Spectroscopic and microscopic analysis of arrestin-rhodopsin interactions

vorgelegt von Master in molecular chemistry and biochemistry Florent Beyrière aus Pau, Frankreich

Von der Fakultät II- Mathematik und Naturwissenschaften der Technischen Universität Berlin zur Erlangung des akademischen Grades

Doktor der Naturwissenschaften - Dr. rer. nat. -

genehmigte Dissertation

Promotionsausschuss

Vorsitzender: Prof. Dr. rer. Nat. Budisa Gutachter: Prof. Dr. rer. Nat. Hildebrandt Gutachter: Prof. Dr. rer. Nat. Hofmann

Tag der wissenschaftlichen Aussprache: 29. Januar 2015

Berlin 2015 Abstract

2 Abstract

Abstract

G protein coupled receptors (GPCRs) are transmembrane receptors responsible for sensing and transmitting signals across the plasma membrane to G proteins. Agonist binding promotes receptor interaction with G proteins, thereby initiating signal transduction. Rhodopsin serves as a model for understanding GPCRs, as previous investigations have provided detailed information on its structure and interactions. Unlike the other GPCRs, rhodopsin´s ligand, 11-cis-retinal, is already bound to the inactive dark state. Light induced cis/trans isomerisation of the retinal triggers conformational changes in the receptor leading to its activation. Before the decay of the activated receptor into the aporeceptor opsin and free all-trans-retinal, G protein activation is blocked by receptor phosphorylation and binding of arrestin. Although different crystal structures of basal and pre-active arrestin have been reported, the transition to the high affinity arrestin-receptor complex is not yet understood. The first part of this thesis explores the common sequence shared by the four arrestin types and G protein of the rod cells (Gt). The four arrestin types are responsible for blocking the signal throughout the whole GPCR family, and all contain the motif E/DxLxxxGL, also present in the Gt C-terminus. This common motif is speculated to bind the receptor crevice in the same way as the Gt C-terminus. Binding measurements using an UV-Vis spectroscopic functional assay, carried out with a peptide library derived from this common motif, suggest that this common sequence acts as the activated receptor sensor in arrestin. In the second part of this work, the formation of the arrestin complex was investigated by means of Fourier transform infrared (FTIR) spectroscopy. A loss of ~5% of the beta sheet content of arrestin occurs upon the transition from pre-bound arrestin (pre-complex) to the high affinity complex, arguing that a small conformational change occurs upon complex formation in arrestin. Two different binding kinetics were observed upon formation of the high affinity complex. The data thus suggest the presence of two different pre-complex populations. Furthermore, dimerization of these pre-complexes is suggested to play a major role. Half of the high affinity complex population remains stabilized in the active receptor conformation, after decay. The other half of the receptor population decays to

3 Abstract

inactive opsin. The asymmetric complex decay shifts the functional arrestin- rhodopsin stoichiometry from one-to-one to one-to-two. In the last part of this work, organization of receptors in native disc membranes was analysed by means of single molecule technique. Individually fluorescent labelled receptors were tracked with a wide-field microscope. The data show that ~80% of receptors are immobile and ~20% of receptors are mobile with a diffusion coefficient of 0,005 μm2/s. The high number of immobile receptors and the low diffusion rate of the mobile receptors support an organization of rhodopsin in dimers and/or in racks.

4 Abstract in german

Abstract in German

G Protein-gekoppelte Rezeptoren (GPCRs) sind Transmembran-Rezeptoren, die dafür verantwortlich sind, chemische Signale zu empfangen und diese Signale durch die Plasmamembran an G-Proteine weiterzugeben. Die Bindung von aktivierenden Liganden (Agonisten) leitet die Interaktion des Rezeptors mit dem G- Protein ein und löst so die Signaltransduktion aus. Rhodopsin ist der am besten untersuchte GPCR und daher zum Modell dieser Rezeptorklasse geworden. Im Gegensatz zu anderen GPCRs ist der 11-cis-Retinal-Ligand im inaktiven Dunkelzustand bereits am Apoprotein Opsin gebunden. Die lichtinduzierte Isomerisierung des 11-cis-Retinals zum Agonisten all-trans-Retinal führt zu Konformationsänderungen und der Aktivierung des Rezeptors. Vor dem Zerfall des aktivierten Rezeptors in Opsin und freies all-trans-Retinal, wird die G-Protein Aktivierung durch Phosphorylierung des Rezeptors und der nachfolgenden Bindung von Arrestin geblockt. Obwohl Röntgenkristallstrukturen sowohl der basalen als auch der preaktivierten Arrestinkonformationen bekannt sind, ist die molekulare Organisation des Arrestin/Rezeptor Komplexes unklar. Im ersten Teil dieser Arbeit wurde die gemeinsame Sequenz der vier Arrestin Typen und dem G-Protein der Stäbchenzelle (Gt) untersucht. Die vier Arrestin Typen sind dafür verantwortlich, die Signalweiterleitung der gesamten GPCR-Familie zu blockieren und teilen mit dem C-Terminus des Gt ein gemeinsames E/DxLxxxGL Motiv. Es wird spekuliert, dass dieses Motiv des Arrestins in der gleichen Weise in der Bindungstasche des aktiven Rhodopsins bindet wie das Gt C-Terminus. UV-Vis- spektroskopische Bindungsmessungen von Peptiden, die aus diesem Motiv abgeleitet wurden, legen nahe, dass diese gemeinsame Sequenz tatsächlich in Arrestin als Sensor der aktivierten Rezeptoren fungiert. Im zweiten Teil dieser Arbeit wurde die Bildung des Arrestin-Komplexes mit Hilfe von Fourier-Transformations-Infrarot (FTIR) Spektroskopie untersucht. Die Messungen zeigen, dass der Übergang von initial Rezeptor gebundenem Arrestin (pre-complex) zum stabilen Arrestin/Rezeptor-Komplex (high-affinity complex) nur von geringfügigen Konformationsänderungen begleitet ist (5% Verringerung des Beta-Faltblatt-Anteils im Arrestin). Eine kinetische Auswertung dieses Übergangs deutet auf die Existenz von zwei unterschiedlichen initial gebundenen Arrestin

5 Abstract in german

Populationen hin, wobei eine Dimerisierung dieser Arrestin/Rezeptor-Komplexe dabei vermutlich eine wichtige Rolle spielt. Die FTIR-Messungen zeigen zudem, dass beim Zerfall des Arrestin-Komplexes eine Änderung der funktionelle Arrestin-Rhodopsin Stöchiometrie von 1:1 zu 2:1 erfolgt, wobei die eine Hälfte der Rezeptoren in einer Arrestin gebundenen, aktiven Rezeptorkonformation verbleibt, während die andere Hälfte der Rezeptoren zu inaktivem Opsin und all-trans-Retinal zerfällt. Im letzten Teil dieser Arbeit wurde die Diffusion fluoreszenzmarkierter Rhodopsinmoleküle in nativen Diskmembranen mittels Einzelmolekülmikroskopie verfolgt. Die Messungen zeigen, dass etwa 80% der Rezeptoren immobil sind und ca. 20% der Rezeptoren eine geringe Mobilität (Diffusionskonstante = 0,005 μm2/s) aufweisen. Der hohe Anteil immobiler Rezeptoren und die geringe Diffusionsgeschwindigkeit der mobilen Rezeptoren unterstützen Modelle einer Organisation von Rhodopsin in Di- bzw. Oligomeren.

6 Abbreviations

Abbreviations

ABS Arrestin binding spectrum Abs Absorption AFM Atomic force microscopy Arr Arrestin ATP Adenosine 5´-triphosphat Atto 647 Atto 647 N maleimide BSI Bue shifted intermediate cGMP Cyclic guanosine monophosphate CNGC cGMP gated ion channel Fibrolast-like cell line derived from monkey kidney COS cells tissue CT C-terminal tail DMSO Dimethyl sulfoxide DTT 1,4-Dithiothreitol EDTA Ethylenediaminetetraacetic acid EPR Electron paramagnetic resonance Equ Equation Fig Figure FTIR Fourier transformation infrared GC Guanylate Cyclase GCAP Guanylate Cyclase activating proteins GDP Guanosine diphosphate GMP Guanosine monophosphate GPCR G protein coupled receptor GRK1 G protein receptor kinase 1 Gt Transducine Gt alpha CT C-terminal tail Gt alpha peptide Gt alpha HAA High affinity analogue Gt alpha peptide GTP Guanosine triphosphate GTP Guanosine triphosphate Gtα Gt alpha subunit Gβγ Gt beta-gamma heterodimere H8 Helix 8 HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid N,N'-dimethyl-N-(iodoacetyl)-N'-(7-nitrobenz-2- IANBD oxa-1,3-diazol-4-yl)ethylenediamine IPTG Isopropyl 1-thio-ß-D-galactopyranoside Kd Dissociation constant

7 Abbreviations

Meta I Metarhodopsin I Meta II Metarhodopsin II MSD Mean square displacement NMR Nuclear magnetic resonance OD optic density Ops Close conformation of opsin Ops* Active conformation of opsin p44 Splice variant of arrestin-1 PBS Peptide binding spectrum PDB Protein data bank PDE Phosphodiesterase PMSF Phenylmethanesulfonylfluoride p-Ops Phosphorylated opsin P-R* Phosphorylated activated receptor p-rhodopsin Phosphorylared rhodopsin QD Quantum Dot R Receptor R* Activated receptor ROS Rod outer segment RPE Retinal pigment epithelium TM Transmembrane alpha helix Tris-HCl Tris(hydroxymethyl)aminomethane hydrochloride

8 Contents

Contents

Abstract ...... 3 Abstract in German ...... 5 Abbreviations ...... 7 Contents ...... 9 1. Introduction ...... 13 1.1 G Protein-Coupled Receptors ...... 13 1.2 The retina and the rods ...... 14 1.3. Visual signal transduction ...... 16 1.3.1 Activation ...... 16 1.3.2 Recovery ...... 18 1.4 Light induced translocation of signalling proteins ...... 19 1.5 Rhodopsin ...... 20 1.5.1 Rhodopsin structure and microdomains ...... 21 1.5.2 Photointermediates and rhodopsin activation ...... 23 1.5.3 Active conformation of rhodopsin ...... 26 1.5.4 Retinal Channel ...... 28 1.5.5 Rhodopsin organization in disc membrane ...... 29 1.6 Arrestin ...... 31 1.6.1 Basal arrestin structure ...... 32 1.6.2 Arrestin binding ...... 34 1.6.3 Pre-active structure of arrestin ...... 35 1.7 Aim of the project ...... 39 2. Material and Methods ...... 41 2.1 Isolation of Rod Out of Segment ...... 41 2.2 Isolated Membranes ...... 42 2.3 Phosphorylation of Rhodopsin ...... 42 2.4 Arrestin and P44 expression...... 43 2.5 Arrestin purification ...... 43 2.6 P44 purification ...... 44 2.7 Specific labelling of protein ...... 45 2.8 UV-vis absorption spectroscopy ...... 47 2.9 Extra MII assay ...... 49

9 Contents

2.10 Fourier Transform Infrared Spectroscopy ...... 49 2.11 Pulled Down assay ...... 53 2.13 Electron Microscopy ...... 54 2.14 Single Molecule Fluorescence Microscopy ...... 54 3. Results ...... 59 3.1 Peptides derived from arrestin finger loop stabilized activated rhodopsin ..... 59 3.1.1 Arrestin-1 finger loop peptide series ...... 62 3.1.2 Arrestin-2 finger loop peptide series ...... 64 3.1.3 Arrestin-3 and arresti-4 finger loop peptide series ...... 65 3.1.4 G alpha C-terminus peptides ...... 66 3.2 Arrestin complex formation studied by Fourier transform infrared spectroscopy ...... 67 3.2.1 Influence of the phosphorylation on rhodopsin ...... 68 3.2.2 Arrestin binding difference spectrum ...... 72 3.2.3 Arrestin binding spectrum ...... 74

3.2.4 Arrestin binding spectrum (ABS) in D2O ...... 75 3.2.5 Arrestin binding kinetic ...... 77 3.2.6 Complex formation of p44 with phosphorylated rhodopsin ...... 80 3.2.7 Complex formation of p44 with unphosphorylated rhodopsin...... 83 3.2.8 Decay of the arrestin complex ...... 86 3.2.9 To which products decays the complex? ...... 89 3.3 Rhodopsin diffusion in native disk membrane ...... 93 3.3.1 Samples quality control ...... 93 3.3.2 Rhodopsin tracking with single fluorescence microscopy ...... 95 3.3.3 Phosphorylated rhodopsin tracking ...... 96 3.3.4 Tracking of quantum-labelled Opsin ...... 98 4. Discussion ...... 101 4.1 The finger loop derived peptides ...... 101 4.1.1 Finger loop as Meta II sensor ...... 101 4.1.2 Structure of bound arrestin finger loop and its interaction with activated rhodopsin ...... 103 4.2 Formation of high affinity arrestin complex investigated with FTIR spectroscopy ...... 107 4.2.1 At high concentration, arrestin is already pre-complexed in dark state . 107

10 Contents

4.2.2 Assignment of the arrestin binding spectrum (ABS) ...... 109 4.2.2.1 The 1660 cm-1 Band ...... 109 4.2.3 Arrestin loses beta sheet content upon high affinity complex formation ...... 110 4.2.4 High affinity complex is formed in a two step mechanism ...... 112 4.2.5 Possible mechanisms of arrestin binding ...... 116 4.2.6 Stochiometry change during formation and decay of the high affinity complex ...... 119 4.2.7 Model of high affinity complex decay ...... 121 4.3 Rhodopsin organization in disc membrane ...... 123 References ...... 125 Published journal articles ...... 139 Acknowledgements ...... 141 Selbständigkeitserklärung ...... 143

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12 1. Introduction

1. Introduction

1.1 G Protein-Coupled Receptors

G protein-couple receptors (GPCRs) are a large family of transmembrane receptors present in eukaryotic organisms. The entire family of GPCRs share a structurally homologous seven-transmembrane helix (TM) topology and a common activation mechanism (K P Hofmann et al. 2009). This transmembrane structure allows the receptors to connect the extracellular environment to the inner cell. GPCRs are responsible for sensing molecules outside of the cell and trigger several internal signal transduction pathways. They are at the start of several of molecular cascade reactions in a wide variety of cellular systems. These receptors bind to a large variety of ligands such as hormones, neurotransmitters, odorants, light-sensitive compounds or gustatory substances. Agonist binding induces distinct conformational changes in protein and promotes receptor interactions with the heterotrimeric G proteins at the intracellular side of the receptor. Activation and dissociation of G α- and βγ- subunits promote signal transduction through their interactions with various downstream effector proteins thus initiating the signalling cascade (see signal transduction chapter). Activated GPCRs are then phosphorylated by receptor kinase. Once phosphorylated, the protein arrestin, subject of this work, can bind the receptor and thus block further G protein activation. Arrestin interacts with numerous other proteins, initiating other signalling pathways (Xiao et al. 2007). GPCRs represent ~ 4% of the entire protein-coding human genome (Gershengorn and Osman 2001). Due to the strategic role of GPCRs in cell signalling and the many diseases which are caused by these receptors, a great interest has been focused on them in the last decades. GPCRs represent more than 30% of all modern medicinal drug target (Hopkins and Groom 2002). The human GPCR superfamily contains five main GPCR families: rhodopsin-, secretin-, adhesion-, glutamate- and frizzled/taste2-like. The rhodopsin family (or class A GPCRs) is by far the largest. Rhodopsin responds to light and not directly to the binding of ligands. However rhodopsin shares homologous topology and mechanism similarities to ligand activated GPCRs. Rhodopsin, present in high quantity in retinas, easy to extract and accessible to biophysical techniques, became therefore a model for all GPCRs.

13 1. Introduction

1.2 The retina and the rods

Figure 1.1: The eye and the retina. A. The human eye cross-sectional view: 1) cornea, 2) iris, 3) lens, 4) pupil, 5) sclera, 6) retina, 7) vitreous body, 8) optic nerve. B. Schema of retina, section view: 1) rod cells, 2) cone cells, 3) horizontal cells, 4) bipolar cells, 5) amacrine cells, 6) ganglion cells, 7) axons of optic nerve, 8) retinal pigment epithelium, 9) sclera, 10) choroid. The light goes through the transparent inner layer and middle layer of retina and is detected by the photoreceptor rods and cones of the outer layer. The electric signal induced by light absorption is sent through the complex interconnected neurons network to the optic nerve. A modified from www.mjc.edu and B taken from wikispaces.com.

The vertebrate eye, organ of vision (Fig. 1.1, A), is principally composed of a cornea (1), an iris (2), a flexible lens (3) and retina (6). It is surrounded with a white tissue, the sclera (5) and the chamber is filled with a transparent vitreous body (7). The cornea and the lens are in charge of catching and focussing the light on the retina. The iris is responsible for regulating the amount of light by controlling the size of the pupil (4), the hole in the centre of the iris. The retina is a layer of tissues coating the inner surface of the eye and is responsible for detecting the light and transmitting the electric signal to the optic nerve (8) (Rodieck 1998). The retina is composed of a complex system of cells organized in layers. This stack of photoreceptor cells and neuronal cells can be summarised with an assembly of three layers. From the surface of the retina to the bottom of the eye: inner, middle and outer layer (Fig. 1.1, B). The outer layer, in the bottom of the retina, located directly on the sclera (9) and on the choroid cells (10) (meshwork of blood vessels), is constituted of rod cells and cones cells lying on a bed of melanin-containing epithelium cells (8). Rods and cones are light sensitive cells and are called

14 1. Introduction

photoreceptors. They convert the photon they capture into an electric signal. The middle layer, composed of horizontal cells (3) and bipolar cells (4), and the outer layer constituted of amacrine cells (5) and ganglion cells (6) are interconnected by synapses and are charged to transmit and treat the electric signal produced by the photoreceptors. The output takes the form of action potentials in the ganglion cells, whose axons form the optic nerve (7). Light passes through several transparent nerve layers to reach the rods and cones. Both photoreceptors contain visual pigment in their outer segment (Fig. 1.2 A). Visual pigments are composed of a protein, opsin, and a chromophore, 11-cis-retinal. The absorption of a photon by the visual pigment induces a cascade reactions leading to the hyperpolarization of the photoreceptor and induces a signal to the nerves. Rods function mainly in dim light and provide black/white vision while the colours are detected by the cones. Cones are less sensitive but have the possibility to detect different wavelengths. Variations of strategic amino acids of the protein determine its sensitivity to different wavelengths. Each photoreceptor contains only one type of visual pigment protein. There are three different cone cells (L, M and S) with each type containing a different ´´cone rhodopsin´´ allowing each type of cone to detect distinct wavelengths (commonly red, green and blue.). The visual pigment of the rod is called rod rhodopsin, commonly rhodopsin. Bovine rods are 50 μm long cylindrical cells, aligned in direction of the lens on the outer surface of the retina (Fig. 1.2 A). They are composed of an outer segment (ROS) connected through a cilium to the inner segment and finally synaptic terminals connected to neuronal cells (bipolar cells and horizontal cells). The inner segment contains a nucleus and all usual cell organelles cells necessary for the metabolism of the cell. Rhodopsins are synthesized in the inner segment of the rod and transported to the outer segment where they are incorporated into disc membranes (Fig. 1.2, B, C and D). These disks are derived from the plasma membrane of the cell but are disconnected from it. They are stacked, each one on top of the other, forming a parallel lamellar structure of 500 to 2000 double-membrane close disc vesicles (disks) located in the ROS (Daemen 1973). Rhodopsin constitutes ~85% of protein fraction of the ROS. The stacking of the disks, and the high density of rhodopsin, allows for optimal light absorption. It provides the rod with high sensitivity to light, with the capacity to respond to a single photon (Rieke and Baylor 1998).

15 1. Introduction

Figure 1.2: The rod cell. A. Rod cell is composed of outer segment connected to the inner segment through a cilium and finally of synaptic terminals in contact with neurones. The nucleus is located in the inner segment. The outer segment is principally composed of a staking of disks. (Scheme modified from: www.mjc.edu). B. Scheme of a Disk, (cross sectional view). Rhodopsin molecules are embedded in lipid bilayer disk membrane. C. Representation of rhodopsin molecule in lipid bilayer. (PDB: 1U19).

1.3. Visual signal transduction

1.3.1 Activation

Phototransduction serves as a model of signal transduction for most GPCRs. Phototransdution occurs in the outer segment of rods and cones. This mechanism converts the light into a neural signal. It can be summarised in five steps. The first three reactions occur at the disk membrane where the rhodopsin and the membrane associated proteins (transducin, phosphodiesterase) are located. The last two reactions take place in the cytosol and at the plasma membrane (Fig. 1.3).

Rhodopsin absorbs a photon and activates the Gt-protein transducin, which further amplifies the signal. Dissociation of transducin triggers further reactions with downstream effectors leading to the hyperpolarization of the photoreceptor (Lamb and Pugh Jr. 2006).

16 1. Introduction

Figure 1.3: Illustration of phototransduction cascade in rod. After absorption of a photon (hν) by the rhodopsin (R), light activated rhodopsin (R*) catalyzes the exchange of GDP for GTP on the Gtα subunit of transducin (G). Gtα subunit dissociates, migrates and binds the PDE at its γ-subunit. Activated PDE hydrolyzes cGMP reducing drastically the cGMP (cG) concentration in the cytosol which leads to the closure of the cGMP gated ion channel (CNGC) at the plasma membrane. The ion inward flux (Na+, Ca2+) is blocked, reducing the circulating electrical current. The Na+/Ca2+,K+ exchanger continues to work and continues to pomp Ca2+ out leading to the hyperpolarization of the membrane. Taken from (Pugh and Lamb 2000).

The first step of phototranslation is the detection of light and the activation of rhodopsin. Embedded in the disks, rhodopsin is composed of the apoprotein opsin and the covalently bound chromophore 11-cis-retinal. The 11-cis-retinal acts in the dark as a inverse agonist. Absorption of a photon isomerizes the retinal to the strong agonist all-trans-retinal which induces conformational changes in the protein leading to the activated receptor (R*, Metarhodopsin II or Meta II, see rhodopsin chapter for details). The Meta II conformation of rhodopsin results in the presence of a crevice in the receptor on the intracellular side, where transducin can bind. The second step of signal transduction is the activation of transducin. The heterotrimeric form of transducin is composed of α, β and γ subunits where a nucleotide GDP is bound at the Gtα subunit. Interaction of the transducin with Meta II induces the GDP-GTP exchange catalysis at the Gtα subunit and finally the dissociation of the Gtα of the

Gtβγ subunits. The third and fourth parts of the cascade reaction consist of the triggering of the enzyme cGMP phosphodiesterase (PDE) and the hydrolysis of cGMP. Activated Gtα binds PDE which starts to hydrolyse cGMP to inactive 5´-GMP. cGMP is synthesised by guanylate cyclase (GC). The full activation of the

PDE is achieved after the binding of a second Gtα to PDE. The signal is amplified downstream of the reaction cascade. In vitro, one Meta II activates over 1000 molecules transducin per second (M Heck and Hofmann 2001). Two activated G

17 1. Introduction

alpha activate thereafter one PDE molecule. In vitro, one PDE hydrolyzes over 4000 molecules of cGMP per second. In the absence of light, PDE is inactive. The absorption of a photon ultimately induces the activation of over 40,000 cGMP molecules, thereby decreasing drastically the cytosolic cGMP concentration (see Arshavsky, Lamb, and Pugh 2002). The last step of signal transduction is the closure of the ion channel at the plasma membrane and the hyperpolarization of the membrane, leading to the electrical response of the cell. In dark conditions, cGMP binds the cGMP gated ion channel and keeps it open. The decrease of cytosolic cGMP concentration induces the closure of the cGMP gated ion channel. The inward current of Na+ and Ca2+ ions is blocked, reducing the circulating current. However the Na+/Ca2+, K+ exchanger continues to pump cations (Ca2+) out of the cell resulting in the hyperpolarization of the membrane, which reduces the rate of release of glutamate neurotransmitters at the rod synapse, thereby transmitting the signal to the neurons (horizontal and bipolar cells)(Marie E Burns and Pugh 2010; Rodieck 1998).

1.3.2 Recovery

After the light stimulus, all the activated intermediates of the signalling pathways have to be inactivated rapidly and restored to the basal state level so that the absorption of a new photon can activate the signal cascade again (see Arshavsky,

Lamb, and Pugh 2002). Thus, R* and activated Gα bound to PDE have to be inactivated. They are both deactivated through interactions with outer segment proteins. R* is deactivated by arrestin, after the phosphorylation of the receptor with the rhodopsin kinase (GRK1). The binding of arrestin to R* blocks completely the interaction of G protein with the receptor and thus hindered the activation of G protein. The interaction of arrestin with rhodopsin is the object of this thesis and will be described in detail later (see arrestin chapter). After decay of Meta II to apoprotein Opsin and all-trans-retinal ligand, the receptor is dephosphorylated by the proteinphosphatase 2A (Palczewski et al. 1989) and regenerated to rhodopsin with 11-cis-retinal (see chapter 1.5.4). The second component of phototransduction inactivation is the hydrolysis of GTP bound to the Gα subunit, by the intrinsic

GTPase activity. The hydrolysis of GTP to GDP drives the dissociation of the Gα subunit from the PDE, and the subsequent return of the effector to its inactive state. 18 1. Introduction

The inactive PDE is no longer able to hydrolyse cGMP. cGMP feedback is regulated by Ca2+ dependent proteins (GCAP, guanylate cyclase activating protein). Low Ca2+ concentration activates GCAP which in turn activates the guanylate cyclase (GC). Therefore the cellular concentration of cGMP, synthesised by guanylate cyclase (GC), increases rapidly and is restored to its resting state level, leading to the reopening of the cGMP gated ion channel. The Ca2+ concentration in the cell is thus reestablished and the rod cell is depolarized as before the illumination (Hofmann et al. 2006).

1.4 Light induced translocation of signalling proteins

The rod cell is able to sense and amplify the signal for one photon, but it has also the ability to adjust its sensitivity of light to different light levels (see Burns and Baylor 2001). The translocation of signalling proteins inside the cell contribute to the adaptation of the photoresponse to different light conditions. Under bright light, transducin and arrestin translocate into vertebrate rod cells (Arshavsky 2003). In the dark, transducin is present in high concentrations in the outer segment (~80- 90% of total amount) and moves to the inner segment upon light (Fig. 1.4). The higher concentration of transducin in the dark enables rods to produce amplified photoresponses including those induced by single photons.

Figure 1.4: Illustration of light driven translocation of transducin and arrestin. A. In dim light, transducin (blue spheres) is principally located in the outer segment of rod where less than 7% of arrestin (green spheres) is concentrated in the ROS. In dark state, arrestin is mainly located in the inner segment. B. Under bright light condition, transducin migrates to the inner segment to avoid saturation of the phototransduction. Arrestin translocates in the opposite direction to the ROS to quench the transducin activity. Figure modified from (Calvert et al. 2006). 19 1. Introduction

In bright light, as during daytime, transducin migrates out of the ROS. The migration of transducin reduces its concentration in the ROS (to ~10-20%), leading to a reduction of the transducin activation amount (Pearring et al. 2013) In contrast, arrestin has a low concentration in the outer segment in the dark (<7%) and is principally situated in the inner segment and moves in the opposite direction in light. Under bright light, arrestin also contributes to light adaptation and migrates to the ROS where visual signal transduction takes place (Fig. 1.4). A higher concentration of arrestin in ROS (~80%) increases the arrestin binding rate and blocks the binding of transducin. Fewer transducin molecules are thus activated, leading also to a reduction of the photosensitivity of the rod cell (Strissel et al. 2006). The high arrestin concentration in the ROS could also accelerate the recovery of the rhodopsin by speeding the deactivation of activated receptors (Calvert et al. 2006).

1.5 Rhodopsin

Rhodopsin is the light sensitive receptor present in rod cells. It consists of the membrane apoprotein opsin and a covalently bound chromophore, 11-cis-retinal. The retinal, a derivate of vitamin A, is responsible for detecting light and transmitting the signal to opsin. The major difference compared to other GPCRs is that the ligand is already bound to rhodopsin in the inactive dark sate. The rhodopsin is activated only after absorption of a photon by the chromophore, and the isomerisation of the retinal to the agonist form. Nevertheless rhodopsin shares homologous topology and similarities to ligand activated GPCRs (see Rosenbaum et al. 2009)(Hofmann et al. 2009). Sequence alignment of GPCRs shows conserved amino acids, suggesting a key role of these motifs for protein function and stability. The different crystal structures of GPCRs reveal the three dimensional representation of these amino acids. The conserved residues are not close to each other in the amino acid sequence, but are located in a neighbouring special environment, and build interactions together. Upon receptor activation, these motifs can change from inactive to active conformation and thereby form functional domains called microswitches. They are strategic molecular switches that can define conformational changes within the receptor (S. O. Smith 2010).

20 1. Introduction

1.5.1 Rhodopsin structure and microdomains

Bovine rhodopsin is composed of a chain of 348 amino acids featuring seven transmembrane helixes (TM) connected to each other by loops, and a cytoplasmic helix 8 following the transmembrane helix 7. The helix 8 is elongated on the membrane and the palmitoylated cysteines (Cys322 and Cys323) at its end anchor the C-terminus to the disk membrane. The C-terminus plays a major role for arrestin binding (see chapter 1.6.1). At the extracellular side (intradiscal), the N-terminus along with the three extracellular loops, form the ``retinal plug´´. This compact domain is composed of short beta sheets at the extracellular loop 2 and at the N-terminus and serves to cover the retinal binding pocket (Janz, Fay, and Farrens 2003). A conserved disulfide bond in the same region between Cys110 and Cys187 stabilizes the correct folding of rhodopsin. Crystal structure of bovine rhodopsin in the dark state, the inactive state, was the first GPCR structure to be solved (K Palczewski et al. 2000)(Okada et al. 2004). High resolution of the three dimensional structure reveals the overall structure of rhodopsin and the presence of domains of specific interacting residues (Fig. 1.5). The retinal is located in the middle of the protein in a polar pocket. The retinal is covalently bound by a Schiff base to the Lys296 in the middle of TM7. In the dark state, negatively charged Glu113 acts as a counter ion and stabilizes the protonated Schiff base. It forms a salt bridge with the protonated Schiff base which leads to a stabilization of the retinal and maintains the receptor in an inactive conformation. Located next to the beta ionone ring is the TM3-TM5 microdomain. It is composed of a network of neighbouring hydrogen bonds including Glu122, Trp126 and His211 which connect TM3 and TM5. On the intracellular side of rhodopsin, there are two microdomains formed with conserved sequences. The TM3 contains the highly conserved (E/D)RY motif. These residues form an important microdomain, called the ionic lock, connecting TM3 and TM6. It is composed of Arg135 and Glu134 on TM3 which build a hydrogen bond network with residues Glu247 and Thr251 on TM6. The ionic lock holds the TM3 and TM6 closed and thus stabilizes the inactive state of rhodopsin.

The second conserved motif, the NPxxY(x)5,6F, provides two constrains, the TM1- TM2-TM7 network and the TM7-H8 interaction. The amino acids Asn302 on TM7, Asn55 on TM1 and Asp83 on TM2 form a hydrogen bond network connecting the

21 1. Introduction

three helices (TM1-TM2-TM7). Finally the electrostatic interactions between aromatic residues Tyr306 on TM7 and Phe313 on H8 constrain an elbow form of the loop between TM7 and H8.

Figure 1.5: Rhodopsin structure and functional mircrodomains. Rhodopsin is composed of seven transmembrane helixes and a cytoplasmic helix (H8). Two cysteins at the end of H8 are palmytoylated (light pink) and keep C-terminus and H8 anchored in the membrane. 11-cis-retinal (red) is covalently bound by Schiff base to Lys296 (orange). The Schiff base is stabilized by E113 (orange) in dark and by E181 (orange) in activated state which act as counterions. A retinal plug (pink), composed of the N-terminus and intracellular loops, forms short beta sheets and shields the retinal. The whole structure is principally stabilized by four domains. TM3-TM5 network (turquoise) keep TM3 and TM5 together and is composed of a hydrogen bond network between E122, W126 and H211. The conserved motif NPxxY(x)5,6F is involved in the TM1-TM2-TM7 network and in the TM7-H8 microdomain. Y306 and F313 (purple) have an aromatic stacking and constrain TM7 and H8. N55, D83 and N302 build together a hydrogen bond network and keep TM1 TM2 and TM7 together. The ionic lock (green) composed of E134 and R135 at TM3, from the conserved (E/D)RY motif, and the residues E247 and E251 at TM6 build hydrogen bonding network and keep TM3 and TM6 together. Y223 and K231 from the conserved motif Y(x)7K(R) interact after illumination with the ionic lock. PDB Data: 1U19, dark rhodopsin.

22 1. Introduction

1.5.2 Photointermediates and rhodopsin activation

Retinal forms a conjugated π-electron system and absorbs at λmax= 380 nm in solution. In rhodopsin, the 11-cis-retinal is bonded via protonated Schiff base to Lys296. The positive charge of the Schiff base decreases the electron delocalization along the conjugated retinylidene chain and red-shifts the absorption to at 498 nm. The Schiff base counterion Glu113 stabilizes the protonated Schiff base, regulates the wavelength absorption and also maintains the receptor in an inactive conformation. Residues in the retinal binding pocket also have a big influence on the absorption properties and their interactions drive the protein conformational change. The retinal is switched from an inverse agonist to a potent agonist. The energy is transmitted from the chromophore to the protein moiety through the retinal/protein interactions and subsequently the protein moiety undergoes a series of conformational changes leading ultimately to the active signalling state Meta II (Fig. 1.6). Isomerization leads first to a series of consecutive short-lived states: Photorhodopsin (570 nm), Bathorhodopsin (543 nm), BSI (blue shifted intermediate, 477 nm) and Lumirhodopsin (497 nm). These photointermediates appear in the pico to nanosecond timescale. All conformational changes occur at the residues close to the retinal. The reactions are too fast for the amino acid side chains to rearrange in the protein binding site (Ernst et al. 2014).

Figure 1.6: Retinal cycle in rhodopsin. Photon energy is absorbs and used to convert inverse agonist 11-cis-retinal of rhodopsin into full agonist all-trans- retinal. Isomerization leads within femtoseconds to photorhodopsin. The distorted all-trans chromophore relaxes through Bathorhodopsin, BSI and Lumirhodopsin within nanosecond scale to reach finally Meta I (478 nm). Deprotonation of Schiff base and large conformational changes lead within milliseconds to Meta II (380 nm) which is in equilibrium with Meta I. Meta II is the active conformation (R*) and can bind transducin, kinase and later arrestin. Decay Meta II leads to the apoprotein opsin and the all-trans-retinal. Within minute scale, retinal Schiff base is hydrolysed and the retinal released. Meta I can also decay through Meta III (465 nm). Meta III is an anti/syn thermal isomerisation of the retinal Schiff base and decays also to opsin and all-trans-retinal.

23 1. Introduction

Retinal is then restore to 11-cis-retinal into the retina and can thereafter regenerate rhodopsin. Taken from (Ernst et al. 2014). After microseconds the longer-lived Metarhodopsin I (Meta I) states arises followed after milliseconds to Metarhodopsin II states (Meta II). Meta I and Meta II states are in equilibrium. They absorb at 478 nm and 380 nm respectively. Meta I does not involve large backbone movement and remains in a conformation near to that of the rhodopsin dark state (Ruprecht et al. 2004). The deprotonation of the retinal Schiff base and protonation of the counterion Glu113 characterises Meta II and blue-shifts the absorption of the receptor to 380 nm. Meta II is formed of different substates containing the active conformation (R*). Meta II can activate transducin and later bind kinase and arrestin. The MetaI/MetaII equilibrium shifts through pH and temperature (Parkes and Liebman 1984a). In condition favouring Meta I, the binding of Gt alpha, arrestin or Gt alpha analogous peptide, mimicking the Gt alpha C-terminus key binding, can bind the receptor and stabilize the Meta II conformation (Extra Meta II, see Methods)(Herrmann et al. 2006)(K P Hofmann 1985).

Figure 1.7: Activation and photointermediates of rhodopsin. Absorption of light isomerises retinal to its all-trans configuration. Relaxation of the early photointermediates (Batho and Lumirhodopsin) leads to the metharhodopsin conformations. Meta I conformation is reach after a few microseconds and presents only small conformational changes compared to rhodopsin. Proton transfer from the conterion to the Schiff base leads to Meta IIa. A major absorption shift is observed (480nm to 380nm). Tilt out of TM6 and elongation of TM5 conducts to the formation of Meta IIb. Motion of the two helixes builds a crevice into the receptor permitting the binding of binding partners (transducin, arrestin). Protonation of Glu134 at the ERY motif induces the Meta IIbH+ conformation. Both Meta IIb conformations are the activated form of rhodopsin. Metasubstates decay within minutes to the apoprotein opsin. Schiff base is hydrolysed and all-trans-retinal is released. The apoprotein opsin exists also in equilibrium between an inactive and active conformation called opsin and opsin* respectively (Ops and Ops*). They are in a pH-dependent equilibrium and are thought to have similar conformation as the inactive rhodopsin or Meta II conformation but without ligand and 24 1. Introduction

have therefore intrinsic properties of the respective active and inactive conformation. Modified from (Hofmann et al. 2009). Meta IIa, Meta IIb and Meta IIbH+ are in equilibrium and arise sequentially in a few milliseconds (Fig. 1.7.). Larger conformational modification and proton uptake occurs upon Meta I/Meta II transition within the Meta II substates. Fourier transform spectroscopy (FTIR) and electronic paramagnetic resonance (EPR) studies could determine the conformational changes, the key residues and the sequence upon they arise (Altenbach et al. 2008)(Mahalingam et al. 2008)(Knierim et al. 2007). A proton transfers from Schiff base to the counterion Glu113 leads to formation of Meta IIa (M Mahalingam et al. 2008). The Schiff base shifts its centre of interaction from Glu113 to Glu181. TM6 can then rotate and tilt outward, accompanied by a short back movement and an elongation of TM5 which results in Meta IIb (Knierim et al. 2007). The ionic lock is broken and Arg135 interacts with Tyr223, while

Glu134 and Thr251 interact with Lys231. Tyr223 and Lys231 of the Y(x)7K(R) conserved motif on TM5 act as microswitch and fix the TM6 in an outward position and thus stabilize the Meta IIb conformation. Crystal structures also report a structural modification on the NPxxY(x)5,6F motif. The electronic interaction of Tyr306 with Phe313 is disrupted. Tyr306 rotates and interacts with residues below Arg135. It builds a large water-mediated hydrogen bonding network with Ser298, Asp83, Asn302, Met257, Tyr223 and Arg135 permitting stabilization of Arg135. Ultimately Meta IIbH+ is stabilized by proton uptake to Glu134 at the ERY motif on TM3 (M Mahalingam et al. 2008)(K P Hofmann et al. 2009)(Knierim et al. 2007). The apoprotein opsin also exists in equilibrium between an inactive and active conformation called opsin and opsin* respectively (Ops and Ops*). They are in a pH-dependent equilibrium and are thought to have similar conformation to the inactive rhodopsin or Meta II conformation, respectively, but without ligand, and therefore have intrinsic properties of the respective active and inactive conformation (R Vogel and Siebert 2001). Alternatively part of the Meta I population can also decay through Meta III (465 nm). Meta III is an anti/syn thermal isomerisation of the retinal Schiff base and also decays to opsin and all-trans-retinal (M Heck et al. 2003) (Reiner Vogel et al. 2003). Meta II decays to opsin and all-trans-retinal within minutes. Hydrolysis of retinal Schiff base releases all-trans-retinal from the protein. The released all-trans-retinal is reduced to retinol by the dehydrogenase enzyme and then transported into the retinal pigment epithelium cells at the base of the rods, to be recycled to

25 1. Introduction

11-cis-retinol. In these cells, the all-trans-retinol is isomerised to 11-cis-retinol by the retinoid isomerase RPE65. The retinal is finally oxidized to 11-cis-retinol and transported back to the rods to regenerate the receptor to its dark state rhodopsin (Kiser, Golczak, and Palczewski 2014).

1.5.3 Active conformation of rhodopsin

The crystal structure of ligand-free activated opsin (opsin*) alone and in complex with a Gt alpha analogue peptide, and also a structure of opsin soaked with all-trans-retinal (Park et al. 2008)(Scheerer et al. 2008)(Choe et al. 2011) were solved some years after the dark state rhodopsin. They all revealed a similar structure related to the active conformation Meta II. The comparison of the dark and active conformations indicates changes in some domains which take place upon photoactivation. These microdomains have crucial functional role in the activation of the receptor and act as microswitches. In comparison with dark rhodopsin, active conformation structure shows interaction modifications at the retinal binding pocket. The network of hydrogen bonds at the retinal counterion is rearranged upon retinal isomerization and receptor activation. The distance between the Schiff base and Glu113 increases while the distance between the Schiff base and Glu181 decreases (Fig. 1.8, A and B). The Schiff base interaction shifts from Glu113 to Glu181 as supported by FTIR studies which predict a deprotonation of the Schiff base and a protonation of Glu113 upon rhodopsin activation (M Mahalingam et al. 2008).

Most structural changes are observed at the ionic lock and at the NPxxY(x)5,6F which undergo new interaction with the conserved motif Y(x)7K(R) at TM5. The major structural change is the large movement of TM6 and TM5 (Fig. 1.8, C and D). The TM6 tilt out and TM5 is elongated and moved inward. This helix movement forms a deep crevice at the cytoplasmic side of the receptor permitting the interaction with binding partners (G protein and arrestin). This helix rearrangement is accompanied by a reorganisation of the ionic lock region. Arg135 can interchange its interaction partners upon activation and thus stabilize either inactive or active state, thereby playing the role of microswitch.

26 1. Introduction

Figure 1.8: Structural changes in functional microdomains upon light activation. A. B. Side view of rhodopsin between TM7 and TM1 of Schiff base connecting the K296 to the retinal (11-cis in red, all-trans in yellow). Schiff base is stabilized by E113 in dark state. E181 interacts with S186. Upon activation a proton transfers occurs between the Schiff base and the amino acid E113. Schiff base interaction shifts from E113 to E181. It builds a C. D. Top view of the intracellular side of rhodopsin. The ionic lock in dark state is composed of R135 and E134 of TM3 build a hydrogen bond network with E247 and Thr251 of TM6. Light activation induces a break of the ionic lock and a tilt of TM6 and elongation of TM5. Arg135 builds a new interaction with Y223 and E247 builds a new network with K231 and Thr251. E. F. Side view of TM7 and H8. Y306 and Phe313 of NPxxYx5,6F motif have an aromatic stacking interaction and keep a constraint between TM7 and H8 in dark state. Illumination induces rotation of Y306 which builds a new large water-mediated hydrogen bonding network including notably M257 and A260 at TM6 and N302 at TM7. Taken from (Ernst et al. 2014).

27 1. Introduction

The salt bridge, between Arg135, Glu134 and Glu247, accompanied by a hydrogen bond with Thr251, is broken. Arg135 builds a new interaction with Y223 of TM5 and Y306 of TM7. The amino acids Glu247 and Thr251, once the interaction with Arg135 is disrupted, form an interaction with Lys231 on TM5. The residues Tyr223 and Lys231 belong to the conserved motif Y(x)7K(R) in TM5 and stabilize the active conformation by tethering TM5-TM3 and TM5-TM6 through their new interactions.

Active conformation also presents new constraints at the NPxxY(x)5,6F motif (Fig. 1.8, E and F). Tyr306 breaks its interaction with Phe313, rotates and builds a new interaction with residues at TM6 in a space below Arg135. It builds a large water-mediated hydrogen bonding network including Ser298, Asp83, Asn302, Met257, Tyr223 and Arg135 which stabilizes the active conformation. Like Arg135, Tyr306 changes its interactions and can stabilize the active and the inactive conformation and thus acts as a rotamer microswitch (Hofmann et al. 2009).

1.5.4 Retinal Channel

All active structures show the presence of two openings at the retinal binding pocket forming a tunnel through the receptor. The rearrangement of transmembrane helices allows the formation of a thin hydrophobic channel connecting the polar retinal binding pocket to the outside of the receptor (Hildebrand et al. 2009; Piechnick et al. 2012). The channel traverses the receptor coplanar with the retinal binding pocket (Fig. 1.9). The two openings A and B are nonpolar and formed with aromatic residues. The channel is thought to be used by the all-trans-retinal to leave the receptor after decay and hydrolysis of Schiff base and by 11-cis-retinal to regenerate the receptor. It was reported that, in crystal, rhodopsin is also able with a low affinity to uptake all-trans-retinal, to reform a Schiff base and a Meta II-like conformation (Choe et al. 2011). The presence of arrestin facilitates the all-trans-retinal uptake of phosphorylated opsin (Hofmann et al. 1992)(Sommer, Hofmann, and Heck 2012). Curiously, only half of phosphorylated opsin population was able to take up all-trans-retinal in presence of arrestin (see Sommer et al. 2014). The asymmetry of ligand binding was explained by the binding of one arrestin to a receptor dimer. This uptake of all-trans-retinal by the presence of arrestin was physiologically speculated

28 1. Introduction

as a protection mechanism of the rod cell in bright light. After bright illumination, the release of much all-trans-retinal is expected. All-trans-retinal is toxic for the cell (Rotanowska and Sarna 2005) and the high concentration of all-trans-retinal in the cell could cause serious damages. The uptake of all-trans-retinal in presence of arrestin would help to decrease the concentration of all-trans-retinal below the toxic dose.

Figure 1.9: Retinal channel. Coplanar cuts (cytoplasmic view) of Meta II-like conformation (Choe et al. 2011). Opening A is located between TM1 and TM7. Opening B is located between TM5 and TM6. The two openings are hydrophobic while the binding pocket (BP) is polar. Electrostatic surface potentials are contoured with negatively and positively charges surfaces areas in red and blue respectively. Taken from (Piechnick et al. 2012).

1.5.5 Rhodopsin organization in disc membrane

Long-established studies represent rhodopsin as a monomeric molecule that is randomly dispersed in disc membrane with a density of ~30,000 molecules rhodopsin per μm2 (Gupta and Williamst 1990; Kaplan 1984; B. P. A. Liebman, Weiner, and Drzymala 1982; Poo and Cone 1972, 1974; Roof and Heuser 1982; Wey, Cone, and Edidin 1981). They all measured a diffusion coefficient of 0,3- 0,5 µm2/s for amphibian rhodopsin in disc membrane. A diffusion of 0,33±0,12 µm2/s for rhodopsin in bovine disc membrane was reported (Takezoe and Yu 1981). The high mobility of rhodopsin describes a receptor with rotational and lateral brownian motion free to diffuse in a lipidic membrane. Later, in 2003, atomic force microscopic (AFM) studies on mouse disk membranes revealed a high density with 30,000 to 55,000 molecules rhodopsin per μm2 in AFM

29 1. Introduction

samples (Fotiadis et al. 2003; Liang et al. 2004). Rhodopsin is partially present as a dimer organized in rows of densely packed oligomers about 25 nm long embedded in a ROS disk membrane composed of fluid lipids in a bilayer (Fig. 1.10, B). The formation of dimeres/oligomers of rhodopsin or of other GPCRs were also reported (Comar et al. 2014; Dell’Orco 2013; Kasai and Kusumi 2014; Shukolyukov 2009; Suda et al. 2004) although functional coupling of G protein to the receptor does not require a dimerization of the receptors (Chabre, Deterre, and Antonny 2009; Chabre and Maire 2005; O P Ernst et al. 2007). The homogeneity and the organisation of the whole disk remain unclear. Furthermore organization of dimers or oligomers of rhodopsin seems incompatible with free rhodopsin diffusion. This dilemma forces researchers to repeat these diffusion and AFM measurements. Taking in account the latest knowledge on rhodopsin (Govardovskii et al. 2009) confirmed the diffusion coefficient of 0,4 µm2/s for amphibian rhodopsin, but also detected a variable portion of immobile rhodopsin molecules.

Figure 1.10: Organization of rhodopsin in disc membrane. A. Electron microscopy image of negative stained bovine disks with uranium acetate (top view). Scale bar: 1μm. B. Atomic force topograph of mouse disk membrane. Rhodopsin dimers are organized in racks. Scale bars: 50 nm Insert: Magnification of the topograph, Scale bars: 1,6 nm. Figure B taken from (Fotiadis et al., 2003).

The organization of rhodopsin in native membrane stays in debate and a subject of some controversy. AFM study effectuated in bovine discs membrane reported a confinement of loosely packed rhodopsins in the central area of the disk, surrounded by a girdle of lipids (Buzhynskyy, Salesse, and Scheuring 2011). The central region of the disk presented a density of ~26,000 rhodopsin molecules per μm2. The same

30 1. Introduction

study reported that old samples stored at 3°C after two months presented similar dimer rows as reported before, and thus suggested that the presence of dimers is an artefact resulting from the conditions used for sample preparation. A phase transition of the lipids could results to a segregation of the proteins (Chabre, Cone, and Saibil 2003).

1.6 Arrestin

Arrestin belongs to a small protein family comprising only four members in mammals: Arrestin 1, called rod visual arrestin (or previously S-arrestin), arrestin 2 (β-arrestin 1), arrestin -3 (β-arrestin 2) and arrestin 4 (cone arrestin). These four proteins have analogous sequences and the crystal structures of the four types present structural homologies (Hirsch et al. 1999)(Han et al. 2001)(Zhan et al. 2011)(Sutton et al. 2005). They share the same binding mechanism, however they do not bind the same receptors (M J Lohse et al. 1992). Despite their homologies arrestins show specificity towards their respective biological systems (Lohse and Hoffmann 2014). Visual arrestin and cone arrestin are restricted to photoreceptors. Arrestin 4 is only expressed in cone and is highly specific to cone rhodopsin while arrestin 1 is expressed in both photoreceptors and can also bind both receptors, rhodopsin and cone rhodopsins (Nikonov et al. 2008). Nonvisual arrestins (β-arrestin 1 and β-arrestin 2) are universally expressed and interact with many different GPCRs. After light absorption and activation of G protein, the receptor has to be inactivated. Arrestin, a soluble protein of ~48kDa, plays this role. After phosphorylation of the serines and threonines residues at the C-terminus of the receptor by the rhodopsin kinase (also called G protein-coupled receptor kinase 1, GRK1), arrestin subsequently binds the active phosphorylated rhodopsin (Kühn, Hall, and Wilden 1984) and shields its cytoplasmic surface, thereby preventing G protein activation. Arrestin thus regulates signal transduction. The rhodopsin kinase phosphorylates only activated receptors and arrestin binds only a previously phosphorylated receptor. GRK1 uses ATP to phosphorylate the serine and threonine residues of the C-terminal tail of rhodopsin (Fig. 1.11). A maximum of seven residues can be

31 1. Introduction

phosphorylated: three serines (Ser334, Ser338 and Ser343) and four threonines (Thr335, Thr336, Thr340 and Thr342)(McDowell, Nawrocki, and Hargrave 1993).

324 330 340 348 G K N P L G D D E A S T T V S K T E T S Q V A P A

Figure 1.11: Sequence of bovine rhodopsin C-terminus tail. Kinase (GRK1) phosphorylises the serines at position 334, 448 and 343, and the threonines at position 335, 336, 340 of the C-terminus of the receptor. A maximum of seven residues can be phosphorylated by the kinase for bovine rhodopsin. Uniprot: P02699.

In other cells, arrestin can promote other different cellular signals. After its interaction with a GPCR, receptor-bound arrestin induces receptor internalization through interaction with the protein clathrin (Moaven et al. 2013) and thus triggers a secondary signal transduction. Arrestin also interacts with plenty of non-receptor- binding partners (Xiao et al. 2007). It is a multifunctional molecule involved in multiple pathways. In addition to the full length arrestins, short versions of arrestin were discovered for the four arrestin types. They have all a truncated C-terminus tail compared to their respective full length arrestin. These splice variants are the result of the replacement of the final exon which codes the last amino acids with an exon that code for a single alanine residue. The splice variation occurs at different sites but is still confined to the C-terminal (W Clay Smith 2013). In rods, the splice variant of visual arrestin P44 represents ~5-10% of the amount of visual arrestin. Its sequence is identical to the full length but the last 35 amino acids are replaced with an alanine (Krzysztof Palczewski et al. 1994). Crystal structures of this splice variant show a similar structure to the visual arrestin (Joachim Granzin et al. 2012). P44 has the ability to bind both phosphorylated and unphosphorylated activated rhodopsin with high affinity, while arrestin binds only the phosphorylated receptor.

1.6.1 Basal arrestin structure

Visual arrestin was first crystallised in its basal (or inactive) conformation (Granzin et al. 1998)(Hirsch et al. 1999). The structure presents arrestin as an elongated molecule (~90Å long) composed of two domains: C-domain and N-domain, forming two concave cavities (Fig. 1.12, grey and black respectively). Both domains share a similar structural organisation. Each domain is organized in a superposition of seven

32 1. Introduction

beta sheet strands. Four beta sheet strands are packed against three other beta sheet strands. About half of the 404 residues are organized in beta sheets. The solved crystal structures of the other arrestin types as well as the splice variant p44 present the same overall structural organisation. Globally, the two domains have only few contacts between them. One of the major interactions between the domains is a hydrogen-bond network called the ´´polar core´´ (yellow). It consists of interactions of the embedded residues Asp30, Arg175 of the N-domain, Asp296, Asp303 of the lariat loop of the C-domain and Arg382 of the C-terminus tail. The residues involved in the polar core are highly conserved in the arrestin family. Their interaction keeps the two domains bound and oriented, thereby stabilizing the inactive conformation. The residues involved in the polar core are highly conserved in the arrestin family

Figure 1.12: Structure of basal arrestin. Arrestin is composed of two similar domains: N-domain (black) and C-domain (grey). The polar core (yellow) constituted of core Asp30, Arg175, Asp296, Asp303 and Arg382. The interaction of these amino acids constrains the two domains and keeps the arrestin in the inactive conformation. C-terminus tail (red) interacts with the N-domain and especially with the helix-1 and the beta sheet-1 (wheat) of N-domain which together form the ´´three elements interaction´´. Loops in the crest region; finger loop (green), middle loop (pink) blue and lariat loop (blue, including gate loop, cyan) are supposed to interact with the rhodopsin for the high affinity complex. In basal state, the finger loop is stabilized in a close conformation. PDB: 1CF1, molecule D (Hirsch et al. 1999).

The C-terminal tail (residues 372-404, red), missing in P44, is connected to the C-domain and forms a beta sheet interaction with lateral beta sheet of N-domain. Interactions between beta-strand I (Val3-Ile16, wheat) and alpha-helix I (Arg102- Leu111, wheat) of N-domain and with the C-terminus tail (called three-element 33 1. Introduction

interaction) also keep the arrestin inactive. The three-element interaction and the polar core are the two main constraints which keep arrestin inactive. Located in the central crest region, are three loops which are supposed to be involved in arrestin activation and in binding with the receptor ((M. Kim et al. 2012)(M E Sommer et al. 2007) (Hanson et al. 2006)): the finger loop (Gly68-Ser78, green), the middle loop or also called loop 139 (Glu133-Ser142, red) and the lariat loop (Leu283-Asn305, blue/cyan). Note that the lariat loop also includes the gate loop (Asp296-Asn305, cyan) Many studies suggest that the finger loop builds the main interaction with receptor and binds in its crevice (Feuerstein et al. 2009)(Dinculescu et al. 2002)(Hanson et al. 2006)(Shukla et al. 2014). In the inactive conformation, the finger loop is folded down.

1.6.2 Arrestin binding

Arrestin binds activated phosphorylated rhodopsin (P-R*) with a high affinity

(Kd= ~50nM) (Schleicher et al. 1989; Zhuang et al. 2013). Arrestin also specifically interacts with inactive phosphorylated rhodopsin (P-rho) (Kd = ~80μM) (Zhuang et al. 2013) and phosphorylated opsin (P-ops) Kd= 1,5μM (Sommer, Hofmann, and Heck 2012). No binding is detected by dark unphosphorylated rhodopsin. Arrestin binding affinity increases with each phosphorylation. Arrestin requires at least three phosphates on the rhodopsin tail to detect it and to bind it. Bovine rhodopsin can have a maximum of seven phosphates. The addition of more than three phosphates does not increase the stability of the high affinity complex, but increases the affinity of arrestin binding to dark phosphorylated rhodopsin and phosphorylated opsin (Vishnivetskiy et al. 2007). Arrestin recognises and specifically binds light-activated and/or phosphorylated rhodopsin species. Arrestin requires both conditions, the active form of receptor and phosphorylation of the C-terminus tail, to undergo high affinity binding to the receptor. Arrestin binding models suggest that arrestin contains a ´´phosphorylation sensor´´ and an ´´activation sensor´´ which independently detect the activated rhodopsin form and the phosphates at the receptor tail (Gurevich and Benovic 1993). It must be able to discriminate between the active and the inactive form, and between the phosphorylated and unphosphorylated form of the receptor. Binding of arrestin to P-R* is achieved by a multistep binding mechanism (scheme 1.1).

34 1. Introduction

hv * kinase * Arrinactive * * R R Rp Rp  Arrpreactive Rp  Arractive  ATP precomplex highaffinity complex

Scheme 1.1: Phosophorylation and arrestin binding. Arrestin binding to rhodopsin occurs in two steps. After light absorption, C-terminus tail of activated rhodopsin is phosphorylated by rhodopsin kinase. Arrestin (arrinactive) recognises the phosphorylated activated receptor (RP*) and interact with its phosphorylated tail (pre-complex), inducing first a arrestin conformational change into pre-active state (Arrpre-active). A subsequent arrestin rearrangement leads to the formation of the high affinity complex.

Arrestin first recognises the receptor phosphorylated tail and forms the pre-complex in which the arrestin is only bound to the tail of the receptor. Due to multiple exposed positive amino acids (Ostermaier et al. 2014), arrestin builds salt dependent electrostatic interactions with the phosphorylated tail. The C-terminus tail plays a crucial role for discrimination between Rho* to P-Rho*. The phosphorylated receptor tail displaces the C-terminus tail of arrestin. This tail exchange induces conformational changes in arrestin, leading to the pre-active form, where additional arrestin elements make contacts with rhodopsin. Deletion of arrestin C-terminus tail leads to the binding of arrestin to unphosphorylated active receptor such as the case of p44 (Palczewski, Buczylko, et al. 1991). Rearrangement of arrestin into a pre-active state allows arrestin to be prepared for the final conformational changes needed to bind tightly the active receptor (Gurevich et al. 2011). . The conformational rearrangement involves additional arrestin elements, in particular the loops at the centre crest, coming into contact with rhodopsin, and leads to the high affinity binding of arrestin (Gurevich and Gurevich 2004). The extended finger loop conformation allows interactions with the receptor and the adjustment into a fitted form, permitting the introduction of the loop into the receptor crevice to form the high affinity complex and the stabilization of the Meta II conformation (A Schleicher, Kühn, and Hofmann 1989)

1.6.3 Pre-active structure of arrestin

Recently the crystal structure of arrestin-2 co-crystallized with GPCR phosphorylated tail analogue peptide (Shukla et al. 2013) and a new structure of splice variant P44 were reported (Y. J. Kim et al. 2013). Both structures are highly

35 1. Introduction

similar and present the structure of arrestin in a pre-active state. Comparison of the pre-active state with the basal state presents significant structural rearrangements and gives a general idea of the mechanism of arrestin activation (Fig. 1.13). The conformational changes show an overall screw-like twist of 20° between the two domains, a reorganization of the polar core and displacement of the finger, middle and gate loops.

Receptor phosphorylated tail analogue peptide binds the arrestin at a similar location to the arrestin C-terminus tail. The phosphorylated C-terminus tail of the receptor replaces the C-terminus tail of the arrestin (tail exchange) and builds antiparallel beta sheet interactions with the arrestin at the N-domain. The three element interaction is absent and the release of the arrestin C-terminus tail upon arrestin binding was already shown by mutant EPR (Hanson et al. 2006) and NMR (Zhuang et al. 2013) studies. Receptor tail interacts with Arg29, a residue close to the polar core, Arg66 of the finger loop and with Lys300 of the gate loop. A point mutation mutagenesis study (Ostermaier et al. 2014) suggests that Arg29 controls the tail exchange mechanism. It also forms new charge-charge interactions with arginine and lysine side chains (Lys14, Lys15, Lys20, Lys110 and Lys300) without modifying the beta sheet structures of N-domain. The lysines and arginines of the N-domain were already predicted to be the phosphate sensor of arrestin (Vishnivetskiy et al. 2000). Displacement or absence of the C-terminus tail triggers the breaking of the polar core and the activation of the arrestin. The inactive state is principally stabilized by the polar core. The replacement or the absence of the C-terminus tail of arrestin removes the interaction of Arg382 from the polar core and thus disrupts it. The interaction between Arg175 and residues Asp296 and Asp303 of the gate loop are also lost. Binding of phosphorylated receptor tail to arrestin indirectly disturbs the polar core. The receptor tail does not make any direct contact with the polar core. However replacement of the C-terminus tail of arrestin by the receptor tail, or the absence of C-terminus tail such as in the case of P44, retrieves Asn305 from the polar core and breaks the polar core. The disruption of the polar core is required for arrestin activation. The resulting rearrangement of the polar core and of the arrestin loops allows arrestin to bind tightly the phosphorylated activated receptors and thus form the high affinity complex. Mutagenesis of polar core key residues (Arg175 and

36 1. Introduction

Asp296) and thus disruption of the polar core gives the possibility to arrestin to bind unphosphorylated activated receptor as p44 does (Kovoor et al. 1999). Arg29 interacts with the phosphorylated tail and was suggested to have a significant influence on the polar core and to control a switch between the arrestin tail and the receptor tail (Ostermaier et al. 2014). Reorganisation of the polar core is accompanied by conformational changes of the finger, middle and gate loops. In the inactive state, these loops present significant flexibility, however they show a clear conformational change in the pre-active conformation. C-tail displacement triggers conformational changes and formation of new interactions at the polar core region in the pre-active forms. This rearrangement exposes the gate loop so that both finger and gate loop are available as the major interaction site with the activated receptor. Asp303 and Asp296 residues are retrieved from the polar core. Gly297 interacts with the residual amino acids of the polar core (Asp30, Arg175). Asp303 moves and interacts with Pro288 of gate loop. An additional interaction of Lys300 of the gate loop is observed with the phosphorylated peptide. The gate loop moves, twists and stabilizes the pre-active conformation. Compared to the basal state, the finger loop is in an extended conformation which enables better contact with the receptor and engages its binding crevice. The phosphorylated receptor tail peptide interacts with the base of the finger loop (Arg66) and probably contributes to the increase of the stabilization of the finger loop in its extended form. The middle loop also shifts upon arrestin activation. Its position shifts in a significant manner in the pre-active state. It moves away from the binding interface leaving place for rhodopsin interaction. Rearrangement of arrestin into a pre-active state permits arrestin to be prepared for the final conformational changes needed to bind the active receptor tightly. The extended finger loop conformation allows interactions with the receptor and the adjustment into a fitted form permitting the introduction of the loop into the receptor crevice.

37 1. Introduction

Figure 1.13: View of N-domain of basal and pre-activated states. A. Inactive structure of arrestin-1(PDB: 1GMA, molecule A). In red, arrestin C-terminus interacts with the N-domain and is involved in the three elements interaction. The finger loop is in a close conformation (green). The polar core (yellow) keeps the arrestin in the basal conformation. B. Pre-active structure of arrestin-1 with phosphorylated C-terminus receptor analogue peptide (PDB :4JQI.). Phsophorylated C-terminus analogue peptide (turquoise, phosphates marqued) builds antiparallel beta sheet with arrestin N-domain. Polar core is broken (yellow) and the three elements interaction is absent. Finger loop (green) is elongated and middle loop (pink) is retracted to the arrestin. Gate loop (blue) is twisted and builds new interaction with polar core residues. C. In basal state, the polar core is constituted of a salt bridge between Arg175 and Glu297 and a hydrogen-bond network with Asp30, Asp303 and Arg383. The polar core constrains N-domains, the basal loop and the C-terminus tail together. D. In pre-active state (here of p44), the polar core is broken. C-terminus tail is retrieved from the network. Asp303 and Asp296 from the gate loop get also back from the interaction. Asp303 builds a new interaction with Arg288 of the gate loop and Asp296 forms a new interaction with the residual amino acids of the polar core and stabilizes the pre-active conformation. Thereby, the lariat loop moves and twists. Figures C and D taken from (Y. J. Kim et al. 2013).

38 1. Introduction

1.7 Aim of the project

Different studies on arrestin rhodopsin interactions (M. Kim et al. 2012; Ostermaier et al. 2014; Shukla et al. 2014; Sinha et al. 2014; W C Smith et al. 2004; Martha E Sommer, Hofmann, and Heck 2012) have shown that the finger loop builds the major interactions with the receptor. Moreover, the arrestin family contains only four members which are responsible for desensitizing the large GPCR family. The following questions arise: How does the finger loop interact with the receptor crevice? Do the arrestins share a common finger loop motif to bind their receptors? Do the four arrestins have the same binding interactions as the Gt protein? Aim of this study was, alongside crystallographic studies, to provide functional measurements to quantify the capacity of peptides derived from the different finger loops of the four arrestins to stabilize Meta II. Since no crystal structure of the high affinity arrestin/receptor complex is yet available, the transition to the high affinity complex remains obscure. Two contradictory models on arrestin binding have been reported. EPR experiment indicate that the overall shape of arrestin remains essentially the same, and changes are only localized to flexible loops in the central crest region (M. Kim et al. 2012). In contrast, NMR experiments suggest that arrestin undergoes a global unstructuring and transition to a molten globule (Zhuang et al. 2013). Aim of this study was thus to explore the formation and decay of arrestin-rhodopsin complex by FTIR. This technique provides information of structural changes occurring upon formation and decay of the complex. Dimerisation of rhodopsin (as also shown for other GPCRs)(Dimitrios Fotiadis et al. 2003)(Mansoor, Palczewski, and Farrens 2006)(Jastrzebska et al. 2006) is suspected to influence in the binding and decay mechanism of the arrestin complex (Sommer, Hofmann, and Heck 2012)(Sommer, Hofmann, and Heck 2011). The objective of the last part of this study was thus to investigate the diffusion of rhodopsin in native disc membranes and to explore the mutual influence of the rhodopsin dimerisation on the arrestin-rhodopsin interaction with single fluorescence microscopy.

39 1. Introduction

40 2. Material and Methods

2. Material and Methods

2.1 Isolation of Rod Out of Segment

Isolation of Bovine Rod outer Segments (ROS) was preceded as described by Sommer (Sommer et al, nature communications, 2012). ROS were mechanically extracted from the retina (W.L. Lawson Company, USA) and separated from the other cells using a suitable sucrose 3-step-gradient (ρ=1.105; g/mL ρ=1.115 g/mL and ρ=1.135 g/mL). The separation results from the difference in density between ROS and the other cells of the retina. All sucrose solutions were prepared in ROS buffer (70 mM Potassium phosphate, 1 mM MgCl2, 0,1 mM EDTA, 1 mM DTT, 1 mM DTT, 0,5 mM PMSF, pH: 7,0) and the density of each sucrose solution was tested and corrected using hydrometers. Under dim red light, 100 frozen retinae were thawed. 90 mL of 45% (wh/vol) sucrose solution were added to the retinae. The solution was vigorously shaken during 4 min to separate the ROS from the rod cells. The suspension was centrifuged (5,000 rpm, 5 min) and the supernatant was filtered through a cheesecloth and diluted with an equivalent volume of ROS buffer. The suspension was then centrifuged at 7,000 rpm during 7 min. The supernatant was carefully poured off and resuspended in a 25,5% (w/v) sucrose (ρ=1.105 g/mL) with a end volume of 40 mL and carefully layered onto fours gradients. The gradients where prepared in 4 polyallomer thin-wall tubes (Beckman) composed of 14 mL of 27.125% (w/v) sucrose (ρ=1.115 g/mL) underlayed with 14 mL 32.25% (w/v) sucrose (ρ=1.135 g/mL). The layered gradients were centrifuged at 25,000 rpm during 30 min in a swinging bucket (Beckman SW 28 rotor) with mild acceleration and deceleration mode. After centrifugation, the ROS was localized at the 27,125% and 32.25% interface in a thin band. Each tube was pierced with a syringe and only the ROS band and the solution directly on the upper face of the band was extracted from the tube. Collected ROS was diluted 1:1 with ROS buffer and centrifuged at 25,000 rpm during 30 min. The ROS pellet was resuspended in 10 mL ROS buffer, snap frozen and stored in dark at -80°C

41 2. Material and Methods

2.2 Isolated Membranes

Isolated membranes are discs membranes containing rhodopsins extracted from ROS and washed with hypotonic buffer. The hypotonic condition allows through osmotic pressure to burst the ROS and to liberate the disc membranes and also allows to separate all proteins bound to the discs. In low salt condition all different proteins bound to the discs are dissociated from the membrane (Kühn et al., 1982) without to disturb neither the rhodopsin nor the lipids of the disc membrane. All the preparation occurs under dim red light. ROS was thawed, resuspended in a salt less 100mM HEPES buffer pH: 7.0 (4-(2- hydroxyethyl)-1-piperazineethanesulfonic acid) and dounced to burst the ROS, liberating the disc membranes and to homogenized them. The membrane suspension was centrifuged at 16,000 rpm during 20 min. The resulting pellet was resuspended in salt less HEPES buffer and dounced. Several similar washing steps (centrifuge steps followed by resuspension of the pellet in HEPES buffer) washed all bound proteins (G protein, Arrestin, Kinase etc...) from the disc membranes. After 5 washed steps, the isolated disc membranes were aliquoted and stored in dark at -80°C.

2.3 Phosphorylation of Rhodopsin

Rhodopsin was phosphorylated through light induced phosphorylation of the andogenous rhodopsin kinase present in the ROS as published (Sommer, Hofmann, and Heck 2012). Addition of ATP to the ROS and illumination activate the rhodopsin kinase. ROS was thawed, carefully homogenized with a glass douncer to burst the plasma membrane and phosphorylated in 100 mM potassium phosphate buffer (pH: 7,4) in the presence of 2 mM MgCl2 and 8 mM ATP. The reaction occurred two hours at room temperature under desk lamp. 30 mM Hydroxilamine was then added to the phosphorylated ROS to convert all rhodopsin photoproduct to opsin and free retinal. After 10 minute reaction, phosphorylated opsin was centrifuged at 16,000 rpm during 20 min and resuspended with ROS buffer (70 mM potassium phosphate,

1 mM MgCl2 , 0,1 mM EDTA, pH: 7,0 + 1 mM DTT and 0,5 mM PMSF). The phosphorylated opsin was then washed 3 times with the same procedure with

42 2. Material and Methods

standard HEPES Buffer (100 mM HEPES, pH: 7,0). This hypotonic buffer allows the dissociation of all different proteins (G protein, Arrestin, Kinase etc...) from the discs membrane. The membrane pellet was finally resuspended in HEPES Buffer at pH: 8,0 with a final phosphorylated opsin of 100-200 μM. The membrane suspension was homogenized for a last time with the douncer, aliquoted, snap-frozen in liquid N2 and stored at -80 °C. For measurement, the phosphorylated opsin was regenerated up to one hour at room temperature with a three-fold molar excess of 11-cis-retinal. The efficiency of the phosphorylation was verified by an Extra MII essay. Only rhodopsin preparations with high phosphorylation rate were used for arrestin binding.

2.4 Arrestin and P44 expression

Cysteine less and tryptophan less bovine arrestin mutant (C63A, C128S, C143A and W194F) analogue to arrestin wild type and P44 wild type were expressed in E.Coli BL21(D3) in LB medium. 5 flasks of 800 mL of LB medium with Ampicillin were inoculated with each 80 ml of a pre-culture of transformed E.Coli BL21(DE3) containing the arrestin plasmid (pET15b). After ~3 hours incubation at 30°C, 30 µM of IPTG (isopropyl 1-thio-ß-D-galactopyranoside) were added to initiate the expression of arrestin. The cells were inoculated overnight at 28°C. The next day, cells were harvest by centrifugation (5,000 rpm, 5 min). The pellet was resuspended in 25 mL C-Buffer (10 mM Tris-HCl, 2 mM EDTA, 0,1 M NaCl, pH: 7.0) and the cells were lysed through ultrasonification.

2.5 Arrestin purification

Arrestin was purified first by ammonium sulfate precipitation and then through an affinity Heparin sepharose column followed by an anion exchange Q-sepahrose column as describe in (M E Sommer, Smith, and Farrens 2006). Arrestin has a plenty of specific binding partners. One of them is Heparin so that an affinity column with Heparin can be used to purify Arrestin (K Palczewski, Pulvermüller, et al. 1991). The many negative charges of Arrestin can also be used for the purification through a standard anion exchange column.

43 2. Material and Methods

1 mL of DNAse II and 7 µM MgCl2 was added to the resuspended pellet and the solution was sonificated (2 min, max power). The lysate was cleared by centrifugation at 18,000 rpm during 30 min and 0,32 mg/mL of ammonium sulfate was carefully added to the supernatant to precipitate the proteins. The precipitate was centrifuged (15,000 rpm, 30 min). The precipitate pellet was resuspended in hight salt C-Buffer (10 mM Tris-HCl, 2 mM EDTA, 1 M NaCl, pH: 7.0) and centrifuged during 30 min at 18,000 rpm. The supernatant was filtered and loaded onto a 3 x 5 mL HiTrap Heparin Sepharose column (Pharmacia) equilibrated with salt less C-Buffer (10 mM Tris-HCl, 2 mM EDTA, pH: 8.5). Arrestin was washed and eluted with a linear gradient (from 0.1 to 0.5 M NaCl over 300 mL). Fraction containing arrestin were pooled and filtered and loaded onto a 5 mL Q-Sepharose anion exchange column (Pharmacia) equilibrated with salt less Q-Buffer (10 mM Tris-HCl, 2 mM EDTA, pH: 8.5). Arrestin was washed and eluted through a linear salt gradient (0 M NaCl to 0.3 M NaCl and then 1 M NaCl). Fractions containing arrestin were pooled, concentrated and the buffer was then exchanged with standard arrestin buffer (50 mM HEPES, 130 mM NaCl, pH: 7.0) with a 10 kDa centricon. Arrestin concentration was determined by measuring the absorbance at 280 nm -1 -1 (εcysless= 20760 M cm ). Arrestin was aliquoted and snap-frozen in liquid nitrogen and stored at -20°C.

2.6 P44 purification

The absence of arrestin tail changes the isoelectric point of arrestin. Purification was performed as previously described for full length arrestin excepted that the ion exchange purification step was replaced by cation exchange purification using cation exchanger SP-Sepharose column (GE healthcare). Briefly, the cell pellet was resuspended with L-Buffer (10 mM HEPES, 100 mM NaCl, pH: 7.5). The cells were lyzed through sonification, centrifuged and filtered as for arrestin. The lysate was then loaded (2mL/min) into the 3x5 mL HiTrap Heparin column (Pharmacia) which was previously equilibrated with L-Buffer. The protein was eluted with a salt gradient (from 0.1 to 1 M NaCl over 400 mL). Fraction containing p44 were pooled and filtered and loaded (2ml/min) onto a 2x5 mL SP-Sepharose anion exchange column (GE healthcare) equilibrated with salt less M-Buffer (10 mM HEPES, pH: 7.5). p44 was eluted with a salt gradient (from 0.1 to 44 2. Material and Methods

1 mM NaCl). Fractions containing p44 were pooled together. The buffer was then exchanged with standard arrestin buffer (50 mM HEPES, 130 mM NaCl, pH: 7.0) and finally concentrated with a 10 kDa centricon (Milipore). p44 concentration was determined by measuring the absorbance at 280 nm -1 -1 (εcysless= 24,870 M cm )(Schubert et al. 1999). p44 was aliquoted and snap-frozen in liquid nitrogen and stored at -20°C.

2.7 Specific labelling of protein

2.7.1 Maleimide reaction

The specific labelling was performed with dye attached to maleimides compounds. Maleimides are electrophilic compounds which react selectively with the thiol group. The labelling of protein can be specific to cysteine residues of the protein. The labelling results to covalent bond between the maleimide and the protein (Figure 2.1).

Figure 2.1: Reaction scheme of maleimide reaction. Maleimide reacts with thiol group of the cysteines of a protein. The double bond of maleimide react with thiol group found on cysteine and forms a covalent bond with the protein.

2.7.2 Rhodopsin labelling

Cys140 and Cys316 are the only two cysteines of rhodopsin present on the cytoplasmic side of the disk suitable for labelling (Fig. 2.1)(Chen and Hubbell 1978). The dyes, Fluorescein maleimide (sigma aldrich) and Atto 647 N maleimide (Atto 647, Atto-tech) were first solubilized in dimethyl sulfoxide (DMSO) and then in 100 mM HEPES, pH: 7 buffer to a end concentration of 100 mM. The reaction occurs during 2h at room temperature or over night at 4°C. The labelled rhodopsin was then washed several times with 100mM HEPES buffer to removed free dyes. A wash step consists of a centrifugation 14,000 rpm during 30 min. The pellet is the resuspended with buffer.

45 2. Material and Methods

Atto 647 has a maximal absorption at 647 nm. It has a high fluorescence quantum yield and a high photo-stability. It was used for single molecule fluorescence microscopy. Single molecule labelling was effectuated at low concentration to label at most one rhodopsin per disk. 60 nM Atto 647 for 100 μM of rhodopsin Figure 2.2. Schema of labelled disk. Many rhodopsin molecules (grey was the best labelling ratio for single molecule spot) are labelled with fluorescein measurement. In contrast, fluorescein was used (green). Atto 647 N maleimide was used for single labelling. Only one as control at high concentration (50 μM) to label rhodopsin per disk was labelled with Atto 647 N maleimide. many rhodopsins per disks in way to detect the disks (Fig. 2.2). Fluorescein absorbs at 494 nm but has a weaker photo-stability. The labelling of Atto 647 and the fluoresceine were proceeded simultaneously (Fig. 2.2). The labelling with fluorescein, and its illumination with a 532 nm diode laser, was only used to control if the single molecule labelling with Atto 647 into a disk which was excited with a 633 nm diode laser. The fluorescence of fluorescein does not over lap the absorption of Atto 647.

2.7.2 Quantum Dot single labelling strategy

Quantum dots are extremely photostable and emit a bright fluorescence. The quantum dot, QD 655 (life technologies) used for single fluorescence microscopy emits at ~655 nm and has a bright absorption spectrum. QD 655 was excited with a 532 nm. QD 655 used for rhodopsin labelling does not contain maleimide. The QD 655 is coated with streptavidin. Streptavidin has a very high binding affinity for biotin. The binding of biotin to streptavidin is one of the strongest non-covalent interactions known in nature and can be used for a stable labelling. The single labelling of rhodopsin occurs in two steps. First, a biotin-PEG-maleimide linker (sigma aldrich) was used to labelled the rhodopsin. Subsequently, the QD 655 was added to the sample to bind the biotin linker. The aim is to have less than one rhodopsin QD 655 labelled per disc membrane. 60 nM of biotin-PEG-maleimide was added to 100uM opsin. The reaction occurs 2 h at room temperature. The labelled was washed 5 times to removed all free labels.

46 2. Material and Methods

100pM of QD 655 was then added to the sample for the final labelling step. The sample was washed 10 times to removed all free labels.

2.7.3 Arrestin labelling

Cysteine less and tryptophan less bovine arrestin mutant (C63A, C128S, C143A and W194F) with A360C single cysteine mutant was labelled with IANBD (N,N'- dimethyl-N-(iodoacetyl)-N'-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)Ethylenediamine, life technology) as describe (M E Sommer, Smith, and Farrens 2006). The dye was solubilised in DMSO. Arrestin samples were buffer-exchanged (100 mM HEPES, 150 mM NaCl, pH 6.5) and concentrated, and IANBD was added in 10-fold molar excess to the single cystein arrestin mutant. Note that the concentration of DMSO must be below 5%. Reaction occurs during 3h at room temperature. The labelled arrestin was then passed over a size exclusion column (500µl, Sephadex G-15) to remove the free labels. The sample were finally buffer-exchanged into 100 mM HEPES, 150 mM NaCl, pH 7.0. NBD has a maximal absorption at 500 nm with an extinction -1 -1 ε500 = 25,000 cm M .

2.8 UV-vis absorption spectroscopy

Spectroscopy is the interaction of molecules with electromagnetic radiation. Molecules with electrons in delocalized aromatic systems (π electron system) absorb light in the ultraviolet-visible region (UV-vis: 150-800 nm). The absorption of the UV-vis light induces electronic transitions. The energy of light is used to excite these electrons and to promote them from the ground state to an excited state. The absorption spectroscopy allows to detect different pigments and conformational changes and to measure concentrations with the Lambert-Beer law (Equ. 1).

 I  A  log     c l (1)  I0 

47 2. Material and Methods

The absorbance A (in OD = optical density) corresponds to the difference of light intensity I and I0 after passing through a solution with and without the analyte. The absorbance is also in a direct correlation to the concentration C, the pathway length l and to the extinction coefficient ε of the analyte.

The different rhodopsin conformations where identified through this absorption technique. The photoproducts have different chromophore isomers and are characterised with specific absorptions for each Meta species. Rhodopsin spectrum is characterised with a 280 nm maximum absorption due to the absorption of the intrinsic aromatic residues (tryptophan and tyrosine) and a maximum absorption at 498 nm corresponding to the protonated Schiff base between the 11-cis-Retinal linked to Lys296. After illumination, 11-cis-Retinal is isomerized to all-trans-Retinal and the Meta I and Meta II conformation are formed. The Meta I has a 478 nm maximum absorption while the deprotonation of the Schiff base leads to the Meta II conformation with a shift of maximum absorption to 380 nm. Opsin like all rhodopsin absorbs only at 280 nm. In absence of retinal and Schiff base, no pigment is detected. Free retinal absorbs around 380 nm (11-cis-Retinal : 380nm, all-trans-Retinal: 383nm). All spectra were recorded with a Varian Cary 50 UV/Vis-Spectrometer with 2 nm resolution. The usual sample volume was 100 μL and recorded in a 1 cm pathlength cuvette at room temperature. The determinations of the concentration of Rhodopsin were proceed in a 20 times dilution and recorded after and before the illumination during 10s with a long-pass filter (λ > 480 nm, Schott GG495). The concentration was calculated with 500 nm minimum of the difference spectrum (light-dark) and -1 -1 the extinction coefficient of rhodopsin (ε500nm: 40,800 M cm ).

48 2. Material and Methods

2.9 Extra MII assay

After illumination, Meta I and Meta II are in equilibrium. This equilibrium shifts through pH and temperature (Parkes and Liebman, 1984). The binding of peptide or arrestin within receptor cavity forces and stabilizes the Meta II conformation. In condition favouring Meta I (0°C, pH: 8.0) the increase of formation of Meta II engender by a biding is easily detectable with the specific absorption of Meta II. All measurements were recorded with a Shimadzu UV3000 two-wavelength spectrometer. 380 nm and 417 nm wavelengths were simultaneously recorded reducing/minimizing scattering artefacts. Samples consisted of 200 μL of 5-10 μM Rhodopsin in 100 mM HEPES buffer pH 8.0 were measured in 2 mm path length cuvette at 0°C. The peptides (Genscript, Selleckchem, >95% purity) were solved in water to a final concentration of 12 mM and the pH adjusts to pH: 8.0. The reaction was induced with a photoflash with a green filter (500nm) activating 18-20% of the rhodopsin. The difference of absorption of Meta II at 380 nm and the 417 nm were measured simultaneously upon time. The isobestic point between Meta I and Meta II at 417 nm was used to subtract light-scattering artefacts from the MII signal at 380 nm (Schleicher et al., 1989). The final amplitudes of each peptide concentration were then plotted in function of peptide concentration. The signal was compared to minimum Meta II formation (rhodopsin alone) and the maximum Meta II formation in presence of high affinity Gt alpha C-terminus peptide (Gt HAA).

The titration data were fit to a hyperbolic expression (Bmax*x/Kd+x), and the resulting apparent Kd was corrected for Meta I / Meta II equilibrium at pH 8.0, 0°C (12,5% of Meta II).

2.10 Fourier Transform Infrared Spectroscopy

Infrared light has longer wavelengths and lower frequencies than those of visible light. In comparison with UV-Vis spectroscopy, infrared radiation does not have enough energy to induce electronic transitions. The absorption of infrared radiation excites vibrational transitions of molecules. The light is absorbed when the frequency of light matches with the vibration of the molecule and the molecular dipole moment change during the vibration.

49 2. Material and Methods

Instead of the wavelength, the wavenumber ν, the inverse of wavelength, is used as unit. The mid-infrared range, between (2.5 to 50 μm = 4000 to 400 cm-1), is used to study fundamental vibrations and associated rotational-vibrational structure. There are different kinds of molecular vibrational modes: symmetric and antisymmetric stretching which change the bond length and scissoring, rocking, wagging and twisting.

Nowadays infrared spectrometers are equipped with interferometer which permits the measurement of all frequencies simultaneously. The spectra can be calculated through a Fourier transformation, from the interferogram obtained from the detector. This technique, called Fourier-transform Infrared (FTIR) is able to record time resolved spectra at high resolution. Probes for FTIR measurement were prepared as following. Depending on the measurement, the membranes containing rhodopsin or phosphorylated rhodopsin were washed with 100 mM HEPES buffer at an appropriate pH. Similarly, arrestin or p44, were buffer exchanged with the appropriate buffer. At the same time, a 10 kDa centricon (Milipore) was used to bring the concentration to to 1 mM. The membranes containing rhodopsin or phosphorylated rhodopsin were centrifuged at 14,000 rpm for 30 min. The supernatant was removed and the pellet was resuspended with arrestin, p44 or peptide. The resulting suspension was centrifuged at 14,000 rpm for 30 min. The supernatant was removed and the pellet was transferred to a BaF2 cuvette for FTIR measurements. FTIR spectroscopy was performed with a Bruker ifs66/v vacuum spectrometer equipped with a HgCdTe Detector (PV-MCT), as described in (E Ritter et al. 2007). Highly concentrated samples of 300 to 500 μM were loaded in a BaF2 cuvette ´´sandwich´´ with a 4 μm spacer. The resulting optical pathlength was 4 μm. All spectra were measured in rapid-scan mode with a time-resolution of 200 ms.

Illumination of rhodopsin was performed with 4 Orange LEDs (λmax = 590 nm) for 5 s, which achieved a homogeneous irradiation that activated ~100% of the receptors. Before measurement, the samples were equilibrated for one hour in the spectrometer. A set of time-dependent IR-transmission spectra was recorded 2 minutes before, during illumination and thereafter.

50 2. Material and Methods

2.10.1 Difference spectrum

Due to the many overlapping bands in the spectra, the chemical structure of a protein cannot be deduced. Only changes in the chemical structure can be detected and monitored by infrared spectroscopy. To observe the light induced conformational changes, the initial state spectrum before illumination was averaged and subtracted from whole data set. This allowed all changes occurring during the transition to be monitored. In order to obtain a more precise representation of the transition, a difference spectrum was calculated. The spectra of the initial state (before illumination) were subtracted from the spectra of the final state (after illumination). The resulting spectrum is called the difference spectrum Δ:

Δ = SLight - SDark (2) Or Δ = {MetaI/Meta II - Rho} (3)

The resulting calculated spectrum shows the changes induced by the light, in which negative bands indicate vibrational modes of the dark states while positive bands represent vibrations of the respective illuminated states. Typical difference spectra of rhodopsin representing the transition of dark state to Meta I and to Meta II are shown in figure 2.3.

Figure 2.3: Example of standard difference spectrum. Difference spectra of rhodopsin (light minus dark) representing the transition of dark state to Meta IIbH+ (red, 20°C, pH: 5.0)) and to Meta I (blue, 10°C, pH: 9.5)). Taken from (Mohana Mahalingam et al. 2008).

51 2. Material and Methods

The data sets were corrected for slight temperature drifts. A base line correction was calculated based on the spectra recorded before illumination. The data sets were then normalized based on the chromophore band present at 1238 cm-1. The fingerprint pattern around this band was integrated between 1220 and 1620 cm-1 to avoid small deviations of the 1238 cm-1 band. The difference spectra give us global information about the conformation and photointermediates of the protein, and provide detailed information about the chromophore and all other functional groups. The amide I region (around 1650 cm-1) and amide II (region 1560 cm-1) indicate small distortions of the protein backbone, while the region at 1230 cm-1 provides information on the chromophore and the Schiff base. Without the assignation of all bands, the difference spectra cannot be interpreted at an atomic level. Nevertheless, motifs and specific bands of these amide regions can be used to describe the different conformational transitions. Finally, spectra describing pure transition components and the corresponding kinetic constants of these components were derived by a combination of singular value decomposition (SVD) and global fitting, as described (M Elgeti, Ritter, and Bartl 2008). Small contribution of Meta I were estimated by global fitting and subsequently subtracted.

2.10.2 Double difference spectrum

To highlight differences between the samples containing only rhodopsin, or containing rhodopsin with arrestin, p44 or Gt alpha HAA analogue peptide, double differences ΔD were calculated (equ. 4). A double difference spectrum is the difference of two difference spectra (ΔD= Δa - Δb). Difference spectra without arrestin were subtracted from the difference spectrum obtained in the presence of arrestin (or p44, or Gt alpha HAA).

ΔD = Δwith arrestin – Δwithout arrestin (4) or D  MetaII  Arr* Rho  ArrMetaII  Rho (5)

The contributions of Meta I and residual Meta II, depending on the pH and on the residual unbound rhodopsins were estimated by global fitting and by the intensity of

52 2. Material and Methods

the 1768 cm-1 Meta II marker band and subsequently subtracted from the difference spectra. The positive bands in the double difference spectra represent the vibrations of the high-affinity complex, while negative bands result form vibrations of the arrestin pre-complex. With the expression:   Rho  ArrRho  Arr, which corresponds to the formation of the pre-complex, the double difference spectrum can be written as:

D  MetaII  Arr* MetaII  Rho (6)

The double difference spectrum represents all changes that occur during transition from Meta II and free unbound arrestin to their complexed form, except for all changes that already have occurred in the formation of the pre-complex (δ). Considering that all rhodopsin molecules are already pre-complexed with arrestin in the dark, the contribution of δ is equal to zero and thus the double difference spectrum represents only the transition of the pre-complex to the high affinity complex.

2.11 Pulled Down assay

Pulled down assay was proceeded as described in (Sommer, Hofmann, and Heck 2011). Briefly, 20 μM NBD labeled arrestin (A366C-NBD, 95% labelling) was mixed with 8 μM phosphorylated rhodopsin (100 μl volume). Samples were fully photoactivated (with a long-pass filter λ > 480 nm, Schott GG495) and then immediately centrifuged 14,000 rpm, 30min. Following centrifugation, 70 μl of the supernatant was carefully removed from each sample, and the absorbance was measured using a Varian Cary 50 spectrometer (small window cuvette, 1 cm path length). The amount of arrestin “pulled down” with the membranes was simply the difference between the total concentration of arrestin in the sample and the concentration of arrestin remaining in the supernatant, -1 -1 which was determined by the NBD absorbance at 500 nm (ε500=25,000 cm M ).

53 2. Material and Methods

2.13 Electron Microscopy

3,5 μL droplet of 2 μM of rhodopsin in disc membrane were disposed on a copper grid coated with continuous carbon layer (Quantifoll). After one minute, the droplet was absorbed with a filter paper and the disks laying on the grid were then stained with a 3,5 μL droplet of 2% uranium acetate (sigma Aldrich). After one minute the droplet of uranium acetate was discarded with a piece of filter paper. The grid was then introduce into the Morgagni 268 D electronic microscope to take electron micrographs.

2.14 Single Molecule Fluorescence Microscopy

The single molecule fluorescence tracking was measured at the LCPPM (laboratory of physical chemistry of polymers and membranes) laboratory at the EPFL. Single molecule experiments were performed at room temperature on a laser illuminated wide-field microscope. The setup consists of an excitation system, a microscope and a detection system. The excitation system is composed of a 532 nm and a 633 nm Helium-neon solid state diode lasers controlled by computer. The laser beam is directed to the sample with a dichroic mirror (412/532/633 nm, Chroma Corp) and a water immersion objective (63x / 1.2 W corr, Zeiss) mounted on a Zeiss Axiovert 200 wide-field microscope. Fluorescence emitted by the sample is collected back through the same pathway and directed to the detection system. A suitable filter avoids the residual excitation light of the laser to enter into the camera. Detection system consists of two cameras: an EM-CCD camera (electron multiplier charge- coupled device, Andor iXon 887 BV) and a CMOS fast camera (complementary metal oxide semiconductor, Andor Neo).

54 2. Material and Methods

Figure 2.4: Wide-field microscopy setup. The set up is composed of three main parts. Solid state diode laser, 532 nm or 633 nm, to excite the system. A microscope and a camera (EM-CCD or CMOS) for detection. Laser beam is reflected by the dichroic mirror with appropriate wavelength and collimated by the objective and illuminates a wide region (wide-field) of the sample. The fluorescence emitted by the sample, on the cover slip, is filtered and imaged by the camera.

A 12 μl droplet of sample consisted of ~10 μM of labelled rhodopsin in disks in HEPES 100 mM pH: 7.0, was laid on a standard glass cover slip. After 10 minutes, the disks sediment and stick automatically to the glass through hydrophobic interactions. A droplet of water was placed on the immersion objective and the cover slip was laid on the objective. The 633 nm laser was used to excite the Atto 647 N maleimide dye while the 532 nm laser was used to excite fluorescein dye and the QD 655. The single molecule fluorescence emitted by the sample was then recorded with the EM-CCD camera for the measurements at 10 Hz and a CMOS camera for the measurements at 30 Hz.

55 2. Material and Methods

Spots resulting of the detection of single fluorescence molecule are extracted for each frame to localize in two dimensions the single molecule with the software Igor Pro. From the sequence of images, it is possible to track the motion of the rhodopsin in time. A molecule´s position in the first frame is compared to the position of the next frame. The resulting traces of single molecule were extracted from the 30 seconds videos. Several parameters can be extracted from the molecule trajectories like diffusion coefficient (D0), and length of confinement (Lc). To analyze a single-molecule trajectory, the mean square displacements (MSD) were calculated for each time interval (tlag) and plotted for all time intervals. The mean square displacement is a measure of the average distance of the average distance a molecule travels.

2 2 MSD(tlag )  ri t  ri t ri 0 (7)

In this equation, ri is the position of molecule i at time t. ri(t)-ri(0) is the (vector) distance travelled by molecule i over some time interval of length t, and the squared magnitude of this vector is averaged (as indicated by the angle brackets) over many such time intervals. This quantity is averaged also over all molecules in the system, summing i from 1 to N and dividing by N (Michalet 2010)(Schütz, Schindler, and Schmidt 1997).

Single molecules undergoing Brownian motion are characterized by a linear relation between the MSD and the diffusion coefficient D0 and the time lag.

MSD  4 D0 tlag (8)

In living cells, plasma membrane proteins motions are often hindered by the presence of other membrane proteins or membrane components. Their free diffusion is confined in domains. The confined molecules exhibit an asymptotic behaviour of the MSD vs tlag plot. In the confined diffusion model, diffusion of the molecule is free within a square of side length Lc, which is surrounded by an impermeable, reflecting barrier. In such a model the mean square displacement depends on Lc and the initial diffusion constant D0, and varies with tlag(Lommerse et al. 2004).

56 2. Material and Methods

L2   12 D t  MSD t  c 1 exp 0 lag  (9)  lag    2  3   Lc 

Fit of the MSD vs Tlag plot with the equation (9) yields the values of the diffusion D0 of the molecule and for the length of confinement Lc of its diffusion.

57 3. Results

58 3. Results

3. Results

3.1 Peptides derived from arrestin finger loop stabilized activated rhodopsin

The aim of the work, present in this chapter, is to study how can arrestin bind the cytoplasmic crevice of the active receptor and desensitize the receptor. What are the constraints used between the finger loop and the receptor crevice and what are the intramolecular interactions in the finger loop. The arrestin family contains only four members which share a homologous sequence and similar structures. The arrestin finger loop is suggested to be the main loop interacting with the receptor (Sommer, Hofmann, and Heck 2012)(M. Kim et al. 2012)(Shukla et al. 2014). The C-terminus of Gt alpha is known to be the key binding to bind the receptor and stabilize Meta II. The question arises if there is a shared motif between arrestin and Gt alpha C-terminus and if this share motif would be also the key binding for arrestin to bind the receptor crevice. Is this motif the Meta II sensor of the four arrestin types? We therefore performed a sequence alignment of the finger loop region of the four arrestins and Gt alpha C-terminus. The sequence alignment shows a conserved sequence (E/D)x(I/L)xxxGL between the four arrestins and the Gt alpha C-terminus (Fig. 3.1.1).

res. Arr-1 visual arrestin 64 A F R Y G Q E D I D V M G L S F 79 Arr-2 ß-arrestin 1 60 A F R Y G R E D L D V L G L T F 75 Arr-3 ß-arrestin 2 61 A F R Y G R E D L D V L G L S F 76 Arr-4 Cone arrestin 56 A F R Y G H D D L D V I G L T F 71 Similarity * * * * * : : * : * * : * * : *

res. Gt Gt alpha 336 T D I I I K E N L K D C G L F 350 common seq. E/D x L x x x G L

Figure 3.1.1: Finger loop region alignment of all four bovine arrestin types and C-terminus of Gt alpha. Sequence alignment of Arrestin-1 (visual arrestin, residues 64-79, UniProt P08168), Arrestin-2 (ß-arrestin 1, residues 60-75, UniProt P17870), Arrestin-3 (ß-arrestin 2, residues 61-76, UniProt P32120), Arrestin-4 (Cone arrestin, residues 56-71, UniProt Q9N0H5) and C-terminus of Gt alpha (UniProt P04695). Alignment performed with clustalo program. Star and grey colour represent a total homology, colon represents high similarity between the different arrestins. Blue colour shows conserved sequence between all arrestins and Gt alpha terminus.

59 3. Results

res. 340 342 344 346 348 350 Gt alpha I K E N L K D C G L F Gt alpha HAA I L E N L K D C G L F

res. 384 386 388 390 392 394 Gs alpha Q R M H L R Q Y E L L

Figure 3.1.2: The Gt and Gs alpha C-terminus analogue peptide sequence alignment. Sequence alignment of the eleven last amino acids of bovine Gt and Gs alpha C-terminus (Uniprot entree P04695, residues 340-350 and P04896, residues 384-394 respectively). High affinity analogue Gt peptide (HAA) is analogue to the native peptide. Only on residue is mutated at position 341. Both Gt alpha peptide have the (E/D)x(I/L)xxxGL indicted in light blue.

In collaboration with Dr. Sczcepek and Dr. Scheerer the functionality and the structure of finger loop interacting with rhodopsin were investigated. Based on the observation of the conserved sequence between the arrestin and C-terminus of Gt alpha, different peptides mimicking the arrestin finger loop (Fig. 3.1.3) were investigated to detect the functionality of the peptides and to locate which amino acids or sequence of amino acids from the arrestin finger loop are required to bind the receptor. In parallel, Dr. Sczcepek and Dr. Scheerer co-crystallized and analyzed the structure of these peptides bound to active receptor. Part of this work is published in the Nature Communication journal (Szczepek, Beyrière, Hofmann, Elgeti, Kazmin, Rose, Bartl, et al. 2014a). Constrains for designing the peptides are as follows: First of all, peptides should not exceed eleven amino acids, because Gt alpha peptides exceeding eleven amino acids protrude past the crystal lattice and thus do not crystallize. Short peptides under 11 amino acids were avoided, because they are assumed to have not enough binding energy to bind the crevice. In the case of Gt alpha, the interacting part with the receptor is a C-terminus. In contrast, arrestin engage a loop (Shukla et al. 2014) so that some peptides were extended with a maximum of two amino acids in the extension of the Gt alpha C-terminus.

60 3. Results

Arrestin 1 res. 68 70 72 74 76 78 Kd ArrFL-1 (69-79) Q E D I D V M G L S F - ArrFL-1 (68-79) G Q E D I D V M G L S F - ArrFL-1 (67-79) Y G Q E D I D V M G L S F - ArrFL-1 (67-77) Y G Q E D I D V M G L 0,8 mM ArrFL-1 (69-77) Q E D I D V M G L -

Arrestin 2 res. 64 66 68 70 72 74 ArrFL-2 (65-75) R E D L D V L G L T F - ArrFL-2 (64-75) G R E D L D V L G L T F 0,6 mM ArrFL-2 (63-75) Y G R E D L D V L G L T F 0,35 mM ArrFL-2 (63-74) Y G R E D L D V L G L T 1,95 mM ArrFL-2/3 (63-73) Y G R E D L D V L G L 1,26 mM

Arrestin 3 res. 64 66 68 70 72 74 76 ArrFL-3 (66-76) R E D L D V L G L S F - ArrFL-3 (65-76) G R E D L D V L G L S F 1,18 mM ArrFL-2/3 (64-74) Y G R E D L D V L G L 1,26 mM

Arrestin 4 res. 60 62 64 66 68 70 ArrFL-4 (59-69) Y G H D D L D V I G L -

Figure 3.1.3: Table of the finger loop arrestin peptide series derived from the four arrestin types. The different peptides derived from the finger loop of the four different arrestin families. Residues highlighted in blue represent the (E/D)x(I/L)xxxGL motif. Green colour, the conserved residues and salmon colour the deviated residues. In red, missing residues suspected to play a role in the binding. The real Kd are summarised in the table. Unbound peptide are depict with a dash (-).

Three to five peptides per arrestin type were selected (Fig. 3.1.3), except for arrestin-4. This arrestin is known to be specific for cone rhodopsin only, and our first measurement with ArrFL-4 (59-69) (Fig. 3.1.5, D) shows no binding and suggests that the arrestin 4 derived peptides would not bind. The arrestin 2 and 3 share a common sequence for the finger loop (Arr-2: 63-73, Arr-3: 64-74). The arrestin-2/3 peptide represents this sequence and thus belongs to the two series. In parallel peptides derived of C-terminus of G alpha subunit of Gt and Gs peptides were selected. These peptides consist of the eleven amino acids of G alpha C-terminal of Gs and Gt (Fig. 3.1.2) and were used to compare this common sequence between different G alpha proteins. Gs alpha subunit is known not to bind activated rhodopsin like its C-terminus homologous peptide (Natochin et al. 2000)(Cerione et al. 1985). It does not contain the (E/D)x(I/L)xxxGL motif. In contrast, Gt alpha is the associated G protein subunit for rhodopsin. The Gt alpha C-terminus peptide binds and stabilizes Meta II (Herrmann et al. 2006). The

61 3. Results

structure of the peptide within the rhodopsin crevice was reported (Scheerer et al. 2008). Finally, the Gt C-terminal high affinity analogue peptide (Gt HAA: ILENLKDCGLF) contains one amino acid exchange at position 341 which give it the capacity to bind Meta II with higher affinity (Herrmann et al. 2006). Gt HAA was used as control. Functionality of the peptides was measured through the Extra Meta II assay (see methods). This technique permits to measure the formation of the 380 nm-absorbing active Meta II state induced by specific binding of the peptide to the receptor as do the full length arrestin and Gt. Measurements were performed at pH 8.0 and 0°C and the binding was initiated with a flash. Recorded traces represent the time dependency of absorption differences (A380–A417nm; the isosbestic point between Meta I and Meta II is at 417 nm, Fig. 3.1.4, A) from samples containing 10 μM phosphorylated rhodopsin and increasing concentrations of peptide up to 6 mM. The final amplitudes of each peptide concentration were then plotted in function of peptide concentration. The maximum amount of Meta II of the sample was controlled with the record of 300 μM of Gt alpha HAA. Maximal Meta II formation with Gt alpha HAA is 0,01 OD. The titration data were fit to a hyperbolic expression

(Bmax*x/Kd+x) and the apparent Kd was corrected for the MetaI/MetaII equilibrium to get the real Kd (Parkes and Liebman 1984b).

3.1.1 Arrestin-1 finger loop peptide series

The Extra Meta II assay with the ArrFL-1 peptides, derived from the finger loop of visual arrestin, shows that only the ArrFL-1 (67-77) binds the receptor (Fig. 3.1.4, A and B). The resulting binding curve yields a value for the real Kd of 0,8 mM similar to a previous study (Feuerstein et al. 2009). The ArrFl-1 (68-69) peptide data points suggest a weak Meta II increasing upon peptide concentration. However, the slope of the binding curve is too shallow to be analyzed reliably (Fig. 3.1.4, E). The other peptides do not stabilize Meta II at all (Fig. 3.1.4, C, D and F).

62 3. Results

Figure 3.1.4: Extra Meta II formation induced by titration of ArrFL-1 peptide series with phosphorylated ROS. A. The signal represents the absorption change at 380 nm minus the absorbance change at 417 nm over time for different ArrFL-1 (67-77) peptide concentration at 0°C, pH: 8.0. In red, trace of HAA Gt alpha peptide (300 μM) representing the maximal amount of Meta II. B. Plot of the final amplitudes of the ArrFL-1 (66-77) titration from A. Different symbols represent different data sets. Solid line represents data points fit which yields an apparent Kd of 0,8mM. The Kd are summarized in Fig. 3.1.2 C. Titration of ArrFL-1 (69-79). D. Titration of ArrFL-1 (69-77). E. Titration of ArrFL-1 (68-79). F. Titration of ArrFL-1 (67-79).

63 3. Results

3.1.2 Arrestin-2 finger loop peptide series

In comparison the peptides derived from the finger loop of arrestin-2, arrestin-3 and arrestin-4 were titrated against 10 μM rhodopsin and Meta II formation induced by peptide binding was measured. The finger loop from arrestin-2 differs from arrestin-1 by three residues: Arg65, Leu68 and Thr74.

Figure 3.1.5: Extra Meta II formation induced by titration of ArrFL-2 peptide series. Titration of different peptides against 10 μM phosphorylated rhodopsin. Difference absorption between 380 nm and 417 nm was measured in function of the peptide concentration. A. Titration of ArrFL-2 (64-75). True Kd corrected with the MetaI/MetaII equilibrium is 0,59 mM. B. Titration of ArrFL-2 (63-74) presents a true Kd of 1,9 mM. C. Titration of ArrFL-2 (65-75). The fitted curves plateaus at 0,002 and should not speak for a specific Meta II stabilization D. Titration of ArrFL-2 (63-75) presents a true Kd of 0,36 mM.

The binding curves of ArrFL-2 peptides yield a real Kd of 0,36 mM, 0,59 mM and 1,9 mM for ArrFL-2 (63-75), ArrFL-2 (64-75) and ArrFL-2 (63-74) respectively (Fig. 1.3.5, A, B and D). Strangely, the ArrFL-2 (65-75) plateaus at 0,002 OD whereas the maximal Meta II formation of the HAA peptide is 0,011 OD (Fig. 3.1.5,

64 3. Results

C). This curve does not correspond to a standard Meta II stabilization. Despite the fact that the peptides were purchased with a purity of over 95%, we get different batches that had different colours. Thereby, the formation of Meta II upon peptide concentration could results from the impurities from the peptide synthesis disturbing the Meta I/Meta II equilibrium. Another possibility is that the peptide can bind Meta I and only a part of the peptide binds Meta II. No additional measurements have been proceeded to discover the reason of this behaviour.

3.1.3 Arrestin-3 and arresti-4 finger loop peptide series

Figure 3.1.6: Extra Meta II formation induced by titration of ArrFL-3 peptide series and arrestin-4 finger loop derivate. Each peptide was titrated with 10 μM phosphorylated rhodopsin in 100 mM HEPES, pH 8.0, 0°C. A. Titration of ArrFL-2/3 (64-74). Different symbols represent different data sets. Fitted curves yield a real Kd of 1,3 mM B. Titration of ArrFL-3 (65-75) yields a Kd of 1,18 mM. C. Titration of ArrFL-3 (66-76). D. Titration of ArrFL-4 (59-69) presents no binding of the arrestin-4 peptide.

65 3. Results

The finger loop of arrestin-2 and arrestin-3 are quite identical. Only the threonine at position 74 of arrestin-2 differs from the serine of arrestin-3. Thus the arrestin 2 and 3 share a sequence, so that the ArrFL-2/3 peptide belongs to the two arrestin types.

The arrestin-3 peptides have a real Kd of 1,18mM for ArrFL-3 (65-76) and 1,3 mM for the ArrFL-2/3 (64-74) (Fig. 3.1.6, A and B). No binding was detected with the ArrFL-3 (66-76) and for ArrFL-4 peptide derivates (Fig. 3.1.6, C and D).

3.1.4 G alpha C-terminus peptides

Peptides derived from Gt and Gs alpha subunits were also titrated against 10 μM phosophorylated rhodopsin. The Gs alpha subunit does not contain the (E/D)x(I/L)xxxGL motif and does not bind activated rhodopsin (Fig. 3.1.7, C). The absence of Gs alpha binding confirms that G alpha protein requires (E/D)x(I/L)xxxGL sequence to bind activated rhodopsin.

Titration of native Gt alpha yields a real Kd of 0,18 mM. The high affinity peptide analogue has a real Kd of 2,38 μM (Fig. 3.1.7). These two Gt alpha peptides bind with a higher affinity to activated receptor compared to the arrestin derived peptides. The Gt high affinity analogue peptide (HAA) contains one amino acid exchange at position 341. Exchange of lysine residue to leucine increases drastically the affinity of the peptide to the receptor (Herrmann et al. 2006). As does transducin, these two Gt alpha peptides bind activated receptor.

66 3. Results

Figure 3.1.7: Extra Meta II formation induced by titration of G alpha homologue peptides. Each peptide was titrated with 10 μM phosphorylated rhodopsin in 100 mM HEPES, pH 8.0, 0°C. A. Titration of native Gt alpha peptide (340-350). Fitted curves presents a real Kd of 0,18 mM B. Titration of the Gt alpha high affinity analogue peptide (HAA). Real Kd of 2,4 μM. C. Titration of native Gs alpha C-terminus analogue peptide (340-350). Different triangle symbols identify different data sets.

3.2 Arrestin complex formation studied by Fourier transform infrared spectroscopy

Arrestin binding to activated phosphorylated rhodopsin occurs via a stepwise process (see Introduction, chapter 1.6.2). Arrestin recognizes the phosphorylated tail of the receptor and forms the pre-complex. Replacement of arrestin C-terminus by the receptor tail induces conformational changes in arrestin leading to its activation. This intramolecular reorganization of arrestin allows a following conformational change to fit and bind the activated receptor and finally form the high affinity complex. Crystal structure of pre-active state of arrestin (Shukla et al. 2013)(Y. J. 67 3. Results

Kim et al. 2013) presents a twist of the two domains and a displacement of the loops a the central crest region and gives a general idea of structural organization of the pre-complex. However, no structure is available for the high affinity complex. In this part of the project, the interaction of arrestin with rhodopsin is monitored with Fourier transform infrared (FTIR) spectroscopy. This method allows to monitor functional changes in proteins without labelling. The transition between different conformations can be detected and compared to the different Meta substate spectra already known (Mohana Mahalingam et al. 2008)(Zaitseva, Brown, and Vogel 2010)(Matthias Elgeti et al. 2011). This project was effectuated in collaboration with Dr. Eglof Ritter. The primary analysis of the row-data, composed of base line correction, singular value decomposition (SVD) and global fit (see Methods), was done by Dr. Ritter. Most part of this project was submitted in Journal of Biological Chemistry.

3.2.1 Influence of the phosphorylation on rhodopsin

The properties of unphosphorlated rhodopsin are well known and its Meta substates are spectroscopically well characterized. No FTIR investigation on phosphorylated rhodopsin was done till now. Thus the contribution of the phosphate on their FTIR spectra is unknown. In order to determine the influence of the phosphorylation on rhodopsin structure and its Meta II state, formation of Meta II was measured for rhodopsin and phosphorylated rhodopsin. Measurements were performed at pH: 7.0 at 20°C, conditions under which the Meta I/Meta II equilibrium is shifted towards Meta II (~80% of Meta II for unphosphorylated rhodopsin, (Parkes and Liebman 1984b)). The absorption spectrum of dark phosphorylated rhodopsin at ~350 µM in isolated membrane presents five usual distinct vibration regions (Fig. 3.2.1, A). Vibrations of protonated carbonyl groups show up between 1700 and 1800 cm-1. Vibration of amide I, amide II and amide III are located near the 1650, 1550 and ~1450 cm-1 respectively. Vibration patterns for phosphate are located around 1200 and 1100 cm-1.

68 3. Results

Figure 3.2.1: Influence of the phosphorylation on Meta II conformation. A. Absorption spectrum of phosphorylated dark rhodopsin. Vibrations of protonated carbonyl groups show up between 1700 and 1800 cm-1 (I). Vibration of amide I (II), amide II (III) and amide III (IV) are located near the 1650, 1550 and ~1450 cm-1 respectively. Vibration patterns for phosphate are located around 1200 and 1100 cm-1 (V). B. Difference spectra (light minus dark) of rhodopsin (blue) and phosphorylated rhodopsin (red) at 20°C, pH: 7.0. C. Double difference of the two difference spectra highlights the deviation between both spectra. The spectra were normalized with the retinal isomerization band at 1238 cm-1 of the difference spectra.

69 3. Results

The difference spectra (light minus dark, Fig. 3.2.1) were calculated for unphosphorylated rhodopsin (blue) and for phosphorylated rhodopsin (red). It indicates a mixture of Meta II substates spectrum with a majority of Meta IIbH+ spectrum. Negative bands indicate vibration modes of the dark states of the receptor while positive bands represent vibration modes of the illuminated rhodopsin in Meta II state. Meta II features include the prominent marker bands at 1768 cm-1 (C=O -1 mode of protonated Asp83; Meta IIa), 1727/ 1748 cm (protonated Asp83 and Glu122; Meta IIb/ Meta IIbH+), 1713 cm-1 (protonation of Glu113, counter ion of the Schiff base, Meta IIa) and 1644 cm-1 (amide I mode of Meta IIb/ Meta IIbH+) (Siebert 1995)(Siebert, Mantele, and Gerwert 1983)(Mahalingam et al. 2008). Comparison of these two difference spectra, representing the transition of dark rhodopsin to Meta II in unphosphorylated or in the phosphorylated case, reveals almost no deviations. A double difference (Fig. 3.2.1. C, green) of both difference spectra was calculated to highlight the small differences due to the phosphorylation. Although the big intensity (~1 mOD) of bands in the amide I region, these vibration do not represent significant structural changes. Comparison of two different samples induces deviations due to the slight deviation of pH and of the different water content of the samples and/or of a slight shift of MetaI/MetaII equilibrium due to the phosphorylation (see further). Moreover, these deviations might be caused by detector non-linearities. These data show that the phosphorylation does not induce significant structural deviations on Meta II conformation. No changes are also observed between 1000 and 1200 cm-1 where vibration of phosphate groups occur (Liu and Barth 2003)(Allin et al. 2001)(Klähn et al. 2004) which signify that the phosphates at the C-terminus of rhodopsin do not affect the overall structure of Meta II. To determine the influence of the phosphorylation on the Meta I/Meta II equilibrium, the same experiment was performed at pH: 8.0 at 1°C. At these conditions, the Meta I conformation is favoured (90% of Meta I for unphosphorylated rhodopsin (Parkes and Liebman 1984b)). The difference spectrum of unphosphorylated rhodopsin (Fig. 3.2.2, blue) presents a Meta I conformation (1662 and 952 cm-1) whiled phosphorylated rhodopsin (red) presents significant deviations indicative for increased contribution of the Meta II intermediate (1768, 1747, 1713, 1552 and 1644 cm-1)(Siebert 1995)(Siebert, Mantele, and Gerwert

70 3. Results

1983)(Reiner Vogel and Siebert 2003) (for typical Meta II spectrum, see Introduction Fig. 2.3 or Fig. 3.2.3). The double difference (phosphorylated rhodopsin minus unphosphorylated rhodopsin, green) clearly presents a spectrum typical for the Meta I → Meta II transition which indicates a shift of the Meta I/Meta II equilibrium towards Meta II due to the phosphorylation of receptor tail. Illumination results in 75% of Meta I and 25% of Meta II with phosphorylated rhodopsin whereas unphosphorylated rhodopsin forms only 10% of Meta II. No other additional species besides the mixture of Meta I and Meta II conformations are present. These data confirm previous studies (Gibson et al. 1998)(Gibson, Parkes, and Liebman 1999) showing the phosphorylation dependent shift of the Meta I/Meta II equilibrium.

Figure 3.2.2: Influence of the phosphorylation on the Meta I/Meta II equilibrium. Difference spectrum (light minus dark) of rhodopsin (blue) and phosphorylated rhodopsin (red) at 1°C, pH: 8.0. Both spectra were normalized with the retinal isomerization band at 1238 cm-1. Phosphorylated rhodopsin contains more Meta II as the unphosphorylated rhodopsin. Double difference spectrum of both difference spectra (bottom, green) presents a clear Meta I → Meta II transition spectrum. 71 3. Results

3.2.2 Arrestin binding difference spectrum

A mixture of phosphorylated rhodopsin with arrestin was prepared and centrifuged. The resulting pellet, with a concentration of ~350 μM phosphorylated rhodopsin and excess of arrestin was used for FTIR measurements. At this concentration, the arrestin is already pre-bound to the phosphorylated receptor (see discussion, chapter 4.2.1). Complex formation was monitored with 200 ms time-resolution before, during and after the five-second orange light illumination. Arrestin binding to phosphorylated rhodopsin was first investigated under conditions favouring Meta II. At pH: 8.0 and 30°C this equilibrium is shifted to Meta II (Parkes and Liebman 1984b). Over 90 % of all light activated phosphorylated rhodopsin are in Meta II conformation at these conditions. To represent the light activation transition occurring in the sample, a difference spectrum (light minus dark) was calculated (Fig. 3.2.3, red). Negative bands indicate vibration modes of the dark states of the rhodopsin mixed with arrestin while positive bands represent vibrational modes of the illuminated state of rhodopsin and changes induced in arrestin upon complex formation.

Figure 3.2.3: FTIR difference spectrum of arrestin complex formation. Difference spectrum (light minus dark) of phosphorylated rhodopsin in presence of arrestin (red) at 30°C, pH: 8.0. Difference spectrum of phosphorylated rhodopsin, at 30°C, pH: 5.0 (blue), represents a typical Meta IIbH+ spectrum. Both spectra were normalized with the retinal isomerization 1238 cm-1 band.

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The difference spectrum of arrestin binding was compared with a standard Meta II spectrum (blue) of rhodopsin, recorded in absence of arrestin at pH: 5.0, 30°C, optimal conditions for the formation of the Meta IIbH+ state (M Mahalingam et al. 2008)(Matthias Elgeti et al. 2013) and normalized on the amplitude of the 1238 cm-1 integrated band region, representing the isomerisation of the retinal (Siebert, Mantele, and Gerwert 1983). Comparison of difference spectrum of arrestin binding and pure Meta IIbH+ shows high similarities. Difference spectrum of arrestin binding shares the same typical Meta II features as the pure Meta IIbH+. Meta II features include the prominent -1 marker bands at 1768 cm (C=O mode of protonated Asp83; Meta IIa), 1727/ 1748 cm-1 (protonated Asp83 and Glu122; Meta IIb/ Meta IIbH+), 1713 cm-1 (protonation of Glu113, counter ion of the Schiff base, Meta IIa) and 1644 cm-1 (amide I mode of Meta IIb/ Meta IIbH+) (Siebert 1995)(Siebert, Mantele, and Gerwert 1983)(Mahalingam et al. 2008). The protuberant band at 1661 cm-1 identifies formation of Meta IIbH+ (Zaitseva, Brown, and Vogel 2010)(Matthias Elgeti et al. 2013). These marker bands are identical for both spectra and suggest that no deviations of the hydrogen bonded networks involving these residues exist. These first measurements confirm that arrestin binds Meta II (Schleicher et al. 1989). However major deviations are observed between 1700 and 1500 cm-1 and attest of the formation of the arrestin complex. The difference spectrum in presence of arrestin is a superposition of the difference spectrum of receptor activation (formation of Meta II) and of all conformational changes induce in both proteins upon complex formation. A positive band at 1661 cm-1 indicates an additional vibrational mode in the illuminated sample. Additionally, the strong negative band at 1694 cm-1, accompanied by the negative band at 1624 cm-1, suggest a loss of a particular vibrational mode upon illumination. In the amide-II region, the negative band 1555 cm-1 and positive band 1533 cm-1 are more pronounced. Interestingly, no deviation in the region below 1300 cm-1 where the vibrational modes of the phosphate groups are expected (Klähn et al. 2004)(Allin et al. 2001) is observed. This absence of changes in the phosphate region suggests that no alterations occur at the phosphorylated tail upon high affinity complex formation and confirms that the arrestin is already pre-bound in the dark.

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3.2.3 Arrestin binding spectrum

To analyse the specific contributions of arrestin to the spectrum, the Meta II contribution induced by arrestin (“Extra-Meta II”) was retrieved of the spectrum. A pure Meta II spectrum, whose intensity corresponds to the Meta II amount in the spectrum, was substracted of the different spectrum (see Methods). The double difference spectrum is called ´´arrestin binding spectrum´´(ABS) (Fig. 3.2.4, red). The double difference spectrum is obtained by subtracting the difference spectrum with arrestin from the difference spectrum without arrestin (see methods). The double difference spectrum allows the extraction of the additional specific conformational changes to either of the proteins induced by complex conformation. The arrestin binding spectrum is the fingerprint for arrestin conformational changes and for conformational changes in Meta II due to arrestin binding.

Figure 3.2.4: Arrestin binding spectrum (ABS). Double difference of arrestin binding spectrum (red) representing all conformation changes induced on arrestin and on the Meta II due to the complex formation. The peptide binding spectrum (PBS) of Gtα protein C-terminus analogue peptide (green) represents all conformational changes of peptide and of Meta II conformation due to peptide binding.

The ABS reveals new patterns specific for the arrestin complex formation. Positive bands represent vibrational modes gained upon arrestin binding and the negative ones, the vibrational modes loosed upon complex formation. They represent structural changes occurring in arrestin and in Meta II upon complex formation. The activation of rhodopsin, the formation of Meta II, is already retrieved in the ABS so 74 3. Results

that the ABS represent only changes induces by arrestin binding (in arrestin and in Meta II). A significant positive band is present in the carbonyl region of the ABS at 1716 cm-1. The amide I region presents two large negative bands at 1964 and 1625 cm-1 and three pronounced positive bands at 1685, 1660 and 1642 cm-1. A positive and a negative band at the amide II region are also present. Many bands are overlapped and it is a real challenge to assign bands to a specific structural change or, even more difficult, to specific residues. A peptide binding spectrum (PBS, Fig. 3.2.2 green) was measured for comparison with the ABS. Dark rhodopsin samples were prepared with the Gt alpha HAA analogue peptide and measured under the same conditions as the ABS. Difference spectrum and double difference spectrum were calculated as in case of arrestin. The ABS shares with the PBS the large positive band at 1660 cm-1. The G-protein analogue peptide was already studied and its binding signature well characterized (Reiner Vogel et al. 2007). A global isotopic labelling of the peptide was used to assign the large 1660 cm-1 band to conformational changes of rhodopsin induced by peptide binding and also to a structural rearrangement of the peptide upon binding. It was shown that the binding of the Gt alpha analogue peptide induces conformational changes in the rhodopsin and in the peptide. The 1661 cm-1 band in the ABS could also represent a helix formation upon high affinity binding, however, due to the overlap of many different bands in this region, the assignation of this 1661 cm-1 band is uncertain (see discussion). The absence of changes in the region of phosphate bands (1200-1100 cm-1) confirms the observation of the difference spectrum measured at (Fig. 3.2.1). No alterations are occurring at the phosphorylated tail upon high affinity complex formation confirming that the arrestin is already pre-bound in the dark.

3.2.4 Arrestin binding spectrum (ABS) in D2O

The two large negative bands, 1625 and 1694 cm-1, occur in a spectral region where the vibrational modes of beta sheets are observed (Barth 2007). Beta sheet structures can be characterized by two vibrational modes (high- and low-frequency) (Table 3.2.1).

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Table 3.2.1: Assignment of amide I band positions to secondary structure. Vibration patterns for the different secondary structures in water and deuterium. The two vibrations characteristic for beta sheet are highlighted in blue. Table from (Barth 2007).

To prove the origin of these two bands and to confirm their assignment to beta sheet,

H2O was exchanged to D2O in the sample. The mass difference between the isotopes induces a small change in the vibration frequency of the molecules in particular in amide I region. Characteristic shifts of hydrogen coupled vibrations allow the assignment of these bands to peptide groups or to specific secondary structure (Kapoor et al. 2011).

Figure 3.2.5: Influence of deuterium on arrestin binding spectrum. The ABS spectrum in D2O -1 (black) presents a shift of the bands 1694, 1660 and 1625 cm compared to the ABS in H2O (red).

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Compared to the arrestin binding spectrum in H2O, the ABS in D2O (Fig. 3.2.5, black) presents specific shifts for the bands at 1694, 1661 and 1625 cm-1 while the other bands are conserved. The shift of the negative bands at 1694 and 1625 cm-1 in -1 H2O (red) to 1684 and 1622 cm respectively in D2O corresponds to the values expected for beta sheet (table 3.2.1)(Barth 2007). Crystal structure of arrestin-1 (Hirsch et al. 1999) shows that half of arrestin structure is composed of beta sheets and it is not surprising that a beta sheet rearrangement occurs upon complex formation. These negative beta sheet markers mean that a loss of beta sheet structure occurs upon arrestin binding. Comparison of the amplitude of the 1629 cm-1 band of the ABS with those of the absorption spectra of arrestin rhodopsin mixture in dark yields a change of ~5% of beta sheet due to the complex formation. Finally, the large -1 positive band at 1660 cm shifts for ~5 wavenumbers in D2O and could correspond to a helix formation upon the reaction (see discussion). The analysis of the ABS shows that the different binding spectrum of arrestin binding (Fig. 3.2.3, red) contains the usual marker for Meta substates of rhodopsin (1768, 1748, 1713, 1661 and 1644 cm-1) and accompanied of distinct beta sheet markers (1694 and 1624 cm-1), which are directly linked to arrestin (see discussion). The simultaneous presence of these distinct markers for both proteins gives a powerful opportunity to monitor in the same time, and from both sides, arrestin and receptor, the high affinity complex formation triggered by light.

3.2.5 Arrestin binding kinetic

In order to monitor the receptor changes induced by the binding of arrestin, the complex formation was measured under Meta I conditions, pH: 8.0, 0°C. A UV-Vis spectroscopy study showed already that arrestin binding stabilizes the receptor in its Meta II conformation (Schleicher et al. 1989). However, this technique is not able determine which Meta II substate is favored. FTIR is powerful technique to determine the conformational changes, and thus ideal to determine the Meta substates of rhodopsin. Measurements of arrestin complex formation at Meta I condition present an identical spectrum as for Meta IIbH+ conditions (Fig. 3.2.1) proving that the receptor is stabilized in Meta IIbH+ conformation. This data confirms that arrestin binds and stabilizes Meta II conformation (extra Meta II). However, global analysis of the time-dependent datasets of light-activation can

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extract the different component occurring upon reaction. In the case of arrestin binding, a biphasic behavior is observed. A fast component (Fig. 3.2.6, A, red), completed within the illumination period, with a half time of less than one second and a slow component (Fig. 3.2.6, B, red) with a half time of ~20 s. Both component present the same amplitude so that the total reaction is composed of 50% of fast component and 50% of slow component. To determine the nature of these two transitions, same experiment was conducted with phosphorylated rhodopsin without arrestin.

Figure 3.2.6: The two components of the arrestin binding. A. Fast component of arrestin binding (red). Transition of rhodopsin to Meta I/Meta II (blue). Spectrum amplitudes were normalized to the 1238 cm-1 band. B. Slow component of arresting binding (red). Spectrum of Meta I to Meta II transition (blue). C. Double difference of the slow component (slow component minus transition of Meta I to Meta II, light green) and ABS (dark green).Measure conditions: 0°C, pH: 8.0.

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Global analysis of the time-dependent dataset of light-activation of phosphorylated rhodopsin reveals a fast monophasic transition (t1/2 < 1 s). It represent the transition of rhodopsin to a mixture of 75% Meta I and 25% Meta II (Fig. 3.2.6, B, blue). The fast transition of the complex formation shows a similar spectrum and the analysis reveal a transition of ~50% to Meta I. The increased Meta II content, in the fast transition, is seen by the marker bands at 1768, 1748 and 1713 cm-1. The prominent band at 1661 cm-1 identifies formation of Meta IIbH+ (Matthias Elgeti et al. 2013). In the same spectrum (Fig. 3.2.6 A, red) arrestin binding is also clearly identifiable by the marker bands bands at 1694, 1624 and 1555 cm-1. The extra Meta II and the presence of the arrestin markers signify that ~50% of the phosphorylated activated rhodopsin are bound by arrestin and are stabilized in the Meta II conformation. + The subsequent slow component (t1/2= ~20s) was compared with a pure Meta IIbH minus Meta I difference spectrum (Fig. 3.2.6, B, blue). It represents the transition of Meta I to Meta IIbH+ (extra Meta II). The slow component spectrum presents the typical pattern of Meta II formation (1768, 1748, 1713, 1644 and 1661 cm-1) and arrestin binding (1684, 1624 and 1555 cm-1). The carbonyl pattern (1800- 1700 cm-1), where specific band for Meta II are present, matches with the calculated spectrum and represents the transition of the 50% of residual Meta I which is converted to Meta II by arrestin binding. The double difference of the slow component minus transition of Meta I to Meta II (Fig. 3.2.6, C, light green) was calculated and compared to the ABS (Fig. 3.2.6, C, dark green). Both are similar indicating that the slow and the fast component have the same spectral signature and thus represent the same transition: from pre-bound arrestin to illuminated high affinity complex. However, these two similar transitions occur with two different time courses. The reaction of light activation is composed of cis/trans isomerization (characterized by the band at 1238 cm-1), Meta IIb formation (1748 cm-1) and the loss of arrestin beta-sheet associated with high-affinity binding (1625 cm-1). The overall kinetics of arrestin binding is plotted in figure 3.2.7. High affinity complex formation (red traces) is identified by the bands reflecting structural changes leading to Meta II conformation (1748 cm-1, red) and by the loss of beta sheet structuring (1695 and 1625 cm-1, red). Both occur simultaneously with the same biphasic behaviour (slow and fast kinetics) as shown by the two components of the global analysis. This kinetic similarity indicates that the loss of arrestin beta-sheet corresponds to the

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stabilization of Meta II and that these two events are correlated as the global fit already shows (Fig. 3.2.6, A and B, red). In comparison, in absence of arrestin, phosphorylated rhodopsin (blue traces) bands for Meta II formation (1747 cm-1, blue) occurs with a fast monophasic transition. Note that retinal isomerisation, depicted by the 1238 cm-1 band, is monophasic and does not depend on the presence of arrestin. The isomerisation occurs immediately after illumination. The reaction is so quick that only one component was detectable in the global analysis.

Figure 3.2.7: Biphasic behaviour of arrestin binding. Time-courses of the marker bands of Meta II formation (1748 cm-1, magnified four times) and of beta sheet marker (1625 cm-1) in presence (red) and absence (blue) of arrestin. Spectra were normalized with the chromophore band (1238 cm-1). Measurement at 30°, pH: 8.0. Illumination at t=0s.

All these data show that arrestin binds the active receptor conformation and forms the high affinity complex through two time courses. Essentially all accessible light activated receptors were converted into Meta II (> 90%). Arrestin binds quickly half of the accessible activated rhodopsin population and stabilizes its Meta II conformation. The other half of receptor, initially in Meta I conformation, reacts slower and is bound and stabilized by arrestin in a slower manner.

3.2.6 Complex formation of p44 with phosphorylated rhodopsin

In order to better understand the arrestin binding, measurements with p44, the splice variant of arrestin, and (phosphorylated) rhodopsin were performed. This truncated arrestin is already in a preactivated form and does not require phosphorylation to

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bind activated receptor. p44 binds phosphorylated activated rhodopsin and as unphosphorylated activated rhodopsin. p44 binds phosphorylated rhodopsin with very high affinity and is already prebound to the dark-state phosphorylated rhodopsin even at low concentrations (Schröder, Pulvermüller, and Hofmann 2002). p44 was purified with the help of Dr. Michael Szczepek. Samples containing p44 and phosphorylated rhodopsin where prepared like for the arrestin case and FTIR measurements were effectuated under the same conditions favour Meta I (pH: 8.0, 0°C). Binding spectrum (light minus dark, Fig. 3.2.8, blue) of p44 is similar to the binding spectrum of the full length arrestin (red). As arrestin, p44 stabilizes the rhodopsin in the Meta IIbH+ conformation (1768, 1748, 1713, 1661 and 1644 cm-1). No differences are observable at the phosphate vibrations region (1250-1150 cm-1) suggesting that all p44 are already pre-bound to the receptor phosphorylated tail in the dark state.

Figure 3.2.8: Difference spectrum of p44 with phosphorylated rhodopsin. Difference spectrum (light minus dark) of phosphorylated rhodopsin in presence of p44 (blue) at 0°C, pH: 8.0. Difference spectrum of phosphorylated rhodopsin with arrestin (red), at 0°C, pH: 8.0 (blue). Both present the typical Meta II markers. The spectra were normalized with the retinal isomerization 1238 cm-1 band.

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The double difference of p44 (p44 binding spectrum) was calculated like the ABS and is nearly similar as the ABS (Figure 3.2.9). It presents the same loss of beta sheet upon high affinity complex formation (1694 and 1625 cm-1). The amide I and amide II regions are also similar.

Figure 3.2.9: p44 binding spectrum and arrestin binding spectrum (ABS).Double difference spectrum of p44 with phosphorylated rhodopsin (purple) and arrestin binding spectrum of arrestin with phosphorylated rhodopsin (ABS). Positive bands represent vibrational modes gained upon arrestin binding and the negative ones, the vibrational modes loosed upon complex formation

Single value decomposition and global fit reveal, like for arrestin, a biphasic behaviour of the p44 binding. 58±5 % of the p44 binds the receptor and form the complex quickly (t1/2 = <1seconds) while the other 41±5 % of p44 binds slowly

(t1/2= ~25seconds). These two components are nearly identical to the components of the arrestin binding (Fig. 3.2.10). The fast one represents a superposition of the transition of dark rhodopsin to Meta I and to the formation of the complex. The slow component represents the binding of p44 to the residual phosphorylated rhodopsin in Meta I and stabilization of the Meta IIbH+ conformation (transition Meta I to Meta II•p44 complex).

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Figure 3.2.10: The two binding components of p44 and arrestin. The fast component (red) and slow component (blue) of p44 (top) and of arrestin (bottom) are respectively nearly identical. Spectra were normalized with the chromophore band at 1238 cm-1.

3.2.7 Complex formation of p44 with unphosphorylated rhodopsin p44 does not need the phosphorylated tail to bind the receptor, therefore p44 also binds unphosphorylated rhodopsin. Interaction of p44 with unphosphorylated receptor was measured and analyzed with the same protocol as with arrestin. The difference spectrum of p44 with rhodopsin is similar to those of p44 to phosphorylated receptor (Fig. 3.2.11, turquoise and blue) and present the usual Meta IIbH+ markers (1768, 1748, 1713, 1661 and 1644 cm-1). However, a slight different in the intensity of the 1625 cm-1 band representing the beta sheet marker is observable. The double difference spectrum, the binding spectrum of unphosphorylated rhodopsin with p44 (Fig. 3.2.11, orange) was calculated and compared to the binding spectrum of phosphorylated rhodopsin with p44 (Fig. 3.2.11, purple) to highlight deviations. The amide I and amide II regions are similar as p44 with phosphorylated rhodopsin. The biggest deviation is seen in 1625 cm-1 negative band. In absence of phosphorylation its intensity is halved. Surprisingly the second associated band 1694 cm-1 seems not be affected and has the same amplitude as in the case of phosphorylated receptor. 83 3. Results

Figure 3.2.11: Difference spectrum and binding spectrum of p44 with phosphorylated rhodopsin and with unphosphorylated rhodopsin. The difference spectrum (light minus dark) of p44 binding rhodopsin (turquoise) and phosphorylated rhodopsin (blue) at 0°C, pH: 8.0. Both difference spectra present the typical Meta II markers (1768, 1748, 1713, 1661 and 1644 cm-1). Double difference spectra of rhodopsin with p44 (orange) and phosphorylated rhodopsin with p44 (purple) represent all conformation changes induced on p44 and on the Meta II due to the complex formation. Positive bands represent vibrational modes gained upon arrestin binding and the negative ones, the vibrational modes loosed upon complex formation. The spectra were normalized with the retinal isomerization band at 1238 cm-1.

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Single value decomposition and global fit of p44 binding to rhodopsin reveals a biphasicity as in case of phosphorylated rhodopsin and arrestin. 60±5 % of the p44 bind activated rhodopsin quickly with a half time of <1 seconds (Fig. 3.2.12, A, red). While the other 40±5 % of p44 binds slower with a half time of ~20-30 s (Fig. 3.2.12, A, blue). The quick component represents the transition of dark rhodopsin to high affinity complex. The second component represents the binding of p44 to the residual rhodopsin in Meta I and stabilization of the Meta IIbH+ conformation. These two components are nearly identical to the components of the arrestin and p44 binding to phosphorylated rhodopsin (Fig. 3.2.10 and Fig. 3.2.12, B).

Figure 3.2.12: The two binding components of p44 with rhodopsin and phosphorylated rhodopsin. A. The fast component (red) and slow component (blue) of p44 with rhodopsin. B. The fast component (red) and slow component (blue) of p44 with phosphorylated rhodopsin. Spectra were normalized with the chromophore band at 1238 cm-1.

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3.2.8 Decay of the arrestin complex

Decay of the arrestin high affinity complex was already studied with fluorescence techniques (M E Sommer and Farrens 2006)(Sommer, Smith, and Farrens 2006)(Sommer, Hofmann, and Heck 2012). Surprisingly, it was shown that arrestin traps half of the retinal population in the binding pocket. However, the techniques used for these studies were not able to identify the rhodopsin species involved. The FTIR technique is able to monitor simultaneously Meta intermediates, arrestin and the retinal release during the decay of the complex. It also can identify the different conformations of the decay products formed during the retinal release. In this part of the project, decay of arrestin complex was investigated with time-dependent FTIR measurements. Samples consisting of phosphorylated rhodopsin with and without arrestin were measured at 30°C, pH: 8.0. Activation of rhodopsin and arrestin complex formation was initiated with light. After formation of the complex, which occurs rapidly at this temperature, the decay was investigated. The marker bands for arrestin (1694 and 1625 cm-1) and those for Meta IIb/Meta IIbH+ (1644cm-1) were followed upon time to monitor the formation (Fig. 3.2.6) and the decay of the complex with and without arrestin (Fig. 3.2.13).

Figure 3.2.13: Kinetic traces of arrestin complex decay. A. Time course of Meta II formation marker (1644 cm-1) with arrestin (red) and in absence of arrestin (blue). B. Time course 1694 and 1625 cm-1 in presence (red, the two beta sheet marker) and absence (blue) of arrestin. Measurements at 30°C, pH: 8.0. Illumination at time = 0s.

The amplitude of 1644 cm-1 in presence of arrestin (Fig. 3.2.13, A, red) is almost doubled compared to the traces in absence of arrestin. The binding of arrestin shifts the Meta IIb and residual Meta I to Meta IIbH+ resulting in higher amplitude of the

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1644 cm-1 in presence of arrestin. In absence of arrestin (Fig. 3.2.13, A, blue), the activated rhodopsin decays completely to the level seen before illumination. In presence of arrestin, the amplitude decays only to the half of the illuminated level. The decay of Meta II in presence and in absence of arrestin is monophasic and with similar kinetics. Decay in presence and absence of arrestin has a halftime of t1/2 = 100 s and of t1/2 =110 s, respectively. The beta sheet markers (1694 and 1625 cm-1) associated to arrestin complex formation are following the same time course (Fig. 3.2.13, B, red) and are correlated to the MetaIIb/MetaIIbH+ marker. They share the same kinetic and they also plateaued at ~50% of the initial light-activated level. In comparison, in the case of absence of arrestin, the beta sheet markers are insignificant (Fig. 3.2.13, B, blue).

Figure 3.2.14: Doubles difference spectra of early and late spectra. Double difference spectra (without arrestin – whith arrestin) calculated directly after illumination (green, 5-10s) and at the end of the decay process (orange, 1200-1300s). 1768, 1748, 1661, 1664 cm-1 bands represent the Meta II conformation, 1348 cm-1 band the Meta III formation in absence of arrestin.

From this time dependent data set, two different spectra were extracted. The first spectrum was extracted directly after illumination (5-10 s) and the second spectrum after decay when the stationary plateau war reached (1200-1300 s). This operation was effectuated for the measurements in presence and in absence of arrestin. Double difference spectra (with arrestin minus without arrestin) were then effectuated

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between the spectrum with and without arrestin, immediately after illumination (Fig. 3.2.14, 5-10 s, green) and after decay period (Fig. 3.2.14, 1200-1300 s, orange).

The double difference spectrum immediately after illumination (Fig. 3.2.14, green) shows clearly the presence of the Meta IIb/Meta IIbH+ bands (1768, 1748, 1644 cm-1) resulting from the of the Meta equilibrium shift to Meta IIbH+ due to the arrestin binding. The presence of Meta IIb/Meta IIbH+ bands (1768, 1748, 1644 cm-1) in the late double difference spectrum (Fig. 3.2.14, orange), but with a smaller amplitude compared to the early double difference spectrum (Fig. 3.2.14, green), signifies that the Meta IIbH+ is also present at the end of the decay period. It confirms that only a part (~50%) of the total complex population formed after illumination decayed and that the fraction of residual receptors, which are not decayed, are still in the Meta IIbH+ conformation. The arrestin binding and decay are also accompanied by the beta sheet markers (1694, 1625 cm-1) and the 1661cm-1 specific band for complex formation highlighted in the ABS. The amplitude of these bands is also smaller at the end of the decay. Unfortunately, these beta sheet marker specific for arrestin binding are overlapped with rhodopsin bands and it is difficult to determine the amount of complex involved in the decay process (see Fig. 3.2.14). However, the comparison of the difference spectrum with and without arrestin described above (Fig. 3.2.13) gives a reliable estimation of the fraction of arrestin and rhodopsin remained in an activated form. Half of complex decays while the other half is stabilized in Meta IIbH+ after decay. The analysis of this data via singular value decomposition (SVD) and global analysis shows two components. The first component occuring directly after illumination is called activation spectrum. It reflects the formation of the complex (Fig. 3.2.15, red) and is nearly identical to the difference spectrum of arrestin binding. At 30°C, the two binding kinetics (depicted in figure 3.2.6 for 0°C) are too fast to be distinguished. The subsequent component, called deactivation spectrum, occurs after the formation of arrestin complex and represents the decay of the complex (Fig. 3.2.15, purple). The two components are similar but are a mirror images of one another. The deactivation spectrum has however a smaller amplitude compared to the activation spectrum. The relative amplitude of its peaks is approximately 50% of those of the activation spectrum. Together with the kinetics

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traces, these data show that only the half of the arrestin complex that formed after illumination remains in a stabilized Meta IIbH+ form, while the other half decays. No changes are observable at the region around 1300 cm-1 where the phosphate group vibrational bands are located. The absence of changes argues that the arrestin stays bound to the phosphorylated receptor tail during the decay, like during the transition from the pre-complex to high affinity complex.

Figure 3.2.15: Activation and deactivation spectrum. Two components were determined with singular value decomposition and global analysis. The activation spectrum (red) reflects the complex formation upon light activation. The deactivation spectrum (purple) reflects the changes occurring in the activation spectrum during the decay process.

3.2.9 To which products decays the complex?

In absence of arrestin, decay of activated rhodopsin is reflected in a band at 1348 cm-1 (Fig. 3.2.16, blue and Fig. 3.2.14, orange). This band is associated to the Meta III intermediate and represents the syn/anti isomerization of the retinal (Bartl, Ritter, and Hofmann 2001; Eglof Ritter, Elgeti, and Bartl 2008; Eglof Ritter et al. 2007; Reiner Vogel et al. 2003). Meta III is an inactivated decay product of Meta II (Martin Heck et al. 2003) and consists of a protonated Schiff base with

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all-trans 15-syn retinal. Accordingly, the kinetic trace of 1348 cm-1 band represents the formation of Meta III during decay. In absence of arrestin, Meta III appears within t1/2=20 min (Fig. 3.2.16, blue). In presence of arrestin (red), no Meta III is formed. These FTIR data confirm thus that arrestin prevents the formation of Meta III (M E Sommer and Farrens 2006).

Figure 3.2.16: Formation of Meta III upon decay. Time traces of 1348 cm-1 band, Meta III marker, for phosphorylated rhodopsin in presence (red) and absence (blue) of arrestin. Measured at 30°C, pH: 8.0.

The published measurements of arrestin complex decay could not clearly identify the conformation of the aporeceptor at the end of the decay (Sommer, Hofmann, and Heck 2012). To determine which of the ops or ops* conformation decayed the complex, decay of arrestin complex was measured in presence of hydroxylamine. This molecule is small enough to enter in the retinal pocket of activated rhodopsin. Hydroxylamine is a strong nucleophile and cleaves the retinal Schiff base (Sakmar, Franke, and Khorana 1991)(Piechnick, Heck, and Sommer 2011). It converts retinal to retinal oxime and thus converts Meta II to the aporeceptor opsin. Samples consisting of a mixture of phosphorylated rhodopsin and 100 mM Hydroxylamine in the presence and the absence of arrestin were measured at pH: 8.0 and 30°C.

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Figure 3.2.17: Stabilization of Meta II in presence of hydroxylamine. A. Fast component of the difference spectrum of arrestin binding in absence of hydroxylamine (red) and in presence of hydroxylamine (purple). Phosphorylated rhodopsin decayed to opsin in presence of hydroxylamine (blue). The intense 1348 cm-1 band could represent an intermediate state in the hydroxylamine- induced cleavage of the retinal Schiff base (Reiner Vogel et al. 2003).The measurements were effectuated at 0°C, pH: 8.0. B. Decay of arrestin complex in presence of hydroxylamine from A. It represents the transition of the residual arrestin stabilized Meta II decaying to opsin.

After illumination and activation of rhodopsin, the hydroxylamine can enter the retinal pocket and react with the retinal Schiff base resulting of release of retinal and formation of opsin. In absence of arrestin, hydroxylamine reacts quickly after light activation of rhodopsin and cleaves the retinal from the receptor. SVD and global fit of difference spectrum (light minus dark) of phosphorylated rhodopsin in presence of hydroxylamine presents only one component. Both, the formation of Meta II and

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subsequent reaction with hydroxylamine was so fast that it could not be resolved with the available time resolution (<<1s). Consequently, only the global transition of dark state rhodopsin to a stable state of inactive opsin and retinal oxime could be extracted (Fig. 3.2.17, A, blue). In the case of phosphorylated rhodopsin with arrestin and hydroxylamine, the SVD and global fit of the difference spectrum (light minus dark) present two components.

The first one is fast and includes the formation of the arrestin complex (t1/2 < 1.5 s, Fig. 3.2.17, A, purple). The formation of Meta II and of the complex is clearly detectable by the Meta II (1768, 1745, 1713 and 1664 cm-1) and arrestin binding markers (1661, 1694 and 1623 cm-1). However, the intensity of the spectrum was approximately half of the amplitude reached in absence of hydroxilamine (red), suggesting that the other half of receptor is also converted to opsin and retinal oxime as in absence of arrestin. The second component is slower and represents the decay of the complex to opsin, through the reaction with hydroxylamine, of the residual arrestin rhodopsin complex population (t1/2= ~20 s, Fig. 3.2.17, B, purple). It represents the transition of the half residual receptor population in high affinity complex decaying to opsin. This component is basically the same as the ´´deactivation spectrum´´ (Fig. 3.2.15, purple). Only small differences due to different samples and measuring conditions are observable. The addition of the two components, spectra of formation and decay of the complex, leads to the same spectrum as the spectrum of the transition of rhodopsin to opsin (Fig. 3.2.17, A, blue). The experiments with hydroxylamine show that arrestin can not prevent the reaction of hydroxylamine with the Schiff base of Meta II-arrestin complex but slow down the process. It supports the biphasic behaviour of arrestin binding previously measured (Fig. 3.2.6). Immediately after illumination, half of the arrestin population binds fast enough (< 1 s) to slow down retinal Schiff base cleavage by hydroxilamine while the other half of unbound rhodopsin is readily cleaved by hydroxilamine and can not form the complex. The high concentration of hydroxilamine (100 mM) is sufficient to quickly hydrolyse all Schiff bases and thus should not be, at this concentration, a limiting factor (Piechnick, Heck, and Sommer 2011)(K P Hofmann, Emeis, and Schnetkamp 1983).

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3.3 Rhodopsin diffusion in native disk membrane

The organisation and the behaviour of the receptors within the disk membrane are important to understand the binding mechanism of protein to the receptor. Different studies, based on atomic force microscopy and electron microscopy, have shown that rhodopsin is present in dimers and forms racks of dimers in disk membrane (Dimitrios Fotiadis et al. 2003)(Buzhynskyy, Salesse, and Scheuring 2011). They however differ with respect to the distribution and the homogeneity of the receptors. Dimers or racks of dimers densely packed should have a low diffusion coefficient. Nevertheless, diffusion of bovine rhodopsin was already measured and yields a high diffusion coefficient (0,38 μm2/s) (Takezoe and Yu 1981). The biphasic behaviour of arrestin binding and the asymmetric decay suggest the presence of rhodopsin dimers and a contribution of these dimers to the formation and the decay of arrstin- receptor complex (see discussion). The aim of this part of the project is to measure and determine the diffusion of the receptor and to determine the influence of arrestin on it. This work was effectuated in collaboration with Professor Horst Vogel of the Swiss Federal Institute of Technology of Lausanne (EPFL). Disc membranes were extracted from ROS, labelled and washed in Berlin and finally measured in Lausanne on wide field microscope. Extraction of the traces from the videos and its analysis were effectuated by Horst Vogel co-worker Dr. Piguet.

3.3.1 Samples quality control

The diameter of the disk membranes has an average of ~500-800 nm which is at the limit of the optic resolution of the microscope (~500 nm). Thus, it is impossible to observe the disks and to distinguish them from cell debris (plasma membrane debris, aggregates, etc…) under the microscope with normal light (transmission). To determine the quality of the disk preparation, negative stain picture of the preparations were recorded with an electronic microscope.

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Figure 3.4.1: Negative stained Electron micrograph of discs membrane. A. Overview of the disk preparation shows a distribution of disk size and small amount of cell debris component and burst disks. Scale bar: 5 μm. B. Size distribution of intact disks. Disks look like deflated footballs. Scale bar: 1 μm.

The disks have a size distribution from 200 nm to 1 μm. They are flat and look like deflated footballs (Fig. 3.4.1). Destroyed disks and cell debris like plasma membrane and aggregates are also detected on the electron micrograph. Extraction protocol was optimized to minimize the amount of cell debris into the sample. The disks are released from ROS by lysing the plasma membrane. This operation is effectuated through the freezing of the ROS and through shear stress of a douncer. Electron micrographs showed that fresh samples which were not frozen have less cell debris and had the most intact and clean disks, so that no freezing step was used to burst the plasma membrane and the disks were keep fresh at 4°C. Only preparations with minimal cell debris were used for single molecule fluorescence.

Figure 3.4.2: Microscope picture of labelled disks. A. Atto 647 N maleimide labelled rhodopsin in disks illuminated with 633 nm diode laser. Fluorescence emitted by the dyes reveals a donut shape of the disks and a distribution of disk size. Intense spots are probably aggregates. Scale bar: 10 μm. B. Magnification of A shows clearly the donut shape of the disks. Scale bar: 2 μm.

The labelling of small part of rhodopsin with Atto 647 N Maleimide (Atto 647) permits to detect the disks. Many rhodopsins per disk are labelled so that the disk is 94 3. Results

identifiable. Illumination with a 633 nm laser reveals the donut shape of the disks (Fig. 3.4.2). High density of light suggests the presence of aggregates. Note that the full labelling of rhodopsin would produce much fluorescence und thus only an intensive spot would appear.

3.3.2 Rhodopsin tracking with single fluorescence microscopy

Single-molecule receptor tracking was performed with Atto 647 photostable organic dye. Illumination with a 633 nm diode laser was used to excite the dye. The rhodopsin activation is minimized at this wavelength. The videos were recorded with an electron-multiplying charge-couple device (EM-CCD) camera at 10 Hz (99ms acquisition time) during 60 sec. Disks containing single labelled rhodopsin in 100 mM HEPES buffer pH: 7.0 were measured before and after 30 s illumination with 532 nm laser at room temperature. Analysis of the video and the calculation of the mean square displacement (MSD, see methods) of rhodopsin traces permit to determine the different modes of diffusion of rhodopsin in dark and after illumination (Fig. 3.4.4). It also gives the mobility and the domain size in which the receptor is diffusing.

Figure 3.4.4: Examples of mean square displacement plot of rhodopsin traces. Mean square displacements were calculated base on the molecule traces extracted from the videos. Three types of diffusion are detected. Immobile (dark blue), confined in a small domain (red) and mobile confined in a larger domain, representing the disk size (blue).

Three modes of motion were detected for the dark rhodopsin (Fig. 3.4.5, C). 62% of molecules are restricted in domains less than 30 nm (limit of the spatial resolution) and thus immobile. 21% of the molecules are restricted in domains smaller than ~150 nm and finally 17% of the rhodopsins are mobile and free to diffuse. They are

95 3. Results

only restricted by the size of the disk. Calculation of the mean square displacement yields a maximum value of 0,005 μm2/s for rhodopsin mobility. No difference between the mobile and the confined mode were detected. With activated rhodopsin, a similar diffusion to dark rhodopsin, <0,005 μm2/s, was observed but presents a change in the mobility modes. 78% of molecules are immobile while 14% are confined and 8% mobile. The immobile rhodopsin molecules stay immobile upon activation but the part of the mobile molecule becomes confined.

3.3.3 Phosphorylated rhodopsin tracking

The same experiments were effectuated with phosphorylated rhodopsin in presence and absence of arrestin. In absence of arrestin, in dark state 76% of phosphorylated rhodopsin molecules are immobile while 24% are mobile. After illumination part of these mobile molecules are confined (Fig. 3.4.5, A). The presence of two fold excess arrestin does not modify the distribution of the rhodopsin at the dark state. After illumination, the presence of arrestin, through its binding, largely confined the mobile phosphorylated rhodopsin molecules (Fig. 3.4.5, B). 73% of the molecules become immobile while 10% remains mobile and 27% mobile. Mean square displacement of dark and activated phosphorylated rhodopsin yields a maximum value of 0,005 μm2/s for receptor mobility. Most of the rhodopsin or phosphorylated rhodopsin molecules are immobile. Approximately 73 % of them are already immobile in dark state. After illumination most of mobile rhodopsin became immobile or confined. In presence of arrestin and after illumination, no mobile rhodopsins were detected. The binding of arrestin immobilizes almost all rhodopsin. Only 10% of the receptor are confined. The second important point concerns the confined molecules. The limit of detection of the Atto 647 could not distinguish if these confined molecules are really mobile molecules in small domains or if they are immobile molecules and the small confinement detected results of the low resolution of the system. The camera and photostability of Atto 647 do not allow the record of long videos with higher acquisition time. Although Atto 647 is a photostable dye, it stability does not allow the record of long traces. Its temporal definition is low. Thus, in the condition of the measurements, the dye does not emit light enough quickly to measure with higher

96 3. Results

acquisition time and bleaches after few seconds. The (EM-CDD) camera used for these experiments does not allow measurement with higher acquisition time. Another astonish point is that all these mobile or confined rhodopsins present the same mobility of < 0,005 μm2/s. In comparison with previous studies on rhodopsin mobility (Poo and Cone 1974)(Govardovskii et al. 2009), the mobility measured is hundred fold slower. One presumption is the way to prepare the samples and the acquisition time of the measurements. Finally, it is suspected that the light used for exciting the dye may be strong enough to activate the receptor. The 640 nm laser used is far away from the maximal absorption of rhodopsin but the high intensity of light (~0,5 kWatt/cm2) may be sufficient to activate the receptor.

Figure 3.4.5: Distribution of rhodopsin diffusion modes. A. phosphorylated rhodopsin B. phosphorylated rhodopsin in presence of arrestin C. unphosphorylated rhodopsin. Immobile fraction of receptors are coloured in dark blue, confined in small domains (<~150 nm) in red and the mobile fraction (but confined by the size of the disks) is coloured in light blue.

97 3. Results

3.3.4 Tracking of quantum-labelled Opsin

To improve the accuracy of the measurement, longer traces are needed with a higher acquisition time. Atto 647 is not suitable for higher acquisition time measurements. It emits intermittent light. When a single Atto 647 is in a excited state, it can not absorb a new photon before relaxation. Therefore, de dye blinks. The emission of a single fluorophore is not continuous; there is a minimum delay between emission of subsequent photons of a few nanoseconds. This delay for Atto 647 is too long and does not allow faster measurements. Opsin molecules were labelled (see methods) with the quantum dot 655 nm (QD 655). The QD 655 has bright absorption spectra and emits light at 655 nm. It has a higher temporal definition and a better spatial resolution. For this experiment the QD 655 was excited with a 532 nm diode laser. At this wavelength, the rhodopsin is activated; so that the measurements were proceed with the apoprotein opsin. The experiments were effectuated at room temperature in 100 mM HEPES buffer pH: 7.0 and recorded with Neo Andor quick camera at 30 Hz. Quantum dot allows the tracking of opsin over long time without bleaching (Fig. 3.4.6).

Figure 3.4.6: Example of mobile trace of opsin labelled with QD 655. A. Mean square displacement of a opsin labelled with QD 655. The receptor presents a mobility (D) of 0,0048 μm2/s and a confinement (L) of 260 nm. B. Trajectory of the mobile tracked receptor.

Tracking of opsin labelled with QD 655 presents similar results as the experiment on rhodopsin and phosphorylated rhodopsin with Atto 647. More than 80% of all measured opsin are immobile while 17% of the opsins are mobile with a mobility of <0,005 μm2/s (Fig.3.4.7). The quantum dot, coupled to a fast camera, allows a higher lateral resolution and thus specifies the diffusion of the single molecule.

98 3. Results

Figure 3.4.7: Fraction of opsins in the two type of diffusion. Tracking of opsin labelled with quantum dots presents a large amount of immobile receptors. 17% of opsin in disc membrane are mobile and 83 % are immobile.

Surprisingly all tracking experiment present a highly amount of immobile receptor. Approximately 70 to 80% all receptors are not diffusing at all. The absence of confined molecules in the measurements with QD suggests that the confined molecules measured with the standard dye were artefacts. The low lateral resolution of Atto 647 and the background noise of the detection system could not determine if the detected mobile molecules in small confinement were immobile or indeed mobile in a confinement. The measurements with QD suggest that the confined molecule are effectively mobile molecule in a small confinement. In all recorded mobile traces recorded with Atto 647 or with QD 655, no significant changes in diffusion coefficient were observed. They all present a diffusion coefficient under 0,005 μm2/s. Fast measurements with QD confirmed the diffusion measured with the Atto 647.

99 3. Results

100 4. Discussion

4. Discussion

4.1 The finger loop derived peptides

4.1.1 Finger loop as Meta II sensor

All arrestin finger loop - derived peptides which demonstrated binding to activated rhodopsin, bound with low affinity (Fig. 3.1.3 and Fig. 3.1.4-6). The arrestin peptide with the highest affinity was the thirteen amino acid long peptide ArrFL-2 (63-75)

(YGREDLDVLGLTF) with a Kd of 0,35 mM. In comparison, the native Gt alpha C-terminus (Gt alpha CT) peptides have a higher affinity of 0,19 mM, while the high affinity Gt alpha (Gt alpha HAA) peptide analogue, derived from the native Gt alpha, has a Kd of 1,76 μM. Note that in case of Gt alpha peptide, the exchange of only one residue drastically increases the affinity. The stabilization of Meta II at 0°C, pH 8.0 demonstrates the specific binding of these peptides to the activate conformation of rhodopsin. Like full length arrestin, Gt alpha and Gt alpha C-terminal peptides, the arrestin finger loop peptides functionally bind the active state of the receptor and have a common receptor-binding site (see below). This implies that the finger loop is at least one of the Meta II sensors of arrestin. The importance of the finger loop for the formation of the arrestin high affinity complex has already been reported (Ostermaier et al. 2014)(Shukla et al. 2014). Comparison of the affinity between the different peptides provides some information about the nature of binding and which amino acids are required. The finger loops of arrestin-1, arrestin-2 and arrestin-3 demonstrate high homology; only three functionally relevant amino acid exchanges are observed: the Gln/Arg exchange at position 69 of arrestin-1, Met/Leu at position 75 and Ser/Thr at position 78 (Fig. 3.1.2, orange highlighted). Peptides ArrFL-1 (67-77) and ArrFL-2/3 contain the residue exchanges at positions 69 and 75. ArrFL-1 binds with higher affinity than ArrFL-2/3. The exchange of methionine to leucine does not confer a considerable difference in binding, as these two amino acids are similar with respect to size and polarity. A change which confers a greater effect occurs at position 69 between glutamine and arginine. The presence of a glutamine residue therefore may increase the affinity of the peptide to activated receptor.

101 4. Discussion

The finger loops of the two non visual arrestins are nearly identical, except for the threonine residue at position 74 of arrestin-2, which is mutated to serine in arrestin-3 (at position 75). Comparison between ArrFL-2 (64-75) and ArrFL-3 (65-76), shows that this residue exchange is the only difference in sequence, located at the penultimate position of the peptide sequences. The presence of threonine instead of serine increases the affinity of the peptide to the activated receptor (Fig. 3.1.5, D).

Of the finger loop peptide series used in this study, six peptides do not bind. First, the arrestin-4 peptide, ArrFL-4 (59-69), probably does not bind due to the histidine in position 61. It is the only relevant amino acid exchange, when compared to the ArrFL-2/3 peptide. The presence of this bulky residue may disturb the binding. Arrestin-4 is known to bind with high affinity to cone rhodopsin. Its affinity to other GPCRs is low and weak to rhodopsin (Sutton et al. 2005). This residue may lead to the formation of a different secondary structure and thus possibly be the key which allows arrestin-4 to discriminate the binding to activated rod rhodopsin.

Figure 4.1.1: Structure of finger loop of the arrestin-2 in its pre-active state. The finger loop is composed of beta sheet strands at its base (PDB:4JQI). The binding of the phosphorylated receptor tail (orange) forms and stabilises the beta sheet with residues Ser74 Phe75 with Tyr63 and eventually with Gly64 (blue). The residues Gly72 and Leu71 (red) interact directly with the receptor. Residues Leu73 and Asp69 (green) form an intramolecular interaction in the high affinity complex (Szczepek et al. 2014). Insert: Arr-2 structure with finger loop highlighted.

102 4. Discussion

The ArrFL-1 (69-79), ArrFL-2 (65-75) and ArrFL-3 (66-76) peptides are homologues and do not stabilise Meta II. They are derived from the first three arrestin types and have only three different amino acids (Fig. 3.1.2, orange highlighted). They all share the absence of the amino acids tyrosine and glycine at their N-terminus (Fig. 3.1.2, red highlighted). The correlation between the absence of these two residues and the abrogation of binding, suggests that these residues are required for the binding of the finger loop peptides. The crystal structure of pre-activated arrestin, indicates that, in the peptide, these two N-terminal residues could form a beta sheet contact with the residues Ser74 (or Thr in Arr-1) and Phe75 on the C-terminal side of the peptide (Fig. 4.1.1). The partial Meta II stabilisation even with excess of ArrFL-2 (65-75) peptide suggest destabilization of the Meta I/Meta II equilibrium. Further investigation into the possibility of peptide binding to Meta I was not carried out. The intramolecular beta sheet interaction at the base of the finger loop may be essential to stabilize the secondary structure of the finger loop, facilitating the binding to the receptor crevice. This may be the reason why the long version ArrFL-2 (63-75) displays the highest affinity of the whole arrestin peptide series. This notion suggests that the peptide ArrFL-1 (67-79) may also bind with similar affinity. With the exception of ArrFL-1 (67-77), none of the ArrFL-1 peptide binds Meta II.1 However, the absence of the two C-terminal amino acids (Thr/Ser74 and Phe75 in Arr-2) does not block the binding of the peptide arguing, that these are not required. In contrast, the two N-terminal residues (Tyr63 and Gly64 in Arr-2) possibly develop some interactions with the receptor (see above).

4.1.2 Structure of bound arrestin finger loop and its interaction with activated rhodopsin

The ArrFL-1 was co-crystallized with the active receptor by collaborators (Szczepek et al. 2014b). Unfortunately the structure revealed only the seven residues 69-DIDVMGL-77. The electronic density of the other residues was not sufficient to

1 Many batches of these peptides were ordered from different companies, but there are difficulties with synthesising and correctly purifying these peptides. The presence of a glutamine residue at the N-terminus of the peptide can cause spontaneous cyclization to form a pyroglutamic acid. The instability of these peptides could explain why they do not bind. 103 4. Discussion

be interpretable. The structure presents the arrestin finger loop peptide within the activated receptor crevice (Fig. 4.1.2). The activated receptor in the Meta II form has the same conformation as Ops*(Park et al. 2008). ArrFL-1 forms a helical structure with a reverse turn-like structure (or C-cap-like). Strong internal hydrogen bonding from Asp73 and Leu77 and the flexibility of Gly76 facilitate the formation of its reverse turn-like structure. The peptide interacts principally with two highly conserved constraints of the receptor. It forms two hydrogen bonds: one between Met75 and Arg135 of the E(D)RY ionic-lock motif and the second between Gly76 and Lys311 of NPxxY(x)5,6F motif. C-terminal extension of the finger loop would be located between TM6 and TM7/H8. The N-terminus would extend in the direction of cytoplasmic loop 2 of the activated receptor.

Figure 4.1.2: ArrFL-1 structure bound to activated receptor. Cytoplasmic view of the crevice of the upside down receptor. The C-cap formation of ArrFL-1 is stabilized by the residues Asp73 and Leu77 (green) which form an intramolecular hydrogen bond and the hydrogen bonds between two residues of the peptides (red) and two residues of the receptor (orange). The amino acids Met75 and Gly76 (red) form a hydrogen bond to Arg135 of the E(D)RY motif and to Lys311 of the NPxxY(x)5,6F motif, respectively. (PDB: 4PXF).

104 4. Discussion

Superposition and comparison of the crystal structures of Ops*-bond Gt alpha HAA (340-ILENLKDCGLF-350) derived peptide (Scheerer et al. 2008) with ArrFL-1 show a similar shape of both peptides (Fig. 4.1.3, blue and purple respectively). Arr-1 shows a similar C-capping motif as previously observed for Gt alpha peptide. In both cases, the conserved residues glycine and leucine at their C-termini allows the formation of the reverse turn-like (or C-cap, Fig. 4.1.3, sticks). However, Gt alpha CT forms an alpha helix which is not the case with ArrFL-1 peptide. Both peptides interact in a similar manner with the receptor and stabilize Ops* (black and orange). Residues Lys345 and Cys347 (yellow) of Gt alpha CT peptide interact with the Arg135 of the E(D)RY motif and Gln312 of the NPxxY(x)5,6F motif. However, the Gt peptide has a contact to TM5 and TM6 which is too far in the case of ArrFL-1.

Figure 4.1.3: Superposition of ArrFL-1 and Gt alpha HAA peptides. Top view of crystal structure of opsin with ArrFL-1 (black, PDB: 4PXF) and with Gt alpha HAA (orange, PDB:3DQB) are nearly identical. The structures of ArrFL-1 (blue) and Gt alpha HAA (purple) are also similar. In both cases, the residues Gly and Leu (GL motif) allow the formation of the C-cap (Gly76 and Leu77 for ArrFL-1 and Gly348 and Leu349 for Gt alpha CT, sticks). Residues Gly76 and Met75 (red) of ArrFL-1 interact with the receptor while for Gt alpha CT, Cys347 and Lys345 (yellow) interact with the receptor. The intramolecular hydrogen bond between Asp73 and Leu77 (green) stabilizes the reverse turn of ArrFL-1. In the case of Gt alpha CT, more intramolecular hydrogen bonds are present and form the alpha helix.

105 4. Discussion

Altogether the functional Extra Meta II assay and the similar crystal structures of the ArrFL-1 and Gt alpha CT peptides show that the common (E/D)x(I/L)xxxGL motif of arrestin and Gt alpha protein can recognize the active state of rhodopsin within the E(D)RY and NPxxY(x)5,6F motifs. The four arrestins bind rhodopsin with different affinities (Gurevich et al. 1995)(Sutton et al. 2005). Therefore, the (E/D)x(I/L)xxxGL motif confers the finger loop the functional Meta II sensor of arrestin but is not the key for the specificity of the arrestin types to different GPCRs. The functional Extra Meta II assay shows that the presences of the residues Tyr67, Gly68, Ser78 and Phe79 at the base of the ArrFL-1, which form beta sheet contacts in the pre-active conformation, increase the affinity of the peptide for Meta II. This is more intriguing due to the distance between these residues and the main area of interaction with the receptor (E(D)RY and NPxxY(x)5,6F). It could suggest that these residues build additional interactions with the receptor and thus increase the affinity of the peptide to the receptor. However, such interactions are not identified in the crystal structure. Another explanation is that (E/D)x(I/L)xxxGL motif requires the stabilization of the secondary structure of the finger loop to bind into the crevice. The structure of the reverse turn presents only one intramolecular hydrogen bond which might be insufficient for the finger loop to form the constraint alone. Therefore, the finger loop might need additional stabilization and orientation through the adjacent beta sheet at its base. The finger loop in basal arrestin is highly flexible (Fig. 4.1.4, A). Comparison of crystal structures of basal and pre-complexed arrestin suggests that the structural flexibility of the finger loop is apparently essential for the ability of arrestin to adopt a new conformation that is capable of interacting with the receptor to form the high-affinity complex (Fig. 4.1.4). The phosphorylated receptor C-terminal analogue peptide forms an anti-parallel beta sheet adjacent to the finger loop which extends the beta strands present in the basal state of arrestin (Fig. 4.1.4. B). Two phosphate moieties of the phosphorylated peptide build hydrogen bonds and extend the beta sheet at the base of the finger loop. Elongation of the finger loop, principally stabilized by the phosphorylated receptor tail - derived peptide, is observed in the pre-active structure. The new position and structure of the finger loop suggests that this conformation initiates or facilitates the insertion of the finger loop into the receptor and formation of the C-cap. The crystal structure of the peptide-receptor complex shows that the finger loop is inserted deep into the crevice

106 4. Discussion

which, suggests that the finger loop is more elongated upon binding to the receptor. Thus the interaction should be accompanied with a melting of the beta sheet at the base of the finger loop (Szczepek et al. 2014). The phosphorylated receptor tail involved in this region might also reorganize slightly upon high affinity complex formation.

Figure 4.1.4: Arrestin finger loop conformational change upon activation and binding. A. Crystal structure of arrestin-1 in its basal state shows a labile and highly flexible finger loop present in two conformers: open (dark-green) and closed (olive) conformation. B. Pre-active structure of arrestin-2 finger loop (blue) bound to phosphorylated receptor C-terminal peptide (red). An anti- parallel beta sheet is formed with the peptide which stabilizes the loop. C. Opsin (orange) co- crystallized with finger loop peptide ArrFL-1 (purple) shows ArrFL-1 within the crevice of activate receptor. Taken from(Szczepek et al. 2014).

4.2 Formation of high affinity arrestin complex investigated with FTIR spectroscopy

4.2.1 At high concentration, arrestin is already pre-complexed in dark state

At high concentrations, arrestin self-associates and is present in an inactive form as dimers and tetramers. These arrestin oligomers dissociate in the presence of phosphorylated rhodopsin to bind the receptor and form the pre-complex. Only the monomeric form of arrestin binds the receptor through electrostatic interactions to the phosphorylated tail of rhodopsin (M. Kim et al. 2011)(Hanson, Van Eps, et al.

2007). The affinity of arrestin to phosphorylated rhodopsin (Kd of ~80 μM) is comparable to the arrestin dimerization affinity; therefore the presence of phosphorylated rhodopsin affects the arrestin monomer-dimer equilibrium (Zhuang et al. 2013). The samples used for FTIR measurements contained a mixture of excess arrestin with highly concentrated phosphorylated rhodopsin at 300-500 μM.

107 4. Discussion

At these high concentrations, due to the high level of rhodopsin phosphorylation and the high affinity of arrestin to phosphorylated receptor, almost all receptors are likely to be pre-bound with arrestin in the dark state. Hence, the FTIR measurements reflect the transition of pre-bound arrestin to the high affinity complex triggered by light (Fig. 4.2.1).

R  Arr h R*  Arr  R*  Arr  p preactive p preactive p active dark precomplex light precomplex highaffinity complex

Figure 4.2.1: Formation of high affinity complex from dark pre-complex. After illumination the arrestin is in equilibrium between pre-active in the light - pre-complex and the high affinity complex. This equilibrium is largely shifted to the high affinity complex. Light absorption activates rhodopsin in the pre-complex and thus forms the light pre-complex. Dark and light pre-complexes are possibly similar. Only the receptor probably undergoes a conformational change, hence the name the pre-complexes.

The data are consistent with the notion that, under the conditions in vitro, all rhodopsins are already pre-complexed to arrestin in the dark state. In all FTIR difference spectra, no bands reflecting phosphate vibrations were observed. Phosphate groups normally induce strong vibration patterns around 1200 - 1000 cm-1 (Klähn et al. 2004)(Allin et al. 2001). The absence of these bands argues that no light induced binding of free arrestin to activated phosphorylated rhodopsin occurs during the experiment. The absence of phosphate vibration bands during the stabilization of Meta II by arrestin binding indicates that all receptors were already in a pre-complexed arrangement in the dark. It also indicates a very limited role of phosphate groups in the transition from pre-complex to high-affinity complex. The pre-activated arrestin splice variant p44 shows a binding spectrum with phosphorylated rhodopsin and unphosphorylated rhodopsin that is almost identical to the ABS (Fig. 3.2.9) and also exhibits the same biphasic binding kinetics (Fig. 3.2.10). p44 binds phosphorylated dark state rhodopsin with very high affinity and is thus completely bound to phosphorylated dark rhodopsin even at low micromolar concentrations (Schröder, Pulvermüller, and Hofmann 2002). The similar spectrum further supports the hypothesis that arrestin was pre-complexed to phosphorylated rhodopsin before illumination.

108 4. Discussion

4.2.2 Assignment of the arrestin binding spectrum (ABS)

The interpretation of the arrestin binding spectrum is challenging. It is almost impossible to assign the bands to specific amino acids without measurement of mutants and site directed isotopically labelled samples. However, the changes of secondary structure occurring upon arrestin binding can be detected in the amide I region (Arrondo et al. 1993)(Barth 2007). This is the case for the two correlated bands at 1694 and 1625 cm-1 and for the positive band at 1660 cm-1 (Fig. 3.2.4, red). The ABS in deuterium helps to determine the secondary structures originating with these bands. Deuterium exchange induces distinct spectral shifts of molecular vibrations coupled to protonation which can allow their assignment.

4.2.2.1 The 1660 cm-1 Band

-1 The large positive band at 1660 cm in the ABS (Fig. 3.2.5, red) is also present with a slight shift in the ABS with deuterium (black), and could represent the formation of helical or turn structures upon formation of high affinity complex. Due to the superposition of many different bands, its assignation with certainty is not possible. However, the following evidence shows that this band could represent the formation of helical structures or turns. This band is similar to the dominant band seen in the binding spectrum of Gt alpha CT HAA peptide (Fig. 3.2.4, green) and of the arrestin finger loop analogue peptide (ArrFL-1, Fig. 4.2.2)(Szczepek et al. 2014). FTIR study of isotopically labelled Gt alpha CT peptide shows that this molecular vibration is assigned to the formation of helix structures in rhodopsin and in the Gt alpha peptide upon binding of the peptide (R Vogel et al. 2007). The formation of helical structures was later confirmed with the crystal structure of the Gt alpha HAA peptide co-crystallized with opsin (Scheerer et al. 2008)(Fig. 4.1.3). In comparison, the crystal structure of rhodopsin co-crystallized with ArrFL-1 shows that the finger loop derived peptide forms a C-cap-like structure when it binds within the rhodopsin crevice (Szczepek et al. 2014).

109 4. Discussion

Figure 4.2.2: FTIR measurement of ArrFL-1 peptide. Difference spectrum (light minus dark, grey) ArrFL-1 measured at pH: 5.8 shows a typical MetaIIbH+ spectrum. Double difference spectrum of ArrFL-1 (red) shows a 1660 cm-1 band similar as Gt alpha CT HAA peptide and arrestin (Fig. 3.2.4). Modified from (Szczepek et al. 2014).

As for the Gt alpha CT peptide, the 1660 cm-1 band in the ABS may also results from transmembrane helix elongation in rhodopsin due to the binding of arrestin, and could also represent the formation of the C-cap of the arrestin finger loop upon the formation of the high affinity complex. The 1660 cm-1 band is more intense with the Gt alpha HAA peptide than it is for ArrFL-1 (Fig. 3.2.4). Gt alpha peptide forms an alpha helix which includes approximately 5 hydrogen bonds. In contrast, ArrFL-1 has only one intramolecular hydrogen bond. The difference of intramolecular hydrogen bonds between the Gt alpha HAA and the ArrFL-1 peptides may thus reflect the difference in intensity of the 1660 cm-1 band of their respective binding spectra (Fig. 3.2.4).

4.2.3 Arrestin loses beta sheet content upon high affinity complex formation

-1 Confirmed by the D2O experiment (Fig. 3.2.5), the two bands at 1694 cm and 1625 cm-1 are assigned to a loss of beta-sheet structure upon high affinity complex formation. This loss of beta sheet is essentially assigned to arrestin as follows. Rhodopsin contains only short beta sheets at the ´´retinal plug´´ at the extracellular side of the membrane. This retinal plug covers the hydrophobic retinal, to protect it from spontaneous hydrolysis (Janz, Fay, and Farrens 2003)(Filipek et al. 2003). The retinal plug remains identical for all rhodopsin conformations crystallized. 110 4. Discussion

Moreover, no beta sheet changes were observed in FTIR for transitions between all rhodopsin conformations known so far. Thus the beta sheets at the retinal plug should not melt during arrestin binding. In the absence of arrestin, the phosphorylated tail is unstructured, (Getmanova et al. 2004)(Werner et al. 2008)(Langen et al. 1999) so that no loss of beta sheet can occur at the rhodopsin C-terminus. Since there is no significant beta sheet structural contribution from rhodopsin alone, this alteration must occur in arrestin alone, or at the interface between rhodopsin and arrestin in the pre-complex. Almost half of the arrestin is composed of beta sheets. The crystal structure of the finger loop peptide engaged into the rhodopsin crevice reveals the conformation of the arrestin finger loop in its bound conformation (Szczepek et al. 2014) and suggests that, during the formation of the high affinity complex, the beta sheets at the base of the finger loop melt to allow the arrestin finger loop to fit into the receptor crevice (Fig. 4.1.4). Another possible contribution is the beta sheet formed in pre-complex at the interface between arrestin and the phosphorylated tail of the receptor. In the presence of arrestin in the pre-bound form, the phosphorylated C-terminus binds arrestin as an anti-parallel beta strand. The crystal structure of pre-bound arrestin-2 co-crystallized with a phosphorylated receptor C-terminus peptide, shows the C-terminus of arrestin replaced by the phosphorylated peptide (Shukla et al. 2013). The phosphorylated peptide essentially replaces the interactions formed with the C-tail of basal arrestin but also forms a new anti parallel beta sheet at the base of the finger loop and thus extends the beta sheet adjacent to the finger loop (Fig. 4.1.1). At least two residues of the phosphorylated receptor tail are involved in this beta strand. Part of the loss of beta sheet upon complex formation could result from a reorganization of the phosphorylated receptor tail. Slice variant p44 does not require the phosphorylated receptor tail to form the high affinity complex (Pulvermüller et al. 1997). The binding spectrum of p44 with unphosphorylated rhodopsin (Fig. 3.2.11, purple) presents a less intense beta sheet marker (1625 cm-1). The difference in intensity of this band could represent the contribution of the phosphorylated tail to the total amount of loss of beta sheet upon complex formation. However, no differences in the phosphate bands were detected in all arrestin binding spectra, showing that the phosphorylated tail remains bound. Only a

111 4. Discussion

small reorganization might occur upon the formation of the high affinity complex, which might modestly contribute to the loss of beta sheets. The FTIR data reveal that at most ~5% of the beta sheets are lost in the rearrangement occurring during the formation of the high affinity complex, which represents a maximum of 8 residues. This loss of beta sheet upon arrestin binding is consistent with previous studies, which already predicted a conformational change of arrestin upon binding (K Palczewski, Pulvermüller, et al. 1991)(Schleicher, Kuhn, and Peter 1989)(Gurevich and Gurevich 2004). NMR and EPR studies suggest a partial or major reorganization of arrestin upon formation of the high affinity complex (M. Kim et al. 2012)(Zhuang et al. 2013). Pulsed EPR experiments indicate that the overall shape of arrestin remains the same, and changes are localized to flexible loops in the central crest region. In contrast, NMR experiments suggest arrestin undergoes a global unstructuring and transition to a molten globule. The small contribution in the loss of beta sheet supports the EPR results, and rejects the molten globule hypothesis of the NMR-study.

4.2.4 High affinity complex is formed in a two step mechanism

The Extra Meta II experiment allows the formation of Meta IIbH+ to be followed, as well as the arrestin conformational change, and thus to monitor the formation of the complex over time (Fig. 3.2.6 and Fig. 3.2.7). Interestingly, kinetic evaluation of the spectral changes reveals a biphasic behaviour of the formation of the high affinity complex from the pre-complex. Arrestin in pre-active form changes its conformation and binds the activated receptor through a fast and a slow pathway (Fig. 3.2.6 and Fig. 3.2.7). Moreover, the experiments with hydroxylamine show that arrestin cannot prevent the reaction of hydroxylamine with the retinal Schiff base (Fig. 3.2.17), but slows down the rate of the decay of Meta II induced by hydroxylamine. They also support the biphasic behaviour of arrestin binding. Immediately after illumination, half of receptor population binds arrestin fast enough (t1/2= < 1 s, i.e. rate of Meta II formation) to slow down retinal Schiff base hydrolysis by hydroxylamine while the other half of pre-complexed rhodopsin reacts readily with hydroxylamine (see chapter 3.2.9).

112 4. Discussion

The measured kinetics with FTIR are similar to the kinetics previously measured in a fluorescence study (Kirchberg et al. 2011). Kirchberg et al. observed three distinct kinetic transitions occurring after flash activation of rhodopsin at a low concentration (1 µM rhodopsin, 10 µM arrestin) at 20°C. They assigned the first transition (t1/2= 0,3s) to the conformational changes of pre-bound arrestin to affinity complex. The second transition (t1/2= 7,6s) to the formation of the pre-complex by soluble arrestins and the third transition (t1/2= 25s) to the formation of the high affinity complex from the soluble arrestin. Although they detect three transitions, they interpret these transitions on the basis of a two individual binding reactions. Arrestin first binds the phosphorylated receptor to form the pre-complex and then undergoes a further structural rearrangement to form the high affinity complex. Their analysis implies that the amount of arrestin already pre-bound in the dark would determine the ratio between the two kinetics. Thus a sample containing only arrestin which is already pre-bound in the dark to phosphorylated rhodopsin should result in only one fast kinetic phase, which represents the transition of pre-bound arrestin to the high affinity complex. The FTIR data suggest an alternative interpretation. In the highly concentrated sample used for FTIR measurements, all arrestin is most likely already pre-bound to the receptors (see chapter 4.2.1) so that the two kinetic transitions could only refer to the transition of pre-bound arrestin to the high affinity complex. In other words, half of the pre-complexed population transitioned to the high-affinity complex quickly

(t1/2= <1s), while the other half of the population transitioned slowly (t1/2= 20s). The overall rates of the transitions measured by FTIR are slower than those of Kirchberg et al. because they were measured at lower temperature (0°C). The absence of a third component and the fact that all rhodopsins already pre-bound in the dark suggest that the third (slowest) transition measured by Kirchberg and colleagues correspond to the migration of arrestin to the membrane. Additionally, two kinetics are observable by UV-Vis spectroscopy at a lower concentration but with otherwise the same measurement conditions as for FTIR measurements (100 mM HEPES buffer at 0°C, pH: 8.0, full bleach). Extra Meta II experiments were carried out with samples containing 8 μM phosphorylated rhodopsin and 20 μM arrestin. At this concentration the pre-bound amount is lower and can be quantified through a pull down assay. The pull down assay indicates that ~54% of rhodopsins were already pre-bound in the dark (Fig. 4.2.3, insert).

113 4. Discussion

Extra Meta II assay measured with UV-Vis spectroscopy showed biphasic kinetics. Previous measurements of Extra Meta II formation in the presence of arrestin at a low overall concentration have already shown a biphasic transition to the high affinity complex (Pulvermüller et al. 1997)(Schleicher, Kuhn, and Peter 1989). Fit of the data traces to a bi-exponential expression yields that 46% of the rhodopsin react quickly (t1/2= 5,2s) and the residual 54% of rhodopsin react slower (t1/2= 57s). The slow binding transition is assigned to the soluble arrestins which form the high affinity complex, similar to the third transition of Kirchberg et al.

0,035

0,030

0,025

0,020 100 0,015 80 60 0,010 40 20 0,005 0

Percentage binding [%] binding Percentage Dark Light Abs. Diff. (380-417 nm) (380-417 Diff. Abs. 0,000

-0,005 0 50 100 150 200 250 300

Time [s] Figure 4.2.3: High affinity complex formation measured through UV-Vis Extra Meta II assay. Extra Meta II formation of samples in 100 mM HEPES at 0°C, pH: 8.0 containing 8 µM phosphorylated rhodopsin with and without 20 µM arrestin (blue and green traces respectively). Sample consisted of 8 µM phosphorylated rhodopsin with 300 µM Gt alpha HAA peptide (red trace) represent the maximal Meta II formation of 0,032 OD. Arrestin amplitude does not reach the maximal Meta II formation due probably to the formation of isorhodopsin (rhodopsin inactive conformation with 9-cis-retinal, favoured by alkaline pH). The quick binding of Gt alpha peptide avoids the formation of isorhodopsin. Samples were 100% bleached with 7 seconds orange light illumination (580 nm long pass filter, started at t= 0 s). Saturation of detector during illumination time was subtracted. Solid line through the data points represent the best-fit to a mono-exponential expression (black) for Gt alpha HAA and for a bi-exponential expression (grey) for arrestin. Insert: Pulled-down assay to determine the amount of pre-bound arrestin. Phosophorylated rhodopsin exhibits 54% of pre-bound arrestin in the dark. Illuminated sample exhibits a 100% arrestin binding. Samples similar as for the Extra Meta II experiment consisting of 8 μM phosphorylated rhodopsin with 20 μM of A366C labelled with NBD (see methods, 2.11).

No clear consensus can be found for the assignation of the transitions measured with FTIR (this work), UV-Vis (this work and (Schleicher et al., 1989)(Pulvermüller et

114 4. Discussion

al. 1997)) and fluorescence (Kirchberg et al. 2011), the FTIR data shows that arrestins are already pre-bound in dark state at high concentration (see above, chapter 4.2.1) and thus the two transitions measured under these conditions can not be assigned to the formation of high affinity complex from the soluble arrestin fraction. The double difference spectra (Fig. 3.2.6, C) of the fast and slow components are nearly identical and similar to the ABS, arguing that both components are originated from the very similar arrestin-rhodopsin pre-complex and resulted in the very similar high-affinity complex. Moreover, the spectrum of each component displays the same amplitude. In this new paradigm, the fast component occurs during the illumination time and reflects the transition of rhodopsin to Meta IIbH+ and thus of the high affinity complex. The slow component represents the formation of the high affinity complex of the residual pre-complexed rhodopsins. To support this suggestion, arrestin binding was compared with p44 binding to phosphorylated rhodopsin and unphosphorylated rhodopsin. The splice variant p44 is known to have a stronger interaction with dark phosphorylated rhodopsin than arrestin. Its interaction with unphosphorylated rhodopsin is weaker than with the phosphorylated species so p44 should present a low pre-bound rate with unphosphorylated rhodopsin (Schröder, Pulvermüller, and Hofmann 2002). This difference in affinity to dark phosphorylated receptor may increase the pre-bound rate. However, high affinity complex formation of p44 to phosphorylated rhodopsin and to unphosphorylated rhodopsin presents the same characteristics. The binding spectrum of p44 to phosphorylated rhodopsins is similar to the ABS (Fig. 3.2.9) and p44 also displays the same biphasic behaviour and similar ratio between the two components than arrestin (Fig. 3.2.10). The binding spectrum of p44 to unphosphorylated rhodopsins is similar to the ABS excepted that the intensity of the 1625 cm-1 band is weaker2 (Fig. 3.2.11). However, the kinetics and the ratio between the two transitions (Fig. 3.2.12) are similar to those of arrestin and p44 with phosphorylated rhodopsin. Thus like arrestin, half of the p44 pre-complexed population reacts quickly and the second half of the p44 pre-complexed reacts slowly.

2 The absence of phosphorylation may remove the beta sheet interaction of the receptor tail with the base of the finger loop (Fig. 4.1.1). 115 4. Discussion

4.2.5 Possible mechanisms of arrestin binding

Different explanations for the observed biphasicity of arrestin binding can be envisaged. First, space limitations on the rod disk membrane due to the native high concentration of rhodopsin could restrict arrestin binding. Rhodopsin has diameter of ~48 Ǻ and in the case of the high affinity complex, all receptors are tightly bound by one arrestin molecule with a ~95 Ǻ width (Hirsch et al. 1999). It is evident that the size of arrestin and also the close vicinity of two rhodopsins with their respectively bound arrestins induce a steric clash between the two arrestins. In this case, about half of the pre-complexed arrestins would bind quickly to form the high affinity complex, thereby limiting space on the membrane surface for the remaining pre-complexed arrestins to bind. Formation of high affinity complex of the remaining arrestins would depend on slow rearrangements of both binding partners on the membrane surface (Fig. 4.2.4, A). The affinity of arrestin to activated phosphorylated receptor is high (Kd= 2,2 nM, (M E Sommer, Smith, and Farrens 2006)) and may be strong enough to overcome the steric obstruction. A rearrangement of the system would explain why arrestin binds the receptor with two distinct kinetics, but there is no evidence to support why half of the arrestin population reacts quickly and the second half population slower. A subsequent explanation is needed to explain why the two kinetics are so clearly distributed in two ratios with the same amplitude. The double difference spectra of the fast and slow components are nearly identical (Fig. 3.2.6), arguing that both components originated from the structurally very similar arrestin-rhodopsin pre-complex and resulted in the structurally very similar high-affinity complex. A suggestion would propose the possible existences of two or more types of pre-complex within the population, which are spectroscopically indistinguishable but form the high-affinity complex at different rates. The duality of the binding mechanism suggests that half of the arrestin rhodopsin pre-complex population is discriminated from the other half and binds quickly while the second reacts slower. In this case, the binding may require a contact between both receptors and/or between both arrestins to discriminate between the two pre-complexes. It would suggest a contribution from rhodopsin dimers or arrestin dimers involved in the pre-complex units (Fig. 4.2.4, B and C).

116 4. Discussion

Atomic force microscopy studies have already shown that rhodopsins are present as dimers in murine disks (D Fotiadis et al. 2003) and the dual pathway of the binding suggests a contribution of these rhodopsin dimers to the arrestin binding mechanism. In the case where receptors are dimerized, the first pre-complexed arrestin may form the high affinity complex quickly while the other pre-complexed arrestin requires a slow dissociation of the dimer to bind and produces the one to one arrestin- rhodopsin binding (see below). Dissociation of the dimer would provide enough space to allow the second arrestin to bind with a time delay (Fig. 4.2.4, B). Two arrestins pre-bound to two neighbouring receptors or dimers could, due to the vicinity and their size, interact to form an arrestin dimer where the two arrestins are in pre-active conformation and bound to the phosphorylated tail of the receptor. The affinities of arrestin for dark phosphorylated receptor and for homodimerization are quite similar (Kd of approximately 35 and 80 μM respectively (Hanson, Van Eps, et al. 2007)(Zhuang et al. 2013)). At a concentration of 300 μM (native condition) and in the absence of phosphorylated rhodopsin, almost all arrestin is present as tetramers or dimers. In the presence of phosphorylated receptors, the oligomers dissociate and the arrestins bind the tail of the receptors (Hanson, Van Eps, et al. 2007). It is conceivable that arrestins pre-bound to receptor phosphorylated tails also retain the ability to form arrestin dimers. Crystal structures of basal arrestin show arrestin as tetramers or dimers, with different conformers (Hirsch et al. 1999). This flexibility suggests that pre-bound arrestin could also form yet other types of dimers. If this were the case, one of the arrestin dimers, due to its orientation, would react quickly with the receptor, while the second would have to dissociate and rearrange before binding, conferring a delay which would lead to a slower binding kinetic.

117 4. Discussion

Figure 4.2.4: Hypothetical arrestin binding mechanism. A. General model of arrestin binding. All arrestins (N-domain, blue, C-domain, green) are pre-bound to rhodopsin in dark state (red) interacting with phosphorylated C-terminal tail of the receptor (purple). Two different pre-complex populations are present (α and β). Illumination activates the receptors (yellow) and opens the receptor crevice. One of the pre-complex populations (α) forms the high affinity complex quickly. Arrestin introduces its finger loop within the receptor crevice and forms the high affinity complex. Due to the vicinity of receptors, arrestin obstructs the side of the neighbouring receptor, disturbing the binding of the second arrestin pre-complex population. A slow rearrangement is required to the system to allow the binding of the latter. B - C. From the general scheme A, contribution from rhodopsin or arrestin dimers is suggested. The two different pre-complex populations interact with rhodopsin or arrestin homodimers in dark state. Illumination activates the receptors and allows the fast formation of high affinity complex of one pre-complex population. The second pre-complex population requires a slow dissolution of the rhodopsin or arrestin dimer to form the high affinity complex.

118 4. Discussion

4.2.6 Stochiometry change during formation and decay of the high affinity complex

Due to the high concentration of the FTIR samples, it is not possible to analyze the stochiometry of the complex by titration. However, the FTIR data can indirectly determine the stochiometry of the arrestin complex. The arrestin difference spectrum (Fig. 3.2.3) reveals simultaneous changes in specific marker bands for arrestin (1694 cm-1 and 1625 cm-1) and for rhodopsin (all Meta II markers, notably 1661 and 1664 cm-1). These bands represent the conformational changes occurring in both proteins upon the transition from pre-complex to high affinity complex. Changes of these markers occur at the same time and are correlated (Fig. 3.2.7). A few minutes after illumination, all accessible receptors are bound by arrestin and stabilized in the Meta IIbH+ conformation (see chapter 3.2.2). The emergence of Meta IIbH+ is accompanied by changes of the arrestin marker band. Under conditions of the measurement (pH: 8.0, 0°C) the Meta I is stabilized so that the formation of Meta IIbH+ can only be the result of the binding of arrestin. To be stabilized in Meta IIbH+, each activated phosphorylated rhodopsin requires the insertion of the finger loop of an arrestin leading to a one to one stochiometry. The interaction of one arrestin for one receptor is consistent with previous studies (Hanson, Gurevich, et al. 2007)(Tsukamoto et al. 2010)(Bayburt et al. 2011)(Schleicher, Kuhn, and Peter 1989)(Sommer, Hofmann, and Heck 2011).

During decay, arrestin stabilizes half of the receptor population as Meta IIbH+, while the other half decays through hydrolysis of retinal Schiff base to inactive opsin with a rate similar to that in the absence of arrestin (Fig. 3.2.13, A). Simultaneously, FTIR bands monitoring changes in beta sheet indicate that during decay, half of the arrestin stays bound while the other half of the population recovers the initial beta sheet lost upon dissociation of high affinity complex (Fig. 3.2.13, B and Fig. 3.2.14). The population of arrestin that is released, concurs with the hydrolysis and decay of half of the Meta IIbH+ population. The hydroxylamine experiment (Fig. 3.2.17) shows that Meta IIbH+ decays to the inactive opsin. This receptor state is in a closed conformation, similar to Meta I (Vogel and Siebert 2001) and does not form the high affinity complex.

119 4. Discussion

The data suggest that the half of arrestin is released from the high affinity complex and returns to its initial basal conformation. The absence of changes in the phosphate vibration region and the lack of additional changes in the FTIR spectrum during decay suggest that the arrestin released from the high-affinity complex reverts back to its pre-complexed state and probably remains bound to the phosphorylated tail of the receptor. It has already been reported that arrestin induces trapping of half of active receptors after decay (Sommer and Farrens 2006)(Sommer, Smith, and Farrens 2006). More recently, it was shown that arrestin traps all-trans-retinal in a Schiff-base linked form in half the population of phosphorylated opsin following Meta II decay (Sommer, Hofmann, and Heck 2012). The FTIR data presented here confirm that only half of the complexes decayed and identify the receptor species that remain after decay as inactive Ops. Moreover, the data implies an asymmetry of the decay and a shift in functional arrestin-receptor stoichiometry from one to one for light-activated Meta II to one-to- two following decay. The overall stoichiometry remains as one arrestin for one receptor, but there is a change in the functional stoichiometry. Before illumination, data suggest that each dark phosphorylated rhodopsin is pre-bound with an arrestin pre-complexed in a one to one stoichiometry. Illumination induces the activation of the receptors and arrestin can form the high affinity complex through the engagement of the finger loop into the receptor crevice (Fig. 4.2.5). At this point the FTIR data clearly show that one arrestin is bound to one activated rhodopsin as a high affinity complex. After decay, each rhodopsin remains associated with its arrestin, resulting still in an overall one to one stoichiometry, but subsequently this overall stochiometry is changed. Half of the arrestin population stay bound in the high affinity complex with a receptor in a Meta II conformation, while the other half of the arrestin population of arrestin is released from its previous high affinity complex. The absence of phosphate band modification signifies that the decayed arrestin population stays pre-bound to the phosphorylated tail of opsin. Decay results in one activated arrestin for two receptors. One arrestin couples to a receptor dimer composed of one opsin and one Meta II, while an additional arrestin interacts with the phosphorylated receptor C-terminus, as in the pre-complex.

120 4. Discussion

Figure 4.2.5: Model of binding and decay of high affinity arrestin complex. Arrestin (N-domain blue, C-domain, green) is pre-complexed to rhodopsin in dark state (red) interacting with the phosphorylated C-terminal tail of the receptor (purple). Absorption of light activates the receptor in its Meta II conformation (yellow) and induces formation of the high affinity complex. Arrestin changes its conformation and introduces its finger loop into receptor crevice. A fast (k1) and slow (k2) kinetic constants are observed for the transition from pre-complex to high affinity complex. The resulting high affinity has a functional one to one stoichiometry. After decay of Meta II, half of the receptor population remains as Meta II (yellow) while the other half decays to opsin (grey). Decay of Meta II to opsin is associated with a change of arrestin binding modes and an arrestin conformational change. Arrestin retrieves its finger loop of the receptor crevice but stays bound to the phosphorylated C-terminal tail. The change of arrestin binding mode shifts the functional stoichiometry from one - to - one to one - to - two. Note that the overall stoichiometry remains as arrestin interacting with one rhodopsin.

4.2.7 Model of high affinity complex decay

Previous studies on arrestin receptor complex decay have already suggested the binding of one arrestin to a receptor dimer (M. Sommer, Hofmann, and Heck 2014)(Sommer, Hofmann, and Heck 2012)(see Introduction). All-trans-retinal at high concentrations is toxic for the rod cell. The formation of dimers and trapping of 50% of all-trans-retinal was speculated to be a physiologically relevant protection mechanism under conditions of bright light (see Introduction, chapter 1.5.4). The finger loop of arrestin engages the activated Meta II conformation and a loop of the arrestin C-domain engages on receptor and thereby shields the neighbouring receptor (this study and (Sommer, Hofmann, and Heck 2012)). The FTIR data can refine this model. First, the closed opsin conformation avoids a deep insertion of the arrestin C-domain loop within the receptor crevice, although a weak interaction with the opsin surface may be possible. Secondly, the absence of phosphophate vibrational bands argues that the decayed arrestin stays bound to the phosphorylated C-terminal tail of opsin. Moreover, the affinity of arrestin is similar to that of phosphorylated opsin but slightly higher than that of phosphorylated dark state (see

121 4. Discussion

Introduction, chapter 1.6.2), thus arrestin may stay bound to the phosphorylated opsin. The previous studies propose distinct binding modes to explain the arrestin functional stoichiometry change. However, the mechanism of the transition of the two different stoichiometries and binding modes remains unknown. The FTIR data suggest that the two kinetic constants obtain for arrestin binding and asymmetry of the decay could arise due to the same reason. Two different populations, spectroscopically identical in both the pre-complex and high affinity complex, have different behaviour. A contact between the two different populations is probably required to produce such concerted binding and decay. In the case of decay, two different populations of high affinity arrestin complexes are tightly packed in the membrane. All arrestin molecules are packed tightly together, and therefore competing for space. One population of the complex decays and loses its retinal and simultaneously loses its tight interaction with arrestin. The remaining high affinity complex population is therefore no longer restricted for space, which stabilizes arrestin binding (Fig. 4.2.6).

Figure 4.2.6: Hypothetical arrestin complex decay mechanism. Two different population of high affinity arrestin complex are formed upon light activation. Due to the size of the arrestin molecule and the high density of receptor, the arrestins are hampered. Hence one population of the complex releases its all-trans-retinal, and the arrestin withdraws its tight interaction with the receptor. The arrestin still stays bound to the phosphorylated tail of opsin. The second arrestin has more space and thus can stabilize its interaction with the receptor.

122 4. Discussion

4.3 Rhodopsin organization in disc membrane

The FTIR data, as well as other arrestin binding studies, (Sommer, Hofmann, and Heck 2012)(Sommer, Hofmann, and Heck 2014) suggest that the organization of rhodopsin in the disc membrane could play a major role in the interaction between arrestin and the receptor. The organization of rhodopsin in native membranes is a subject of some controversy (see Introduction, chapter 1.5.5). In spite of the high receptor concentration, early experiments reported rhodopsin as a mobile molecule free to diffuse in the disc membrane, which led to the conclusion that rhodopsin is monomeric (Poo and Cone 1974)(P. A. Liebman and Entine 1974). Although arrestin and Gt protein can bind to one functional monomeric rhodopsin (Ernst et al. 2007)(Hanson, Gurevich, et al. 2007), many studies show that rhodopsin, like many other receptors, has the propensity for dimerization and oligomerization (Angers, Salahpour, and Bouvier 2002)(Mansoor, Palczewski, and Farrens 2006)(Jastrzebska et al. 2011)(Lohse 2010)(Lohse, Maiellaro, and Calebiro 2014)(Kasai and Kusumi 2014). Atomic force microscopy studies reported that rhodopsin is organized in racks of rhodopsin dimers (Fotiadis et al. 2003; Liang et al. 2004) or in nanodomains (Whited and Park 2014). Recently, a study on bovine opsin dimerization in a live COS cell plasma membrane reported that opsin is organized into dimers at low concentrations. (Comar et al. 2014). The bovine opsin has a diffusion rate of 0,38 μm2/s, similar to the value previously reported for bovine rhodopsin in native disc membranes (Takezoe and Yu 1981) and for rhodopsin in frog native discs membrane (Poo and Cone 1974). These values would be consistent with monomer and small oligomer diffusion. Comar and colleagues calculated that in the case of a high concentration of receptor, similar to those found in disc membranes, 87% of the total receptor population would be in a dimeric complex, and also suggests that oligomerization of rhodopsin dimers could occur. Our single molecule fluorescence microscopy data on rhodopsin in native disc membranes show that more than 70% of receptors (rhodopsin and phosphorylated rhodopsin, Fig. 3.4.5) are immobile. The binding of arrestin to phosphorylated activated rhodopsin increases the amount of immobile receptors (Fig. 3.4.5, B)

123 4. Discussion

suggesting that formation of high affinity arrestin complexes increases the immobility of the receptor. Tracking of the ~20% of the mobile receptors, shows a diffusion rate of <0,005 μm2/s. This value is 80 times lower than the values previously measured on similar membranes (Takezoe and Yu 1981). However, tracking of opsin with quantum dot measures 83% of opsins as immobile molecule (Fig. 3.4.7). This value is near to the 87% of dimers at high concentrations, as reported by (Comar et al. 2014). These data support the model of immobile dimer racks or nanodomains, and contradict the monomeric mobile rhodopsin model. Finally the presence of arrestin to the activated phosphorylated rhodopsin increases the amount of immobile receptors. ~90% of the receptors are immobile compared to the 76% prior to the formation of high affinity complex. Thus, arrestin can influence the organization of the receptors within the membrane. The question arises as to why different researchers get different results with the similar types of samples? A partial phase transition of the lipids and a correlated segregation of the protein was previously observed for bovine disc membranes cooled to 5°C (Chabre, Cone, and Saibil 2003). The isolation of the disc membrane with isotonic buffer could also disturb the disc membrane organization. Furthermore, due to Bovine Spongiform Encephalopathy, German law obligates to freeze the bovine retinae. For all these reasons, criticism arises arguing that the formation of nanodomains or racks of dimers in disc membranes could be an artefact due to the sample preparation. However dimerization of bovine rhodopsin has also been measured in different systems, such as liposomes, detergent micelles and in different cells (Mansoor, Palczewski, and Farrens 2006)(Jastrzebska et al. 2006)(Comar et al. 2014)(Suda et al. 2004). Dimerization of different GPCRs have also been reported (see Kasai & Kusumi 2014)(see Angers et al. 2002)(see Lohse 2010). The organization of the receptor seems to play a major role to the arrestin binding mode and arrestin binding mechanism. In contrast, the arrestin may also have a contribution to the formation of dimers of rhodopsin. Most interesting question is when and how are the receptor dimers formed? What is the influence of arrestin on receptor dimerization during and after the binding of arrestin? The answer to these questions may help to understand the physiological role of receptor oligomerization such as the regulation of the toxic all-trans-retinal clearance from the rod cell.

124 References

References

Allin, C, M R Ahmadian, A Wittinghofer, and K Gerwert. 2001. “Monitoring the GAP Catalyzed H-Ras GTPase Reaction at Atomic Resolution in Real Time.” Proc Natl Acad Sci U S A 98(14): 7754–59.

Altenbach, C, A K Kusnetzow, O P Ernst, K P Hofmann, and W L Hubbell. 2008. “High-Resolution Distance Mapping in Rhodopsin Reveals the Pattern of Helix Movement due to Activation.” Proc Natl Acad Sci U S A 105(21): 7439–44.

Angers, Stephane, Ali Salahpour, and Michel Bouvier. 2002. “Dimerization: An Emerging Concept for G Protein-Coupled Receptor Ontogeny and Function.” Annual review of pharmacology and toxicology 42(1): 409–35.

Arrondo, JL, A Muga, J Castresana, and FM Goni. 1993. “Quantitative Studies of the Structure of Proteins in Solution by Fourier-Transform Infrared Spectroscopy.” Progress in biophysics and molecular biology 59(1): 23.

Arshavsky, Vadim Y. 2003. “Protein Translocation in Photoreceptor Light Adaptation: A Common Theme in Vertebrate and Invertebrate Vision.” Science’s STKE : signal transduction knowledge environment 2003(204): PE43.

Arshavsky, Vadim Y, Trevor D Lamb, and Edward N Pugh. 2002. “G Proteins and Phototransduction.” Annual review of physiology 64: 153–87.

Barth, Andreas. 2007. “Infrared Spectroscopy of Proteins.” Biochimica et Biophysica Acta 1767(9): 1073–1101.

Bartl, F J, E Ritter, and K P Hofmann. 2001. “Signaling States of Rhodopsin: Absorption of Light in Active Metarhodopsin II Generates an All-Trans-Retinal Bound Inactive State.” J Biol Chem 276(32): 30161–66.

Bayburt, Timothy H, Sergey a Vishnivetskiy, Mark a McLean, Takefumi Morizumi, Chih-Chin Huang, John J G Tesmer, Oliver P Ernst, Stephen G Sligar, and Vsevolod V Gurevich. 2011. “Monomeric Rhodopsin Is Sufficient for Normal Rhodopsin Kinase (GRK1) Phosphorylation and Arrestin-1 Binding.” The Journal of biological chemistry 286(2): 1420–28.

Burns, M E, and D A Baylor. 2001. “a ctivation , d eactivation , and a daptation in v ertebrate p hotoreceptor c ells.” Annu Rev Neurosci 24: 779–805.

Burns, Marie E, and Edward N Pugh. 2010. “Lessons from Photoreceptors: Turning off G-Protein Signaling in Living Cells.” Physiology (Bethesda, Md.) 25(2): 72–84.

Buzhynskyy, Nikolay, Christian Salesse, and Simon Scheuring. 2011. “Rhodopsin Is Spatially Heterogeneously Distributed in Rod Outer Segment Disk Membranes.” Journal of molecular recognition : JMR 24(3): 483–89.

125 References

Calvert, P D, K J Strissel, W E Schiesser, E N Pugh Jr., and V Y Arshavsky. 2006. “Light-Driven Translocation of Signaling Proteins in Vertebrate Photoreceptors.” Trends Cell Biol 16(11): 560–68.

Cerione, R a, C Staniszewski, J L Benovic, R J Lefkowitz, M G Caron, P Gierschik, R Somers, a M Spiegel, J Codina, and L Birnbaumer. 1985. “Specificity of the Functional Interactions of the Beta-Adrenergic Receptor and Rhodopsin with Guanine Nucleotide Regulatory Proteins Reconstituted in Phospholipid Vesicles.” The Journal of biological chemistry 260(3): 1493–1500.

Chabre, Marc, Richard Cone, and Helen Saibil. 2003. “Is Rhodopsin Dimeric in Native Retinal Rods?” 426(November): 30–31.

Chabre, Marc, Philippe Deterre, and Bruno Antonny. 2009. “The Apparent Cooperativity of Some GPCRs Does Not Necessarily Imply Dimerization.” Trends in pharmacological sciences 30(4): 182–87.

Chabre, Marc, and Marc Maire. 2005. “Current Topics Monomeric G-Protein- Coupled Receptor as a Functional Unit.” 44(27).

Chen, Y S, and W L Hubbell. 1978. “Reactions of the Sulfhydryl Groups of Membrane-Bound Bovine Rhodopsin.” Membr Biochem 1(1-2): 107–30.

Choe, Hui-Woog, Yong Ju Kim, Jung Hee Park, Takefumi Morizumi, Emil F Pai, Norbert Krauss, Klaus Peter Hofmann, Patrick Scheerer, and Oliver P Ernst. 2011. “Crystal Structure of Metarhodopsin II.” Nature 471(7340): 651–55.

Comar, William D, Sarah M Schubert, Beata Jastrzebska, Krzysztof Palczewski, and Adam W Smith. 2014. “Time-Resolved Fluorescence Spectroscopy Measures Clustering and Mobility of a G Protein-Coupled Receptor Opsin in Live Cell Membranes.” Journal of the American Chemical Society.

Daemen, Franz J.M. 1973. “Vertebrate Rod Outer Segment Membranes.” 300: 255– 88.

Dell’Orco, Daniele. 2013. “A Physiological Role for the Supramolecular Organization of Rhodopsin and Transducin in Rod Photoreceptors.” FEBS letters 587(13): 2060–66.

Dinculescu, A, J H McDowell, S A Amici, D R Dugger, N Richards, P A Hargrave, and W C Smith. 2002. “Insertional Mutagenesis and Immunochemical Analysis of Visual Arrestin Interaction with Rhodopsin.” J Biol Chem 277(14): 11703–8.

Elgeti, M, E Ritter, and F Bartl. 2008. “New Insights into Light-Induced Deactivation of Active Rhodopsin by SVD and Global of Time-Resolved UV/Vis- and FTIR-Data.” Zeitschrift für Physikalische Chemie in press.

Elgeti, Matthias, Roman Kazmin, Martin Heck, Takefumi Morizumi, Eglof Ritter, Patrick Scheerer, Oliver P Ernst, Friedrich Siebert, Peter Hofmann, and Franz J Bartl. 2011. “Conserved Tyr223 5 . 58 Plays Different Roles in the Activation and G-Protein Interaction of Rhodopsin.” 7159–65. 126 References

Elgeti, Matthias, Alexander S Rose, Franz J Bartl, Peter W Hildebrand, Klaus-Peter Hofmann, and Martin Heck. 2013. “Precision vs Flexibility in GPCR Signaling.” Journal of the American Chemical Society 135(33): 12305–12.

Ernst, O P, V Gramse, M Kolbe, K P Hofmann, and M Heck. 2007. “Monomeric G Protein-Coupled Receptor Rhodopsin in Solution Activates Its G Protein Transducin at the Diffusion Limit.” Proc Natl Acad Sci U S A 104(26): 10859– 64.

Ernst, Oliver P, David T Lodowski, Marcus Elstner, Peter Hegemann, Leonid S Brown, and Hideki Kandori. 2014. “Microbial and Animal Rhodopsins: Structures, Functions, and Molecular Mechanisms.” Chemical reviews 114(1): 126–63.

Feuerstein, Sophie E, Alexander Pulvermüller, Rudolf Hartmann, Joachim Granzin, Matthias Stoldt, Peter Henklein, Oliver P Ernst, Martin Heck, Dieter Willbold, and Bernd W Koenig. 2009. “Helix Formation in Arrestin Accompanies Recognition of Photoactivated Rhodopsin.” Biochemistry 48(45): 10733–42.

Filipek, Sławomir, Ronald E Stenkamp, David C Teller, and Krzysztof Palczewski. 2003. “G Protein-Coupled Receptor Rhodopsin: A Prospectus.” Annual review of physiology 65(2): 851–79.

Fotiadis, D, Y Liang, S Filipek, D A Saperstein, A Engel, and K Palczewski. 2003. “Atomic-Force Microscopy: Rhodopsin Dimers in Native Disc Membranes.” Nature 421(6919): 127–28.

Fotiadis, Dimitrios, Yan Liang, Slawomir Filipek, David A. Saperstein, Andreas Engel, and Krzysztof Palczewski. 2003. “Rhodopsin Dimers in Native Disc Membranes.” 421(January): 127–28.

Gershengorn, M C, and R Osman. 2001. “Minireview: Insights into G Protein- Coupled Receptor Function Using Molecular Models.” Endocrinology 142(1): 2–10.

Getmanova, E, A B Patel, J Klein-Seetharaman, M C Loewen, P J Reeves, N Friedman, M Sheves, S O Smith, and H G Khorana. 2004. “NMR Spectroscopy of Phosphorylated Wild-Type Rhodopsin: Mobility of the Phosphorylated C- Terminus of Rhodopsin in the Dark and upon Light Activation.” Biochemistry 43(4): 1126–33.

Gibson, S K, J H Parkes, and P A Liebman. 1999. “Phosphorylation Alters the pH- Dependent Active State Equilibrium of Rhodopsin by Modulating the Membrane Surface Potential.” Biochemistry 38(34): 11103–14.

Gibson, Scott K, John H Parkes, Paul A Liebman, and Recei V April. 1998. “Phosphorylation Stabilizes the Active Conformation of Rhodopsin.” American chemical society 37(33): 11393–98.

127 References

Govardovskii, Victor I, Darya a Korenyak, Sergei a Shukolyukov, and Lidia V Zueva. 2009. “Lateral Diffusion of Rhodopsin in Photoreceptor Membrane: A Reappraisal.” Molecular vision 15(April): 1717–29.

Granzin, J, U Wilden, H W Choe, J Labahn, B Krafft, and G Buldt. 1998. “X-Ray Crystal Structure of Arrestin from Bovine Rod Outer Segments.” Nature 391(6670): 918–21.

Granzin, Joachim, Anneliese Cousin, Moritz Weirauch, Ramona Schlesinger, Georg Büldt, and Renu Batra-Safferling. 2012. “Crystal Structure of p44, a Constitutively Active Splice Variant of Visual Arrestin.” Journal of molecular biology 416(5): 611–18.

Gupta, B Y Banshi D, and Theodore P Williamst. 1990. “lateral diffusion of visual pigments in toad (bufo marinus) rods and in catfish.” (1990): 483–96.

Gurevich, Vsevolod V, and Jeffrey L Benovics. 1993. “Visual Arrestin Interaction with Rhodopsin sequential multisite binding ensures.” 268(16): 11628–38.

Gurevich, Vsevolod V, S B Dion, J J Onorato, J Ptasienski, C M Kim, R Sterne- Marr, M M Hosey, and L Benovics. 1995. “Arrestin Interactions with G Protein-Coupled Receptors.” 720–91.

Gurevich, Vsevolod V, and Eugenia V Gurevich. 2004. “The Molecular Acrobatics of Arrestin Activation.” Trends in pharmacological sciences 25(2): 105–11.

Gurevich, Vsevolod V, Susan M Hanson, Xiufeng Song, Sergey a Vishnivetskiy, and Eugenia V Gurevich. 2011. “The Functional Cycle of Visual Arrestins in Photoreceptor Cells.” Progress in retinal and eye research.

Han, M, V V Gurevich, S A Vishnivetskiy, P B Sigler, and C Schubert. 2001. “Crystal Structure of Beta-Arrestin at 1.9 A: Possible Mechanism of Receptor Binding and Membrane Translocation.” Structure 9(9): 869–80.

Hanson, S M, N Van Eps, D J Francis, C Altenbach, S A Vishnivetskiy, V Y Arshavsky, C S Klug, W L Hubbell, and V V Gurevich. 2007. “Structure and Function of the Visual Arrestin Oligomer.” EMBO J 26(6): 1726–36.

Hanson, S M, D J Francis, S A Vishnivetskiy, E A Kolobova, W L Hubbell, C S Klug, and V V Gurevich. 2006. “Differential Interaction of Spin-Labeled Arrestin with Inactive and Active Phosphorhodopsin.” Proc Natl Acad Sci U S A 103(13): 4900–4905.

Hanson, S M, E V Gurevich, S A Vishnivetskiy, M R Ahmed, X Song, and V V Gurevich. 2007. “Each Rhodopsin Molecule Binds Its Own Arrestin.” Proc Natl Acad Sci U S A 104(9): 3125–28.

Heck, M, and K P Hofmann. 2001. “Maximal Rate and Nucleotide Dependence of Rhodopsin-Catalyzed Transducin Activation: Initial Rate Analysis Based on a Double Displacement Mechanism.” The Journal of biological chemistry 276(13): 10000–9. 128 References

Heck, M, S A Schädel, D Maretzki, F J Bartl, E Ritter, K Palczewski, and K P Hofmann. 2003. “Signaling States of Rhodopsin. Formation of the Storage Form, Metarhodopsin III, from Active Metarhodopsin II.” J Biol Chem 278(5): 3162–69.

Heck, Martin, Sandra A Schädel, Dieter Maretzki, Franz J Bartl, Eglof Ritter, Krzysztof Palczewski, and Klaus Peter Hofmann. 2003. “Signaling States of Rhodopsin. Formation of the Storage Form, Metarhodopsin III, from Active Metarhodopsin II.” The Journal of biological chemistry 278(5): 3162–69.

Herrmann, R, M Heck, P Henklein, C Kleuss, V Wray, K P Hofmann, and O P Ernst. 2006. “Rhodopsin-Transducin Coupling: Role of the Galpha C-Terminus in Nucleotide Exchange Catalysis.” Vision Res 46(27): 4582–93.

Hildebrand, P W, P Scheerer, J H Park, H W Choe, R Piechnick, O P Ernst, K P Hofmann, and M Heck. 2009. “A Ligand Channel through the G Protein Coupled Receptor Opsin.” PLoS One 4(2): e4382.

Hirsch, J A, C Schubert, V V Gurevich, and P B Sigler. 1999. “The 2.8 A Crystal Structure of Visual Arrestin: A Model for Arrestin’s Regulation.” Cell 97(2): 257–69.

Hofmann, K P. 1985. “Effect of GTP on the Rhodopsin-G-Protein Complex by Transient Formation of Extra Metarhodopsin II.” Biochim Biophys Acta 810(2): 278–81.

Hofmann, K P, D Emeis, and P P Schnetkamp. 1983. “Interplay between Hydroxylamine, Metarhodopsin II and GTP-Binding Protein in Bovine Photoreceptor Membranes.” Biochim Biophys Acta 725(1): 60–70.

Hofmann, K P, P Scheerer, P W Hildebrand, H W Choe, J H Park, M Heck, and O P Ernst. 2009. “A G Protein-Coupled Receptor at Work: The Rhodopsin Model.” Trends Biochem Sci 34(11): 540–52.

Hofmann, K P, C M Spahn, R Heinrich, and U Heinemann. 2006. “Building Functional Modules from Molecular Interactions.” Trends Biochem Sci 31(9): 497–508.

Hofmann, Klaus Peter, Alexander Pulvermiillers, Janina Buczylkog, and Preston Van Hoosers. 1992. “The Role of Arrestin and Retinoids in the Regeneration Pathway of Rhodopsin*.” 15701–6.

Hofmann, Klaus-Peter, P. Scheerer, P. , Hildebrand, HW Choe, JH Park, M. Heck, and O. P. Ernst. 2009. “A G Protein-Coupled Receptor at Work: The Rhodopsin Model.” Trends Biochem Sci 34(11): 540–52.

Hopkins, Andrew L, and Colin R Groom. 2002. “The Druggable Genome.” Nature reviews. Drug discovery 1(9): 727–30.

129 References

Janz, J M, J F Fay, and D L Farrens. 2003. “Stability of Dark State Rhodopsin Is Mediated by a Conserved Ion Pair in Intradiscal Loop E-2.” J Biol Chem 278(19): 16982–91.

Jastrzebska, Beata, Aleksander Debinski, Slawomir Filipek, and Krzysztof Palczewski. 2011. “Role of Membrane Integrity on G Protein-Coupled Receptors: Rhodopsin Stability and Function.” Progress in lipid research 50(3): 267–77.

Jastrzebska, Beata, Dimitrios Fotiadis, Geeng-Fu Jang, Ronald E Stenkamp, Andreas Engel, and Krzysztof Palczewski. 2006. “Functional and Structural Characterization of Rhodopsin Oligomers.” The Journal of biological chemistry 281(17): 11917–22.

Kaplan, M W. 1984. “Rhodopsin Lateral Diffusion as a Function of Rod Outer Segment Disk Membrane Axial Position.” Biophysical journal 45(4): 851–53.

Kapoor, Shobhna, Gemma Triola, Ingrid R Vetter, Mirko Erlkamp, Herbert Waldmann, and Roland Winter. 2011. “Revealing Conformational Substates of Lipidated N-Ras Protein by Pressure Modulation.”

Kasai, Rinshi S, and Akihiro Kusumi. 2014. “Single-Molecule Imaging Revealed Dynamic GPCR Dimerization.” Current opinion in cell biology 27: 78–86.

Kim, Miyeon, Susan M Hanson, Sergey a Vishnivetskiy, Xiufeng Song, Whitney M Cleghorn, Wayne L Hubbell, and Vsevolod V Gurevich. 2011. “Robust Self- Association Is a Common Feature of Mammalian Visual Arrestin-1.” Biochemistry 50(12): 2235–42.

Kim, Miyeon, Sergey a Vishnivetskiy, Ned Van Eps, Nathan S Alexander, Whitney M Cleghorn, Xuanzhi Zhan, Susan M Hanson, Takefumi Morizumi, Oliver P Ernst, Jens Meiler, Vsevolod V Gurevich, and Wayne L Hubbell. 2012. “Conformation of Receptor-Bound Visual Arrestin.” Proceedings of the National Academy of Sciences of the United States of America 109(45): 18407– 12.

Kim, Yong Ju, Klaus Peter Hofmann, Oliver P Ernst, Patrick Scheerer, Hui-Woog Choe, and Martha E Sommer. 2013. “Crystal Structure of Pre-Activated Arrestin p44.” Nature 497(7447): 142–46.

Kirchberg, Kristina, Tai-Yang Kim, Martina Möller, Darko Skegro, Gayathri Dasara Raju, Joachim Granzin, Georg Büldt, Ramona Schlesinger, and Ulrike Alexiev. 2011. “Conformational Dynamics of Helix 8 in the GPCR Rhodopsin Controls Arrestin Activation in the Desensitization Process.” Proceedings of the National Academy of Sciences of the United States of America 108(46): 18690– 95.

Kiser, Philip D, Marcin Golczak, and Krzysztof Palczewski. 2014. “Chemistry of the Retinoid (visual) Cycle.” Chemical reviews 114(1): 194–232.

130 References

Klähn, Marco, Gerald Mathias, Carsten Ko, Marco Nonella, Klaus Gerwert, and Paul Tavan. 2004. “IR Spectra of Phosphate Ions in Aqueous Solution : Predictions of a DFT / MM Approach Compared with Observations.” 108: 6186–94.

Knierim, B, K P Hofmann, O P Ernst, and W L Hubbell. 2007. “Sequence of Late Molecular Events in the Activation of Rhodopsin.” Proc Natl Acad Sci U S A 104(51): 20290–95.

Kovoor, a., J. Celver, R. I. Abdryashitov, C. Chavkin, and V. V. Gurevich. 1999. “Targeted Construction of Phosphorylation-Independent -Arrestin Mutants with Constitutive Activity in Cells.” Journal of Biological Chemistry 274(11): 6831–34.

Kühn, H, S W Hall, and U Wilden. 1984. “Light-Induced Binding of 48-kDa Protein to Photoreceptor Membranes Is Highly Enhanced by Phosphorylation of Rhodopsin.” FEBS letters 176(2): 473–78.

Lamb, T D, and E N Pugh Jr. 2006. “Phototransduction, Dark Adaptation, and Rhodopsin Regeneration the Proctor Lecture.” Invest Ophthalmol Vis Sci 47(12): 5137–52.

Langen, R, K Cai, C Altenbach, H G Khorana, and W L Hubbell. 1999. “Structural Features of the C-Terminal Domain of Bovine Rhodopsin: A Site-Directed Spin-Labeling Study.” Biochemistry 38(25): 7918–24.

Liang, Y, D Fotiadis, T Maeda, A Maeda, A Modzelewska, S Filipek, D A Saperstein, A Engel, and K Palczewski. 2004. “Rhodopsin Signaling and Organization in Heterozygote Rhodopsin Knockout Mice.” J Biol Chem 279(46): 48189–96.

Liebman, By Paul A, Henry L Weiner, and R E Drzymala. 1982. “Lateral Diffusion of Visual Pigment in Rod Disk Membranes.” 81(1976): 660–68.

Liebman, P A, and G Entine. 1974. “Lateral Diffusion of Visual Pigment in Photoreceptor Disk Membranes.” (August): 457–59.

Liu, M, and A Barth. 2003. “Mapping Interactions between the Ca2+-ATPase and Its Substrate ATP with Infrared Spectroscopy.” J Biol Chem 278(12): 10112– 18.

Lohse, M J, S Andexinger, J Pitcher, S Trukawinski, J Codina, J P Faure, M G Caron, and R J Lefkowitz. 1992. “Receptor-Specific Desensitization with Purified Proteins. Kinase Dependence and Receptor Specificity of Beta- Arrestin and Arrestin in the Beta 2-Adrenergic Receptor and Rhodopsin Systems.” J Biol Chem 267(12): 8558–64.

Lohse, Martin J. 2010. “Dimerization in GPCR Mobility and Signaling.” Current opinion in pharmacology 10(1): 53–58.

131 References

Lohse, Martin J, and Carsten Hoffmann. 2014. 219 Arrestins - Pharmacology and Therapeutic Potential. ed. Vsevolod V. Gurevich. Berlin, Heidelberg: Springer Berlin Heidelberg.

Lohse, Martin J, Isabella Maiellaro, and Davide Calebiro. 2014. “Kinetics and Mechanism of G Protein-Coupled Receptor Activation.” Current Opinion in Cell Biology 27: 87–93.

Lommerse, P H, G A Blab, L Cognet, G S Harms, B E Snaar-Jagalska, H P Spaink, and T Schmidt. 2004. “Single-Molecule Imaging of the H-Ras Membrane- Anchor Reveals Domains in the Cytoplasmic Leaflet of the Cell Membrane.” Biophys J 86(1 Pt 1): 609–16.

Mahalingam, M, K Martinez-Mayorga, M F Brown, and R Vogel. 2008. “Two Protonation Switches Control Rhodopsin Activation in Membranes.” Proc Natl Acad Sci U S A 105(46): 17795–800.

Mahalingam, Mohana, Karina Martínez-mayorga, Michael F Brown, and Reiner Vogel. 2008. “Two Protonation Switches Control Rhodopsin.” 105(46): 17795– 800.

Mansoor, S E, K Palczewski, and D L Farrens. 2006. “Rhodopsin Self-Associates in Asolectin Liposomes.” Proc Natl Acad Sci U S A 103(9): 3060–65.

McDowell, J H, J P Nawrocki, and P A Hargrave. 1993. “Phosphorylation Sites in Bovine Rhodopsin.” Biochemistry 32(18): 4968–74.

Michalet, Xavier. 2010. “Mean Square Displacement Analysis of Single-Paricle Trajectories with Localization Error: Brownian Motion in Isotropic Medium.” Phys Rev E Stat Nonlin Soft Matter Phys 82.

Moaven, Hormoz, Yukihiro Koike, Christine C Jao, Vsevolod V Gurevich, Ralf Langen, and Jeannie Chen. 2013. “Visual Arrestin Interaction with Clathrin Adaptor AP-2 Regulates Photoreceptor Survival in the Vertebrate Retina.” Proceedings of the National Academy of Sciences of the United States of America 110(23): 9463–68.

Natochin, M., K. G. Muradov, R. L. McEntaffer, and N. O. Artemyev. 2000. “Rhodopsin Recognition by Mutant Gs Containing C-Terminal Residues of Transducin.” Journal of Biological Chemistry 275(4): 2669–75.

Nikonov, Sergei S, Bruce M Brown, Jason a Davis, Freddi I Zuniga, Alvina Bragin, Edward N Pugh, and Cheryl M Craft. 2008. “Mouse Cones Require an Arrestin for Normal Inactivation of Phototransduction.” Neuron 59(3): 462–74.

Okada, T, M Sugihara, A N Bondar, M Elstner, P Entel, and V Buss. 2004. “The Retinal Conformation and Its Environment in Rhodopsin in Light of a New 2.2 A Crystal Structure.” J Mol Biol 342(2): 571–83.

132 References

Ostermaier, Martin K, Christian Peterhans, Rolf Jaussi, Xavier Deupi, and Jörg Standfuss. 2014. “Functional Map of Arrestin-1 at Single Amino Acid Resolution.” Proceedings of the National Academy of Sciences of the United States of America: 1–6.

Palczewski, K, J Buczylko, N R Imami, J H McDowell, and P A Hargrave. 1991. “Role of the Carboxyl-Terminal Region of Arrestin in Binding to Phosphorylated Rhodopsin.” J Biol Chem 266(23): 15334–39.

Palczewski, K, T Kumasaka, T Hori, C A Behnke, H Motoshima, B A Fox, I Le Trong, D C Teller, T Okada, R E Stenkamp, M Yamamoto, and M Miyano. 2000. “Crystal Structure of Rhodopsin: A G Protein-Coupled Receptor.” Science 289(5480): 739–45.

Palczewski, K, a Pulvermüller, J Buczyłko, and K P Hofmann. 1991. “Phosphorylated Rhodopsin and Heparin Induce Similar Conformational Changes in Arrestin.” The Journal of biological chemistry 266(28): 18649–54.

Palczewski, Krzysztof, Janina Buczylko, Hiroshi Ohguro, Roland S Annan, Steven A Carr, John W Crabb, Michael W Kaplan, Richard S Johnson, and Kenneth Walsh. 1994. “Characterization of a Truncated Form of Arrestin Isolated from Bovine Rod Outer Segments.” Protein Science: 314–24.

Palczewski, Krzysztof, J Hugh Mcdowells, Scott Jake, Thomas S Ingebritsen, and Paul a Hargrave. 1989. “Regulation of Rhodopsin.” 264(27): 15770–73.

Park, J H, P Scheerer, K P Hofmann, H W Choe, and O P Ernst. 2008. “Crystal Structure of the Ligand-Free G-Protein-Coupled Receptor Opsin.” Nature 454(7201): 183–87.

Parkes, J H, and P A Liebman. 1984a. “Temperature and pH Dependence of the Metarhodopsin I-Metarhodopsin II Kinetics and Equilibria in Bovine Rod Disk Membrane Suspensions.” Biochemistry 23(21): 5054–61.

Parkes, J H, and P a Liebman. 1984b. “Temperature and pH Dependence of the Metarhodopsin I-Metarhodopsin II Kinetics and Equilibria in Bovine Rod Disk Membrane Suspensions.” Biochemistry 23(21): 5054–61.

Pearring, Jillian N, Raquel Y Salinas, Sheila a Baker, and Vadim Y Arshavsky. 2013. “Protein Sorting, Targeting and Trafficking in Photoreceptor Cells.” Progress in retinal and eye research 36: 24–51.

Piechnick, Ronny, Martin Heck, and Martha E Sommer. 2011. “Alkylated Hydroxylamine Derivatives Eliminate Peripheral Retinylidene Schi Ff Bases but Cannot Enter the Retinal Binding Pocket of Light-Activated Rhodopsin.”

Piechnick, Ronny, Eglof Ritter, Peter W Hildebrand, Oliver P Ernst, Patrick Scheerer, Klaus Peter Hofmann, and Martin Heck. 2012. “Effect of Channel Mutations on the Uptake and Release of the Retinal Ligand in Opsin.” Proceedings of the National Academy of Sciences of the United States of America 109(14): 5247–52. 133 References

Poo, Mu-Ming, and Richard Cone. 1972. “Lateral Diffusion of Rhodopsin in Necturus Rods.” 17: 503–7.

Poo, Mu-Ming, and Richard Cone. 1974. “Lateral Diffusion of Rhodopsin in the Photoreceptor Membrane.” Nature 247: 438–41.

Pugh, E N Jr., and T Lamb. 2000. “Phototransduction in Vertebrate Rods and Cones: Molecular Mechanism of Amplification, Recovery and Light Adaptation.” In Molecular Mechanism in Visual Transduction, eds. D G Stavenga, W J DeGrip, and E N Jr. Pugh. Amsterdam: Elsevier science Publisher BV, 183–255.

Pulvermüller, a, D Maretzki, M Rudnicka-Nawrot, W C Smith, K Palczewski, and K P Hofmann. 1997. “Functional Differences in the Interaction of Arrestin and Its Splice Variant, p44, with Rhodopsin.” Biochemistry 36(30): 9253–60.

Rieke, F, and D A Baylor. 1998. “Origin of Reproducibility in the Responses of Retinal Rods to Single Photons.” Biophys J 75(4): 1836–57.

Ritter, E, M Elgeti, K P Hofmann, and F J Bartl. 2007. “Deactivation and Proton Transfer in Light-Induced Metarhodopsin II/metarhodopsin III Conversion: A Time-Resolved Fourier Transform Infrared Spectroscopic Study.” J Biol Chem 282(14): 10720–30.

Ritter, Eglof, Matthias Elgeti, and Franz J. Bartl. 2008. “Activity Switches of Rhodopsin.” Photochemistry and photobiology 84(4): 911–20.

Ritter, Eglof, Matthias Elgeti, Klaus Peter Hofmann, and Franz J. Bartl. 2007. “Deactivation and Proton Transfer in Light-Induced Metarhodopsin II/metarhodopsin III Conversion: A Time-Resolved Fourier Transform Infrared Spectroscopic Study.” The Journal of biological chemistry 282(14): 10720–30.

Rodieck, R W. 1998. The First Steps in Seeing. Sunderland, Massachusetts: Sinauer Associated, Inc.

Roof, D J, and J E Heuser. 1982. “Surfaces of Rod Photoreceptor Disk Membranes: Integral Membrane Components.” J Cell Biol 95(2 Pt 1): 487–500.

Rosenbaum, D M, S G Rasmussen, and B K Kobilka. 2009. “The Structure and Function of G-Protein-Coupled Receptors.” Nature 459(7245): 356–63.

Rotanowska, Maigorzata, and Tadeusz Sarna. 2005. “Invited Review Light-Induced Damage to the Retina : Role of Rhodopsin Chromophore Revisited.”

Ruprecht, J J, T Mielke, R Vogel, C Villa, and G F Schertler. 2004. “Electron Crystallography Reveals the Structure of Metarhodopsin I.” EMBO J 23(18): 3609–20.

Sakmar, T P, R R Franke, and H G Khorana. 1991. “The Role of the Retinylidene Schiff Base Counterion in Rhodopsin in Determining Wavelength Absorbance and Schiff Base pKa.” Proc Natl Acad Sci U S A 88(8): 3079–83.

134 References

Scheerer, P, J H Park, P W Hildebrand, Y J Kim, N Krauss, H W Choe, K P Hofmann, and O P Ernst. 2008. “Crystal Structure of Opsin in Its G-Protein- Interacting Conformation.” Nature 455(7212): 497–502.

Schleicher, A, H Kühn, and K P Hofmann. 1989. “Kinetics, Binding Constant, and Activation Energy of the 48-kDa Protein-Rhodopsin Complex by Extra- Metarhodopsin II.” Biochemistry 28(4): 1770–75.

Schleicher, Andreas, Hermann Kuhn, and Klaus Peter. 1989. “Kinetics, Binding Constant, and Activation Energy of the 48-kDa Protein-Rhodopsin Complex by Extra-Metarhodopsin II.” 1770–75.

Schröder, K, A Pulvermüller, and K P Hofmann. 2002. “Arrestin and Its Splice Variant Arr1-370A (p44). Mechanism and Biological Role of Their Interaction with Rhodopsin.” J Biol Chem 277(46): 43987–96.

Schubert, C, J A Hirsch, V V Gurevich, D M Engelman, P B Sigler, and K G Fleming. 1999. “Visual Arrestin Activity May Be Regulated by Self- Association.” J Biol Chem 274(30): 21186–90.

Schütz, G J, H Schindler, and T Schmidt. 1997. “Single-Molecule Microscopy on Model Membranes Reveals Anomalous Diffusion.” Biophysical journal 73(2): 1073–80.

Shukla, Arun K, Aashish Manglik, Andrew C Kruse, Kunhong Xiao, Rosana I Reis, Wei-Chou Tseng, Dean P Staus, Daniel Hilger, Serdar Uysal, Li-Yin Huang, Marcin Paduch, Prachi Tripathi-Shukla, Akiko Koide, Shohei Koide, William I Weis, Anthony a Kossiakoff, Brian K Kobilka, and Robert J Lefkowitz. 2013. “Structure of Active Β-Arrestin-1 Bound to a G-Protein-Coupled Receptor Phosphopeptide.” Nature 497(7447): 137–41.

Shukla, Arun K., Gerwin H. Westfield, Kunhong Xiao, Rosana I. Reis, Li-Yin Huang, Prachi Tripathi-Shukla, Jiang Qian, Sheng Li, Adi Blanc, Austin N. Oleskie, Anne M. Dosey, Min Su, Cui-Rong Liang, Ling-Ling Gu, Jin-Ming Shan, Xin Chen, Rachel Hanna, Minjung Choi, Xiao Jie Yao, Bjoern U. Klink, Alem W. Kahsai, Sachdev S. Sidhu, Shohei Koide, Pawel a. Penczek, Anthony a. Kossiakoff, Virgil L. Woods Jr, Brian K. Kobilka, Georgios Skiniotis, and Robert J. Lefkowitz. 2014. “Visualization of Arrestin Recruitment by a G- Protein-Coupled Receptor.” Nature.

Shukolyukov, S a. 2009. “Aggregation of Frog Rhodopsin to Oligomers and Their Dissociation to Monomer: Application of BN- and SDS-PAGE.” Biochemistry. Biokhimiia͡ 74(6): 599–604.

Siebert, Friedrich. 1995. “Application of FTIR Spectroscopy to the Investigation of Dark Structures and Photoreactions of Visual Pigments.” Israel Journal of Chemistry 35: 309–23.

Siebert, Friedrich, Werner Mantele, and Klaus Gerwert. 1983. “The Problem of the Protonation State of the Retinylidene Schiff Base Re-Investigated.” 127: 119– 27. 135 References

Sinha, Abhinav, Amber Mary Jones Brunette, Jonathan F Fay, Christopher T Schafer, and David L Farrens. 2014. “Rhodopsin TM6 Can Contact Two Separate and Distinct Sites on Arrestin: Evidence for Structural Plasticity and Multiple Docking Modes in Arrestin-Rhodopsin Binding.” Biochemistry.

Smith, Steven O. 2010. “Structure and Activation of the Visual Pigment Rhodopsin.”

Smith, W C, A Dinculescu, J J Peterson, and J H McDowell. 2004. “The Surface of Visual Arrestin That Binds to Rhodopsin.” Mol Vis 10: 392–98.

Smith, W Clay. 2013. 118 Progress in molecular biology and translational science The Role of Arrestins in Visual and Disease Processes of the Eye. 1st ed. Elsevier Inc.

Sommer, M E, and D L Farrens. 2006. “Arrestin Can Act as a Regulator of Rhodopsin Photochemistry.” Vision Res 46(27): 4532–46.

Sommer, M E, D L Farrens, J H McDowell, L A Weber, and W C Smith. 2007. “Dynamics of Arrestin-Rhodopsin Interactions: Loop Movement Is Involved in Arrestin Activation and Receptor Binding.” J Biol Chem 282(35): 25560–68.

Sommer, M E, W C Smith, and D L Farrens. 2006. “Dynamics of Arrestin- Rhodopsin Interactions: Acidic Phospholipids Enable Binding of Arrestin to Purified Rhodopsin in Detergent.” J Biol Chem 281(14): 9407–17.

Sommer, Martha E, Klaus Peter Hofmann, and Martin Heck. 2011. “Arrestin- Rhodopsin Binding Stoichiometry in Isolated Rod Outer Segment Membranes Depends on the Percentage of Activated Receptors.” The Journal of biological chemistry 286(9): 7359–69.

Sommer, Martha E, Klaus Peter Hofmann, and Martin Heck. 2012. “Distinct Loops in Arrestin Differentially Regulate Ligand Binding within the GPCR Opsin.” Nature communications 3: 995.

Sommer, Martha E, W Clay Smith, and David L Farrens. 2006. “Dynamics of Arrestin-Rhodopsin Interactions: Acidic Phospholipids Enable Binding of Arrestin to Purified Rhodopsin in Detergent.” The Journal of biological chemistry 281(14): 9407–17.

Sommer, Martha, Klaus Peter Hofmann, and Martin Heck. 2014. “Not Just Signal Shutoff: The Protective Role of Arrestin-1 in Rod Cells.”

Strissel, K J, M Sokolov, L H Trieu, and V Y Arshavsky. 2006. “Arrestin Translocation Is Induced at a Critical Threshold of Visual Signaling and Is Superstoichiometric to Bleached Rhodopsin.” J Neurosci 26(4): 1146–53.

Suda, K, S Filipek, K Palczewski, A Engel, and D Fotiadis. 2004. “The Supramolecular Structure of the GPCR Rhodopsin in Solution and Native Disc Membranes.” Mol Membr Biol 21(6): 435–46.

136 References

Sutton, R B, S A Vishnivetskiy, J Robert, S M Hanson, D Raman, B E Knox, M Kono, J Navarro, and V V Gurevich. 2005. “Crystal Structure of Cone Arrestin at 2.3A: Evolution of Receptor Specificity.” J Mol Biol 354(5): 1069–80.

Szczepek, Michal, Florent Beyrière, Klaus Peter Hofmann, Matthias Elgeti, Roman Kazmin, Alexander Rose, Franz J. Bartl, David von Stetten, Martin Heck, Martha E. Sommer, Peter W. Hildebrand, and Patrick Scheerer. 2014a. “Crystal Structure of a Common GPCR-Binding Interface for G Protein and Arrestin.” Nature Communications 5(May): 4801.

Takezoe, Hideo, and Hyuk Yu. 1981. “Lateral Diffusion of Photopigments in Photoreceptor Disk Membrane Vesicles by the Dynamic Kerr Effect.” 20: 5275–81.

Tsukamoto, Hisao, Abhinav Sinha, Mark DeWitt, and David L Farrens. 2010. “Monomeric Rhodopsin Is the Minimal Functional Unit Required for Arrestin Binding.” Journal of molecular biology 399(3): 501–11.

Vishnivetskiy, S A, D Raman, J Wei, M J Kennedy, J B Hurley, and V V Gurevich. 2007. “Regulation of Arrestin Binding by Rhodopsin Phosphorylation Level.” J Biol Chem 282(44): 32075–83.

Vishnivetskiy, S A, C Schubert, G C Climaco, Y V Gurevich, M G Velez, and V V Gurevich. 2000. “An Additional Phosphate-Binding Element in Arrestin Molecule. Implications for the Mechanism of Arrestin Activation.” J Biol Chem 275(52): 41049–57.

Vogel, R, S Martell, M Mahalingam, M Engelhard, and F Siebert. 2007. “Interaction of a G Protein-Coupled Receptor with a G Protein-Derived Peptide Induces Structural Changes in Both Peptide and Receptor: A Fourier-Transform Infrared Study Using Isotopically Labeled Peptides.” J Mol Biol 366(5): 1580– 88.

Vogel, R, and F Siebert. 2001. “Conformations of the Active and Inactive States of Opsin.” The Journal of biological chemistry 276(42): 38487–93.

Vogel, Reiner, Swetlana Martell, Mohana Mahalingam, Martin Engelhard, and Friedrich Siebert. 2007. “Interaction of a G Protein-Coupled Receptor with a G Protein-Derived Peptide Induces Structural Changes in Both Peptide and Receptor: A Fourier-Transform Infrared Study Using Isotopically Labeled Peptides.” Journal of molecular biology 366(5): 1580–88.

Vogel, Reiner, and F Siebert. 2003. “Fourier Transform IR Spectroscopy Study for New Insights into Molecular Properties and Activation Mechanisms of Visual Pigment Rhodopsin.” Biopolymers 72(3): 133–48.

Vogel, Reiner, Friedrich Siebert, Gerald Mathias, Paul Tavan, Guibao Fan, and Mordechai Sheves. 2003. “Deactivation of Rhodopsin in the Transition from the Signaling State Meta II to Meta III Involves a Thermal Isomerization of the Retinal Chromophore C[double bond]D.” Biochemistry 42(33): 9863–74.

137 References

Werner, Karla, Christian Richter, Judith Klein-Seetharaman, and Harald Schwalbe. 2008. “Isotope Labeling of Mammalian GPCRs in HEK293 Cells and Characterization of the C-Terminus of Bovine Rhodopsin by High Resolution Liquid NMR Spectroscopy.” Journal of biomolecular NMR 40(1): 49–53.

Wey, C L, R a Cone, and M a Edidin. 1981. “Lateral Diffusion of Rhodopsin in Photoreceptor Cells Measured by Fluorescence Photobleaching and Recovery.” Biophysical journal 33(2): 225–32.

Whited, Allison M, and Paul S-H Park. 2014. “Nanodomain Organization of Rhodopsin in Native Human and Murine Rod Outer Segment Disc Membranes.” Biochimica et biophysica acta.

Xiao, K, D B McClatchy, A K Shukla, Y Zhao, M Chen, S K Shenoy, J R Yates 3rd, and R J Lefkowitz. 2007. “Functional Specialization of Beta-Arrestin Interactions Revealed by Proteomic Analysis.” Proc Natl Acad Sci U S A 104(29): 12011–16.

Zaitseva, Ekaterina, Michael F Brown, and Reiner Vogel. 2010. “Sequential Rearrangement of Interhelical Networks upon Rhodopsin Activation in Membranes: The Meta II(a) Conformational Substate.” Journal of the American Chemical Society 132(13): 4815–21.

Zhan, Xuanzhi, Luis E Gimenez, Vsevolod V Gurevich, and Benjamin W Spiller. 2011. “Crystal Structure of Arrestin-3 Reveals the Basis of the Difference in Receptor Binding between Two Non-Visual Subtypes.” Journal of molecular biology 406(3): 467–78.

Zhuang, Tiandi, Qiuyan Chen, Min-Kyu Cho, Sergey a Vishnivetskiy, Tina M Iverson, Vsevolod V Gurevich, and Charles R Sanders. 2013. “Involvement of Distinct Arrestin-1 Elements in Binding to Different Functional Forms of Rhodopsin.” Proceedings of the National Academy of Sciences of the United States of America 110(3): 942–47.

138 Published journal articles

Published journal articles

Parts of this thesis are published in the following journal articles:

- Michal Szczepek, Florent Beyrière, Klaus Peter Hofmann, Matthias Elgeti, Roman Kazmin, Alexander Rose, Franz J. Bartl, David von Stetten, Martin Heck, Martha E. Sommer, Peter W. Hildebrand, and Patrick Scheerer. Crystal structure of a common GPCR-binding interface for G protein and arrestin. (2014) Nature Communications, 5, 4801.

- Florent Beyrière, Martha E.Sommer, Michal Szczepek, Franz J. Bartl, Klaus Peter Hofmann, Martin Heck, and Eglof Ritter. Formation and decay of the arrestin-rhodopsin complex in native disc membrane. Submitted to Journal of biological chemistry, October 2014.

139 Acknowledgements

140 Acknowledgements

Acknowledgements

I would like to express my gratitude to everyone who supported me throughout the course of this doctor thesis. I am thankful for their aspiring guidance, constructive criticism and friendly advice during my work. I am sincerely grateful to Prof. Dr. Peter Hildebrandt of the Technischen Universität Berlin, who supervised this work and to Prof. Dr. Klaus Peter Hofmann who gave me the possibility to work in the Institut für Medizinische Physik und Biophysik (IMPB) at the Charité Universitätsmedizin Berlin. I would like to thank them for their thesis corrections and remarks. I am especially grateful to Dr. Martin Heck who directly supervised my work and developed a lot of ideas, thereby making this dissertation possible. His expertise and feedback was extremely helpful to interpret the data. Special thanks to Dr. Eglof Ritter who performed the analysis of the FTIR row-data and supported me with performing FTIR measurements. I would also like to thank Dr. Michal Szczepek and Dr. Patrick Scheerer for their successful collaborations with the crystallography project and their great support for general questions and discussions. I express my warm thanks to Prof. Dr. Horst Vogel for giving me the opportunity to perform the single fluorescence microscopy experiments in his laboratory at the Swiss Federal Institute of Technology in Lausanne and to his collaborator Dr. Joachim Piguet who analyzed the videos and the single molecule traces. I would also like to thank Dr. Martha Sommer for her kindness and her support on all arrestin questions and also Justus Lorke, Ciara Lally and Dr. Ronny Piechnick of the IMPB and to Dr. Mathias Prigge, Anton Rösler, Dr. Blaise Petitpierre for their friendship, support and precious advice. It would not have been possible to work in the laboratory without the help of the technicians of the IMPB. I am in particular grateful to Brian Bauer and Anja Koch for their help and logistical support and to Andreas von Garnier who helped me to overcome all electronic failures of the spectrometer. My parents and brother have given me their unequivocal support throughout, as always, for which my mere expression of thanks likewise does not suffice. I modestly dedicate this work to them.

141 Acknowledgements

142 Selbständigkeitserklärung

Selbständigkeitserklärung

Eidesstattliche Erklärung

Name: Florent Beyrière Matrikelnummer: 346120

Hiermit versichere ich an Eides Statt, dass die vorliegende Arbeit, bis auf die offizielle Betreuung, selbstständig und ohne fremde Hilfe durch mich angefertigt wurde und benutzte Quellen und Hilfsmittel vollständig angegeben sind.

Berlin, November 2014

Florent Beyrière

143