flip of the middle-domain and the C-terminal helices relatively to the LG-domain and in line with molecular dynamics simulations. We can describe most observations by a two state system. However, the amplitude decay of the relaxation time spectrum suggests that the conformational change happens on a rugged energy landscape. We previously showed that the GTPase- and the C-terminal helices of two hGBP1s associate in presence of GDP-AlFx. If the protein remains in its major state at room temperature an association of the helices a13 is sterically impossible. Hence, we conclude that the minor state at room temperature or a transient state populated during the domain flip is relevant for association of the C-terminal helices a13.

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Parallel evening discussion groups

All abstracts of the parallel sessions are printed under their corresponding number in the poster section of the book of abstracts.

Standards and measurement protocols

Discussion leaders: Thorsten Hugel, Claus Seidel

Narain Karedla A‐P17: Molecular model of the fluorescent protein based FRET sensor with multiple acceptors

Peter Nagy B‐P46: Understanding and reducing the error in the evaluation of intensity‐based microscopic FRET experiments in the presence of low signal‐to‐noise ratio

Probe and Biosensor Development

Discussion leaders: Dorus Gadella Jr., Alexander Savitsky

Jasper van der Velde A‐P35: Enhancing single‐molecule FRET studies with photostabilizer‐dye conjugates

Asko Uri B‐P48: Organic photoluminescent probes possessing triplet –singlet energy transfer by Förster mechanism

Zongwen Jin E‐P80: Multiplexed micro‐RNA assays using time‐resolved FRET with biospectral correction

Nanoparticles

Discussion leaders: Niko Hildebrandt, Igor Medintz

Kateryna Trofymchuk E‐P85: Exploiting fast exciton diffusion in dye‐doped polymer nanoparticles to engineer efficient photoswitching through FRET

Igor Medintz E‐P81: Energy transfer‐based sensitization of luminescent gold nanoclusters

Jurriaan Zwier E‐P88: Time gated FRET microscopy‐ and plate based assays to study G‐Protein Coupled Receptors using bright Eu3+ and Tb3+ donors.

Lauren Field E‐P74: Semiconductor quantum dots as Förster Resonance Energy Transfer donors for intracellularly‐based biosensors

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3D N-‐way FRET Microscopy -‐ Seeing more molecular interactions throughout the living cell

Adam D. Hoppe* and Brandon L. Scott*

*South Dakota State University, Brookings, SD*BioSNTR, Brookings, SD

[email protected]

Förster Resonance Energy Transfer (FRET) is a powerful tool for analyzing protein interactions within living cells. For FRET microscopy to reach its full potential in the analysis of biochemical networks, more than one molecular interaction must be imaged at a time. Additionally, the three-dimensional resolution of FRET microscopy must be improved to capture the true spatial distribution of these interactions. I will describe a new strategy for unmixing overlapping fluorescent spectra based on a linear algebra formalism and empirical calibration that allows estimation of concentrations of bound and free molecules for theoretically any number of FRET partners[1]. With this approach, called N-way FRET, we have compared fluorescent proteins to find an optimal trio for 3-way FRET imaging [2]. We improved the 3D-resolution of N-way FRET via iterative image reconstruction[3]. This approach, 3D N-Way FRET, can be applied to widefield, confocal and linear superresolution microscopies and advances FRET microscopy’s ability to capture the subcellular orchestration of biochemical pathways.

Fig. 1 N-Way FRET can be defined in terms of a spectral superposition on an excitation and emission landscape – shown here for CFP, YFP and RFP. A calibrated linear algebra model is then used to estimate concentrations of bound and free fluorophores. 3D N-Way FRET can reconstruct the 3D distributions of bound and free molecules – shown here as a 3D rendering of HIV CFP-Gag, YFP-Gag and YFP-Gag/RFP-Gag complexes.

1. Hoppe, A.D., et al., N-‐way FRET microscopy of multiple protein-‐protein interactions in live cells. PLoSOne, 2013. 8(6): p. e64760.

2. Scott, B.L. and A.D. Hoppe, Optimizing fluorescent protein trios for 3-‐Way FRET imaging of proteininteractions in living cells. Sci Rep, 2015. 5: p. 10270.

3. Hoppe, A.D., et al., Three-‐dimensional FRET reconstruction microscopy for analysis of dynamicmolecular interactions in live cells. Biophys J, 2008. 95(1): p. 400-‐18.

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Homogenous FRET in Molecular Photonic Wires

Sebastián A. Díaz*, Susan Buckhout-White*, Mario G. Ancona**, Joseph S. Melinger**, and Igor L. Medintz*

* Center for Bio/Molecular Science and Engineering, U.S. Naval Research Laboratory, Washington DC, USA

** Electronic Science and Technology Division, U.S. Naval Research Laboratory, Washington DC, USA

[email protected]

Molecular photonic wires (MPWs) precisely position dyes using structural DNA, exploiting Förster resonance energy transfer (FRET) to direct photonic energy over nm distances.[1] Although versatile, the number of donor-acceptor dye pairs available and the downhill nature of FRET combine to limit the size and efficiency of current MPWs. HomoFRET between identical dyes should provide zero energy loss but at the cost of random transfer directionality. We investigated the use of HomoFRET as a means to extend MPWs. Steady-state-, lifetime-, and fluorescence anisotropy measurements along with computational models were utilized to characterize various 3-, and 5-dye MPW constructs containing from 1-6 HomoFRET repeat sections. Results show that HomoFRET can be extended up to 6 repeat dyes/5 steps with only a ~55% energy transfer efficiency decrease while doubling the longest MPW length to a remarkable 30 nm.[2] Even with non-directionality, the introduction of a repeated-optimized HomoFRET transfer dye is preferable compared to additional less efficient dye species. HomoFRET further provides the benefit of having a higher energy output. The extended MPWs may be exploited in applications such as light harvesting, biosensing, and molecular electronics.

Fig. 1. (Left) Schematic of MPWs. The colored circles represent the dyes, while each DNA strand is presented by a different color. Arrows represent possible energy transfer steps and their directionality. (Right) End-to-end transfer efficiency of the 5-dye MPW as a function of the length of the construct.

[1] Heilemann, M., Kasper, R., Tinnefeld, P., Sauer M., “Dissecting and reducing the heterogeneity of excited-state energy transport in DNA-based photonic wires”, J. Am. Chem. Soc., 133, 4193, (2011). [2] Díaz, S., Buckhout-White, S., Ancona, M. et al. “Extending DNA-Based Molecular Photonic Wires with Homogeneous Förster Resonance Energy Transfer”, Adv. Opt. Mat., (2016) DOI: 10.1002/adom.201500554 .

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farFRET: Extending the Range in Single-Molecule FRET Experiments beyond 10nm Georg Krainer*,**, Andreas Hartmann* and Michael Schlierf*

* B CUBE – Center for Molecular Bioengineering, TU Dresden, 01307 Dresden** Molecular Biophysics, University of Kaiserslautern, 67663 Kaiserslautern

[email protected]

Single-molecule Förster resonance energy transfer (smFRET) has become a powerful nanoscopic tool in studies of biomolecular structures and nanoscale objects; however, conventional smFRET measurements are generally blind to distances above 10 nm thus impeding the study of long-distance phenomena. Here, we report the development of farFRET, a technique that extends the range in single-molecule FRET (smFRET) measurements beyond the 10 nm line by enhanced energy transfer using multiple acceptors. We demonstrate that farFRET can be readily employed to quantify FRET efficiencies and conformational dynamics using double-stranded DNA molecules, RecA-filament formation on single-stranded DNA and Holliday junction dynamics. farFRET allows quantitative measurements of large biomolecular complexes and nanostructures thus bridging the remaining gap to superresolution microscopy [1].

Fig. 1. Artistic illustration of farFRET. The technique uses the enhancement in FRET from multiple acceptors to break the 10 nm limit in smFRET experiments. Donor excitation energy is transferred to a

bundle of four acceptors on double-stranded DNA. The presence of multiple acceptors increases the probability of energy transfer and thereby amplifies the observed FRET efficiency, enabling the

observation of long-distance phenomena on individual molecules.

[1] Krainer G., Hartmann A., Schlierf M. “farFRET: Extending the Range in Single-Molecule FRET Experiments beyond 10 nm”, Nano Letters, 15, 5826-5829, (2015).

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Testing Biomolecular Coordination: Quantitative Analysis of Single-Triad FRET Data Anders Barth*, Lena Voith von Voithenberg*, Don C. Lamb1

* Chemistry Department, NIM, CIPSM, CeNS, Ludwig-Maximillians-Universtität Mü[email protected]

With the development of single-molecule spectroscopy, it has become possible to measure the conformation and dynamics of biomolecules on the single-molecule level. Single-pair FRET (spFRET) experiments give unprecedented insights into how biomolecules function. However, spFRET experiments are limited to measuring one distance at a time. Three distances can be measured simultaneously in a single molecule by adding a third fluorophore and performing single triad FRET (stFRET), which was introduced over 10 years ago. A quantitative analysis of 3cFRET burst analysis data has many challenges that make a quantitative analysis of the experiments difficult. To deal with the broad FRET distributions and the inherent correlation of counts in three-color experiments, we have developed a three-color Photon-Distribution Analysis (3cPDA) based on the corresponding method for spFRET [1]. With 3cPDA, shot noise and the distribution of FRET efficiencies are incorporated into the analysis and made it possible to detect the presence of coordinated motions within biomolecules. In this presentation, the method of stFRET and 3cPDA will be introduced. The reliability of the newly developed analysis approach was first explored using simulated data (Figure). As a second test, stFRET data was collected on double-stranded DNA where fluorophores were attached at known positions and the distances between the fluorophores determined. We then applied stFRET and 3cPDA to investigate the presence of coordinated motions in the Heat-Shock Protein 70, a chaperone protein that helps nascent proteins fold and reach their final destination within a cell. The Hsp70 is constructed of two domains, a substrate binding domain (SBD) and the nucleotide-binding domain (NBD). Using non-natural amino acids and cysteine mutations, we were able to label Hsp70 with three fluorophores and investigate the conformation of the lid in the SBD as well as the interdomain distance. Coordinated motions were observed for the endoplasmic reticulum Hsp70, BiP, but not in the mitochondrial Hsp70.

Fig. Single triad FRET. (A) Inset. A three-color FRET system was simulated for the given geometry using a Förster Radius of 50 Å. The BG stFRET histogram (A) and BR stFRET histograms (B) are shown. The FRET efficiencies distributions are broad with values below zero and above one for the BR FRET efficiency histogram. (C) A two-dimensional plot demonstrating the intrinsic correlation

observed between the two FRET efficiencies. Data is shown as grey bars in A/B and as a surface plot in C. Fits are shown as black lines. The observed shot-noise broadening can be well described by 3C-

PDA. In C, the surface is colored according to the weighted residuals as shown in the legend.

[1] M. Antonik, S. Felekyan, A. Gaiduk, C.A.M. Seidel “Separating structural heterogeneities from stochastic variations in fluorescence resonance energy transfer distributions via photon distribution analysis”, J Phys Chem B 110, 6970 (2006).

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Deciphering CaMKII multimerization using holoenzyme assembly mutants, FCS, and concurrent homo- and hetero-FRET analysis.

Pabak Sarkar*, Jithesh V. Veetil**, Kaitlin Davis***, Henry L. Puhl III*, Tuan A. Nguyen* & Steven S. Vogel*

* Laboratory of Molecular Physiology, National Institute on Alcohol Abuse and Alcoholism,National Institutes of Health, Maryland, USA ** Foundation for Advanced Education in the Sciences, National Institutes of Health, Maryland, USA *** University of Maryland, Maryland, USA

Contact: [email protected]

Abstract

Monitoring changes in molecular conformation is essential for studying protein interactions within and between complexes. Current methods such as FRET or FCS can only reveal limited aspects of these changes, which in many cases is insufficient for interpretation. Fluorescent Polarization and Fluctuation Analysis (FPFA), a time-correlated single-photon counting technique that combines homo-FRET and FCS was developed to address this problem. In FPFA, changes in mass and/or shape of a protein complex, the number of fluorescent subunits per complex, as well as subunit proximity (1 – 10 nm), are simultaneously detected. Here we have extended FPFA by incorporating simultaneous hetero-FRET analysis so that additional structural changes can be monitored concurrently. This multi-modal approach was then used to observe changes in CaMKII holoenzyme in response to activation and subsequent interactions with NR2B. Revised FPFA revealed that catalytic-domain pairing is the fundamental structural motif of the holoenzyme organization. A genetically engineered dimeric-CaMKII functioned like native holoenzyme (8-14 subunits) in terms of activity, T286 autophosphorylation, structural changes triggered by calcium-calmodulin, and those associated with interaction with NR2B.

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On new classes of genetically-encoded fluorescent proteins optimized for fluorescence polarisation and FRET

Gerard Marriott Department of Bioengineering, University of California, Berkeley, CA 94720

[email protected]

1,8-Dimethylaminonaphthalenesulphonyl chloride (Dansyl-Cl) was introduced by Gregorio Weber in the late 1940’s as a probe for fluorescence polarisation (anisotropy; r or FA) based analysis of protein hydrodynamics. The rotational correlation time (τc) for a spherical protein increases by 1ns for every ~2500 Dalton increase in mass, ie ~12ns for a 30kD protein. According to the Perrin-Weber equation [ro / r = (1 + τf / τc )], the rotational properties of a 30kD protein are best determined when τc = τf , ie FA = ~0.2, or half the limiting value (ro). The τf for Dansyl is 12~14ns, which makes it suitable to study the tumbling motions of proteins from ~10kD~100kD. In spite of almost 70 years of FA research, 1,8-Dansyl is still the best performing probe to quantify protein hydrodynamics. CFP and GFP are unsuitable for FA-sensing of protein hydrodynamics because of their short excited state lifetime, ~2.2ns, and large mass (~30kD) with unbound CFP fusion proteins having FA values close to ro. Recently we introduced new genetically-encoded fluorescent proteins (LUMP and fLov2 ) whose long lifetimes (5ns~14ns) and small mass (~10-20kD) make them suitable probes for FA-based analysis of protein hydrodynamics. These features allow us to append capture groups as large as ~30kD onto LUMP or fLov2 and still generate large differences in FA between their free and bound states.

Our new FA-probes offer significant advantages over FRET-probes in the design and performance of sensors for new target molecules. First, the binding of a known target molecule to a LUMP- or fLov2 FA-sensor results in a predictable change in FA value, with the unbound sensor tumbling more rapidly (low FA) compared to the larger target-bound sensor (high FA); Second, a single FA-measurement is sufficient to calculate the amounts of both the free- and target-bound states of the FA-sensor in the sample; Third, LUMP and fLov2 function as universal sensors of protein hydrodynamics ie their FA-value changes in a direct and predictable fashion with their mass. This latter feature has allowed us to implement a simple and universal strategy in the design of FA-sensors for new target molecules, where a capture sequence specific for a larger target molecule is appended to LUMP or fLov2. The FA value of the FA-sensor increases predictably on binding to the target molecule and unlike FRET sensors based on CFP-YFP the capture group does not need to undergo a target-induced conformational change. Finally, we will show how the smaller mass and surface location of the fluorescent probe in LUMP and fLov2 improves their performance as donor probes in FRET with YFP and mRuby, compared to CFP-YFP and GFP-mRuby respectively. In particular we have measured FRET efficiencies for LUMP-YFP and fLov2-mRuby2 connected by “zero-length” linkers that are up to 2-fold higher than the equivalent VFP-YFP and GFP-mRuby2 fusion proteins.

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Genetically Targeted and Activated Fluorogenic FRET-based IndicatorsBrigitte F. Schmidt*, Matharishwan Naganbabu*,**, Lydia A. Perkins*,*** and Marcel P.

Bruchez*,**,*** * Molecular Biosensor and Imaging Center, Carnegie Mellon University, Pittsburgh, PA

** Department of Chemistry, Carnegie Mellon University, Pittsburgh, PA *** Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, PA

[email protected]

Fluorogen activating proteins that activate the fluorescence of triarylmethane dyes have been demonstrated as practical tags for both cell surface and intracellular labelling, with applications ranging from single-molecule imaging to whole-animal optogenetics. The binding of a fluorogenic dye can result in thousands-fold activation, serving as a binding-mediated optical switch, which activates fluorescence from otherwise dark molecules. Synthesis of various fluorescent donors linked to a far-red excitable fluorogen at distances far shorter than the Forster radius of the dyes has established a new family of FRET-based multi-excitation fluorogenic dyes, with tunable excitation and emission properties suitable for use with a wide variety of conventional and superresolution microscopy methods. Use of environmentally sensitive donor dyes produces targeted and activated ratiometric fluorescent indicators, enabling optical physiology at and beyond the diffraction limit. I will discuss applications of pH sensors in living cells for measurement of endolysosomal trafficking and development and validation of K+ and ROS sensor dyes for use in live cells and model organisms.

Fig. 1. Stimulated Emission Depletion Microscopy of Ratiometric pH Fluorogen Probes on Endosomes in Live Cultured Cells.

STED

STED

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Peptide molecular dynamic beacon for FRET-sensors

Savitsky A.P.*, Goryashchenko A.S.*, Khrenova M.G.**, Zherdeva V.V.*, Ivashina T.V.***

* A.N. Bach Institute of Biochemistry, Research Center of Biotechnology of the RussianAcademy of Sciences, Moscow, Russia

** M.V. Lomonosov Moscow State University, Department of Chemistry, Moscow, Russia *** Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy

of Sciences, Pushchino, Moscow Region, Russia [email protected]

The use of the novel peptide linker optimizes the conformation of the sensor to the beacon-like shape due to the hydrophobic interactions and thereby adjust donor to acceptor and increases the FRET efficiency. We introducing new structural elements so that novel linker dominantly exist in a loop conformation with C and N termini close to each other and DEVD motif recognized by caspase-3 is exposed to the solution for its higher accessibility (peptide molecular beacon conformation). Novel linker has two hydrophobic regions that tend to interact with each other rather than be exposed to solution according to molecular dynamic simulation. Prolines before and after DEVD motif provide for the turns of the loop. We develop the TR-M4-K peptide beacon-like sensor based on red fluorescent proteins TagRFP and KFP. TR-M4-K sensor is successfully hydrolyzed by caspase-3 in vitro and in tumor cell line Hep2. Detection of the hydrolysis can be carried out by a change in the fluorescence intensity, or by a change in the fluorescence lifetime of the donor. We propose a new method for estimating the efficiency of hydrolysis of FRET-sensors. Primary evaluation is carried out by change of the FRET efficiency calculated from average values of fluorescence lifetime. After that, it is possible to get a quantitative evaluation by the change of the amplitudes of fluorescence decay components. Sensor TR-M4-K has a dynamic range of measurements equal to 4.56 in the case of measurements of the ratio of amplitudes of the components corresponding to closed and open conformations of sensor, which exceeds the results previously described in the literature. In addition to photo physical advances novel sensor have also genetic advances while possessing more compact size, which reduces the genetic load at the step of transfection. This work was supported by Russian Science Foundation, grant № 15-14-30019. M.K. acknowledges the use of supercomputer resources of the M.V. Lomonosov Moscow State University and of the Joint Supercomputer Center of the Russian Academy of Sciences.

Fig. 1. Peptide molecular dynamic beacon. Hydrophobic amino acids tend to interact with each other rather then solvent

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Decoding Molecular Plasticity in the Dark Proteome

Edward A Lemke

Structural and Computational Biology Unit, Cell Biology and Biophysics Unit

EMBL, 68161 Heidelberg, Germany

E-mail: [email protected]

The mechanisms by which intrinsically disordered proteins (IDPs) engage in rapid and highly selective binding is a subject of considerable interest and represents a central paradigm to nuclear pore complex (NPC) function, where nuclear transport receptors (NTRs) move through the NPC by binding disordered phenylalanine-glycine-rich nucleoporins (FG-Nups). In the first part of my talk, I will present a combined single molecule and ensemble spectroscopy approach that paired with atomic simulations revealed that a rapidly fluctuating FG-Nup populates an ensemble of conformations that are prone to bind NTRs with diffusion-limited on-rates. This is achieved using multiple, minimalistic, low affinity binding motifs that are in rapid exchange when engaging with the NTR, allowing the FG-Nup to maintain an unexpectedly high plasticity in its bound state. Since site-specific labeling of proteins with small but highly photostable fluorescent dyes inside cells remains the major bottleneck for directly performing such high resolution studies in the interior of the cell, I will demonstrate an approach how to overcome this limitation in the second part of my talk. We have now developed a semi-synthetic strategy based on novel artificial amino acids that are easily and site-specifically introduced into any protein by the natural machinery of the living cell. Expressed proteins only differ from their natural counterparts by very few atoms, constituting a ring-strained cyclooctyne or cyclooctene functional group. This allowed rapid, specific “click” labeling and even multi-color studies of living cells and subsequent super resolution microscopy.

References:

Milles S, Mercadante D, Aramburu IV, Jensen MR, Banterle B, Koehler C, Tyagi S, Clarke J, Shammas S, Blackledge M, Gräter F, Lemke EA. Plasticity of an ultrafast interaction between nucleoporin and transport receptors, Cell. 2015 Oct 22;163(3):734‐45. doi: 10.1016/j.cell.2015.09.047. Epub 2015 Oct 8.

Nikic I, Plass T, Schraidt O, Szymanski J, Briggs JA, Schultz C, Lemke EA*. Dual‐color super‐resolution microscopy via genetic code expansion technology and tuned click reactions. Angew Chem Int Ed Engl, 2014, Feb, 53(8), 2245‐2249 (2014)

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Concentric FRET with Quantum Dots: Energy Transfer Pathways and Application to Multiplexed Biological Sensing

W. Russ Algar * * Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver,

British Columbia, V6T 1Z1, Canada. [email protected]

For more than a decade, quantum dots (QDs) have been utilized as superior FRET donors [1]. This superiority arises not only from their unique photoluminescence (PL) properties, which can be tuned to match the properties of an acceptor, but also from their surface area, which permits the assembly of the QD with multiple copies of that acceptor. Building on this foundation, we have developed QD-based concentric FRET (cFRET) configurations where multiple copies of two or more different acceptors are co-assembled per QD (see Fig. 1).

Fig. 1. An example of concentric FRET imaging for multiplexed protease assays [3].

Our prototypical cFRET configuration is a QD assembled with peptides labelled with Alexa Fluor 555 (A555) and Alex Fluor A647 (A647), where energy transfer occurs from the QD to both dyes and between the two dyes. Sequential and competitive FRET in this configuration [2] will be discussed along with examples of its application in biological sensing, including multiplexed imaging of protease activity [3] and two-pronged detection of protease activity and concentration [4]. The A555/QD and A647/QD PL emission ratios provide a unique combination of signals that permits multiplexed analysis. Other cFRET configurations have also been developed and will be compared with the prototypical configuration. Two examples are (i) a multi-donor cFRET configuration that can measure protease activity through a PL excitation ratio (cf. emission ratio) [5], and (ii) a long-wavelength cFRET configuration that combines QD600 with Alexa Fluor 633 (A633) and Alexa Fluor 680 (A680) dyes for multiplexed DNA hybridization assays. The different properties of A633 and A680 versus A555 and A647 results in a different balance between competitive and sequential FRET, and a different set of benefits and liabilities. The success of this initial research promises exciting future opportunities for biological sensing and imaging with cFRET probes.

[1] I.L. Medintz, H. Mattoussi. “Quantum dot-based resonance energy transfer and its growing application in biology.” Phys. Chem. Chem. Phys., 11, 177 (2009). [2] M. Wu, M. Massey, E. Petryayeva, W.R. Algar. “Energy Transfer Pathways in a Quantum Dot-Based Concentric FRET Configuration.” J. Phys. Chem. C., 119, 26183 (2015). [3] M. Wu, W.R. Algar. “Concentric Förster Resonance Energy Transfer Imaging.” Anal Chem. 87, 8078 (2015). [4] M. Wu, E. Petryayeva, W.R. Algar. “Quantum Dot-Based Concentric FRET Configuration for the Parallel Detection of Protease Activity and Concentration.” Anal. Chem, 86, 11181 (2014). [5] H. Kim, C.Y.W. Ng, W.R. Algar. “Quantum Dot-Based Multidonor Concentric FRET System and Its Application to Biosensing Using an Excitation Ratio.” Langmuir, 30, 5676 (2014).

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International Discussion Meeting “Förster Resonance Energy transfer in Life Sciences 2" (FRET 2)


Abstracts of poster presentations (for each topic in alphabetical order of presenting author)

Topics:

A Single-molecule and ensemble in vitro FRET studies of Proteins, Nucleic Acids and Membranes; FRET spectroscopy

B FRET Theory, Analysis and Instrumentation

C FRET microscopy

D Hybrid Approaches: FRET combined with Modelling and other Spectroscopies

E Biosensors, Probes and Nanoparticles

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Probing the folding dynamics of human telomeric G-quadruplex with single-molecule FRET

a,b, Victoria Birkedala,c, Birgit Schiøtta,b, Sofie Noera,c, Siri Søndergaarda,bMikayel Aznauryan

a Interdisciplinary Nanoscience Center (iNANO) and b Centre for DNA Nanotechnology (CDNA), Aarhus University, Gustav Wieds Vej 14, 8000 Aarhus, Denmark; c Department of

Chemistry, Aarhus University, Langelandsgade 140, 8000 Aarhus, Denmark

[email protected]

Guanine-rich sequences consisting of several tandem repeats of TTAGGG are abundant in human telomeric DNA. They have been shown to fold into unique secondary structures under physiological conditions. One of these is the G-quadruplex structure that can undergo complex folding pathways [1-3]. Single-molecule fluorescence microscopy combined with Förster resonance energy transfer (FRET) allows probing the conformation and dynamics of individual molecules without time and population averaging and contributed to uncover the large conformational diversity of G-quadruplexes [4].

Here we utilize single-molecule FRET microscopy to probe the K+-induced folding dynamics of human telomeric G-quadruplex DNA. Our single-molecule experiments identify several transient folded states formed at the early stages of the folding process. These states are in dynamic interchange with each other and with the unfolded state and, in the course of several hours, converge into long-lived folded conformation. A combination of experimental results and MD simulations provides a comprehensive structural description of the folded conformations observed along the folding of G-quadruplexes. Our work thus offers a novel view of the process and timescales of folding of human telomeric G4 structures in the presence of K+.

[1] Bochman, M. L., Paeschke, K., Zakian, V. A. “DNA secondary structures: stability and function of G-quadruplex structures”, Nature Reviews Genetics, 13, 770, (2012). [2] Biffi, G., Tannahill, D.; McCafferty, J.; Balasubramanian, S. “Quantitative visualization of DNA G-quadruplex structures in human cells”, Nature Chemistry, 5, 182, (2013). [3] Gray, R. D., Trent, J. O., Chaires, J. B. “Folding and unfolding pathways of the human telomeric G-quadruplex”, J. Mol. Biol., 426, 1629, (2014). [4] Lee, J. Y. Okumus, B., Kim, D. S., Ha, T. J., “Extreme conformational diversity in human telomeric DNA”, Proc. Natl. Acad. Sci. U.S.A., 102, 18938, (2005).

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3D Light microscopy of protein structure with Angstrom resolution

Siegfried Weisenburger1, Daniel Boening1, Benjamin Schomburg2, Stefan Becker2, Christian Griesinger2, and Vahid Sandoghdar1

1Max Planck Institute for the Science of Light and Department of Physics, University of Erlangen-Nuremberg, 91058 Erlangen, Germany2Max Planck Institute for Biophysical Chemistry, 37077 Goettingen, Germany

The significance of super-resolution microscopy beyond the diffraction barrier was honored by the Nobel Prize in Chemistry last year. One such popular method employs pinpointing the position of single fluorophores, whereby the center of the point-spread function can be determined with arbitrary localization precision depending on the available signal-to-noise ratio. At room temperature, the signal of a fluorophore is limited by photobleaching resulting in typical localization precisions on the order of ten nanometers. We have already demonstrated Angstrom localization precision made possible by the substantial enhancement of the molecular photostability at cryogenic temperatures [1]. We also verified the feasibility of colocalization and cryogenic distance measurements by resolving two fluorophores on the backbone of a double-stranded DNA at nanometer separation [2].

Here, we present our results on resolving the positions of multiple fluorophores attached to proteins using cryogenic colocalization microscopy [3]. By applying algorithms borrowed from cryogenic electron microscopy, we can reconstruct a three-dimensional density map for the positions of the fluorescent labels with a resolution of several Angstrom, yielding excellent agreement with the expected crystal structure (Figure 1). Our technique pushes optical resolution by nearly two orders of magnitude beyond the state-of-the-art conventional super-resolution microscopy. It allows us to gain structural information that might not be accessible via existing analytical methods such as x-ray scattering or magnetic resonance spectroscopy. We discuss the potential of the technique for further improvement and its important challenges.

Figure1: 3D reconstruction of the streptavidin-biotin binding sites. Overlay of the reconstructed fluorophore positions (red) with the crystal structure of streptavidin.

[1] S. Weisenburger, B. Jing, A. Renn, and V. Sandoghdar, Proc. SPIE 8815, 88150D (2013).

[2] S. Weisenburger, B. Jing, D. Hänni, L. Reymond, B. Schuler, A. Renn, and V. Sandoghdar, ChemPhysChem 15, 763–770 (2014).

[3] S. Weisenburger, D. Boening, L. Wei, and V. Sandoghdar, in preparation.

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Conformational mechanisms of homing-to-target in protein-DNA binding and characterization of binding partners with different affinities

Milagros Castellanos1, Silvia Zorrilla2, Víctor Muñoz1

1 Centro Nacional de Biotecnología (CNB-CSIC) and 2 Centro de Investigaciones Biológicas(CIB-CSIC). Madrid, Spain. [email protected]

Proteins that bind DNA specifically need to find a short target sequence within the enormous pool of binding sites provided by genomic DNA. Before reaching their targets, DNA-binding proteins encounter nonspecific DNA first and bind to it, although with weaker affinity than DNA specific sequence [1]. As observed first in 1970, DNA binding proteins binds its target ~100 times faster than allowed by the 3D diffusion limit [2]. This faster-than-diffusion binding was explained by the facilitated-diffusion model, in which a DNA-binding protein interacts with nonspecific DNA before reaching its target [3-5] by three main mechanism: sliding, where protein slides in 1D (keeping contact) along nonspecific DNA; hopping, where the protein dissociates from DNA briefly, performing free 3D diffusion, and lands back on DNA at other location, shorter that the DNA persistence length; and jumping, where the protein’s DNA landing location is not correlated to the dissociation site [6]. Facilitate diffusion has been studied theoretically, computationally and experimentally using single molecule methods. However, very little is known from the protein viewpoint. We are using the DNA-binding engrailed homeodomain (engHD), a ~60-residue helix bundle module that binds with extremely high affinity to DNA sequence TAATTA/G [7]. Our results, obtained mainly by fluorescence spectroscopy methods, indicate that DNA-protein binding happens at the pM-nM range, increasing conformational stability of the protein and that nonspecific binding also takes place and seems dependent on DNA length.

[1] Revzin A (1990) The biology of nonspecific DNA protein interactions. CRC Press, London. [2] Riggs AD, Bougeois S, Cohn M (1970) The lac repressor-operator interaction. 3. Kinetic studies. J Mol Biol 53:401–417. [3] Adam G, Delbruck M (1968) Reduction of dimensionality in biological diffusion process. In: Rich A, Davidson N (eds) Structural chemistry in molecular biology, Freeman, San Francisco, pp. 198–215. [4] Winter RB, von Hippel PH (1981) Diffusion-driven mechanisms of protein translocation on nucleic acids. 2. The Escherichia coli repressor-operator interaction: equilibrium measurements. Biochemistry 20:6948–6960. [5] Gowers DM, Halford SE (2003) Protein motion from non-specific to specific DNA by threedimensional routes aided by supercoiling. Embo J 22:1410–1418. [6] Halford SE, Marko JF (2004) How do site-specific DNA-binding proteins find their targets? Nucleic Acids Res 32:3040–3052. [7] Ades SE, Sauer RT (1995) Specificity of minor-groove and major-groove interactions in a homeodomain-DNA complex. Biochemistry 34(44):14601-8.

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Unravelling the domain contributions to the folding mechanism of a multi-domain protein.

Cerminara M *, Schöne A*, Kempe D**, Gabba M***, Züchner T and Fitter J*,**

*Forschungszentrum Jülich GmbH, Institute of Complex Systems ICS-5 MolecularBiophysics, Jülich, Germany

** Physikalisches Institut (IA), AG Biophysik, RWTH Aachen University, Aachen, Germany

*** Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Groningen, Netherlands

[email protected]

Deciphering how the amino acid code translates into 3D structures is the key to understand how proteins really work. In this context, the two-domain protein phosphoglycerate kinase (PGK) has proven to be an excellent model for multi-domain proteins. It is known that both domains of PGK interact during folding. The N-terminal domain only gains its native structure in presence of the C-terminal domain. The C-domain is able to fold individually but the process is facilitated by the N-domain. In addition, intermediate states are involved. [1,2] A detailed picture of these intermediates and to what extent they are populated is still missing. It is therefore challenging to unravel the mechanisms of tertiary structure formation, especially since subpopulations are hard to identify in ensemble methods. To avoid averaging over all conformations, single-molecule methods are a perfect tool to distinguish and quantify such subpopulations. We established a set of PGK cysteine variants for site-specific labeling with fluorescent dyes for single molecule fluorescence resonance energy transfer (FRET). We verified that secondary and tertiary structures were not affected by cysteine mutations applying circular dichroism (CD) spectroscopy and tryptophan fluorescence. In addition all PGK cysteine mutants were catalytically active. The native states of the double labelled PGK variants were thoroughly characterized by fluorescence correlation spectroscopy (FCS) and single molecule FRET. Our System is designed to follow motions in between and within the individual domains displayed by distance changes of fluorophores during unfolding/folding transitions under denaturing conditions.

[1] S. Osváth, J. J. Sabelko, M. Gruebele “Tuning the Heterogeneous Early Folding Dynamics of Phosphoglycerate Kinase”, J Mol Biol, 333, 187, (2003).

[2] J.-H. Han, S. Batey, A. A. Nickson, S. A. Teichmann, J. Clarke, “The folding and evolution of multidomain proteins”, Nat Rev Mol Cell Bio 8, 319 (2007).

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Asymmetric conformational states of the ribosome recycling factor ABCE1

probed by single-molecule FRET

Kristin Kiosze-Becker*, Giorgos Guoridis** , Marijn de Boer** , Bianca Hetzert*, Thorben Cordes** , Robert Tampé*

*Institute of Biochemistry, Biocenter, Goethe University Frankfurt, Max-von-Laue-Str.9, D-60438 Frankfurt a.M., Germany **Molecular Microscopy Research Group and Single-molecule Biophysics, Zernike Institute for Advanced Material, University of Groningen, The Netherlands.

[email protected]

It is hypothesized that the ABC-type ATPase ABCE1 uses two conformational states for its mechanochemical work that includes splitting of posttranslational complexes, i.e., ribosome recycling. This essential process allows for processive protein synthesis.The transition between the conformational states is driven by binding and hydrolysis of ATP and is accomplished in two independent and structurally distinct ATP binding sites. These sites are formed between two head-to-tail NBDs (Nucleotide Binding Domains) linked by a hinge. Additionally, ABCE1 contains an iron-sulfur-cluster domain (FeS) which performs mechanical displacement of the ribosomal subunits. The mechanism by which the conformational transitions in the two ATP sites is regulated remains elusive. In this study, we characterise the conformational states and dynamics of ABCE1 in its two ATP binding sites using single-molecule FRET. The conformational states were probed by measuring FRET efficiency between a different donor- and acceptor pairs attached at strategic positions in the two ATP binding sites. Solution-based smFRET experiments with a confocal microscope allow to probe conformational states in the two ATP sites independently and at the same time monitor the interaction of ABCE1 with the ribosome by analysing the diffusion properties of the molecules while transiting through the confocal volume. For the first time we have shown that each ATP site of ABCE1 transits between three conformational states, instead of two. ABCE1 is found to be inherently in dynamic equilibrium between different conformational states in the two ATP binding sites, instead of being frozen in an open state. Binding interactions with nucleotides and the ribosome alter these equilibria in a different way and on a different timescales. Moreover, the FeS-cluster domain is crucial not only for function as it affects both the inherent and ligand regulated equilibria between the conformational states. Surprisingly, nucleotide binding, rather than ribosome binding, is involved in the rate limiting steps towards equilibria. From our experiments the complexity of ABCE1 emerged (being a structurally relatively simple protein) able to acquire up to 18 different states. This complexity might explain the enormous functional diversity of the ABCE1.

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ACCURATE DETERMINATION OF THE RNA JUNCTIONS VIA SINGLE-MOLECULE HIGH-PRECISION FRET MEASUREMENTS

Olga Doroshenko*, Hayk Vardanyan*, and Sascha Fröbel*, Stanislav Kalinin*, Simon Sindbert*, Oleg Opanasyuk*, Christian Hanke**, Sabine Müller***, Holger Gohlke**, Claus

A. M. Seidel*

* Chair of Molecular Chemistry, Heinrich-Heine-Universität, Universitätsstraße 1, Geb26.32,40225 Düsseldorf, Germany

** Institute of Pharmaceutical and Medicinal Chemistry, Heinrich-Heine-University, Universitätsstr. 1, 40225 Düsseldorf, Germany

*** Institute for Biochemistry, 3Ernst-Moritz-Arndt-Universität Greifswald, Felix-Hausdorff-Straße 4, 17487, Greifswald, Germany

[email protected]

Förster-Resonance-Energy-Transfer (FRET) restrained high-precision structural modeling is a powerful tool for analyzing the biomolecular structure. We apply multi-parameter fluorescence detection (MFD) of single molecules and ensemble Time-Correlated Single Photon Counting measurements (eTCSPC) to perform FRET study on RNA three- and four-way-junctions (4WJs and 3WJs) which are derived from the hairpin ribozyme. Overall 283 FRET pairs were measured with single-molecule MFD and analyzed with the analysis toolkit [1] that includes probability distribution analysis (PDA) for FRET distance determination and FRET position and screening (FPS) toolkit for structural model generation. In order to study the influence of the junction on the RNA structure we studied the functional junction part with prolonged helices. Bulge and sequence variations were considered as dominant factors influencing junction conformations for RNA 3WJ. Six different RNA 3WJ with different sequences were studied, two of which have two and five unpaired nucleotides in the junction region. We found three different conformers for RNA 4WJ. However RNAs 3WJ have only one predominant conformer. Furthermore we report that bulges in the junction region determine orientation and rotation of helices, inducing coaxial stacking. Noteworthy the stacked helices are different for the 3WJs with different bulges. Our results show that small changes in the sequence make dramatic changes in RNA 3WJ tertiary structures which are expected to have significant impact on the functionality.

[1] Kalinin, S. et al, “A toolkit and benchmark study for FRET-restrained high-precision structural modeling”, Nature Methods, 9, 1218–1225 (2012)

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Point mutations reveal specific intradomain interactions essential for group II intron ribozyme folding

l*and Roland K. O. Sige ***Danny Kowerko, **Lucia Cardo, *Richard Börner, *Erica Fiorini *Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich,

Switzerland; **Institute of Biomedical Research, University of Birmingham, Birmingham, B15 2TQ, UK; ***Department of Computer Science, Chemnitz University of Technology,

Strasse der Nationen 62, 09111 Chemnitz, Germany.

[email protected]

Group II introns are among the largest ribozymes known. Their structural analysis suggests that they have evolved into ribonucleoproteins generating the eukaryotic nuclear spliceosome. They are found in the genome of bacteria, plants, and lower eukaryotes [1]. These self-splicing ribozymes are active upon formation of specific long-range tertiary interactions that define a precise conformation influenced by co-factors such as Mg2+ [2, 3]. We study the folding pathway of an engineered but active Sc.ai5γ group II intron variant which allows to study conformation rearrangement trough single molecule Fluorescent Resonance Energy Transfer (smFRET) (Figure, left) [3,4]. Moreover, point mutations and/or domain deletion were inserted in specific sequence essential for inter-domain docking. Combining bulk activity assays and single molecule experiments we test the effect of these mutations on the catalytic activity and the folding pathway of this ribozyme. From smFRET experiments we discovered the presence of four reoccurring FRET states, which correspond to four different structural conformations [3, 4]. This technique also allowed us to quantify the differences in the relative population of each conformation; especially a clear shift towards the unfolded conformations when the catalytic domain (D5) is mutated was observed (Figure, right) [5]. Although even a drastic mutation still allows for folding into the most compact state, addressed to be the active one, no catalytic activity is present if the interaction which involves the catalytic core formation is disturbed (Figure, centre). Even introducing high Mg2+ concentration and crowding environment, the catalytic activity cannot be restored [4]. From the obtained results we can assign a change in conformation/FRET state to a particular event in the folding pathway. Moreover, we obtain a deeper understanding of the role of the single domains in the folding into the native state and the stabilization of this conformation.

[1] A.M. Pyle, “The tertiary structure of group II introns: implications for biological function and evolution”, Crit. Rev. Biochem. Mol. Biol., 2010, 45, 215-32. [2] R.K.O. Sigel, “Group II intron ribozymes and metal ions - A delicate relationship”, Eur. J. Inorg. Chem., 12, 2281-92, (2005). [3] M. Steiner, K. Karunatilaka, R.K.O. Sigel, D. Rueda, “Single-molecule studies of group II intron ribozymes”, Proc. Natl. Acad. Sci. USA, 105, 13853–58, (2008). [4] E. Fiorini, R. Börner, R.K.O. Sigel, “Mimicking the in vivo environment – The effect of crowding on RNA and biomacromolecular folding”, Chimia, 69, 207-12, (2015). [5] S.L.B. König, M.C.A.S. Hadzic, E. Fiorini, R. Börner, D. Kowerko, W.U. Blanckenhorn, R.K.O. Sigel, “BOBA FRET: bootstrap-based analysis of single-molecule FRET data”, PLoS ONE, 8:e84157, (2013).

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65

Folding Thermodynamics and Kinetics of the Outer Membrane Phospholipase A

investigated by Single-Molecule FRET Spectroscopy

Pablo Gracia*, Georg Krainer*, **, Andreas Hartmann*, Sandro Keller**, Michael Schlierf*

*B CUBE Center for Molecular Bioengineering, Technische Universität Dresden, Germany ** Molecular Biophysics, Technische Universität Kaiserslautern, Germany

[email protected]

[email protected]

[email protected]

Membrane protein folding is a major challenge in biology. The molecular processes by which an unfolded polypeptide chain assumes its three-dimensional native structure and inserts into the complex, anisotropic milieu of the lipid bilayer remain unclear. The transmembrane β-barrel forming Outer Membrane Proteins (OMPs) from Gram-negative bacteria constitute a model system of particular interest due to their ability to self-insert into lipid membranes or membrane-mimetic environments such as detergent micelles. Here, we studied the folding mechanism of the Outer Membrane Phospholipase A (OmpLA) from E. coli using diffusion-based single-molecule Förster Resonance Energy Transfer (smFRET). By probing intramolecular distance changes upon equilibrium chemical denaturation using guanidine hydrochloride (GdnHCl) in the presence of lauryldimethylamine-N-oxide (LDAO) detergent micelles, we identify different FRET populations that reflect the different conformational states of OmpLA across the gradient of GdnHCl. From such experiments we obtain quantitative thermodynamic information that determines the folding mechanism of OmpLA. Furthermore, the kinetic rates of reversible folding and unfolding of OmpLA become accessible by real-time monitoring such reactions using smFRET. Our observations endorse the potential of smFRET and OmpLA as a powerful tandem to cast new light on the long-standing puzzle of membrane protein folding.

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Engineering FRET pairs to study the dynamics of protein conformational changes with a frequency-domain perturbative technique involving temperature oscillations

D. Kostrz,* T. Strick,** and C. Gosse*

* Laboratoire de Photonique et de Nanostructures, LPN-CNRS, Marcoussis, France

** Institut Jacques Monod, Université Paris Diderot/CNRS, Paris, France

[email protected]

We have developed an instrument which enables to measure the rate constants of reaction involving biomolecules using the response of the latter species to small thermal modulations (Fig. 1) [1]. Such kind of technique has already provided valuable information regarding the pairing between oligonucleotides [1,2] and we wish to apply it to the measurement of conformational change kinetics in proteins (upon heating or ligand binding). We have selected the folding/unfolding of the phosphoglycerate kinase enzyme as a starting point since a FRET construct enabling dynamics investigations with a T-jump setup already exist [3]. In the initial configuration a green fluorescent protein, AcGFP1, was used as a donor and a red fluorescent protein, mCherry, as an acceptor. Herein we present how the FRET pair was modified in order to improve the system reliability and its response upon unfolding. In particular, relying on a SNAP-tag conjugated to an organic chromophore and acting as an acceptor has proved valuable.

Fig. 1. Principle of the experiment proposed to measure conformational change dynamics. (A) The protein of interest is labeled with genetically encoded tags involved in FRET. (B) The reactive system

is introduced in a microfluidic chamber heated by an ITO resistor and placed on the stage of an epifluorescence microscope. (C) Injection of a sinusoidal current (––) generates temperature

oscillations. In turn, the chemical system rate constants are also modulated, as well as the species concentrations. The resulting fluorescence oscillations (–– et ––) are analyzed by lock-in detections

and the variations of the amplitude and phase of the response with the frequency yields the thermokinetic parameters characterizing the investigated chemical system.

[1] K. Zrelli, T. Barilero, E. Cavatore, H. Berthoumieux, T. Le Saux, V. Croquette, A. Lemarchand, C. Gosse, L. Jullien. “Temperature modulation and quadrature detection for selective titration of two-state exchanging reactants”. Anal. Chem. 83, 2476 (2011); K. Bournine, X. Zhao, C. Gosse. “Kinetic measurements using the frequency response of interacting biomolecules subjected to a thermal modulation”. Proceedings MicroTAS 2013, pp. 1800-1802. [2] D. Braun, A. Libchaber. “Lock-in by molecular multiplication”. Appl. Phys. Lett. 83, 5554 (2003); I. Schoen, H. Krammer, D. Braun. “Hybridization kinetics is different inside cells”. Proc. Natl. Acad. Sci. USA, 106, 21649 (2003). [3] S. Ebbinghaus, A. Dhar, D. McDonald, M. Gruebele. “Protein folding stability and dynamics imaged in a living cell”. Nature Methods 7, 319 (2010).

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Two step millisecond kinetics of the fourU RNA thermometer revealed by single-molecule FRET

Andreas Hartmann1, Frederic Berndt2 and Michael Schlierf1

1 B CUBE Center for Molecular Bioengineering, Technische Universität Dresden, Germany 2 Max-Planck-Institut für molekulare Zellbiologie und Genetik, Dresden, Germany

[email protected]; [email protected]; [email protected]

Monitoring and reacting to changes of the ambient temperature is essential for bacterial and eukaryotic survival. A prominent example is the fourU RNA thermometer found in Salmonella enterica, a non-coding mRNA hairpin structure, which suppresses protein expression by base-pairing with the Shine-Dalgarno sequence. A fine tuned base pairing sequence, which melts very cooperatively at elevated temperatures and subsequently allows ribosome binding and expression of the heat shock protein AgsA. The high cooperativity of the 34bp long RNA sequence is unusual and till date not well understood. Here, we study the fourU RNA opening and closing dynamics using single-molecule FRET combined with infrared heating of the confocal volume. Hairpin opening and closing occurred on the millisecond timescale as previously only observed for significantly shorter sequences. We further studied a destabilizing single base-pair mutation (G13A-C24U), which led to a reduced melting temperature and cooperativity. Additionally, we found a significant population of an on-path intermediate state for both variants at low temperatures indicating a basal expression of AgsA below the global melting temperature of the fourU.

Fig. 1. Molecular structure of the closed and fully open conformation of the fourU RNA hairpin. The temperature dependent opening and closing of the hairpin was monitored by single-molecule FRET.

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Tools in protein chemistry for multi-subunit and multidomain enzymes in smFRET

Simon Hartmann*, Airat Gubaev*, and Dagmar Klostermeier*

*University of Muenster, Institute of Physical Chemistry, Corrensstraße 28/30, 48149 Muenster

[email protected]

Single-molecule Foerster Resonance Energy Transfer (smFRET) has become a widely used tool to study conformational changes in proteins [1]. Though these proteins can be labeled statistically by modification of two amino acids with the same kind of chemical reaction, e.g. the reaction of cysteines with maleinimides, questions may arise where the site-specific introduction of probes becomes critical. An example is the investigation of conformational changes in multi-subunit and multi-domain enzymes, and of ATP-dependent enzymes that often undergo a cascade of concerted conformational changes during their catalytic cycle [2, 3].

Here we present different tools that can be used to address questions specific for the type II topoisomerase DNA gyrase by smFRET. Gyrase is a heterotetramer that consists of its two subunits in the arrangement (GyrB)2(GyrA)2. The heterotetramer catalyzes the ATP-dependent introduction of negative supercoils into DNA. To prevent subunit dissociation under single-molecule conditions we also used a GyrB-GyrA fusion protein [3]. Using tandem-affinity-chromatography we produced heterodimers, consisting either of two different subunits GyrA, GyrB-GyrA or of GyrB-GyrA and GyrA. These proteins could be labeled selectively on one subunit. Furthermore we generated the GyrB-GyrA fusion protein by expressed protein ligation (EPL). In this way, one could accomplish selective subunit labeling without dissociation under single molecule conditions. We also incorporated unnatural amino acids into gyrase that can be modified with fluorophores in orthogonal reactions. By a combination of these techniques, labeling schemes can be realized that allow simultaneous monitoring of multiple movements. Such experiments will contribute to our mechanistic understanding of gyrase activity and will also be relevant for other symmetric, multimeric proteins.

[1] Airat Gubaev and Dagmar Klostermeier, “The mechanism of negative DNA supercoiling: A cascade of DNA-induced conformational changes prepares gyrase for strand passage”, DNA Repair 16, 23—34 (2014).

[2] Bettina Theissen, Anne R. Karow, Jürgen Köhler, Airat Gubaev, and Dagmar Klostermeier, “Cooperative binding of ATP and RNA induces a closed conformation in a DEAD box helicase RNA helicase”, Proc. Natl. Acad. Sci. 105, 548—553 (2008).

[3] Airat Gubaev and Dagmar Klostermeier, “DNA-induced narrowing of the gyrase N-gate coordinates T-segment capture and strand passage”, Proc. Natl. Acad. Sci. 108, 14085—14090, (2011).

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69

Investigating Structural Dynamics of Mega-Enzymes by smFRET

Christina S. Heil, Alexander Rittner and Martin Grininger Institute for Organic Chemistry and Chemical Biology, Buchmann Institute for Molecular

Life Sciences, Cluster of Excellence “Macromolecular Complexes”, Goethe University Frankfurt, Max-von-Laue-Str. 15, 60438 Frankfurt am Main, Germany

[email protected]

Type I polyketide synthases (PKSs) are gigantic mega-enzymes, responsible for the biosynthesis of polyketide natural products. Iterative PKSs reuse one set of enzymatic domains repeatedly, whereas modular PKSs are organized in a linear arrangement of modules, used in a serial manner. Fatty acid synthases (FASs) use an iterative mode for fatty acid biosynthesis and are structurally and functionally similar to PKSs. Since the structure of modular PKSs is poorly characterized, the mammalian FAS represents an appealing model of structural organization of individual PKS modules. Single-particle EM studies revealed large conformational changes in the mammalian FAS, making it a very dynamic system. [1] To study those conformational dynamics we aim to perform single molecule FRET on these complex enzymes. Fluorophore FRET pairs are introduced site-specifically over the entire protein using amber codon suppression and incorporation of non-natural amino acids (nnAAs), which are suitable for click chemistry. We successfully generated FRET probes of FAS and PKS constructs using this novel technique and exceeded the limit of 100 kDa protein size for nnAA incorporation. With these valuable probes in hand, we want to address the following questions: (a) How much conformational flexibility is given within and between different modules? (b) What is the impact of conformational freedom on biosynthesis? (c) Can we map the movement of an acyl carrier protein? (d) What are the time frames of domain-domain interactions? (e) Do affinity and catalytic efficiency correlate? (f) Is there a substrate driven directionality of the ACP domain?

Fig. 1. Cartoon representation of the mammalian FAS. Catalytic domains arranged in one polypeptide chain and structurally organized in the homodimer. FRET pairs are introduced at different catalytic

domains and reveal the conformational dynamics of the enzyme.

[1] Brignole, E.J., Smith, S., and Asturias, F.J. (2009). Conformational flexibility of metazoan fatty acid synthase enables catalysis. Nat Struct Mol Biol 16, 190–197.

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Mechanistic insights into the multi-component Hsp90 machinery from smFRET

Ratzke C.*, Hellenkamp B.** and Hugel T.**

* Department of Physics, MIT, Cambridge, Massachusetts 02139, USA.

** Institute of Physical Chemistry, University of Freiburg, 79104 Freiburg, Germany

[email protected]

In living organisms, most proteins work in complexes to form multi-component protein machines. The function of such multicomponent machines is usually addressed by dividing them into a collection of two state systems at equilibrium - but many molecular machines work far from equilibrium by utilizing the energy of ATP hydrolysis. Here we show how multi-color single-molecule Förster resonance energy transfer (FRET) enables us to observe the ATP-dependent multi-component machine heat shock protein 90 (Hsp90) out of equilibrium [1]. In particular, we show how cochaperones modify the coupling between ATP hydrolysis and kinetic steps involved in the Hsp90 system.

[1] C. Ratzke et al., “Four-colour FRET reveals directionality in the Hsp90 multicomponent machinery”, Nature Communications 5:4192 (2014)

A-P13

71

Investigation of Energy Transfer in Light Harvesting Antenna Systems

Damla Inan*, Rajeev K. Dubey**, Ernst J. R. Sudhölter**, Wolter F. Jager**,

Ferdinand C. Grozema*

* Optoelectronic Materials

** Organic Materials and Interfaces, Chemical Engineering Department, Delft

University of Technology, Julinalaan 136, 2628 BL Delft, The Netherlands

E-mail: [email protected]

Development of new light harvesting antenna systems plays an important role to convert sunlight to chemical energy using a water splitting process such as artificial photosynthesis. Prior artificial photosynthetic devices have limited quantum efficiency and exhibits rapid photo-degradation. In order to meet the requirements, we have developed dendritic structures where the center consists of a perylene dye, surrounded by naphthalene moieties with a blue-shifted absorption.[1,2] These dendritic light-harvesting structures show Förster energy transfer to the center of the structure. Energy transfer of these structures was studied before the assembly on the surface by time resolved florescence spectroscopy and transient absorption spectroscopy. These results demonstrate that the energy transfer shows a marked dependence on the redox properties of donor and acceptor moieties, and the polarity of the solvent.

Fig.1: Schematic Representation of FRET from naphthalene moieties to perylene

group.

[1] Sengupta, S.; Dubey, R. K.; Hoek, R. W. M.; van Eeden, S. P. P.; Gunbaş, D. D.; Grozema, F. C.; Sudhölter, E. J. R.; Jager W. F. J. Org. Chem. 2014, 79, 6655–6662.

[2] Dubey, R. K.; Westerveld, N.; Grozema, F. C.; Sudhölter, E. J. R.; Jager W. F. Org. Lett. 2015, 17, 1882–1885.

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Simultaneous Measurements of Thickness and Diffusion of Lipid Bilayers using MIET and 2f-FLCS

Sebastian Isbaner*, Falk Schneider**, Narain Karedla*, Ingo Gregor*, and Jörg Enderlein*

* Drittes Physikalisches Institut, Georg-August-Universität Göttingen, Germany

** Weatherall Institute of Molecular Medicine, John Raddcliffe Hospital, University of Oxford, United Kingdom

[email protected]

Diffusion plays a key role for passive transport and signaling in cell membranes. A lipid bilayer is a simple model system for these membranes which has been used extensively for diffusion studies. As the thickness of a bilayer is just a few nanometers, the diffusion coefficient is usually an average over both leaflets. We report here the first simultaneous measurement of thickness and diffusion coefficients for both leaflets of a bilayer by combining metal-induced energy transfer (MIET) and two focus fluorescence lifetime correlation spectroscopy (2f-FLCS). MIET describes the phenomenon that a fluorophore is quenched when it is close to a metal surface. Similar to Förster resonance energy transfer (FRET), the quenching is distance dependent, and from the measured lifetime of the fluorophore the distance to the surface is obtained with nanometer precision. 2f-FLCS uses different fluorescence lifetimes to separate the contribution of spectrally indistinguishable fluorophores to the intensity correlations. Because of the MIET effect, fluorophores in the two leaflets will have different lifetimes for which 2f-FLCS can determine individual diffusion coefficients.

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73

Single-molecule FRET experiments for investigating the binding mechanism of the enzyme PARP-1 to DNA single-strand breaks

Eleni Kallis*, David Neuhaus**, Sebastian Eustermann*** and Jens Michaelis*

* Ulm University, Institute of Biophysics, Albert-Einstein Allee 11, 89081 Ulm, Germany

** MRC Laboratory of Molecular Biology, Francis Crick Avenue, Cambridge CB2 0QH, UK

*** Ludwig-Maximilians-University Munich, Gene Center and Department of Biochemistry, Feodor-Lynen-Straße 25, 81377 Munich, Germany

[email protected]

Combining the techniques of pulsed interleaved excitation (PIE) and multiparameter fluorescence detection (MFD) allows for the separation of various subpopulations in heterogeneous samples according to several characteristic parameters such as FRET efficiency, lifetime, anisotropy or labeling stoichiometry. Therefore, the method is ideally suited to study the dynamics of protein-DNA complexes [1,2]. Here, we report on recent single-molecule FRET experiments investigating the binding mechanism of Poly-(ADP-ribose)-polymerase-1 (PARP-1) to DNA single-strand (ss) breaks. PARP-1 is a nuclear enzyme involved in processes like DNA repair and transcription regulation. It is a key DNA damage signaling protein that detects DNA ss-breaks because of the DNA’s ability to adopt a highly kinked conformation [3]. We use intramolecular FRET to visualize the protein-induced conformational change of a fluorescently labeled DNA with a ss-break in the middle (see figure 1A). First results indicate that only a minor population of DNA is kinked in solution, while upon addition of PARP-1 the DNA undergoes a conformational change into a kinked structure (see figure 1B).

Fig. 1. A: Schematic of the double-labeled DNA dumbbell with ss-break. F1-F2 (cyan) represents a fragment of PARP-1 consisting of its two N-terminal zinc fingers. B: 2D-histograms of FRET-

efficiency and stoichiometry for measurements with the DNA shown in A (left) or with the same DNA plus the F1-F2 fragment (right).

[1] V. Di Cerbo et al. “Acetylation of histone H3 at lysine 64 regulates nucleosome dynamics and facilitates transcription”, eLife, 3:e01632, (2014). [2] W. Kügel, A. Muschielok and J. Michaelis. “Bayesian-Inference-Based Fluorescence Correlation Spectroscopy and Single-Molecule Burst Analysis Reveal the Influence of Dye Selection on DNA Hairpin Dynamics”, ChemPhysChem, 13, 1013-1022, (2012). [3] S. Eustermann et al. “Structural Basis of Detection and Signaling of DNA Single-Strand Breaks by Human PARP-1”, Molecular Cell, 60, 742-754, (2015).

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74

Single-Molecule Metal Induced Energy Transfer (smMIET): Resolving nanometer distances at single molecule level

Karedla N., Chizhik A. I., Gregor I., Chizhik A. M., Schulz O. and Enderlein J.*

*III. Institute of Physics, Georg August University, 37077 Göttingen, Germany [email protected]

We present a new concept for measuring distance values of single molecules from a surface with nanometer accuracy using the energy transfer from the excited molecule to surface plasmons of a metal film [1]. We measure the fluorescence lifetime of individual dye molecules deposited on a dielectric spacer as a function of a spacer thickness. By using our theoretical model [2], we convert the lifetime values into the axial distance of individual molecules. Similar to Förster resonance energy transfer (FRET), this allows emitters to be localized with nanometer accuracy, but in contrast to FRET the distance range at which efficient energy transfer takes place is an order of magnitude larger. smMIET can be potentially used as a tool for measuring intramolecular distances of biomolecules and complexes by combining fluorescence lifetime measurements on single molecules together with their orientations [3].

Fig. 1. Fluorescence lifetime of a single molecule as a function of the distance from a metal surface. The bottom right inset shows the lifetime distributions of single molecules on top of various spacer thicknesses.

[1] Karedla, N., Chizhik, A.I., Gregor, I., Chizhik, A.M., Schulz, O., Enderlein, J. “Single‐Molecule Metal‐Induced Energy Transfer (smMIET): Resolving Nanometer Distances at the Single‐Molecule Level”, ChemPhysChem, 15, 705-711 (2014).

[2] Enderlein J. "A theoretical investigation of single molecule fluorescence detection on thin metallic layers", Biophys. J. 78, 2151-8 (2000).

[3] Karedla, N., Stein, S. C., Hähnel, D., Gregor, I., Chizhik, A., & Enderlein, J.‚“Simultaneous Measurement of the Three-Dimensional Orientation of Excitation and Emission Dipoles”, Phys. Rev. Lett. 115, 173002 (2015).

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75

Selective bioorthogonal labeling of protein complexes in cell extracts for fast single-molecule pulldown assays

Kramm K.*, Gust A.*, Tinnefeld P.**, Grohmann D.*

*Universität Regensburg, Institut für Genetik, Biochemie und Mikrobiologie, Lehrstuhl fürMikrobiologie, Universitätsstraße 31, 93053 Regensburg

**Technische Universität Braunschweig, Institut für Physikalische und Theoretische Chemie, NanoBiosciences Workgroup, Hans-Sommer-Straße 10, 38106 Braunschweig

[email protected]

Single-molecule fluorescence energy transfer (smFRET) measurements are ideally suited to explore the structure-function-dynamics relationships of biomolecular complexes. Setting up a smFRET experiment often requires elaborate purification and in vitro reconstitution steps that strip a biomolecule of its natural binding partners. In this work, we combined a set of sophisticated biochemical and biophysical techniques into a single workflow that allows for fast site-specific fluorescent labeling of protein complexes in cellular extracts and subsequent smFRET analysis of immobilized molecules under near physiological conditions. A GST-tagged variant of the heterodimer Rpo4/7, a subcomplex of the archaeal RNA polymerase, was used as a model system to explore the limits of this workflow. First, applying the amber suppression strategy [1] we incorporated the unnatural amino acid p-azidophenylalanine into each subunit of the complex during recombinant co-expression of the proteins in E. coli. Second, after cell lysis, the Staudinger-Bertozzi-Ligation [2] was employed to stochastically label the complex with the organic fluorophors DyLight 550 and DyLight 650 directly in the crude cell extract. This bioorthogonal labeling scheme was highly specific yielding almost exclusively fluorescently labeled Rpo4/7. Third, the labeled complex was immobilized for single molecule TIRF microscopy measurements employing the single-molecule pulldown technique [3] without prior removal of the excessive dye from the labeling reaction. Even with a high background of free dye, the inter dye distance of Rpo4/7 could be precisely determined via smFRET and was in good agreement with the distance derived from the crystal structure. Hence, the newly developed workflow allows the fast preparation of native protein complexes for smFRET measurements yielding quantitative distance information in less than a day.

[1] T. S. Young, I. Ahmad, J. A. Yin, and P. G. Schultz, "An Enhanced System for Unnatural Amino Acid Mutagenesis in E. coli", J. Mol. Biol., vol. 395, no. 2, pp. 361–374, (2010).

[2] K. L. Kiick, E. Saxon, D. a Tirrell, and C. R. Bertozzi, "Incorporation of azides into recombinant proteins for chemoselective modification by the Staudinger ligation", Proc. Natl. Acad. Sci. U. S. A., vol. 99, no. 1, pp. 19–24, (2002).

[3] A. Jain, R. Liu, B. Ramani, E. Arauz, Y. Ishitsuka, K. Ragunathan, J. Park, J. Chen, Y. K. Xiang, and T. Ha, "Probing cellular protein complexes using single-molecule pull-down", Nature, vol. 473, no. 7348, pp. 484–488, (2011).

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Fluorescence spectroscopy reveals large-amplitude conformational dynamics of the multifunctional protein GABARAP

Jakub Kubiak*, Christina Möller**, Thomas-Otavio Peulen*, Philipp Neudecker**, Claus A.M. Seidel*

* Institut für molekulare physikalische Chemie, Heinrich-Heine-Universität Düsseldorf,40225 Düsseldorf, Germany

** ICS-6 (Structural Biochemistry), Forschungszentrum Jülich, 52425 Jülich, Germany

[email protected]

The γ-aminobutyrate type A receptor-associated protein (GABARAP) from H. sapiens belongs to the MAP1 LC3 family of ubiquitin-like proteins, involved in vesicle transport and fusion events, in autophagy, and apoptosis [1]. Structure determination of GABARAP by NMR and X-ray crystallography suggested conformational heterogeneity in the micro- to millisecond timescale [2,3]. We apply our fluorescence spectroscopy toolkit to investigate conformational dynamics in a broad time range. The FRET distance landscape is investigated by analysis of fluorescence decay and FRET dynamics by correlation techniques (filtered FCS). We show how the MFD toolkit [4] can be applied to a dynamic system, providing a complete set of fluorescence spectroscopic parameters from a single measurement. We find unexpected conformational dynamics of the N-terminal domain at tens of microsecond time-scale and a new thermally-excited state. This process may have implications for the regulation of GABARAP interactions and its function as a hub protein.

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Fig. 1. Single Molecule FRET-MFD analysis of GABARAP variant E7C-F62C labelled with Alexa488 and Alexa647. A. Single molecule MFD histogram [4] shows relation between donor

lifetime and donor-acceptor fluorescence ratio, indicating “dynamic shift” of the FRET population from the “static FRET-line” (orange) caused by fast mixing of FRET states (limiting states showed with horizontal lines). B. 1D-FRET histogram analysis (dynamic PDA [5]) provides FRET states linked with kinetic rate constants. C. Fluorescence decay analysis provides static donor-acceptor distance distribution. D. Filtered FCS [6] provides information on timescales of FRET-dynamics.

[1] Mohrluder, J., M. Schwarten, et al. “Structure and potential function of gamma-aminobutyrate type A receptor-associated protein”,FEBS J, 276, 4989 (2009). [2] Kouno, T., K. Miura, et al. “1H, 13C and '5N resonance assignments of GABARAP, GABAA receptor associated protein” J Biomol NMR, 22, 97 (2002). [3] Schwarten, M., M. Stoldt, et al. “Solution structure of Atg8 reveals conformational polymorphism of the N-terminal domain” Biochem Biophys Res Commun, 395, 426 (2010). [4] Sisamakis, E., A. Valeri, et al. “Accurate Single-Molecule Fret Studies Using Multiparameter Fluorescence Detection” Method Enzymol, 475, 455 (2010). [5] Kalinin, S., A. Valeri, et al. “Detection of structural dynamics by FRET: a photon distribution and fluorescence lifetime analysis of systems with multiple states” J Phys Chem B, 114, 7983 (2010). [6] Felekyan, S., S. Kalinin, et al. “Filtered FCS: Species Auto- and Cross-Correlation Functions Highlight Binding and Dynamics in Biomolecules” Chemphyschem, 13, 1036 (2012).

A-P19

77

Effect of dye-linker dynamics on the interpretation of FRET experiments

Stanislav Kalinin1$, Simone Fulle2§, Ralf Kühnemuth1, Simon Sindbert1, Suren Felekyan1, Holger Gohlke2, Claus A.M. Seidel1

1Institut für Physikalische Chemie, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany.

2Institut für Pharmazeutische und Medizinische Chemie, Heinrich-Heine-Universität, Düsseldorf, Germany.

$Present address: Carl Zeiss Microscopy GmbH, Carl-Zeiss-Promenade 10, 07745 Jena, Germany

§Present address: BioMed X GmbH, Im Neuenheimer Feld 583, D-69120 Heidelberg,Germany.

[email protected]

Site specific coupling of fluorophores with long flexible linkers to biomolecules is widely used to study their structure and dynamics by measuring interdye distances via Förster resonance energy transfer (FRET). In a model study employing double-stranded RNA site specifically labeled with Alexa488 and Cy5 dyes we investigated the dynamics of restrained fluorophore motions and its impact on the quantitative interpretation of fluorescence anisotropy and FRET experiments. To this end we performed molecular dynamics (MD) simulations as well as ensemble and single-molecule fluorescence experiments. The dyes explore their sterically accessible volume on four major time scales: (i) very fast (~0.1 ns) local rotations, (ii) fast (a few ns) local motions, (iii) slower exchange between free and temporarily trapped dye configurations, and (iv) lateral fluorophore diffusion within the accessible volume on a ~100 ns time scale, as also confirmed by full fluorescence correlation spectroscopy of the FRET signal. In FRET analysis, this temporal behavior is well approximated as static distance and dynamic orientation averaging. Our findings rationalize and give rules for the use of simple and successful approximations such as accessible volume calculations and ⟨κ2⟩ = 2/3 in quantitative FRET analysis.

A-P20

78

An off-pathway route in Escherichia Coli transcription initiation

Lerner Eitan*, Chung Sangyoon*, Allen Benjamin**, Shuang Wang***, Jookyung J. Lee ****, Shijia Winson Lu*, Grimaud Wilson Logan*, Ingargiola Antonino*, Alhadid Yazan*, Borukhov Sergei****, Strick

Terence***, Taatjes J. Dylan**, Weiss Shimon*

* Dept. of Chemistry & Biochemistry, University of California Los Angeles, Los Angeles

** Dept. of Chemistry & Biochemistry, University of Colorado, Boulder

*** Institut Jacques Monod, Centre National de la Recherche Scientifique and University of Paris Diderot and Sorbonne Paris Cité

**** School of Osteopathic Medicine, Rowan University

[email protected]

Abstract

An essential and highly regulated step in gene expression is transcription initiation. After promoter binding and DNA unwinding (‘bubble opening’) and in the presence of nucleoside triphosphates (NTPs), the RNA polymerase (RNAP)-promoter initial transcribing complex (RPITC) engages in ‘abortive initiation’, a process in which RNAP cycles between synthesis and release of short RNA transcripts. In abortive initiation, RPITC is believed to undergo a sequence of transitions between different initiation sub-states. The kinetics of the production of a full RNA transcript starting at a late initiation state (e.g. after production of a 7-base transcript) is expected to be similar to that measured from an earlier initiation state (e.g. an n-base transcript; n<7). We developed an in vitro quenched kinetics assay using single molecule detection and quantification of single run-off transcripts. Using this assay and magnetic tweezer assays, we found that run-off transcription kinetics starting from late initiation states is slower than kinetics starting from earlier initiation states. The transcription elongation factor GreA, however, was able to accelerate run-off transcription kinetics starting from a late initiation state. Gel-based kinetics assays focusing on abortive initiation under NTP starvation suggest that the off-pathway route involves backtracking of one base of RNA transcript. Our results suggest that RPITC can enter an off-pathway state in which the nascent RNA is in a backtracked, temporally paused position. Moreover, the transition into this state is sensitive to NTP levels, suggesting that this off-pathway state could help regulate gene expression under stressed conditions.

Figure 1. Quenched kinetics derived from DNA FRET probe hybridization to an RNA transcript

A-P21

79

Folding pathways of designed protein polyhedra

Ajasja Ljubetič*, Igor Drobnak*, Jana Aupič*, Helena Gradišar*, Roman Jerala*

* National Institute of Chemistry, Hajdrihova 19, Ljubljana, Slovenia

Corresponding author: [email protected]

Protein origami1, inspired by the success of DNA origami, uses modular components with

exactly defined binding partners to circumvent the lack of well-defined complementarity rules

between amino acids. The topology of the interacting building blocks (i.e. their location on the

polypeptide chain) determines the three dimensional shape of such TOPOFOLD proteins. As

an example, a tetrahedron has been constructed out of six coiled coil pairs2.

The folding pathway is crucial for a successful origami design as topofold proteins do not

possess a compact hydrophobic core. Therefore the folding pathway is determined by the

topology of building blocks and the order in which these building blocks assemble. Recently

the folding pathway of a single-stranded DNA pyramid has been constructued3, showcasing

how the control of the folding pathway enables the design of knotted structures.

Here we have examined the folding of the protein tetrahedron and square pyramid using all-

atom structure based (Gō) simulations. We are using these simulations to test how different

arrangements of building blocks in the protein sequence affect its folding pathway. This

allows us to design optimized versions of topofold proteins with smooth folding pathways in

order to avoid misfolding and aggregation. From the simulations it is also possible to predict

where FRET labels can be placed in order to gain maximum experimental information.

We have also examined the folding pathways using stop-flow techniques. The chevron plot

shows a weak dependence on denaturant concentration, which is compatible with multistate

folding pathways observed in Gō simulations.

The folding pathway of a protein tetrahedron obtained from Gō simulations. Each ellipse represents a state

with a varying number of formed edges. Thicker arrows represent a higher number of observed transitions.

Orange coloured ellipses represent the most frequently observed states.

Refrences:

[1] Kočar, V. et al. “TOPOFOLD, the designed modular biomolecular folds: polypeptide-based molecular

origami nanostructures following the footsteps of DNA.” WIREs Nanomed Nanobiotechnol (2014).

doi:10.1002/wnan.1289

[2] Gradišar, H. et al. “Design of a single-chain polypeptide tetrahedron assembled from coiled-coil segments.”

Nat. Chem. Biol. 9, 362–366 (2013).

[3] Kočar, V. et al. “Design principles for rapid folding of knotted DNA nanostructures.” Nat. Commun. In press,

(2016).

A-P22

80

Structural dynamics of metabotropic glutamate receptors by single-molecule FRET Anne-Marinette Cao*, Fataneh Fatemi*, Linnea Olofsson*, Suren Felekyan**, Etienne

Doumazane***, Pauline Scholler***, Ludovic Fabre***, Jurriaan M. Zwier****, Philippe Rondard***, Claus A.M. Seidel**, Jean-Philippe Pin*** and Emmanuel Margeat*

* Centre de Biochimie Structurale, CNRS UMR5048, INSERM U1054, Université deMontpellier, 34090 Montpellier, France

** Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität, 40225 Düsseldorf, Germany

*** Institut de Génomique Fonctionnelle, UMR 5203 CNRS, INSERM U661, Université de Montpellier, 34090 Montpellier, France

**** Cisbio Bioassays, 30200 Codolet, France

Contact : [email protected]

G protein-coupled receptors (GPCRs) constitute the most abundant protein family in mammalian genome andare the target for about 30% of the drugs on the market. Metabotropic glutamate receptors (mGluRs) belong to the class C of GPCRs and are involved in controlling synaptic transmission and in various central nervous system disorders such as pain, Parkinson’s disease, schizophrenia, etc…They are homodimers, and each protomer consists of a heptahelical transmembrane domain linked to a bilobate Venus flytrap domain (VFT) by a Cystein-rich domain. Understanding the conformational changes of mGluRs is essential to decipher the allosteric transitions associated with their activation. Since certain ambiguities and discrepancies about the resting and active states of mGluRs have been observed by X-ray crystallography, we first used Multi-parameter Fluorescence Detection (MFD) and Pulsed Interleaved Excitation (PIE) on isolated VFTs in solution. We observed that the mGluR VFTs oscillate between the resting and active states in a time range of 50-100 µs, and explained the molecular basis of the activity of partial agonists [1]. Now, in order to gain additional insights into mGluR activation, we investigate full-length receptors, solubilized in detergents or reconstituted into liposomes. Our current results confirm the structural dynamics obtained on the mGluR VFTs, and suggest a stabilizing role of the transmembrane domain. Further studies on allosteric modulation and G protein effect are on-going.

[1] Olofsson et al. “Fine tuning of sub-millisecond conformational dynamics controls metabotropic glutamate receptors agonist efficacy”, Nature Communications, 5, 5206, (2014)

A-P23

81

FRET in Fluorophore-Labeled DNA Crystals Joseph S. Melinger *, Ruojie Sha**, Nadrian Seeman**, and Mario G. Ancona*

* Naval Research Laboratory, Washington DC 20375 ** Department of Chemistry, New York University, New York, NY

[email protected]

Structural DNA nanotechnology provides a flexible means for controlling the structure of matter on a nanometer scale [1]. A recent example is the demonstration of self-assembled DNA crystals based on a DNA motif referred to as a tensegrity triangle [2]. The facile addressability of DNA through established chemical attachment methods further allows these DNA crystals to be functionalized in a variety of ways including with fluorescent dye molecules [3]. Here, we extend prior work by focusing on the optical and fluorescence resonance energy transfer (FRET) properties of fluorescent DNA crystals. The single unit of the DNA crystal studied here is shown in Fig. 1, and consists of two triangle-like molecules (A and B) that are linked by complementary sticky ends. Fluorophores Cy3 and Cy5 may be covalently attached to the 5’ end of the central DNA strand to form any of the combinations ACy3+B, A+BCy5, or the FRET pair ACy3+BCy5 that is depicted in Fig. 1a. These units are then self-assembled into rhombohedral lattices containing one or both fluorophores. For the latter case, Fig. 1b shows an optical micrograph of the purple ACy3+BCy5 crystals suspended in a drop of mother liquor. The optical responses of the dye-labeled crystals are characterized by steady-state and picosecond time-resolved fluorescent spectroscopy. A first set of plots compares the fluorescent transients for Cy3 (Fig. 1c) when free in solution versus when incorporated in a DNA crystal. That the in-crystal spectra are nearly unchanged, the lifetime is longer, and the decay more purely exponential show that being in the crystal has no deleterious effects and that the local dye environment is more uniform. In Fig. 1d we demonstrate that FRET does indeed occur in the ACy3-BCy5 crystal by showing complementary Cy3 donor decay and Cy5 acceptor induction signals. These may be compared to the fluorescence decay of the donor-only ACy3+B crystal. These and other experiments will be discussed along with interpretations derived from numerical simulations of the FRET dynamics including both hetero- and homo-FRET processes.

[1] N.C. Seeman, Structural DNA Nanotechnology (Cambridge University Press, 2016). [2] J. Zheng et al., “From molecular to macroscopic via the rational design of a self-assembled 3D DNA crystal,” Nature, 461, 74-77 (2009). [3] T. Wang, R. Sha, J. Birktoft, J. Zheng, C. Mao, and N. C. Seeman “A DNA crystal designed to contain two molecules per asymmetric unit”, J. Am. Chem. Soc., 132, 15471, (2010).

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Fig. 1. (a) DNA bi-molecularasymmetric triangle unit (A+B) with covalent 5’ attached fluorophores. (b) An optical micrograph of ACy3+BCy5 crystals in the mother liquor. (c) Comparison of Cy3 transients in solution and in crystal (ACy3+B), with each unit labeled, and 1 out of 64 units labeled. (d) Time-resolved FRET of ACy3+BCy5 crystals. Red lines are fits using exponential response functions.

A-P24

82

Probing the same inter-dye-distance of 6 nm by superresolution DNA-PAINT and FRETon DNA Origami

Julia Molle*, Mario Raab*, Ija Jusuk*, Sarah Schulz, Dina Grohmann** and Philip Tinnefeld*

*Technische Universität Braunschweig, Institut für Physikalische und Theoretische ChemieNanoBioSciences, Hans-Sommer-Straße 10, 38106 Braunschweig, Germany

**Department Mikrobiologie & Archaeenzentrum, University of Regensburg, Universitätsstraße 31, 93053 Regensburg, Germany

[email protected],

Distance measurements in the biomolecular relevant distance range of 1 -10 nm are typically carried out using Förster resonance energy transfer (FRET) experiments. FRET has been instrumental to determine the structure and to observe the dynamics of biomolecular complexes. However, with the advent and rapid development of super-resolution (SR) microscopy, distances as low as 6 nm could be resolved [1,2]. Hence SR microscopy advances to the distance range of a few nanometers characteristic for proteins and nucleic acids which hitherto has been inaccessible for standard fluorescence microscopy approaches.

Using rectangular DNA Origami as programmable nanostructure [3] with varying binding sites we were able to carry out comparative FRET and SR microscopy measurements on the samebiomolecular object. We measured the same defined distance of 6 nm between two dyes employing the SR microscopy technique DNA-PAINT [1] as well as FRET to evaluate the accuracy of the distance determination. We show that the results obtained with both techniques are highly consistent and that the resolution gap between FRET and SR microscopy is now truly closed.

[1] M. Raab, J.J. Schmied, I. Jusuk, C. Forthmann, P. Tinnefeld, Fluorescence microscopy with 6 nm resolution on DNA origami, ChemPhysChem, 15 (2014) 2431–2435.[2] R. Jungmann, C. Steinhauer, M. Scheible, A. Kuzyk, P. Tinnefeld, F.C. Simmel, Single-molecule kinetics and super-resolution microscopy by fluorescence imaging of transient binding on DNA origami, Nano Letters 10 (2010) 4756–4761.[3] Rothemund, Paul W K, Folding DNA to create nanoscale shapes and patterns, Nature 440 (2006) 297–302

A-P25

83

A Comparison of MreB Conformations upon Interactions with GroEL/ES and Tail-less

Complex Polypeptide 1 Ring Complex (TRiC) Chaperonins

Satish Babu Moparthi1, Uno Carlsson2, Renaud Vincentelli3, Bengt-Harald Jonsson2,

Per Hammarström2, & Jérôme Wenger1

1CNRS, Aix Marseille Université, Centrale Marseille, Institut Fresnel, 13013 Marseille, France. 2IFM, Department of Chemistry, Linköping University, 581 83 Linköping, Sweden. 3Architecture et Fonction des Macromolécules Biologiques (A.F.M.B), UMR7257 CNRS, Université Aix-Marseille, Case 932, 163 Avenue de Luminy, 13288 Marseille Cedex 9, France.

Correspondence email: [email protected]

GroEL/ES and tail-less complex polypeptide 1 ring complex (TRiC) systems are major

chaperonins ensuring correct functional folding of proteins in cells. Despite their crucial role

and the large scientific interest they have raised, their mechanisms of action and substrate

recognition is still open to fundamental questions. Here we investigate the mechanisms of

action of assisted protein folding of GroEL/ES and TRiC systems with the MreB substrate

protein. MreB is a homologue to actin in prokaryotes both structurally and functionally, and

plays a central role to control cell shape, division or locomotion. The strength of our approach

is to take advantage on complementary time-resolved fluorescence techniques (FCS,

anisotropy and FRET) to monitor the conformational rearrangements of MreB occurring in

GroEL/ES and TRiC assisted refolding. The significance of our work is two-fold. First, we

clearly establish that during folding MreB forms complexes with TRiC, GroEL and GroES

independently and in concert. Second, we also shown an unexpected role of GroES acting as

an unfoldase to induce a dramatic expansion of MreB and facilitate refolding in the GroEL/ES

system. GroES has a strikingly larger effect than GroEL or TRiC alone. Our analysis

importantly provides quantitative distance information about the MreB conformation

expansions for both GroEL/ES and TRiC systems.

A-P26

84

Excited-state annihilation reduces power dependence of single molecule FRET experiments

Daniel Nettels*, Dominik Haenni*, Sacha Maillot**, Moussa Gueye**, Anders Barth***, Verena Hirschfeld***, Christian G. Hübner***, Jérémie Léonard**, and Benjamin Schuler*

* Department of Biochemistry, University of Zurich

** Institut de Physique et Chimie des Matériaux de Strasbourg & Labex NIE, Université de Strasbourg

*** Institute of Physics, University of Lübeck

[email protected]

Single-molecule Förster resonance energy transfer (FRET) experiments are an important method for probing biomolecular structure and dynamics. The results from such experiments appear to be surprisingly independent of the excitation power used, in contradiction to the simple photophysical mechanism usually invoked for FRET. Here we show that excited-state annihilation processes are an essential cause of this behavior. Singlet-singlet annihilation (SSA) is a mechanism of fluorescence quenching induced by Förster-type energy transfer between two fluorophores while they are both in their first excited singlet states (S1S1), which is usually neglected in the interpretation of FRET experiments. However, this approximation is only justified in the limit of low excitation rates. We demonstrate that SSA is evident in fluorescence correlation measurements (Fig.1) for the commonly used FRET pair Alexa 488/Alexa 594, with a rate comparable to the rate of energy transfer between the donor excited state and the acceptor ground state (S1S0) that is exploited in FRET experiments. Transient absorption spectroscopy shows that SSA occurs exclusively via energy transfer from Alexa 488 to Alexa 594. Excitation-power dependent microsecond correlation experiments support the conclusion based on previously reported absorption spectra of triplet states that singlet-triplet annihilation (STA) analogously mediates energy transfer if the acceptor is in the triplet state. The results indicate that both SSA and STA have a pronounced effect on the overall FRET process and reduce the power dependence of the observed FRET efficiencies. The existence of annihilation processes thus seems to be essential for using FRET as a reliable spectroscopic ruler at the high excitation rates commonly employed in single-molecule spectroscopy [1].

Fig. 1. Singlet-singlet annihilation between FRET dyes is evident in nanosecond fluorescence cross-correlation measurements.

[1] D. Nettels et al., Phys.Chem.Chem.Phys. , 17, 32304 (2015)

A-P27

85

Position dependent fluorescent properties of coupled fluorescent dyes in large double-stranded RNA

Aiswaria Prakash*, Olga Doroshenko*, Hayk Vardanyan* Sascha Fröbel*, Stanislav Kalinin*, Simon Sindbert* and Claus A. M. Seidel*

*Chair of Molecular Physical Chemistry, Heinrich-Heine-Universität, Universitätsstraße 1,40225 Düsseldorf, Germany

[email protected]

Non-protein coding RNAs perform essential functions in living organisms. They commonly exhibit helical junctions as main architectural building blocks of RNA tertiary arrangements which define a dynamic ensemble of pre-existing conformational states. They are also widely used as building blocks and functional components in nanotechnology applications. However, the knowledge about the three dimensional structure of RNA junctions is rather limited. Förster-Resonance-Energy-Transfer (FRET) restrained high-precision structural modeling can be used to determine the structure of these junctions [1]. It has been shown that the dye environment influences the quenching of the fluorophores and thus the quantum yields [2].

Quantitative FRET studies were conducted to characterize the position dependent fluorescent properties of coupled fluorescent dyes in large double-stranded RNA. Two types of investigations were involved: The evaluation of labeling sites and the statistical analysis of the FRET distance deviation. The optimal choice for the labeling sites of the donor Alexa488 dye and the acceptor Cy5 dye is of great importance. Here we study 73 single labeled molecules at 15 donor and 16 acceptor labeling sites in 4 RNA three- way junctions, 1 RNA four-way junction molecule and 2 reference duplexes. We provide the statistical analysis of fluorescence quantum yields measured under the same local environments in different RNA junctions and their duplexes. Fluorescence anisotropy measurements were also performed to study the restriction of dye movement in the local environment. The results of our study show that labeling positions at the ends of the helices or at the junction region might bring unexpected behavior of donor and acceptor dyes. In the next stage, the results of quantitative FRET studies on 283 FRET pairs were compared. Systematic deviations of the calculated donor-acceptor distances from the model generated distances were statistically analyzed. This allowed us to identify the problematic labeling positions. Thus we could improve the accuracy of the FRET method by careful selection of labeling sites.

[1] Sindbert S, et al., “Accurate distance determination of nucleic acids via Förster resonance energy transfer: implications of dye linker length and rigidity”, J. Am. Chem. Soc. 133, 2463, (2011) [2] Seidel, C.A.M., “Nucleobase-Specific Quenching of Fluorescent Dyes. 1. Nucleobase One-Electron Redox Potentials and Their Correlation with Static and Dynamic Quenching Efficiencies”, J. Phys. Chem., 100 (13), 5541, (1996)

A-P28

86

Amyloid-like fibers and the role of lipids: Multibilayer structure and protein oligomerization from FLIM-FRET and homo-FRET

methodologies

1Manuel Prieto ,1Fábio Fernandes, 3,4, Luís Loura1,2, Ana Coutinho1Ana Melo 1. Centro de Química-Física Mol., I.S.T, UTL, Av. Rovisco Pais, 1049-001 Lisboa, Portugal2. Departamento Química e Bioquímica, FCUL, Campo Grande, 1749-016 Lisboa, Portugal

548 Coimbra, Portugal-Faculdade de Farmácia, Universidade de Coimbra, 3000 3 535 Coimbra, Portugal-bra, 3004Centro de Química de Coimbra, Universidade de Coim 4

[email protected]

Protein/peptide self-assembly into highly ordered β-sheet-rich fibrils plays a key role in various diseases, e.g., Alzheimer, Parkinson and Type II diabetes mellitus. Specifically, it has been suggested that the initial binding of amyloidogenic proteins/peptides to anionic membrane lipids can promote their pathological conversion into amyloidogenic assemblies. Surprisingly, it has also been proposed that this type of membranes can trigger rapid “amyloid-like” fibril formation of several non-amyloidogenic proteins, such as lysozyme and cytochrome c. In order to elucidate the key factors that govern the formation of these lipid-protein supramolecular complexes, lysozyme was here used as a model protein.

A detailed structural study of the fibers via advanced time-resolved FRET methodologies was carried out. Two different experimental designs were used: the derivatized-protein with the Alexa probe was used as donor to derivatized lipids as acceptors, or intra and interbilayer FRET involving only derivatized lipids. Data was obtained both in bulk solution (ensemble average), that allowed fitting the exact solutions for interplanar FRET, and in addition FRET efficiency was also determined at the single-fiber level via FLIM. It was concluded that these supramolecular complexes are multi-stacked pinched membranes, with proteins in-between [1].

The variation of the mean fluorescence lifetime of Alexa488-fluorescently-labelled lysozyme (Ls-A488) as a function of the surface coverage of the anionic liposomes was analyzed within the framework of a three-state cooperative partition/oligomerization model, revealing the reversible formation of oligomers with at least 6 monomers [2]. An energy migration (homo-FRET) study further established that the membrane-bound Lz-A488 oligomer stoichiometry is 6 1 [3]. The pronounced decrease detected in the fluorescence anisotropy of Lz-A488 always correlated with the system reaching a high membrane surface density of the protein (at a low lipid-to-protein (L/P) molar ratio). The occurrence of energy homo transfer-induced fluorescence depolarization was further confirmed by measuring the anisotropy decays of Lz-A488 under these conditions. Funding from FCT (Portugal), Project FAPESP/20107/2014 is acknowledged.

[1] A. Melo et al., “Electrostatically driven lipid-lysozyme mixed fibers display a multilamellar structure without amyloid features” Soft Matter 10 (6), 840 – 850 (2014) [2] A. Melo et al., Fluorescence detection of lipid-induced oligomeric intermediates involved in lysozyme “amyloid-like” fiber formation driven by anionic membranes. J. Phys. Chem B. 117(10):2906-17 (2013) [3] A. Melo et al., “Exploring homo-FRET to quantify the oligomer stoichiometry of membrane-bound proteins involved in a cooperative partition equilibrium ”, Phys. Chem. Chem. Phys. 16 (34), 18105-18117 (2014)

A-P29

87

Following the Structural Dynamics of Elongation Factor G during Ribosomal Translocation

Enea Salsi, Elie Farah, Jillian Dann, Zoe Netter & Dmitri N. Ermolenko

Department of Biochemistry and Biophysics & Center for RNA Biology, University of Rochester Medical Center (Rochester, NY, USA)

Contact: [email protected]

During protein synthesis, translocation of mRNA and tRNAs through the ribosome is catalyzed by a

universally conserved elongation factor (EF-G in prokaryotes and EF-2 in eukaryotes). EF-G binds to the

ribosome in a GTP-bound form and subsequently undergoes ribosome-stimulated GTP hydrolysis. Previous

studies have suggested that EF-G undergoes large-scale structural rearrangements that promote translocation.

We follow the movement of domain IV of EF-G, which is critical for translocation, relative to protein S12 of

the small ribosomal subunit[1] and also relative to domain II of EF-G[2] using single-molecule Förster

resonance energy transfer (smFRET) and total internal reflection fluorescence (TIRF) microscopy. We show

that ribosome-bound EF-G adopts distinct conformations corresponding to the pre- and post-translocation

states of the ribosome. Our results suggest that, upon ribosomal translocation, domain IV of EF-G moves

toward the A site of the small ribosomal subunit facilitating the movement of peptidyl-tRNA from the A to the

P site. We found no evidence of direct coupling between the movement of domain IV of EF-G and GTP

hydrolysis.

[1] Salsi, E., Farah, E., Dann, J. and Ermolenko, D.N. “Following movement of domain IV of elongation factor G during ribosomal

translocation”, Proc Natl Acad Sci USA, 111, 15060-5 (2014).

[2] Salsi, E., Farah, E., Netter, Z., Dann, J., and Ermolenko, D.N. “Movement of elongation factor G between compact and extended

conformations”, Journal of Molecular Biology, 427, 454-67 (2015).

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qAN4: a second generation adenine analog as fluorescent probe to understand conformation and dynamics of nucleic acids.

Sangamesh Sarangamath *, Anders Foller Larsen*, Mattias Bood#, Moa Sandberg Wranne*, Morten Grøtli#, L Marcus Wilhelmsson*

* Department of Chemistry and Biochemistry, Chalmers University of Technology, Sweden.# Department of Chemistry and Molecular Biology, University of Gothenburg, Sweden.

[email protected]

Fluorescent nucleobase analogs constitute tools that can provide a detailed understanding of the structure and dynamics of nucleic acids. Our group has previously reported the cytosine analogue tCO which together with another cytosine analog tCnitro constitute the first nucleobase FRET-pair[1] which has opened up for new detailed studies of DNA.[2] To follow up on this interesting development of C-analogs, synthesis and photophysical studies of a quadracyclic adenine analog, qA, were performed.[3] However, the quantum yield of this adenine analog inside duplex DNA turned out to be significantly quenched (Φf<0.01). To enhance the photophysical properties of this A-analog, we recently used quantum chemically aided design and synthesized a second generation of quadracyclic adenine analogs.[4] qAN4 is one of these analogs with a significantly higher quantum yield (0.32 in water) compared to qA (Φf=0.07) and highly interesting fluorescence properties. In preliminary studies we have found that when qAN4 is incorporated into DNA, it is able to differentiate between neighboring bases through its fluorescence properties and reaches quantum yields inside duplexes that are more than 10 times higher than the previously developed qA. As for most other fluorescent base analogs, the quantum yield of qAN4 on average decreases going from a single- to double-stranded environment. Purines seem to have the smallest quenching effect on the fluorescence quantum yield of qAN4 and contrary to most fluorescent nucleobase analogs, qAN4 shows relatively minor quenching when incorporated with surrounding guanines. Importantly, circular dichroism (CD) and thermal melting studies show that qAN4 does not significantly affect the structure of DNA B-form helices. In fact the duplex is slightly stabilized compared to a non-modified DNA, most probably because of better stacking interactions between qAN4 and the neighboring bases. Moreover, qAN4 along with a recently developed adenine analog, qAnitro, will constitute a good adenine-adenine FRET donor-acceptor pair. In conclusion, qAN4, a second generation quadracyclic adenine analog, is a promising probe for the analysis of nucleic acid structures and dynamics.

Fig. 1. Structure of qAN4

[1] Karl Börjesson, et al. “Nucleic Acid Base Analog FRET-Pair Facilitating Detailed Structural Measurements in Nucleic Acid Containing Systems” J. Am. Chem. Soc., 131, 4288, (2009). [2] Yonghong Shi, et al. “Mammalian Transcription Factor A is a Core Component of the Mitochondrial Transcription Machinery” Proc. Natl. Acad. Sci. USA., 109, 16511, (2012). [3] Anke Dierckx, et al. “Quadracyclic Adenine: A Non-Perturbing Fluorescent Adenine Analogue” Chem. Eur. J., 18, 5987-5997, (2012). [4] Blaise Dumat, et al. “Second-Generation Fluorescent Quadracyclic Adenine Analogues: Environment-Responsive Probes with Enhanced Brightness”, Chem. Eur. J., 21, 4039-4048, (2015).

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The environment shapes the inner vestibule of LeuT

Azmat Sohail*, Kumaresan Jayaraman*, Santhoshkannan Venkatesan*, Kamil Gotfryd***, Markus Daerr****, Ulrik Gether**, Claus J. Loland**, Klaus T. Wanner****, Michael

Freissmuth*, Harald H. Sitte*, Walter Sandtner* and Thomas Stockner*.

*Medical University of Vienna, Center for Physiology and Pharmacology, Institute ofPharmacology, Waehringerstrasse, 13A, 1090 Vienna, Austria

**University of Copenhagen, Faculty of Health and Medical Sciences Denmark, Department of Neuroscience and Pharma cology, Blegdamsvej 3B, 2200 Copenhagen N, Copenhagen,

Denmark

***University of Copenhagen, Faculty of Health Sciences Denmark, Department of Biomedical Sciences, Blegdamsvej 3B, 2200 Copenhagen N, Denmark

****Ludwig Maximilians University Munich, Department of Pharmacy, Center of Drug Research, Butenandtstraße 7, D-81377 Munich, Germany

[email protected]

The role of human neurotransmitter transporters is fast termination of signal transmission by rapid removal of neurotransmitter molecules from the synaptic cleft. The transporters for the monoamines serotonin, dopamine and norepinephrine are members of the solute carrier family 6 (SLC6), located on the presynaptic neurons. The bacterial orthologue LeuT served since the first crystallization as a structural and functional paradigm for the SLC6 transporter family. LeuT was resolved in three conformations: outward-open, outward-occluded and inward-open. We investigated here the inward-open state of LeuT. We compared LeuT in membranes and micelles using molecular dynamics simulations and applied the lanthanide-based resonance energy transfer (LRET) for distance measurements. Simulations revealed a stable and widely open inward-open conformation of micelle solubilised LeuT that was similar to the crystal structure [1]. However, this conformation was unstable in a membrane environment and the first transmembrane helix (TM1A) partitioned out of the hydrophobic membrane core. We performed distance measurements using LRET and showed that the distance across the inner vestibule between the C-terminus that carries the lantanide binding tag and helix TM1A differed between the membrane environment and the micelle environment. The measured changes in distance were compatible with the computational predictions and showed that the conformation of the inward-open state of LeuT is dependent on the environment.

Krishnamurthy H, Gouaux E. X-ray structures of LeuT in substrate-free outward-open and apo inward-open states. Nature 481: 469-474 (2012).

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90

Pressure unfolding of a model folding protein followed by smFRET

Sven Schneider*, Erik Hinze**, Kim Reiter*, Christian Hübner* *Institute of Physics, University of Lübeck, Ratzeburger Allee 160, D-23562 Lübeck

**Max-Planck-Research Unit for Enzymology of Protein Folding, Weinbergweg 22, D-06120 Halle

[email protected]

Single molecule FRET (smFRET) has proven a powerful method for the study of the unfolded state of folding model proteins [1]. Unfolding by temperature and chemical denaturants has been elucidated by smFRET. Unfolding by high pressure, however, has not been demonstrated to date.

We introduce a method for smFRET measurements under high pressures (up to 3 kbar) based on a silica square bore capillary as a sample container [2]. The dimensions of the capillary with a wall thickness comparable to the thickness of standard microscope cover slips allow for their use on microscopes with high NA water immersion microscope objectives. We characterize the influence of optical aberrations in the microscope/capillary system by fluorescence correlation spectroscopy (FCS) analyses. We improve the imaging properties by placing the capillary on a silica coverslip with a thickness of 90 µm while immersing the capillary in an optical gel of suitable refractive index and dispersion.

We present results from smFRET unfolding experiments of the model folding protein CspA in solution under the application of pressure. The measurements allow for following the folding/unfolding transition as well as for assessing the dimensions of the unfolded chain. In contrast to unfolding by denaturants and temperature, where the size of the unfolded chain is increasing with increasing denaturant concentration or temperature, pressure leads to only subtle changes of the size of the unfolded chain.

Fig. 1. High-pressure setup

[1] Schuler, Lipman, Eaton” Probing the free-energy surface for protein folding with single-molecule fluorescence spectroscopy“, Nature, Vol 419, 743-747, (2002).

[2] Tekmen, Müller” High-pressure cell for fluorescence fluctuation spectroscopy”, Review of scientific instruments, Vol 75, 5143-5148, (2004).

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Dissecting carbocyanine photophysics in the context of RNA

Fabio D. Steffen, Roland K. O. Sigel, Richard Börner

Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland

[email protected]

Carbocyanine dyes have long been recognized as versatile probes in single-molecule spectroscopy due to their high photostability and large spectral separation [1]. A profound understanding of their photophysical properties in a specific macromolecular environment is essential for the interpretation of Förster resonance energy transfer (FRET) experiments [2,3]. We report on the interweaving of fluorophores, covalently bound to RNA, and metal ions in terms of the fluorescence lifetime, quantum yield and dynamic anisotropy [4]. Divalent cations like Mg2+ promote the interaction propensity of cyanines with nucleic acids and establish a dynamic equilibrium of tumbling and stacked fluorophores. Tracking the dye mobility on an atomic level by means of molecular dynamics allows to disentangle various interaction modes and to monitor the sterical constraints imposed by the RNA environment. Our hybrid approach combining time-correlated single photon counting (TCSPC) and computer simulations will benefit the analysis of absolute distance measurements by smFRET. The authors thank Hauke Paulsen and Christian Hübner (University of Lübeck) for access to the computational infrastructure. Financial support from the University of Zurich and the UZH Forschungskredit grant (FK-14-096 to RB) is gratefully acknowledged.

Fig.1. Fluorescence lifetime decay (left) and dynamic anisotropy (right) of Cy3, either freely diffusing or coupled to an RNA hairpin, namely the exon binding site (EBS1) of the group II intron

Sc.ai5γ (middle, PDB: 2m24 [5]).

[1] R. Roy, S. Hohng and T. Ha, "A practical guide to single-molecule FRET", Nat. Methods, 5, 507-516 (2008). [2] M. Levitus and S. Ranjit, "Cyanine dyes in biophysical research: the photophysics of polymethine fluorescent dyes in biomolecular environments", Q. Rev. Biophys, 44, 123-151 (2011). [3] E.M.S. Stennett, M.A. Ciuba and M. Levitus, "Photophysical processes in single molecule organic fluorescent probes", Chem. Soc. Rev., 43, 1057-1075, (2014). [4] F.D. Steffen, R.K.O. Sigel, R.Börner, to be submitted. [5] D. Kruschel, M. Skilandat and R.K.O. Sigel, "NMR structure of the 5′ splice site in the group IIB intron Sc.ai5γ – conformational requirements for exon–intron recognition", RNA, 20, 295-307 (2014).

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92

International Discussion Meeting – FRET in Life Sciences II– 3rd - 6th April 2016

Enhancing single-molecule FRET studies with photostabilizer–dye conjugates.

Jasper H. M. van der Velde1, Jens Oelerich2, Jingyi Huang3, Jochem H. Smit1, Atieh Aminian Jazi1, Silvia Galiani4, Kirill Kolmakov5, Giorgos Guoridis1, Christian

Eggeling4, Andreas Herrmann3, Gerard Roelfes2, Thorben Cordes1,

1 Molecular Microscopy Research Group, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands; 2 Stratingh Institute for Chemistry, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The

Netherlands; 3 Department of Polymer Chemistry, Zernike Institute for Advanced Materials, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands; 4 MRC

Human Immunology Unit, Weatherall Institute of Molecular Medicine, University of Oxford, Headley Way, OX3 9DS Oxford, UK; 5 Department NanoBiophotonics, Max-Planck-Institute

of Molecular Medicine, Am Fassberg 1, 37077 Goettingen, Germany [email protected]

FRET is a valuable tool to study conformational states and dynamics of biomolecules. Various FRET-applications but especially single-molecule based experiments are limited by signal fluctuations (blinking) and irreversible photodestruction (photobleaching). Up to date, photostabilizing buffer additives are the method of choice to enhance signal brightness and stability of fluorophores. This strategy has, however, various drawbacks such as associated toxicity of the photostabilizers and unreliable performance depending on the biological target. In this contribution, organic fluorophores with intramolecular photostabilization were tested for their performance in smFRET and ALEX spectroscopy. Unnatural amino acids were used as a flexible scaffold to bind organic fluorophores to a photostabilizer and a (bio)molecular target, i.e., here a periplasmic binding protein of an ABC transporter.1 We study the influence of intramolecular photostabilization of donor and acceptor fluorophores in smFRET and ALEX. Our results show that the simple addition of a single photostabilizer to the acceptor fluorophore is sufficient to gather reliable results on the conformational states of the protein from solution-based smFRET. We also find that the addition of a stabilizer on the donor-fluorophore can increase the overall available photon budget (linked to data quality), but the acceptor strictly requires a solution-based or covalently-linked photostabilization. The obtained results show that intramolecular photostabilization is an effective method to increase the photostability of fluorophores used for smFRET and Alex microscopy. Further unpublished results with covalently linked photostabilizers provides important information on observed differences in photostabilization efficiencies between DNA and proteins that have high relevance for smFRET studies even when using buffer-based photostabilization.

Fig. 1. Removal of photophysical artefacts in single-molecule FRET with photostabilizer-dye conjugates.

[1] van der Velde, J. H. M. et al., A simple and versatile design concept for fluorophore-derivatives with intramolecular photostabilization. Nature Communications 7 , 1–15 (2016).

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Studying the function of BAP in the nucleotide cycle of BiP by spFRET using MFD-PIEDaniela Wengler *, Mathias Rosam**, Jelle Hendrix*,***, Johannes Buchner** and

Don C. Lamb*

* Physical Chemistry, Department of Chemistry, Ludwig-Maximilians-Universität München,Center for Nanoscience (CeNs), Center for Integrated Protein Science Munich (CIPSM),

Nanosystems Initiative Munich (NIM), 81377 Munich, Germany

** Department Chemistry, Technische Universität München, Center for Integrated Protein Science Munich (CIPSM), 85748 Garching, Germany

*** Current address:Laboratory of Photochemistry and Spectroscopy, Division of Molecular Imaging and Photonics, KU Leuven, B-3001 Leuven, Belgium

[email protected]

Molecular chaperones, like Hsp70 and Hsp90, have different functions in a cell. One of their major tasks is to assist nascent proteins to reach their final functional structure in the appropriate location within the cell. Different disorders, such as Alzheimer or Parkinson’s disease, are linked to unfolded or missfolded proteins. Thus understanding the underlying mechanisms of chaperone-assisted protein folding is important for the development of treatment against these disorders. All Hsp70s consist of two domains that are connected by a short linker, the nucleotide binding domain (NBD) and the substrate binding domain (SBD), containing a flexible, alpha helical lid that can open or close the binding pocket of the SBD. When a client protein enters the ER, its charged regions are initially protected by the Hsp70 located in the endoplasmic reticulum (ER), called BiP, to avoid non-specific interactions and therefore missfolding. One of the regulating factors in this process is the BiP associated protein, BAP, a nucleotide exchange factor (NEF) of BiP. BAP accelerates the ATP/ADP exchange and therefore controls the binding and release of the nascent protein. In order to gain molecular insights into the function of BAP, we employed solution-based single-pair Förster Resonance Energy Transfer (spFRET) experiments using a Multiparameter Fluorescence Detection (MFD) setup combined with Pulsed Interleaved Excitation (PIE) [1]. BiP was fluorescently labeled with the FRET pair ATTO 532 and ATTO 647. We used three different spFRET mutants of BiP to monitor the conformation of the lid of BiP and to identify distance changes between the NBD and SBD upon binding of BAP in the presence and absence of a substrate. Our results suggest that BAP acts as a NEF and therefore enhances the rate of the nucleotide dependent chaperone cycle of BiP. We detect that without BAP, the lid opens twice during one nucleotide cycle, first upon binding of ATP and secondly when phosphate is released. By adding BAP, the lid stays in its open conformation during hydrolysis and BAP cannot bind to BiP, when non hydrolysed ATP is present. Furthermore BAP increases the dissociation constant of ADP and changes the nucleotide binding pocket of BiP to increase the rate of phosphate release after hydrolysis.

[1] Kudryavtsev, V.; Sikor, M.; Kalinin, S.; Mokranjac, D.; Seidel, C. A. & Lamb, D. C. “Combining MFD and PIE for accurate single-pair Forster resonance energy transfer measurements”, Chemphyschem, 13, 1060-78, (2012).

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Fluorescent nucleobase analogue development and their use in FRET-investigations L. Marcus Wilhelmsson *

* Department of Chemistry and Chemical Engineering/Chemistry and Biochemistry,Chalmers University of Technology, S-41296 Gothenburg, Sweden.

[email protected]

Fluorescent base analogues is a class of molecules that is rapidly increasing in importance for investigating systems containing DNA and RNA in biology and nanotechnology.[1] We use quantum chemical calculations in designing novel fluorescent base analogues with new or improved properties. The aim is to decrease the time and synthetic effort needed to find promising new candidate molecules and with these to develop fluorescent analogues for all natural nucleobases that can be used in FRET. Recently we have used this approach on a class of quadracyclic adenines, qAs,[2] and we are currently investigating their possible use in FRET measurements. We also utilize our previously developed family of molecules called tricyclic cytosines, tC, tCO, and tCnitro. In contrast to other reported fluorescent base analogues tCO, for example, has i) a high quantum yield (φf≈0.2) in duplex that is virtually insensitive to neighboring base combination, ii) an emission after incorporation into DNA being characterized by a single exponential decay in double stranded systems, and iii) an average brightness of the base analogues in duplex DNA being among the highest reported so far and up to 50 times higher than the most commonly used fluorescent base analogue 2-aminopurine.[3] Importantly, we have utilized tCO as a donor and developed tCnitro as an acceptor and, thus, established the first nucleic acid base analogue FRET-pair.[4] To allow optimized use of this kind of FRET-pair we have developed a freeware called FRETmatrix[5], suited for rigidly positioned probes (high control of orientation factor, κ2), that globally fit a set of time-resolved FRET-data to obtain the best overall structure/dynamics of the nucleic acid structure under investigation. Recently we have successfully utilized our FRET-pair in studies on DNA structural changes[6] and have ongoing investigations using our methodology to study both DNA-/RNA-conformations as well as using the emission from our fluorescent base analogues in the development of drug development assays for pharma industry. We envision our method, possibly in combination with single-molecule FRET on longer distances, to be a powerful complement for techniques like NMR and X-ray crystallography.

Fig. 1. Energy transfer for FRET-pair tCO-tCnitro in DNA as a function of base-base separation.

[1] Wilhelmsson, L.M. “Fluorescent Nucleic Acid Base Analogues” Q. Rev. Biophys., 43, 159, (2010). [2] Dumat, B. et al. ”Second-generation fluorescent quadracyclic adenine analogues: Environment-responsive probes with enhanced brightness” Chem. Eur. J., 21, 4039, (2015). [3] Sandin, P. et al. “Characterisation and Use of an Unprecedentedly Bright and Structurally Non-perturbing Fluorescent DNA Base Analogue” Nucleic Acids Res., 36, 157, (2008). [4] Börjesson, K. et al. “A Nucleic Acid Base Analog FRET-pair Facilitating Detailed Structural Measurements in Nucleic Acid Containing Systems” J. Am. Chem. Soc., 131, 4288, 2009. [5] Preus, S. et al. “FRETmatrix: A General Methodology for the Simulation and Analysis of FRET in Nucleic Acids” Nucleic Acids Res., 41, e18, (2013). [6] Shi, Y. et al. “Mammalian Transcription Factor A is a Core Component of the Mitochondrial Transcription Machinery” P.N.A.S., 109, 16510, (2012).

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95

Probes to study AS-membrane interactions

Volodymyr Shvadchak1, Oleksandr Kucherak1, Kseniia Afitska1 and Dmytro Yushchenko1,2

1 Institute of Organic Chemistry and Biochemistry (IOCB), Flemingovo nám. 2, 16610 Prague, Czech Republic

2 Cell Biology & Biophysics Unit, European Molecular Biology Laboratory (EMBL), Meyerhofstraße 1, 69117 Heidelberg, Germany

[email protected] address: -E

Aggregation of protein alpha-synuclein (AS) is a hallmark of Parkinson’s disease. The

mechanism of AS aggregation is not yet well understood. However, there is ample evidence

that modulation of AS-membrane interaction significantly affects the propensity of AS to

aggregate. Solvatochromic dyes may serve as suitable probes to study protein-membrane

interaction due to the dramatic change of their fluorescence properties upon interaction with

lipid bilayers. However, no systematic study of their sensitivity and structure optimization for

protein-membrane interaction was performed so far. We developed a series of optimized

covalent solvatochromic probes which we applied to study interaction of AS with lipid

membranes. Some of these dyes are also useful to study AS oligomerization and fibrilization.

We apply these probes in combination with commercial dyes for investigation of AS

oligomerization in the presence of lipid membranes. The propensity of AS to oligomerize we

quantify by FRET.

A-P38

96

Site-specific labeling of large RNA with fluorophores for the application in single molecule FRET studies

Meng Zhao, Fabio D. Steffen, Richard Börner, Eva Freisinger, and Roland K. O. Sigel

Department of Chemistry, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland

[email protected]

While solid-phase synthesis can be used for RNA sequences shorter than 60 nucleotides, a facile method for the site-specific labeling of large RNAs at internal sites is still unavailable. The demand for such a method is high since non-coding RNAs were found to have important biological functions. Our interest focuses on the btuB riboswitch from E. coli which is >200 nucleotides in length. Their re-structuring upon metabolite binding regulates gene expression and metabolite maintenance.[1] Recently, we developed a post-transcriptional method that enable the site-specific labeling of large RNA.[2] By applying this method, we prepared several btuB riboswitch constructs with varying labeling schemes (Fig. 1a). Each of these constructs carries Cy3 as Förster resonance energy transfer (FRET)-donor and Cy5 as FRET-acceptor fluorophore. These differently labeled btuB riboswitches were characterized at the single molecule level. The observed FRET traces (Fig. 1b) point to successful application of our labeling method to large RNAs. Details will be presented during conference. Financial support from the Swiss National Science Foundation (EF), the UZH Forschungskredit (FK-14-096, RB), the European Research Council (RKOS), the University of Zurich (EF, RKOS), and the SBFI (COST Action CM1105; EF, RKOS) is gratefully acknowledged.

Fig. 1. Site-specific labeling of the btuB riboswitch and the characterization at the single molecule level. (a) Schematic illustration of the btuB riboswitch and the chosen labeling sites. Donor and

acceptor are both positioned at two different sites, resulting in four different FRET labeling schemes (D1, A1), (D1, A2), (D2, A1), and (D2, A2). (b) Representative FRET traces (time resolution 0.1 s) of the

four different constructs.

[1] Mironov, A. S.; Gusarov, I.; Rafikov, R.; Lopez, L. E.; Shatalin, K.; Kreneva, R. A.; Perumov, D. A.; Nudler, E. “Sensing Small Molecules by Nascent RNA: A Mechanism to Control Transcription in Bacteria” Cell, 111, 747 (2002). [2] Zhao, M.; Steffen, F. D.; Börner, R.; Freisinger, E.; Sigel, R. K. O. “Site-specific labeling of large RNAs for single-molecule FRET” in preparation.

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97

PIE analysis in MATLAB (PAM) – A software package for the analysis of fluorescence experiments using pulsed interleaved excitation

Anders Barth *, Waldemar Schrimpf*, Jelle Hendrix** and Don C. Lamb*

* Physical Chemistry, Department of Chemistry, Ludwig-Maximilians-University, and Centerfor Integrated Protein Science (CIPSM) and Center for Nanoscience (CeNS), München

** Laboratory of Photochemistry and Spectroscopy, KU Leuven, Leuven [email protected]

In Pulsed Interleaved Excitation (PIE), alternating excitation of different fluorophores is performed on the nanosecond timescale using pulsed lasers [1]. PIE is an expansion of the method of Alternating Laser Excitaiton (ALEX) developed in the group of Shimon Weiss [2]. Using the information of arrival time with respect to the excitation pulses, photons can additionally be sorted based on excitation source. PIE has originally been developed for fluorescence cross-correlation spectroscopy, where the information about the excitation source of each photon enables quantitative and artifact-free analysis [1]. PIE has also been applied to diffusion-based single molecule single-pair FRET experiments and raster image correlation spectroscopy [2,3]. In our lab, we have developed a software package in MATLAB specifically designed for the analysis of experiments performed using PIE. The capabilities of PAM currently includes various fluorescence fluctuation techniques (FCS, FCCS, FLCS, filtered-FCS, RICS, TICS, STICS, iMSD, pair-correlation analysis), fluorescence lifetime and time-resolved anisotropy analysis, single-pair and single-triad FRET burst analysis (PIE-MFD) with photon distribution analysis (PDA), and fluorescence lifetime imaging using the phasor approach (Phasor-FLIM). All of these methods are integrated with the photon sorting capabilities that are available with PIE, providing an extensive analysis framework for researchers looking into upgrading their fluorescence microscope with PIE.

[1] Müller, B. K., Zaychikov, E., Bräuchle, C. & Lamb, D. C. “Pulsed interleaved excitation”, Biophys J, 89, 3508–3522 (2005) [2] Lee, N. K., Kapanidis, A. N., Wang, Y., Michalet, X., Mukhopadhyay, J., Ebright, R. H. & Weiss, S. “Accurate FRET Measurements within Single Diffusing Biomolecules Using Alternating-Laser Excitation”, Biophys J 88, 2939–2953 (2005) [2] Kudryavtsev, V., Sikor, M., Kalinin, S., Mokranjac, D., Seidel, C. A. M. & Lamb, D.C. “Combining MFD and PIE for Accurate Single-Pair Förster Resonance Energy Transfer Measurements”, ChemPhysChem 13, 1060–1078 (2012) [3] Hendrix, J., Schrimpf, W., Höller, M. & Lamb, D. C. “Pulsed Interleaved Excitation Fluctuation Imaging”, Biophys J 105, 848–861 (2013)

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98

Extended excitation FLIM for rapid FLIM-FRET determinations of EGFR conformational dynamics and association states

Nathan P. Cook*, Donna J. Arndt-Jovin*, Thomas M. Jovin** Laboratory of Cellular Dynamics, Max Planck Institute for Biophysical Chemistry,

37077 Göttingen, Germany

[email protected] The function of the epidermal growth factor receptor (EGFR) in living cells is driven by conformational transitions and clustering processes mediated by a variety of intermolecular interactions. We have explored these processes with ACP tagged CHO cells [1,2] in combination with other agents constituting FRET donor-acceptor pairs. A new, rapid FLIM method (eeFLIM) was used to determine the FRET efficiency (see also presentation Jovin et al.).

eeFLIM employs excitation schemes based on extended rectangular light pulses (e.g. 10-60 ns) and recording the integrated emission of a fluorophore or mixture of fluorophores with a gated intensified camera. The method provides pixel-by-pixel determinations of intensity weighted mean lifetimes of arbitrarily heterogeneous populations and the data evaluation is very simple (linear calculations), non-iterative and thus rapid. The utilization of all the light emitted per pulse provides a high signal SNR, allowing the acquisition of sequential 1K×1K lifetime images in studies of living cells.

Cells with EGFR labeled with Abberior Star440SX yielded high quality FLIM images with only 3 sequential images using increasing gate widths of detection (Fig. 1A,B). Upon addition of the NR12S acceptor, which incorporates into the outer leaflet of the plasma membrane, the lifetimes decrease (Fig. 1C), as was observed in previous studies [1,2]. The effects of ligand binding and of agents altering membrane potential have been studied in a collaboration with colleagues of the Univ. of Debrecen, Hungary, using this system [3]. We are also exploring EGFR clustering and internalization by eeFLIM-FRET using 2-color ACP tags and labeled EGF.

[1] Ziomkiewicz I., Loman A., Klement R., Fritsch C., Klymchenko A.S., Bunt G., Jovin T.M., and Arndt-Jovin D.J. Cytometry A, 2013, 83 794; [2] Valley C.C., Arndt-Jovin D.J., Karedla D.N, Steinkamp M.P., Chizhik A.I., Hlavacek W.S., Wilson B.S., Lidke K.A., and Lidke D.S., Molec. Biol. Cell, 2015, doi: mbc.E15-05-0269. [3] Kovács T., Nagy P, et al., manuscript in preparation.

Figure1.FRETbetweenEGFRandplasmamembrane(seetext).A)CHOcellslabeledwithACPdonor;ROIsegmentationforanalysis(panelB).B)Fluorescentlifetimesfrom3pointmeasurementsofdonoronlyincellsandinPBS(redbar).C)Donor

quenchinginpresenceofNR12Sacceptor.

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B-P41

99

Multiple FRET in molecular ensembles: site-photoselection effects Alexander P. Demchenko

Palladin Institute of Biochemistry, Kiev 01030, Ukraine [email protected]

FRET is often considered as the process occurring between two fluorophores – the donor and the acceptor. Meantime in dye-doped nanoparticles, nanocomposites, multi-Trp proteins and highly dye-loaded living cells the situation can be different. The energy exchange between the donors as well as between the acceptors may become efficient. Even if these populations are composed of identical fluorophores, the so called Red Edge effects can be observed. In essence, they are the effects of photoselection due to variations of distances, orientations and interaction energies in the ground and excited states. In rigid and highly viscous fluorophore environments, i.e. in the conditions, where the dynamics of averaging the fluorophore environments is slower than the emission, a number of spectacular spectroscopic effects can be observed. Particularly:

• With the increase of dye concentration when the average distance between dyemolecules becomes shorter that the Förster Ro the fluorescence spectra shiftprogressively to longer wavelengths. This shift stops to be observed at the red-edgeexcitation.

• In the presence of FRET acceptor as the quencher in a system with high donorconcentration the fluorescence may re-appear at the red-edge excitation.

• Due to inhomogeneous broadening, thepositions of excitation wavelength maximadepend on emission wavelength and this dependence is manifested in the case ofFRET showing the dye concentration dependence. This dependence disappears at thedetection of emission at the blue edge.

• The fluorescence depolarizing effect of FRET is removed at the red edge of excitationor the blue edge of emission. This allows distinguishing between the cases ofdepolarization due to fluorophore rotation and due to FRET.

The understanding of these effects and practical operation with them is particularly important in the development of sensing and imaging technologies, in which fluorescent nanoparticles are made of dyes or doped with concentrated amounts of dyes. The consistent model that explains all these phenomena will be presented being illustrated by our own and literature data [1-3].

[1] A.P. Demchenko. “Red-Edge effects: thirty years of exploration (review)”. Luminescence, 17, 19-42, (2002). [2] A.P. Demchenko. “Site-selective Red-Edge effects”. Meth. Enzymol. 450, 59-78, (2008). [3] A.P. Demchenko. “Fluorescent nanocomposites”. In: Introduction to fluorescence sensing, 2-nd ed. Springer Verlag, pp. 263-300, (2015).

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A specialized Sampling Engine for the Bayesian Inference Software Fast-NPSEilert T, Drecksler F, Beckers M and Michaelis J

Ulm University, Institue of Biophysics, Albert-Einstein Allee 11, 89081 Ulm, Germany

[email protected]

Single-molecule spectroscopy can be used to measure the efficiency of fluorescence resonance energy transfer between dyes. Fast-NPS is a software package that aims to infer information about the structure of macromolecular complexes via FRET-measurements between dyes with a Bayesian model [1,2]. In Bayesian probability theory the posterior is the product of likelihood and prior. In Fast-NPS the likelihood is a multivariate normal distribution centered at the measured FRET-efficiencies. In the classic model the prior for the dye orientations constitutes the uniform distribution over the hemisphere. The dye positions are uniformly distributed in volumes discretized via a collection of boxes established with the information of e.g. X-ray structures. Thus, the posterior is a high-dimensional, discontinuous probability measure with possibly many separated cavities. Here, we present problems and their solutions Fast-NPS has to cope with. In order to extract the structural information encapsulated within the posterior, we implemented a Gibbs sampler, which draws from the conditional distributions with a Metropolis algorithm. It is known that the posterior oftentimes exhibits multimodality induced by likelihood and/or prior. Thus, we equipped the sampling engine with a parallel tempering scheme. We discuss further how alternate models influence the precision and consistency of the structural inference [3]. Finally a method is presented, that introduces spatial dependencies between dyes attached at complex ligands.

Fig. 1. (A) NPS is based on data from different FRET-measurements between dyes attached to different positions on a macromolecule. The dye molecules may reside anywhere within their accessible volumes (AVs). (B) Classic model. Within the AV, the dye occupies one unknown

position. At this position, the dye molecule can rotate within a cone. (C) Iso model. As in classic, but dynamical averaging over all orientations is assumed. (D) Meanpos-iso model. The dye molecule is

free to rotate, but the center of the AV volume is used in the analysis. (E) Var-meanpos-iso model. As in meanpos-iso, but the position is variable. (F) Var-meanpos model. As in classic, but the dye

dynamically switches between different positions.

[1] Muschielok A, Andrecka J, Jawhari A, Brückner F, Cramer P, Michaelis J. A nano-positioning system for macromolecular structural analysis. Nat Methods. 5, 965, 2008. [2] Muschielok A, Michaelis J. Application of the nano-positioning system to the analysis of fluorescence resonance energy transfer networks. J Phys Chem B. 115, 11927, 2011. [3] Beckers M, Drechsler F, Eilert T, Nagy J, Michaelis J. FD2015-Single Molecule Microscopy:Quantitative structural information from single-molecule FRET. Faraday Discuss. 2015

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N-FRET in Assemblies and Clusters by Steady State Anisotropy: Theory and Practice Zahra Zolmajd-Haghighi,* Zahra Gholami,** and Quentin S. Hanley**

* Institute of Biochemistry and Biophysics, University of Tehran, Tehran, Iran

** School of Science and Technology, Nottingham Trent University, Nottingham, UK

[email protected]

The study of homo-FRET between multiple fluorophores was pioneered by Weber and Daniel [1] followed later by a distance dependent treatment from Runnels and Scarlatta [2]. The theory presented in these pioneering studies allows the assessment of cluster size subject to a number of assumptions. Importantly, the distance (R) between fluorophores must be less than 0.8 R0, the fluorophores in proximity must not quench or enhance, and they must behave as integer countable species. A wide range of theoretically and empirically understood behaviour will cause these assumptions to be broken. Examples include non-random orientation and quenching. Further, many real systems in biology do not have R < 0.8 R0 and in systems involving fluorescent proteins it is difficult to achieve.

We have constructed a range of clustered and assembled systems to investigate the characteristics of N-FRET on anisotropy [3-5]. Resulting N-FRET ranged from 2 fluorophores up to 24 over a range of distances. Steady state anisotropy and intensity data collected from the clusters and assemblies were used to test and refine existing theories. These systems have exhibited behaviour consistently breaking widely used assumptions. From this we have reached a number of conclusions: i) observing titrations producing assemblies indicate some assemblies form stochastically while others form in step-wise fashion and observing anisotropy behaviour allows this to be determined; ii) quenching affects apparent cluster size and measurement of both intensity and anisotropy allows this to be corrected; iii) enumeration of large clusters is favoured by the use of incompletely labelled units made up of fluorophores that exhibit high amounts of distance dependent quenching.

More recent work on a range of labelled DNA assemblies have allowed us to determine that: iv) the distance dependent behaviour described by Runnels and Scarlatta is affected byquenching and if this accounted for both distance and cluster size may be assessed when R > 0.8 R0; and v) applying known models yields behaviour in which the measured number n in an N-FRET cluster differs from the known N suggesting non-integral behaviour relative to existing models.

We will present data providing an overview of homo-N-FRET systems including: the correspondence between data and theory, adjustments to that theory allowing improved assessment of cluster sizes over a range of distances, approaches to achieving improvements in practice, and opportunities for further development.

[1] Weber, G., and E. Daniel. 1966. Biochemistry 5:1900-1907. [2] Runnels, L. W., and S. F. Scarlata. 1995. Biophysical Journal 69:1569-1583. [3] Zolmajd-Haghighi Z. and Q. S. Hanley. Biophys. J. 2014 106:1457-1466. [4] Gholami, Z. and Q. Hanley. Bioconjug. Chem. 2014 25:1820-1828. [5] Gholami Z., L. Brunsveld, and Q. Hanley. Bioconjug. Chem. 2013 24:1378-1386.

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Förster Resonant Energy Transfer (FRET) in Orthogonally Arranged Chromophores.

Limitations of the Theory

H. Langhals*, A. J. Esterbauer*, D. Dietl*, I. Pugliesi**, E. Riedle**, R. de Vivie-Riedle*

and P. Kölle*

* LMU University of Munoch, Department of Chemistry,

Butenandtstr. 13, D-81377 Munich, Germany

** LMU University of Munich, Lehrstuhl für BioMolekulare Optik,

Oettingenstrasse 67, D-80538 Munich, Germany

[email protected]

We investigated the ultrafast resonant energy transfer of a perylene bisimide dyad by pump-

probe spectroscopy, chemical variation and calculations. This dyad undergoes transfer with

near unit quantum efficiency although the transition dipole moments of donor and acceptor

are in a perfectly orthogonal arrangement to each other in the equilibrium geometry.

According to the point dipole approximation used in Förster theory no energy transfer should

occur. Experimentally we do, however, find an ultrafast transfer time of 9.4 ps.

N

O

OO

N

O

N

N OO

O N O

O

O

hν

hν‘FRET

9.4 ps

N

O

OO

N

O

N

N OO

O N O

O

O

hν

hν‘FRET

9.4 ps

Fig. 1. Dyad with fast FRET and orthogonal transition moments (left). Formulas according to

Förster’s theory (right).

With the transition density cube approach we show that in the orthogonal arrangement the

Coulombic interactions do not contribute to the electronic coupling. Through the change of

the spacer both in length and chemical character we can clearly exclude any Dexter type

energy transfer. The temperature effects on the FRET rate demonstrate that energy transfer is

enabled through low frequency ground state vibrations, which break the orthogonal

arrangement of the transition dipole moments. The dyads presented here therefore are a first

example that shows with extreme clarity the decisive role molecular dynamics plays in energy

transfer processes [1-3].

[1] P. Kölle, I. Pugliesi, H. Langhals, R. Wilcken, A. Esterbauer, R. de Vivie-Riedle, E. Riedle "Hole-transfer

induced energy transfer in perylene diimide dyads with a donor-spacer-acceptor motif", Phys. Chem. Chem.

Phys. 2015, 17, 25061-25072; DOI: 10.1039/c5cp02981c.

[2] H. Langhals, A. J. Esterbauer, A. Walter, E. Riedle, I. Pugliesi "Förster resonant energy transfer in

orthogonally arranged chromophores", J. Am. Chem. Soc. 2010, 132, 16777-16782.

[3] H. Langhals, S. Poxleitner, O. Krotz, T. Pust, A. Walter "FRET in orthogonally arranged chromophores",

Eur. J. Org. Chem. 2008, 4559-4562.

( ) ( ) ( )ˆ ˆˆ ˆ ˆ ˆ3κ = × − × × ×D A D DA DA Aμ μ μ R R μ

κ = 0, Φ ≈100%; ??? !!

65

2

128

)10(ln1000

DADA

DDAFRET

N

ΦJk

R××××

××××=

τπ

κ

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Understanding and Reducing the Error in the Evaluation of Intensity-based Microscopic FRET Experiments in the Presence of Low Signal-to-noise Ratio

Peter Nagy*, Ágnes Szabó*,**, Tímea Váradi*, Tamás Kovács*, Gyula Batta*, János Szöllősi*,**

* Department of Biophysics and Cell Biology, University of Debrecen, Debrecen, Hungary

** MTA-DE Cell Biology and Signaling Research Group, University of Debrecen, Debrecen, Hungary

[email protected] (PN)

Intensity-based measurement of FRET in microscopy remains one of the most widely used methods for assessing protein clustering in living cells. In this approach samples labeled with or expressing donor-tagged and acceptor-tagged targets of interest are measured in the donor, FRET and acceptor channels, and the FRET efficiency is evaluated after the determination of spectral overspill factors. Errors in the measurement of the intensities propagate into the FRET values resulting in significant widening of FRET histograms and potentially nonsense FRET values depending on the signal-to-noise ratio. The signal-to-noise ratio is often artificially increased by overexpressing the targets of interest generating biological artifacts. Measurement of FRET in the absence of overexpression of targets is made difficult by the lack of understanding and appreciation of the effect of error spreading from the measured intensities into the FRET and spectral overspill factors.

We generated intensities doped with Poissonian noise in the donor, FRET and acceptor channels simulating different assumed FRET efficiencies and determined the FRET efficiency using pixelwise calculation, maximum likelihood estimation (MLE) and from summed intensities. In the MLE approach intensities measured in the donor, FRET and acceptor channels were all assumed to follow Poisson statistics and the joint probability of photon numbers detected in the three channels was derived. The FRET efficiency generating the measured photon numbers with the largest likelihood was determined iteratively providing a single FRET value for all pixels in the calculation [1]. Pixelwise calculation turned out to be the least reliable estimate for the simulated FRET efficiencies under conditions of low fluorescence intensity, while calculation from summed intensities and MLE were robust. A similar conclusion was reached for the determination of spectral overspill factors. The simulations also revealed that the reported intensity dependence of spectral overspill factors can at least partially be attributed to the statistical nature of photon detection.

Given the importance of the error spread-induced widening of FRET histograms we developed a Matlab-based evaluation tool for analyzing intensity-based FRET experiments since previously available programs provided only limited functionality to observe the error in the FRET distribution related to low signal-to-noise ratio. Evaluation of FRET in cells labeled with donor- and acceptor-tagged antibodies confirmed the conclusions reached in the simulations. MLE turned out to be superior for determining low FRET efficiencies. In conclusion, careful deliberations must be made before analyzing intensity-based FRET experiments in the presence of low signal-to-noise ratio.

[1] Nagy P, Szabó Á, Váradi T, Kovács T, Batta G, Szöllősi J “Maximum likelihood estimation of FRET efficiency and its implications for distortions in pixelwise calculation of FRET in microscopy”, Cytometry A, 85A., 942/952, (2014).

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Quantitative Protein Kinetics From smFRET Time TracesSonja Schmid, Markus Goetz and Thorsten Hugel

Institute of Physical Chemistry, University of Freiburg, Germany [email protected]

Single molecule FRET (smFRET) time traces provide unique information on the kinetic state sequence of the system under study. Notably many fundamental thermodynamic, as well as, kinetic questions cannot be addressed based on steady-state histograms alone, e.g. energy coupling and resulting directed flow.

These key aspects of smFRET time traces are not considered in most published studies. This is because of experimental limitations associated with surface immobilized smFRET, such as photo-bleaching, inter-fluorophore variation and the limited time window. Therefore, analysis tools beyond dwell-time histograms are required to quantify smFRET kinetics.

Here we present a new maximum likelihood-based approach custom-tailored for experimental smFRET data. Our analysis tool handles multiple input traces with all their experimental short-comings, performs unbiased model selection, and includes multiple complementary evaluation procedures. We demonstrate the robustness of rate determination and model selection on the basis of experimental and simulated data. Furthermore, this analysis uncovers the energy landscape of conformational states (see Fig. 1), as well as, out of equilibrium effects.

Fig. 1. Energy landscape with conformational states of the heat-shock protein Hsp90 deduced by smFRET time traces.

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ORGANIC PHOTOLUMINESCENT PROBES POSSESSING TRIPLET-SINGLET ENERGY TRANSFER BY FÖRSTER MECHANISM

Asko Uri, Kadri Ligi, Ganesh babu Manoharan, Taavi Ivan, and Erki Enkvist

Institute of Chemistry, University of Tartu, 50411 Tartu, Estonia

[email protected]

We have reported [1] novel protein binding-responsive organic photoluminescent probes [ARC-Lum(Fluo) probes] whose association with a PK leads to green, orange or red luminescence with long (τ = 20-250 µs at room temperature in water solution) decay time upon excitation of the probe with a pulse of near-UV radiation (Fig.1). An ARC-Lum(Fluo) tandem probe in addition to a sulfur- or selenium-comprising heteroaromatic phosphor that specifically binds to the ATP-pocket of the target protein kinase (PK), incorporates an orange or red fluorescent dye whose absorption spectrum overlaps with phosphorescence emission spectrum of the phosphor.

Efficient Förster-type resonant energy transfer (FRET) from the excited triplet electron state of the low-QY donor phosphor (3D*) to the acceptor fluorophore A (3D* + 1A → 1D + 1A*) [2,3] leads to emission of light from the fluorescent dye. If the absorption spectrum of the acceptor fluorescent dye has good overlap with the phosphorescence emission spectrum of the donor phosphor and the dye possesses high brightness, substantial (20 – 500-fold) sensitization of the phosphorescence emission signal takes place [4]. Long lifetime of the emission mediated by the short-lifetime fluorescent dye is due to slow prohibited resonant energy transfer from 3D* to the acceptor dye. As the emission is mediated by the fluorescent dye, the delayed (50 μs) photoluminescence spectrum of the tandem dye coincides with fluorescence emission spectrum of the attached dye. We have used ARC-Lum(Fluo) probes for analysis of PKs in biological samples, screening of inhibitors in biochemical assays, and for mapping and monitoring of PKs in living cells using time-gated luminescence microscopy [5].

Fig. 1. Organic photoluminescent ARC-Lum(Fluo) probes with microsecond-scale decay time [6].

1. Enkvist E., Vaasa A., Kasari M., Kriisa M., Ivan T., Ligi K., Raidaru G., Uri A. ACS Chem Biol. 6, 1052 (2011).2. Bennett R.G., Schwenker R.P., Kellogg R.E. J. Chem. Phys. 41, 3040 (1964).3. Galleyt W.C., and Stryert L. Biochemistry 8, 1831 (1969).4. Maliwal B.P., Gryczynski Z., Lakowicz J.R. Anal Chem. 73, 4277 (2001).5. Vaasa A., Ligi K., Mohandessi S., Enkvist E., Uri A., Miller L.W. Chem Commun (Camb). 48, 8595 (2012)6. Kasari M., Ligi K., Williams J.A., Vaasa A., Enkvist E., Viht K., Pålsson L.O., Uri A. Biochim Biophys Acta1834, 1330 (2013).

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Advanced spectral FRET approaches for quantitative Imaging reveal importance of receptor oligomerization in serotonergic signalling

Andre Zeug*, Volodymyr Cherkas*,** and Evgeni Ponimaskin*

* Hannover Medical School, Hannover Germany ** Bogomoletz Institute of Physiology, Kyiv, Ukraine

[email protected]

Förster Resonant Energy Transfer (FRET) is often used in quantitative molecular microscopy, which aims at investigating the interaction of proteins at distances beyond diffraction-limited resolution. The exact determination of the FRET signals, which are often only fractions of the fluorescence signals commonly, requires high experimental effort. Moreover, the correct interpretation of FRET measurements as well as FRET data-based modelling represents an essential challenge in microscopy and biophysics. Previously, we have developed 'linear unmixing FRET' (lux-FRET) approach [1]–[3] on various spectrally resolving imaging system, covering filter based wide field and TIRF imaging, confocal LSM and multi-foci systems such as Yokogawa Spinning Disk microscope. For that, we developed advanced acquisition and calibration strategies to acquire all necessary information with a minimum on experimental effort. Essential for successful application for lux-FRET in confocal systems is the spatial calibration for that we developed our eSIP approach [4]. We further demonstrate the challenging aspects of pixel based analysis, which leads to improper evaluation over ROI-based analysis. Finally, we extended our lux-FRET strategy to overcome the so fare required prior knowledge of fluorescence quantum yields of the FRET fluorophores in standard lux-FRET. We demonstrate the potential of the quantitative lux-FRET approach by elucidating the complexity of serotonergic downstream signalling. In this study we combined lux-FRET with the simultaneous application of FRET based biosensors [5]. Using this strategy, we analyzed the oligomerization between 5-HT1A and 5-HT7 receptor and its biological impact on the cAMP pathway in molecular detail at quantitative level in living cells. On basis of these data we developed a biophysical model of receptor-receptor interaction [6] and its correlation to cAMP downstream signaling in both recombinant system as well as in intact neurons to reveal the complex signaling characteristics of endogenous serotonin receptors.

References: [1] J. Wlodarczyk, A. Woehler, F. Kobe, E. Ponimaskin, A. Zeug, and E. Neher, “Analysis of FRET Signals in

the Presence of Free Donors and Acceptors,” Biophys. J., vol. 94, no. 3, pp. 986–1000, Feb. 2008. [2] S. Prasad, A. Zeug, and E. Ponimaskin, “Chapter 14 - Analysis of Receptor–Receptor Interaction by

Combined Application of FRET and Microscopy,” in Methods in Cell Biology, vol. 117, P. M. Conn, Ed. Academic Press, 2013, pp. 243–265.

[3] A. Zeug, A. Woehler, E. Neher, and E. G. Ponimaskin, “Quantitative Intensity-Based FRET Approaches—A Comparative Snapshot,” Biophys. J., vol. 103, no. 9, pp. 1821–1827, Nov. 2012.

[4] M. Butzlaff, A. Weigel, E. Ponimaskin, and A. Zeug, “eSIP: A Novel Solution-Based Sectioned Image Property Approach for Microscope Calibration,” PLoS ONE, vol. 10, no. 8, p. e0134980, Aug. 2015.

[5] P. S. Salonikidis, M. Niebert, T. Ullrich, G. Bao, A. Zeug, and D. W. Richter, “An Ion-insensitive cAMP Biosensor for Long Term Quantitative Ratiometric Fluorescence Resonance Energy Transfer (FRET) Measurements under Variable Physiological Conditions,” J. Biol. Chem., vol. 286, no. 26, pp. 23419–23431, Jul. 2011.

[6] U. Renner, A. Zeug, A. Woehler, M. Niebert, A. Dityatev, G. Dityateva, N. Gorinski, D. Guseva, D. Abdel-Galil, M. Fröhlich, F. Döring, E. Wischmeyer, D. W. Richter, E. Neher, and E. G. Ponimaskin, “Heterodimerization of serotonin receptors 5-HT1A and 5-HT7 differentially regulates receptor signalling and trafficking,” J. Cell Sci., vol. 125, no. 10, pp. 2486–2499, May 2012.

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Establishment of Sensitive Probes for Radiation-Induced Chromatin Decondensation Using Fluorescence Lifetime Imaging Microscopy

Elham Abdollahi1, Gisela Taucher-Scholz1, Marco Durante1, 2, Burkhard Jakob1

1 Department of Biophysics, GSI Helmholtz Center for Heavy Ion Research, Planckstrasse 1, 64291 Darmstadt, 2 Trento Institute for fundamental physics and applications, Via Sommarive1438123 Trento, Italy

[email protected]

Fig.1. Decondensation of heterochromatic DNA after charged particle irradiation. (a) Intensity image of a NIH3T3 nucleus where a chromocenter was traversed by a low energy uranium ion. DNA

staining with 1µM Hoechst 34580 after fixation. (b) Corresponding color coded lifetime image of Hoechst 34580. Insets in (a) and (b) represent the magnified hit chromocenter. (red rectangular) (c) Intenstiy image of the repair protein XRCC1 marking the ion traversal site (immunostaining; Alexa

514).

The repressive environment of heterochromatin makes the processing of heterochromatic double-strand breaks (DSBs) and the maintenance of genomic stability a challenging task for the cellular repair system after a radiation insult. Recently, we were able to show that ion irradiation induces a local decondensation of heterochromatin in murine chromocenters at the sites of ion traversal. This decondensation is accompanied by a relocation of the induced double-strand breaks (DSB) to the adjacent euchromatin, which might be a requirement for efficient DNA repair [1]. As it proved difficult to reveal minor changes in radiation-induced chromatin decondensation in intensity based measurements of living cells, we started to exploit Fluorescence Lifetime Imaging Microscopy (FLIM) as a promising alternative technique. For this purpose, a confocal FLIM scanner was coupled to the GSI accelerator beamline microscope [2]. In addition, several DNA dyes were screened on their suitability to serve as potential chromatin compaction probes. The indications of a pronounced compaction dependent lifetime contrast as well as sufficient photostability were the most important characteristics of the promising candidates. By enzymatic/chemical changes of chromatin structure, promising probes like Syto13 or Hoechst 34580 were benchmarked against the known FRET-sensor pair H2B-GFP/-RFP [3] expressed in the same cell line. In addition, we were able to present first evidence of detecting radiation induced heterochromatic chromatin decondensation in ion irradiated murine chromocenters using FLIM [2]. [1] B. Jakob, J. Splinter, S. Conrad, K. Voss, D. Zink, M. Durante, M. Löbrich and G. Taucher-Scholz, NAR, 39, 15, 6489-6499 (2011).

[2] E. Abdollahi, G. Taucher-Scholz, Marco Durante, B.jakob, NIMB, 365, 626-630 (2015)

[3] D. Llѐres, J. James, S. Swift, D.G. Norman, A.L. Lamoned, Cell Biol. 187, 4, 481-496 (2009)

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Quantitative three-color FRET for the study of coordinated intramolecular motion

Ganesh Agam1, Anders Barth1, Lena Voith von Voithenberg1 and Don C. Lamb1

Single‐pair FRET (spFRET) has become a standard tool to monitor conformational changes ofbiomolecules in vitro. In solution‐based burst analysis, the burst of photons emanating frommolecules diffusing through the confocal volume gives access to the underlying distribution of the FRET efficiencies. Multiple intramolecular distances can be explored in individual experiments, however information about the coordination of motion is not available. A promising approach to address the coordination of motion within complex biological systems is multicolor FRET, allowing multiple distances to be monitored simultaneously. While several three‐ and four‐color FRETstudies have already been performed, quantitative analysis of multicolor single molecule dataremains challenging. In three‐color FRET, the contribution of shot noise is increased due to thedistribution of the signal into three channels, the FRET efficiency calculations are more complicated and the experimental imperfections more significant. Multicolor FRET is thus still limited to the extraction of average values due to the complexity of the analysis, losing the additional information encoded in the distribution of FRET efficiencies. For spFRET, a statistical description of photon emission (Photon Distribution Analysis, PDA) allows the shot‐noise broadening of the FRETefficiency histogram to be distinguished from physically relevant heterogeneities of the sample,e.g. due to the existence of conformational substates.We have developed an extension of PDA to three‐color FRET systems, granting access to thethree‐dimensional distance distribution and allowing the question of coordinated motion withinsingle biomolecules to be directly addressed. The new method of threecolor PDA is used to investigate allosteric effects in heat‐shock proteins (Hsp). Hsp70 consists of a nucleotide‐bindingdomain and a substrate‐binding domain (NBD/SBD), which are connected by a small linker. Usingthree‐color FRET, we seek to answer the question of how a change in the nucleotide state of theNBD effects the conformation of the SBD.

1Chemistry Department, NIM, CIPSM, CeNS,Ludwig-Maximilians-University Munich

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FRET-FLIM as a tool to investigate interactions of hyaluronic acid with the skin and implications for the dermal delivery of biomacromolecules

Alexander Boreham*, Robert Brodwolf*, Madeleine Witting**, Wolfgang Friess**, Sarah Hedtrich*** and Ulrike Alexiev*

* Department of Physics, Institute of Experimental Physics, Freie Universität Berlin, Berlin,Germany, ** Department of Pharmaceutical Sciences, Ludwig-Maximilians-Universität,Munich, Germany, *** Institute for Pharmaceutical Sciences, Freie Universität Berlin,

Berlin, Germany [email protected]

We investigated the skin absorption of hyaluronic acid (HA) as well as the interactions of HA

with proteins by combining fluorescence lifetime imaging microscopy (FLIM) and Förster

resonance energy transfer (FRET). Besides moisturizing the skin and improving wound

healing, HA facilitates the skin absorption of drugs. In particular, FRET-FLIM was used to

characterize the ability of HA hydrogels as a drug delivery system for labile

biomacromolecules, such as proteins, in topical treatment. We used bovine serum albumin

(BSA) as a model protein. HA and BSA were fluorescently labeled with N-methylisatoic

anhydride (HA-MANT) and rhodamine B isothiocyanate (BSA-RhoB), respectively. We

detected dermal absorption of HA-MANT in cryosections of skin, characterized by an

intraepidermal accumulation. Interestingly, a colocalization with the model protein BSA

resulted in altered skin absorption due to intense interactions between HA and the protein that

were verified by FRET. Moreover, FLIM measurements revealed a formulation dependent

epidermal delivery of proteins. Significantly higher skin penetration of BSA-RhoB was

observed when applying HA fragments; 5 kDa HA being superior. In summary, our FRET-

FLIM data substantiate the hypothesis of a dermal drug co-transport [1].

Fig. 1. FRET-FLIM as a tool to investigate HA as a dermal delivery system for biomacromolecules

[1] Witting et al. “Interactions of hyaluronic acid with the skin and implications for the dermal delivery of biomacromolecules”, Mol. Pharmaceutics, 12, 1391-1401, (2015).

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NexttoCENP-AalsotheH3variantsH3.1andH3.3havedefinedcell-cycledependentlocationsincentromericchromatin

StephanDiekmann,ChristianHoischen,PaulKamweruandChristianAbendroth

Leibniz-InstitutfürAlternsforschung(FLI),Beutenbergstr.11,07745Jena,Germany

Contactauthor:[email protected]

Abstract

Kinetochore proteins assemble onto centromeric chromatin and regulate DNAsegregation during cell division. The inner kinetochore proteins bind centromeresduring the whole cell cycle while most outer kinetochore proteins assemble atcentromeresduringmitosis,connectingthecomplextomicrotubules.Thecentromere-kinetochorecomplexcontainsspecificnucleosomesandnucleosomalparticles:CENP-Areplaces canonicalH3 in centromeric tetrameric and octameric nucleosomes, definingcentromeric chromatin. Next to CENP-A, the Constitutive Centromere-AssociatedNetwork (CCAN)multi-protein complex settleswhich contains CENP-T/W/S/X. Thesefourproteinsmayformanucleosomalparticleatcentromeres.During S-phase after replication, the histone variants H3.1 and H3.3 are deposited at centromeres as ‘placeholders’ between CENP-As, pending deposition of new CENP-A in late mitosis and early G1. After replication we found by cell-cycle dependent in vivo FRET that H3.3, but not H3.1 or H3.2, is loaded next to CENP-A. Here, FRET studies are essential since currently, these three H3 variants cannot be distinguished from each other by immuno-staining. We found that the H3.3 chaperone DAXX is located in close proximity to H3.3. When DAXX or other H3.3 chaperones (i.e. HIRA, ATRX) were knocked-down in human HEp-2 cells or knocked-out in mouse cell lines, H3.3 remained in close proximity to CENP-A at centromeres. However, when pairs of H3.3 chaperones were knocked-down, H3.1 was found close to CENP-A. Furthermore, in H3.3 knock-out mouse cells, we found H3.1 next to CENP-A. When however expressing H3.3 in this H3.3 knock-out cell line, again H3.3 is preferred and placed next to CENP-A, clearly indicating the strong preference of H3.3 over other H3 variants. Since the H3.3 chaperone DAXX binds to CENP-C, we speculate that CENP-C directs the loading of H3.3 next to CENP-A via H3.3 chaperones. Also by cell-cycle dependent in vivo FRET we found the CENP-T C-terminus and theCENP-S termini next to histone H3.1 but not to CENP-A, suggesting that the CCANbridgesCENP-A-andH3-containingnucleosomes.ThisproximitybetweenCENP-TandH3 is specific forH3.1butneither for theH3.1mutantsH3.1C96A andH3.1C110Anor forH3.2 or H3.3. Thus we found CCAN proteins next to a specifically H3.1-containingcentromericnucleosome.During particular periods of the cell cycle, we detected H3.1 and H3.3 at specificlocationsincentromericchromatin.WhiletheH3variantCENP-Ashapesacentromere-specificnucleosomeanddefinescentromerelocationonthechromosomes,twootherH3variantsadoptdefinedlocal,andprobablyalsofunctional,positions.WespeculatethatthreeH3variantsdefine a cell-cycledependent local centromeric chromatin structurewhichcontributestocentromerefunction.

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Using FLIM-FRET microscopy to study protein-specific glycosylation in living cellsFranziska Doll*, Annette Buntz* and Andreas Zumbusch*

* University of Konstanz, Department of Chemistry and Konstanz Research School ChemicalBiology (KoRS-CB), Universitätsstraße 10, 78464 Konstanz, Germany

[email protected]

The post-translational modification of proteins with β-linked N-acetylglucosamine, termed O-GlcNAcylation, is widespread and plays an important role in regulating the structure and function of proteins. Dysregulation of protein O-GlcNAcylation is related to severe ailments such as type 2 diabetes and Alzheimer’s disease [1]. Thus, imaging O-GlcNAcylation of specific proteins inside living cells is of great interest. To achieve this, we established a novel approach by combining metabolic glycoengineering with Fluorescence Lifetime Imaging-Förster Resonance Energy Transfer (FLIM-FRET) microscopy [2]. Thereby, EGFP is genetically fused to a protein of interest and a recently developed N-acetylglucosamine derivative bearing a cyclopropene-tag (Ac4GlcNCyoc) [3] is incorporated into the cellular glycome in order to target protein O-GlcNAcylation. This sugar-derivative reacts in a second step with a dye-tetrazine conjugate in a bioorthogonal reaction inside living cells. The glycosylation of a specific protein then leads to FRET between the EGFP donor and the acceptor dye. The advantage is that FRET can be detected with high contrast even in presence of a large excess of acceptor fluorophores via a reduction of the fluorescence lifetime of EGFP (see Fig. 1). We successfully employed this strategy to visualize the glycosylation of the proteins O-GlcNAc-transferase, the forkhead transcription factor Foxo1, the tumor suppressor p53 and the serine/threonine kinase 1 Akt1. Our approach provides an innovative tool for a better understanding of how glycosylation regulates the localization and function of intracellular proteins which is generally applicable, given that the EGFP tag can be fused in close proximity to the glycosylation site of the protein of interest.

Fig. 1. Experimental strategy.

[1] Lefebvre, T. et al. “Dysregulation of the nutrient/stress sensor O-GlcNAcylation is involved in the etiology of cardiovascular disorders, type-2 diabetes and Alzheimer's disease”, Biochim. Biophys. Acta Gen. Subj., 1800, 67, (2010). [2] Doll, F. et al. “Visualization of Protein-Specific Glycosylation inside Living Cells”, Ange. Chem. Int. Ed., DOI: 10.1002/anie.201503183, (2016); Ange. Chem., DOI: 10.1002/ange.201503183, (2016). [3] Späte, A.-K. et al. “Expanding the scope of cyclopropene reporters for the detection of metabolically engineered glycoproteins by Diels–Alder reactions“, Beilstein J. Org. Chem., 10, 2235, (2014).

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Mapping biochemical networks in single living cells by FLIM and multiplexed FRET: a Systems Microscopy approach

Haas K, Fries MW, Venkitaraman AR* and Esposito A*

The Medical Research Council Cancer Unit at the University of Cambridge,

Hutchison/MRC Research Centre, Box 197, Biomedical Campus

Cambridge, CB2 0XZ, United Kingdom

*shared senior authors

[email protected]

Biochemical networks cooperate to determine cellular decisions and maintain cellular functions. However, techniques capable to map several biochemical reactions within a single living cell are rather limited. Therefore, we have developed a biochemical sensing platform based on novel imaging technologies and a family of novel FRET pairs that permit the simultaneous quantification of three biochemical reactions with the low invasiveness and high spatiotemporal resolution characteristic of fluorescence microscopy. To maximize our biochemical resolution we develop advanced FLIM imaging modality and data processing algorithms, Hyper-Dimensional Imaging Microscopy (HDIM), fast spectral FLIM and fast multi-colour FLIM, imaging techniques that can attain biochemical multiplexing and high biochemical resolution [1-2].

[1] Popleteeva et al, “Fast and simple spectral FLIM for biochemical and medical imaging “, Opt. Express, Vol. 23, Issue 18, pp. 23511-23525 (2015)

[2] A. Esposito et al., “Maximizing the Biochemical Resolving Power of Fluorescence Microscopy” PLOS ONE DOI: 10.1371/journal.pone.0077392 (2013)

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Bacterial toxin induced cAMP gradients shape Gap Junction responses.Majoul I *, Eugenia Butkevich** and Rainer Duden*

* Institute of Biology, Centre for Structural and Cell Biology in Medicine, University ofLübeck, Ratzeburger Allee 160, D-23562 Lübeck, Germany

** Drittes Physikalisches Institut, Biophysik, Georg-August-Universität, D-37077 Göttingen, Germany

[email protected]

Using super-resolution Bessel beam plane illumination microscopy and FRET analyses of fluorescent protein-tagged connexin isoforms we showed the formation of CDRs (Connexin Depleted-regions) inside the Gap Junction plaques which are induced by AB5 toxins. CDR-responses of Gap Junction (GJ) plaques were first described by us in Majoul et al., 2013; PNAS (1). In-depth study of cellular CDR-responses revealed further details of this process. Interestingly, bacterial toxins are not bound directly to the GJ plaques, but instead upon touching the glycosphingolipid receptors at the plasma membrane (PM) generate a wave of tension across the PM that most likely results in a phase-separation condition inside the GJ plaque. CDRs appear in GJ plaques within milliseconds and create microdomains of dense lipids such as cholesterol, in addition to the overall modulation of PM fluidity. Surprisingly, measurable electrical coupling between cells remains unchanged during formation and recovery of CDRs. Due to the fast millisecond-to-second range nature of the process of CDR formation, their temporal evolution is so far only partially characterized. In living cells, GJ plaques are curved and have an intricate 3-D structure that complicates the interpretation of imaging data. FRET analysis revealed CDR-induced changes in the levels of the second messengers cAMP, Ca2+, and IP3 in contacting cells. Obtained data were correlated to the spatial re-organization of GJ plaques, CDR-responses and their temporal dynamics. Analyses of lateral diffusion and mobility of the 4-transmembrane connexin channel clusters inside the GJ plaque revealed their rather heterogeneous composition. FRET data provided an unbiased live cell-based interpretation of second-messenger fluctuations between GJ-coupled cells during CDR-response. Our data confirm broad applicability of live cell FRET in resolving both the gradients of second messengers and the spatio-temporal organization of molecular interactions in gap junction proximity. .

Fig. 1. (left) Gap Junctions channels formed Cx43, (middle) Membrane topology of connexins and assembly of Gap junction channels, (right) CDRs visualized inside the GJ plaques.

[1] Majoul I, Gao L, Betzig E, Onichtchouk D, Butkevich E, Kozlov Y, Bukauskas F,Lippincott-Schwarz J, Duden R. Fast structural responses of gap junction membrane domains to AB5 toxins. . Proc. Natl. Acad. Sci USA 110(44): E4125-4133, (2013)

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Simultaneous imaging of three FRET sensors by fluorescence anisotropy microscopy

,**Agustín Corbat, *, Piotr Liguzinski*Klaus C. Schuermann*, Yvonne Radon

Hernán E. Grecco** and Peter J. Verveer*

* Department of Systemic Cell Biology, Max Planck Institute of Molecular Physiology, Otto-Hahn-Str. 11, 44227 Dortmund, Germany

** Laboratorio de Electrónica Cuántica, Departamento de Física, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires and IFIBA, CONICET, Argentina.

[email protected]

We developed cleavage-based sensors that can be used to follow the activities of three caspases (Caspase 3, 8 and 9) simultaneously in the same cell. The sensors are based on measuring the fluorescence anisotropy of either homo-FRET pairs or fluorescent proteins whose spectra are strongly overlapping, yielding the same effect. In contrast to ensemble based measurements, we observe the activity of caspases directly and can monitor the timing between those. We could see that the onset of caspase activity across cells has a large variance, while the timing of the individual caspases is strongly correlated.

Fig. 1. Sensor is cleaved by Caspase activity removing FRET coupling (A), which we employed with three different flourophore combinations across the spectrum using both homo-FRET and hetero-FRET (B) to monitor apoptosis and the resulting change in fluorescence anisotropy over time on a

single-cell level

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Revealing structural features and affinities of protein complexes in living cells by MFIS-FRET analysis

Qijun Ma1, Marc Somssich3, Suren Felekyan1, Stanislav Kalinin1, Thomas Peulen1, RalfKühnemuth1, Yvonne Stahl3, Rüdiger Simon2,3, Stefanie Weidtkamp-Peters2,

Claus A.M. Seidel1,2

Heinrich Heine University, Düsseldorf, Germany: 1Chair for Molecular Physical Chemistry,2Center for Advanced Imaging, 3Institute for Developmental Genetics.

[email protected]

Due to its sensitivity of distance Förster resonance energy transfer (FRET) has been widely used to investigate the structure and interaction of biomolecules. Multiparameter fluorescence image spectroscopy (MFIS)[1,2] provides particular advantages to FRET imaging because all the fluorescence parameters are monitored simultaneously with picosecond accuracy, which allows for a comprehensive analysis on biological systems. Traditionally, a reduction in average donor lifetime or an increase of average FRET efficiency was used as an indicator for molecular interaction. However, such changes observed in FRET-imaging can have two reasons: (1) the conformational change or (2) change in fraction of FRET-active species. To resolve this ambiguity, we introduce a new sub-ensemble analysis method to directly visualize and quantitatively analyze both factors. Characterization of true FRET efficiency enabled us to detect even subtle FRET variations and provided crucial information about the structural properties of molecular complexes. Furthermore, from determined fraction of FRET-active species, utilizing the intrinsic cell-to-cell variations of protein concentration, we show that dissociation constant (KD) of membrane-receptor interactions [3,4] and protein oligomerisation [5] can be characterized in living cells.

[1] Kudryavtsev, V., Felekyan, S., Woźniak, A. K., König, M., Sandhagen, C., Kühnemuth, R., Seidel C. A.M., Oesterhelt, F.; Multiparameter fluorescence imaging to monitor dynamic systems, Anal. Bioanal. Chem. 387, 71-82 (2007); [2] Weidtkamp-Peters, S., Felekyan, S., Bleckmann, A., Simon, R., Becker, W., Kühnemuth, R., Seidel, C. A. M.; Multiparameter Fluorescence Image Spectroscopy to study molecular interactions. Photochem. Photobiol. Sci., 8, 470-480 (2009). [3] Stahl, Y., Grabowski, S., Bleckmann, A., Kühnemuth, R., Weidtkamp-Peters, S., Pinto, K. G., Kirschner, G. K., Schmid, J. B., Wink, R. H., Hülsewede, A., Felekyan, S., Seidel, C. A. M., Simon, R.; Moderation of Arabidopsis root stemness by CLAVATA1 and ARABIDOPSIS CRINKLY4 receptor kinase complexes. Current Biology 23, 362–371 (2013). [4] Somssich, M., Ma, Q, Weidtkamp-Peters, S., Stahl, Y., Felekyan, S., Bleckmann, A., Seidel, C A. M., Simon, R.; Differential real-time dynamics of peptide ligand dependentreceptor complex formation in planta. Sci. Signal 8, ra76 (2015). [5] Kravets, E.*, Degrandi, D.*, Ma, Q.*, Peulen, T.-O., Klümpers, V. Felekyan, S., Kühnemuth, R., Weidtkamp-Peters, S., Seidel, C. A.M., Pfeffer, K. Guanylate binding proteins (GBPs) directly attack T. gondii via supramolecular complexes. eLife 5, e11479 (2016).

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FRET-FLIM-Imaging: From 2D to 3D Cell Culture Systems Petra Weber, Sarah Schickinger, Thomas Bruns, Michael Wagner and Herbert

Schneckenburger

Institute of Applied Research, Aalen University, Beethovenstr. 1, 73430 Aalen, Germany

[email protected]

In previous experiments we could show that Förster Resonance Energy Transfer (FRET) in combination with Fluorescence Lifetime Imaging (FLIM) seems to be appropriate to assess molecular interactions of proteins, relevant in Alzheimer’s disease [1] but can also in sensing of apoptosis [2]. In the second case energy transfer from an enhanced cyan fluorescent protein (ECFP) to an enhanced yellow fluorescent protein (EYFP) is used for detection of apoptosis in 3-dimensional cell cultures after application of staurosporine. Emission spectra of the donor (ECFP) and acceptor (EYFP) as well as fluorescence lifetime of the donor revealed to be appropriate parameters. Using different methods of fluorescent microscopy, for example Light Sheet Fluorescence Microscopy (LSFM) it is possible to get additional information about inner layers of cellular tumor models. This appears relevant for investigation of pharmaceutical agents to monitor apoptosis.

Hela-Mem-ECFP-DEVD-EYFP Control

Hela-Mem-ECFP-DEVD-EYFP + Staurosporine 1 M, 2.5 h

Fig. 1. Transillumination (A), fluorescence intensity (λ ≥ 515 nm; B) and donor lifetime (450–490 nm; C) of HeLa cervical carcinoma cells transfected with a Mem-ECFP-

DEVD-EYFP encoding vector prior to and subsequent to application of staurosporine (2 µM, 2.5 h). Light sheet based fluorescence microscopy of individual layers of a

multicellular spheroid with 6–10 µm thickness each. Excitation wavelength: 391 nm. Scale bars: 50 µm.

[1] von Arnim CA(1), von Einem B, Weber P, Wagner M, Schwanzar D, Spoelgen R, Strauss WL, Schneckenburger H, “Impact of cholesterol level upon APP and BACE proximity and APP cleavage”, Biochem Biophys Res Commun. 370(2):207-12(2008). [2] Weber P, Schickinger S, Wagner M, Angres B, Bruns T, “Monitoring of apoptosis in 3D cell cultures by FRET and light sheet fluorescence microscopy”, Int J Mol Sci.;16(3):5375-85(2015).

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Toolkit for Multi-conformation Biomolecular Structure Determination by High-precision FRET and Molecular Simulations

Mykola Dimura1, Stanislav Kalinin1, Thomas Peulen1, Holger Gohlke2, Claus A. M.Seidel1

1 Physikalische Chemie II, Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany, [email protected]

2 Institut für Pharmazeutische und Medizinische Chemie, Heinrich-Heine-Universität Düsseldorf, Germany, [email protected]

A comprehensive methodology and a toolkit for FRET-restrained modeling of biomolecules and their complexes is presented. The demonstrated approach enables to recover representative conformations for multiple observed states of the investigated biomolecular system. The toolkit[1] is available from the authors including:

1. FRET-restrained docking and Metropolis Monte Carlo simulation tool for assemblingstructural units with high precision and determining the confidence levels of thegenerated models.

2. FRET-screening too1[1] for assessing an arbitrary set of conformations (from MDtrajectories, crystal structures etc.) with respect to their agreement with FRETmeasurements[2].

3. NMSim[3][4] geometric simulations for extensive sampling of the conformational spaceovercoming time-limitations of MD simulations.

4. FRET-restrained MD simulations for refining conformations generated by coarse-grained sampling, e.g. by NMSim simulations.

5. Experiment planning tool for determining efficient labeling positions and distance pairsfor FRET measurements based on a priori knowledge on conformational changes.

Using these tools, formerly unknown conformations of various biomolecules were determined[5].

An example on T4 Lysozyme with 24 FRET restraints will be presented.

[1] http://www.mpc.hhu.de/software/fps.html [2] Sindbert S., Kalinin S. et al. (2011) Accurate Distance Determination of Nucleic Acids via

Forster Resonance Energy Transfer: lmplications of Dye Linker Length and Rigidity. J. Am. Chem. Soc. 133, 2463-2480

[3] Krüger D. M., Aqeel A., and Gohlke H. (2012) NMSim web server: Integrated approach for normal mode-based geometric simulations of biologically relevant conformational transitions in proteins. Nucl. Acids Res., 40, pp. W310-W316.

[4] http://cpclab.uni-duesseldorf.de/nmsim [5] Kalinin S., Peulen T., Sindbert S. et al. (2012) A toolkit and benchmark study for

FRET- restrained high-precision structural modeling. Nat Methods 9, 1218-1225.

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Homeopathy – Applied Quantumphysics

Lenger Karin, Institute for Scientific Homeopathy, Kaiserstr.28, D-63065 Offenbach

[email protected]

Until now it hasn’t been possible to explain the fundamental principles of homeopathy. It can be clarified since Lenger’s detection of magnetic photons in homeopathic remedies with frequencies in the MHz-region by the two magnetic resonance methods: the Tesla-flatcoil system [1] and delayed luminescence using a modified photomultiplier [2, 4]. Separation of photons from their carrier substance ethanol or saccharose globules took place when the measuring system has a bigger resonating frequency field than the field between carrier substance and photons. The potency levels were measurable by the number of photons [2, 4] and/or by the characteristic size of the magnetic frequency field separating the photons in dependence on the potency level and on the type of potentized substance [1]; the results showed that the necessary strength of the magnetic frequency field increases with increasing potencies. Frequency spectra were obtained by stimulating the remedy by one of their resonance frequencies, and then the frequency spectrum of the other ones can be uptaken. Finally it was possible to identify six unknown remedies with their potency levels by applying delayed luminescence [4]. The obtained results [1,2,4] link to the fundamental principles of homeopathy: the law of similars (Hahnemann 1796): the frequencies of the patient must match the frequencies of the remedy to get in resonance; concerning the production of remedies: a substance has e.g. a frequency I in the visible region of light; its potentized homeopathic remedy prepared by dilution and succussion creates additional frequencies e.g. II, III, IV, V till to the MHz region; provings to get a symptom picture: a healthy person is poisoned by a substance with frequency I and creates more frequencies II, III, IV, V , as well as an ill body and psychological symptoms; a sick person: he/she has symptoms as if he/she was poisoned by the substance with the frequency I and developed the frequencies II, III, IV,V; healing: it means to take the potentized remedy with the frequencies I, II, III, IV, V. to get in resonance with the ill making frequencies of the pathological pathways which have the bigger resonating magnetic field to attract the photons from their carrier substance: This is an application of the same principle of Lenger’s measuring methods [1,2,4]. Resonance takes place: the ill making frequencies of the patient are attenuated or the amplitude is enhanced. Firstly healing takes place on the energy level, then on the biochemical level. Förster told that chemical reactions occur only in excited states. As Lenger had shown [3, 5] healing can be achieved by using the photons of potentized substrates, enzymes, receptors and its inhibitors of the pathological pathways. It is concluded that homeopathy is a regulation therapy healing both by resonance effect: hyperfunction and hypofunction of a pathological pathway and simultaneously psychological symptoms, because body and consciousness are treated as one which is in agreement that homeopathy is a holistic medicine. This confirms F.A.Popp’s quantumphysical theory about life and illness: a normal living organism has an energy level of about 50% excited states by the uptake and emission of photons. Illness means the uptake of too many photons or the emission of too many photons[6,7].The detection of magnetic photons enables an explanation of the principles of homeopathy.

[1] Lenger K.,Homeopathic potencies identified by a new magnetic resonance method. Homeopathy-an energetic medicine. Subtle Energies & Energy Medicine, 15, No. 3:225-243, 2006. [2] Lenger K., Bajpai R.,Drexel M.,Delayed luminescence of high homeopathic potencies on sugar globuli. Homeopathy; 97, 134-140, 2008 [3] Lenger K.,A new biochemical of homeopathic efficacy in patients with chronic diseases.Subtle Energies & Energy Medicine, 19(3), 9-41, 2010. [4] Lenger K., Bajpai R., Spielmann M.,Identification of unknown homeopathic remedies by delayed luminescence. Cell Biochem. Biophys , 68; 321-334, 2014. [5] Lenger K, Lang G.,Photons detected by magnetic resonance are efficious in homeopathy. A critical review. OA Alternative Medicine, 2(1):4, 2014. [6] F.A. Popp Biophotonen-Neue Horizonte in der Medizin, Haug-Verlag, MVS, Stuttgart, 2006. [7] Bischof Marco, Biophotonen, Das Licht in unseren Zellen, Zweitausendeins, Frankfurt a. M.1996

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PIFE meets ALEX: observing both binding and conformational changes in unlabelled proteins

Ploetz E.a,b,#, Lerner E.c,#, Husada F.a, Roelfes M.a, Chung S.c, Hohlbein J.d,

Weiss S.c and Cordes T.a a Molecular Microscopy Research Group, University of Groningen, The Netherlands

b Department of Chemistry, Ludwig-Maximilians-University Munich, Germany c Department of Chemistry and Biochemistry, University of California Los Angeles, USA

d Laboratory of Biophysics, Wageningen UR, Wageningen, The Netherlands # both authors have equally contributed.

[email protected]

Advanced microscopy methods have emerged as important tools for monitoring structural changes in biomolecules. Current implementations of molecular rulers employ photophysical properties such as fluorophore brightness or lifetime to retrieve (real-time) information on the structure of biomolecules. Limitations of such molecular rulers based on FRET, PIFE, PET etc. are their restricted distance range, the need for labeling with fluorescent dyes and most importantly the accessible information, which is at best a quantitative but yields an averaged one-dimensional distance. The latter restriction prohibits monitoring coordinated motion between different protein domains or their interaction with nucleic acids.

In this contribution, we present a hybrid single-molecule technique, dubbed PIFE-FRET, combining two photophysical effects into one powerful two-dimensional assay. We provide a theoretical framework and data analysis routine to allow simultaneous and quantitative read-out of the two different photophysical parameters: donor brightness (PIFE) and energy transfer efficiency (FRET). In proof-of-concept experiments we show that PIFE-FRET can monitor the interaction between unlabeled proteins (BamHI, EcoRV) with freely diffusing doubly labeled dsDNA due to changes in donor brightness via alternating laser excitation. We further investigated the spatial sensitivity of PIFE-FRET for binding of DNA-binding enzymes with respect to the PIFE- and FRET-ruler aspects. We finally outline possible applications of PIFE-FRET both in studies with diffusing and immobilized molecules to unravel the full potential and limitations of the technique for mechanistic investigations for any biomolecular interaction.

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Determination of nanodomain sizes by FRET Radek Šachl*, Mariana Amaro*, Alena Koukalová*, Gokcan Aydogan*, Ilya Mikhalyov**,

Jana Humpolíčková* and Martin Hof*

* J. Heyrovský Institute of Physical Chemistry, Academy of Sciences of the Czech Republic, v.v. i., Dolejškova 3, 18223 Prague 8, Czech Republic

** Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry of the Russian Academy of Science, Moscow, GSP-7, Russian Fed

[email protected]

The techniques achieving the highest resolution only can characterize membrane heterogeneities on the lowest molecular level. In this contribution, we will show that combination of Förster resonance energy transfer (FRET) with Monte Carlo (MC) simulations allows for determination of nanodomain sizes (2-40 nm) and the area fraction these domains occupy in a lipid bilayer. Whereas other techniques start being less efficient at such short distances FRET is most efficient in this region. After explaining principles and limitations of this technique [1,2] and giving clues on how to plan a MC-FRET experiment to achieve the highest resolution in the determination of domain sizes [3] usefulness of this approach will be demonstrated on the examples presented below. i) It will be shown that GM1 co-clusters into 6 nm large pools in pure DOPC bilayer [4]. Thenature of clusters does not change as cholesterol is added but the domains grow in size with sphingomyelin addition. At 10 % of Sph, the domain size reaches 26 nm in radius. The pools are still fluid, disordered and highly dynamic. ii) Domains of similar nature were also revealed in model vesicles consisting of DOPC,sphingomyelin and cholesterol at vesicle compositions being close to the phase separation boundary. Because of the dynamic nature of these heterogeneities, they may rather represent local temporal fluctuations rather than stable long-living phases. Finally, perspectives will be given on how to determine sizes and concentrations of bilayer pores by MC-FRET technique [3].

[1] Šachl R, Humpolíčková J, Štefl M, Johansson LBÅ and Hof M Limitations of electronic energy transfer in the determination of lipid nanodomain sizes, Biophys. J. 101, L60, (2011) [2] Amaro M, Šachl R, Jurkiewicz P, Coutinho A, Prieto M and Hof M Time-Resolved Fluorescence in Lipid Bilayers: Selected Applications and Advantages over Steady State, Biophys. J., 107, 2751–60, (2014) [3] Šachl, R. and Johansson LBÅ Heterogeneous Lipid Distributions in Membranes as Revealed by Electronic Energy Transfer. Reviews in Fluorescence 2015. Eddited by Chris D. Geddes (2016) [4] Šachl R, Amaro M, Aydogan G, Koukalová A, Mikhalyov I I, Boldyrev I A, Humpolíčková J and Hof M On multivalent receptor activity of GM1 in cholesterol containing membranes, Biochim. Biophys. Acta, 1853, 850–7, (2015)

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Studying dynamics of protein interactions by FRET-FRAPJohannes A. Schmid and Bernhard Hochreiter

Center for Physiology and Pharmacology, Med. Univ. Vienna, Austria [email protected]

Macromolecular complexes are not rigid entities but in equilibrium with their single components in a process of repetitive association and dissociation. We developed a technique, which allows measuring the dynamics of complexes in living cells by combining FRET microscopy with FRAP (Fluorescence Recovery After Photobleaching), a method, which is usually used to determine the mobility of molecules. Bleaching of a FRET-donor within a protein complex followed by recording FRAP in the donor- and the raw-FRET-channel revealed a different recovery kinetics for the two channels, whereas the kinetics was identical for non-interacting proteins (Fig. 1). This indicates that information on k-on and k-off rates of the interaction are embedded in the different recovery curves. For a further validation of this technique, we performed it with mutually binding antibodies labelled with FRET-capable fluorophores in vitro and compared the results with stopped-flow fluorometry of the FRET effect in the microsecond range and the mixing of a constant amount of donor with increasing amounts of acceptor. The latter revealed the k-on and k-off values of the binding and implied that the FRET-FRAP microscopy technique is suited to determine k-off values of protein-interactions in living cells and thus the half-lives of protein complexes. To extend our approach to physiological expression levels, we are currently tagging endogenous interaction partners using CRISPR/Cas9 genome editing.

Fig. 1. Example of FRET-FRAP microscopy. A) Image of live cells expressing green- and red fluorescent proteins. The donor fluorophore was bleached in the region indicated by the arrow. A time

series of the fluorescence recovery in the donor and the raw-FRET channel is shown below. B) Interacting proteins exhibit different recovery curves for the raw-FRET and the donor channel.

C) Non-interacting proteins show superimposable donor- and raw-FRET curves (with the latter beinga consequence of spectral bleed-through of donor fluorescence into the raw-FRET channel.

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Determination of the Conformation of LeuT: Using an Integrated Computational and Experimental Approach

Azmat Sohail1, Kumaresan Jayaraman1, Santhoshkannan Venkatesan1, Ulrik Gether2, Claus J.Loland2, Klaus T. Wanner3, Michael Freissmuth1, Harald H. Sitte1, Walter Sandtner1, and

Thomas Stockner1

1) Medical University of Vienna, Center for Physiology and Pharmacology, Institute ofPharmacology, Waehringerstrasse, 13A, 1090 Vienna, Austria

2) University of Copenhagen, Faculty of Health and Medical Sciences Denmark, Departmentof Neuroscience and Pharmacology, Blegdamsvej 3B, 2200 Copenhagen N, Copenhagen,

Denmark 3) Ludwig Maximilians University Munich, Department of Pharmacy, Center of Drug

Research, Butenandtstraße 7, D-81377 Munich, Germany

[email protected]

An essential step of neurotransmission is the fast clearance of neurotransmitters from the synaptic cleft, carried out by pre-synaptic transporters mainly from the SLC6 family. These secondary active transporters couple substrate uptake to the transmembrane sodium gradient. Crystal structures of the bacterial homolog LeuT were solved in three states of the transport cycle: occluded, outward and inward. The recent structure of the dopamine transporter from drosophila melanogaster confirmed the relevance of the bacterial LeuT structures. The inward facing structure of LeuT shows a conformation in which the first helix (TM1A) does not seem to be compatible with the membrane environment. We investigated the conformation of TM1A and combined molecular dynamics simulations with distance measurements using the lanthanide based FRET variant LRET. Comparison of results obtained from a membrane bilayer with detergent micelle data showed that the conformation of LeuT was environment dependent and only the micelle data were consistent with the crystal structure. Helix TM1A moved towards the membrane-water interface. In contrary, TM1A was stable in its position in the micelle simulations. The same results were obtained by distance measurements using LRET for LeuT solubilized in micelles and LeuT reconstituted into POPC liposomes. Movement of TM1A seems to be driven by partitioning of the polar and charge groups of TM1A out of the hydrophobic core of the membrane. Potential of mean force calculations confirmed an energetic favourable process with an energy difference of ~15 kJ/mol.

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The E. coli Sec reaction pathway for cellular protein sorting under a single molecule loupe

Niels Vandenberk *, Lily Karamanou **, Johan Hofkens *, Tassos Economou ** and Jelle Hendrix *

* Laboratory for Photochemistry and Spectroscopy, Department of Chemistry, KU Leuven,Belgium

** Laboratory for Molecular Bacteriology, Department of Microbiology and Immunology, KU Leuven, Belgium

[email protected]

Protein targeting and secretion is an essential biological process in all life forms. In E. coli, the Sec translocase machinery consists of a conserved protein-conducting channel (SecYEG), which associates with cytoplasmic partners such as SecA [1]. Although busily studied, important mechanistic details on bacterial protein targeting and secretion are still lacking. Since protein conformation and dynamics seems to be key to the function of translocases, we therefore carried out quantitative studies on the dynamic conformation of SecA in vitro using single-pair Förster resonance energy transfer (FRET) on a home-built multiparameter fluorescence detection microscope with pulsed interleaved excitation (MFD-PIE) [2,3]. Using FRET restrained structural modeling (FPS), we selected suitable FRET pairs on SecA [4]. The different positions were mutated to Cysteines, and optimal conditions for labeling with maleimide dyes were determined. Preliminary FRET measurements were performed that provide insight in the stoichiometry of SecA in solution. The perspective is to study all the SecA mutants to create a dynamic 3D model of SecA. In a second part, we perform a biophysical analysis of the Sec pathway in vivo in E. coli. To do this, we have selected plasmids that allow controllable expression of the proteins. Then, we employed number and brightness analysis and raster image correlation spectroscopy to study the stoichiometry and dynamics of SecA in E. coli using a confocal laser scanning microscope [5,6].

[1] Chatzi KE, Sardis MF, Economou T, Karamanou S, “SecA-mediated targeting and translocation of secretory proteins”, Biochimica et Biophysica Acta, 1843., 1466-1474, (2014). [2] Müller BK, Zaychikov E, Brauchie C, Lamb DC, “Pulsed interleaved excitation”, Biophys J, 89(5), 3508-22, (2005). [3] Kudryavtsev V, Sikor M, Kalinin S, Mokranjac D, Seidel C, Lamb D.C, “ Combining MFD and PIE for Accurate Single-Pair Förster Resonance Energy Transfer Meaurements”, ChemPhysChem, 13, 1060-1078, (2012). [4] Kalinin S, Peulen T, Sindbert S, Rothwell PJ, Berger S, Restle T, Goody RS, Gohlke H, Seidel C, “A toolkit and benchmark study for FRET-restrained high-precision structural modelling”, Nature methods, 9(12), 1218-1227, (2012). [5] Digman M, Dalal R, Horwitz A, Gratton E, “Mapping the number of molecules and Brightness in the laser scanning microscope”, Biophysical Journal, 94, 2320-2332, (2008). [6] Digman M, Brown C, Sengupta P, Wiseman P, Horwitz A, Gratton E, “Measuring fast dynamics in solutions and cells with a laser scanning microscope”, Biophysical journal, 89, 1317-1327, (2005).

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Multiplexed Detection of Three Different Tumor Biomarkers in a Rapid Time-Gated Terbium-to-Quantum Dot Homogeneous FRET Immunoassay

S. Bhuckory, K. D. Wegner, X. Qiu, Y. Wu, N. Hildebrandt NanoBioPhotonics (nanofret.com), Institut d’Electronique Fondamentale, Université Paris-

Sud, CNRS, Université Paris-Saclay, 91405 Orsay, France [email protected]

Quantum dot nanocrystals (QDs) are fluorophores with exceptional photophysical and photochemical properties, such as size-tunable absorption and photoluminescence (PL) spectra, efficient and spectrally broad excitation, narrow PL bands, resistance to photobleaching, and stability in biological media. Size-dependent color-coding of QDs makes them ideal candidates for optical multiplexing. In clinical diagnostics, sensitive and specific detection of multiple biomarkers are of utmost importance, and therefore QDs can provide many advantages for clinical immunoassays [1, 2]. One of the main obstacles to overcome for a successful application of QDs in immunoassays for clinical diagnostics is a functional bioconjugation and stability in serum-based samples. In this study, we demonstrate a triplexed homogeneous Förster Resonance Energy Transfer (FRET) immunoassay for the simultaneous detection of the cancer biomarkers PSA (prostate specific antigen), NSE (neuron-specific enolase) and CEA (carcinoembryonic antigen). Monoclonal antibody pairs against the three different markers were conjugated to specific QDs (PL maxima at 605, 655, and 705 nm, respectively) and luminescent terbium complexes (Lumi4-Tb) for the use in sandwich immunoassays. Tb was used as a universal FRET donor for the three different QDs (as FRET acceptors) and time-gated PL detection on a commercial clinical fluorescence plate reader (KRYPTOR) allowed for multiplexed quantification of the three tumor markers from a single 50 µL serum sample at clinically relevant concentrations with sub-nanomolar limits of detection.

Fig. 1. Homogeneous Tb-to-QD FRET immunoassays.

[1] S. Bhuckory, O. Lefebvre, X. Qiu, K. D. Wegner, and N. Hildebrandt “Evaluating quantum dot performance in homogeneous FRET immunoassays for prostate specific antigen” Sensors, 16(2), 197 (2016). [2] K. D. Wegner, Z. Jin, S. Lindén, T. L. Jennings, and N. Hildebrandt “Quantum-Dot-Based Förster Resonance Energy Transfer Immunoassay for Sensitive Clinical Diagnostics of Low-Volume Serum Samples”, ACS Nano, 7 (8), 7411–7419, (2013).

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Near-Infrared FRET imaging reveals the fate and integrity of lipid nanocarriers in healthy and tumor-bearing mice

Redouane Bouchaala1,2, Luc Mercier2, Bohdan Andreiuk1, Ievgen Shulov1, Yves Mély1, ThierryVandamme4, Jacky G. Goetz3, Nicolas Anton4, Klymchenko Andrey1

1) CNRS UMR 7213, Laboratoire de Biophotonique et Pharmacologie, University of Strasbourg, 74 route du Rhin, 67401Illkirch Cedex, France

2) Laboratory of Photonic Systems and Nonlinear Optics, Institute of optics and fine mechanics, University of Sétif 1,19000 Algeria.

3) Inserm U1109, MN3T, unistra Strasbourg, F-67200, France,4) CNRS UMR 7199, Laboratoire de Conception et Application de Molécules Bioactives, University of Strasbourg, 74

route du Rhin, 67401 Illkirch Cedex, France

[email protected];

Lipid nanocarriers emerged as promising candidates for drug delivery and cancer targeting because of their low toxicity, biodegradability and capacity to encapsulate a drug or a contrasting agent (1). However, cause of poor understanding of their in vivo fate and integrity, their translation from laboratory to biomedical applications is limited. In this work, we exploited the Förster Resonance Energy Transfer (FRET) technique for real time investigation of their stability in vivo. Using our recently developed approach of hydrophobic counterion (TPB) (2), we encapsulated two NIR cyanine dyes (Cy 5.5/TPB and Cy 7.5/TPB) inside a lipid nanocarrier of 100 nm size (3). After validation of our FRET nanocarriers in vitro, they were retro-orbitally injected into healthy and tumor bearing mice. Using two-color whole animal NIR imaging, we could quantify the content of nanoparticles in different compartments of the mice, observing that the particles remain stable in the blood circulation for at least 6h. The changes in the FRET signal, i.e. disintegration, was observed after ~4h and finished after 24h. Finally, we found a fast accumulation of nanocarriers inside tumors with the loss of the particle integrity after 1.30 h. In conclusion, we developed a FRET system that allows directly visualization and quantification of nanocarrier integrity in vivo.

Figure. (A) Schematic presentation of FRET system inside lipid nanocarrier encapsulating NIR cyanine dyes. (B) Emission spectra of intact nanocarriers in water and after addition of dioxane. (C) NIR in vivo imaging in

living mice using 100-nm FRET nanocarriers at 0h and 24 h.

1) Klymchenko, A.; Roger, E.; Anton, N.; Anton, H.; Shulov, I.; Vermot, J.; Mely, Y.; Vandamme, T. F. Highly LipophilicFluorescent Dyes in Nano-Emulsions: Towards Bright Nonleaking Nano-Droplets. RSC Adv. 2012, 2, 11876–11886.

2) Kilin VN, Anton H, Anton N, Steed E, Vermot J, Vandamme TF, Mely Y, Klymchenko AS. Counterion-enhancedcyanine dye loading into lipid nano-droplets for single-particle tracking in zebra fish Biomaterials, 6 (2014), pp. 4950–4957.

3) Bouchaala R. et al. in preparation.

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Longitudinal investigation of the calcium concentration in retinal neurons during chronic inflammation in vivo - a FRET-based approach

Daniel Bremer*, Helena Radbruch**, Anja Hauser* and Raluca Niesner*

* German Rheumatism Research Center (DRFZ) Berlin

** Charité University of Medicine Berlin

[email protected]

Preventing pathogenesis and developing selective therapeutic strategies are the main goals in modern biomedicine. Therefore, it is absolutely necessary to probe cellular and tissue functions in the living organism. FRET based systems have proved to be a reliable tool for observing changes through optical systems in vivo, e. g. by means of 2-photon microscopy [1]. The FRET-based calcium indicator troponin-C expressed by neuronal cells of the transgenic reporter mouse strain CerTN L15 is suitable for observing changes in the calcium concentrations in neurons of the brain stem in vivo, in health and disease, as we have previously shown [2]. That means ratiometric FRET allows for probing early neuronal dysfunction preceding morphological changes and finally neuronal death. However, imaging the brain stem implicates terminal experiments and therefore longitudinal measurements in one and the same animal at several time points are not possible. The retina gives a great opportunity to overcome this limitation. First, the eye is the only optical based system in mammals allowing non-invasive measurements of the retina. Second, during embryonic development, the retinal tissue is built from brain tissue meaning observing the retina gives conclusion about the physiologic and pathologic processes in the brain. Thus, observing changes in the mouse retina longitudinally, during autoimmune diseases, e.g. in animal models of multiple sclerosis and uveoretinitis, is suitable for developing therapeutic strategies and is subject of this study.

Fig. 1. Blue (466 nm), green (525 nm) and red (593 nm) fluorescence signals of the FRET-based calcium indicator (Cerulean and Citrine) expressed by neurons in the retina of a healthy CerTN L15 x

LysM tdRFP mouse acquired with a 2-photon microscope

[1] Agata A. Mossakowski et al., “Tracking CNS and systemic sources of oxidative stress during the course of chronic neuroinflammation”, Acta Neuropathologica, (2015)

[2] Helena Radbruch et al., “Intravital FRET: Probing Cellular and Tissue Function in Vivo“, International Journal of Molecular Sciences, 16, 11713-11727, (2015)

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Intra and Extracellular Biosensing Using Time-Gated Terbium to Quantum Dot FRET

M.P. van Bergen P. ,*uX. Qi, **T. Chen ,*Lindén S. ,**H.S. Afsari, *M. Cardoso Dos Santosen Henegouwen***, T.L. Jennings****, K. Susumu*****, I.L. Medintz******,

N. Hildebrandt *, L.W. Miller **

* NanoBioPhotonics (nanofret.com), Institut d'Electronique Fondamentale, Université Paris-Sud, CNRS, Université Paris-Saclay, Orsay, France. ** Department of Chemistry, University

of Illinois at Chicago, USA. *** Cell Biology, Department of Biology, Utrecht University,Netherlands. ****Affymetrix, Inc., San Diego, CA, USA. ***** Sotera Defense Solutions,

Columbia, MD, USA. ****** Center for Bio/Molecular Science and Engineering, Code 6900, US Naval Research Laboratory, Washington, DC, USA.

[email protected]

Time-gated FRET using the unique material combination of long-lifetime terbium complexes (Tb) and semiconductor quantum dots (QDs) provides many advantages for highly sensitive and multiplexed biosensing. Although time-gated detection can efficiently suppress sample autofluorescence and background fluorescence from directly excited FRET acceptors, Tb-to-QD FRET has been rarely exploited for biomolecular imaging. In this study, we highlight Tb-to-QD time-gated FRET biosensing that can be applied for intra and extracellular imaging (Fig.1). Immunostaining of different epitopes of the epidermal growth factor receptor (EGFR) with Tb and QD conjugated antibodies and nanobodies allowed for efficient Tb-to-QD FRET on A431 cell membranes. The broad usability of Tb-to-QD FRET was further demonstrated by intracellular multicolor Tb-to-QD FRET or Tb-to-QD-to-dye FRET using microinjection as well as cell penetrating peptide-mediated endocytosis with HeLa cells. Multiplexed biosensing at very low concentrations (~nM), and the rapid and sensitive detection void of FRET acceptor background fluorescence are highly important advantages for advanced live cell imaging of biomolecular interactions [1].

Fig. 1. Schematic presentation of the various Tb-to-QD FRET imaging approaches [1].

[1] H.S. Afsari, M. Cardoso Dos Santos, S. Lindén, T. Chen, X. Qiu, P. van Bergen, T. Jennings, K. Susumu, I.L. Medintz, N. Hildebrandt, L. Miller, “Time-gated terbium to quantum dot FRET nanoassemblies for fast and sensitive intra and extracellular fluorescence imaging”, Science Advances, submitted (2016).

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Utilizing Photochromic FRET to Develop Color Switching Quantum Dots Sebastián A. Díaz*, Florencia Gillanders**, Kimihiro Susumu***, Eunkeu Oh***, Igor L.

Medintz*, and Thomas M. Jovin**** * Center for Bio/Molecular Science and Engineering, U.S. Naval Research Laboratory, Washington DC, USA

** Center for Investigation in Bionanosciences (CIBION-CONICET),Buenos Aires, Argentina

*** Optical Sciences Division, , U.S. Naval Research Laboratory, Washington DC, USA

**** Laboratory of Cellular Dynamics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

[email protected]

Photoswitchable probes are of great utility in fluorescence microscopy, permitting numerous determinations, including molecular localization super-resolution based on their modifiable emission intensity and spectra.[1] We have associated a 9 nm diameter quantum dot (QD) with blue emission (425 nm) with a diheteroarylethene (PCf), of which the closed form isomer presents absorption and emission maxima at 440 and 520-530 nm respectively, functioning as a fluorescent acceptor for the QD donor in photochromic Förster resonance energy transfer (pcFRET).[2,3] The transition from the non-absorbing, non-fluorescent open state to the fluorescent closed state is achieved by irradiation with near-UV (340 nm), with the reverse transition achieved with >500 nm light. The PCf is coupled to an amphiphilic polymer that stably coats the QD, thereby creating a water soluble color switching QD (csQD) emitting in the blue after visible light irradiation and in the green after UV irradiation. The resulting emission ratio change (QD emission/Dye emission) is up to 280%. The csQD can undergo multiple photocycles with minimal fatigue.

Fig. 1. (Left) Schematic of color switching quantum dots (Right) Fluorescence spectra of color switching quantum dot in the open (visible irradiation) and closed (UV irradiation) state.

[1] Hell, S., Sahl, S., Bates, M., Zhuang, X., et al. “The 2015 Super-Resolution Microscopy Roadmap”, Journal of Physics D: Applied Physics, 48, 35, (2015). [2] Díaz, S., Gillanders, F., Jares-Erijman, E., Jovin, T. “Photoswitchable Semiconductor Nanocrystals with Self-Regulating Photochromic Förster Resonance Energy Transfer Acceptors”, Nat. Comm, 6, (2015). [3] Gillanders, F., Giordano, L., Díaz, S., Jovin, T., Jares-Erijman, E. “Photoswitchable Fluorescent Diheteroarylethenes: Substituent Effects on Photochromic and Solvatochromic Properties”, Photoch. Photobio. Sci., 13, 603, (2014).

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Resolving Interactions Inside Living Cells With Engineered Upconversion Nanoparticles

Drees C*, Raj AN**, Kurre R*, You C*, Haase M** and Piehler J*

* University of Osnabrück, Dept. Biology, Barbarastr. 11, 49076 Osnabrück

** University of Osnabrück, Dept. Inorganic Chemistry I, Barbarastr. 7, 49069 Osnabrück

[email protected]

Upconversion nanoparticles (UCNP) are efficiently excited by sequential multiphoton absorption of NIR light and emit photons in the UV/VIS regime and therefore can be detected with negligible background [1]. Moreover, lanthanide resonance energy transfer (LRET) from UCNP to molecules in immediate proximity opens exciting possibilities as spectroscopic reporters or photoactuators with very high spatial resolution [2]. However, even though strategies for improving the optical properties of UCNP emerged [3], the determinants of UCNP-based LRET as well as its application in a biological context are still poorly resolved. We have engineered biofunctional UCNP optimized for LRET as novel reporters for spatially-resolved protein interaction analysis within living cells. To this end, we implemented microscopic techniques for UCNP excitation and synthesized various nanoparticle species to systematically improve UCNP emission and energy transfer efficiency. These phenomena strongly profited from power densities far beyond commonly published values – in agreement with recent studies on this topic [3]. Strikingly, we observed further enhancement of LRET when breaking with traditional paradigms of UCNP design. In order to exploit these unique optical properties in a biological context, we established a biofunctional coating with an anti-GFP nanobody for selective targeting in the cytoplasm of living cells. With the mitochondrial TOM complex as model system we demonstrated specificity of UCNP functionalization by colocalization and further confirmed this interaction by LRET-sensitized dye emission.

Fig. 1. UCNP functionalized with anti-GFP specifically bind to mitochondrial Tom20-EGFP-HaloTag. Once excited by NIR light, UCNP luminesce and transfer energy to the acceptor dye label of the HaloTag.

[1] Wang M, Abbineni G, Clevenger A, Mao C & Xu S “Upconversion nanoparticles: synthesis, surface modification and biological applications”, Nanomedicine : nanotechnology, biology, and medicine, 7, 710-729, (2011). [2] Wang F, Banerjee D, Liu Y, Chen X & Liu X “Upconversion nanoparticles in biological labeling, imaging, and therapy”, Analyst 135(8):1839-1854, (2010). [3] Gargas DJ, Chan EM, Ostrowski AD, Aloni S, Altoe MVP, Barnard ES, … Schuck PJ “Engineering bright sub-10-nm upconverting nanocrystals for single-molecule imaging”, Nature Nanotechnology, 9, 300-305 (2014).

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Semiconductor Quantum Dots as Förster Resonance Energy Transfer Donors for Intracellularly-Based Biosensors

James ,1Igor L. Medintz, 3Eunkeu Oh, 3, Kimihiro Susumu1, Scott A. Walper1Lauren D. Field*1B. Delehanty

U.S. Naval Research ineering, Code 6900, ce and EngCenter for Bio/Molecular Scien1

Laboratory, 4555 Overlook Ave, S.W. Washington, DC 20375, USA Optical Sciences Division, Code 5600 U.S. Naval Research Laboratory Washington, DC 2

20375, USA , USAlumbia, MD 21046Sotera Defense Solutions, Inc. 7230 Lee DeForest Drive Co3

Contact author: [email protected]

Förster resonance energy transfer (FRET)- based assemblies currently form a significant portion of intracellular sensors. Although extremely useful, the fluorescent protein pairs typically utilized in such sensors are still plagued by many photophysical issues including significant direct acceptor excitation, small changes in FRET efficiency, and limited photostability. Luminescent semiconductor nanocrystals or quantum dots (QDs) are characterized by many unique optical properties including size-tunable photoluminescence, broad excitation profiles coupled to narrow emission profiles, and resistance to photobleaching, which can cumulatively contribute to overcoming many of the issues associated with use of fluorescent protein FRET donors. Utilizing QDs for intracellular FRET-based sensing still requires a significant development of materials optimization, cellular delivery, bioconjugation and assay designs. We are currently developing several QD-based FRET sensors for intracellular application. This includes sensors targeting intracellular proteolytic activity along with those based on theranostic nanodevices for monitoring drug release. The protease sensor is based on a unique design where an intracellularly expressed fluorescent acceptor protein substrate assembles onto a QD donor following microinjection to form the active sensor. For the latter, the QD is conjugated to a carrier protein and a drug analogue to visualize real-time intracellular release of the drug from its carrier. The focus of this talk will be on the design, properties, photophysical characterization and preliminary cellular application of these sensor constructs.

Fig. 1. Schematic of a Quantum Dot protein complex with depicted FRET.

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Genetically encoded caspase-3 FRET-sensor based on terbium chelate and red fluorescent protein

Goryashchenko A.S.*, Khrenova M.G.**, Bochkova A.A.**, Ivashina T.V.***, Vinokurov L.M.**** and Savitsky A.P.*

* A.N. Bach Institute of Biochemistry, Research Center of Biotechnology of the RussianAcademy of Sciences, Moscow, Russia;

** M.V. Lomonosov Moscow State University, Department of Chemistry, Moscow, Russia;

*** Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino, Moscow Region, Russia;

**** Branch of Shemyakin and Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Pushchino, Moscow Region, Russia;

[email protected]

FRET-based biosensors are widely used to study the enzymatic activity in living cells. Fluorescent complexes of lanthanides, which have a microsecond fluorescence, can be used as donors in FRET-pair, and while combined with time delay spectroscopy allow to eliminate short-lived autofluorescence of biomolecules and scattered light. In addition, measurements with time delay detect only sensitized fluorescence of the acceptor, that’s why it solves the problem of simultaneous fluorescence excitation of the donor and acceptor and increases the dynamic range of measurements and the accuracy of the FRET efficiency determination.

Here we describe the genetically encoded caspase-3 FRET-sensor Tb3+-TBP-19-TagRFP based on the terbium-binding peptide YIDTNNDGWYEGDELLA as a donor, cleavable linker VDGGSGGDEVDGWGGSGLD with caspase-3 recognition site DEVD and red fluorescent protein TagRFP as an acceptor. The engineered construction performs two induction-resonance energy transfer processes: from tryptophan of the terbium-binding peptide to Tb3+ and from sensitized Tb3+ to acceptor - the chromophore of TagRFP.

For the first time it was shown that fluorescence resonance energy transfer between sensitized terbium and TagRFP in the engineered construction can be studied via detection of microsecond TagRFP fluorescence intensities. Direct measurement of sensitized TagRFP fluorescence using time gated technique eliminates such problems as direct acceptor excitation due to spectral cross-talk between donor and acceptor. Previously, FRET efficiency of 44% was detected by change of donor (terbium ion) fluorescence lifetime from 0.33 ms for photobleached form of Tb3+-TBP-19-TagRFP to 0.18 ms for initial form. This method alsoallows to exclude the signal cross-talk, but fluorescence lifetime of terbium ion can vary because of changes in its coordination sphere, decreasing the accuracy of the results. Direct detection of sensitized TagRFP fluorescence using time gated approach allows to get rid of both problems mentioned above, because we measure only the signal that is connected to FRET, while the background fluorescence and signal from directly excited TagRFP were already gone to zero during the delay time. We also calculated the Kd value of the Tb3+ andTBP-19-TagRFP complex and found it to be 17±7 µM. Finally, the molecular dynamics modeling allowed us to calculate the distribution of possible distances between the donor and acceptor in Tb3+-TBP-19-TagRFP sensor. Calculated FRET efficiency is equal to 43% whichin good agreement with the experimental value of 44%.

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Single Terbium-Quantum Dot FRET Pair for Time-GatedLifetime Multiplexing of microRNA

Niko Hildebrandt , andJiajia Guo ,Xue QiuNanoBioPhotonics (nanofret.com), Institut d'Electronique Fondamentale, Université Paris-

Sud, CNRS, Université Paris-Saclay, 91405 Orsay Cedex, [email protected]

Quantification of several biomarkers at very low concentrations is an important requirement for many optical biosensing applications. Förster resonance energy transfer (FRET) using luminescent terbium complexes (Tb) as donors provides several unique advantages for multiplexed biomolecular sensing [1]. The various narrow photoluminescence (PL) emission bands over a relatively large spectra range allow for FRET to several different acceptors, a concept which has been exploited for spectral (or color) multiplexing [2-4]. Using the extremely long PL lifetimes of Tb (ms) and different donor-acceptor distances for the same type of donor-acceptor pair would also allow for designing different FRET-quenched and FRET-sensitized PL decay times, which could open the possibility of lifetime multiplexing. Here, we demonstrated the development of a lifetime-duplexed microRNA assay using a single Tb-quantum dot FRET pair. By applying different QD self-assemblies, we were able to control the distances of the Tb-donor and QD-acceptor and to obtain distinguishable PL decay curves. Time-gated FRET detection was used to perform a rapid and homogeneous lifetime-multiplexed assay to detect hsa-miRNA-20a-5p and hsa-miRNA-20b-5p from a single 150 µL sample. The assay was carried out at room temperature and could be accomplished within only 35 minutes. Limits of detection were 0.43 nM and 0.88 nM for miRNA-20a and miRNA-20b, respectively. This proof-of-concept of time-gated temporal multiplexed miRNA detection opens a new route for higher order multiplexed clinical diagnostics.

Fig. 1. Tb-to-QD FRET with tuned Tb-QD distances for temporal multiplexed detection of miRNA.

[1] N. Hildebrandt, K. D. Wegner, and W. R. Algar “Luminescent Terbium Complexes: Superior Förster Resonance Energy Transfer Donors for Flexible and Sensitive Multiplexed Biosensing“, Coordination Chemistry Reviews, 273–274, 125–138, (2014). [2] D. Geißler, S. Stufler, H.-G. Löhmannsröben and N. Hildebrandt “Six-Color Time-Resolved Förster Resonance Energy Transfer for Ultrasensitive Multiplexed Biosensing”, Journal of the American Chemical Society, 135, 1102-1109, (2013). [3] X. Qiu and N. Hildebrandt “Rapid and Multiplexed MicroRNA Diagnostic Assay Using Quantum Dot-Based Förster Resonance Energy Transfer”, ACS Nano, 9 (8), 8449-8457, (2015). [4] Z. Jin, D. Geißler, X. Qiu, K. D. Wegner, and N. Hildebrandt “Rapid, Amplification-Free, and Sensitive Diagnostic Assay for Single-Step Multiplexed Fluorescence Detection of MicroRNA”, Angewandte Chemie International Edition, 54, 10024-10029, (2015).

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April 2016th 6 - 3rd –IIFRET in Life Sciences –tional Discussion Meeting Interna

Using FRET to follow protein oligomerization throughout a viruses’ life cycle Doortje Borrenberghs*, Lieve Dirix*, Flore De Wit**, Susana Rocha*, Jolien Blokken**,

Stéphanie De Houwer**, Rik Gijsbers**, Frauke Christ**, Johan Hofkens*, Jelle Hendrix* and Zeger Debyser**

* Laboratory for Photochemistry and Spectroscopy, Department of Chemistry, KU Leuven, Celestijnenlaan 200F, 3001 Leuven, Belgium.

** Laboratory for Molecular Virology and Gene Therapy, Department of Pharmaceutical and Pharmacological Sciences, KU Leuven, Kapucijnenvoer 33, 3000 Leuven, Belgium

[email protected] Nuclear entry is a selective, dynamic process granting the HIV-1 pre-integration complex (PIC) access to the chromatin. Classical analysis of nuclear entry of heterogeneous viral particles only yields averaged information. We now have employed single-virus particle fluorescence methods to follow the journey of single viral pre-integration complexes (PICs) during infection by visualizing HIV-1 integrase (IN). Nuclear entry is associated with a reduction in the number of IN molecules in the complexes while the interaction with LEDGF/p75 enhances IN oligomerization in the nucleus. Addition of LEDGINs, small molecule inhibitors of the IN-LEDGF/p75 interaction, during virus production, prematurely stabilizes a higher-order IN multimeric state, resulting in stable IN multimers resistant to a reduction in IN content and defective for nuclear entry. This suggests that a stringent size restriction determines nucleopore entry. Taken together, this work demonstrates the power of single-virus imaging providing crucial insights in HIV replication and enabling mechanism-of-action studies.

Fig. 1. (left) Acceptor photobleaching (AP)-FRET measurements, the fluorescence intensity of the

donor (mTFP1; D,pre) in viral complexes containing donor and acceptor capable of FRET is quenched by the proximal acceptor (mVenus; A,pre). After photobleaching of the acceptor (mVenus; A,post),

the fluorescence intensity of the donor is dequenched and its brightness increases (D, post). (middle) A representative image of a lamin stained (blue) HeLaP4 cell infected with HIVIN-mTFP1+IN-

mVenus is shown. Scale bars represent 5 µm. (right) A 2D Gaussian function showing an increase in fluorescence intensity of the donor after (D, post) photobleaching of the acceptor compared to the

intensity of the donor before photobleaching (D, pre) [1]. [1] Borrenberghs et al. “Dynamic integrase oligomerization orchestrates HIV nuclear entry”, submitted.

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Multiplexed MicroRNA Detection Using Terbium to Quantum Dot FRETNiko HildebrandtXue Qiu and

NanoBioPhotonics (nanofret.com), Institut d'Electronique Fondamentale, Université Paris-Sud, CNRS, Université Paris-Saclay, 91405 Orsay Cedex, France

[email protected]

The detection of next generation microRNA (miRNA) biomarkers has become a highly important aspect for clinical diagnostics. Here we present an enzyme-free, simple, rapid, and homogeneous multiplexed assay-technology for the sensitive detection of multiple miRNAs in a single small-volume buffer or serum-based sample measured at room temperature. The assay is based on time-gated FRET (TG-FRET) from the luminescent terbium complex Lumi4-Tb (Tb) to different commercial QDs. It does neither require any washing or separation steps nor the addition of enzymes and has been designed for its application on an approved clinical diagnostics fluorescence microplate reader (KRYPTOR). In order to demonstrate full multiplexing functionality of our assay we have detected the three miRNAs hsa-miR-20a-5p, hsa-miR-20b-5p, and hsa-miR-21-5p in a single 150 µL sample at constant temperature, all with ca. 1 nM (sub-pmol) detection limits. We also demonstrate precise multiplexed measurements of these miRNAs at different and varying concentrations and the feasibility of adapting the technology to point-of-care testing (POCT) in buffer containing 10 % serum. Our assay does not only demonstrate an important milestone for the integration of quantum dot nanoparticles to multiplexed clinical diagnostics but also a unique rapid miRNA detection technology that is complimentary to the rather complicated high-throughput and high-sensitivity approaches that are established today [1].

Fig. 1. Triplexed detection of three different microRNAs (miRNAs) from a single sample using Tb-to-QD FRET and time-gated luminescence detection [1]. Copyright 2015 American Chemical Society.

[1] X. Qiu and N. Hildebrandt “Rapid and Multiplexed MicroRNA Diagnostic Assay Using Quantum Dot-Based Förster Resonance Energy Transfer”, ACS Nano, 9(8), 8449-8457, (2015).

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Time-resolved FRET measurements of CFP-YFP-based biosensors Henning Höfig*,**, Daryan Kempe*, Martina Pohl*** and Jörg Fitter*,**

* I. Institute of Physics (IA), Biophysics Group, RWTH Aachen, Aachen, Germany

** Institute of Complex Systems, Molecular Biophysics (ICS-5), Forschungszentrum Jülich GmbH, Jülich, Germany

*** Institute of Bio- and Geosciences, Biotechnology (IBG-1), Forschungszentrum Jülich GmbH, Jülich, Germany

[email protected]

Genetically encoded FRET-based biosensors consist of two fluorescent proteins (donor and acceptor) and a sensing domain. If a biological stimulus acts on the sensing domain this is converted into a change in energy transfer [1]. During the last years, the variety of FRET-based biosensors and also the combinations of fluorescent proteins have become numerous. A frequently used FRET pair is cyan fluorescent protein (CFP) as donor and yellow fluorescent protein (YFP) as acceptor and their variants. We use a confocal microscope with 440nm and 510nm laser excitation to perform time-resolved FRET measurements on CFP-YFP-based biosensors. The readout of FRET-based biosensors usually utilizes the ratio of fluorescence emission intensities of the donor and the acceptor upon donor excitation. Often, the sensor signal is calibrated in an in vitro measurement. The obtained calibration curve is then used to assign a measured sensor signal in an in vivo experiment to a certain physical quantity, e.g. a metabolite concentration. However, it was shown that the sensor readout can vary remarkably with changing environmental conditions (e.g. pH, salt and metabolite concentrations) [2]. Also the presence of macromolecular crowding inside the cell might have an impact on the sensor performance. We carried out time-resolved FRET measurements on the glucose binding sensor FLII12Pglu600µ [3]. The glucose binding protein MglB forms the sensing domain of this sensor because it undergoes a conformational change upon glucose binding. The donor of the sensor is ECFP and the acceptor is Citrine. Sensor optimization showed that a larger signal change of the sensor is observed for a design with internally fused fluorescent proteins in comparison to terminally fused fluorescent proteins [3]. Deuschle et al. attribute the higher signal change to a reduction of rotational mobility of the fluorescent proteins. Consequently, the relative orientation of the fluorescent proteins alters when the sensor undergoes the conformational change. We perform single-molecule FRET and time-resolved anisotropy measurements to investigate the sensing mechanism of FLII12Pglu600µ. Additionally, we introduced a double cysteine mutation of MglB and labelled it with the artificial dye pair Alexa 488/647. This approach rules out the uncertainty of relative fluorophore orientation because the dyes have a high rotational mobility and rotational averaging is valid. Thus, absolute distance changes upon glucose binding can be measured.

[1] Haley J Carlson and Robert E Campbell “Genetically encoded FRET-based biosensors for multiparameter fluorescence imaging”, Curr. Opin. Biotechnol., 20, 19-27, (2009). [2] Roland Moussa et al. “An evaluation of genetically encoded FRET-based biosensors for quantitative metabolite analyses in vivo”, J. Biotechnol., 191, 250-259, (2014). [3] Deuschle et al. “Construction and optimization of a family of genetically encoded metabolite sensors by semirational protein engineering”, Protein Sci., 14, 2304-2314, (2005).

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Multiplexed micro-RNA assays using time-resolved FRET with biospectral correction Zongwen Jin *, Daniel Geißler**，Niko Hildebrandt***

* Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences (China)

** BAM, Federal Institute for Materials research and Testing (Germany) *** Université Paris-Saclay, Université Paris-Sud, CNRS (France)

Contact authors: [email protected] and [email protected]

Applications based on Förster Resonance Energy Transfer (FRET) play an important role for the determination of concentrations and distances within nanometer-scale systems in vitro and in vivo in many fields of the life sciences.[1] Using time-resolved optical spectroscopy and microscopy for the analysis of FRET systems offers several advantages concerning sensitivity and specificity. Luminescent lanthanide complexes, and in particular Tb complexes, exhibit extremely long luminescence lifetimes and multiple narrow emission peaks over a broad spectral range. These photophysical features make them highly interesting FRET donors in combination with different FRET acceptors, such as organic dyes. Such FRET pairs have been successfully used for the multiplexed and highly sensitive detection of protein, peptide, DNA, and RNA biomarkers.[2] In this contribution we will present organic acceptor dye based homogeneous single-step FRET biosensors for the sensitive and specific detection of three different micro-RNAs or ssDNAs from a single low-volume sample. In particular, we will discuss problems of spectral crosstalk and biological interferences in our multi-color and multi-analyte assays and present an efficient correction algorithm to efficiently overcome both problems.[3] These novel FRET biosensors provide a rapid, simple, selective, and sensitive tool for multiplexed detection of various oligonucleotides, which makes them highly interesting for clinical diagnostics and other biosensing applications. [1],[4]

Fig. 1. Multiplexed assay for detecting micro-RNAs using Time-resolved FRET

[1] I. Medintz and N. Hildebrandt (editors). “FRET – Förster Resonance Energy Transfer”， From Theory to Applications, Wiley-VCH, Germany 2014, ISBN 978-3-527-32816-1. [2] N. Hildebrandt, K. D. Wegner, and W. R. Algar. “Luminescent Terbium Complexes: Superior Förster Resonance Energy Transfer Donors for Flexible and Sensitive Multiplexed Biosensing”, Coordination Chemistry Reviews, 273–274, 125–138, 2014. [3] Z. Jin, D. Geißler, X. Qiu, K. D. Wegner, and Niko Hildebrandt. “Rapid, Amplification-Free, and Sensitive Diagnostic Assay for Single-Step Multiplexed Fluorescence Detection of MicroRNA” Angewandte Chemie International Edition, 54, 10024-10029, 2015. [4] D. Geißler , S. Stufler, H.-G. Löhmannsröben and N. Hildebrandt. “Six-Color Time-Resolved Förster Resonance Energy Transfer for Ultrasensitive Multiplexed Biosensing” Journal of the American Chemical Society , 135, 1102-1109, 2013.

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Energy Transfer-Based Sensitization of Luminescent Gold Nanoclusters

Eunkeu Oh*, Alan L. Huston*, Andrew Shabaev**, Alexander Efros**,

Kimihiro Susumu*, and Igor L. Medintz***

*Optical Sciences Division, Code 5600;**Center for Computational Material Science, Code6390; **Center for Bio/Molecular Science and Engineering, Code 6900;

U.S. Naval Research Laboratory, Washington, DC 20375 USA

[email protected]

Luminescent gold nanocrystals (AuNCs) are a recently developed material with strong potential for biological and optoelectronic applications.[1] Although the exact method by which they luminesce is still not fully understood, they have already demonstrated energy transfer properties and especially as an acceptor that undergo sensitization. The observed energy transfer processes have been primarily ascribed to Förster resonance energy transfer (FRET) and, to a lesser extent, nanosurface energy transfer (NSET).[2] We have been investigating AuNC acceptor capabilities with three structurally/functionally-distinct classes of donors including organic dyes, metal chelates and semiconductor quantum dots (QDs). A significant level of donor quenching was observed for every donor-AuNC acceptor pair although AuNC acceptor sensitization was only observed when paired with the long-lifetime metal-chelates and the QDs. The application of FRET theory largely underestimated the observed energy transfer while use of NSET-based damping models provided somewhat better fits but could not reproduce the experimental data. Cumulatively, this suggests that AuNC sensitization is not by classical FRET or NSET and, along with discussing additional contributing factors, we provide a simplified ET model to fit such experimental data.

Wavelength (nm)

PL (A

U)

Fig. 1. TEM of semiconductor quantum dots assembled with AuNCs and sensitization

[1] Oh, E.; Fatemi, F.; Currie, M.; Delehanty, J.B., Pons, T.; Fragola, A.; Lévêque-Fort, S.; Goswami, R., Susumu, K.; Huston, A. and Medintz, I.L. “PEGylated Luminescent Gold Nanoclusters: Synthesis, Characterization, Bioconjugation and Application to One- and Two-Photon Cellular Imaging.” Particle and Particle Systems Characterization 30, 453-466 (2013). [2] Pons, T., Medintz, I.L., Sapsford, K.E., Higashiya, S., Grimes, A.F., English, D.S., and H. Mattoussi. On the quenching of semiconductor quantum dot photoluminescence by proximal gold nanoparticles. Nano Letters 7, 3157-3164 (2007).

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Homogeneous Proximity-Ligation FRET Immunoassays for VEGF DetectionHamsika Parimi, Xue Qiu, and Niko Hildebrandt

NanoBioPhotonics (nanofret.com), Institut d'Electronique Fondamentale, Université Paris-Sud, CNRS, Université Paris-Saclay, 91405 Orsay Cedex, France

[email protected]

Biomarker recognition and the monitoring of biomolecular interactions to understand their interplay in complex biological systems is of fundamental importance to the life sciences. Förster resonance energy transfer (FRET) is a non-radiative energy transfer between a luminescent donor molecule and a suitable acceptor molecule. Applying the high speed and sensitivity of luminescence to the study of nanoscale interactions, FRET is widely used to develop homogeneous assays. In this study we aimed at a combination of proximity ligation assays (PLA) and FRET to develop versatile and sensitive homogeneous immunoassays. Vascular endothelial growth factor (VEGF), which plays a key role in the regulation of angiogenesis, vasculogenesis, skeletal growth, and reproductive function in adults, and has been implicated in a range of human malignancies as well as cardiovascular and neural dysfunction [1, 2], was used as a model biomarker to develop and validate PLA-FRET assays. VEGF-specific antibodies were functionalized with PLA oligonucleotides (PLA arms), which were further used to hybridize complementary Tb donor and dye (Cy5.5) acceptor oligonucleotides. Binding of the antibodies to VEGF leads to spatial proximity and ligation of the different PLA arms. In turn, this proximity ligation leads to FRET between Tb and dye, which can be used for VEGF quantification. Several different hybridization, ligation, and biorecognition schemes, and steady-state and time-resolved fluorescence spectroscopy were used to investigate the PLA-FRET assay design.

Fig. 1. Combination of To-to-dye FRET and proximity ligation assays for the detection of VEGF.

[1] Berendsen AD, Olsen BR “How vascular endothelial growth factor-A (VEGF) regulates differentiation of mesenchymal stem cells”, Journal of Histochemistry and Cytochemistry, 62(2),103-108, (2014). [2] Gao JZ, Wang YL, Li J, Wei LX “Effects of VEGF/VEGFR/K-ras signaling pathways on miRNA21 levels in hepatocellular carcinoma tissues in rats”, Genetic Molecular Research, 14(1), 671-679, (2015).

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Multiplexed and Homogenous FRET Diagnostics of MicroRNAAkram Yahia-Ammar*, Xue Qiu*, Alexandra Petreto*, Zongwen Jin**, Nadia Cherradi***,

Florence Apparailly****, and Niko Hildebrandt* * NanoBioPhotonics (nanofret.com) Institut d’Electronique Fondamentale, Université Paris-Sud, CNRS, Université Paris-Saclay 91405 Orsay Cedex (France). ** CAS Key Laboratory of

Health Informatics, Shenzhen (China). *** INSERM U1036F, Grenoble (France). **** INSERM U1183, CHU Saint Eloi, Montpellier (France)

[email protected]

The importance of microRNA (miRNA) dysregulation for the development and progression of diseases and the discovery of stable miRNAs in peripheral blood have made these short- sequence nucleic acids to next-generation biomarkers. Different methods have been used for detecting miRNAs, such as northern blotting, RT-qPCR, microarrays, and sequencing. However, all of the methods have limitations related to sensitivity, selectivity, multiplexing, or ease-of-use [1]. The development of novel methods, which are applicable to clinical settings and provide simple and rapid detection, are therefore highly important for clinical diagnostics of circulating miRNAs. Here, we show a homogeneous, simple, rapid, sensitive, and multiplexed method to detect different miRNAs from a single 150 µL sample with low picomolar limits of detection. The technology is based on time-gated Förster resonance energy transfer (FRET) from Tb complexes (Lumi4-Tb) to different organic dyes and combines different hybridization and ligation steps for probe–target recognition. The clinical applicability of the FRET assays was evaluated by detecting miRNA-139-5p extracted from human adrenocortical carcinoma cells and miRNA-146a extracted from lipopolysaccharide (LPS)-stimulated cells, and by comparing the results to RT-qPCR measurements.

Fig. 1. Multicolor microRNA detection using Tb-to-dye FRET

[1] Jin, Z.; Geißler, D.; Qiu, X.; Wegner, K. D.; Hildebrandt, N. “A Rapid, Amplification-Free, and Sensitive Diagnostic Assay for Single-Step Multiplexed Fluorescence Detection of MicroRNA”, Angewandte Chemie International Edition, 54, 10024–10029, (2015).

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Developing fluorogen activating protein-fluorescent protein FRET pairs to investigate signalling of the high affinity IgE receptor (FcεR1)

Genevieve K. Phillips*, Samantha L. Schwartz*, Diane S. Lidke*, and Marcel P. Bruchez**

* University of New Mexico, Health Sciences Center, Pathology Department** Carnegie Mellon University, Molecular Biosensor and Imaging Center

[email protected]

Fluorogen activating proteins (FAPs) are genetically encoded tags made from single chain antibody fragments (scFv) designed to bind fluorogens with high specificity [1]. Both the fluorogen and FAP can be modified to provide flexibility in properties such as affinity, membrane permeability, spectra, and quantum yield [2]. The fluorogen Malachite Green (MG) has two excitation peaks, the maximum at 630 nm and a secondary peak at 450 nm. The emission spectra of blue-emitting fluorescence proteins, such as mCerulean (mCer), overlap with the MG secondary peak, generating a FRET pair with large Stokes shift emission. Using 405 nm excitation of mCer, we observe acceptor sensitized emission at >650 nm with no spectral crosstalk between the donor and acceptor channels. Additionally, we can control when FRET occurs since there is no acceptor until after the addition of fluorogen, providing intra-cellular controls.

We have characterized the FAP-FRET system using proof of principle constructs: FAP-mCer-transmembrane (TM) as a positive FRET control and FAP-TM-mCer as a negative FRET control. We express these constructs either in HeLa cells or RBL cells. Multiple MG derivatives were compared and imaging parameters were optimized to determine the optimal FRET pair. Analysis was performed using code written in Matlab to mask the cell membrane and quantify FRET efficiencies, based on donor intensity before and after addition of fluorogen. Data from membrane impermeable MG-βTau showed high energy transfer efficiency (0.234) with the FAP-mCer-TM construct compared to negligible FRET (0.021) for FAP-TM-mCer. (Figure 1).

Our ultimate goal is to apply FAP-FRET to investigate the spatiotemporal events of IgE receptor signalling. Experiments to investigate the recruitment of Syk, a cytoplasmic kinase, to the IgE receptor tail are underway using both ensemble and single molecule FRET measurements.

Fig. 1. FRET is distance-dependent. The FAP-mCer-TM (positive control) and FAP-TM-mCer (negative control) constructs show markedly different FRET ratio images (405 nm ex; 640-740 nm

em/450-550 nm em). The FRET efficiency for the proximal construct is ~0.23 while the FRET efficiency for the membrane-separated construct is ~0.

[1] Szent-Gyorgyi, C., et al. “Fluorogen-activating single-chain antibodies for imaging cell surface proteins”, Nature Biotechnology, vol. 26(2), 235-40, (2008). [2] Saurabh, S., et al. “Kinetically tunable photostability of fluorogen-activating peptide-fluorogen complexes”, ChemPhysChem, vol. 16(14), 2974-2980, (2015).

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Exploiting fast exciton diffusion in dye-doped polymer nanoparticles to engineer efficient photoswitching through FRET

Kateryna Trofymchuk*, Andreas Reisch*, Luca Prodi**, Jochen Mattay***, Yves Mely*, Andrey S. Klymchenko*

* Laboratoire de Biophotonique et Pharmacologie, UMR 7213 CNRS, Université deStrasbourg, Faculté de Pharmacie, 74, Route du Rhin, 67401 ILLKIRCH Cedex, France

** Dipartimento di Chimica “Giacomo Ciamician”, Università degli Studi di Bologna, via Selmi 2, 40126Bologna, Italy

*** Organic Chemistry I, Department of Chemistry, Bielefeld University, Universitätsstr. 25, 33615 Bielefeld, Germany

[email protected]

Photoswitching of bright fluorescent nanoparticles opens new possibilities for bioimaging with superior resolution. However, efficient photoswitching of nanoparticles is hard to achieve using Forst r resonance energy transfer (FRET) to a photochromic dye, because the particle size is usually larger than the Forst r radius and encapsulation of a large number of dyes leads to aggregation-caused quenching. Recently, we proposed a solution to these problems by exploiting bulky hydrophobic counterions that prevent self-quenching and favour communication of octadecyl rhodamine B dyes by exciton diffusion inside a polymer matrix of poly(D,L-lactide-co-glycolide), resulting in 40nm NPs that were ~6-fold brighter than quantum dots (QD605) [1]. We exploited the exciton diffusion within the FRET donor dyes to boost photoswitching efficiency in dye-doped polymer nanoparticles. To this end, we coencapsulated into PLGA NPs rhodamine B dye salt with four different bulky hydrophobic counterions as well as a photochromic acceptor of the diarylethene family. It was found that the counterion favouring the exciton diffusion enables light-driven variation of fluorescence intensity up to 20-fold, while for all other counterions, the switching efficiency was much lower. The performance of the new NPs was validated at the single particle level, where reversible photoswitching was demonstrated [2]. The proposed concept paves the way to new efficient photoswitchable nanomaterials.

0 1 2 3 40

5

10

15

20

25B) high

low

Re

lative

in

ten

sity (

I/I 0

off)

UV/ Vis cycles

Dye-dye comunicationA)

Fig. 1. . A) Schematic presentation of the photoswitching concept in dye-doped NPs. B) Photoswitching of fluorescent NPs with high and low level of dye-dye communication.

[1] Reisch, A., Didier, P., Richert, L., Oncul, S., Arntz, Y., Mely, Y., Klymchenko, A. S. “Collective fluorescence switching of counterion-assembled dyes in polymer nanoparticles”, Nat. Commun, 4089, (2014). [2] Trofymchuk, K., Prodi, L., Reisch, A., Mely, Y., Altenhoener, K., Mattay, J., Klymchenko, A.S.” Exploiting Fast Exciton Diffusion in Dye-Doped Polymer Nanoparticles to Engineer Efficient Photoswitching”, J.Phys.Chem.Lett. 6, 2259, (2015).

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Towards functional imaging in vivo A FRET-based Genetically Encoded Calcium Indicator in B Cells

Carolin Ulbricht*,** , Lars Nitschke*** , Raluca Niesner** , Helena Radbruch*, Anja E. Hauser*,** * Charité University of Medicine Berlin

** German Rheumatism Research Center (DRFZ) Berlin *** Friedrich-Alexander-University Erlangen

[email protected] A key feature of the adaptive immune response is the production of high affinity antibodies by plasma cells stemming from B cells of germinal centers (GC) in secondary lymphoid organs. To achieve this, the B cell receptor (BCR) that recognizes antigen has to be edited by somatic hypermutation following selection for the highest reactivity against antigen bound on follicular dendritic cells (FDCs) within the GCs. In order to elucidate the selection in vivo, we want to image this process in living mice. Available reporter systems rely on the intensity enhancement of a single fluorophore in response to calcium [1], but this does not allow for a quantitative approach in determining calcium flux as an activation marker. To circumvent this problem, we used a B cell-specific, FRET-based genetically encoded calcium indicator (GECI). A similar biosensor has already been used in our intravital studies investigating the function of neurons [2]. We were able to detect an increased signal ratio between acceptor (Citrine) and donor (eCFP) fluorophore after stimulation in vitro by flow cytometry and confocal microscopy. Furthermore, we established an in vivo two-photon imaging setup to analyze B cell activation. Calcium flux was detected in stationary B cells contacting FDCs in GCs of the popliteal lymph node, while motile B cells in the same area did not display a change in signal ratio. We plan to further quantify calcium flux in this system using fluorescence lifetime imaging in vivo.

Fig. 1. Schematic overview of calcium flux coupled signal detection

[1] Ziv Shulman, Alexander D. Gitlin, Jason S. Weinstein, Begoña Lainez, Enric Esplugues, Richard A. Flavell, Joseph E. Craft, Michel C. Nussenzweig „Dynamic signaling by T follicular helper cells during germinal center B cell selection“, Science, 345, 1058 (2014) [2] Helena Radbruch, Daniel Bremer, Ronja Mothes, Robert Günther, Jan Leo Rinnenthal, Julian Pohlan, Carolin Ulbricht, Anja E. Hauser, Raluca Niesner “Intravital FRET: Probing Cellular and Tissue Function in Vivo“, International Journal of Molecular Sciences, 16, 11713-11727, (2015)

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Development and characterization of tandem heterodimeric fluorescent proteins Matthew D. Wiens *, Yi Shen*, Xi Li*, Mohammad Salem*, Nick Smisdom, Alex Brown*,

Nils Peterson*, and Robert E. Campbell*

* University of Alberta

** Universiteit Hasselt

[email protected]

Fluorescent protein development has primarily focused on making the bright and monomeric protein tags that fold and form the chromophore with high efficiency. As the majority of fluorescent proteins in nature are tetramers, this means that the first step in almost any evolution program is to break the tetramer down to a dimer and then to a monomer. This process invariably decreases brightness and folding, requiring further efforts to regain these characteristics. In this work I have retained the dimeric oligomerization interface in an effectively monomeric “tandem dimer”. In tandem dimer proteins, two copies of the same protein are genetically fused together with a linker that allows the dimerization to occur intramolecularly. This allows two fluorophores to be in close and stable proximity in positions and orientations that should be effectively identical to those observed in the x-ray crystal structures of oligomeric variants. It has previously been found that the DsRed fluorescent protein can get trapped in a dead-end green during the autogenic post-translational modifications that generally lead to a red fluorophore. We were able to create a variant of DsRed protein that favours the green state through directed evolution. With this green variant we were able to engineer GRvT, “Green Red vine Tomato”, which is a tandem heterodimer composed of a green DsRed variant linked to a red DsRed variant. GRvT has fascinating properties such as near perfect predicted and measured FRET efficiency in the range of 90-100% and a quantum yield for the FRET acceptor that is close to 100%.

Fig. 1. Cartoon representation of GRvT

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Time gated FRET microscopy- and plate based assays to study G-Protein Coupled .donors 3+Tb and 3+Eu bright usingReceptors

Jurriaan Zwier *, Pauline Scholler*/**, Laurent Lamarque*, Orestis Faklaris**, Philippe Rondard**, Thierry Durroux**, David Parker***, Jean-Philippe Pin** and Eric Trinquet*

* Cisbio Bioassays, Codolet, France

** Institut de Génomique Fonctionnelle, Montpellier, France

*** Department of Chemistry, Durham University, Durham, United Kingdom

[email protected]

Ever since the first appearance of time gated microscopy based on phosphorescence or lanthanide based luminescence [1] this type of microscopy has, until recently, had a moderate impact on the life-sciences. The emergence of bright, kinetically stable, luminescent lanthanide complexes however has changed the game. Time-gated (resolved) FRET (TR-FRET) technologies based on luminescent lanthanide cryptates such as HTRF® [2] are now well established. The translation of this technology in a microscopy setting was only established recently [3] due to the development of the very bright Lumi4®-Tb complex [2]. This terbium complex has now triggered a renewed interest in this type of microscopy [4] hosting, in principle, several advantages over established FRET microscopy techniques. Here we present multi-color TR-FRET microscopy with Lumi4®-Tb as the donor and either small molecule or quantum-dots as acceptors for the study of G-protein coupled receptor (GPCR) organisation and internalisation in living cells [4a]. Furthermore, we show the development of tuneable, very bright europium complexes and their use in TR-FRET microscopy [5]. Finally, live cell TR-FRET sensors with high dynamic ranges (up to 1000 %) to monitor conformational changes in GPCR’s upon ligand binding are presented [6].

[1](a) H. B. Beverloo, A. van Schadewijk, S. van Gelderen-Boele, H. J. Tanke. “Inorganic phosphors as new luminescent labels for immunocytochemistry and time-resolved microscopy.” Cytometry 11, 784 (1990) (b) G. Marriott, R. M. Clegg, D. J. Arndt-Jovin, T. M. Jovin. “Time resolved imaging microscopy. Phosphorescence and delayed fluorescence imaging” Biophys. J. 60, 1374 (1991). (c) L. Seveus et al. “Time-resolved fluorescence imaging of europium chelate label in immunohistochemistry and in situ hybridization” Cytometry 13, 329 (1992) [2] J. M. Zwier, H. Bazin, L. Lamarque, G. Mathis. “Luminescent lanthanide cryptates: from the bench to the bedside.” Inorg. Chem. 53, 1854 (2014). [3] H. E. Rajapakse et al.”Time-resolved luminescence resonance energy transfer imaging of protein-protein interactions in living cells. “Proc. Nat. Acad. Sci. USA 107, 1582 (2010). [4](a) O. Faklaris et al. “Multicolor time-resolved Förster Resonance Energy Transfer microscopy reveals the impact of GPCR oligomerization on internalization processes.” FASEB J. 29, 2235 (2015). (b) S. Linden et al. “Terbium-based time-gated Förster resonance energy transfer imaging for evaluating protein interactions on cell membranes” Dalton Trans. 44, 4994 (2015). (c) M. Rajendran & L. W. Miller. “Evaluating the performance of time-gated live-cell microscopy with lanthanide probes.” Biophys. J. 109, 240 (2015). [5](a) M. Delbianco et al. “Bright, highly water-soluble triazacyclonane europium complexes to detect ligand binding with time-resolved FRET microscopy.” Angew. Chem. Int. Ed. 53, 10718 (2014). (b) S. J. Butler et al. “Eurotracker® dyes: design, synthesis, structure and photophysical properties of very bright europium complexes and their use in bioassays and cellular optical imaging.” Dalton Trans. 44, 4791 (2015). [6](a) E. Doumazane, P. Scholler, L. Fabre, J. M. Zwier, E. Trinquet, J.P. Pin, P. Rondard. “Illuminating the activation mechanism and allosteric properties of metabotropic glutamate receptors.” Proc. Nat. Acad. Sci. USA 110, E1416 (2013). (b) P. Scholler et al, submitted.

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List of Participants

Surname Name

Contact Email

Abdollahi Elham

GSI Helmholtzzentrum fuer Schwerionenforschung GmbH Biophysics Darmstadt, Germany

[email protected]

Agam Ganesh

Ludwig-Maximilians-University Munich Chair of Physical Chemistry I. Munich, Germany

[email protected]

Alexiev Ulrike

Freie Universitaet Berlin Institute of Experimental Physics Berlin, Germany

[email protected]

Algar Russ

University of British Columbia Chemistry Vancouver, Canada

[email protected]

Arndt-Jovin Donna

Max Planck Institute for Biophysical Chemistry Research Group - Cellular Dynamics Goettingen, Germany

[email protected]

Aznauryan Mikayel

Aarhus University Interdisciplinary Nanoscience Center Aarhus, Denmark

[email protected]

Barth Anders


[email protected]

Bastiaens Philippe

Max Planck Institute of Molecular Physiology Dept. of Systemic Cell Biology Dortmund, Germany

[email protected]

Bergman Trygve

FCI-Frenzel Business Development Mainz, Germany

[email protected]

Bhuckory Shashi

Paris-Sud University Dept. of Physics Orsay, France

[email protected]

Birkedal Victoria

Aarhus University iNANO center Aarhus, Denmark

[email protected]

Böning Daniel

Max Planck Institute for the science of light Nanophotonics Erlangen, Germany

[email protected]

Borst Janwillem

Wageningen University Laboratory of Biochemistry Wageningen, Netherlands

[email protected]

Bouchaala Redouane

University of Strasbourg Laboratory of Biophotonics and Pharmacology Illkirch, France

[email protected]

Bremer Daniel

German Rheumatism Research Center Biophysical Analytics Berlin, Germany

[email protected]

147

Surname Name

Contact Email

Brodwolf Robert


[email protected]

Bruchez Marcel

Carnegie Mellon University Molecular Biosensor and Imaging Center Pittsburgh, USA

[email protected]

Cardoso Dos Santos Marcelina


[email protected]

Castellanos Milagros

Centro Nacional de Biotechologia - CSIC Macromolecular Stucture Madrid, Spain

[email protected]

Cerminara Michele

Forschungszentrum Juelich GmbH Institute of Complex Systems ICS-5 Molecular Biophysics Juelich, Germany

[email protected]

Chizhik Alexey

Georg-August-University Goettingen 3. Physical Institute - BiophysicsGoettingen, Germany

[email protected]

Chung Hoi Sung

National Institutes of Health Laboratory of Chemical Physics Bethesda, USA

[email protected]

Clarke David

STFC Central Laser Facility Research Complex at Harwell Didcot, UK

[email protected]

Coates Alexandra

Andor Technology Sales Belfast, UK

[email protected]

Cook Nathan

Max Planck Institute for Biophysical Chemistry Research Group - Cellular Dynamics Goettingen, Germany

[email protected]

Cordes Thorben

University of Groningen Zernike Institute for Advanced Materials Groningen, Netherlands

[email protected]

Craggs Tim

University of Bristol School of Biochemistry Bristol, UK

[email protected]

de Boer Marijn

University of Groningen Faculty of Natural Sciences Molecular Biophysics Groningen, Netherlands

[email protected]

Demchenko Alex

National Academy of Sciences of Ukraine Palladin Institute of Biochemistry Kyiv, Ukraine

[email protected]

Deussner Nina

Goethe-University Frankfurt/Main Institute of Physical and Theoretical Chemistry Frankfurt/Main, Germany

[email protected]

148

Surname Name

Contact Email

Diaz Sebastian

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Osnabrueck University Biology Osnabrueck, Germany

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Aarhus University Molecular Biology and Genetics Aarhus, Denmark

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National Institutes of Health Biophysical Chemistry Sestion Bethesda, USA

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Naval Research Laboratory and University of Maryland Bioengineering College Park, USA

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Technical University Dresden B CUBE - Center for Molecular Bioengineering Dresden, Germany

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University of California, Irvine Biomedical Engeneering Irvine, USA

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Albert-Ludwigs-University Freiburg Institute of Physical Chemistry Freiburg, Germany

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University of Debrecen Dept. of Biophysics and Cell Biology Debrecen, Hungary

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University of California, Los Angeles Chemistry and Biochemistry Los Angeles, USA

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Arizona State University School of Molecular Sciences Tempe, USA

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University of New Mexico Pathology Albuquerque, USA

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Shenzhen Institutes of Advanced Technology Res. Centre for Biosen. and Med. Instrum. Shenzhen, P.R. China

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Clemson University Physics and Astronomy Clemson, USA

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University of Strasbourg Laboratory of Biophotonics and Pharmacology Illkirch, France

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Ulbricht Carolin

Charité University of Medicine Berlin German Rheumatism Research Center Berlin Berlin, Germany

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Uri Asko

University of Tartu Institute of Chemistry Tartu, Estonia

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Western Kentucky University Physics Bowling Green, USA

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University of Groningen Zernike Institute for Advanced Materials Groningen, Netherlands

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Chalmers University of Technology Chemistry and Chemical Engineering Gothenburg, Sweden

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Institute of Organic Chemistry and Biochemistry Research Group - Organic Chemistry Prague, Czech Republic

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Hannover Medical School Cellular Neurophysiology Hannover, Germany

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University of Zurich Dept. of Chemistry Zurich, Switzerland

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University of Tehran Institute of Biochemistry and Biophysics Tehran, Iran

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158

Abstracts Index Name Presenting author

Abstract number (page number) Other contributions

Abstract number (page number)

Elham Abdollahi C-P50 (108) Ganesh Agam C-P51 (109) Ulrike Alexiev C-P52 (110) Russ Algar T39 (57) Donna Arndt-Jovin T19 (36), B-P41 (99) Mikayel Aznauryan A-P1 (59) Anders Barth B-P40 (98) T33 (51), A-P27 (85), C-P51 (109) Philippe Bastiaens T10 (27) T23 (40) Shashi Bhuckory E-P68 (126) Victoria Birkedal T17 (34) T2 (19), A-P1 (59) Daniel Boening A-P2 (60) Redouane Bouchaala E-P69 (127) Daniel Bremer E-P70 (128) Robert Brodwolf C-P52 (110) Marcel Bruchez T36 (54) E-P84 (142) Marcelina Cardoso Dos Santos E-P71 (129) Milagros Castellanos A-P3 (61) Michele Cerminara A-P4 (62) Alexey Chizhik T9 (26) T-19 (36), A-P17 (75) Hoi Sung Chung T7 (24) David Clarke T12 (29) Nathan Cook B-P41 (99) T24 (41) Thorben Cordes A-P5 (63), A-P35 (93),

D-P63 (121) Tim Craggs T26 (43) Marijn de Boer A-P5 (63) Alex Demchenko B-P42 (100) Sebastian Diaz T31 (49), E-P72 (130) Stephan Diekmann C-P53 (111) Mykola Dimura D-P61 (119) T29 (46) Franziska Doll C-P54 (112) Olga Doroshenko A-P6 (64) A-P28 (86) Christoph Drees E-P73 (131) Mateusz Dyla T2 (19) A-P21 (79) Rene Ebrecht C-P55 (113) Tobias Eilert B-P43 (101) T27 (44) Joerg Enderlein T3 (20), T9 (26), A-P15 (73),

A-P17 (75) Alessandro Esposito T13 (30) C-P56 (114) Lauren Field E-P74 (132) Erica Fiorini A-P7 (65) Theodorus Gadella T15 (32) T14 (31) Pablo Gracia A-P8 (66) Irina Gopich T5 (22) Alexander Goryashchenko E-P75 (133) T37 (55)

159

Name Presenting author Abstract number (page number)

Other contributions Abstract number (page number)

Charlie Gosse A-P9 (67) Timo Graen T3 (20) Enrico Gratton T11 (28) Helmut Grubmüller T3 (20) Jiajia Guo E-P76 (134) Kalina Haas C-P56 (114) T13 (30) Christian Hanke A-P6 (64) Quentin Hanley B-P44 (102) Andreas Hartmann A-P10 (68) T32 (50), A-P8 (66) Simon Hartmann A-P11 (69) Christina Heil A-P12 (70) Bjoern Hellenkamp T28 (45) A-P13 (71) Katherina Hemmen T4 (21) Jelle Hendrix E-P77 (135) A-P36 (94), B-P40 (98),

D-P67 (125) Niko Hildebrandt E-P78 (136) E-P68 (126), E-P71 (129),

E-P76 (134), E-P80 (138), E-P82 (140), E-P83 (141)

Bernhard Hochreiter E-P65 (123) Martin Hof D-P64 (122) Henning Hoefig E-P79 (137) Johannes Hohlbein T20 (37) T26 (43), D-P63 (121) Wolf Holtkamp T18 (35) Adam Hoppe T30 (48) Thorsten Hugel A-P13 (71) T28 (45), B-P47 (105) Gerhard Hummer T25 (42) Damla Inan A-P14 (72) Sebastian Isbaner A-P15 (73) Kees Jalink T14 (31) Zongwen Jin E-P80 (138) E-P83 (141) Thomas Jovin T24 (41) B-P41 (99), E-P72 (130) Eleni Kallis A-P16 (74) Narain Karedla A-P17 ( 75) T9 (26), T19 (36), A-P15 (73) Bettina Keller T6 (23) Reinhard Klement T3 (20) Dagmar Klostermeier T1 (18) A-P11 (69) Tamás Kovács B-P46 (104) Benedikt Kraemer T22 (39) Kevin Kramm A-P18 (76) Jakub Kubiak A-P19 (77) Ralf Kuehnemuth A-P20 (78) T4 (21), C-P59 (117) Don Lamb T33 (51) A-P36 (94), B-P40 (98),

C-P51 (109) Heinz Langhals B-P45 (103) Edward Lemke T38 (56) Karin Lenger D-P62 (120) Eitan Lerner A-P21 (79) D-P63 (121) Marcia Levitus T16 (33)

160



Diane Lidke T19 (36) E-P84 (142) Ajasja Ljubetič A-P22 (80) Irina Majoul C-P57 (115) Emmanuel Margeat A-P23 (81) Gerard Marriott T35 (53) Marisa Martin-Fernandez T12 (29) Martin Masip T23 (40) Igor Medintz E-P81 (139) T31 (49), E-P71 (129),

E-P 72 (130), E-P74 (132) Joseph Melinger A-P24 (82) T31 (49) Jens Michaelis T27 (44) A-P16 (74), B-P43 (101) Julia Molle A-P25 (83) Satish Babu Moparthi A-P26 (84) Peter Nagy B-P46 (104) Daniel Nettels A-P27 (85) Tuan Nguyen T34 (52) Hamsika Parimi E-P82 (140) Alexandra Petreto E-P83 (141) Thomas Peulen T29 (46) A-P19 (77), C-P59 (117),

D-P61 (119) Genevieve Phillips E-P84 (142) Evelyn Ploetz D-P63 (121) Aiswaria Prakash A-P28 (86) Manuel Prieto A-P29 (87) Sarah Rauscher T3 (20) Alexander Rittner A-P12 (70) Radek Šachl D-P64 (122) Peter Salonikidis Hugo Sanabria T4 (21) Enea Salsi A-P30 (88) Sangamesh Sarangamath A-P31 (89) Alexander Savitsky T37 (55) E-P75 (133) Max Schelski Michael Schlierf T32 (50) A-P8 (66), A-P10 (68) Sonja Schmid B-P47 (105) Johannes Schmid D-P65 (123) Herbert Schneckenburger C-P60 (118) Sven Schneider A-P33 (91) Klaus Schuermann C-P58 (116) Benjamin Schuler T8 (25) A-P27 (85) Claus Seidel C-P59 (117) T4 (21), T29 (46), A-P6 (64),

A-P19 (77), A-P20 (78), A-P23 (81), A-P28 (86), D-P61 (119)

Volodymyr Shvadchak A-P38 (96) Azmat Sohail A-P32 (90) D-P66 (124) Fabio Steffen A-P34 (92) A-P39 (97) Thomas Stockner D-P66 (124) A-P32 (90) Kateryna Trofymchuk E-P85 (143)

161



Carolin Ulbricht E-P86 (144) Asko Uri B-P48 (106) Wieb van der Meer T21 (38) Jasper van der Velde A-P35 (93) Niels Vandenberk D-P67 (125) Steven Vogel T21 (38), T34 (52) Petra Weber C-P60 (118) Daniela Wengler A-P36 (94) Matthew Wiens E-P87 (145) Marcus Wilhelmsson A-P37 (95) A-P31 (89) Dmytro Yushchenko A-P38 (96) Andre Zeug B-P49 (107) Meng Zhao A-P39 (97) Zahra Zolmajd B-P44 (102) Jurriaan Zwier E-P88 (146) A-P23 (81)

162

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