Protein-membrane interactions investigated with time-resolved FTIR spectroscopy

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Protein-membrane interactions investigated with time-resolved FTIR spectroscopy

DISSERTATION for the academic degree of

doctor of natural sciences (Dr. rer. nat.) presented by

Michael Jawurek submitted to

Mathematisch-Naturwissenschaftliche Sektion Fachbereich Chemie

Date of the oral examination: 20.12.2017

First referee: Prof. Dr. Karin Hauser

Second referee: Prof. Dr. Andreas Zumbusch

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Abstract

Membrane proteins are key players for various biological processes and are essential for a living cell. The interaction of these membrane proteins with their surrounding lipid membranes is of high importance for their functionality. Retinal proteins are membrane proteins, whose function can be triggered with light and are used here as model systems to investigate these protein-lipid interactions. Therefore, Bacteriorhodopsin and Prote- orhodopsin were reconstituted into liposomes by systematically varying lipid physical properties. Both retinal proteins are light-driven proton pumps, which pump one proton across the membrane following light excitation. A step-scan FTIR experimental setup was established, which allows to monitor time-resolved dynamics of the photoreceptors.

In particular the dynamics of the primary proton transfer step in the core of the protein were probed to monitor the influence of the membrane on the function of the protein.

The primary proton transfer step involves deprotonation of the retinal Schiff base and protonation of Asp85 for Bacteriorhodopsin, Asp97 for Proteorhodopsin, respectively.

Variation of the lipid chain length, led only to minor changes on the protonation dynamics, whereas the degree of saturation and the lipid phase had a significant influence.

Temperature-dependent measurements below and above the phase transition temperature of the lipids strengthened these observations. It is concluded that protonation of Asp85 for Bacteriorhodopsin is accelerated in the lipid liquid phase followed by a longer-lived M intermediate as compared to the lipid gel phase. For Proteorhodopsin, both protonation as well as deprotonation dynamics of Asp97 are accelerated in case of the lipid liquid phase. Variation of lipid headgroups revealed a strong influence on the protonation dynamics of Bacteriorhodopsin displaying in particular slower dynamics for positively charged lipids, whereas for Proteorhodopsin no alterations on the dynamics could be resolved.

Furthermore, conformational dynamics of the retinal proteins as well as dynamics of the lipids were explored. It was demonstrated that structural changes of the protein correlate with protonation dynamics. Deuterated lipids were introduced to monitor dynamics of the lipid chains in a region free from the absorption of other groups. Finally, pH-dependent measurements for Proteorhodopsin reveal the pH-dependent protonation dynamics in different lipid environments.

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Kurzfassung

Membranproteine spielen eine Schlüsselrolle für verschiedenste biologische Prozesse und sind essentiell für eine lebende Zelle. Die Interaktion dieser Membranproteine mit ihren Membranumgebungen ist von enormer Wichtigkeit für Ihre Funktionalität. Retinalpro- teine sind Membranproteine, deren Funktion mit Licht ausgelöst wird und die hier als Modellsysteme zur Untersuchung von Protein-Lipid Wechselwirkungen verwendet wurden.

Dazu wurden Bacteriorhodopsin und Proteorhodopsin in Liposomen rekonstituiert und die physikalischen Eigenschaften der Lipide systematisch variiert. Beide Retinalproteine sind lichtgetriebene Protonenpumpen, welche nach Lichtanregung ein Proton durch die Membran pumpen. Ein step-scan FTIR Versuchsaufbau wurde etabliert, welcher es erlaubt, die Dynamik der Photorezeptoren zeitaufgelöst zu beobachten. Insbesondere die Dynamik des ersten Protonentransfer Schrittes im Inneren des Proteins wurde untersucht, um den Einfluss der Membran auf die Funktion des Proteins zu verfolgen. Der erste Protonentransfer Schritt beinhaltet die Deprotonierung der Schiffschen Base und Pro- tonierung von Asp85 für Bacteriorhodopsin, beziehungsweise Asp97 für Proteorhodopsin.

Eine Variation der Lipidkettenlänge führte nur zu geringen beobachtbaren Änderungen der Protonierungsdynamik, wohingegen der Sättigungsgrad und die Lipidphase einen sig- nifikanten Einfluss hatten. Temperaturabhängige Messungen unterhalb und oberhalb der Phasenübergangstemperatur der Lipide bestärkten diese Beobachtungen. Es wurde gefol- gert, dass in der flüssigen Lipidphase die Protonierung von Asp85 für Bacteriorhodopsin beschleunigt und gefolgt von einem längerlebenden M Intermediat im Vergleich zur Gel-Lipidphase stattfindet. Für Proteorhodopsin sind sowohl Protonierungs- als auch Deprotonierungsdynamiken von Asp97 in der flüssigen Phase beschleunigt. Variation der Lipidkopfgruppen führte zu einem starken Einfluss auf die Protonierungsdynamik von Bacteriorhodopsin, beobachtbar insbesondere durch langsamere Dynamiken positiv geladener Kopfgruppen, wohingegen für Proteorhodopsin keine Änderungen der Dynamik aufgelöst werden konnten. Des Weiteren wurde die Konformationsdynamik der Retinal- proteine, sowie die Dynamik der umgebenden Lipide erforscht. Es konnte gezeigt werden, dass strukturelle Änderungen des Proteins mit der Protonierungsdynamik korrelieren.

Deuterierte Lipide wurden eingeführt, um die Dynamik der Lipidketten in einer spek- tralen Region beobachten zu können, die frei von der Absorption von anderen Gruppen ist. Schließlich konnten pH-abhängige Messungen an Proteorhodopsin die pH-abhängige Protonierungsdynamik in unterschiedlichen Membranen aufzeigen.

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Publications

M. Jawurek, C. Glaubitz, K. Hauser, Impact of the lipid environment on the protonation dynamics of bacteriorhodopsin studied with time-resolved step-scan FTIR spectroscopy, Biomed. Spectrosc. and Imaging 5 (2016) 167-174.

M. Jawurek, J. Dröden, B. Peter, C. Glaubitz, K. Hauser, Membrane influence on protonation dynamics of photoreceptors monitored with time-resolved FTIR spectroscopy, submitted to Chemical Physics.

Poster Presentations

M. Jawurek, S. Schmidt, C. Glaubitz, K. Hauser, Influence of the lipid environment on the kinetics of photoreceptors studied by step-scan FTIR spectroscopy, Meeting of the Dutch and German Biophysical Society, Hünfeld, 2015.

M. Jawurek, S. Schmidt, C. Glaubitz, K. Hauser, Time-resolved FTIR spectroscopy to monitor the influence of the lipid environment on retinal proteins, 16th European Conference on the Spectroscopy of Biological Molecules (ECSBM), Bochum, 2015.

M. Jawurek, C. Glaubitz, K. Hauser, Dynamics of photoreceptors in different lipid environments studied with time-resolved FTIR spectroscopy, Joint meeting of the membrane sections of the French and German Biophysical Societies, Bad Herrenalp, 2016.

M. Jawurek, J. Dröden, C. Glaubitz, K. Hauser, Influence of the lipid environment on the function of Proteorhodopsin monitored with time-resolved FTIR spectroscopy, 17th International Conference on Retinal Proteins, Potsdam, 2016.

M. Jawurek, J. Dröden, B. Peter, C. Glaubitz, K. Hauser, Effect of membrane lipid properties on protonation dynamics of photoreceptors studied with time-resolved FTIR spectroscopy, Time-resolved vibrational spectroscopy meeting (TRVS), Cambridge (UK), 2017.

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Abbreviations

AC Alternating current

ADC Analog-to-digital converter ADP Adenosine diphosphate Asp Aspartic acid

ATP Adenosine triphosphate BR Bacteriorhodopsin

CMC Critical micellar concentration DC Direct current

DDM n-dodecyl-β-D-maltoside

DMoPC 1,2-dimyristoleoyl-sn-glycero-3-phosphocholine DMPA 1,2-dimyristoyl-sn-glycero-3-phosphate

DMPC 1,2-dimyristoyl-sn-glycero-3-phosphocholine

DMPG 1,2-dimyristoyl-sn-glycero-3-phospho-(1’-rac-glycerol) DMTAP 1,2-dimyristoyl-3-trimethylammonium-propane

DNA Deoxyribonucleic acid

DOPC 1,2-dioleoyl-sn-glycero-3-phosphocholine DOPE 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine DOPG 1,2-dioleoyl-sn-glycero-3-phospho-(1’-rac-glycerol) DOPS 1,2-dioleoyl-sn-glycero-3-phospho-L-serine

DOTAP 1,2-dioleoyl-3-trimethylammonium-propane DPPC 1,2-dipalmitoyl-sn-glycero-3-phosphocholine DSPC 1,2-distearoyl-sn-glycero-3-phosphocholine E.coli Escherichia coli

EPR Electron paramagnetic resonance FTIR Fourier-transform infrared

Glu Glutamic acid

IR Infrared

Lys Lysine

MCT Mercury cadmium telluride

MES 2-(N-morpholino)ethanesulfonic acid MSP Membrane scaffold protein

Nd:YAG Neodynium-doped yttrium aluminum garnet OG n-octyl-β-D-glucoside

PC Phosphatidylcholine

POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine PR Proteorhodopsin

ssNMR Solid-state nuclear magnetic resonance UV-vis Ultraviolet-visible

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1. Introduction

The examination of life on earth corresponds probably to one of the most fundamental and fascinating fields of research. Amazingly, all life on this planet is basically built up by the same four building blocks of life: nucleic acids, proteins, lipids and carbohydrates.

While nucleic acids in form of DNA code the genetic information, proteins execute the biological processes in a cell and are the key players performing a massive range of functions. Lipid molecules form membranes, which are selective permeable barriers and allow compartmentalization and a higher degree of organization of the cell. Proteins in the lipid membrane are responsible for the communication between the intracellular and extracellular environment. All kinds of signaling or transport processes across the membrane are mediated by these membrane proteins. Among these, the large family of G-protein-coupled receptors constitutes of around half of all known drug targets [1]. A thorough understanding of the interactions between lipids and these membrane proteins during protein function is of fundamental importance. Retinal proteins, which share the conserved structural pattern of G-protein-coupled receptors are a good model system to investigate these interactions. Following light excitation, retinal proteins pass through a photocycle. Time-resolved spectroscopy allows the characterisation of the photoreaction and gives therefore the opportunity to analyze the influence of the membrane environment on the kinetics of the photocycle. In Chapter 2, a variety of retinal proteins are first introduced, ranging from microbial rhodopsins to the visual rhodopsin in the human eye. The focus is then set onto structural and functional properties of Bacteriorhodopsin and Proteorhodopsin, the two retinal proteins used within this work. Basic properties of lipid membranes are presented and the current state of research on protein-lipid interactions is summarized. Chapter 3 motivates the following experimental methods used in this project which are presented in Chapter 4.

Bacteriorhodopsin purification and reconstitution into proteoliposomes is described and fundamentals of Fourier-transform infrared (FTIR) spectroscopy are recapitulated. A brief overview on basic aspects of vibrational spectroscopy is given. The approach of

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1. Introduction

reaction-induced FTIR difference spectroscopy and the assignment of protein and lipid bands is covered in this chapter. Time-resolved step-scan FTIR spectroscopy including the method and established experimental setup is explained in detail. Chapter 5 shows time-resolved step-scan FTIR data for both Bacteriorhodopsin and Proteorhodopsin. The variation of the lipid physical properties of the membrane environment and its influence on the function of the two photoreceptors lies in the focus of this chapter. Moreover, the attempt of recording lipid and protein dynamics simultaneously is described. Finally, pH-dependent protonation dynamics of Proteorhodopsin are explored.

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2. Current state of research

2.1. Retinal proteins

Retinal proteins with seven transmembrane α-helices and the chromophore retinal covalently bound to the protein play an outstanding role as photoreceptors in cells. They are abundant in all kingdoms of life [2] and include the visual protein rhodopsin which converts light into an electrical nerve stimulus in our eyes. Microbial rhodopsins found in archaea or bacteria also belong to the group of retinal proteins, sharing the same structural features, but can perform a wide range of different functions in the cells. Using light energy, they are able to act as pumps or channels for ions across the membrane or as sensors for e.g. phototaxis [3]. The family of retinal proteins can hence be classified into two different types. Type I retinal proteins transport different substrates across the membrane or function as phototaxis receptors while type II retinal proteins consist of photosensitive receptor proteins in animal eyes including human rod and cone visual pigments such as visual rhodopsin located in the retina of the human eye. Rhodopsin is denoted as the entity of retinal and protein, with the latter being termed opsin. For the retinal, a strong dependency of the molecular environment on the maximum of light absorption can be observed, also known as Opsin-shift. The absorption spectra of retinal proteins are modulated by the interaction of the retinal with the amino acids located close to the retinal. The maximum of absorption can thereby vary between 360 nm and 570 nm [4]. Table 2.1 lists selected retinal proteins with their corresponding wavelength of maximal absorption, isomerization state of the retinal in the ground state and type of retinal protein. The type I proteins Bacteriorhodopsin (λ_max = 568 nm) and Proteorhodopsin (λ_max = 525 nm) are light-driven proton pumps and will be discussed in detail in the following chapters. Halorhodopsin (λ_max = 570 nm) is a light-driven chloride ion pump. It is found like Bacteriorhodopsin in Halobacterium salinarium and has an inwardly directed pumping direction. Another archaeal retinal protein is sensory-Rhodopsin II (λ_max = 487 nm) which in contrast acts as a sensor for phototaxis,

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2. Current state of research

meaning that it triggers movement of the whole organism towards or away from light.

Channelrhodopsin-2 (λ_max = 470 nm) from the algae Chlamydomonas reinhardtii [5]

functions as an ion channel as the name implies. Its discovery opened the fascinating new field of optogenetics. Channelrhodopsin-2 is expressed in neurons and by exposing the neurons to pulses of light, they can be activated in a temporally precise manner [6]. For all type I retinal proteins, the retinal is in the all-trans conformation in the ground state and isomerizes to 13-cis through light excitation. The absorbed light energy drives then the conformational changes of the protein and respective functional tasks.

Intermediate states are formed during this process and finally, the protein relaxes back to the ground state which is usually shown in form of a photocycle. After passing through the photocycle, absorption of new photons can trigger the photocycle in a repetitive fashion. The photocycle of Bacteriorhodopsin lasts e.g. around 10 ms [7]. During this photocycle, one proton is pumped across the membrane. Due to the ease of triggering the function of type I retinal proteins with light and the fast repeatability of the process, these proteins are ideal candidates to study general mechanisms and functions of membrane proteins. In contrast for type II retinal proteins the retinal is in the 11-cis conformation in the ground state and isomerizes to all-trans when absorbing light (Figure 2.1). Type II visual rhodopsin (λ_max = 500 nm) in vertebrate eyes as well follows a photocycle with distinct intermediates but undergoes a slow process of regeneration that involves release of the photo-isomerised all-trans retinal and binding of a new 11-cis retinal [8]. For all retinal proteins the retinal is covalently bound to a lysine residue via a Schiff base linkage in the core of the protein. The protonated Schiff base is usually surrounded by a counter-ion complex and plays a key role for the light-triggered function of the protein.

Table 2.1.: Properties of selected retinal proteins showing the wavelength of maximal absorption [9–13] as well as the state of isomerization of the retinal before light triggering.

For type 1 retinal proteins the retinal isomerizes from all-trans to 13-cis whereas for type 2 retinal proteins the retinal isomerizes from 11-cis to all-trans.

Protein Wavelength Abs_max [nm] Retinal Isomerization Type

Bacteriorhodopsin 568 all-trans I

Proteorhodopsin 525 all-trans I

Halorhodopsin 570 all-trans I

Sensory Rhodopsin II 487 all-trans I

Channelrhodopsin-2 470 all-trans I

Visual Rhodopsin 500 11-cis II

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2.2. Bacteriorhodopsin

CH₃

CH₃ CH₃ CH₃ CH₃

O

O CH₃ CH₃

CH₃ CH₃ CH₃

CH₃

CH₃CH₃ CH₃ CH₃

O CH₃

CH₃ CH₃ CH₃

O CH₃

13

11

Figure 2.1.: Isomerization of the retinal from all-trans to 13-cis (left) and from 11-cis to all-trans (right). Molecular bonds involved in the isomerization are highlighted in green.

2.2. Bacteriorhodopsin

2.2.1. Overview and history of Bacteriorhodopsin

Bacteriorhodopsin is an integral membrane protein discovered in 1973 fromD. Oester- heltandW. Stoeckenius [14]. It was found in the purple membrane ofHalobacterium salinarium which is an archaea bacterium found in waters with very high salt concentration. Bacteriorhodopsin weighs 27 kDa and is the only membrane protein in the purple membrane. Using light, it can pump protons across the cell membrane producing a proton gradient which Halobacteria use as a source of energy production in form of ATP synthesis or drive e.g. flagellar motility. Therefore, even under anaerobic conditions, light is sufficient for the Halobacterium to maintain its energy demand [15]. Bacteriorhodopsin is a very robust protein and served since its discovery as a model for membrane proteins.

It is considered to be the best-understood membrane protein.

2.2.2. Structure and function of Bacteriorhodopsin

Bacteriorhodopsin (BR), containing 248 amino acids, folds into the membrane as a seven α-helices transmembrane receptor.The α-helices are arranged in a parallel fashion, all spanning the whole range across the membrane. These seven α-helix bundles are a conserved structural pattern for retinal proteins and G-protein-coupled receptors [16].

Figure 2.2 shows the structure of BR according to the protein data bank ID 1FBB. The

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loop in between helix B and helix C forms an antiparallel β-strand colored in yellow.

The retinal chromophore depicted in cyan is centered in the core of the protein and is connected via a protonated Schiff base linkage to the lysine residue K216 (orange) [17]. It separates a cytoplasmic half-channel from an extracellular half-channel which are composed of amino acids playing an important role for the proton transfer. Key residue Asp85 of the extracellular channel is depicted in blue in Figure 2.2. Asp85 first accepts the proton from the retinal Schiff base (primary proton acceptor). Asp96 which subsequently reprotonates the retinal Schiff base is marked in gray (primary proton donor). The oligomerization state of BR in the native purple membrane is a trimer packed in a hexagonal lattice, but the monomer in detergent has been shown to be functional as well [18].

Figure 2.2.: 3D structure of Bacteriorhodopsin according to protein data bank ID: 1FBB.

Secondary structure elements are shown in form of a cartoon with α-helices in red, antiparallel β-strand in yellow and loops connecting the α-helices in green.

Highlighted are the retinal depicted in cyan, primary proton acceptor D85 in blue, primary proton donor D96 in gray and K216 in orange.

BR undergoes a cyclic reversible photocycle induced by light with several spectral distinct photointermediates termed J, K, L, M, N and O (Figure 2.3). The indices indicate

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the wavelength of maximal absorption of the retinal for the respective intermediate.

Figure 2.3.: Photocycle of Bacteriorhodopsin, adapted from [19].Upon light excitation, the retinal isomerizes from all-trans to 13-cis and the protein undergoes a series of transient intermediate states J, K, L, M, N and O before relaxing back to the ground state. Indices indicate the wavelength of maximal absorption in nanometers for the respective intermediate. Depicted times show the time constants, with which the respective intermediates rise and decay, respectively.

After light absorption, photoisomerization of the retinal from all-trans to 13-cis takes place, yielding via the ultrashort J intermediate the early photointermediate K of the photocycle of BR. This photoisomerization of the retinal belongs to the fastest known biological photoreactions. Relaxation of the configurational strain of the distorted 13-cis retinal in K leads then to the L intermediate [17]. These structural transitions around the retinal pave the way for the following proton transfer reactions (Figure 2.4). During M intermediate formation, the first proton transfer step takes place from the retinal Schiff base to the primary proton acceptor Asp85 (step 1). In order to allow vectorial proton transport, de- and reprotonation of the Schiff base have to take place from different sides of the membrane. Therefore, the accessibility of the Schiff base is switched towards Asp96 during the transition from M₁ to M₂. Simultaneously, proton release occurs to the

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extracellular side (step 2), whereas Asp85 stays protonated to the end of the photocycle.

The following N intermediate is characterized by the reprotonation of the Schiff base from Asp96 (step 3). Reprotonation of Asp96 (step 4) and following thermal relaxation of the retinal from 13-cis back to all-trans leads to the final O intermediate. Finally, Asp85 deprotonates (step 5) to Asp212 which directly transfers the proton to the proton release complex [19], consisting primarily of Glu194 and Glu204 (step 6). A second accessibility switch of the Schiff base from the cytoplasmic side back to the extracellular side completes the photocycle and results in the restoration of the initial ground state. The absorption maximum of the retinal strongly shifts during the photocycle. The strongest shift is observed during the M intermediate. Thereby the deprotonated Schiff base leads to a blue-shifted absorption maximum of the retinal at 412 nm.

Figure 2.4.: Corresponding amino acids involved in proton transfer steps (steps 1 to 6) of Bacteriorhodopsin, adapted from [19]. The first proton transfer step occurs from the protonated Schiff base (Schiff base linkage between retinal and Lys216) to the primary proton acceptor Asp85. Subsequent steps are discussed in the text.

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2.2.3. The purple membrane

BR is found as a trimer in the purple membrane ofHalobacterium salinarium which in turn is packed in a hexagonal arrangement as shown in Figure 2.5. The purple membrane consists of 75% BR by weight which is the only membrane protein present and the remainder is a mixture of lipids. It contains amongst others the polar lipids phosphatidyl- glycerophosphate methyl ester (24%), glycolipid sulphate (30%), phosphatidylglycerol (10%), archaeal glycocardiolipin (10%) and the neutral lipid squalene (20%) which is a precursor for steroids. Cholesterol is not present in the purple membrane. Figure 2.6 gives an overview of the lipids detected in a lipid extract of the purple membrane by NMR analyses [20]. The purple membrane has a high negative charge because of the presence of many phosphate and sulphate lipid headgroups [21]. The protein-lipid stoichiometry reveals 30 lipids per BR trimer [21, 22] with 6 lipids located in the center of the trimer and the other 24 surrounding the trimer. These archaeal lipids feature rather phytanyl chains than fatty alkyl chains and the interfaces to the glycerol backbone are by ether rather than ester bonds [21].

Figure 2.5.: Bacteriorhodopsin trimers packed in a hexagonal arrangement in the purple membrane, taken from [23]. BR monomers are painted in green, blue and red, lipids enclosed in the center of the trimers in yellow and surrounding lipids in gray colors. The lipid-protein stoichiometry is 10 lipids per BR monomer.

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Figure 2.6.: Lipid composition of the purple membrane, taken from [20].

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2.3. Proteorhodopsin

2.3.1. Overview and history of Proteorhodopsin

Proteorhodopsin (PR) is like BR a photoreceptor membrane protein and acts also as a light-driven proton pump. It was discovered in the year 2000 through genomic analyses of sea water at Monterey Bay in California [24]. The discovery came as a surprise, since it was the first evidence of a bacterial retinal-based photoreceptor [10]. The former known photoreceptors such as e.g. Bacteriorhodopsin and Halorhodopsin were found in archaea.

The name Proteorhodopsin was given due to the first identification in the subgroup SAR86 ofγ-proteobacteria which is ubiquitous and abundant in surface ocean samples [25]. In the meantime, many more PR-like sequences have been identified and it is known nowadays that Proteorhodopsins are the most abundant retinal-based photoreceptors on earth. They are even considered to be the most abundant phototrophic system on this planet [26]. Samples of PR have been also found in freshwater as well as sea ice and in all three kingdoms of life, bacteria, archaea and eukaryotes [10] and even in viruses [27]. PRs can be classified into two major classes, a green absorbing form (λ_max = 525 nm) which has been studied in greater detail and which was used in this project and a blue absorbing form (λ_max = 490 nm) [28]. A single mutation of an amino acid close to the retinal L105Q is responsible for the observed blue-shift of the wavelength of maximal absorption.

This spectral tuning is a specific property observed for PR. The blue-absorbing form is found in the ocean at greater depths, whereas the green-absorbing form is predominantly found closer to the surface of the ocean in the photic zone. Blue light can penetrate deeper in the ocean which explains the biological relevance of the amino acid mutation.

PR expressed inE.coli has been shown to enable photophosphorylation [29] and drive flagellar motility under anaerobic conditions [30]. Figure 2.7 shows an illustration of the fundamental biological processes of PR in the cell membrane [31]. Through light excitation, PR pumps protons to the extracellular side of the cell membrane at alkaline pH, creating a proton motive force. This proton gradient can be used to drive flagellar motility or is used by the ATP-synthase, another membrane protein, to generate ATP.

For acidic pH, PR changes the vectoriality of proton pumping described in subsection 2.3.3.

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Figure 2.7.: Artist view of PR in the membrane, taken from [31]. Through light excitation, PR (1) pumps protons across the membrane and produces a proton gradient (2).

The proton gradient is used by the ATP synthase (3) which phosphorylates ADP (4) to the universal energy currency ATP (5).

2.3.2. Structure of Proteorhodopsin

In 2011, the first 3D backbone structure of PR in diC7PC micelles has been solved with solution-NMR spectroscopy [32] and is available under the protein data bank ID 2L6X.

Figure 2.8 shows a structural model based on these findings. The seven transmembrane α-helices arrange in a similar way like for Bacteriorhodopsin or other rhodopsins and are connected with short loops. No antiparallelβ-sheet which is formed for Bacteriorhodopsin (compare Figure 2.2), is observed for Proteorhodopsin.

The oligomerization state of Proteorhodopsin is strongly dependent on the environment [33]. In a lipid bilayer or proteoliposomes, the protein oligomerizes mainly in form of a pentamer or hexamer which in turn assembles in form of donut-shaped complexes revealed with atomic-force microscopy imaging [34]. In OG or Triton-X 100 detergent, PR is monomeric, whereas in DDM detergent it assembles like in a lipid environment as a pentamer [33]. It is speculated, whether the radial arrangement of the pentamers in form of donut-shaped complexes is beneficial for harvesting light in the sea in order to achieve a higher quantum yield [10]. Comparing BR with PR, several differences can be observed. Very importantly, PR in contrast to BR has a Histidine H75 in transmembrane helix B close to the primary proton acceptor D97. H75 forms a pH-dependent hydrogen

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2.3. Proteorhodopsin

Figure 2.8.: 3D structure of Proteorhodopsin according to protein data bank ID: 2L6X.

Secondary structure elements are shown in form of a cartoon withα-helices in red and loops connecting the α-helices in green. Highlighted are the retinal depicted in cyan, primary proton acceptor D97 in blue, primary proton donor E108 in gray and K231 in orange.

bond with D97 explaining the unusually high pK_a (~7.5) of the primary proton acceptor for PR [35]. Further differences between BR and PR are the above mentioned spectral tuning properties for PR (L105) as well as the absence of a proton release group typical for BR [10]. Figure 2.9 shows important amino acid residues of PR close to the retinal.

K231 (orange) is covalently bound to the retinal (cyan) via a Schiff base linkage. For alkaline pH, D97 (primary proton acceptor) accepts the proton from the Schiff base and E108 (primary proton donor) subsequently reprotonates the Schiff base, whereby PR follows a BR-like photocycle, pumping one proton to the extracellular side. D227 and D97 act both as complex counter-ion to the protonated Schiff base.

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retinal

D97

E108 L105

H75 D227

K231

His-Asp Cluster

Figure 2.9.: Important amino acids close to retinal for PR and highlighted His-Asp cluster.

Histidines close to the primary proton acceptor are not present in most other retinal proteins. For PR, His75 undergoes a pH-dependent hydrogen bonding with the primary proton acceptor Asp97 yielding the unusual high pK_a of Asp97.

2.3.3. PH-dependent vectoriality of proton transport in Proteorhodopsin

As a special feature, the direction of the proton transfer for PR is dependent on the pH in contrast to BR (Figure 2.10). For alkaline pH values, PR pumps protons outwardly in a similar fashion like BR, whereas for acidic pH values PR turns into an inwardly pumping proton pump. This variability could be demonstrated using electrophysiological measurements [36]. Thereby, photocurrent measurements of proteoliposomes adhering onto black lipid membranes were performed. The vectoriality change of proton pumping was questioned first by other research groups [37], not observing proton pumping at low pH in oriented membrane fragments or in proteoliposomes. Inverse proton pumping at low pH was then reconfirmed inXenopus oocytes [38] and with fluorescence spectroscopy [39]. The inward current flow at acidic pH is estimated to be around one magnitude smaller compared to the outward flow at alkaline pH. The molecular mechanism for the inverse proton pumping at acidic pH is not yet understood.

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2.4. Properties of lipid membranes

alkaline pH acidic pH

cytoplasmic extracellular H⁺

H⁺

Figure 2.10.: pH-dependent proton pumping vectoriality of PR. For alkaline pH, PR pumps protons like BR from the cytoplasmic side to the extracellular side. For acidic pH, an inverted proton transfer direction is observed from the extracellular side to the cytoplasmic side for PR.

2.4. Properties of lipid membranes

Biological membranes play an important role for a huge range of biological processes in the cell such as transport and signaling. They act as selective permeable barriers and lead to compartmentalization of the cell. A thorough understanding of the molecular organization of these membranes is crucial for a better understanding of these important biological processes.

2.4.1. Structure of lipids

Lipids are the building blocks of a lipid bilayer which is the most fundamental structure of cell membranes [40]. The phospholipids are a major component of all cell membranes.

They possess an amphiphilic structure having a polar phosphate headgroup and usually two fatty acid tails. Headgroup and fatty acid tails are linked together by a glycerol molecule. The charge of the lipid headgroups can vary as well as the number of carbon atoms for the fatty acid tails. When all the bonds between the carbon atoms in the tails are single bonds, the lipid is classified as "saturated". Lipids which contain one or more double bond(s) between the carbon atoms of the hydrophobic chain are termed

"unsaturated" or("polyunsaturated"). Figure 2.11 displays the structure of 1,2-dioleoyl-sn- glycero-3-phosphocholine abbreviated as 18:1 (∆ 9-cis) or DOPC. The first abbreviation

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is the easiest way of classifying lipids. Thereby, the first number before the colon indicates the number of carbon atoms in the hydrophobic chain, while the second number after the colon indicates the number of double bonds found in the chain [41]. DOPC has 18 carbon atoms in the fatty acid chain and onecis double bond at carbon position 9 counted from the first carbon of the C=O ester group which lies at the interface between headgroup and lipid tail. The position of the double bond is therefore denoted by the ∆ nomenclature.

Apart from phospholipids, sterols such as cholesterol play an important role in the fatty acid composition of cell membranes. Cholesterol is a rigid molecule having a steroid ring structure and a polar head. It plays an outstanding role in the maintenance and regulation of membrane structural integrity and fluidity of animal plasma membranes.

N O P O O O O

O H O

O

Figure 2.11.: Structure of 1,2-dioleoyl-sn-glycero-3-phosphocholine abbreviated as 18:1 (∆

9-cis) or DOPC. The polar phosphate headgroup is highlighted in blue and the two hydrophobic lipid tails in gray, both containing one cis double bond at carbon position 9.

2.4.2. Lipid phase transitions

Soft matter like lipid bilayers undergoes phase transitions just like basically any other kind of matter [41]. As a very simple example, water in form of ice melts to liquid water and finally boils to water vapour with increasing temperature. Likewise lipid bilayers can be in a more ordered gel phase and undergo a phase transition to a more fluid liquid crystalline phase with increasing temperature. The temperature at which the transition occurs is defined as the phase transition temperature T_m. Phosphatidylcholines (PC) are the most abundant class of lipids in mammalian membranes. Figure 2.12 schematically shows the phase transitions of this lipid class with increasing temperature.

Due to the large area requirement of the bulky polar headgroup of phosphatidylcholines [43], the fatty acid chains are tilted 30^◦ with respect to the bilayer normal in the gel

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Gel phase Ripple phase Liquid crystalline phase

Temperature

Figure 2.12.: Lipid phases of phosphatidylcholines, adapted from [42]. With increasing temperature, the lipids undergo a phase transition from the ordered gel phase to the more disordered liquid crystalline phase via a ripple phase. The bilayer thickness substantially decreases in the liquid crystalline phase, due to the higher disorder of the hydrocarbon chains.

phase and show a pretransition to the ripple phase. With further increasing temperature, the main transition to the liquid crystalline phase can then be observed, in which the fatty acid chains are no longer tilted, since the bulky headgroups are further apart from each other [44]. The lipid fatty acid chains convert from a rigid extended all-trans conformation in the gel phase to the more flexible disordered liquid crystalline phase characterized by the presence of gauche conformations [45].These gauche bonds result in a kink of the hydrocarbon chain being responsible for the higher disorder in the liquid crystalline phase (Figure 2.13). Hence, the hydrophobic bilayer thickness is reduced in the liquid crystalline phase. The phase transition temperature is determined by a competition of the entropically favoredgauche conformation of the hydrocarbon chains and the attractiveVan der Waals interactions between neighboring chains [44]. The fatty acid chain length is therefore proportional to the phase transition temperature, since longer chain lengths result in increased van der Waals interactions leading to a higher phase transition temperature. The introduction of a cis double bond drastically reduces the phase transition temperature having a maximal effect when the cis double bond is located in the middle of the chain [44]. Thecis double bond induces a kink in the fatty acid chain which hinders an efficient molecular packing required for the gel phase.

Infrared spectroscopy is one of the powerful tools to monitor the phase transition of lipids.

Thereby, thetrans-gaucheisomerization from the gel phase to the liquid crystalline phase can be monitored in form of a frequency shift of the CH₂ stretching vibrations of the hydrocarbon chains [46, 47].

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Potential energy

0 60 120 180 240 300 360 Dihedral angle

g t g ⁺ ^-

CH₃

H H

CH₃

H H

CH₃

H H

R

H H

R

H

H R

H H

...ttt...

...g⁺tg^-...

cis-double bond g⁺

g^- t

Figure 2.13.: Upper panel: Potential energy for rotation around a carbon-carbon bond.

Trans-conformation t is the lowest energy state with a dihedral angle of 180^◦. Further local minima aregauche-conformations g⁺ and g^- with dihedral angles of 60^◦ and 300^◦. Middle panel: Newmanprojections of the respective conformations. Lower panel: Examples of lipid chain configurations (all-trans ttt, first-order kink g⁺tg^- and monounsaturated lipid tail with double cis-bond).

Adapted from [44].

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2.4.3. Artificial model membranes

The lipid environment surrounding a membrane protein can interact with the protein and has a high impact on its function. Investigation of this influence requires profound knowledge about the physical properties of the respective lipids surrounding the protein.

In vivo, membranes are typically built up by many different lipids with distinct physical properties, making it tremendously difficult to analyze the effect of the membrane on the function of the protein. Therefore, in vitro experiments usually try to reduce the complexity by using membrane mimetic systems with only one sort of lipid or a few lipids with precise physical properties in order to obtain a better understanding how these properties can affect the functionality of membrane proteins. Highly applied membrane mimetic systems are micelles or liposomes (Figure 2.14). More recently lipid nanodiscs have also become a membrane mimetic tool [48]. Membrane scaffold proteins (MSP) are used in this case to stabilize the disk-shaped lipid bilayers. By varying the length of the MSP, precise control of the oligomerization state of the membrane protein of interest can be achieved.

Liposome Micelles

Hydrophilic head

Hydrophobic tails

Figure 2.14.: Membrane mimetic systems. Liposomes are spherical vesicles built up by a lipid bilayer enclosing an aqueous environment. Micelles are closed lipid monolayers which start to assemble at the critical micelle concentration (CMC). Both systems can assemble spontaneously. These systems are largely used to imitate a specific lipid environment with precise lipid physical properties for membrane proteins.

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2.5. Protein-lipid interactions

2.5.1. Influence of the membrane on the function of the protein

Protein-lipid interactions are key determinants for the comprehension of the function of membranes. SingerandNicholsonfirst characterised the organization of membranes in 1972 formulating the standard fluid-mosaic model [49]. Thereby, the biological membrane is described as a two-dimensional solution consisting of oriented lipids and globular proteins. According to the model, lipids and proteins can diffuse freely in lateral direction.

This mobility is required for biological functions to take place which rely on molecular motions. The lipids thereby only provide the necessary fluid environment. In the last decades however, a stronger coupling between the lipids and proteins has been observed and lipids are in the mean time thought to highly influence the function of membrane proteins [21, 50–53]. Erich Sackmannproposed in 1984 that the function of integral membrane proteins is influenced by lipid bilayer properties [54]. First evidence for the impact of the lipid membrane on the function of membrane proteins came for the Ca²⁺- ATPase [55, 56] and rhodopsin [57–59]. The fatty acid chain length highly influences the activity of the Ca²⁺-ATPase [56]. The activity was shown to be highest for a carbon chain length of 18 and was greatly reduced with shorter or longer chain length. The bilayer thickness can therefore play an important role for protein functionality. Hydrophobic mismatch occurs when the hydrophobic part of the membrane protein does not match the hydrophobic chain length of the lipids. Too long hydrophobic lipid chains result in a necessary stretching of the protein whereas too short hydrophobic chains lead to a compression of the protein. This phenomenon of hydrophobic mismatch is illustrated in Figure 2.15.

Figure 2.15.: Schematic illustration of hydrophobic mismatch, taken from [50]. For large lipid chain lengths, the α-helices of the membrane protein need to stretch, while they need to compress for short lipid chain lengths.

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2.5. Protein-lipid interactions

The ATPase activity of the Ca²⁺-ATPase was also monitored at temperatures below and above the phase transition temperature of different lipids [55, 60]. At temperatures below the phase transition temperature, the activity of the Ca²⁺-ATPase strongly declined. The low activity in the gel phase is explained by a slow rate of formation of the phosphorylated intermediate of the Ca²⁺-ATPase [61]. The increased packing order of the lipids in the gel phase can be assumed to hinder conformational changes of the enzyme, but also the increased bilayer thickness in the gel phase has to be taken into account. For rhodopsin, the equilibrium between the photointermediates metarhodopsin I and metarhodopsin II depends also on the length of the lipid hydrocarbon chains. An increase of the fatty alkyl chain length leads to a favorable formation of metarhodopsin II [62]. Rhodopsin is located in the retina of the eye in the rod outer segments which consist of disk membranes surrounded by a plasma membrane of the rod cells. Rhodopsin is in both membranes the most abundant protein. The lipid bilayer composition of the rod outer segments is critical for visual transduction [63]. The amount of cholesterol is high in the plasma membrane and also high for newly synthesized disk membranes. However, during the apical displacement of the disk membranes, the cholesterol amount decreases and the amount of unsaturated phospholipids increases. The high cholesterol content in the plasma membrane inhibits the activation of rhodopsin, whereas the lower concentration of cholesterol in the apically displaced disk membranes allows activation of the photoreceptor [64]. Cholesterol increases the order of the lipid bilayer and might therefore hinder the conformational changes of rhodopsin which are necessary for activation. Moreover it binds also directly to rhodopsin which might also influence protein activation. These two options can serve as an example to generalize a fundamental question in the field of protein lipid interactions: Whether chemical specificity or rather bulk lipid biophysical properties give rise to the impact of the lipids on the functionality of the membrane proteins [51].

2.5.2. Retinal protein-membrane interactions

Retinal proteins such as rhodopsin or microbial rhodopsins including Bacteriorhodopsin and Proteorhodopsin are excellent candidates to study general aspects of protein-lipid interactions since their function can be very easily controlled using light. FTIR spectroscopy significantly contributed in identifying a specific interaction of a lipid molecule with rhodopsin [65]. The influence of lipid bilayer properties on photocycle dynamics

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of Bacteriorhodopsin and Proteorhodopsin has been so far most often investigated with transient UV-vis spectroscopy. Thereby, the photocycle of the protein gets triggered with a short laser flash and the absorption changes are probed with continuous monochromatic light in the visible region. Rise and decay of the M Intermediate of Bacteriorhodopsin can for example be monitored by the absorbance change at 412 nm. Many different membrane mimetic systems ranging from micelles over liposomes to nanodiscs were used for these investigations. Reconstitution of Bacteriorhodopsin into liposomes with various lipid headgroup charges was shown to have an impact on the decay kinetics of the M intermediate [66]. Also in lipid nanodiscs, the dynamics of Bacteriorhodopsin can be influenced for different ratios of neutral and negatively charged lipids [67]. For Proteorhodopsin, photocycle dynamics in nanodiscs are twice as long as compared to the dynamics in liposomes with identical lipid composition [33]. Dynamics were found to differ also strongly in detergent, mixed micelles (lipid+detergent) and liposomes with Proteorhodopsin having fastest kinetics in liposomes, intermediate in mixed micelles and slowest in detergent. The effect of various lipid compositions in mixed micelles were also investigated [68]. Both the membrane mimetic system and the lipid composition affect the function of the protein. In nanodiscs, the dynamics of Proteorhodopsin can be tuned through the variation of the discoidal diameter of the nanodiscs. Furthermore, longer unsaturated alkyl chains (POPC) reveal a faster photocycle of Proteorhodopsin as compared to shorter saturated chains (DMPC) [69]. Similarly, for Bacteriorhodopsin in nanodiscs, ground-state recovery in DOPC nanodiscs is faster than in DMPC nanodiscs.

Apart from the bilayer thickness and the charge of the lipid headgroups, the phase transition temperature of the lipids is speculated to affect the observed altered kinetics [67].

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3. Motivation and aim of the project

A better understanding of the interaction of proteins with membranes is of fundamental importance in membrane protein research. The current status of research for protein-lipid interactions is summarized in section 2.5. By now there is a general consensus that the membrane lipid composition can have an influence on the function of membrane proteins and plays a more active role than originally thought. However, analyzing these interactions while the membrane protein performs its task remains challenging. Ideally, both the dynamics of the protein and the dynamics of the surrounding lipids should be monitored while the protein is in an active state. Photoreceptors are very good candidates to investigate the function of membrane proteins, since their function can be easily triggered with light. So far investigations of the lipid environment on the function of photoreceptors were mainly carried out with UV-vis transient absorption spectroscopy. The protein is triggered thereby with a short nanosecond laser pulse and the absorption changes in the visible region of the protein are monitored for different lipid environments. In this project, a new approach was chosen to investigate the interactions between photoreceptors and surrounding lipids by monitoring the absorption changes in the infrared spectral region. This approach offers several advantages: due to the high sensitivity of FTIR spectroscopy, the protonation dynamics of single amino acids can be directly followed. Conformational dynamics of the protein in form of movements of the polypeptide backbone become accessible in the amide I and amide II region of the FTIR spectrum. While the membrane protein performs its function, dynamics of the surrounding lipids absorbing in the infrared can in principle be monitored simultaneously.

In order to realize this approach, a first objective was to establish the step-scan approach for time-resolved FTIR measurements. Furthermore, expression and purification of the membrane protein Bacteriorhodopsin and reconstitution of Bacteriorhodopsin and Proteorhodopsin into different artificial membrane environments. A systematic analysis of the influence of lipid physical properties such as lipid chain length, degree of saturation and charge of lipid headgroups on the function of the photoreceptors was then the

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3. Motivation and aim of the project

ultimate goal. Moreover, the conformational changes of the protein and the dynamics of the surrounding lipids should be explored simultaneously as well as the pH-dependent protonation dynamics of Proteorhodopsin for different membrane environments.

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4. Experimental methods

4.1. Protein expression

4.1.1. Cultivation of Halobacterium salinarium

Bacteriorhodopsin was isolated and purified several times during this project according to the protocol of Oesterhelt and Stoeckenius[70]. Halobacterium salinarium cells of the retinal deficient strain JW5 were kindly provided from Clemens Glaubitz, Goethe University Frankfurt. Halobacterium salinarium has a very slow generation time of 8 hours [70] in contrast to e.g. Escherichia coli which has a generation time of approximately 20 minutes. Table 4.1 lists the culture medium used for cultivation of Halobacterium salinarium, containing a very high concentration of sodium chloride which accounts for the halophilic properties ofHalobacterium salinarium.

Table 4.1.: Culture medium (1 Litre) used for the cultivation of Halobacterium salinarium cells.

Sodium chloride 250 g Oxoid Bacterial Peptone 10 g Magnesium sulfate 20 g Sodium(III) citrate 3 g Potassium chloride 2 g

At these high salt concentrations autoclavation of the culture medium is no longer absolutely required but was still performed. Alternatively the culture medium can be filtered using sterile millipore filters of 0.45µm pore size. Cultivation of the halobacterium cells takes several weeks, starting with a preculture. Thereby, one to two culture beads were used for inoculation in around 20 to 30 mL of culture medium shaken at 180 rpm at 37^◦C. After approximately one week, grown cells were transferred in 150 mL culture medium and after successful growth for about one more week, cells were splitted and cultivated in 10 L culture medium. Thereby, either ten 2 L flasks containing each 1 L

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4. Experimental methods

culture medium were shaken in an incubator at 150 rpm and 37 ^◦C or a fermentor was used which is preferable for efficient growth of the cells. Growth of the cells was optically monitored by measuring the absorption at 600 nm. 6 mg all-trans retinal (Sigma-Aldrich, Germany) dissolved in 2 mL ethanol was added when growth entered the stationary phase. The dissolved retinal and the cultured cells have to be kept at all times in the dark to prevent denaturation of the retinal. The retinal deficient JW5 cells incorporate the retinal into Bacteriorhodopsin in the purple membrane.

4.1.2. Isolation and purification of the purple membrane

Optimally an optical density of OD₆₀₀ in between 1.5 and 2 is reached, when cells can be harvested via centrifugation at 7000 rpm for 15 minutes at 4 ^◦C. The obtained purple pellet was resuspended in basal salt which is essentially the culture medium without Oxoid bacterial peptone. DNase was added to prevent excessive viscosity from liberated DNA and the cells were dialyzed overnight against 2 L of 0.1 M NaCl which separates cell fragments from the membranes. The obtained lysate was washed several times with 0.1 M NaCl and ultimately with H₂O by centrifugation at 20000 rpm for 40 minutes at 4^◦C until the supernatant became nearly colorless. Halobacterium salinarium does not possess any intracellular membranes and the cell membrane consists of a purple membrane and a red membrane. The color of the red membrane arises from the carotenoid bacterioruberin which protects the cells against DNA damage from UV radiation. In order to separate the purple membrane from the red membrane, sucrose density gradient centrifugation was applied. Thereby, a linear sucrose gradient between 30% and 50% was generated using a gradient mixer and the final sediment was layered on top of the sucrose gradient.

Centrifugation was performed at 17000 rpm for 17 hours at 15^◦C giving rise to a purple band and a red band as illustrated in Figure 4.1. The purple band representing the purified purple membrane including Bacteriorhodopsin was collected and washed several times with H₂O by centrifugation at 45000 rpm for 1 hour at 4 ^◦C and resuspended eventually in a couple of mL of H₂O and stored at 4 ^◦C. Figure 4.2 displays an absorption spectrum of the purified BR in purple membrane sample, revealing a high degree of purity of the isolated purple membrane and the absorption maximum of BR located at 568 nm.

The static FTIR absorption spectrum of purified BR in purple membrane reveals the exceptional high wavenumber of the amide I band at 1659 cm^-1 (Figure 4.3) for BR trimers reported in the literature [71]. Bands of the purple membrane lipid archaeol

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4.1. Protein expression

chains are located at 2959 cm^-1, 2929 cm^-1 and 2872 cm^-1.

Figure 4.1.: Separation of the red and purple membrane via sucrose density gradient centrifugation.

250 300 350 400 450 500 550 600 650 700 0.0

0.1 0.2 0.3 0.4 0.5

568

absorbance/OD

wavelength / nm 280

Figure 4.2.: UV-vis absorption spectrum from 250 to 700 nm of purified BR in purple membrane sample. The maximum at 280 nm corresponds to the aromatic amino acid absorption and the maximum at 568 nm to the retinal absorption. The ratio Abs₂₈₀/Abs₅₆₈= 2.1 indicates a high degree of purity of the isolated purple membrane [70].

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3500 3000 2500 2000 1500 1000

0.0 0.1 0.2 0.3 0.4

1546 1659

2959

2872

A/OD

wavenumber / cm -1 2929

Figure 4.3.: Static FTIR absorption spectrum from 1000 cm^-1 to 3500 cm^-1 of purified BR in form of a dry purple membrane film.

4.2. Reconstitution into proteoliposomes

Liposomes form spontaneously in an aqueous environment due to the amphiphilic nature of phospholipids. Self-assembly occurs in form of a double layer, whereby the phospholipid polar headgroups are exposed to the aqueous environment and the hydrophobic tails remain shielded from contact with water. In general multiple stacked layers are formed and techniques such as sonication and extrusion can be used to generate unilamellar liposomes.

Liposomes serve as an excellent tool in form of a biomimetic artificial membrane system.

Thereby, a minimal model system reduces the complexity of biological membranes.

Reconstitution of membrane proteins into liposomes resulted in a major breakthrough for the understanding of the function of membrane proteins, such as transport or signaling processes across the cell membrane [72, 73]. Figure 4.4 schematically illustrates the detergent-mediated reconstitution of a membrane protein into liposomes. Due to the hydrophobicity of membrane proteins, either lipids or detergent have to protect the protein from the aqueous environment. Solubilization of a membrane protein from a native membrane therefore involves the addition of detergent which yields the membrane protein of interest surrounded by a micellar-like detergent environment. Mixing with preformed unilamellar liposomes and detergent removal via dialysis or addition of polystyrene

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4.2. Reconstitution into proteoliposomes

beads leads then to the membrane protein incorporated into liposomes which is called a proteoliposome. The following three subsections describe the employed detergent- mediated reconstitution process in detail and Figure 4.5 shows the structures of the lipids used within this work (all purchased from Avanti-Polar Lipids, USA).

Figure 4.4.: Schematic representation of membrane protein reconstitution into proteoliposomes, taken from [74]. The membrane protein of interest is solubilized first with a detergent out of the native membrane and purified. Reconstitution into proteoliposomes can be executed in several ways, e.g. through addition of detergent-adsorbing polystyrene beads.

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O N P O O O O

O H O

O

P O O O O O

O H O

O

OH OH

HO

H H H

O N P O O O O

O H O

O

O N P O O O O

O H O

O

O N P O O O O

O H O

O a)

b)

c)

d)

e)

f)

g)

N O

O H O

O

O N P O O O O

O H O

O D2 D

2 D

2 D 2 D

2 D

2

D₂ D₂ D2

D₂ D₂ D₂ D₂ D₂ D₂ D2 D

2 D

2

D₂ D₂ D₂ D₂ D₂ D₂ D₂ D₂ D₃

D₃ D2

h)

Figure 4.5.: Lipids used for reconstitution a) DSPC b) DOPC c) DMPC d) DMoPC e) DMPG f) DMTAP g) 18:0 PC-d70 (deuterated lipid chains) h) Cholesterol.

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4.2. Reconstitution into proteoliposomes

4.2.1. Preparation of liposomes

10 mg of desired lipid were dissolved in 400µL chloroform (CHCl₃) and dried for several hours under a stream of nitrogen. The lipid film was then rehydrated in 2 mL Tris-HCl, pH 7.4 to achieve a concentration of 5 mg/mL. The sample was shaken for 2 hours and sonicated several times for 10 minutes, resulting in multilamellar liposomes. Extruding the sample through a membrane with 100 nm pore size (Avanti-Polar Lipids, USA) yielded unilamellar liposomes of defined size which could be stored at 4 ^◦C.

4.2.2. Solubilization of Bacteriorhodopsin and Proteorhodopsin

Solubilization of BR in purple membrane (2 mg/mL) was performed using 0.8% Triton X- 100 detergent and 50 mM phosphate buffer, pH 7.0. After addition of detergent, the sample had to be shielded from light radiation and kept in the dark at all times since solubilized BR is susceptible to light. After 20 minutes of sonification and incubation overnight, the sample was centrifuged at 100000 g at 4 ^◦C for 1 hour. The purple supernatant containing solubilized BR was collected and ideally the remaining membrane pellet was no longer purple but white for an efficient solubilization. Typically, the incubation time period before the centrifugation lasted approximately 24 h, but increasing the incubation time to several days can improve the efficiency of the solubilization. Solubilized BR can be spectrally confirmed by a blue shift of the absorption maximum to 550 nm. Solubilized PR samples were received from Clemens Glaubitz laboratory, Goethe University Frankfurt. PR was solubilized in 0.05% DDM, 500 mM imidazole, 50 mM MES and 300 mM NaCl at pH 7.0.

4.2.3. Preparation of proteoliposomes

A lipid to protein ratio of 2:1 (w/w) was chosen for reconstitution which corresponds e.g.

for DSPC to a molar ratio of 68 lipids per protein. Solubilized protein was added to the preformed liposomes and incubated for at least 30 minutes. After addition of 80 mg/mL detergent adsorbing beads (SM2 Bio-Beads, Bio-Rad), the sample was incubated over night. During the next day, Bio-Beads were subsituted twice with new Bio-Beads for 2 hours each time and finally removed. Centrifugation at 100000 g at 4^◦C for 1 hour led to a transparent supernatant and purple pellet which contained the desired proteoliposomes.

Therefore, the supernatant was discarded and the proteoliposome pellet was resuspended

Protein-membrane interactions investigated with time-resolved FTIR spectroscopy