X-ray crystallographic analysis of three membrane proteins:
The nicotinic acetylcholine receptor from Torpedo californica, Omp85 and
TtoA from Thermus thermophilus HB27
Dissertation
submitted to the
Department of Biology, University of Konstanz, Germany
for the degree of
Doctor of Natural Sciences
presented by
Dipl. Biol. Alexander Brosig
First referee: Prof. Wolfram Welte Second referee: Prof. Winfried Boos
Date of oral examination: 15. April 2009
Konstanzer Online-Publikations-System (KOPS) URN: http://nbn-resolving.de/urn:nbn:de:bsz:352-opus-82348
URL: http://kops.ub.uni-konstanz.de/volltexte/2009/8234/
Contents
Summary xi
Zusammenfassung xiii
1 General introduction 1
2 Methods of X-Ray structure determination 7
2.1 Crystallization of proteins . . . 8
2.2 Protein crystals . . . 11
2.3 X-ray diffraction by crystals . . . 12
2.4 Structure determination by X-ray crystallography . . . 15
2.4.1 The electron density function . . . 15
2.4.2 The phase problem . . . 16
2.4.3 Model building and refinement . . . 21
3 Purification and crystallization of the nicotinic acetylcholine receptor from Torpedo californica, characterization of microcrystals 25 3.1 Introduction . . . 25
3.2 Materials and methods . . . 29
3.2.1 Solubilization and purification of nAChRs from nAChR-enriched mem- branes fromT. californica . . . 29
3.2.2 Crystallization . . . 30
3.2.3 Data collection and data analysis . . . 31
3.2.4 MALDI analysis of microcrystals . . . 31
3.2.5 Two-dimensional polyacrylamide gel electrophoresis . . . 32
3.3 Results . . . 33
3.3.1 Characterization of microcrystals: X-ray crystallographic analysis . . . 33
3.3.2 Characterization of microcrystals: MALDI mass spectrometry . . . 35
3.3.3 Influence of detergents on homogeneity of nAChR-preparations . . . . 38
3.3.4 Crystallization experiments with solutions of purified nAChR . . . 41
3.4 Discussion . . . 44
4 Purification, crystallization and in-vitro characterization of TtOmp85 from Thermus thermophilusHB27 49 4.1 Introduction . . . 49
4.2 Materials and methods . . . 52
4.2.1 Bacterial strains and growth conditions . . . 52
4.2.2 Purification of native TtOmp85 . . . 52
4.2.3 Protein analysis . . . 53
4.2.4 Crystallization . . . 53
4.2.5 Data collection and data analysis . . . 54
4.2.6 In-vitrocharacterization of purified TtOmp85 . . . 54
4.3 Results . . . 55
4.3.1 Overexpression of TtOmp85 inT. thermophilusand purification in na- tive form from OM preparations . . . 55
4.3.2 Oligomeric form of purified TtOmp85 in the presence of detergents . . 56
4.3.3 Crystallization of purified TtOmp85 and crystallographic analysis . . . 58
4.3.4 In-vitrocharacterization of purified TtOmp85: TEM, FFEM and single channel conductance measurements . . . 60
4.4 Discussion . . . 63
5 Crystal structure of TtoA, a major outer membrane protein fromThermus thermophilusHB27 67 5.1 Introduction . . . 67
5.2 Materials and methods . . . 70
5.2.1 Cell fractionation and OM preparation . . . 70
5.2.2 Expression and purification of His-TtoA . . . 70
5.2.3 Crystallization and data collection . . . 71
5.2.4 Heavy atom derivatization . . . 71
5.2.5 Selenomethionine labeling . . . 71
5.2.6 Phasing, structure solution, model building and refinement . . . 72
5.2.7 Bioinformatic analysis and figure preparation . . . 73
5.2.8 Data deposition . . . 73
5.2.9 In-silicoandin-vitrocharacterization, cloning of TtoA . . . 73
5.3 Results . . . 74
5.3.1 TtoA is an outer membrane protein . . . 74
5.3.2 Purification of His-tagged TtoA inT. thermophilusHB8 . . . 74
5.3.3 Crystallization and data collection . . . 76
5.3.4 Structure solution and model building of TtoA . . . 81
5.3.5 Overall structure of TtoA . . . 81
5.3.6 Transmembrane part . . . 82
5.3.7 Extracellular part . . . 83
5.3.8 Crystal contacts . . . 86
5.3.9 Homology with known structures of other eight strandedβ-barrel OMPs 87 5.3.10 Evidence for TtoA as assembly substrate of TtOmp85 . . . 89
5.4 Discussion . . . 91
References 95 Appendix 107 5.5 List of abbrevations . . . 107
5.6 Amino acid nomenclature . . . 110
List of publications 111
Danksagungen 113
List of Figures
1.1 Porin fromRhodobacter capsulatus . . . 2
1.2 Potassium channel KscA fromStreptomyces lividans . . . 3
1.3 Scheme of the fluid mosaic membrane model . . . 4
2.1 Diffraction pattern of a protein crystal . . . 8
2.2 Phase diagram of protein crystallization experiments . . . 9
2.3 Lattice planes and Bragg’s law . . . 14
2.4 The Ewald construction . . . 15
2.5 Harker construction for SIR . . . 20
2.6 Harker construction for MIR . . . 20
2.7 Harker construction for SIRAS . . . 22
2.8 Experimental electron density map . . . 23
3.1 Stereo representation of the AChBP protomer . . . 26
3.2 4.0 Å model of muscle-type nAChR fromTorpedo marmorata . . . 27
3.3 Microcrystals grown from preparations of detergent-solubilized nAChR-enriched membranes fromT. californica . . . 33
3.4 Images of diffraction patterns collected from a microcrystal . . . 34
3.5 MALDI-MS of mother liquor . . . 36
3.6 MALDI-MS of washed crystals . . . 37
3.7 MALDI-MS of a batch of crude crystals . . . 37
3.8 Analytic gelfiltration and twodimensional PAGE of nAChR-preparations with nonyl-β-D-glucoside . . . 38
3.9 Twodimensional PAGE of a nAChR-preparation with Cyclofos-7 . . . 39
3.10 Chemical structure ofβ-D-Dodecylmaltoside and Cymal-6 . . . 39
3.11 Preparative gelfiltration and twodimensional PAGE of nAChR-enriched mem- branes solubilized with Cymal-6 . . . 40
3.12 Analytic gelfiltration and twodimensional PAGE of purified monomeric and dimeric nAChR . . . 41
3.13 Needle shaped crystals obtained from protein purified in Cymal-6 . . . 42
3.14 Analytic gelfiltration and twodimensional PAGE of crystal-producing fraction of peak 3 . . . 43
4.1 Crystal structure of FhaC . . . 51
4.2 Expression, purification, secondary structure and stability of TtOmp85 . . . 56
4.3 Purified TtOmp85 is primarily monomeric . . . 57
4.4 Crystals of purified TtOmp85 . . . 58
4.5 Diffraction images of crystallized TtOmp85 . . . 59
4.6 Negative-stain TEM images of TtOmp85 reconstituted into lipid vesicles . . . . 61
4.7 FFEM of reconstituted TtOmp85 . . . 61
4.8 Single-channel conductance of TtOmp85 in black lipid films . . . 62
5.1 Model for OMP biogenesis inE.coli . . . 68
5.2 TtoA is a HB27 specific major OMβ-barrel protein . . . 75
5.3 Expression and localization of TtoA . . . 75
5.4 Crystals of native TtoA . . . 77
5.5 Diffraction images of native TtoA crystals . . . 77
5.6 Crystals of SeMet-labeled TtoA . . . 78
5.7 Diffraction images of SeMet-labeled TtoA crystals . . . 79
5.8 Overall structure of TtoA . . . 82
5.9 Lumen of theβ-barrel with the barrier for hydrophilic compounds . . . 83
5.10 Stereo representation of the extracellular part of TtoA . . . 84
5.11 Omit map of the divalent cation . . . 85
5.12 Charge distribution on the surface of TtoA . . . 85
5.13 Crystal contact of putative physiologic interest . . . 86
5.14 Structural alignment of TtoA . . . 88 5.15 Pull-down assay showing binding of native TtOmp85 to denatured His-TtoA . . 89
Summary
The work presented here was focused on the purification, crystallization and X-ray structure determination of three different transmembrane proteins - the nicotinic acetylcholine recep- tor (nAChR) fromTorpedo californica and two outer membrane proteins fromThermus ther- mophilusHB27, TtOmp85, and one of its potential substrates, TtoA.
The nAChR is a ligand-gated ion channel which belongs to the superfamily of “Cys-loop receptors” and plays a keyrole in synaptic transmission of neuronal signals. nAChRs consist of five homologous subunits composed as tissue-specific homo- or heteropentamers. So far, only a structure of 4.0 Å resolution derived from electron microscopy on tubular 2D-crystals is available for whole nAChR pentamers fromTorpedo marmorata. Albeit two high-resolution structures of prokaryotic homologues were published recently, a high-resolution structure of the nAChR is still missing. Microcrystals obtained from preparations of detergent-solubilized nAChR-enriched membranes from the electric organ of the pacific rayTorpedo californiawere reported already in 1988, but the properties of those crystals remained uncharacterized. In the work presented here, such crystals were examined by synchroton radiation and diffraction patterns typical for protein crystals were recorded. Diffraction of those crystals was limited to 20 Å resolution, possible spacegroup and unit-cell parameters were determined with the recorded data. Neither refinement of the crystallization conditions nor optimization of the pu- rification protocol allowed to grow crystals of solubilized nAChR suitable for high resolution X-ray structure determination.
Proteins belonging to the Omp85 family are involved in the assembly ofβ-barrel outer mem- brane proteins or in the translocation of proteins across the outer membrane in bacteria, mi- tochondria and chloroplasts. The thermophilic bacteriumThermus thermophilusrepresents an intermediate between Gram-positive and Gram-negative bacteria with an only roughly charac-
terized outer membrane. It encodes one Omp85-like protein, TtOmp85 which represents an ancestral type of this family. TtOmp85 was overexpressed inT. thermophilusstrain HB27 and natively purified from preparations of the outer membrane. In the presence of detergent, purified TtOmp85 existed mainly as a monomer, composed of two stable protease-resistant modules.
Detergent-solubilized TtOmp85 was used for crystallization experiments and microcrystals ap- peared under crystallization conditions including PEG400 as precipitant. Crystals diffracted to 5.0 Å resolution and belonged to spacegroup C222 with unit cell dimensions of a = 96.3 Å, b=416.1 Å,c=94.3 Å;α =β =γ =90°. Only a partial dataset could be collected because diffraction was anisotropic, depending on the orientation of the plate-shaped microcrystals in the X-ray beam, and crystals rapidly accumulated radiation damage during proceeding datacol- lection. The structure of TtOmp85 could not be solved with the available data. Preparations of detergent solubilized TtOmp85 were provided for accompanying studies in collaborating working groups to further characterize TtOmp85.
Only a few further outer membrane proteins from T. thermophilus are described. TtoA, a new β-barrel outer membrane protein, was identified by bioinformatic analysis of the genome sequence of strain HB27. Results of anin vitrobinding assay where TtoA bound to TtOmp85 suggest that insertion of proteins in the outer membrane inT. thermophilusmight be similar to the mechanisms found in modern Gram-negative bacteria. TtoA was successfully crystallized and its structure was determined to a resolution of 2.8 Å, representing the first crystal structure of an outer membrane protein from a thermophilic bacterium. Crystals belonged to space group P3121 with unit cell parameters a = b = 166.7 Å, c = 97.5 Å; α = β = 90°, γ = 120°.
TtoA consists of an eight-strandedβ-barrel with a large extracellular part to which a divalent cation is bound. A five-stranded extracellularβ-sheet protrudes out of the membrane-embedded transmembrane barrel and is stabilized by a disulfide bridge. The edge of this β-sheet forms crystal contacts with neighbouring TtoA molecules that could mimic physiologic interactions with other proteins.
Zusammenfassung
Die hier vorgestellte Arbeit konzentrierte sich auf die Reinigung, Kristallisation und Rönt- genstrukturanalyse dreier unterschiedlicher Transmembranproteine - des nikotinischen Acetyl- cholinrezeptors (naChR) ausTorpedo californicaund zweier Außenmembranproteine ausTher- mus thermophilus, TtOmp85, und eines seiner potentiellen Substrate, TtoA.
Der nAChR ist ein zur Superfamilie der “Cys-loop Rezeptoren” gehörender ligandenge- steuerter Ionenkanal, der eine Schlüsselrolle in der synaptischen Weiterleitung neuronaler Sig- nale spielt. nAChRs bestehen aus fünf zueinander homologen, gewebsspezifisch als Homo- oder Heteropentamere zusammengesetzten, Untereinheiten. Bislang ist lediglich eine Struk- tur mit 4.0 Å Auflösung eines Rezeptorpentamers ausTorpedo marmoratavorhanden, welche durch Elektronenmikroskopie an zweidimensionalen tubulären Kristallen erhalten wurde. Auch wenn hochaufgelöste Strukturen zweier homologer, prokaryotischer Varianten kürzlich veröf- fentlicht wurden, so fehlt bis heute immer noch eine Struktur vergleichbarer Qualität für den nAChR. Es wurde zwar bereits im Jahre 1988 von Mikrokristallen berichtet, welche mit Präpa- rationen Detergenz-solubilisierter, nAChR-reicher Membranen aus dem elektrischen Organ des pazifischen RochensTorpedo californica erhalten wurden, die Eigenschaften dieser Kristalle wurden jedoch nicht weiter untersucht. In der hier vorgestellten Arbeit wurden solche Kristalle mittels Synchrotronstrahlung charakterisiert und für Proteinkristalle typische Beugungsbilder wurden aufgenommen. Die Röntgenbeugung dieser Kristalle war auf eine Auflösung von 20 Å beschränkt, mögliche Raumgruppen und Einheitszellparameter wurden anhand der gesam- melten Daten bestimmt. Weder eine Verfeinerung der Kristallisationsbedingungen, noch die Optimierung des Reinigungsprotokolls führten zu für hochauflösende Röntenstrukturbestim- mung geeigneten Kristallen.
Der Omp85-Familie zugehörige Proteine sind in Bakterien, Mitochondrien und Chloroplasten an der Faltung und Insertion von β-Barrel-Proteinen in oder der Translokation von Proteinen über die Außenmembran beteiligt. Das thermophile Bakterium Thermus thermophilus stellt eine Übergangsform zwischen Gram-positiven und Gram-negativen Bakterien dar, dessen Aus- senmembran bislang lediglich grob charakterisiert ist. T. thermophilus besitzt mit TtOmp85 einen archaischen Vertreter der Omp85-Familie. TtOmp85 wurde imT. thermophilus Stamm HB27 überexprimiert und nativ aus Präparationen der Außenmembran gereinigt. In der Anwe- senheit von Detergenz existierte TtOmp85 hauptsächlich als, aus zwei Protease-stabilen Mo- dulen zusammengesetztes, Monomer. Detergenz-solubilisiertes TtOmp85 wurde für Kristalli- sationsversuche verwendet und Mikrokristalle unter Bedingungen mit PEG400 als Fällungsmit- tel erhalten. Diese Kristalle beugten Röntgenstrahlen bis zu einer Auflösung von 5.0 Å und gehörten der Raumgruppe C222 mit den Einheitszelldimensionen vona =96.3 Å, b=416.1 Å,c=94.3 Å;α=β=γ =90° an. Aufgrund der, von der Orientierung der plättchenförmigen Kristalle im Röntgenstrahl abhängigen, ungleichmäßigen Beugungseigenschaften und während der Datensammlung akkumuliertem Strahlenschaden konnte lediglich ein unvollständiger Da- tensatz aufgenommen werden. Mit den vorhandenen Daten konnte die Struktur von TtOmp85 nicht gelöst werden. Präparationen von Detergenz-solubilisiertem TtOmp85 wurden kooperier- enden Arbeitsgruppen für begleitende Studien bereitgestellt, um TtOmp85 weitergehend zu charakterisieren.
Wenige andere Außenmembranproteine wurden bislang für T. thermophilus beschrieben.
TtoA, ein neues β-Barrel-Außenmembranprotein, wurde durch bioinformatische Analyse der Genomsequenz des Stammes HB27 identifiziert. Die Resultate eines in vitro Bindungstests, in dem sich TtoA an TtOmp85 binden ließ, legen die Vermutung nahe, daß die Insertion von Proteinen in die Außenmembran vonT. thermophilusnach einem ähnlichen Mechanismus funk- tioniert wie in modernen Gram-negativen Bakterien. TtoA wurde erfolgreich kristallisiert und dessen Kristallstruktur als erstes Außenmembranprotein eines thermophilen Bakteriums bis zu einer Auflösung von 2.8 Å bestimmt. Die Kristalle gehörten der Raumgruppe P3121 an, mit Einheitszellparametern von a = b = 166.7 Å, c = 97.5 Å; α = β =90°, γ =120°. TtoA besteht aus einem achtsträndigen β-Barrel mit einem großen extrazellulären Bereich, an dem ein zweiwertiges Kation gebunden ist. Ein fünfsträndigesβ-Faltblatt ragt aus dem in die Außen-
membran eingelagertenβ-Barrel hervor, welches durch eine Disulfidbrücke stabilisiert wird.
Die Kante diesesβ-Faltblattes bildet Kristallkontakte mit benachbarten TtoA-Molekülen aus, die unter Umständen physiologische Interaktionen mit anderen Proteinen nachahmen.
1 General introduction
Biological membranes establish the interfaces between living cells and their surroundings and separate cellular compartments. They consist of phospholipid molecules which, due to their am- phiphilic architecture, self-organize into bilayers of phospholipid planes: the polar headgroups face the aqueous medium while the apolar moieties orient towards the centre of the lipid bi- layer. Thereby, they form a barrier for hydrophilic molecules which can hardly diffuse through the hydrophobic part of the membrane. The physical characteristics of a biological membrane are defined by its actual composition of various different kinds of lipid molecules. The specific biological function of a membrane, however, is defined by proteins which are inserted in the lipid bilayer. These proteins often span the entire membrane plane and are then called “trans- membrane proteins”. Transmembrane proteins fulfill a variety of different functions, contribute about 50% of the mass of biological membranes and act as channels, transporters, receptors, enzymes, anchors and more (von Heijne, 2007).
The architecture of transmembrane proteins is specifically adapted to the amphiphilic envi- ronment of biological membranes. According to this, the protein surface facing the membrane is significantly hydrophobic while the non-transmembrane parts are usually hydrophilic. Due to their architecture, transmembrane proteins possess a unique orientation in the membrane and do not flip within the lipid bilayer. Because the peptide bonds in the backbone of proteins are po- lar, they need to be shielded from the surrounding apolar lipid bilayer in membrane embedded parts of proteins. This is achieved by the formation of secondary structure elements, mainlyα- helices orβ-strands, in which hydrogen bonds are established between carbonyl-oxygen (C=O) and amid-hydrogen (N–H) atoms of two peptide bonds in close proximity.
These two main forms of secondary structures differ in the way how the hydrogen bonds are established: inα-helices the hydrogen bonds are formed between residues within the same
Figure 1.1: Porin fromRhodobacter capsulatus. Ribbon representation of a porin monomer. Porin occurs as a homotrimer with 301 amino acid residues per subunit in the outer membrane (Weisset al., 1991).
α-helix four residues apart in the amino acid sequence of the protein. In contrast,β-strands are formed by hydrogen bonds between residues of neighbouring strands which can be many resi- dues apart from each other in the amino acid sequence. Consequently,β-strands always exist in arrangements of two or moreβ-strands, the so-calledβ-sheets. Due to this special arrangement of the hydrogen bonds,β-sheets can only move as whole entities, whereasα-helices can slide against each other. Therefore, the membrane spanning parts of transmembrane proteins which undergo large conformational changes during their functional cycles, e.g. receptors which trans- fer a signal received on one side of the membrane to the other side, usually are composed of bundles ofα-helices (von Heijne, 2007).
Proteins with membrane spanning parts composed of β-strands form a closed form of β- sheets, a so-called β-barrel (fig. 1.1). Because the hydrogen bonds of β-barrels bind each β-strand to its two neighbours,β-barrels are very rigid structures. They often act as channels for water molecules, ions, sugars and other components of small molecular weight. Substrate specificity is achieved by the presence of appropriate amino acid side chains lining the lumen of the β-barrel. Some β-barrel proteins, however, also act as receptors or fulfill enzymatic
Figure 1.2: Potassium channel KscA fromStreptomyces lividans. Orientation of the KscA tetramer in the membrane in ribbon representation. Aromatic amino acids on the membrane-facing surface are displayed in black (Doyleet al., 1998).
functions. Transmembraneβ-barrels are found in outer membranes of bacteria, mitochondria and chloroplasts (Schulz, 2002). On the opposite, transmembrane proteins in eukaryotic cells and bacterial plasma membranes are constructed byα-helices (fig. 1.2).
Biological membranes are no static arrangements of lipid molecules and transmembrane pro- teins but rather a twodimensional, semi-fluid composition of their components (Singer and Nicolson, 1972). The molecules of a biological membrane diffuse laterally in the membrane plane, though, this movement can be restricted to defined areas of a membrane (fig. 1.3). This is either achieved by components within the membrane itself, usually by transmembrane proteins arranged as some kind of a fence, or by membrane-underlying components of the cytoskeleton.
These mechanisms confine transmembrane proteins to specific regions of the continuous lipid bilayer, which is obviously important for organizing cells in organs.
Transmembrane proteins are the keyrole players in cellular communication and thus respon- sible for meeting one of the common life-defining criteria: namely the ability to respond to stimuli, often referred to as “excitability”. In contrast to their major contributions to the func- tionality of single- or multicellular organisms, their restriction to the amphiphilic environment
Figure 1.3: Schematic drawing of a biological membrane according to the fluid mosaic membrane model of Singer & Nicolson (Singer and Nicolson, 1972).
makes transmembrane proteins hard to be studied. Compared to water soluble proteins, they can hardly be obtained in substantial amounts and in isolated, functional form for bological ex- periments. This also affects attempts to crystallize transmembrane proteins for X-ray structure solution. Still nowadays, despite constant scientific and methodological progress, functional and structural knowledge about transmembrane proteins is still limited compared to the vast of data available for soluble proteins.
X-ray structure analysis is of great importance to understand the mechanisms of how biolog- ical processes work and how diseases can be cured: According to recent estimates, 46% of the human targets for small-molecule drugs currently on the market are transmembrane proteins, 30% of those are G-protein coupled receptors (Hopkins and Groom, 2002; Russ and Lampel, 2005). The atomic structure of a protein is the ultimate chemical information of biological macromolecules. Chemistry relies on the arrangement of electronic shells and bonds as the atoms themselves are conserved in all biological and chemical processes. The arrangement and covalent connection of all atoms of a protein therefore is the basis for understanding the chemical and dynamical processes in proteins.
The work presented here was focused mainly on crystallographical, but also functional, ana- lysis of three purified transmembrane proteins: a) The nicotinic acetylcholine receptor (nAChR)
prepared from a natural source, the electric tissue of the pacific ray Torpedo californica. b) The outer membrane protein Omp85 fromThermus thermophilus HB27, homologously over- expressed herein. c) The outer membrane protein TtoA fromT. thermophilusHB27, heterolo- gously overexpressed in the closely related, but TtoA-negative, strainT. thermophilusHB8.
The nAChR of eukaryotic organisms is one of the main components of synaptic communica- tion between neurons. Despite its emersed importance in understanding the molecular details of how this communication is realized in neuronal cells as well as in understanding the related numerous diseases observed, no high-resolution structure of either variant of nAChR is avail- able so far. The nAChR fromT. californicawas chosen as subject for crystallization attempts because (Hertling-Jaweedet al., 1990) it is one of the best characterized nAChR-isoforms, its purification from native sources is well established but no alternative recombinant sources are available, yet, to obtain nAChRs in sufficient amounts for crystallization attempts.
Proteins of the Omp85 family mediate the insertion ofβ-barrel proteins into the outer mem- brane of mitochondria, chloroplasts, Gram-negative and hereto related bacteria. Omp85 pro- teins themselves are located in the outer membrane and are often found in complex with ac- cessory peripheral membrane proteins to fulfill their function (Boset al., 2007b). An ancestral form of Omp85 family proteins exists in the thermophilic bacteriumT. thermophiluswhich be- longs to one of the oldest branches of evolution. Regarding its outer membrane, not much is known about the present integral membrane proteins. Considering the evolutionary relation- ships between thermophilic bacteria like T. thermophilus and mitochondria, chloroplasts and the branch of Gram-negative bacteria, the Omp85 protein encoded by the genome ofT. ther- mophilusHB27, TtOmp85, represents an ancestral type of its family. Proteins of thermophilic organisms usually display a higher degree of stability, due to thermal stress which makes them preferred objects for crystallization attempts for X-ray structure determination. Taking these two points together, TtOmp85 is a promising target to study the mechanism of protein insertion into outer membranes.
Only a few outer membrane proteins ofT. thermophilusare characterized and no structure of such a protein was available so far. TtoA, aβ-barrel outer membrane protein recently identified by searching the genome sequence of strain HB27 for proteins meeting criteria featured by
other outer membrane proteins, can serve as a model to study the features of those proteins in thermophilic organisms and their insertion into the outer membrane by TtOmp85.
2 Methods of X-Ray structure determination
To obtain magnified images of protein molecules, atomic distances have to be optically resolved.
Covalently bonded atoms are 1–2 Å apart, therefore it is necessary to use electromagnetic waves of wavelengths no larger than this scale to optically resolve the atomic structure of a protein.
Electromagnetic waves of such wavelengths are known as X-rays. The shorter the wavelength of X-rays is, the less they interact with matter and the less they become scattered. Radiation of wavelentghs between 1.0 Å and 2.0 Å, so-called “soft X-rays”, is used for X-ray crystallogra- phy.
Crystals provide a regular lattice of multiple copies of the (macro-)molecule of interest. Soft X-rays are scattered by atoms contained in that lattice, resulting in destructive and constructive interference of the scattered photons. Still, they penetrate a crystal sufficiently so that the whole crystal volume contributes to that scattering. For monochromatic X-rays constructive interfer- ence leads to amplified, defined signals which are finally observable as diffraction patterns (fig.
2.1). In contrast to electromagnetic waves used in microscopy, X-rays can not be focused by lenses. Thus, no direct image of the scattering object can be optically retrieved during the exper- iment. However, a threedimensional image of the scattering macromolecule can be calculated by examination of datasets of diffraction patterns which are obtained by consecutive rotation of a crystal in a monochromatic X-ray beam. .
As the presented work is focused on X-ray crystallographic analysis of the membrane pro- teins nAChR, TtOmp85 and TtoA, the general steps involved in the process of X-ray structure determination shall be schematically introduced in this chapter.
Figure 2.1: Diffraction pattern of a protein crystal. Shown is a diffraction pattern of a crystal of the outer membrane protein TtoA fromThermus thermophilusobtained during rotation of the crystal of 0.5° around itsϕ-axis.
2.1 Crystallization of proteins
The fundamental requirement for X-ray structure determination of proteins is to grow crystals of the protein of interest. This step is the least manageable as crystal growth depends on prop- erties of the yet unknown protein structure and therefore is mainly a trial-and-error procedure.
Methodological improvements have been achieved to overcome the initial problem of identify- ing successful crystallization conditions with a preferably low amount of purified protein: The application of “sparse-matrix-screens” of crystallization conditions led to a significantly de- creased number of buffer compositions to be screened to identify successful crystallization con- ditions. These screens cover the statistically most successful crystallization conditions for pro- teins and systematically explore the composition of crystallization buffers (Jancarik and Kim, 1991; Brzozowski and Walton, 2001). Additionally, nano-drop robotic automatization of set- ting up crystallization experiments was developed to minimize the amount of purified protein required for crystallization experiments (Sulzenbacheret al., 2002).
Figure 2.2: A schematic phase diagram showing the solubility of a protein in solution as a function of the concentration of the precipitant (McPherson, 1999).
Apart from such major improvements, resulting in a constantly growing number of new pro- tein structures deposited in the Protein Database (PDB) per annum, the principle of protein crystallization remained the same: A higly pure solution of native protein at a concentration of several milligrams of protein per milliliter of buffer solution is brought to supersaturation to cause crystal nucleation. This is done by mixing a protein solution with a crystallization buffer which provides a precipitant and/or salts at a defined pH. The solubility of the protein might then be lowered in this new buffer environment, the solution becomes supersaturated.
As a consequence, small aggregates might be formed whose intermolecular contacts resemble those found in the final crystal lattice, the crystal contacts. These highly regular clusters have to reach a critical size in order to function as nuclei for crystal growth which is then thermody- namically favoured over random precipitation (Jullienet al., 1994). Since there is this energy barrier, nucleation takes time. If the supersaturation is too small, the nucleation rate will be so slow that no crystals form in any reasonable amount of time. The corresponding area of a phase diagram, plotting protein concentration versus precipitant concentration (fig. 2.2), is known as the “metastable zone”. In the “labile zone”, the supersaturation is large enough that sponta- neous nucleation is possible. If the supersaturation is too large, disordered structures, such as aggregates or precipitates, may form. Thus, the “precipitation zone” is unfavorable for crystal formation, because the aggregates and precipiates form faster than the crystals (Asherie, 2004).
However, in most protein-buffer-mixtures the precipitation kinetics only lead to such disor- dered aggregates or precipitates. Therefore, the success of protein crystallization depends on
the number of different crystallization conditions tested. Still, a large number of different crys- tallization conditions alone does not guarantee for growth of crystals. In the first place, the intrinsic characteristics of the protein and the solution it is provided in decide upon the success of crystallization experiments (D’Arcy, 1994; Garavitoet al., 1996; Derewenda, 2004).
Crystallization experiments are commonly done using the vapour-diffusion method: the ac- tual crystallization experiment takes place in a preferably small volume (≤ 1 µl) in which crystallization buffer is mixed with the solution of the purified protein. This volume is placed next to a reservoir of pure crystallization buffer. The set-up is then sealed from the environ- ment. The volume of the reservoir is two or three orders of magnitude larger than the actual crystallization experiment. The differences in concentrations of the various components of the two buffer compartments lead to diffusion of volatile components, mainly water molecules from the drop into the reservoir. In consequence, the concentration of precipitant in the protein drop slowly increases. The drop gets supersaturated and either protein precipitation or nucleation with subsequent crystal growth occurs (McPherson, 2004).
Proteins that are naturally membrane-embedded or membrane-associated cause additional problems because they are insoluble in normal aqueous solutions. This makes the application of conventional protein crystallization based on aqueous buffers obviously problematic. Vari- ous types of detergents are used to substitute lipids attached to membrane embedded parts of protein surfaces (Ostermeier and Michel, 1997): detergents are amphiphilic compounds, their hydrophobic part binds to hydrophobic surface areas of proteins and renders them hydrophilic by exposing their hydrophilic part towards the solvent. Care has to be taken that the detergent acts mild enough towards the membrane protein. It should only attach to hydrophobic surface parts and not penetrate into the core of the membrane protein which would unfold and denature its defined threedimensional native structure.
The inevitable use of detergents in crystallization attempts of membrane proteins adds an additional dimension to the matrix of conditions to be screened for identification of success- ful crystallization conditions: not each detergent works with every membrane protein so that various potentially useful detergents have to be tested. Some detergents function best when ac- companied by small amphiphilic molecules which adds yet another dimension to the screening task (McPherson, 2004). Additionally, the membrane protein surface covered by the disordered
detergent micelle usually does not participate in forming crystal contacts, restricting the number of possible arrangements of membrane proteins in crystal lattices.
Specialized techniques have been developed for crystallization of membrane proteins, for example crystallization in lipidic-cubic phases: a membrane system consisting of lipids, water and purified protein in appropriate proportions is established. These components form a struc- tured threedimensional lipidic array, which is pervaded by a network of aqueous channels and is thought to provide a suitable arrangement for crystal contact interactions between membrane proteins (Landau and Rosenbusch, 1996). A recent modification of this approach is crystalliza- tion in lipidic sponge phases which represent the liquid analogue of the cubic phase allowing its application in conventional vapour-diffusion crystallization experiments (Wadstenet al., 2006).
Nevertheless, these methods represent no routine methods for crystallization of membrane pro- teins as they require specialized equipment and broad experience with the system. By far the most frequently used system in membrane protein crystallization still is the vapor-diffusion method applied to mixed micellar protein-detergent solutions.
Given the fact that membrane proteins usually can be prepared only in small amounts, their crystallization remains a challenging task in X-ray protein structure determination. No matter, this adventure is still feasible and each year dozens of new and unique structures of membrane proteins are solved – of the∼46000 crystal structures deposited in the PDB, by end of the year 2008, 400 represent structures of membrane proteins (nearly 200 unique structures).
2.2 Protein crystals
Unlike crystals of salts or small molecules, protein crystals contain a high content of solvent, usually between 30% and 70% (Matthews, 1968). The protein occupies the remaining volume of the crystal. Thus, instead of a solid corpus the entire crystal is more of an ordered gel.
The loose packing of proteins is permeated by interstitial spaces through which solvent and other small molecules can freely diffuse. In proportion to its molecular mass, the number of interactions that a biological macromolecule forms with its neighbours in the crystal lattice – salt bridges, hydrogen bonds and hydrophobic interactions – is very low (McPherson, 2004).
As a consequence, the growth of protein crystals is a quite rare event. A characteristic of
protein crystals which is important for crystal treatment and handling is that they disintegrate if allowed to dehydrate, they are temperature sensitive and crush easily. Compared to salt crystals they diffract X-rays less strongly: protein crystals are usually smaller in size, they contain less unit cells per volume and their internal order is decreased.
The resolution of the diffraction pattern of a crystal depends on the degree of internal order of the crystal and the diffraction volume, i.e. the total size of the crystal. The internal order of protein crystals easily suffers from the relatively large spaces between adjacent molecules and the weak lattice forces. Consequently, the protein molecules in the crystal do not occupy exact equivalent orientations and positions, they rather vary slightly within the crystal lattice.
These imperfections in location of the protein molecules is called “mosaicity”. With increased mosaicity the level of detail to which atomic positions can be determined by subsequent crys- tal structure analysis is limited. The degree of mosaicity therefore should be minimized by refinement of the crystallization conditions to influence the kinetics of crystal growth.
A crystal is a threedimensional repetition of identical copies of a single building block, the
“unit cell”. The dimensions of the unit cell are defined by three vectors~a,~band~c. According to the building block analogy, a whole crystal is defined by multiples of these three vectors. Within the unit cell, a crystal can contain further symmetry elements – rotation, reflection and inver- sion – dividing it into several “asymmetric units” which form the most basic structural element within the crystal. The geometry of the unit cell defines together with the possible symmetry op- erations the “space group” of the crystal. Among 230 possible space groups belonging to seven crystal systems (table 2.1), only 65 are enantiomorphic and thus feasible for chiral molecules like proteins – hence, space groups comprising reflections or inversions are not possible. Iden- tification of the correct space group is essential for correct indexing of diffraction patterns and it is therefore the first step to solve a crystal structure. Conventionally, the unit cell with the shortest edges and the highest possible symmetry is chosen (Blundell and L. N. Johnson, 1976).
2.3 X-ray diffraction by crystals
Upon interaction with the crystal lattice, the oscillating electrical field of an X-ray photon in- duces an oscillation of equal frequency in the electron shell of the atoms. In elastic or coherent
Crystal system Minimum symmetry Conventional choice Constraints on interaxial
requirement of axes angles and axial lengths
triclinic None No constraints None
monoclinic One 2-fold axis aandcperpendicular αandγ=90°
to 2-fold
bparallel to 2-fold
orthorhombic Three perpendicular a,bandcparallel α,β,γ=90°
2-fold axes to the 2-fold axes
trigonal One 3-fold axis aandbperpendicular αandγ=90°,β=120°
to 3-fold aandbof equal length cparallel to 3-fold
tetragonal One 4-fold axis aandbperpendicular α,β,γ=90°
to 4-fold aandbof equal length cparallel to 4-fold
hexagonal One 6-fold axis aandbperpendicular αandγ=90°,β=120°
to 6-fold aandbof equal length cparallel to 6-fold
cubic Four 3-fold axes a,bandcrelated α,β,γ=90°
by 3-fold axis a,bandcof equal length
Table 2.1: The seven crystal systems (Blow, 2002).
diffraction (also known as Thomson scattering), the electrons act as oscillating dipoles emitting secondary radiation of the same frequency as the incident radiation. The single waves that orig- inate from any point of the finite electron density sum up to a total intensity of the secondary radiation of zero (destructive interference), except if the path difference between the waves is an integer multiple of their wavelength (constructive interference). Diffraction of X-rays from the real lattice of a crystal creates a reciprocal threedimensional lattice of diffraction maxima, called “reflections”. In the reciprocal lattice the geometric properties are inverse to those of the real crystal.
A convenient way to describe diffraction by a crystal lattice is to imagine every single diffrac- tion spot to be a reflection of the incident beam on a family of imaginary planes which is iden-
Figure 2.3: Lattice planes and Bragg’s law. a) Lattice planes that allow for constructive interference of diffracted waves are those that divide the unit cell edges into integer fractions. The number of those fractions is used to index the plane. The family of lattice planes shown would have the Miller indices (2,3,3). b) Bragg’s law: two waves that are reflected by a family of lattice planes with distance d have a difference in path length that is equal to2dsinθ, as it can be derived from the scheme. A prerequisite for constructive interference is that this difference in path is an integer multiple of the wavelength used:
2dh,k,lsinθ=nλ(Blow, 2002).
tified by the “Miller indices”(h, k, l)(fig. 2.3 a)). The Miller indices represent an unequivocal numbering scheme of all reflections.
Constructive interference from a set of lattice planes with distancedwill occur at an angle of θif the path difference between the diffracted waves is an integer multiple of the wavelengthλ and thus they are in phase. This relation between diffraction angle and lattice plane distance is known as “Braggs’ Law” (fig. 2.3b)):
2dh,k,lsinθ =nλ
The “Ewald sphere” (fig. 2.4 a)) defines reciprocal lattice points where Bragg’s law is ful- filled. It is a sphere of a radius of 1/λ with the crystal in its centre. The points where the incident beam enters the sphere and the origin O of the reciprocal lattice are on opposite sides of the centre C. Bragg’s law is fulfilled for every reciprocal lattice point that is intersected by the Ewald sphere. A rotation of the crystal results in an equal rotation of the reciprocal lattice around its origin O. During rotation, different reciprocal lattice points move through the Ewald sphere. If a reciprocal lattice point intersects with the sphere, a reflection is observed on the X- ray detector plate (fig. 2.4 b)). Hence, for a specific orientation of a crystal a defined ensemble of reflections is observed – referred to in crystallography as “diffraction pattern”.
Figure 2.4: a) The Ewald construction. The crystal (C) is placed in the centre of a sphere (here, in two dimensions, a circle) with radius1/λ. The origin of the reciprocal lattice, i.e. reflection (0,0,0), is located in (O). The reciprocal lattice (dots) will rotate as the crystal is rotated. When the reciprocal lattice point crosses the surface of the sphere, the trigonometric condition1/d = (2/λ)sinθis fulfilled. This is the three-dimensional illustration of Bragg’s law. Thus, those reciprocal lattice points that intersect with the Ewald sphere are in diffraction condition and will be recorded by the detector. b) A still exposure with a stationary crystal contains only a small number of reflections arranged in a set of narrow ellipses. (Dauter, 1999)
As every recorded reflection represents one ensemble of lattice planes(h, k, l), the measure- ment of the positions of the spots is sufficient to deduce the geometry of the crystal lattice and in most cases also the space group by examining the symmetry observed in the diffraction pattern (Evans, 2006). The intensity of the diffraction spots is determined by the content of the unit cell.
The result of data collection will primarily be the knowledge about space group, unit cell dimenions and the intensity measurementI(h, k, l)for every reflection(h, k, l).
2.4 Structure determination by X-ray crystallography
2.4.1 The electron density function
The aim of a crystallographic experiment is to calculate the distribution of electron density in the asymmetric unit – and thus also the unit cell – in order to deduce an atomic model of the crystallized molecule.
The scattering from all atoms in the unit cell is the sum of all atomic scattering factorsf, taking into account individual phase shifts. For every single reflection(h, k, l)this summation leads to a structure factorF(h, k, l):
F(h, k, l) = X
i
fie2πi(hxi+kyi+lzi)
The sum of all structure factors is the Fourier transform of the electron distribution in the crystal. This means that the electron density ρ(x, y, z) for every point in real space can be calculated as a Fourier summation over all structure factors:
ρ(x, y, z) = 1 V
X
h,k,l
F(h, k, l)e2πi(hx+ky+lz)
2.4.2 The phase problem
Each structure factor F(h, k, l) represents a reflection by a family of lattice planes. It is de- scribed by a wave function with an amplitude,|F(h, k, l)|and a phase angle,α(h, k, l):
F(h, k, l) = |F(h, k, l)|eiα(h,k,l)
The structure factor amplitude|F(h, k, l)|is obtained experimentally as it is the square root of the measured intensityI:
I(h, k, l) =|F(h, k, l)|2
Unfortunately there is no way to observe the time of arrival of the amplitudes of the scattered X-rays so that the information about the phase shift in respect to the incoming wave is lost.
The determination of the phases is therefore a crucial step in any crystal structure determination once diffracting crystals have beeen obtained and a data set has been recorded. It is commonly referred to as the “phase problem”. Different approaches are used to overcome this problem:
• Molecular replacement
• Single or multiple isomorphous replacement (SIR/MIR)
• Single isomorphous replacement with anomalous scattering (SIRAS)
• Single- or multiple-wavelength anomalous dispersion (SAD/MAD)
• Direct methods (only applicable for high-resolution data)
During structure solution of the outer membrane protein TtoA from Thermus thermophilus in chapter 5, two of those methods were used and thus shall be introduced here in more detail:
Phase determination by molecular replacement
If a known structure of a protein is very similar to the yet unsolved structure of the target protein or similar to a major part of it, the known structure can be used as a model to estimate the missing phase information: a unit cell is assembled which contains – instead of the unknown protein structure – the model structure. However, this unit cell has to be of the same dimensions and geometry as deduced from the collected data set of diffraction images. Then the model structure has to be placed into it in the correct number of molecules in their respective correct orientations.
The tool to place the model structures into the unit cell is the “Patterson function” (PF), which is the Fourier transform of the scattering intensities (Patterson, 1934). The amplitude of a scattered wave can be retrieved from the intensity of the diffracted X-ray beam: this intensity is proportional to the square of the amplitude of the corresponding wave. Therefore, they can be thought of as the product of the amplitudes of the scattered wave with itself (Blow, 2002).
Importantly, a PF can also be obtained from the convolution of a scattering structure with itself.
This allows to calculate PFs as well for experimental scattering data as for any arrangement of protein molecules in any unit cell. The peaks in the PF represent the intra- and interatomic distances of pairs of atoms, weighted by the product of the number of electrons in the atoms concerned. The resulting Patterson vectors are arranged in a volumeU around the origin of the PF with a radius equal to the dimension of the molecule. A twodimensional plot of the PF is called “Patterson map“ (PM).
According to this, the correct orientation of the model structure can be found by systemati- cally exploring the PF of the experimentally measured intensities. This is achieved by observing
the correlation of the PM of the measured intensities with the PM of the search model while rotating and translating the search model in threedimensional space. Due to computational reasons, this sixdimensional search is divided into two subsequent threedimensional searches:
rotation and translation.
First, the rotational search is carried out. The rotation of the molecule leads to a modification of its corresponding PM. The search molecule is rotated to any possible rotational orientation defined by the three rotational angles. At each angular position the actual functional values are correlated with those of the experimental PM all through the volumeU. The correlation of the PMs has maxima if the orientation of the intramolecular vector sets are coincident (Rossmann and Blow, 1962).
Once the correct orientation of the model structure has been determined, the correct transla- tion of the molecule can be obtained from translation searches. The translations in the volume of the unit cell are evaluated by comparison of the resulting crystallographic R-factors, Patterson correlation and packing criteria.
If more than one copy of the model structure has to be placed in the unit cell, the described rotation- and translation searches has to be carried out for each additional molecule.
If finally the correct orientations of the model structures in the unit cell are known, the elec- tron density of the unknown structure can be calculated from structure factors composed of the experimental amplitudes and the calculated phase angles of the correctly oriented model structures (Rae, 1977).
Phase determination by isomorphous replacement: SIR, MIR and SIRAS
MIR is the most common experimental approach to solve the phase problem in X-ray crys- tallography if phase determination by molecular replacement is not successful. MIR is done by soaking crystals of the protein of interest with heavy-atom solutions or by co-crystallizing the protein with heavy-atoms. The heavy-atoms have to bind to specific sites of the crystal- lized protein and must not affect the crystal form or unit cell dimensions in comparison to native crystals without heavy-atoms. This means, crystals of native protein and the heavy-atom
derivative should be isomorphic. The structure factorFPH for a heavy-atom derivative crystal then becomes
FPH =FP+FH
where FP is the structure factor of the native protein and FH is the contribution of heavy- atom substructure to the structure factor of the derivative. The isomorphous difference,FH = FPH−FPcan be estimated from experimental data sets of the native and derivative protein.
If a dataset of a derivative crystal is collected, the positions of the heavy-atoms can be local- ized: while it is not possible to calculate an electron density map without phase information, the intensities of the native and the derivative datasets can be used to calculate PMs. The differ- ence between these two PMs reveals the PM of the heavy-atom substructure, i.e. the location of the heavy-atoms bound to the protein. This is done by correlation studies of the PMs and is feasible as long as the number of heavy-atom positions is not too high (which is usually the case). Once the heavy-atom positions have been determined and refined, the phase angles for the heavy-atom substructure can be determined by the so-called ”Harker construction“. These phase angles serve as a first estimation for the phases of the protein structure.
If only one heavy-atom derivative is available (SIR), the determination of phase information leaves an ambiguity, the Harker construction for SIR is shown in figure 2.5: FH is calculated from the identified heavy-atom positions. Circles are drawn with the radiiFPandFPH around pointsOandA, respectively. The intersection points of both circles then determine two possible phase angles forFP.
This phase ambiguity can be overcome if two or more isomorphous heavy-atom derivatives are available which possess different heavy-atom substructures (MIR). A Harker construction with two heavy-atom derivatives is shown in figure 2.6. The point which is the closest to the intersections of all three cirlces,ϕM, serves as the initial phase angle forFP.
Another way to avoide phase ambiguity if only one isomorphic heavy-atom derivative is available is phase determination via SIRAS. The ambiguity is overcome by making use of anomalous scattering: particularly heavy-atoms give rise to anomalous scattering. The effect becomes maximal if the energy of the incident X-rays is close to an absorption edge of the used
Figure 2.5: Harker construction for SIR.ϕ1 andϕ2 are the most probable phase angles for FP (ITfC, 2006).
Figure 2.6: Harker construction for MIR. Harker construction for double isomorphous replacement.ϕM
is the most probable phase angle for FP (ITfC, 2006).
type of heavy-atom. As the X-ray beam provided by synchrotrons is tunable, strong anomalous scattering of heavy-atoms can be caused on purpose. The wavelength of the used X-ray beam is tuned close to the absorption edge of the heavy-atom which is bound to the protein in the derivative crystal.
For most other wavelentghs the atomic scattering factorf for an atom is proportional to the number of electrons that it possesses. For wavelengths that approximate the one for which an atom strongly absorbs radiation, the scattering factor undergoes a change due to anomalous dispersion. The dispersion not only affects the magnitude of the factor but also imparts a phase shift in the elastic collision of the photon, leading to breakdown of ”Friedel’s law“. Friedel’s law states that related pairs of reflexesh, k, l andh, k, l, the so called ”Friedel pairs“, have the same value for their structure factor amplitudes,|F(h, k, l)|and|F(h, k, l)|but opposite values for their phase angles. The atomic scattering factor of the atom in question can be expressed as
f =f0+ ∆f0+if00 =f0+if00
wheref0is the scattering factor far from an absorption edge, and∆f0andf00are the correction terms which arise due to dispersion effects. The quantity ∆f0, in phase with f0, is usually negative andf00, the imaginary part, is alwaysπ/2 ahead of the phase of the real part(f0+ ∆f0).
In consequence to breakdown of Friedel’s law, the structure factor amplitudes of the respective Friedel pairs differ: FPH+ 6=FPH- andFH+ 6=FH-. This allows to set up a Harker construction which resolves the phase ambiguity and allows the identification of the phase information for the present set of heavy-atoms. A Harker construction for SIRAS is shown in figure 2.7.
However, it has to be mentioned that due to increased radiaton absorption during data col- lection close to absorption edges of heavy-atoms, the accompanying radiation damage occuring in the crystal is also increased. This leads to accelerated decay of scattering of X-rays by the crystal with ongoing exposure to radiation, introducing a new problem into data collection.
2.4.3 Model building and refinement
After initial phase information is retrieved by one of the aforementioned methods, a first electron density map can be calculated. If heavy-atom derivatives were used, this electron density is
Figure 2.7: Harker construction for SIRAS (ITfC, 2006).
called ”experimental electron density map“ as it is deduced from experimentally obtained data.
An electron density map is a threedimensional representation of the location probability of the electrons in the unit cell and thus effectively a representation of the atomic structure of the protein. As shown in figure 2.8, the protein structure is visible as continuous tubes of electron density.
In such initial maps usually not the whole protein structure is clearly visible because the phase angles are still inaccurate. Additionally, flexible areas in proteins result in a decreased electron density because the location of those electrons is less defined.
In practice, the model of a protein structure is created by placing amino acids into continuous and clear defined areas of electron density according to the amino acid sequence of the protein.
Usually, the protein mainchain is better defined in electron density maps than sidechains. This is because the mainchain is less flexible than the amino acid sidechains. Side chains mainly establish non covalent atomic interactions and thus remain more flexible than the covalently bonded atoms of the mainchain.
The more atoms of a protein are defined by the atomic model, the more accurately the phase angles can be determined during refinement. As a consequence of more accurately determined phase angles the calculated electron density maps improve. Thus, model building is a cyclic process of model improvement according to the available electron density map and refinement
Figure 2.8: Experimental electron density map. The location of the mainchain of a protein (TtoA, see chapter 5) is visible as continuous areas of electron density, represented by meshed areas. For the left molecule the Cα-backbone is drawn into the respective electron density.
of the phase angles followed by further improvement of the subsequently calculated electron density map.
3 Purification and crystallization of the nicotinic acetylcholine receptor from Torpedo californica, characterization of microcrystals
3.1 Introduction
Rapid communication of neuronal signals across cell membranes makes use of a coordinated network of transmembrane proteins. Ion pumps generate local membrane potentials by selective ion transport. Upon incoming signals these membrane potentials become modified by signal activated ion channels. These act as selective pores for passive but rapid flow of ions down their electrochemical gradients. Depending on the selectivity for either anions or cations they depolarize or hyperpolarize membrane potentials. The various types of signal activated ion channels in nervous systems can be distinguished by the nature of the signal they receive and how the signal is translated into the flow of ions: ligand-gated ion channels, voltage-gated ion channels, mechanoreceptors and G-protein coupled receptors (Kandelet al., 2000).
Amongst ligand-gated ion channels three superfamilies are known: the pentameric ligand- gated ion channels, ionotropic glutamate receptors and the Adenosin-5’-triphosphate (ATP) - gated channels. The basic principle of ligand gating is that the ligand binds differently to the open and the closed conformations of the channels. If it binds more tightly to the open state the ligand acts as a channel activator, if binding to the closed conformation is stronger it will be an inhibitor (Ashcroft, 2006).
Figure 3.1: Stereo representation of the AChBP protomer as viewed from outside the pentameric ring (Brejcet al., 2001). The ribbon representation is coloured as a rainbow gradient, from blue to red (N- to C-terminus). Disulphide bridges are indicated in green ball-and-stick representation. In a complete ion channel the N-terminus points towards the synaptic cleft and the C-terminus enters the membrane at the bottom, continuing into the first transmembrane helix M1.
Pentameric ligand gated ion channels, or as called since more recently “Cys-loop receptors”
(CLRs), exist for a number of ligands such as acetylcholine and 5-hydroxy-tryptamine (sero- tonin), both permeable for cations, and forγ-aminobutaric-acid and glycine, both permeable for anions. CLRs consist of five homologuous subunits composed as homo- or heteropentamers.
The actual composition of a specific CLR depends on its localization in the nervous system and results in a variety of tissue-specific receptor subtypes.
Major progress in the understanding of CLR function in general and of the nicotinic acetyl- choline receptor (nAChR) in particular was achieved in the recent years when two protein struc- tures were solved: First, the 2.7 Å crystal structure of the homopentameric acetylcholine bind- ing protein (AChBp) from the marine snailLymnea stagnalis(Brejcet al., 2001), homologous to extracellular domains of nAChRs (fig. 3.1). Second, the 4.0 Å structure derived from electron microscopy on tubular 2D-crystals of the muscular nAChR-subtype from Torpedo marmorata (Unwin, 2005) consisting of a heteropentamer of two α- and each one β-, γ- and δ-subunit (“α2βγδ”, fig. 3.2).
Additional crystal structures of an AChBp from Aplysia californica (Hansen et al., 2005) or AChBPs in complex with agonist, various toxins and other structural analogous compounds (Celieet al., 2004; Bourneet al., 2005; Ulenset al., 2006) revealed detailed information about ligand binding and conformational changes of the extracellular domain of CLRs. Recently,
Figure 3.2: 4.0 Å model of muscle-type nAChR fromTorpedo marmorata(Unwin, 2005). (a) view from the synaptic cleft and (b) view parallel with the membrane plane. Ligand-binding domains are located betweenα- andγ- orα- andδ-subunits, repectively, highlighted by a tryptophane residue of theα-subunit (yellow). Only the front two subunits are highlighted in (b) (α, red;β, green;γ, blue;δ, light blue). Also shown are the the main immonologic region (MIR) and the membrane (horizontal bars; E, extracellular; I, intracellular). The apex of the C-loop ofαδ(broken trace in (a)) was not visible in the densities.
crystal structures of prokaryotic CLRs from Gloeobacter violaceus, “GLIC” (Bocquet et al., 2008), and Erwinia chrysanthemi, “ELIC” (Hilf and Dutzler, 2008), were published. These channels represent ancestral examples of CLR structures and were crystallized in open (GLIC) and closed (ELIC) conformation.
The subunits of CLRs can be divided topologically into a N-terminal extracellular domain which contains the ligand binding site and a C-terminal transmembrane domain. The actual channel is formed by symmetric arrangement of the five transmembrane domains. The accu- mulating structural information led to a functional model how CLRs might translate chemical signals into the flow of ions (Sine and Engel, 2006): The extracellular side of CLR-subunits contains the ligand-binding site and is almost exclusively composed of β-strands. It contains two opposingβ-sheets, a singleα-helix at the very top and a number of connecting loops (fig.
3.1). Notably two loops play important roles in ligand-binding (“C-loop”) and in coupling of
the chemical signal to channel-opening (“Cys-loop”). Ligand binding takes place at the inter- face betweenα-subunits and their respective counterclockwise neighbouring subunit (fig. 3.2) and subsequently leads to a major movement of the C-loop (Brejcet al., 2001). The movement of the C-loop is proposed to be propagated to the Cys-loop via rigid body movements of the two opposing β-sheets of the extracellular domain which causes an altered orientation of the Cys-,β1-β2- andβ8-β9-loops. The displacement of these three loops then couples the ligand binding to channel opening through interactions with the loop connecting the transmembrane helices M2 and M3 (Unwin, 2005).
The transmembrane domain of each single nAChR subunit consists of fourα-helices, “M1”–
“M4”, of which only M2 lines the channel pore (fig. 3.2). In the pentameric receptor the M2-helices of the five subunits fit together like the staves of a barrel. Ion selectivity is achieved by charged residues of each helix M2. These residues point towards the channel lumen and are located at the extracellular and intracellular ends of M2 (Imotoet al., 1988). Additionally, residues of the intracellular helix, located between helices M3 and M4 (fig. 3.2), are also contributing to ion selectivity (Kelleyet al., 2003). Upon ligand binding, the induced movement of the loop between M2 and M3 leads to a reorientation of the M2-helices towards each other.
As a consequence the distances between the sidechains of the ion selectivity filters change.
The connective part between the lower end of the M3-helix and the lower end of the intra- cellular “MA”-helix, 70–100 residues for each subunit, is not resolved in the final 4 Å model of muscle-type nAChRs fromT. marmorata(Unwin, 2005; Sine and Engel, 2006).
Before such detailed knowledge about Cys-loop receptor function became available, already in 1988 three dimensional crystals obtained from preparations of detergent-solubilized nAChR- enriched membranes from the electric organ of the pacific rayTorpedo californiawere reported by the group of F. Hucho, Freie Universität Berlin (Hertling-Jaweedet al., 1988). The crystals were of column-like shape with dimensions of 5µm×5µm×20µm. The work on the nAChR fromT. californicareported here was done in cooperation with the group of F. Hucho with the aim a) to analyze the present but yet uncharacterized microcrystals by X-ray crystallography and biochemical methods and b) to reproduce the crystals and to refine the crystallization conditions to obtain crystals suitable for X-ray structure analysis.
3.2 Materials and methods
3.2.1 Solubilization and purification of nAChRs from nAChR-enriched membranes from T. californica
Isolated membranes from the electric organ of T. californica enriched in nAChR were pro- vided by the group of F. Hucho. Membrane fragments were prepared by continuous density gradient centrifugation (25–50% sucrose) as described (Hertling-Jaweed et al., 1990). Frac- tions of sucrose gradients containing membrane fragments with nAChRs as the main com- ponent were chosen by SDS-PAGE analysis (Laemmli, 1970) as starting material for further nAChR-purification. The total amount of protein was determined by the Bradford protein assay (Bradford, 1976) as this assay is less influenced by lipids. According to the published initial purification protocol (Hertling-Jaweedet al., 1990) used for the production of microcrystals, sucrose gradient-fractions corresponding to 6 mg of total protein were incubated for 1 h in 15 ml of 50 mM Tris (pH 8.1), 10 mM ethylene glycol tetraacetic acid (EGTA) and 10 mM lithium 3,5-diiodosalicylate (LIDS) to remove proteins peripherally bound to the membranes and subsequently centrifuged at 100000 × g for 1 h at 4°C. Pellets were resuspended in 50 mM Tris(hydroxymethyl)aminoethane (Tris) (pH 7.4), 10 mM EGTA and 0.5% nonyl-β-D- glucoside (NG) and incubated at gentle agitation for 12 h at 4°C followed by centrifugation at 100000 × g for 1 h at 4°C. The supernatant was applied to 5 ml of blue sepharose medium (CL6B, Amersham) to remove a cosolubilized Na+/K+-ATPase subunit and passed through the column via gravity flow. The flow through was collected and the medium was washed with an- other 2 ml of buffer as used in the solubilization step and added to the flow through. The whole volume of flow through was concentrated to a protein concentration of 15 mg/ml by ultrafil- tration in Centricon-concentrators (100 kilodaltons [kDa] molecular weight cut-off [MWCO];
Millipore) and used for crystallization experiments.
In order to optimize this initial purification protocol various other detergents were tested.
Screened detergents were different members of the most common families of non-ionic and zwitter-ionic detergents. Their concentration during solubilization of the membrane fragments in buffer containing 50 mM Tris (pH 7.4) and 200 mM NaCl was chosen between 3–100 times
