Structural aspects of molybdenum-transhydroxylase from Pelobacter acidigallici and tungsten-acetylene hydratase from Pelobacter acetylenicus
Dissertation submitted to
Fachbereich Biologie, Universität Konstanz, Germany
for the degree of
Doctor of Natural Sciences
by
Dipl.-Biol. Holger Niessen
Konstanz, February 2004
Dissertation der Universität Konstanz Datum der mündlichen Prüfung: 10. Mai 2004
Referent: Prof. Dr. P.M.H. Kroneck Co-Referent: Prof. Dr. A. Messerschmidt
Für meine Frau und meine Eltern
Table of Contents I
Table of Contents
TABLE OF CONTENTS...... I
ZUSAMMENFASSUNG ...... IV
1. Acetylenhydratase aus Pelobacter acetylenicus...... IV
2. Transhydroxylase aus Pelobacter acidigallici ...... V
SUMMARY...... VII
1. Acetylene hydratase from Pelobacter acetylenicus ...... VII
2. Transhydroxylase from Pelobacter acidigallici...... VIII
1. INTRODUCTION...... 1
1.1 Physical and chemical properties of molybdenum and tungsten...... 1
1.2 The mononuclear molybdenum and tungsten enzymes...... 3
1.3 The molybdenum cofactor...... 6
1.4 Iron-sulfur centers...... 8
1.5 Pelobacter acetylenicus acetylene hydratase ...... 11 1.5.1 Metabolism of acetylene by P. acetylenicus ...... 12 1.5.2 Molecular properties of acetylene hydratase...... 13
1.6 Pelobacter acidigallici transhydroxylase...... 14 1.6.1 Metabolism of gallic acid by Pelobacter acidigallici ...... 14 1.6.2 Molecular properties of transhydroxylase...... 15
1.7 Scope of the study...... 17
2. MATERIALS AND METHODS...... 19
2.1 Chemicals and biochemicals...... 19
2.2 Organisms ...... 20 2.2.1 Pelobacter acetylenicus...... 20 2.2.2 Pelobacter acidigallici ...... 20
2.3 Cultivation of bacteria ...... 20 2.3.1 Pelobacter acetylenicus...... 20
2.4 Glycerol cryo cultures...... 26
2.5 Enzyme purification...... 26 2.5.1 Acetylene hydratase...... 27 II Table of Contents
2.5.2 Transhydroxylase ...... 28
2.6 Enzyme activity ...... 28 2.6.1 Acetylene hydratase...... 28 2.6.2 Transhydroxylase ...... 29 2.6.3 Alcohol dehydrogenase...... 30
2.7 UV/Vis spectroscopy ...... 30
2.8 Analytical methods...... 30 2.8.1 ICP-MS...... 30 2.8.2 Protein ...... 30 2.8.3 Polyacrylamide gel electrophoresis...... 31
2.9. Crystallography...... 31 2.9.1 Theoretical background...... 31 2.9.2 Crystal growth...... 32 2.9.3 Crystals...... 34 2.9.4 X-ray diffraction by crystals ...... 34 2.9.5 The electron density function...... 36 2.9.6 The phase problem ...... 38 2.9.7 MAD with metalloproteins...... 39 2.9.8 Crystallisation under exclusion of dioxygen...... 42 2.9.9 Cryocrystallogragphy...... 43 2.9.10 Substrate and inhibitor complexes ...... 44 2.9.11 Data collection...... 44 2.9.11 Transformation of crystals ...... 45 2.9.12 Data analysis...... 46
3 RESULTS...... 48
3.1 Acetylene hydratase of Pelobacter acetylenicus ...... 48 3.1.1 Growth of Pelobacter acetylenicus under various conditions ...... 48 3.1.2 Purification of acetylene hydratase ...... 49 3.1.3 Crystallization and three-dimensional structure of acetylene hydratase...... 51 3.1.4 SAD data collection at ESRF...... 52
3.2 Transhydroxylase of Pelobacter acidigallici...... 54 3.2.1 Growth of Pelobacter acidigallici...... 54 3.2.2 Purification of transhydroxylase...... 56 3.2.3 Metal content of tungstate cultivated P. acidigallici Transhydroxylase...... 59 3.2.4 Activity measurements of molybdenum Transhydroxylase...... 60 3.2.5 Crystallization of Transhydroxylase ...... 60 3.2.6 Unit cell parameters of transhydroxylase...... 61 3.2.7 Transformation of transhydroxylase crystals ...... 62 3.2.8 Data collection of transhydroxylase crystals...... 63 3.2.9 Heavy metal soaks of transhydroxylase crystals...... 63 3.2.10 MAD and SAD data collection of transhydroxylase crystals ...... 63 3.2.11 Structure determination...... 65 3.2.12 Description of the structure...... 66 3.2.12.1 Overall structure of transhydroxylase ...... 66 3.2.12.2 α-subunit of transhydroxylase...... 67 3.2.12.3 β-subunit of transhydroxylase ...... 72 Table of Contents III
3.2.12.4 Transhydroxylase with substrate pyrogallol bound ...... 74 3.2.12.5 Transhydroxylase with inhibitor 1,2,4-trihydroxy-benzene bound...... 75
4. DISCUSSION ...... 77
4.1 Molybdenum versus tungsten in enzymes ...... 77
4.2 Cultivation of bacteria and enzyme purification...... 78 4.2.1 Pelobacter acetylenicus...... 79 4.2.2 Pelobacter acidigallici ...... 79
4.3 Crystallization and structural analysis of acetylene hydratase ...... 80
4.4 Structural aspects of transhydroxylase...... 83 4.4.1 Crystallization experiments...... 83 4.4.2 MAD experiments at DESY...... 85 4.4.3 SAD experiment at the ESRF...... 86 4.4.4 Overall structure of transhydroxylase ...... 87 4.4.5 Towards the reaction mechanism of transhydroxylase ...... 90 4.4.5.1 Former proposed rection mechanisms...... 91 4.4.5.2 New reaction mechanism ...... 94 4.4.5.3 Is the reaction molybdenum transhydroxylase efficient?...... 97 4.4.5.4 The role of the β-subunit and its [4Fe-4S]-clusters...... 99
5. REFERENCES...... 101
6. APPENDIX ...... 110
6.1 Abbreviations...... 110
6.2 Amino acids...... 111
6.3 Nucleic acid bases...... 112
6.4 International System of Units (SI) ...... 112
6.5 Figures and Tables ...... 113
6.6 Supplementary Table...... 115
6.8 Curriculum vitae ...... 119
6.9 Publications...... 120
6.10 Conference abstracts...... 121
IV Zusammenfassung
Zusammenfassung
1. Acetylenhydratase aus Pelobacter acetylenicus
P. acetylenicus ist ein mesophiles, strikt anaerob lebendes Bakterium, das in der Lage ist, auf Acetylen als einziger Kohlenstoff- und Energiequelle zu wachsen. Die Metabolisierung von Acetylen wird durch das W/Fe-S abhängige Enzym Acetylenhydratase eingeleitet, wobei in einer ungewöhnlichen Reaktion Acetylen zu Acetaldehyd hydratisiert wird.
Das Enzym Acetylenhydratase wurde aus P. acetylenicus zur Homogenität gereinigt. Es handelt sich um ein Monomer mit einer molekularen Masse der Aminosäurekette von 81.9 kDa. Das Enzym gehört zur Familie der DMSO-Reduktasen. Acetylenhydratase ist ein thermostabiles Enzym, dessen Temperaturoptimum im Bereich von 50 bis 55°C liegt. In einer Stick- stoff/Wasserstoff Atmosphäre bei 6°C konnte das Enzym 3 Monate gelagert werden, ohne daß ein Aktivitätsverlust festgestellt wurde (Abt, 2001). Obwohl die Acetylenhydratase keine Redox- Reaktion katalysiert, enthält sie ein [4Fe-4S] Zentrum und einen W-bisMGD Kofaktor.
Kristalle der W-Acetylenhydratase wurden in Anwesenheit und in Abwesenheit (N2 : H2 = 94 : 6 v/v) von Luftsauerstoff erhalten. Jedoch nur die in Abwesenheit von Luftsauerstoff erhaltenen Kristalle waren in der Lage Röntgenstrahlen zu beugen. Dithionit-reduziertes Enzym ergab unter Ausschluß von Luftsauerstoff Kristalle, die am Deutschen Elektronen Synchrotron (DESY) in Hamburg vermessen wurden und bis zu einer Auflösung besser als 2,5 Å streuten. Ebenfalls wurden Kristalle am ESRF in Grenoble an der Wolfram L-Kante vermessen, jedoch konnte dieser Datensatz aufgrund sehr hoher Mosaizität und einer Abnahme der Auflösung nicht benutzt werden die 3-dimensionale Struktur zu ermitteln. Dieser Kristall war nicht stabil genug der Synchrotronstrahlung standzuhalten.
Mit dem in Hamburg aufgenommenen nativen Datensatz wurde ein molecular replacement durchgeführt und ein erstes Model der Acetylenehydratase konnte mit der Hilfe von Dr. Oliver Einsle berechnet werden. Für das molecular replacement wurde die Wolfram- Formatdehydrogenase FDH-T (Raaijmakers et al., 2002) verwendet und eine Elektronendichtekarte berechnet. Jedoch ist eine vernünftige Verfeinerung dieses Models, mit einer Auflösung von bis zu 2.4 Å, bislang noch nicht möglich gewesen. Zusammenfassung V
2. Transhydroxylase aus Pelobacter acidigallici
P. acidigallici ist ein strikt anaerob lebendes Bakterium, das in der Lage ist, mit Gallussäure (3,4,5-Trihydroxybenzoesäure), Pyrogallol (1,2,3-Trihydroxybenzol), Phloroglucin (1,3,5-Tri- hydroxybenzol) oder 2,4,6-Trihydroxybenzoesäure als einziger Kohlenstoff- und Energiequelle zu leben. Ein entscheidender Schritt während der Metabolisierung von decarboxylierter Gallussäure (Pyrogallol) ist die Transhydroxylierung des Pyrogallols zum Phloroglucin. Diese Reaktion wird von dem Mo/Fe-S abhängigen Enzym Transhydroxylase (Pyrogallol:Phloroglucin Hydroxyltransferase E.C. 1.97.1.2) katalysiert.
Es handelt sich um ein Heterodimer, das aus einer großen Untereinheit (100,4 kDa) und einer kleinen Untereinheit (31,3 kDa) besteht. Das Enzym ist eng mit Mitgliedern der DMSO- Reduktase Familie verwandt. Obwohl die Gesamtreaktion der Transhydroxylase keine Redoxreaktion ist, enthält das Enzym einen Mo-bisMGD Redoxkofaktor und verschiedene Eisen-Schwefel Zentren.
12 der 13 Cysteine der kleinen Untereinheit der Transhydroxylase sind hochkonserviert. Einige davon sind als [4Fe-4S] Ferredoxine beschrieben worden. Die 15 Cysteine der großen Untereinheit lassen sich nicht mit den Cysteinen anderer Proteine abgleichen. Aus diesem Grund ist es wahrscheinlich, daß die Eisen-Schwefel Zentren sich in der kleinen Untereinheit befinden.
Experimente mit „as isolated“-Transhydroxylase führten zu Kristallen, die im Röntgenstrahl nicht beugten. Die Kristallisation von Dithionit-reduzierter Transhydroxylase unter anoxischen Bedingungen in einer Stickstoff/Wasserstoff-Atmosphäre führte zu Kristallen, die mit Synchrotonstrahlung eine Auflösung von mehr als 2,5 Å ereichten.
Diese Kristalle konnten mit Synchrotronstrahlung vermessen und die 3-dimensional Struktur der Transhydroxylase ermittelt werden. Ebenfalls konnten hochaufgelöste Strukturen der Transhydroxylase mit Pyrogallol und Inhibitor (1,2,4-Trihydroxybenzol) mit einer Auflösung von bis zu 2.0 Å ermittelt werden. Diese Ergebnisse ließen Schlussfolgerungen über einen neuartigen möglichen Reaktionsmechanismus zu. In diesem Falle hat das Molybdän die Funktion das Pyrogallol an dessen C1 Position zu koordinieren. Katalytische Funktionen haben hierbei die Aminosäuren Asp A174, His A144 und Tyr A404 in der unmittelbaren Nähe zum aktiven Zentrum. VI Zusammenfassung
Die Funktion der kleineren β-Untereinheit mit den 3 [4Fe-4S]-Klustern ist unklar. Die Distanz vom [4Fe-4S]-1 zum Molybdän ist mit 23.4 Å ist für einen effizienten Elektronentransfer zu groß.
Summary VII
Summary
1. Acetylene hydratase from Pelobacter acetylenicus
P. acetylenicus is a strictly anaerobic and mesophilic bacterium that is able to grow on acetylene as single energy and carbon source. The first step in the metabolization of acetylene is the transformation of acetylene to acetaldehyde. This addition of water is catalyzed by the W/Fe-S dependent enzyme acetylene hydratase.
Acetylene hydratase from P. acetylenicus was purified to homogeneity. It is a monomer with a molecular mass of the amino acid chain of 81.9 kDa. BLASTP searches revealed that the enzyme is highly similar to enzymes of the DMSO-reductase family. Acetylene hydratase is a thermostable enzyme with a temperature optimum between 50 and 55°C. It is a very stable enzyme when stored under exclusion of dioxygen in a nitrogen/hydrogen atmosphere at 6°C (Abt, 2001). Within three months, there was no detectable loss of acetylene hydratase activity from tungstate-grown P. acetylenicus. Although acetylene hydratase catalyzes no redox reaction, it contains one [4Fe-4S] center and one W-bisMGD as redox-cofactors.
Crystals of the w-acetylene hydratase were obtained both in presence and in absence (N2: H2 = 94: 6 v/v) of dioxygen. Only the crystals grown in absence of dioxygen were able to diffract X- ray-radiation. Dithionite-reduced enzyme crystals obtained under exclusion of dioxygen, could be measured at the Deutsches Elektronensynchrotron (DESY) in Hamburg up to resolution better then 2,5Å. Also crystals were measured at the ESRF in Grenoble at the tungsten L-edge, however, this dataset could not be used to solve the three dimensional structure because of high mosaicity and decreasing of resolution. This crystal was not stably enough to stand the measurement in the synchrotron radiation.
A molecular replacment of the native dataset collected in Hamburg was performed and a preliminary model of acetylene hydratase could be calculated with the help of Dr. Oliver Einsle. For molecular replacment the tungsten containing formate dehydrogenase (FDH-T) (Raaijmakers et al., 2002) was used and electron density calculated. A suitable refinment of this model with a resolution limit of 2.4 Å was not possible till now.
VIII Summary
2. Transhydroxylase from Pelobacter acidigallici
P. acidigallici is a strictly anaerobic bacterium that is able to live on gallic acid (3,4,5- trihydroxybenzoic acid), pyrogallol (1,2,3-trihydroxybenzene), phloroglucinol (1,3,5-trihydroxy- benzene), or 2,4,6-trihydroxybenzoic acid. A crucial step in the fermentation of decarboxylated gallic acid (pyrogallol) is the transhydroxylation of pyrogallol to phloroglucinol. This reaction is catalyzed by the Mo/Fe-S dependent enzyme transhydroxylase (pyrogallol:phloroglucinol hydroxyltransferase E.C. 1.97.1.2).
Transhydroxylase from P. acidigallici is a heterodimer consisting of a large subunit (100.4 kDa) and a small subunit (31.3 kDa). This enzyme is closely related to enzymes of the DMSO- reductase family. Although the overall reaction of transhydroxylase is no redox reaction it contains different iron-sulfur centers and one Mo-bisMGD as redox-cofactors.
12 of the 13 cysteines in the small β-subunit are highly conserved. Some of them are referred to the [4Fe-4S] ferredoxins. The 15 cysteines of the big subunit do not align with the cysteines of related iron-sulfur proteins. Therefore, it is unlikely that an iron-sulfur center is located in the large subunit. It is more likely that there are three [4Fe-4S] clusters located in the small subunit.
Crystals of as isolated transhydroxylase were not able to diffract X-ray radiation. Crystallization of dithionite reduced transhydroxylase under exclusion of dioxygen led to crystals which diffracted to resolution limits higher than 2.5 Å with synchrotron radiation. These crystals were measured with synchrotron radiation and the three-dimensional structure of transhydroxylase was solved. Even structures of transhydroxylase in complex with pyrogallol and inhibitor (1,2,4-trihydroxybenzene) were solved at high resolutions up to 2.0 Å
These result led to a new possible reaction mechanism. Hereby the function of the molybdenum ion is to coordinate the pyrogallol at its C1 position. The amino acids Asp A174, His A144 and Tyr A404 near the active site seem to have catalytic function.
The role of the small β- subunit containing the 3 [4Fe-4S] clusters is not clear. The distance between the [4Fe-4S]-1 and the molybdenum with 23.4 Å is too large for an efficient elctron transfer.
Introduction 1
1. Introduction
Molybdenum and tungsten are the only elements of the second and third row transition series to have known biological functions (Pilato and Stiefel, 1999; Johnson et al., 1996). This results from its good bio-availability, as well as from its suitability to undergo a two-electron reduction. Molybdenum has been recognized since the 1930s for its role in nitrogen fixing enzyme systems (Stiefel, 1997). From 1953 on it was realized that molybdenum is essential for diverse aspects of metabolism in a wide range of organisms (De Renzo et al., 1953). Tungsten was first identified in 1983 in the NADP-dependent formate dehydrogenase from the thermophilic organism Clostridium thermoaceticum (Yamamoto et al., 1983) and has been extensively studied in bacteria and hyperthermophilic archaea since then. Molybdenum and tungsten enzymes are found throughout the biological world and catalyze critical reactions in the metabolism of purines, aldehydes, carbon monoxide, as well as nitrogen– and sulfur containing compounds (Hille, 1999; Stiefel, 1997). With the exception of nitrogenase, the molybdenum and tungsten enzymes share a structural unit at their catalytic sites. This component is called the molybdenum cofactor (moco, Figure 1.3) and binds molybdenum as well as tungsten. There are now a large series of molybdenum and tungsten enzymes with known three- dimensional structures, and this new structural information has provided the basis for an increasingly detailed understanding of the reaction mechanisms of these enzymes. An overview of the diverse structures and functions of the molybdenum and tungsten-enzymes is given in a number of recent publications: Hille, 2000; Pilato and Stiefel, 1999; Kisker et al., 1999; Hille et al., 1999; Hille, 1999; Rees et al., 1997; Stiefel, 1997; Johnson et al., 1996; Kletzin and Adams, 1996; Hille, 1996.
1.1 Physical and chemical properties of molybdenum and tungsten
Although the chemistry of molybdenum and tungsten is variable and complex because of the range of possible oxidations states (-II to +VI), only the +IV, +V, and +VI oxidation states of both elements appear biologically relevant (Kletzin and Adams, 1996). But Yandulov et al. (2003) showed a theoretical reaction where the molybdenum in nitrogenases could have also the oxidation state +III. The similarity in their chemical properties is well established (Table 1.1). 2 Introduction
The atomic radii of Mo and W, as well as their electron affinity, are virtually the same. Radioactive isotopes suitable for biological research are available for both elements (99Mo and 185W), as well as stable nuclear spin isotopes for the study of hyperfine interactions by various spectroscopic techniques (95Mo, I = 5/2 and 183W, I = 1/2). Both, W and Mo, are relatively rare in nature. The abundance in the earth’s crust is only 1.2 ppm for both elements. The concentrations in seawater are ≈ 100 nM for Mo (Frausto da Silva and Williams, 1991) and ≈ 1 pM for W. In freshwater, the Mo concentration is in the range of 5 – 50 nM and the W concentration is less than 500 pM (Table 1.1). IV Molybdenum is mostly present in jordesite and molybdenite (both Mo S2) and seldom in the
+VI oxidation state as wulfenite (PbMoO4) or powellite (CaMO4; M=Mo or W), Greenwood and Earnshaw, 1990).
Tungsten is usually found in oxo-rich minerals (oxidation state +VI) either as scheelite (CaWO4) IV or wolframite ([Fe/Mn]WO4), whereas the more reduced tungstenite (W S2) is very rare, in part because WS2 is readily solubilized.
2- + - WS2 + 4H2O WO4 + 2H2S + 4H + 2e
Molybdenum (Mo) Tungsten (W) Atomic number 42 74 Average atomic weight 95.94 183.85 Electronic configuration of the outer shell 4d5 5s1 4f14 5d4 6s2 Atomic radii (Å) 1.40 1.40 Ionic radii for +IV oxidation state (Å) 0.65 0.66 Ionic radii for +V oxidation state (Å) 0.61 0.62 Ionic radii for +VI oxidation state (Å) 0.59 0.60 Electronegativity 1.8 1.7 2- - pKa of oxo acid (MO4 /HMO4 ) 3.87 4.60 Concentration in seawater ≈ 100 nM ≈ 1 pM Concentration in freshwater ≈ 5 – 50 nM ≈ 500 pM M = O bond length (Å) 1.76 1.76
Table 1.1 Physical and chemical properties of molybdenum and tungsten. Compiled from Kletzin and Adams (1996) and Greenwood and Earnshaw, (1990). M represents Mo or W.
Introduction 3
You can find molybdenum in enzymes in two basic forms: a.) as a component of the FeMoco active center of nitrogenases, and b.) in the mononuclear centers of oxomolybdenum enzymes (Moco), which are therefore called molybdenum enzymes (Hille 1996).
1.2 The mononuclear molybdenum and tungsten enzymes
Since the discovery of the first molybdenum enzyme, over 50 mononuclear molybdenum or tungsten enzymes have been discovered (Hille et al., 1999). They catalyze a variety of hydroxylations, oxygen atom transfer, and other oxidation-reduction reactions, and share the unique molybdenum cofactor. The mononuclear oxomolybdenum enzymes were divided into hydroxylases and oxotransferases due to the catalyzed reactions, the sequence homologies, the comparison of the subunit and/or domain structures, the crystal structures as well as EPR-and EXAFS spectroscopic characteristics (Hille 1996). While hydroxylases are a relatively homogeneous group of enzymes with similar composition of cofactors and a similar amino acid sequence, the oxotransferases becomes further subdivided into the DMSO reductase and the sulfite sulfit-oxidase family. In the nitrogen cycle both molybdenum enzymes nitrogenase and nitrate reductase are key enzymes. In the metabolism of N-heterocycles a large family of molybdenum enzymes encompasses a wide range of substrate specifities that allow hydroxylation of carbon centers in strategic regiospecificity. In the sulfur cycle, molybdenum-dependent sulfite oxidation and dimethyl sulfoxide (DMSO) reduction play crucial roles. In carbon metabolism, both in the formation of methane and oxidation of formate, carbon monoxide, and various aldehydes, the molybdenum enzymes again have a prominent position (Stiefel, 1997). The tungsten enzymes are involved in carbon metabolism and usually have functions related to those of their molybdenum counterparts (Stiefel, 1997). Table 1.2 shows some stoichiometric formulations for substrate reactions of molybdenum enzymes. In table 1.3 a list of molybdenum and tungsten enzymes is given sorted according to their metabolic roles.
4 Introduction
Enzyme Reaction + - Dimethyl sulfoxide reductase (CH3)2SO + 2H + 2e → (CH3)2S + H2O + - Trimethylamine N-oxide reductase (CH3)3NO + 2H + 2e → (CH3)3N + H2O - + - - Nitrate reductase NO3 + 2H + 2e → NO2 + H2O 2- 2- + - Sulfite oxidase SO3 + H2O → SO4 + 2H + 2e + - Formate dehydrogenase HCOOH → CO2 + 2H + 2e - - + - - - Polysulfide reductase S-(S)n-S + 2H + 2e → S-(S)n-1-S + H2S III - V 3- + - Arsenite oxidase As O2 + 2H2O → As O4 + 4H + 2e - + CO oxidoreductase CO + H2O → CO2 + 2e + 2H
Acetylene hydratase C2H2 + H2O → CH3CHO Transhydroxylase 1,2,3 trihydroxybenzene → 1,3,5 trihydroxybenzene
Table 1.2 Stoichiometric formulations for substrate reactions of selected molybdenum and tungsten enzymes.
Several alternative classification schemes have been suggested: Hille (1996, 1999) differentiated three families based on the structure of the molybdenum center in the oxidized Mo(VI) state plus one family for the tungsten containing enzymes (Figure 1.1). According to Dobbek and Huber (2001) the Mo/Cu CO-dehydrogenase from Oligotropha carboxidovorans belongs to the xanthine oxidase family. Based on sequence similarities Kisker et al. (1997) divided the moco containing enzymes into four different families, namely the dimethyl sulfoxide reductase (DMSOR), xanthine oxidase, sulfite oxidase, and aldehyde ferredoxin oxidoreductase (AOR) families.
S S SCys I VI Cu S Mo SOH
CODH
S O (S-Cys) O O OSer O SCys (Se-Cys) S MoVI S MoVI O S MoVI S S WS S OH S S S SS
The Xanthine Oxidase Family The Sulfite Oxidase Family The DMSO Reductase Family The Aldehyde Oxidoreductase Family molybdenum hydroxylases eukaryotic oxotransferases prokaryotic oxotransferases tungsten enzymes
Figure 1.1 The families of mononuclear molybdenum enzymes (Hille, 1996; Hille, 1999; Hille et al., 1999). Note that the recently discovered molybdenum-containing Carbon monoxide dehydrogenase (CODH) from Oligotropha carboxidovorans contains the first binuclear metal center (Dobbek and Huber, 2002).
Introduction 5
Molybdenum enzymes Tungsten enzymes Nitrogen cycle Aldehyde oxidoreductase (carboxylic acid reductase) Nitrogenase (only molybdenum enzyme not Formate dehydrogenase containing the pterindithiolene ligand) Formaldehyde ferredoxin oxidoreductase Nitrate reductase (assimilatory) N-Formyl methanofuran dehydrogenase Nitrate reductase (dissimilatory) Acetylene hydratase Nitrate oxidase Trimethylamine N-oxide reductase
N-Heterocyclic metabolism Isonicotinic acid hydroxylase Nicotinic acid hydroxylase Nicotine hydroxylase Picolinic acid dehydrogenase Pyrimidine oxidase Isoquinoline oxidoreductase Quinaldic acid 4-oxidoreductase Quinoline oxidoreductase Xanthine dehydrogenase Xanthine oxidase
Acid and aldehyde reactions Aldehyde oxidase (retinal oxidase) Aldehyde dehydrogenase Pyridoxal oxidase
Carbon metabolism Formate dehydrogenase Carbon monoxide oxidoreductase N-Formyl methanofuran dehydrogenase 2-Furoyl dehydrogenase
Sulfur metabolism Polysulfide reductase Sulfite oxidase Biotin sulfoxide reductase Dimethyl sulfoxide reductase Tetrathionate reductase
Miscellaneous Transhydroxylase Arsenite oxidase Chlorate reductase
Table 1.3 Molybdenum and tungsten enzymes (Stiefel, 1997).
6 Introduction
Over the years scores of structures of mononuclear molybedum enzymes were solved (Dobbek, 2000) (Figure 1.2).
Mononuclear molybdenum enzymes
Sulfite-oxidase Xanthine-oxidase DMSO-Reductase familiy family family (LMoOS ) (LMoO 2 ) (L 2 MoX) CODH FDH DMSOR SO
TMAOR MOP
Figure 1.2 Molybdenum enzymes. From left to right. Sulfite-oxidase (SO) (Kisker, Schindelin et al. ,1997), CO-dehydrogenase (CODH) (Dobbek, Gremer et al., 1999), Aldehyde- oxidoreductase (MOP) (Romao, Archer et al., 1995), DMSO-Reductase (DMSOR) (Schindelin, Kisker et al., 1996), Formate-dehydrogenase (FDH) (Boyington, Gladyshev et al., 1997), TMAO-Reductase (Dos Santos, Iobbi-Nivol et al., 1998).
1.3 The molybdenum cofactor
The three-dimensional structure of the Mo-molydopterin cofactor (Figure. 1.3A) was first shown 1995 with the structure of the aldehyde oxidoreductase of Desulfovibrio gigas (Romao, Archer et al., 1995), before the tricyclic ring structure was discovered already in the tungsten containing enzyme aldehyde oxidoreductase from Pyrococcus furiosus (Chan, Mukund et al., 1995). In all well-known enzymes the molybdenum cofactor has the same structure, consisting of an organometallo complex made of molybdenum and a pterin derivative. Both are present in the molybdenum hydroxylase and sulfite sulfite-oxidase family in a 1:1 stoichiometry. The pterin is a 2-amino-4-hydroxy-pteridin with a C4-alkyl side chain at the position 6. The pyran ring is formally formed thereby by the condensation of the 3‘-OH-group of the C4 side chain of the pterin ring with the C7-atom of the pterin nucleus. The positions C1‘ and C2‘ of the pyran ring are sp2-hybridisated and carry in cis configuration a sulfur ligand. The sulfur atoms of this Introduction 7
dithiolene function coordinate the molybdenum ion. The C4‘-phosphate-group forms a pyrophosphate bridge with the 5‘-phosphate-group of the 5‘-CMP in the molybdopterin cytosine dinucleotide (MCD) cofactor (Romao, Archer et al. ,1995) (Huber, Hof et al., 1996). The Mo or W ion that binds to the moco is found to be coordinated by three types of ligands: (i) sulfur atoms provided by the moco; (ii) non-protein oxygen or sulfur species, such as oxo, water or sulfido; (iii) (optionally) amino acid side chains (Rees et al., 1997). In bacteria, additional variability of the moco is achieved by conjugation of one of the nucleotides guanosine, adenosine, inosine, or cytidine-5’-monophosphate to the phosphate group of the moco (Figure 1.3 B). The name of the resulting molecule is abbreviated e.g. as MGD (molybdopterin-guanosine-dinucleotide). One or two molecules of e.g. MGD can complex the molybdenum or tungsten atom (Figure 1.3 C). Enzymes with the molybdenum cofactor often incorporate additional cofactors or prosthetic groups such heme, coenzyme B12, or iron-sulfur centers (Stiefel, 1997). The rest that is bound to the phosphate-group of the molybdopterin differs in the families, also in the enzymes, dependening on the organism from that they were isolated. While enzymes from eukaryotic systems possess the cofactor in the form represented above (R=H),one can find the dinucleotide of guanine, cytosine, adenine or hypoxanthine in prokaryontic systems also. The tri cycle is clearly not planar, both in the central pyrazine ring and in the pyrane ring. The pyrane ring is in half chair conformation, derives clearly from the level of the pterine systems and is bent in relation to the level of the pterine systems around ~30-60°. Although the biochemical meaning of the cofactors is not completely understood, it stabilizes obviously the proteine conformation and can be involved in the transfer of electrons to other cofactores. The conformational flexibility of the cofactors permits the modulation of the redox potential of the molybdenum ion by the relative position of the dithiolene ligands to the molybdenum. 8 Introduction
(A) OSH H N SH 4 5 1' HN 3 9 6 2'
2 10 7 3' 2- 1 8 4' OPO3 H2N NNO H O (B) OSH N H N SH 4 5 1' NH HN 3 9 6 2' N
2 10 7 3' O 1 8 4' O O O N H2N NNO P P NH H - - 2 O2 O2 HO OH
HO OH (C) - - Q-MGD O2 O2 H H N 2 P P O N N NH2 N O O O O N HN NH S N N H O S O O S Mo O H O N N S HN NH N O O O O N H2N N N O P P H - - NH2 O2 O2 P-MGD HO OH Figure 1.3 Cofactors of molybdenum and tungsten enzymes. (A) Molybdenum cofactor = molybdopterin = moco. The tricyclic form was observed in all crystal structures of enzymes containing this cofactor (Kisker et al., 1999). (B) Molybdopterin guanosine dinucleotide (MGD) form as found in some bacterial enzymes (Stiefel, 1997). (C) Extended molybdenum cofactor (bisMGD) as found in Alcaligenes faecalis arsenite oxidase (Ellis et al., 2001).
1.4 Iron-sulfur centers
In addition to moco, iron sulfur proteins are used mostly for a electron transfer at negative redox potential and can have redox or not-redox-catalytic function. The sulfide and iron ions are reversibly extractable from ferredoxins and the remaining apoenzyme can be reconstituted with external S2- and Fe2+/3+ . Their occurrence in hyperthermophilic and anaerobic organisms as well as its low redox potential suggest an important role of Fe/S clusters in the early evolution, in reactions, which could stand at the beginning of chemolithoautotrophic metabolism. Iron-sulfur Introduction 9
centers constitute one of the most ancient, ubiquitous, structurally, and functionally diverse class of biological prosthetic groups (Cammack, 1992; Beinert et al., 1997). In several molybdopterin-containing enzymes, such as acetylene hydratase, transhydroxylase, or xanthine oxidase iron-sulfur centers were found. In the simplest case the iron atom is tetrahedrally coordinated by four cysteinyl residues, whereas in the more complex centers several iron atoms are bridged by inorganic sulfide (S2-), the so- called acid-labile sulfur. In a scenario of the origin of life in hot environments (Achenbach- Richter et al., 1987; Wächtershäuser, 1988) the conversion of FeS (Eq. 1) was postulated to serve as the first energy source of primordial life (Wächtershäuser, 1988). Therefore, the iron- sulfur centers found today in proteins might represent remainders of the early past. The most common types of iron-sulfur centers comprise [2Fe-2S], [3Fe-4S], and [4Fe-4S] cores with cysteinyl residues serving as fourth ligand of each iron atom (Figure 1.4).
- + - FeS + HS → FeS2 + H + 2e E°’ = –620 mV
The iron-sulfur proteins fall into two major categories: simple iron-sulfur proteins that contain only one or more iron-sulfur centers, and the complex iron-sulfur proteins that bear such additional active redox centers as flavin, molybdenum, or heme. The most frequent and most stable type of the FeS centers is the [4Fe-4S]-type. It forms in first approximation tetrahedrons with the µ3-sulfidions on the tetrahedron surfaces and serves usually for the electron transfer with negative redox potential (to –0.7 V). The charge of the oxidized cluster is about -2 with 2 pairs of iron dimers with equal isomeric adjustment (Moessbauer spectroscopy), according to an oxidative state of +2.5 for Fe. This means that mixed-valence Fe(II)/Fe(III) pairs with large electron delocalization are present while effective spin pairing. The reason for the delocalization (resonance) with [4Fe-4S]-centers in contrast to the located charge of [2Fe-2S]-centers probably lies in a structurally caused orthogonality of the metal orbitals, which interacts over superexchanging sulfide bridges. A favouring effect for fast electron transfer are small changes of geometry of the center while reduction/oxidation. The number of hydrophobic rests, as well as the decreased accesibility of water seem to determine stability and redox potential the cluster (Stephens, Jollie et al., 1996).
10 Introduction
Rubredoxin [3Fe-4S]
[2Fe-2S] Ferredoxin
[4Fe-4S]
[2Fe-2S] Rieske center
Figure 1.4 Structures of the most common types of iron-sulfur centers. iron atoms are colored in gray, sulfur in yellow, and nitrogen in blue.
The functions of iron-sulfur proteins include electron and proton transfer, Lewis acid-base catalysis, structural determinant, and gene regulation (Johnson, 1994). The optical absorption bands of all iron-sulfur proteins are rather broad and featureless and not suitable for obtaining structural information. On the other hand, electron paramagnetic resonance (EPR) spectra of iron-sulfur centers are distinctive (Figure 1.5). From the spectra one can conclude on the nuclearity and redox state of the iron-sulfur centers (Cammack et al., 1985).
Introduction 11
gz=2.02 gy=1.91 reduced 2+/+ gav=1.91 [2Fe-2S] Rieske g =2.05 z g =1.79 reduced x 2+/+ gav=1.96 [2Fe-2S]
gy=1.95 Ferredoxin
gx=1.89 oxidized gz=2.02 gav=2.01 +/0 [3Fe-4S]
gx,y=2.00
reduced gz=2.06 gy=1.92 gav=1.96 2+/+ [4Fe-4S]
gx=1.88 gx,y=2.04 g =2.12 oxidized z 3+/2+ gav=2.06 [4Fe-4S] HiPIP
300 320 340 360 380 400
Magnetic field [mT]
Figure 1.5 Comparison of EPR properties of different types of iron-sulfur centers. The spectra of different types of iron-sulfur centers differ in shape and g-values. According to (Cammack et al., 1985). HiPIP = high potential iron-sulfur protein.
1.5 Pelobacter acetylenicus acetylene hydratase
Pelobacter acetylenicus strain WoAcy 1 (DSM 3246) is a strictly anaerobic, chemoorganotroph, and gram-negative bacterium that is able to grow on acetylene as sole carbon and energy source (Schink, 1985). It was isolated from a freshwater creek sediment near Konstanz. The cells are rod-shaped with 0.6 – 0.8 x 1.5 – 4 µm in size. The DNA base ratio is 57.1 ± 0.2 mol% G + C (Schink, 1984). 12 Introduction
1.5.1 Metabolism of acetylene by P. acetylenicus
Acetylene is so far the only known hydrocarbon that is metabolized in the absence and presence of molecular oxygen in the same manner (Schink, 1985). P. acetylenicus hydrates acetylene to acetaldehyde. The further disproportionation of acetaldehyde leads to acetate and ethanol Figure 1.6). Though the hydration of acetylene to acetaldehyde is a highly exergonic reaction (Schink, 1985), studies on cell yield show that only the free energy of the acetate kinase reaction (0.5 mol ATP per mol acetylene) is used for growth (Schink, 1985).
0’ -1 C2H2 + H2O → [H2C=C(OH)H] → CH3CHO ∆G = –111.9 kJ mol
The acetylene degradation pathway of P. acetylenicus is shown in figure 1.6.
2 HC CH 2 H O 1 2
O
2 H3C C H CoASH 3
2[H] O 2 H3C C S CoA Pi 4 H C CH OH 3 2 CoASH O
H3C C O P ADP 5
ATP O
H3C C O
Figure 1.6 Acetylene degradation pathway of P. acetylenicus (Rosner 1994, Schink, 1985). (1) Acetylene hydratase, (2) Alcohol dehydrogenase, (3) Aldehyde dehydrogenase, (4) Phosphate acetyltransferase, (5) Acetate kinase. Introduction 13
1.5.2 Molecular properties of acetylene hydratase
Acetylene hydratase has been isolated as a monomeric enzyme with a molecular mass of 72 kDa (SDS-PAGE) versus 85 kDa (MALDI-MS) and it is 730 amino acids long. The N-terminus of the protein shows a sequence motif C-x-x-C-x-x-x-C that could represent a motif for a Fe-S site (Rosner and Schink, 1995). 4.4 ± 0.4 mol Fe and 0.5 ± 0.1 mol W (ICP/MS), 3.9 ± 0.4 mol acid labile sulfur, and 1.3 ± 0.1 mol molybdopterin guanine dinucleotide were found per mol enzyme. Selenium was absent (Meckenstock et al., 1999). The isoelectric point is 4.2, the specific activity of the enzyme is highest between pH 6.0 and 7.0, and the temperature optimum is 50°C. Though the acetylene hydratase reaction (Eq. 2) is not a redox reaction, in the photometric assay a strong reductant like Ti(III)citrate or dithionite has to be used. Meckenstock et al. (1999) showed that acetylene hydratase contains one [4Fe-4S] cluster with a midpoint redox potential of – 410 ± 20 mV (Figure 1.7 A). Enzyme activity also depends on the redox potential of the solution with 50% maximum activity at –340 ± 20 mV (Figure 1.7 B). Acetylene hydratase is slightly oxygen sensitive, the [4Fe-4S] cluster degrades to a [3Fe-4S] cluster when purified under air as shown by EPR-spectroscopy (Meckenstock et al., 1999).
1,0
1,0 0,8
0,8 0,6
0,6 0,4 0,4 0,2 Relative activity Relative 0,2
0,0 0,0 B A -400 -350 -300 -250 -200
Fraction of [4Fe-4S] cluster reduced reduced cluster of [4Fe-4S] Fraction -700 -600 -500 -400 -300 Redox potential [mV] Redox potential [mV]
Figure 1.7 Redox properties of acetylene hydratase (Meckenstock et al., 1999). (A) Redox titration of the [4Fe-4S] center as determined by measuring A430. The maximal A430 was taken as totally oxidized, the minimal as totally reduced which is equal to an absolute difference (∆A430) of 0.33. Enzyme prepared under N2/H2. (B) Dependence of acetylene hydratase activity on the redox potential. Enzyme prepared under N2/H2.
BLASTP searches in GenBank showed that acetylene hydratase shares clearly the highest similarity with a putative molybdopterin oxidoreductase of the hyperthermophilic archaeon 14 Introduction
Archaeoglobus fulgidus, with a sequence identity of about 36% (57% similarity) and a sequence of the low GC Gram positive bacterium Desulfitobacterium hafniense (TIGR microbial database), which show with 37% (56% similarity) a slightly higher sequence identity for the most similar of eight high scoring sequences from this organism than the one from Archaeoglobus (Abt, 2001).
1.6 Pelobacter acidigallici transhydroxylase
Pelobacter acidigallici strain MaGal2 (DSM 2377) is a strictly anaerobic, chemoorganothroph, and gram-negative bacterium that ferments gallic acid, pyrogallol, phloroglucinol, and 2,4,6,- trihydroxybenzoic acid to three molecules of acetate (plus CO2; Schink and Pfennig, 1982; Hille et al., 1999). It was isolated from black, anaerobic marine mud of Rio Marin, a channel about 2.5 m wide and 70 cm deep, located in the city of Venice, Italy. The cells are rod-shaped with 0.5 – 0.8 x 1.5 – 3.5 µm in size. The DNA base ratio is 51.8% ± 0.4 mol% G + C (Schink and Pfennig, 1982).
1.6.1 Metabolism of gallic acid by Pelobacter acidigallici
Aerobic degradation of aromatic compounds involves oxygenase reactions in the primary attack on the mesomeric ring structure. In the absence of dioxygen, the stability of the aromatic nucleus is often overcome by a reductive attack (Evans, 1977; Reichenbecher et al., 1994). Trihydroxybenzenes are common intermediates formed in the degradation of plant materials such as glycosides, flavonoids, tannins, and lignin (Brune et al., 1992). In P. acidigallici gallic acid (3,4,5-trihydroxybenzoic acid) is decarboxylated to pyrogallol and subsequently trans- formed to phloroglucinol in a unique reaction through transhydroxylase (Figure 1.8). Although this hydroxyl transfer between two aromatic compounds is no net redox reaction, the substrate pyrogallol is oxidized in position 5 and the cosubstrate 1,2,3,5-tetrahydroxybenzene is reduced in 18 position 2. Recently it was shown, by incubation with OH2, that there is no oxygen transfer from water in the transhydroxylase reaction, and that the hydroxyl groups are transferred only between the phenolic substrates (Reichenbecher and Schink, 1999). With this, transhydroxylase differs fundamentally from all known hydroxylating molybdenum enzymes, which derive their hydroxyl groups from water. Phloroglucinol, the product of the transhydroxylase reaction, undergoes reductive dearomatization (Schink and Pfennig, 1982; Introduction 15
Brune and Schink, 1990) and subsequent hydrolytic cleavage to 3-hydroxy-5 oxohexanoate that is oxidized and thiolytically cleaved to three acetyl-CoA molecules (Brune and Schink, 1992). COOH
HO OH OH Gallic acid
Gallate decarboxylase -CO2 *OH
Pyrogallol
HO OH HO OH OH Transhydroxylase OH
OH OH
Phloroglucinol
HO OH HO OH *OH
1,2,3,5-Tetrahydroxybenzene - 3 CH3COO
Figure 1.8 Transhydroxylase reaction in the pathway of gallic acid degradation in Pelobacter acidigallici (Brune and Schink, 1992).
1.6.2 Molecular properties of transhydroxylase
Transhydroxylase (pyrogallol:phloroglucinol hydroxyltransferase, E.C. 1.97.1.2) is a hetero- dimeric enzyme, with a molecular mass of 133.3 kDa, composed of a 100.4 kDa and a 31.3 kDa subunit. It contains 11.56 ± 1.72 Fe, 0.96 ± 0.21 Mo (atomic absorption spectroscopy), and 13.13 ± 1.68 acid labile sulfur per heterodimer. Furthermore, two molybdopterin guanine dinucleotide per heterodimer had been postulated (Reichenbecher et al., 1994; Reichenbecher et 16 Introduction
al., 1996; Baas and Rétey, 1999). The isoelectric point is 4.1, the specific activity of transhydroxylase is highest at pH 7.0, and the temperature optimum is between 53 and 58°C (Reichenbecher et al., 1994). Sequence analyses showed that transhydroxylase belongs to the family of the DMSO-reductases (Baas and Rétey, 1999). In all members of this family the coenzyme is a dimeric molybdopterin guanine dinucleotide (Kisker et al., 1999). In contrast to most enzymes of the DMSO-reductase family, transhydroxylase has neither an α,β,γ structure nor a signal sequence and is not anchored in the cell membrane. While the large subunit has relatively few cysteines that are not clustered, the small subunit has 13 cysteines, some of which are clustered. This makes it likely that the Fe- S centers are located on the small subunit, while the entire MGD cofactor is associated with the large subunit. EPR studies showed that there must be at least two different types of [4Fe-4S] centers. Furthermore, the existence of [2Fe-2S] sites could not be excluded (Kisker et al., 1999). Figure 1.9 shows experimental and simulated EPR spectra (9.5 GHz, 14 K, 0.6 mW) of the Fe-S centers in dithionite-reduced transhydroxylase.
Introduction 17
g = 2.08 2.057 1.98 1.95 1.94 1.87
A B
C
D
3200 3400 3600 3800 Magnetic field [G]
Figure 1.9 EPR spectra of dithionite-reduced transhydroxylase from P. acidigallici. X-band, 14 K, 0.6 mW microwave power; (Kisker et al., 1999) (A) Enzyme as isolated under exclusion of air; 4.3 mg ml-1 in 40 mM TEA buffer, 300 mM NaCl pH 7.3, after addition of 3.5 equivalents sodium dithionite. (B) Combined simulations, with signals C and D, weight 2:3. (C) Simulated spectrum, with gx = 1.874, gy = 1.953, gz = 2.057. (D) Simulated spectrum, with gx = 1.869, gy = 1.940, gz = 2.080.
1.7 Scope of the study
In the world of molybdenum and tungsten containing enzymes, Pelobacter acetylenicus W- acetylene hydratase and Pelobacter acidigallici Mo-transhydroxylase catalyze two unusual reactions. Both reactions represent no net redox chemistry although in the case of transhydroxylase the substrate is oxidized and the cosubstrate is reduced at the same time. To achieve a better understanding of the mechanisms of these reactions, and to get a deeper insight into the active sites of these novel molybdopterin enzymes, experiments have been performed aiming at the three-dimensional structure and also structures with substrate and inhibitors bound and a detailed biochemical and spectroscopic picture of the metal centers.
18 Introduction
In the first place, this required an efficient purification with high yields of the two metallo- enzymes for getting well diffracting crystals. In the case of transhydroxylase, the replacement of molybdenum by tungsten has been attempted which might help in solving the phase problem for the X-ray structure. Former crystallization conditions have to be improved to get well diffracting crystals. A new technique to vary the humidity of crystals to get better reflection patterns and resolutions while X-ray experiments was also tested. In the case of acetylene hydratase first crystallization experiments in presence and absence of dioxygen has to be performed to obtain suitable crystals for X-ray experiments to solve the three- dimensional structure.
Materials and Methods 19
2. Materials and Methods
2.1 Chemicals and biochemicals
If not specified chemicals were of p.a. quality and obtained from Merck (Darmstadt), Riedel-de Haën (Seelze), or Fluka (Buchs, CH). Other chemicals, at least in p.a. quality, were purchased from other manufacturers:
Buffers: Roth, Karlsruhe: Tris (Tris-(hydroxymethyl)-aminomethane), HEPES (4-[2-Hydroxyethyl]- piperazine-1-[ethanesulfonic acid]). Sigma, Deisenhofen: MOPS 3-Morpholinopropanesulfonic acid,.
Enzymes: Serva, Heidelberg: Bovine serum albumin (BSA). Boehringer, Mannheim: Yeast Alcohol dehydrogenase (400 U mg-1).
Dyes: Serva, Heidelberg: Coomassie-brilliantblue G-250, bromphenolblue (sodium salt).
Reagents: Sigma, Deisenhofen: BCA (bicinchoninic acid solution). Boehringer, Mannheim: NADH (nicotinamide adenine dinucleotide). Riedel-de-Haën, Seelze: Sodium dithionite; Potassium dihydrogenphosphate; Magnesium chlorid-hexahydrate; Calcium chloride dihydrate; Potassium chloride; Hydrochloric acid 37.5%; 1-Butanol; Sodium hydroxide; Potassium hydroxide; Sodium carbonate dihydrate; Sodium acetate, Sodium chloride
Gas: Messer Griesheim, Krefeld: Argon 5.0, Helium 4.6, Acetylene 2.6, Hydrogen 5.0. Sauerstoff- werk Friedrichshafen: Nitrogen 5.0, N2/CO2 (8:2, v/v), N2/H2 (94:6, v/v).
20 Materials and Methods
Proteinstandards: BioRad, München: Low molecular mass standards (PAGE). Sigma, Deisenhofen: Gel filtration molecular mass markers.
Crystal screen solutions: Crystal screen solutions were obtained from Hampton Research (Laguna Hills, USA). House Factorial Screens were obtained from the Max Planck Institut für Biochemie in Martinsried.
2.2 Organisms
2.2.1 Pelobacter acetylenicus
Pelobacter acetylenicus strain WoAcy 1 was provided by Prof. Dr. B. Schink, Universität Konstanz. The strain is deposited in Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ), Braunschweig, under the number 3246.
2.2.2 Pelobacter acidigallici
Pelobacter acidigallici strain MaGal 2 was provided by Prof. Dr. B. Schink, Universität Konstanz. The strain is deposited in the DSMZ, under the number 2377.
2.3 Cultivation of bacteria
2.3.1 Pelobacter acetylenicus
Batch cultures of P. acetylenicus (0.1, 1, 20 or 50 l) were grown in freshwater medium (Table
2.1) at 30°C. The medium was sterilized at 121°C, cooled under a N2/CO2 (8 : 2, v/v) atmosphere, buffered with 30 mM NaHCO3, and reduced with Na2S. After addition of trace element solutions (Table 2.2, 2.4) and vitamin solution (Table 2.3) the pH was adjusted to 7.0 – 7.4 with 1 M HCl. The redox potential of the medium was monitored by the indicator resazurin (≈ 1 µM). Cultures were inoculated with 10% (by vol.) of a stock culture. The substrate acetylene was continuously supplied with 5 – 10 kPa. Growth was monitored by measuring the Materials and Methods 21
optical density at 578 nm, and the pH of the medium was maintained at pH 7.0 with 2 M
Na2CO3. Cells were harvested at the end of the exponential growth phase after one day
(A578 = 0.6) with a Pellicon ultrafiltration unit (cutoff 100 kDa, Millipore Corporation, Eschborn). The concentrate was centrifuged at 10000 g (30 min, 4°C) and the resulting cell pellets were stored in liquid nitrogen prior to use. Figure 2.1 shows the 50 l batch culture system.
ingredients/ inoculum
C2H2 and / or
N2 / CO2
samples
Figure 2.1 The 50 l batch culture system.
´
22 Materials and Methods
Compound [mM] [g l-1]
NaCl 17.1 1.0
MgCl2 ⋅ 6H2O 2.0 0.4
KH2PO4 1.5 0.2
NH4Cl 4.7 0.25
KCl 6.7 0.5
CaCl2 ⋅ 2H2O 1.0 0.15
Table 2.1 Freshwater medium (Schink, 1985). The compounds were dissolved in water and sterilized at 121°C.
Compound [mM] [mg l-1]
FeCl2 ⋅ 4H2O 7.54 1500
ZnCl2 0.51 70
MnCl2 ⋅ 4H2O 0.79 100
CoCl2 ⋅ 6H2O 0.80 190
CuCl2 ⋅ 2H2O 0.01 2
NiCl2 ⋅ 6H2O 0.10 24
H3BO3 0.10 6
Table 2.2 Modified trace element solution SL10 (Widdel and Pfennig, 1981). The components were dissolved in 10 ml of 25% HCl, the volume adjusted to 1 l, and sterilized at 121°C.
Materials and Methods 23
Compound [µM] [mg l-1]
4-Aminobenzoic acid (Vitamin H1) 365 50
(+)-Biotin (Vitamin H) 41 10
D-Pantothenic acid Calcium salt 52 25
Cyanocobalamin (Vitamin B12) 37 50
Nicotinic acid (Vitamin B) 812 100
Pyridoxamine dihydrochloride 965 250
Thiamine hydrochloride (Vitamin B1) 148 50
Table 2.3 Vitamin solution (Widdel and Pfennig, 1981). Vitamins were dissolved in water and sterilized by filtration (0.2 µm).
Name Compound [µM] [mg l-1]
Na2S solution Na2S ⋅ 9H2O 540000 120000
NaOH 125000 5000
Selenite solution Na2SeO3 ⋅ 5H2O 11 3
NaOH 12500 500
Molybdate solution Na2MoO4 ⋅ 2H2O 1000 242
NaOH 12500 500
Tungstate solution Na2WO4 ⋅ 2H2O 400 132
NaOH 12500 500
Table 2.4 Special trace element solutions (Widdel, 1980; modified). Appropriate dilutions were done with water and were sterilized at 121°C.
24 Materials and Methods
P. acetylenicus was cultivated in freshwater medium (Table 2.1) as described above. Additional ingredients were added according to table 2.5.
Compound [ml l-1] concentration
NaHCO3 (1 M) 30 30 mM
Trace element solution (Table 2.2) 1
Selenite solution (11 µM, Table 2.4) 1 11 nM
Tungstate solution (400 µM, Table 2.4) 2 800 nM
Molybdate solution (100 µM, Table 2.4) 0.06 6 nM
Vitamine solution (Table 2.3) 1.0
Na2S solution (Table 2.4) 2 1 mM
Table 2.5 P. acetylenicus tungstate cultivation; additional ingredients.
2.3.2 Pelobacter acidigallici a, Molybdate cultivation Batch cultures of P. acidigallici (0.1, 1, 20 or 50 l) were grown at 30°C in bicarbonate-buffered, sulfide-reduced saltwater mineral medium (Table 2.6, 2.7) in an 80% N2 – 20% CO2 atmosphere (Brune and Schink, 1990). After addition of trace element solutions (Table 2.2, 2.4), and vitamin solution (Table 2.3) the pH was adjusted between 7.2 and 7.4 with 1 M HCl. The redox potential of the medium was monitored by the indicator resazurin (1 µM). Cultures were inoculated with 10% (v/v) of a stock culture. The substrate gallic acid was dissolved in water under exclusion of dioxygen, neutralized to pH 7.0 with concentrated NaOH, sterilized by filtration (0.2 µm), and fed at the start (5 mM) and twice (5 mM) during cultivation. Growth was monitored by measuring the optical density at 578 nm, and the pH of the medium was maintained at pH 7.2 with 2 M Na2CO3. Cells of a 50 l batch culture (Figure 2.1) were harvested at the end of the exponential growth phase after one day (A578 = 0.8) with a Pellicon ultrafiltration unit (cutoff 100 kDa, Millipore Corporation, Eschborn). The concentrate was centrifuged at 10000 g (30 min, 4°C) and the resulting cell pellet was stored at –70°C prior to use. In Figure 2.1 the 50 l batch culture system is shown. Materials and Methods 25
Compound [mM] [g l-1]
NaCl 342 20.0
MgCl2 ⋅ 6H2O 15 3.0
KH2PO4 1.5 0.2
NH4Cl 4.7 0.2
KCl 6.7 0.5
CaCl2 ⋅ 2H2O 1.0 0.15
Table 2.6 Saltwater medium (Brune and Schink, 1990). The components were dissolved in water and sterilized at 121°C.
Compound [ml l-1] concentration
NaHCO3 (1 M) 30 30 mM
Trace element solution (Table 2.2) 1
Selenite solution (11 µM, Table 2.4) 1 11 nM
Molybdate solution (100 µM, Table 2.4) 1.5 150 nM
Vitamin solution (Table 2.3) 0.5
Na2S solution (Table 2.4) 2 1 mM
Gallic acid solution pH 7 (500 mM) 14 7 mM
Table 2.7 P. acidigallici molybdate cultivation; additional ingredients. b Tungstate cultivation P. acidigallici was cultivated in saltwater medium (Table 2.8) as described above. Additional ingredients were added according to table 2.10. To replace molybdenum by W, the culture was transferred at least 6 times in medium containing 400 nM W and 6 nM Mo. Cells of a 50 l batch culture (Figure 2.1) were harvested after 2 days (A578 = 0.6).
26 Materials and Methods
Compound [ml l-1] concentration
NaHCO3 (1 M) 30 30 mM
Trace element solution (Table 2.2) 1
Selenite solution (11 µM, Table 2.4) 1 11 nM
Molybdate solution (100 µM, Table 2.4) 0.06 6 nM
Vitamin solution (Table 2.3) 0.5
Na2S solution (Table 2.4) 2 1 mM
Gallic acid solution pH 7 (500 mM) 14 7 mM
Tungstate solution (400 µM, Table 2.4) 1 400 nM
Table 2.8 P. acidigallici tungstate cultivation; additional ingredients.
2.4 Glycerol cryo cultures
A microscopically pure overnight 1 l culture of P. acetylenicus or P. acidigallici was harvested by centrifugation at 10000 g (30 min, 4°C). Sterile glycerol solution (80% v/v, in water) was added to the cell pellet (1:1 w/w) and mixed to homogeneity. Aliquots of 500 µl were taken and slowly frozen at –20°C for 24 hours. Final storage was done at –70°C. The frozen cells were tested for growth after 2 weeks storage at –70°C by inoculating a 100 ml batch culture with one 500 µl aliquot.
2.5 Enzyme purification
All chromatographic steps were performed with a Pharmacia FPLC system (pump P-500, gradient controller GP-250, Pharmacia Biotech, Freiburg). Detection was carried out at 280 nm and 405 nm (Uvicord S II, Pharmacia Biotech, Freiburg). Centrifugation was done in a RC 5C centrifuge (Sorvall Instruments, Du Pont de Nemours, Bad Homburg), or in a OptimaTM LE-80K ultracentrifuge (Beckman Instruments Inc., Palo Alto, USA) at 4°C. Materials and Methods 27
2.5.1 Acetylene hydratase
Acetylene hydratase was purified at room temperature (Meckenstock et al., 1999) in an anaerobe chamber (Coy, Grasslake, Michigan, USA; 94% N2, 6% H2), equipped with a Palladium catalyst
(Typ K-0242, 0.5% Pd on Al2O3, ChemPur, Karlsruhe) in order to remove traces of dioxygen, an automatic airlock (Coy), and an oxygen/hydrogen gas analyzer (Coy). The content of dioxygen in the anaerobe chamber was 0 – 10 ppm. Dioxygen from buffers was removed by 8 to 10 cycles of vacuum and flushing with argon according to Beinert et al. (1978). Traces of dioxygen were removed from argon by a catalyst type R3-11 (BASF, Ludwigshafen). All glass and plasticware as well as buffers were stored in the anaerobe chamber for at least 24 hours prior to use in order to equilibrate with temperature and atmosphere. The frozen cell suspension was thawed at 30°C in the anaerobe chamber, and the density was adjusted to A578 = 135 with 50 mM Tris/HCl pH 7.5. Cells were lysed with lysozyme (0.6 mg/ml, 2 mM EDTA) for 30 min at room temperature. DNA was digested with DNase I in the presence of 10 mM MgCl2 for 30 min. The suspension was centrifuged at 10000 g for 30 min and the supernatant (crude extract) was subjected to (NH4)2SO4 precipitation. In the first precipitation step, 4 M (NH4)2SO4 in water was added slowly to a final concentration of 2.3 M. The solution was stirred on ice for 30 min. After centrifugation (30 min, 10000 g) acetylene hydratase was precipitated from the supernatant by a further 4 M (NH4)2SO4 addition to a final concentration of 3.2 M and stirring on ice for 30 min. After another centrifugation step the pellet was dissolved in about 10 ml of 50 mM Tris/HCl pH 7.5 and the saltconcentration was also diluted. The solution was centrifuged at 10000 g for 5 min and was loaded on a Q-sepharose (instead on a Mono Q anion-exchange chromatography column HR 10/16, Pharmacia Biotech, Freiburg) anion-exchange chromatography column equilibrated with 50 mM Tris/HCl pH 7.5. The column was developed with a linear 0 – 0.5 M NaCl gradient. Active fractions were pooled and concentrated to 1.5 ml, using Centicon centrifugal filter devices (YM-30, Millipore, Eschborn). The concentrate was loaded on a HiLoad® 16/60 Superdex 75 column (1.6 cm x 60 cm, Pharmacia Biotech, Freiburg), equilibrated with 50 mM Tris/HCl 200 mM NaCl pH 7.5 and eluted with the same buffer. Active fractions were pooled and concentrated with Centricon centrifugalfilter devices. The pure acetylene hydratase was stored in liquid nitrogen or, under exclusion of dioxygen, in a gas-tight bottle at 4°C.
28 Materials and Methods
2.5.2 Transhydroxylase
Transhydroxylase was prepared at 5°C according to Sommer (1995) and Reichenbecher et al. (1994) with some modifications. Frozen cells were thawed at 30°C in a water bath and suspended in 50 mM triethanolamine
(TEA) pH 7.5. Some DNase I/MgCl2 was added, and the cells were disrupted by 3 passages through a French Press (Aminco 1-FA-073, SLM Instruments, Urbana, Illinois, USA) at 137 MPa. The suspension obtained was centrifuged (13200 g, 30 min, 4°C) in order to remove rough material. The resulting supernatant was centrifuged at 100000 g (60 min, 4°C) to separate the membrane fraction (pellet) from the soluble fraction (supernatant = crude extract). The soluble fraction was kept on ice or frozen at –70°C prior to use. The crude extract was passed through a 100 ml DE 52 anion exchange column ( Whatman, UK) but was replaced with a Q-Sepharase anion exchange column previously equilibrated with 50 mM TEA pH 7.5. After loading, the column was flushed with three bed volumes of buffer containing 175 mM NaCl. A five bed volume linear gradient of buffer with increasing NaCl concentration from 175 mM to 500 mM eluted transhydroxylase. Fractions showing transhydroxylase on a SDS-PAGE gel were pooled and concentrated to a final volume of 2 ml by ultrafiltration (30 kDa cutoff) in a stirring cell (Amicon), loaded on a HiLoad® 26/60 Superdex 200 column (2.6 cm x 60 cm, Pharmacia Biotech,Freiburg),and elutedwith 50 mM TEA 200 mM NaCl pH 7.5 at a flow rate of 1 ml min-1. Active fractions were pooled, concentrated by ultrafiltration (30 kDa cutoff), and stored at –70°C.
2.6 Enzyme activity
2.6.1 Acetylene hydratase
Enzyme activity was measured photometrically in a coupled assay with alcohol dehydrogenase (Meckenstock et al., 1999; Rosner and Schink, 1995). The reaction was performed in glass cuvettes sealed with rubber stoppers under N2/H2. In a typical assay, 10 µl acetylene hydratase solution were added to 960 µl of 1.5 mM titanium (III) citrate in 50 mM Tris/HCl pH 7.5 (Zehnder and Wuhrmann, 1976). After addition of 20 µl of 10 mM NADH and 10 µl of 2000 U ml-1 yeast alcohol dehydrogenase (Y-ADH), the reaction mixture was incubated for 10 min at 50°C. The reaction was started by addition of 1 ml acetylene to the gas phase. Depletion of NADH was followed at 365 nm. Activity was calculated using Beers Law and Materials and Methods 29
-1 -1 ε365 (NADH) = 3.4 mM cm (Ziegenhorn et al., 1976). 1 µmol acetylene converted to acetaldehyde per min corresponds to 1 Unit (U).
2.6.2 Transhydroxylase
(a) Assay of transhydroxylase activity
Enzyme activity was assayed at 25°C in the anaerobe chamber (Chapter 2.5.1.) equipped with a cooler (Coolmatic RA-40, Mobitronic Gleichrichter EPS-100W, Waeco, Emsdetten). A discontinuous test system (Reichenbecher et al., 1994; Brune and Schink, 1990) was used. The reaction was performed in Eppendorf reaction tubes. In a typical assay 775 µl of 100 mM potassium phosphate buffer (KPi), pH 7.0 were mixed with 100 µl pyrogallol (100 µM in KPi), 100 µl 1,2,3,5-tetrahydroxybenzene (100 µM in KPi), and incubated for one minute. The assay was started by addition of 25 µl transhydroxylase solution. Aliquots (100 µl) were taken after
3 and 5 min incubation and immediately added to 100 mM H3PO4 (400 µl). Samples were kept on ice and analyzed quantitatively for pyrogallol and phloroglucinol by Reverse Phase HPLC within 1 h.
(b) Substrate and product analysis
Samples were analyzed by High Pressure Liquid Chromatography (S100 Solvent Delivery System, S5110 Injector Value System, S8100 Low Pressure Gradient Mixer, S3300 UV Detector, S4010 Column Thermo Controller, Sykam, Gilching; Lambda 1000 UV Detector, Bischoff, Leonberg) equipped with a Reverse Phase C18 column (Gromsil GSOD50512S2504, Grom Analytik, Herrenberg) at 40°C, in a methanol/phosphate (12.5% methanol, by vol.; 100 mM KPi pH 2.6) solvent system. Samples (20 µl) were injected and eluted at a flow rate of 1.5 ml min-1. Aromatic compounds were detected with a Sykam S 3300 UV detector (Sykam, Gilching) at 205 nm. Data were analyzed by computer programs (AXXI-Chrom 727, Version 3.8; AXXIOM Chromatography, Calabasas California, USA) and quantified by comparison with external and internal standards of known composition. Peak identification was performed by comparison of retention times and UV spectra with those of standard samples.
30 Materials and Methods
2.6.3 Alcohol dehydrogenase
Enzymatic activity was assayed photometrically by measuring the disappearance of NADH at
365 nm. The reaction was performed in glass cuvettes sealed with rubber stoppers under N2/H2.
Yeast alcohol dehydrogenase (Y-ADH)
In a typical assay 915 µl of 1.5 mM titanium (III) citrate in 20 mM Tris/HCl pH 7.0 were added to 25 µl of 10 mM NADH and 10 µl of 40 U ml-1 Y-ADH. The reaction mixture was incubated at an appropriate temperature for 10 min. The reaction was started by addition of 50 µl of acetaldehyde. Disappearance of NADH was followed at 365 nm. Activity was calculated using -1 -1 Beers Law and ε365 (NADH) = 3.4 mM cm (Ziegenhorn et al., 1976).
2.7 UV/Vis spectroscopy
UV/Vis spectra were obtained on a Cary 50 spectrometer (Varian, Darmstadt), a Lambda 16 spectrometer (Perkin Elmer) or a HP 8452A diode array spectrophotometer (Hewlett Packard, Stuttgart), The Cary 50 spectrometer was equipped with a temperature control unit (80% glycerol/water, v/v), allowing temperatures up to 80°C.
2.8 Analytical methods
2.8.1 ICP-MS
Inductively coupled plasma mass spectrometry (ICP-MS) was performed by Spurenanalytisches Laboratorium Dr. Baumann (Maxhütte-Haidhof). The concentrations of iron, molybdenum, and tungsten were determined by ICP-MS in different samples containing acetylenehydratase and transhydroxylase (100 µl, ≈ 2 mg ml-1). A buffer sample served as a control.
2.8.2 Protein
Protein was determined using bicinchoninic acid (BCA) (Smith et al., 1985). This method is based on the Biuret reaction (Goa, 1994). Bovine serum albumin was used as a standard. The Materials and Methods 31
concentration of BSA standard stock solutions was photometrically determined with -1 -1 -1 ε279nm = 0.667 (mg ml ) cm (Foster and Sterman, 1956). 100 µl of unknown or standard proteins (5 − 20 µg) were mixed with 1 ml of a solution containing 50 : 1 (v/v) BCA and
CuSO4 · 5H2O (4%, w/v). The reaction mixture was incubated for 20 min at 60°C. The calibration curve was recorded on a Hewlett Packard diode array spectrophotometer (Hewlett Packard, Waldbronn) and the absorbance was monitored at 562 nm.
2.8.3 Polyacrylamide gel electrophoresis
SDS-PAGE was carried out with the Mighty Small System (SE 250, 100 x 80 x 0.75 mm, Hoefer Scientific Instruments, San Francisco, USA) according to Lämmli (1970). 10 – 15% acrylamide gels were used. The molecular mass of the subunits was estimated using low molecular mass markers (BioRad Laboratories, München). Gels were stained with coomassie (Zehr et al., 1989) or silver (Rabilloud, 1990; Heukeshoven and Dernick, 1985).
2.9. Crystallography
2.9.1 Theoretical background
The maximal achievable resolution of any microscopic technique is limited by the applied wavelength. The radiation needed to analyze atomic distances (e.g. 1.54Å for a carbon-carbon σ- bond) lies within the spectral range of X-rays. Max von Laue realized in 1912 (Glusker et al., 1994)that the three-dimensional ordered lattice arrangement of a crystal will cause interference of the diffracted photons resulting in discrete maxima whose intensity can be measured in an appropriate experimental setup. 32 Materials and Methods
2.9.2 Crystal growth
Crystallization is one of several means (including nonspecific aggregation/precipitation) by which a metastable supersaturated solution can reach a stable lower energy state by reduction of solute concentration (Weber, 1991). The general processes by which substances crystallize are similar for molecules of both microscopic (salts and small organics) and macroscopic (proteins, DNA, RNA) dimensions. There are three stages of crystallization common to all systems: nucleation, growth, and cessation of growth. Nucleation is the process by which molecules or noncrystalline aggregates (dimers, trimers, etc.) which are free in solution come together in such a way as to produce a thermodynamically stable aggregate with a repeating lattice. Crystallization is known to lower the free energy of proteins by ~3-6 kcal/mole relative to the solution state (Drenth and Haas, 1992). The formation of crystalline aggregates from supersaturated solutions does not however necessitate the formation of macroscopic crystals. Instead, the aggregate must first exceed a specific size (the critical size) defined by the competition of the ratio of the surface area of the aggregate to its volume (Feher and Kam, 1985; Boistelle and Astier, 1988). Once the critical size is exceeded, the aggregate becomes a supercritical nucleus capable of further growth. If the nucleus decreases in size so that it is smaller than the critical size, spontaneous dissolution will occur. The process of formation of nonspecific aggregates and noncrystalline precipitation from a supersaturated solution does not involve the competition between surface area and volume (n-mers add to the aggregate chain in a head to tail fashion forming a linear arrangement), and thus generally occurs on a much faster time scale than crystallization. The degree to which nucleation occurs is determined by the degree of supersaturation of the solutes in the solution. The extent of supersaturation is in turn related to the overall solubility of the potentially crystallizing molecule. Higher solubility allows for a greater number of diffusional collisions. Thus, higher degrees of supersaturation produce more stable aggregates (due to higher probability of collision of diffusing molecules) and therefore increase the likelihood of the formation of stable nuclei. In the case of a finite number of solute molecules, this condition generally results in the production of a large number of small crystals. At lower solute concentrations the formation of individual stable nuclei increases in rarity, thus favoring the formation of single crystals. According to Periodic Bond Chain theory, (Boistelle and Astier, 1988), three different types of growth faces exist: flat faces, stepped faces, and kinked faces. Flat faces require two dimensional nucleation (the formation of growing sheets of molecules) in order to induce growth, and thus Materials and Methods 33
grow the slowest. Stepped faces grow as columns of molecules, which requires only one dimensional nucleation, and thus have intermediate growth rates. Stepped faces typically occur as a result of a crystallographic screw axis causing spiral growth patters to occur at the surface of the crystal. Finally, kinked faces are growth sites which do not require nucleation to promote further growth, and therefore grow faster than the other two face types. Thus, the type of the growing crystal face (flat, stepped, or kinked) strongly influences the rate at which crystal growth occurs. The growth of crystals from nuclei is also strongly influenced by diffusional and convection effects. As with nucleation, increased solubility results in increased growth rates. Again, this is a function of the rate at which protein molecules reach the growing surface of the protein crystal. Feher and Kam (1985), through the use of ultraviolet microscopy, have been able to demonstrate the regions surrounding growing crystals to be lowered in protein concentration relative to the surrounding solution .The rate of diffusion of proteins in and out of these halos around the growing crystal provides a growth limiting factor. Cessation of growth of crystals can occur for a multitude of reasons. The most obvious is the decrease in concentration of the crystallizing solute to the point where the solid and solution phases reach exchange equilibrium. In this case, the addition of more solute can result in continued crystal growth. However, some crystals reach a certain size beyond which growth does not precede irrespective of solute concentration. This may be a result either of cumulative lattice strain effects or poisoning of the growth surface. Lattice strain effects in tetragonal lysozyme crystals have been demonstrated (Feher and Kam, 1985). Halved crystals of hen egg white lysozyme, when placed in fresh crystallization solutions, grew to the exact same size as the original crystal. This suggests that the long range propagation of strain in the lattice effectively prevents addition of molecules to the surface once a certain critical volume is reached. Crystals affected by lattice strain are therefore inexorably size limited. Solubility of the protein, C, can be described as a function of the ionic strength I in the precipitant solution:
AZ 2 I lnC = lnC 0 + − KsI 1+ aB I
C0 is the solubility of protein in water, I is the ionic strength, Z is the total charge of the protein, a is the sum of the radii of protein and salt ion, Ks is an empirical salting out constant and A and B are constants depending on temperature and dielectricity. For high salt concentrations, the salting out term will be relevant, and it can be derived that ions with a high charge density will have stronger influence on the solubility, as described in the hofmeister series in 1888: 34 Materials and Methods
Ks for anions: citrate> tartrate> sulfate> acetate> chloride> nitrate + + + 2+ 2 2+ 3+ KS for cations: Li > K > NH4 > Ca > Sr +> Ba > Al In addition to salts, commonly used precipitants are polyethylene glycols or organic solvents like ethanol, isopropanol or methylpentane diol.
2.9.3 Crystals
A crystal can be regarded as a three-dimensional repetition of a single building block, the unit cell. Within the unit cell, a crystal can contain further symetrie elements dividing into several asymmetric units which form the most basic structural element within the crystal. The geometrie of the unit cell together with the possible symmetry operations defines the space group of the crystal. Although there are 230 space groups in seven crystal systems (triclinic, monoclinic, orthorhombic, tetragonal, trigonal, hexagonal and cubic), only 65 are enantiomorphic and are thus fesiable for chiral molecules such as proteins. Identification of the correct space group is essential for correct indexing of diffraction patterns and therefore the first step to get to a crystal structure.
2.9.4 X-ray diffraction by crystals
Upon interaction with the atoms in a crystal, the oscillating electrical field of an X-ray photon induces an oscillation of equal frequency in the electron hull of the atom. The electrons act as oscillating dipoles emitting secondary radiation of the same frequency as the incident radiation, but with a phase difference of 180°. In this elastic or coherent diffraction, the phase shifts between single waves originating from any point of 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). Given the correct orientation of the crystal, this condition is fulfilled for corresponding positions in all unit cells. Diffraction of X-rays on the real lattice of a crystal thus creates another three-dimensional lattice of diffraction maxima. As the geometric properties of this lattice are inverse to those of the real crystal, it is referred to as the reciprocal lattice. A convenient way to describe diffraction by a crystal lattice is to imagine every single diffraction spot to be a reflection of the incident beam on an imaginary lattice plane which is identified by Materials and Methods 35
the Miller indices (h,k,l) (Figure 2.2a). The normal vectors S of those lattice planes then build up the reciprocal lattice, their length reflecting the reciprocal distance of the planes.
Figure 2.2 Reciprocal lattice planes and Bragg’s law. A) Lattice planes that allow for constructive I interference of diffracted waves are those that divide the unit cell sides into integer fractions. The number of those fractions is used to index the plane. The group of lattice planes shown would have the Miller indices (4 2 3). B) Two waves that are reflected by two adjacent lattice planes with distance d have a difference in path length that is equal to 2d sin θ, as it can easily be derived from the scheme. A prerequisite for constructive interference is, that this difference in path is an integer multiple of the wavelength used: 2d sin θ= nλ (Bragg’s law).
Regarding elastic diffraction on a set of lattice planes with distance d, constructive interference will occur at an angle θ, if the path difference between the diffracted waves is an integer multiple of the wavelength λ. This relation between reflection angle and lattice plane distance is known as Bragg’s law (Figure 2.2B):
2 dhkl sin θ = n λ
The Ewald sphere (Figure 2.3) is a tool for constructing reciprocal lattice points on the basis of Bragg’s law. It is a sphere of radius 1/λ with the crystal in its centre. The point where the r incident beam s0 centers the sphere and the origin O of the reciprocal lattice are on opposite sides of the centre. Bragg’s law is fulfilled for every reciprocal lattice point that lies on the Ewald sphere. A rotation of the crystal rotates the reciprocal lattice in the same way, allowing different reciprocal lattice points to intersect with the sphere. For the given orientation of the crystal, those points are the ones that can be recorded on an X-ray detector. 36 Materials and Methods
Figure 2.3 The Ewald construction. In reciprocal space, the crystal (C) is placed in the center of a sphere (here, in two dimensions, a circle) with radius 1/λ, called the Ewald sphere. The origin of the reciprocal lattice, i.e. reflection (0 0 0), is placed in (O). The reciprocal lattice (grey dots) will rotate as the crystal does and only those reciprocal lattice points that intersect with the Ewald sphere will be in diffraction condition (red dots) and will be recorded on an image plate detector in real space.
As every recorded diffraction spot represents one lattice plane (h,k,l), the measurement of the positions of the spots is sufficient to deduce the geometry of the crystal and in most cases also the space group, as additional symmetry elements can manifest in the form of systematic extinctions of reflections. On the other hand, the content of the unit cell influences the intensity of the diffraction spots. A crystal is described as the convolution of the unit cell content with the three-dimensional lattice. As the diffraction image is the result of multiplying these molecular transformations with the discrete reciprocal lattice, the intensity of a diffraction spot will be high if the intensity of the underlying convolution is high and vice versa.
The result of a data collection on a crystal will primarily be the knowledge about space group and unit cell dimensions, and – based on this – an intensity measurement I(h,k,l) for every reflection (h,k,l).
2.9.5 The electron density function
The goal of a crystallographic experiment is to calculate the distribution of electron density in the asymmetric unit of the crystal in order to be able to place an atomic model of the crystallized molecule therein. Materials and Methods 37
If the electron hull of an atom is considered spherical, its diffraction contribution, the atomic scattering factor, does not depend on the direction of the incident beam. However, because of the finite extent of the atom, the phase difference for photons diffracted at different positions in the electron hull will increase with the diffraction angle θ which corresponds to S and thus to resolution
f = ρ(x, y, z) exp(2πi(hx + ky + lz))dxdydz 0 ∫ x,y,z
A further decrease of diffraction power with increasing angle is caused by thermal movement of the atoms, by statistic disorders, absorption or scaling errors. To compensate for this, the mean square displacement u 2 of the atomic vibration is modeled as the isotropic Debye-Waller temperature factor B = 8π 2u 2 and added to the atomic scattering factor as an additional exponential decay term:
sin2 θ Form: f = f exp− B 0 2 λ
With this, the scattering of all atoms in the asymmetric unit is the sum of all atomic scattering factors, taking in account individual phase shifts. For every single reflection (h,k,l) this summation leads to a structure factor F(h,k,l):
Form:F(h, k,l) = ∑ fi exp(2πi(hx + ky + lz )) i
The reciprocal lattice is the Fourier transform of the electron distribution in the crystal, split up in the form of the structure factors. 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:
1 Form: p(x, y, z) = ∑ F(h, k,l) exp(2πi(hx + ky + lz)) V h,k,l
38 Materials and Methods
2.9.6 The phase problem
Approaching from the side of the diffraction experiment, each structure factor F(h,k,l) also represents a reflection by one lattice plane. It is described by a wave function with an amplitude and a phase angle: Form: F(h,k,l) = F(h,k,l) expiα(h,k,l) 142 431424 434 Amplitude Phase angle
The structure factor amplitude can be obtained experimentally, as it is in principle the square root of the measured intensity:
Form: I(h, k,l) ≈ F(h, k,l) 2
The total energy E(h,k,l) in a diffracted beam for a mosaic crystal rotating with uniform angular velocity ω through the reflecting position is given by Darwin’s equation (Darwin, 1914):
4 2 I 0 3 e 1 + cos 2θ LAVx 2 Form: E(h, k,l) = λ 3 4 2 F(h, k,l) ω me c 2 V
Herein, I0 is the intensity of the incident beam, λ is the wavelength, e and me are charge and mass of the electron and c is the velocity of light. The term 1 + cos2 2θ / 2 is a polarization factor that originates from the fact that an electron does not scatter along its direction of vibration. The Lorentz factor L depends on the data acquisition technique and the absorption factor A takes into account the intensity loss through photoelectric absorption and Rayleigh scattering. Vx is the crystal volume and V is the volume of the unit cell. While the structure factor amplitude can be derived from the measured intensity as described above, information about the phase angle is lost. Without correct phase angles, the calculation of an interpretable electron density is impossible, a dilemma commonly referred to as the phase problem of crystallography.
Materials and Methods 39
Four approaches to overcome this problem are applicable today: • Molecular Replacement (MR) • Multiple Isomorphous Replacement (MIR) • Multiple-wavelength Anomalous Dispersion (MAD) • Direct Methods The method of Molecular Replacement depends on the availability of a sufficiently homologous model structure which is oriented by Patterson search techniques and then used for initial phase calculations (Hoppe, 1957; Huber, 1965; Rossmann and Blow, 1962). Without any previous knowledge of the structure, Multiple Isomorphous Replacement is still the most commonly used method. Herein it is attempted to place heavy atoms on specific sites in the protein either by soaking or by cocrystallization and to identify their positions by comparing the data collected from a native crystal with that of a derived one (Green et al., 1954). MAD depends on precisely tunable synchrotron radiation which has only been available for the last years, but is becoming more and more standard (Hendrickson et al., 1988). Direct methods are the common way to determine phases in small molecule crystallography, but due to the high number of atoms per asymmetric unit and the limited resolution that is obtained from most protein crystals, this approach has rarely been successful for large biomolecules. Most recently the number of protein structures solved by direct methods is increasing and the development is promising, but small molecule size, high resolution and good data quality are still a prerequisite.
2.9.7 MAD with metalloproteins
If the atomic absorption coefficient of an element is plotted as a function of X-ray wavelength, a curve with several sharp edge features is obtained. On these absorption edges, the photon energy of the X-rays is sufficient to eject an electron from the atom and so the absorbed energy will not contribute to the scattering of the atom at this wavelength. A frequently encountered problem in MIR is the derivation of crystals with heavy atoms without destroying the morphology that is essential for comparing derived and native data sets. Metalloproteins per se contain heavy atoms, and measurements around the absorption edge of a specific metal can produce data in which the contribution of this metal is significantly different depending on the exact wavelength, in analogy to a native and a derivative data set. 40 Materials and Methods
Figure 2.4 An Argand diagram of the Gaussian plane. A complex wave function can be described by a rotating vector with a length A (amplitude) and an angle α (phase) with the real axis. + B) The difference between MIR and MAD. In MIR, the structure factor FH of a heavy + atom adds to the one of the protein ( FP ) producing the combined structure factor + FPH that can be measured. The same is true for the Friedel mates, which are identical except for the sign of the phase angle. If anomalous contribution is considered, an additional shift is added to he heavy atom contribution, that can be separated into a real part (∆f) and an imaginary part (i f "), and the Friedel pairs will no longer be equal. In MAD on metalloproteins, only this shift can be used, because protein and heavy atom can + not be separated. Although the anomalous contribution is small compared to FH it can be used for phasing because the data are measured on a single crystal and there is no anisomorphy problem.
In addition, it is no longer correct to consider electrons as free electrons when measuring close to an absorption edge. Inner electrons interact with the nucleus, even more so in heavy elements, and the result is a phase shift of the diffracted beam that differs from the regular 180° and is called anomalous scattering. In an Argand diagram of the Gaussian plane (Figure 2.4A), the atomic scattering vector is rotated counterclockwise, modified by a real contribution f ′ = f + ∆f and an imaginary contribution f ′′ (Figure 2.4B). These two components are related through the Kramers-Kronig equation
2 ∞ ω′f ′′(ω′) f ´(ω) = dω′ Form ∫ 2 2 π 0 ω − ω′ with the respective X-ray frequencies ω and ω’.
+ As a consequence, the structure factors of Friedel pairs, FPH (andh,k,l) = FPH
− FPH (h, k ,l ) = FPH are no longer equal and their difference
Materials and Methods 41
f ′ Form: ∆ F = (F + − F − ) ano PH PH 2 f ′′ can be used separately or in combination with isomorphous replacement to search for the anomalous scatterers’ positions. While the dispersive f ′ contribution of the absorption edge in MAD is not unequivocal (as in the case of a single derivative in MIR), the anomalous scattering information can compensate this ambiguity, such that all the information needed to solve an unknown structure can be obtained from a single crystal containing anomalous scatterers.
The first step then is to localize the atomic positions of the anomalous scatterers. While it is not possible to calculate an electron density map without phase information, the intensities
I = F 2 themselves can be used to calculate a Patterson function (Patterson, 1934), whose maxima are interatomic distance vectors rather than real-space atomic positions:
1 2 Form P(u, v, w) = ∑ F(h, k,l) cos 2π (hu + kv + lw) V h,k,l
If an anomalous difference Patterson map is calculated with ∆ F 2 as coefficients, it will only ( ano ) show maxima at the positions of distance vectors between the anomalous scatterers. Once their positions have been determined and refined, the structure factors of the protein can be derived according to
FP = FPH – FH
with the help of the Harker construction (Harker, 1956). In addition to the dispersive difference of the scattering of the metal near the absorption edge compared to the remote data set, the anomalous contribution can be used to compensate the phase ambiguity for every single reflection that a single derivative will give (Figure 2.5). 42 Materials and Methods
Figure 2.5 A Harker diagram for the deduction of protein phase angles by anomalous
scattering (after Drenth, 1994). FP is the structure factor amplitude for the reference + − (remote) dataset and FPH and FPH for the Friedel mates of the dataset with maximized + anomalous contribution (f "). The contribution of the metal to the structure factor is FH for − one member of the Friedel pair and FH for the other member. Due to the anomalous scattering component, these two structure factors are not symmetric to the horizontal − axis. For clarification, the FH vectors have been drawn with opposite phase angles such that the three circles intersect in a single point. The dashed line then indicates the direction of the nonanomalous scattering part of the heavy atoms.
Although the anomalous contribution of a metal corresponds only to a small fraction of the electrons in the heavy atom, the problem of nonisomorphy, which is always present in MIR, does not occur in MAD. All data sets can be measured from a single crystal in its native form.
2.9.8 Crystallisation under exclusion of dioxygen
Crystals were grown by the method of vapor diffusion where the protein solution was mixed with a precipitant solution and equilibrated against a higher concentrated precipitant reservoir in a closed environment. Under regular conditions, using non-volatile precipitants such as polyethylene glycol or salts, equilibrium is reached by diffusion of water from the protein drop to the reservoir, thus slowly increasing the concentration of all components in the drop (McPherson,1982). Crystals of acetylene hydratase or transhydroxylase were grown by the sitting drop vapor diffusion method (Figure 2.6) using Cryschem plates (Charles Supper Company, Natick, USA) or CrystalClear Strips (Hampton Research, Laguna Hills, USA). Materials and Methods 43
Buffers of low ionic strength were used for crystallization experiments, most commonly 5 mM HEPES/NaOH pH 7.5. Buffer exchange was carried out on NAP 5 gravity flow columns (Pharmacia Biotech, Freiburg). The protein was reduced with dithionite and Titane (III) citrate. Each drop was prepared by mixing 2 µl of the protein solution with the same amount of the crystallization solution in the well of the crystallization device and was equilibrated by vapor diffusion against 500 µl of the solution contained in the reservoir of Cryschem plates or 80 µl in the reservoir of CrystalClear Strips plates respectively. The reservoir and the protein solution in the well were sealed with transparent (crystal clear) tape. In order to screen large numbers of more complex precipitant solutions, the method of sparse matrix sampling was applied (Carter Jr. and Carter, 1979; Jancarik and Kim, 1991) to obtain promising starting conditions. Crystallization experiments of acetylene hydratase and transhydroxylase were performed under
N2/H2 (96 : 4, v/v). sealing tape sample drop
Crystal growth solution Figure 2.6 Schematic view of a sitting drop crystallization unit.
2.9.9 Cryocrystallogragphy
A commonly observed problem in protein crystallography is damaging of crystals in the X-ray beam, especially if intense synchrotron radiation is used. This damage is mainly caused by the formation of water radicals by the X-ray photons which in turn react with the protein molecules, destroying the order of the crystal lattice. To minimize crystal degradation, crystals were cooled to 100 K with a nitrogen stream cooling system (Oxford Cryosystems, Oxford, UK), reducing the mobility of solvent radicals significantly. Protein crystals are not supposed to be frozen in the nitrogen stream, as formation of ice crystals in the solvent leads to strong powder diffraction rings in the diffraction pattern. Addition of cryoprotectants helps to avoid ice formation, but as such reagents are often strongly hygroscopic, they can dehydrate and thus destroy the crystal. To successfully cool transhydroxylase and acetylene hydratase crystals for MAD data collection, 2-methyl-2,4-pentanediol (MPD) was applied. Crystals were transferred from the mother liquor 44 Materials and Methods
into a harvesting buffer containing 22% MPD and incubated for 2 minutes and the crystals showed no ice rings in the diffraction pattern.
2.9.10 Substrate and inhibitor complexes
In order to examine the binding of different substrates and inhibitors to the active site of transhydroxylase, crystals were soaked with the compounds 1,2,3-trihydroxybenzene, 1,2,3,5- tetrahydroxybenzene and 1,2,4-trihydroxybenzene under exclusion of dioxygen. After incubation for a sufficient time to allow for binding, the crystals were cooled and measured. All data sets were measured with synchrotron radiation.
2.9.11 Data collection
Measurements were taken on wiggler beamline 6 (BW6) of the German Electron Synchrotron DESY, Hamburg, on a Mar Research CCD detector and on beamline ID29 of the European Synchrotron radiation facility (ESRF) in Grenoble on a ADSC Q210 2D detector . An X-ray fluorescence scan was carried out around the K-shell absorption edge of iron to determine the optimal wavelengths for data collection. This procedure is neccessary, because although element-specific absorption edges can be calculated according to the theory of Cromer and Liberman (Cromer and Liberman, 1970), the interaction of the scattering atom with its chemical neighbours influences the scattering behaviour considerably. The absorption edge itself is shifted mainly by the oxidation state of the iron atoms, while the local chemical environment introduces a fine structure (EXAFS) to the absorption edge, caused by the interference of the emitted photoelectron with the electron hulls of the surrounding atoms (backscattering). Thus the maximum achievable f ′ can be significantly larger than the theoretical edge jump. On the other hand, the maximum achievable f ′ is smaller than the theoretical value due to limitations of the bandwidth of the X-ray source. Cryotests for transhydroxylase and acetylene hydratase and some substrate complexes of transhydroxylase were taken on a Rigaku Rotaflex rotating anode X-ray generator with
CuKα radiation at a wavelength of λCu = 1.5418 Å with Mar Research 2000 or Mar345 image plate detectors, ususally with rotation angles of 0.5° per image.
Materials and Methods 45
2.9.11 Transformation of crystals
The transformation of transhydroxylase crystals was executed on a free-mounting system. The crystals were mounted on a crystal holder with a loop which will match on a goniometerhead and the head part is freely rotatable or on micropipette.
A stream of humid air with a defined moisture content is produced (Figure. 2.7 The dew point
Tdp of the outcoming mixed gas is measured (MTR 2.12/SA, IL METRONIC Sensortechnik, Ilmenau, Germany). A flexible teflon tube transports the humid air to the crystal holder. The temperature Tcr of the crystal holder is controlled. The temperature of the air stream is adjusted to Tcr by passing through the crystal holder. The relative humidity near the probe follows from the Magnus formula,
with the constants aw = 17.5043 and bw = 241.2 K. Typically, the temperature Tcr is chosen to be at least 1-2 K above the ambient temperature. By the use of two separated valves, V1 and V2, a humidity range of 0-100% can be adjusted within ±0.3%. A personal computer controls the valves V1 and V2 via step motors. Arbitrary humidity time protocols, such as humidity jumps or defined humidity gradients, can be realised. The host computer is connected to a computer network. External processing and evaluation of humidity experiments is therefore possible. (Kiefersauer et al, 2000)
46 Materials and Methods
Figure 2.7 The main functionalities of the humidity apparatus. A line with dry air (1) is split into two; one is saturated with water by blowing onto foam rubber (2). The foam rubber is placed in a perforated plastic vessel in contact with the water reservoir (3). The other line is connected to the outlet of the vessel. The combined humid air stream has a dew point Tdp. The flow in both lines (0.8 l min¡1) is regulated with the valves V1 and V2. The water reservoir is temperature controlled; water losses are replaced by a pump. The humid air stream is connected to the crystal holder (5) via a flexible transport line (4). The tube is heated and isolated to avoid condensation. The temperature of the crystal holder, Tcr, is regulated (Industrieregler 48 _ 48, RS Components, Moerfelden-Walldorf, Germany) by a thermo-foil and a Pt100 ceramic sensor (Minco, Telemeter Electronic GmbH, Donauwoerth, Germany) with an accuracy of 0.1 K. The crystal is fully accessible for irradiation by X-rays or light (Kiefersauer et al, 2000).
With this method the crystals could be transformed anoxic in a nitrogen stream. The purpose of this technique is to improve the resolution and to lower the mosaicity of the crystals while decreasing the humidity.
2.9.12 Data analysis
For determination of the orientation of the crystal, the space group, the parameters of the unit cell and for the integration of the intensities of the reflexions the programs MOSFLM6XX (Leslie, 1991) and DENZO (Otwinowski and Minor, 1996) were used. The data reduction was done with SCALA (Collaborative Computational Project No. 4 1994), TRUNCATE (Collaborative Computational Project No. 4 1994) and SCALEPACK (Otwinowski and Minor, 1996) (Reviews: (Dauter, 1999) (Leslie 1999; Rossmann and van Beek, 1999)).
Materials and Methods 47
Selfrotation functions were executed with the program GLRF (Tong und Rossman, 1990) . Patterson search was done with MOLREP (Collaborative Computational Project No. 4, 1994) und CNS (DeLano and Brünger, 1995) (Brünger, Adams et al., 1998) (Review: (Turkenberg and Dodson, 1996)).
Evaluation and phasing with calculated anomalous and dispersive differences at different wavelenghts were executed with the CCP4-package (FFT, MLPHARE, RSPS) (Collaborative Computational Project No. 4, 1994) (Helliwell, 1997) , SHARP (La Fortelle, Irwin et al., 1997) and Shake `n Bake (Xu et al, 2002)( Review: (Abrahams und De Graaff, 1998)).
The Fe positiones were entered into an ANALYSE run in SOLVE (Terwilliger et al., 1999) and non crystallographic symmetry operators were determined in RESOLVE (Terwilliger, 2000). The electron density map was averaged using AVE (Jones, 1992). The structural superpositions were made with LSQMAN (Kleywegt et al., 1994)
48 Results
3 Results
3.1 Acetylene hydratase of Pelobacter acetylenicus
3.1.1 Growth of Pelobacter acetylenicus under various conditions
In order to obtain large quantities of acetylene hydratase it was necessary to produce high amounts of P. acetylenicus cells. Dietmar Abt (Universität Konstanz, personal communication) described that the expression of the enzyme is increased 10-fold if a 50 l batch culture is fed continuously with 5 – 10 kPa of acetylene and the pH is adjusted between 6.8 and 7.0. The solubility of acetylene in water is 1185 mg ml-1 or 45.5 mM (20°C, 100 kPa; material safety data sheet acetylene, Messer, Griesheim). A 50 l batch culture of P. acetylenicus on 800 nM tungstate led to 48 g wet cell mass. The cells grew within 23 hours to an optical density of 0.77 at 578 nm (Figure 3.1) and were harvested at the end of the exponential growth phase.
0.8
0.7 Harvesting 0.6
0.5
578 0.4 OD 0.3
0.2
0.1
0.0 0 5 10 15 20 25 time [h]
Figure 3.1 Growth of P. acetylenicus. 50 l batch culture, 10 kPa C2H2 as energy and carbon source, freshwater medium with 800 nM tungstate.
Results 49
3.1.2 Purification of acetylene hydratase
Figure 3.2 shows a modified flow chart of the enzyme purification that had been carried out for acetylene hydratase from the tungstate cultivation (Abt, 2001) using here the Q-sepharase anion exchange column instead of the Mono-Q column. In order to avoid degradation of iron-sulfur clusters by dioxygen, all purification steps were done in an anaerobe chamber.
P. acetylenicus cells Lysozyme Pellet 10.000 x g Crude extract 2.3M (NH4)2SO4 10.000 x g Pellet Supernatant
3.2M (NH4)2SO4 10.000 x g Supernatant dialysis Anion exchanger Q-Sepharose
Gelfiltration Superdex
Acetylene Hydratase
Figure 3.2 Purification scheme of P. acetylenicus acetylene hydratase
Ammonium sulfate precipitation: After lysing the cells with lysozyme at room temperature the crude extract was subjected to a 2.3 M ammonium sulfate precipitation. After centrifugation, the whole acetylene hydratase activity was found in the supernatant. About 80% of the contaminating enzymes were removed in this step. The supernatant was subjected to a 3.2 M ammonium sulfate precipitation and the acetylene hydratase activity was found in the pellet. After the second (3.2 M) step about 64% of the initial activity was recovered.
50 Results
Anion exchange chromatography: After diluting of the resuspended pellet, the protein was loaded onto a Q-Sepharose anion exchange column, equilibrated with 50 mM Tris/HCl pH 7.5. With a salt gradient from 0 to 500 mM NaCl, flow rate of 3 ml min-1, acetylene hydratase eluted with 300 mM NaCl. During the elution the absorption of the eluted proteins was measured at 280 nm and 405 nm. About 61% of the initial activity was recovered (Table 3.1).
Gel filtration: The fractions of the Q-Sepharose purification step, showing acetylene hydratase bands on the SDS-PAGE, were pooled, concentrated to 1.5 ml, and loaded onto a Superdex 75 gel filtration column, equilibrated with 50 mM Tris/HCl 200 mM NaCl pH 7.5. Acetylene hydratase eluted, at a flow rate of 0.7 ml min-1, after 77 min. About 24% of the initial activity was recovered after the final gel filtration step. According to the specific activity acetylene hydratase was enriched 29.7 fold with a specific activity of 10.7 U mg-1 (Table 3.1). 20 g of wet cell mass led to 14 mg of pure acetylene hydratase.
Protein Activity Specific activity Yield Yield Enrichment -1 [mg] [U] [U mg ] (Protein) (activity) factor [%] [%] Crude extract 1712 616 0.36 100 100 1 3.2M AS 332 395 1.19 19 64 3.3 Q-Sepharose 86 374 4.35 5 61 12.1 Superdex 14 150 10.7 0.8 24 29.7 Table 3.1 Purification of acetylene hydratase from tungstate grown P. acetylenicus. 25 g of wet cells were used; activity was measured at 20°C. 1 U = 1 µmol acetylene min-1; AS = ammonium sulfate.
Figure 3.3 shows the SDS-PAGE of a typical acetylene hydratase purification. After Coomassie staining the gel was silver-stained and digitized. The enrichment of the acetylene hydratase band (73 kDa) during the purification procedure is clearly documented on the gel.
Results 51
kDa 1 2 3 4 5 6 kDa 94.4 –– 66.2 –– ––73 45.0 ––
31.0 ––
21.5 ––
14.4 ––
Figure 3.3 SDS-PAGE (12,5%) of acetylene hydratase purification from tungstate grown P. acetylenicus. Lane 1: Molecular weight markers; Lane 2: Crude extract (10 µg); Lane 3: Protein after 3.2 M (NH4)2SO4 precipitation (7 µg); Lane 4: Protein after anion exchanger Q-Sepharose (4 µg); Lane 5: Pure acetylene hydratase after gel filtration (2.5 µg); Lane 6: Molecular weight markers.
3.1.3 Crystallization and three-dimensional structure of acetylene hydratase
Crystallization of acetylene hydratase was performed under N2/H2 atmosphere in a Coy anaerobic tent. Crystals grew within one month from 7 mg ml-1 protein, reduced with 2.5 equivalents Na-dithionite, from 0.1 M Na-citrate pH 6.1, 0.1 M (NH4)2SO4, 30% PEG 8000, and -1 0.02% NaN3 and from 10 mg ml protein reduced with 2.5 equivalents Na-dithionite or 1.5 mM
Ti(III)-citrate from 21% PEG 8000, 0.2M Mg(Ac)2 0,1M MES pH 6.5. The second condition led to better well diffracting crystals. These crystals had a size at about 40*40*50 µm3 and they were flash cooled at 100K using 25% MPD as cryprotectant. Well diffracting Pelobacter acetylenicus acetylene hydratase crystals were obtained and belong to space group P21 with cell dimensions of a= 70.1Å, b= 105.3Å and c= 95.7Å and β=101° or to space group C2 with cell dimensions of a= 120.7Å, b= 70.5Å and c= 106.5Å and β=124° with each two monomers in the asymmetric unit.
52 Results
3.1.4 SAD data collection at ESRF
As a prerequisite for SAD data collection an X-ray fluorescence scan was carried out to determine the exact position of the L-shell tungsten absorption edge of Pelobacter acetylenicus acetylene hydratase. The observed edge was shifted to the theoretical expectations for free tungsten. According to this scan the wavelengths for this SAD data collection were chosen, 1.2136Å to maximize the anomalous contribution f`` of the tungsten atoms and a remote dataset at the intensity maximum of the synchrotron radiation at 0.9768Å. Figure 3.4 is showing the diffraction image of the acetylene hydratase crystals during the data collection.
Figure 3.4 Diffraction image of the acetylene hydratase crystal which was used for the SAD experiment. The detector edge corresponds to a limiting resolution of 2.0Å at an X-ray wavelength of 0.9768Å.
Results 53
The datasets were integrated with denzo and scalepack (Table 3.2).
Shell Lower Upper Average Average Norm. Linear Square limit Angstrom I error stat. Chi**2 R-fac R-fac 50.00 6.24 3854.1 145.5 78.1 3.500 0.065 0.074 6.24 4.95 3120.1 134.5 88.7 2.588 0.070 0.078 4.95 4.33 4481.0 177.6 106.6 3.098 0.063 0.068 4.33 3.93 3896.2 170.8 112.7 2.766 0.074 0.081 3.93 3.65 3497.5 167.3 121.6 4.225 0.111 0.119 3.65 3.44 2346.2 142.9 115.6 3.388 0.135 0.150 3.44 3.26 1814.7 128.3 110.2 2.238 0.129 0.136 3.26 3.12 1419.9 119.0 107.1 2.329 0.161 0.156 3.12 3.00 1120.5 111.8 103.6 1.991 0.185 0.171 3.00 2.90 928.2 106.4 100.4 1.959 0.222 0.216 2.90 2.81 784.8 103.0 98.3 1.756 0.243 0.230 2.81 2.73 586.6 98.7 96.0 1.767 0.308 0.304 2.73 2.66 526.1 98.1 95.9 1.680 0.339 0.338 2.66 2.59 450.3 97.4 95.6 1.440 0.365 0.345 2.59 2.53 427.2 95.2 93.6 1.388 0.376 0.376 2.53 2.48 384.2 94.4 93.2 1.318 0.397 0.412 2.48 2.43 330.8 95.3 94.4 1.275 0.456 0.522 2.43 2.38 303.3 94.5 93.7 1.001 0.444 0.435 2.38 2.34 275.5 93.5 92.9 0.925 0.459 0.493 2.34 2.30 149.0 96.2 95.7 0.930 0.916 0.596 All reflections 1554.2 118.9 99.7 2.021 0.142 0.114
Table 3.2 Scaled dataset of acetylene hydratase X-ray experiments
An anomalous difference patterson map was calculated because tungsten has a strong anomalous signal at 1.0Å. There is a tungsten signal at (0.01, 0, 0.55). It is difficult to compare this with replacement solutions because all was calculated in space group C2 where the y axis is not defined. The solution of the replacement has to be translated in a form that tungsten is laying at y=0. For replacement MolRep and Amore were used and as model NapA and Fdh (Raaijmakers et al., 2002) were taken. The c-alpha model of both was used. One solution was calculated but it was not able to refine this model very well (personal communication Dr. Oliver Einsle, Göttingen). The calculated model of acetylene hydratase is shown in Figure 3.5. 54 Results
Figure 3.5 Model of acetylene hydratase
3.2 Transhydroxylase of Pelobacter acidigallici
3.2.1 Growth of Pelobacter acidigallici
a, Molybdate cultivation of P. acidigallici
Cultivation of P. acidigallici in a 50 l batch culture led to 52 g of wet cell mass. The substrate gallic acid was 5 mM initially, and added twice during cultivation. The cells grew within 23 hours to an optical density at 578 nm of 0.89 (Figure 3.6).
Results 55
0,9
0,8 Harvesting 0,7
0,6
578 0,5 OD 0,4
0,3 5mM gallate 5mM gallate 5mM gallate 0,2
0,1 0 5 10 15 20 25 time [h]
Figure 3.6 Molybdate cultivation of P. acidigallici in a 50 l batch culture with gallic acid as energy and carbon source in saltwater medium.
b, Tungstate cultivation of P. acidigallici
This cultivation was performed to replace the molybdenum with tungstate for the X-ray experiments. 400nm tungstate was added to the solution and 6nm molybdate as backround. Tungstate cultivation of P. acidigallici in a 50 l batch culture led to 43 g of wet cell mass. The substrate gallic acid was 5 mM initially, and added twice during cultivation. The cells grew within 24.5 hours to an optical density at 578 nm of 0.77 (Figure 3.7). 56 Results
0,8
0,7 Harvesting 0,6
0,5 578 0,4 OD
0,3
0,2 5mM gallate 5mM gallate 5mM gallate 0,1
0 5 10 15 20 25 time [h]
Figure 3.7 Tungstate cultivation of P. acidigallici in a 50 l batch culture with gallic acid as energy and carbon source in saltwater medium.
3.2.2 Purification of transhydroxylase
Figure 3.8A shows a flow chart of the enzyme purification that was originally developed by Reichenbecher (1995) and modified by Sommer (1995). During chromatofocussing the protein is moving to its isoelectric point (pI), which is 4.1 in case of the transhydroxylase (Reichenbecher, 1995). Many iron-sulfur centers are known to be sensitive to acid pH and oxygen. The iron- sulfur centers of transhydroxylase appeared to degrade during the chromatofocussing step (Sommer, 1995). Thus, a modified protocol was developed (Figure 3.8 B) where the chromatofocussing step was omitted. This led to a highly active and electrophoretically pure enzyme.
Results 57
P. acidigallici cells P. acidigallici cells French Press Pellet French Press Pellet 100.000 x g 100.000 x g Crude extract Crude extract
Anion exchanger Anion exchanger DE 52 Q-Sepharose
Chromatofocussing Gelfiltration Superdex
Gelfiltration Superdex
Transhydroxylase AB Figure 3.8 Purification scheme of P. acidigallici transhydroxylase. (A) Purification scheme reported by Reichenbecher (1995) and Sommer (1995). (B) Modified purification scheme. a, Molybdate cultivation
Anion exchange chromatography: The crude extract was passed through a 100 ml Q-Sepharose anion exchanger column, equilibrated with 50 mM TEA pH 7.5. At these conditions, transhydroxylase did bind on this anion exchanger and was eluted by increasing the NaCl concentration of the buffer. About 42% of the initial activity was recovered (Table 3.3).
Gel filtration: The fractions of the Q-Sepharose purification step showing transhydroxylase bands on SDS- PAGE were loaded on a Superdex 200 gel filtration column. Transhydroxylase eluted, at a flow rate of 1 ml min-1, as a single peak after 150 min. About 65% of the initial activity was recovered after the final gel filtration step. According to SDS-PAGE (Figure 3.9), transhydroxylase was purified to homogeneity and the specific activity was enriched 9.2 fold (Table 3.3). The two subunits of transhydroxylase are at 86 kDa and 38 kDa, respectively.
58 Results
Protein Activity Specific activity Yield (Protein) Yield Enrichment -1 [mg] [U] [U mg ] [%] (activity) factor [%] Crude extract 726 290 0.4 (0.4) 100 100 1 Q-Sepharose 205 123 0.6 (0.7) 28 42 1.5 Superdex 50 190 3.8 (3.1) 6.8 65 9.5
Table 3.3 Purification of molybdate cultivated P. acidigallici transhydroxylase. 25 g of wet cells were used, 1 U = 1 µmol phloroglucinol min-1. Specific activity in brackets is from Reichenbecher et al. (1994).
kDa 1 2 3 4 5 6 kDa
97.4 –– –– 86 66.2 ––
45.0 –– –– 38
31.0 ––
21.5 –– 14.4 ––
Figure 3.9 SDS-PAGE (12,5%) of the purification of molybdenum P. acidigallici transhydroxylase. Lane 1 and 6: Molecular weight markers; Lane 2: Crude extract before ultrafiltration (7 µg); Lane 3: Crude extract after ultrafiltration (6 µg); Lane 4: Transhydroxylase after anion exchanger Q-Sepharose (3.2 µg); Lane 5: Transhydroxylase after gel filtration (2.6 µg).
b, Tungstate cultivation After each purification step no significant transhydroxylase activity could be measured. The purification was controlled with SDS-PAGE. The tungstate cultivation of P. acidigallici led to nearly inactive transhydroxylase. Only after the last purification step specific activity could be detected. Also the yield of transhydroxylase was very low (Table 3.4)
Results 59
Protein Activity Specific activity Yield (Protein) Yield Enrichment -1 [mg] [U] [U mg ] [%] (activity) factor [%] Crude extract 150 - - 100 - - Q-Sepharose 79 - - 53 - - Superdex 6 - <0.1 4 - -
Table 3.4 Purification of tungstate cultivated P. acidigallici transhydroxylase. 25 g of wet cells were used, 1 U = 1 µmol phloroglucinol min-1.
Tungstate cultivated P. acidigallici transhydroxylase could be purified to homogeneity as seen on the SDS-Page (Figure 3.10).
kDa 1 2 3 4 5 6 kDa
97.4 –– ––86 66.2 ––
45.0 –– ––38
31.0 ––
21.5 ––
14.4 ––
Figure 3.10 SDS-PAGE (12,5%) of the purification of tungstate P. acidigallici transhydroxylase. Lane 1 and 6: Molecular weight markers; Lane 2: Crude extract before ultrafiltration (10 µg); Lane 3: Crude extract after ultrafiltration (11 µg); Lane 4: Transhydroxylase after anion exchanger Q-Sepharose (5 µg); Lane 5: Transhydroxylase after gel filtration (4 µg).
3.2.3 Metal content of tungstate cultivated P. acidigallici Transhydroxylase
The active Transhydroxylase from molybdate-cultivated P. acidigallici contains about 12 mol iron and one mol molybdenum per mol enzyme, tungsten was absent (Abt, 2001). Table 3.5 presents the results of the ICP-MS datas of tungstate cultivated P. acidigallici transhydroxylase.
60 Results
Fe/mol Mo/mol W/mol Molybdenum Transhydroxylase 11.56 0.96 - Tungsten Transhydroxylase 1.8 0.13 0.0014 Table 3.5 Metal content of molybdenum transhydroxylase and tungsten transhydroxylase
3.2.4 Activity measurements of molybdenum Transhydroxylase
In one experiment of Reichenbecher and Schink (1999) as little as 0.7 µM (≈ 0.18 nmol) pyrogallol were incubated with 0.13 mg (≈ 1 nmol transhydroxylase) enriched enzyme in a final volume of 0.25 ml. Only under these conditiones formation of tetrahydroxybenzene was observed. A long time activity test with exclusion of dioxygen was carried out if also formation of tetrahydroxybenzene could be observed.. In the first sample 0.1 mg transhydroxylase was incubated with 1 mM pyrogallol at pH 7.0 to a final volume of 1 ml and in the second sample 0.1 mg transhydroxylase was incubated with 1 mM pyrogallol and 1 mM phloroglucinol to a final volume of 1 ml. Each sample was measured after 3h, 6h, 12h, 24h, 36h and 48 hours. All the tests did not show formation of tetrahydroxybenzene.
3.2.5 Crystallization of Transhydroxylase
Transhydroxylase was crystallized both in the “as isolated“ state and after addition of dithionite in the absence of dioxygen. Only the reduced enzyme led to suitable crystals (Figure 3.11). The -1 crystals grew in a N2/H2 atmosphere within three to for weeks from 14 mg ml protein, reduced with 12 equivalents Na-dithionite, from 0.05 M KPi pH 7.5, 20% PEG 8000. A 6% (v/v) additive of Crystal Screen I Factorial #7 (Hampton; 0.1 M sodium cacodylate pH 6.5, 1.4 M sodium acetate trihydrate) was beneficial for crystal growth. The crystals looked like triangular prisms and diffracted to resolutions better than 2.5 Å.
Results 61
Figure 3.11 Crystals of transhydroxylase
3.2.6 Unit cell parameters of transhydroxylase
The crystals belonged to space group P1, with unit cell dimensions of a = 173 Å, b = 179 Å, and c = 180 Å and angles of α = 63.8°, β = 64.1°, and γ = 65°. 12 monomers of transhydroxylase are located in the asymmetric unit. Another possible space group was R3 with only 4 monomers per asymmetric unit. Figure 3.12 is showing the less differences between the unit cells parameters that were calculated for the transhydroxylase crystals.
Figure 3.12 Unit cells of P1 and R3 (P1=green; R3=orange) of transhydroxylase.
62 Results
3.2.7 Transformation of transhydroxylase crystals
For the optimization of scattering characteristics of the crystals and to transform the unit cell into space group R3, several methods were tested. At first transformation of the crystals was done in a capillary (Figure 3.13) to see if the transhydroxylase crystals could be transformed.
Figure 3.13 Transformation of crystals in a capillary (Kiefersauer et al, 2000).
The crystals could be transformed in a nitrogen stream but they were not stable dating the X-ray diffraction experiments. Therefore the crystals had to be transformed in a loop to freeze the crystals after transformation. (Figure 3.14)
Figure 3.14 Transformation of crystals in a loop (Kiefersauer et al, 2000).
First attempts were accomplished thereby with vegetable oil, n-heptane, petroleum gasoline, high-viscosity paraffin oil, highly liquid paraffin oil and silicone oil and the cryoprotectant MPD (Methyl pentane diol) The best results were obtained with 25% MPD in the growth solution. With the help of Dr. Rainer Kiefersauer a two step process was developed to transform the crystals. In the first step the humidity was decreased from 94% to 89.5% within one 1 minute. After 5 minutes the humidity was decreased fromm 89.5% to 86% within one minute. This procedure helped to improve the resolution by 0.3 Å and the length of the axes decreased by about 2%. In addition the mosaicity was lowered to 0.4°. After transformation of the crystal datas were collected, but it was still not possible to index and scale the datas by using space Results 63
group R3. In addition the crystals could not be flash cooled after transformation. However the transhydroxylase crystals could be flash cooled after incubating them 2 minutes in the growth solution with 25% MPD in presence and absence of dioxygen.
3.2.8 Data collection of transhydroxylase crystals
First time data collection was performed on a Rigaku Rotaflex rotating anode X-ray generator with CuKα radiation at a wavelength of λCu = 1.5418 Å with Mar Research 2000 or Mar345 image plate detectors, ususally with rotation angles of 0.5° per image. A full dataset was collected with 360°. Molecular replacement with the data of DMSO-reductase did not help to find the phases and to solve the 3-dimensional protein structure.
3.2.9 Heavy metal soaks of transhydroxylase crystals
The crystals were soaked with heave metals under exclusion of dioxygen in order to perform Multiple Isomorphous Replacement (MIR) experiments. Hereby strongly diffracting heavy metals were attached to the protein crystals without changing the crystal geometry (isomorphy). Then the phases of the heavy metals alone were calculated and used to “phase” the protein. 2+ Soaking experiments were carried out with mercury compounds (HgCl2, Hg2SO4) and Ta6Br12 but the resolution did decrease more then 1 Å or the crystal underwent degradation. Also less spots were observed.
3.2.10 MAD and SAD data collection of transhydroxylase crystals
As a prerequisite for MAD data collection an X-ray fluorescence scan was carried out to determine the exact position of the K-shell iron absorption edge of transhydroxylase. The observed edge was shifted to the theoretical expectations for free iron (as derived from the equation of Kramers and Kronig).
According to this scan, the wavelengths for MAD data collection were chosen: λ1 = 1.7426 Å to maximize the anomalous f ′ contribution of the iron atoms, and λ2 = 1.7363 Å at the peak of the f ′ inflection (Figure 3.15) A third, remote, dataset was collected at the intensity maximum of the synchrotron radiation at λ3 = 1.0500 Å. In order to optimize completeness of Friedel pairs, two
120° oscillation sets were measured at each λ1 and λ2 with a difference of 180°. The rotation 64 Results
angle for every single image was 0.5° to avoid spot overlap. The rotation angle for every single image was 0.5° to avoid spot overlap.
Wavelength [A] 1.7426 1.7363
4
2 f'' 0
-2 Electrons -4
-6
-8 f'
7060 7080 7100 7120 7140 7160 Energy [eV]
Figure 3.15 Choice of wavelengths for the MAD experiment with transhydroxylase from Pelobacter acidigallici. The theoretical values for the iron absorption edge as obtained from the equation of Kramers and Kronig with the help of X-ray fluorescence scan of the transhydroxylase crystals wavelengths for data collection were chosen to maximize the anomalous f" contribution and the dispersive f’ inflection, respectively.
A self-rotation function of the remote data set of P. acidigallici transhydroxylase was calculated with the program GLRF (Tong & Rossman, 1990). A self-rotation function calculated in P1 (Figure 3.16) shows two clear twofold axes and a splitting of a further twofold axis into three parts.
Results 65
1 c* 0 0
. 0 0 .
2 0 0 1 0 0 . 0 . 0 0 0 2
3 0 0 0 .0 . 0 0 3
4 0 0 .0 .0 0 0 4
5 0 00 .0 . 0 50
6 0 .00 .00 0 6
70 .00 0.00 7
8 0.00 0.00 8
90.00 90.00 ψ ψ b// a
0 1 100.0 00.00
0 11 0.0 0.00 11
1 20 .00 .0 20 0 1
1 0 3 .0 0. 30 00 1
1 0 4 .0 0 0 .0 4 0 1
1 0 5 0 . 0 0 .0 5 0 1
1 0 6 0 . 0
0 . 1 0 6 0 7 0 1 0 . 0
0 . 0 7 0 1
k = 180.00
Figure 3.16 Polar plot of a self-rotation function for κ = 180 ° (twofold correlation). Two twofold axes can clearly be seen at Φ = 109°, ψ = 117° and Φ = 31°, ψ = 158°. (Abt et al., 2002)
3.2.11 Structure determination
X-ray data were collected at beam line BW6 at Deutsches Elektronensynchrotron in Hamburg (native data set and complex structure with pyrogallol) and at beam line ID29 at European Synchrotron Radiation Facility in Grenoble (high redundancy SAD data set and complex structure with 1,2,4-trihydroxy-benzene). They were processed with MOSFLM (Leslie,1999), scaled and further reduced using the CCP4 suite of programs (http:www.ccp4.ac.uk). The native structure was solved by the single anomalous diffraction technique using a high redundant data set (2 × 360° rotation angle) at the maximum of the f’’ of the Fe absorption edge (2.8 Å resolution). The positions of 33 of the 36 [4Fe-4S] clusters present in the P1 triclinic unit cell (12 hetero dimers per unit cell) could be located with Shake and Bake (Xu et al., 2002). Initial phases for both hands were calculated with SHARP (La Fortelle et al., 1997) and the residual 3 cluster positions and the correct hand for the phase calculation could be determined. The individual Fe positions within the clusters could not be resolved due to the actual resolution. The Fe positions were entered into an ANALYSE run in SOLVE (Terwilliger and Berendzen, 1999) and non crystallographic symmetry operators were determined in RESOLVE (Terwilliger, 2000) followed by solvent flattening of the electron density. The resulting electron density map showed clearly secondary structure elements. This electron density map was 12-fold averaged using AVE (Jones, 1992). Model building was done in this improved map using O 66 Results
(http://xray.bmc.uu.se/alwyn) and refinement was performed using CNS (Brünger et al., 1998) with native data set collected at BW6 (2.35 Å resolution, Rcryst = 0.199, Rfree = 25.4). Both complex structures were solved with difference Fourier technique using the structural model of native transhydroxylase and were refined with CNS (pyrogallol-complex: 2.20 Å resolution,
Rcryst = 0.179, Rfree = 22.4; inhibitor-complex: 2.00 Å resolution, Rcryst = 0.172, Rfree = 20.2) (for all refinements see Supplementary Table 1). The structural superpositions were made with LSQMAN (Kleywegt et al., 1994).
3.2.12 Description of the structure
3.2.12.1 Overall structure of transhydroxylase
The crystal structure shows that transhydroxylase is a heterodimer of ~ 90 Å × 91 Å × 70 Å, with the α- and ß-subunits consisting of 4 domains and 2 domains, respectively, and the relevant iron- sulfur clusters and the moco (MGD-cofactor)(Figure 3.17).
Figure 3.17. Overall structure of transhydroxylase. α-subunit domains I – IV are coloured magenta, blue, red and cream, respectively. ß- subunit sub-domains 1 and 2 of domain I are coloured orange and pink and domain II green. The Mo and MGD cofactors are shown as ball-and-stick models and the three [4Fe- 4S] clusters as red (Fe) and yellow (S) spheres. The figure was made with BOBSCRIPT (Esnouf, 1997) and RASTER3D (Merrit, 1994).
Results 67
3.2.12.2 α-subunit of transhydroxylase
The α-subunit (875 residues) is in the middle of the range from 755 residues for the dissimilatory nitrate reductase (NIR) and 982 for the formate dehydrogenase N (FDH-N)and does not contain a fifth domain as in FDH-N and the tungsten-containing formate dehydrogenase (FDH-T). The four domains are similar to those of the other DMSO-reductase (DMSOR) family organised around the MGD cofactors (Figure 3.18).
68 Results
Results 69
Figure 3.18. Structure based amino acid sequence alignment for a selection of members of the DMSO reductase family.α-subunit of transhydroxylase. Common secondary structure elements are boxed. Identical residues are coloured red, the molybdenum -coordinating residue yellow or green. The figure was produced with ALSCRIPT (Barton, G.J. 1993).
Figure 3.19 is showing the topology diagram of the α-subunit of transhydroxylase. Here its shown in which way the α-helices, the β-strands and loops are arranged in this subunit. Ist also shown how the MGD-Cofactor is located in the α-subunit and which secondary structure motives are involved in coordination of this cofactor. 70 Results
β18 α24
β17
α3 α5
α α2 4 α6 α22
β β β α27 β α26 β α23 β α7 β β 7 6 21 20 19 8 9 16 α21
α α 20 25 Ser175 MGD2 Mo α28 β22 MGD1 α29 α8 α30 β N β1 23
α1 β15
β2 α31 α32
α33 α19
β24 β3 α18 α35 β14 α34
α9
β10 α36 α10 α39 α11
β29’ α17 α41 β11 β 28 β25 β13 α14
α12 β31
α13
α15
β12
α37 β29’’ α40 β27 β30 β26 α16
α 38 Figure 3.19. Topology diagram of the α-subunit
The fold of Transhydroxylase is completely different between the secondary structure elements ß3 and α1, ß6 and ß7, ß15 and α23, α23 and ß19, ß22 and ß23, α35 and ß25 as well as ß25 and ß26, and concerns about 250 amino acid residues. Many of them take part in the formation of the substrate and co-substrate binding sites, which are accessible from the solvent through a narrow channel (Figure. 3.20). Results 71
Figure 3.20 Molecular surface representation of transhydroxylase displaying the access channel for substrate and co-substrate.
It contains three cis-peptides at Phe A166, Pro A483 and Pro A670. The active site of TH is located in the α-subunit and includes the Mo-binding site. The coordination of the Mo-ion is similar to that in DMSO reductase from Rhodobacter sphaeroides (Li et al, 2000) with six Mo- ligands arranged in a distorted trigonal pyramid (Figure 3.21). There are four sulphur ligands from both MGD moieties (bond distances between 2.39 and 2.46 Å), OG from Ser A175 (1.85 Å) and an oxygen from an acetate molecule (1.78 Å), which originates from the crystallisation buffer. This is a clear evidence that molybdenum transhydroxylase is a member of the DMSO-reductase family. In the acetate-free native structure, this space is probably filled by a hydroxyl or water molecule. The Mo-ion should be in the Mo (IV) oxidation state because the reduction agent sodium dithionite was present in the crystallisation buffer. The side chain of Tyr A560 adopts two different conformations and locks the active site if it is in the right conformation in Figure 3.21.
72 Results
Figure 3.21 Stereo drawing of the active site in transhydroxylase plus 2FO-FC electron density contoured at 1σ for selected residues of native TH with bound acetate molecule. The figure was made with BOBSCRIPT (Esnouf, 1997) and RASTER3D (Merrit et al, 1994).
3.2.12.3 β-subunit of transhydroxylase
The ß-subunit consists of three domains and has one cis-peptide at Pro B57. Domains I and II are ferredoxin-like domains, which together superimpose well with the relevant ferredoxin domains of the ß-subunits of tungsten-containing formate dehydrogenase (FDH-T) (132 Cα atoms, rmsd
1.15 Å ) and formate dehydrogenase N (FDH-N) (144 Cα atoms, rmsd 1.13 Å ) (Figure 3.22)
Results 73
Figure 3.22. Structure based amino acid sequence alignment for a selection of members of the DMSO reductase family of transhydroxylase ß-subunit. Common secondary structure elements are boxed. Identical residues are coloured red. ß-subunit cysteines are displayed in yellow. The figure was produced with ALSCRIPT
Domain II holds one and domain II two Fe-S-clusters as in FDH-T, in contrast to FDH-N which has two clusters in both domains. Domain III starting at residue B190 is folded in a seven- stranded mainly antiparallel ß-barrel. A search with domain III for related 3D structures in the DALI-Server (Holm et al., 1993) revealed the same fold for transthyretin (prealbumin) (Hamilton et al., 1993) and a closely related one for tenascin (third fibronectin type III repeat) (Leahy et al., 1992) (Figure 3.23). The latter is a cell adhesion protein and TH may be associated with the cytoplasmic membrane via this domain. 74 Results
Domain I
C
1 1 FeS 1 1
α5 α1 β8 β1 β9 β2 β3 α6
2 2 N FeS
2 2 3 3 α4 β6 3 FeS α3 3 α 2 Domain II β7
β5
β4
Domain III
β11 C
β14’’ β10 β17 β16 β15 β12 β13
α N 8
Figure 3.23 Topology diagram of the ß-subunit.
3.2.12.4 Transhydroxylase with substrate pyrogallol bound
In the crystal structure of the transhydroxylase substrate complex the pyrogallol binds with its O1 oxygen to the Mo (Figure 3.24, 2.4 Å bond distance) and replaces the acetate or the hydroxyl or water group in the acetate-free enzyme. This reaction is catalysed by His A144 (NE2 in hydrogen bond distance to O1 of pyrogallol), which acts as general base. The other part of the Mo coordination remains unaltered with similar bond distances as in the native structure. Carbon C1 of the pyrogallol is in sp3 state represented by the position of O1 above the plane of the Results 75
pyrogallol benzene ring. O2 is hydrogen bonded to OE2 of Asp A174 and O3 to NH2 of Arg A153. The Mo and the side chain functions of Asp A174 and Arg A153 are the recognition sites for the substrate. The side chain of Tyr A560 is in the open conformation and allows substrate binding. The space of the alternate conformation has been occupied by water molecules 1 – 3. The side chains of Tyr A404 and Tyr A152 are situated on top of the pyrogallol molecule. Their phenol rings are stacked parallel to each other. The OH group of Tyr A404 and the SG of Cys A557 are in hydrogen bonding distances to C5 of pyrogallol and may play a role as general base in catalysis of the hydroxyl transfer from the cosubstrate to the substrate. The space below the benzene ring of pyrogallol is lined by hydrophobic residues like Trp A176, Trp A354 and Phe A468. They create the hydrophobic surface region for binding of the hydrophobic part of the substrate molecule in the active site.
Figure 3.24 Stereo drawing of the active site in transhydroxylase plus 2FO-FC electron density contoured at 1σ for selected residues; transhydroxylase in complex with substrate pyrogallol, labelled as PG.
3.2.12.5 Transhydroxylase with inhibitor 1,2,4-trihydroxy-benzene bound
The crystal structure of the transhydroxylase inhibitor complex shows the 1,2,4-trihydroxy- benzene molecule bound with its O5 atom coordinated to the Mo (Figure 3.25, 2.35 Å bond distance). This additional O5 was formed while the soaking experiment and is showing that the molybdenum atom is binding the substrates in a highly specific way. The other part of the Mo- coordination is identical to the native structure of the transhydroxylase pyrogallol complex with similar bond distances. The side chain of Tyr A560 is in the closed conformation and the OH is bound to O2 of the inhibitor. O4 is hydrogen bonded to Asp A174 and O1 points into the direction of the side chain of Cys A577. Arg A153 cannot contact the inhibitor molecule because the corresponding OH-function is missing in the inhibitor. 76 Results
Figure 3.25 Stereo drawing of the active site in transhydroxylase plus 2FO-FC electron density contoured at 1σ for selected residues; transhydroxylas in complex with inhibitor 1,2,4- trihydroxy-benzene, labelled as INH.
Discussion 77
4. Discussion
This study presents data on two molybdopterin containing enzymes: the tungsten/iron-sulfur containing acetylene hydratase from Pelobacter acetylenicus and the molybdenum/iron-sulfur containing transhydroxylase from Pelobacter acidigallici. Although the overall reaction of both enzymes does not involve a net redox chemistry, they contain a set of complex redox cofactors. Both organisms were cultivated under exclusion of dioxygen and the two enzymes were purified to homogeneity and characterized with biochemical methods and crystallography. Crystallization of both proteins led to crystals suitable for X-ray analysis. The three-dimensional structure of native transhydroxylase was solved at a high resolution limit of 2.35 Å. Even the structures of transhydroxylase in complex with the substrate pyrogallol and the inhibitor 1,2,4-trihydroxy benzene were solved at high resolutions.The solution of the three dimensional structure of acetylene hydratase has been started and is still in progress.
4.1 Molybdenum versus tungsten in enzymes
There are now a number of well structurally characterized tungsten enzymes (Table 1.3). These enzymes have been mostly isolated from thermophilic or hyperthermophilic bacteria, living in extreme environments, such as deep-sea hydrothermal vents (Johnson et al., 1996; Kletzin and Adams, 1996). The tungsten containing acetylene hydratase is from a mesophilic bacterium that was isolated from freshwater creek sediment near Konstanz (Schink, 1984). Tungsten enzymes catalyze reactions that are related to those catalyzed by molybdenum enzymes (Stiefel, 1997). Moreover, the tungsten active sites contain the same type of pterin–ene-dithiolate ligand (molybdopterin; moco, Figure 1.2 A) that the molybdenum enzymes employ (Johnson et al., 1993; Chan et al., 1995; Johnson et al., 1996; Kletzin and Adams, 1996). However, the question why both tungsten and molybdenum containing enzymes exist has not been fully answered. One reason why tungsten has been favored may be related to the chemical differences between molybdenum and tungsten that are selected by the special conditions under which the tungsten enzymes function. For example, tungsten is more difficult to reduce than molybdenum (Stiefel, 1997). The redox potentials of W-complexes are more negative than the potentials of the corresponding Mo-complexes, typically by 300 – 400 mV. (Johnson et al., 1996). Therefore, if an enzyme is required to work at low redox potentials, tungsten might be evolutionarily favored. The greater inertness of tungsten towards substitution reactions can be an advantage in 78 Discussion
hyperthermophilic environments (≈ 100°C) where hydrolytic reactions can compromise the integrity of the active site (Stiefel, 1997). Therefore, the fitness of tungsten over molybdenum in certain cases may involve tungsten’s ability to function better in the chemical or physical environments, in which the tungsten enzymes containing oganisms have been found. Another reason is that the chemical differences of molybdenum and tungsten do not manifest in the different reactivity but in the geochemical environment of the organisms habitats.
Molybdenum deposits are mostly found in nature in the form of MoS2, while the harder tungsten is found as CaWO4 and Fe/MnWO4 (Chapter 1.1.). The chemical difference of the two elements is sufficient to effect geochemical differentiation of these elements. In chemistry it is much more difficult to convert tungstate to tungsten sulfide compounds than it is to convert molybdate to molybdenum sulfide compounds (Greenwood and Earnshaw, 1990; Stiefel, 1997). Thus, in sulfur-rich hydrothermal environments, in which many of the organisms have been detected that contain tungsten enzymes, molybdenum is perhaps not available because it precipitated with the massive sulfide deposits of these habitats. The organisms therefore just can use tungsten. Further studies on the enzymes, the organisms, the biogeochemistry of the habitats, and the coevolution of these enzymes should help to sort out the teleology of the molybdenum versus tungsten (Stiefel, 1997).
4.2 Cultivation of bacteria and enzyme purification
Chemical properties as well as atomic and ionic radii of tungsten and molybdenum are very similar (Table 1.1). Over the last decade, some striking parallels between the nature and function of molybdenum and tungsten centers in enzymes have emerged (Johnson et al., 1996; Hagen and Arendson, 1998). In 1999, Buc et al. successfully substituted tungsten for molybdenum in E. coli trimethylamine-N-oxide reductase (TMAOR) and obtained an active enzyme. Stewart et al. (2000) published the crystal structure of a highly active, tungsten substituted, DMSO-reductase from Rhodobacter capsulatus. In this publication, it was shown that R. capsulatus only can grow on tungstate in the presence of a low concentration (6 nM) of molybdate. The reason for this is not clear. The authors presume that molybdate (i) could be required for the biosynthesis of some other molybdoenzymes, such as nitrate reductase or xanthine dehydrogenase, (ii) that it is maybe needed as an activator for the biosynthesis of the molybdopterin cofactor, or (iii) that molybdate prevents the uptake of tungstate by the cell to a toxic level (Imperial et al., 1998; Grunden et al., 1999; Stewart et al., 2000). The Rhodobacter cells grew well at a concentration of 3 µM tungstate (plus 6 nM Discussion 79
molybdate). Significant inhibition of cell growth was obtained at tungstate concentrations higher than 100 µM, 60 mM tungstate in the medium inhibited cell growth completely.
4.2.1 Pelobacter acetylenicus
P. acetylenicus grew well in medium containing tungstate. Compared to former cultivations (Abt 2001, Meckenstock et al., 1999 ) the tungsten and vitamine concentration were increased two- fold, and molybdate (6 nM) was added. Harvesting the cells after 23h at an optical density of 0.77 at 578 nm led to high wet cell mass of 48 g (Figure 3.1),compared to a harvested 37 g wet cell mass reported by Abt (2001). The additional molybdenum may have a positive influence on the expression of other metabolic enzymes. Isolation of acetylene hydratase led to pure enzyme with a specific activity of 10.7 U mg-1 at 20°C, even though the purification strategy was modified. At this new strategy a Q-Sepharose anion exchange column was used instead of a Mono-Q anion exchange column The specific activity was not significantly lower than the previously reported 14.9 U mg-1 at 30°C. Note that the specific activity of acetylene hydratase depended very much on the individual batch and at the temperature of the activity measurements as shown by Abt (2001) where he measured a high activity of 42.25 U mg-1 at 50°C. However, the yield of acetylene hydratase with 0.8% was significantly lower than the reported 1.5% by Abt (2001).
4.2.2 Pelobacter acidigallici
(a) Molybdate cultivation
P. acidigallici grew well in a medium containing molybdate. Cultivation conditions with 10 µM molybdate and 10 nM tungstate as well as a three-fold increase of trace element solution concentration lead to a remarkable increase in cell yield by more than 100% (Figure 3.6) in contrast to former cultivations (Abt, 2001). The enzyme obtained with the new purification procedure developed in this work was more active (specific activity: 3.8 U mg-1) versus a specific activity of 3.1 U mg-1 reported by Reichenbecher et al. (1994).
80 Discussion
(b) Tungstate cultivation
P. acidigallici cells grew in a medium containing tungstate. Cultivation conditions with 400 nM tungstate and 6 nM molybdate led to 43 g of wet cell mass from a 50l batch culture volume and the cells grew within 24.5 hours to an optical density at 578 nm of 0.77. This is comparable to the results of molybdate cultivated P. acidigallici shown in Figure 3.6. Transhydroxylase from Tungstate cultivated P. acidigallici could be purified to homogeneity. The yield of protein was rather poor with 6mg/25g wet cell mass. Even the amount of protein in the crude extract with 150 mg is more than ten times lower than at the molybdate cultivation of P. acidigallici. The specific activity of the enzyme after the Superdex 200 purification step was less than 0.1 U mg-1 (Table 3.4). Metal analysis of ICP-MS revealed only 1.8 mol Fe, 0,13 mol Mo and 0.0014 mol W per mol tungsten transhydroxylase versus 11.56 mol Fe and 0.96 mol Mo per mol molybdenum transhydroxylase (Table 3.5). Tungstate cultivated P. acidigallici cells maybe do not produce as much protein as observed at molybdate cultivated P. acidigallici cells, even though a high amount of cell wet mass could be harvested (Figure 3.7). It was not possible to get high amounts of active tungsten transhydroxylase in contrast to successfully tungsten substituted trimethylamine-N-oxide reductase from E. coli (Buc et al., 1999). Maybe the metabolism system of P. acidigallici depends specifically on molybdenum, so i.e. the ratio of W Mo in the organism is very crucial. It could be possible that the uptake of tungsten reached a toxic level in P. acidigallici cells.
4.3 Crystallization and structural analysis of acetylene hydratase
Initial crystallization conditions for acetylene hydratase were obtained using the Hampton Screens I and II. After refining these crystallization conditions, crystals suitable for X-ray diffraction experiments grew within three to four weeks under the strict exclusion of dioxygen. Both, Na-dithionite and Ti(III)-citrate could be used as reductants. With Ti(III)-citrate the crystals grew slightly faster but the diffraction properties of theses crystals were not as good as with dithionite as reducing agent. These crystals were rather small and sensitive to dioxygen. A complete SAD dataset at the tungsten K-edge was collected at the ESRF in Grenoble, but the quality of this dataset was not good enough to solve the three dimensional structure of acetylene hydratase satisfactorily. One reason was related to the size of the crystals seen in the rapid decay Discussion 81
of diffraction quality in the synchrotron beam. During exposure the lattice order was destroyed which resulted in a significant decrease of effective resolution as well as an increase in Rmerge. In addition, the mosaicity of this crystal was above 1° which also led to a decrease of the overall data quality. In a second set of measurements a native dataset at a wavelength of 1.05 Å with a high resolution limit of 2.4 Å was collected at the DESY in Hamburg. This dataset allowed to perform molecular replacement using several structures of molybdopterin enzymes. Hereby the tungsten containing formate dehydrogenase (FDH-T) from Desulvofibrio gigas (Raaijmakers et al., 2002) worked as the best model for the replacement (Figure 3.5). Unfortunatly the refinement of this model did not work too well, so that at this point it represents only a preliminary structural model of acetylene hydratase.
Active site of acetylene hydratase
Figure 4.1 shows the active site of acetylene hydratase in the preliminary model. The distance between the tungsten and the [4Fe-4S]-cluster is about 12.4 Å. This distance, taken from the FDH-T would allow an efficient electron transfer. Redox titrations of the iron-sulfur center and of the enzyme activity gave potentials of –410 mV and –340 mV, respectively (Figure 1.7). Thus, acetylene hydratase seems to be active when the iron sulfur center is still in the oxidized [4Fe-4S]2+ state. Below –410 mV the iron sulfur center should be in the reduced [4Fe-4S]+ state which did not change the activity of the enzyme.
Figure 4.1 Active site of acetylene hydratase model (distances are adopt from the tungsten containing formate dehydrogenase (FDH-T) from Desulvofibrio gigas (Raaijmakers et al., 2002)). 82 Discussion
Biometric studies with the W model complexes demonstrated the likely participation of the W(IV) value state in the catalysis of the hydration of acetylene whereas the corresponding W(VI) remained inactive (Figure 4.2; Yadav et al., 1997).
2- 2- NC S O S CN NC S O S CN Na S O W VI 2 2 4 W IV
O NC S S CN NC S S CN
oxidized reduced
VI Figure 4.2 Reduction of [Et4N]2[W O2(mnt)2] (Yadav et al., 1997). mnt = 1,2-dicyanoethylenedithiolate
Reaction mechanism of acetylene hydratase
The reaction mechanism of acetylene hydratase remains unclear. Furthermore, the site for acetylene binding is unknown. The preliminary structure is showing the active site at the surface of the protein. The tungsten can serve as catalytic center where the acetylene binds and were the reaction product acetaldehyde could be released very easily (Figure 4.3).
C2H2
Figure 4.3 Coordination of the Molybdopterin cofactor at the preliminary acetylene hydratase model.
Several possible reaction mechanisms for acetylene hydratase are postulated. In addition to the classical Hg(II)/H+-catalyzed Markownikoff hydration of alkynes (Figure 4.4 A), numerous transition metal catalysts have been developed in recent years for the synthesis of ketones via hydration of alkynes, including the anti-Markownikoff hydration of terminal alkynes catalyzed Discussion 83
by a Ru(II)/phosphane mixture (Tokunaga and Wakatsuki, 1998). Figure 4.4 B shows an addition/elimination process of acetylene hydration according to Tokunaga and Wakatsuki’s Ruthenium catalyzed hydration. Figure 4.4 C shows a vinyl-tungsten intermediate that could undergo a direct OH insertion to vinylalcohol and hence acetaldehyde.
(A) + OH2 OH 2+ + + H OH Hg H2O - H + H HCCHH CCH HCCH HCC H C C CH3CHO 2+ - Hg H H 2+ Hg Hg+ Hg+
(B) alkyne complex vinylidene complex hydroxycarbene acyl intermediate intermediate O IV CH3 W H IV IV + H2O IV red. Elimination WC C W C VI W + CH3CHO H W CH H CCH O 3 H H O HH
W-vinyladduct
(C) H H IV H H Keto Enol W + H+ + OH- IV Tautomerism W + V CH3CHO W H H CCH HO H OH-
Figure 4.4 Possible reaction mechanisms of acetylene hydratase. (A) Hg2+ catalyzed addition of water to alkynes (March, 1992). (B) A putative mechanism, deduced from the anti-markownikoff hydration of terminal alkynes, catalyzed by a Ru(II)/phosphane mixture (Tokunaga and Wakatsuki, 1998). (C) Vinyl intermediate-hydration of acetylene (B.T. Golding, personal communication).
4.4 Structural aspects of transhydroxylase
4.4.1 Crystallization experiments
The crystallization of molybdenum transhydroxylase from P. acidigallici in presence and absence of dioxygen was described by R. Krieger (1997) and D. Abt (2001). The diffraction properties of these small, thin crystals were not suitable to perform further X-ray experiments and only the crystals grown in absence of dioxygen were able to diffract X-ray radiation. Varying the crystallization conditions and using higher protein concentrations and Na-dithionite 84 Discussion
as reductant led to large and well diffracting trigonal crystals with high resolutions limits better than 2.4Å (Figure 3.11). As described in chapter 3.2.6, the reduced molybdenum transhydroxylase crystals, grown in a
N2/H2 atmosphere, belong to space group P1, a space group with no symmetry elements. A native dataset was collected but the three-dimensional structure could not be solved with the method of molecular replacement, even though the amino acid sequence of transhydroxylase (Baas et al, 1999) is about 30% identical to the structurally charcterized DMSO reductase of Rhodobacter capsulatus (Schneider, 1996). The fact that the space group is P1 and the asymmetric unit contains 12 transhydroxylase molecules with 3 [4Fe-4S] clusters per the small β-subunit places 144 iron atoms in the asymmetric unit. Locating such a large number of heavy atoms is not trivial so other ideas were developed to solve the three dimensional structure of transhydroxylase.
Heavy metal soaks
Soaking experiments were performed with Ta6Br122+ (Figure 4.5). Ta6Br122+ is a strongly diffracting heavy metal cluster that has been shown to be extremely useful for phasing large molecules/molecular assemblies.
2+ Figure 4.5 Structure of Ta6Br12 (pink: tantal; green: bromide).
This multimetal was used by Schneider (1996) to solve the three dimensional structure of the
DMSO-reductase from Rhodobacter capsulatus. In this case the coordinates of Ta6Br122+ could be calculated by difference patterson function. Ta6Br122+ formed a stable derivative with the DMSO-reductase. Originally, this metal cluster was used to calculate the phases of the proteasome of Thermoplasma acidophilum (Loewe et al., 1995) and of the GTPCyclohydrolase I from E. coli (Nar et al., 1995). Discussion 85
Soaking the transhydroxylase crystals with this multimetal cluster would lead to locate 12 metal 2+ sites using the Multiple Isomorphous Replacement (MIR) technique in case of one Ta6Br12 binding to one transhydroxylase molecule in contrast to find 144 iron atoms positions by using the multiple wavelength anomalous dispersion (MAD) technique where measurements at different wavelengths can be used to calculate the phases of natural scatteres. During soaking 2+ experiments with Ta6Br12 , the crystals get degraded on the surface and the high resolution limit 2+ decreased. The soaking with Ta6Br12 maybe could have forced large conformational changes in the crystal geometry, and because of the tight packing of 12 transhydroxylase molecules in the asymmetric unit the lattice order could has been destroyed.
Transformation of transhydroxylase crystals
The observed dimensions of the P1 cell were in fact very close to a cell with R3 symmetry. In this space group, the asymmetric unit would only host 4 transhydroxylase molecules and in this case “only” 48 iron atoms would have been present instead of 144 iron atoms in space group P1. We therefore attempted to decrease the humidity of the crystals to achieve a tighter packing, produncing a unit cell with space group R3. Despite changing the humidity of the transhydroxylase crystals in two steps and increasing the resolution from 2.4 Å to 2.1 Å, it was still not possible to scale the measured data in space group R3.
4.4.2 MAD experiments at DESY
If an anomalous scatterer is present, collecting datasets at different wavelengths can be used to calculate phases. Useful anomalous scatterers are almost all naturally occuring metals. Depending on the type of atom, the absorbed energy can be just enough to eject an electron completely from the atom, so that the energy cannot be re-emitted. Measurements on and after this edge are similar to measurements with and without the anomalous scatterer, like in a MIR derivative. In the case of transhydroxylase the iron atoms of the 3 [4Fe-4S] clusters were used as anomalous scatterers.
Data collection at DESY: a, a full dataset at a remote wavelength at 1.0500Å far from the absorption edge as a reference.
86 Discussion
b, a full dataset on the absorption edge, such as to maximize the dispersive contribution, which is like a heavy atom derivative in MIR and is called f ’ at a wavelength of 1.7426Å. c, a full dataset on the absorption edge, slightly shifted to maximize the anomalous contribution, called f “ at a wavelength of 1.7363Å.
For each wavelength, two segments of 120° (1°-120°; 180°-300°) were measured at beamline BW6 for each dataset at a resolution to 2.3 Å. The rotation range of only 120° was chosen to minimize crystal decay during the measurement, thereby accepting the disadvantage of lower data redundancy and therefore also data accuracy.
The datasets were integrated and scaled, but the iron positiones could not be located and so the phases of the protein were not found. Maybe measuring 240° of the crystals did not yield enough observations to locate the position of the anomalous scatterers. However, no degredation of the transhydroxylase crystals could be observed and the crystals were stable during the MAD experiments at 100K. The changes of the parameters of the unit cell were less then 0.5% in axes lengths or angles (Figure 4.6). Accordingly, longer measurements could be performed to get higher redundancy of observed data.
67 184
182 66 c 180 γ 65 b 178 β
176 Angles 64 Axes lengthAxes [nm]
174 a α 63 172
170 62 0 100 200 300 400 500 600 0 100 200 300 400 500 600 Images Images
Figure 4.6 Changes of unit cell parameters (axes length and angles) while the MAD experiment.
4.4.3 SAD experiment at the ESRF
SAD experiments were performed at the ESRF in Grenoble at the beamline ID29. A high redundancy SAD data set at a wavelength of 1.73648 Å was collected. The crystals were measured 720° and the resolution limit was chosen at 2.8 Å. Using this dataset, the positions of 33 of the 36 [4Fe-4S] clusters present in the P1 triclinic unit cell could be located with Shake and Discussion 87
Bake, and subsequently an atomic model of transhydroxylase could be built. Refinement of the model was performed using CNS against the high resolution native data set (2.3 Å) collected at BW6 in Hamburg (all refinements statistics are shown in Supplementary Table 1).
4.4.4 Overall structure of transhydroxylase
The overall structure of transhydroxylase is shown in Figure 3. 17. Based on the nucleotide sequence of its coding gene (Baas, D. and Rétey, J.; 1999) transhydroxylase belongs to the DMSO reductase family. Members of this family share the Mo-containing α-subunit but may also have one or two additional small subunits. The crystal structures of the following enzymes of the DMSO reductase family have been determined: DMSO reductase (DMSOR) from Rhodobacter sphaeroides (Schindelin et al.; 1996), and Rhodobacter capsulatus (McAlpine et al.; 1997; Schneider et al.; 1996), formate dehydrogenase H (FDH-H) from Escherichia coli (Boyington et al.; 1997), dissimilatory nitrate reductase (NapA) from Desulfovibrio desulfuricans (Dias et al.; 1999), all having one α-subunit only compared to the transhydroxylase. Tungsten-containing formate dehydrogenase (FDH-T) from Desulfovibrio gigas (Raaijmakers et al.; 2002), do has an α- and ß-subunit, nitrate reductase A (NarGH) (Bertero et al.; 2003) and formate dehydrogenase N (FDH-N) (Jormakka et al.; 2002) from Escherichia coli, contain an α-, ß-, and γ-subunit. The fold of the transhydroxylase α-subunit is very similar to the fold of the DMSO-reductase (Figure 4.7). The active site of transhydroxylase is located in the α-subunit and includes the Mo- binding site. The ß-subunit with its three domains differs from well known motives of the structures of the DMSO-reductase family. As described in chapter 3.4.12.3 domains I and II are ferredoxin-like domains, which together superimpose well with the related ferredoxin domains of the ß-subunits of FDH-T and FDH-N. Domain III starting at residue B190 is folded in a seven-stranded mainly antiparallel ß-barrel. This region is similar to transthyretin and tenascin and the latter is a cell adhesion protein and transhydroxylase may be associated with the cytoplasmatic membrane via this domain.
88 Discussion
Figure 4.7 Comparison of transhydroxylase to DMSO-reductase from R. capsulatus (Schneider et al., 1996). red: α-subunit of transhydroxylase, green: β subunit of transhydroxylase, blue: α- subunit of DMSO-reductase
The molybdenum coordination is similar to that in DMSO-reductase from Rhodobacter sphaeroides (Li et al.; 2000) and other enzymes of the DMSO reductase family with six Mo- ligands arranged in a distorted trigonal pyramid. There are four sulphur ligands from both MGD moieties an OG from a Ser A175 and an oxygen from an acetate molecule which originates from the crystallisation buffer (Figure 3.2.21).
Is there a tyrosine at the active site?
The three dimensional structures of enzymes of the DMSO-reductase subfamily (Schneider et al., 1996; Schindelin et al., 1997; McAlpine et al., 1997; Czjzek et al., 1998; McAlpine et al., 1998; Stewart et al., 2000; Hung-Kei et al., 2000) have generated interest in the role of a tyrosine during catalytic turnover of DMSO, TMAO, and biotin sulfoxide (BSO) (Johnson and Rajagopalan, 2001). R. sphaeroides and R. capsulatus DMSO-reductases are able to utilize a great variety of substrates including trimethylamine-N-oxide and dimethyl sulfoxide (Satoh and Kurihara, 1987). R. sphaeroides biotin sulfoxide reductase (BSOR) has been shown to use a variety of S- and N-oxides (Pollock and Barber, 1997). E. coli and S. massilia TMAO-reductases functions as the final enzyme in the anaerobic electron pathway and utilizes trimethylamine-N- oxide as terminal electron acceptor. Kinetic analyses of Iobbi-Nivol et al. (1996) have shown that E. coli TMAO-reductase does not efficiently reduce S-oxides. It has been postulated that the substrate specificity of the enzymes corresponds to a tyrosine near the active site (Czjzek et al., 1998; Buc et al., 1999). The DMSO-reductases and the biotin Discussion 89
sulfoxide reductase (BSOR) should have this residue whereas the TMAO-reductases lacks this tyrosine. Johnson and Rajagopalan (2001) showed, by site-directed mutagenesis, that insertion of a tyrosine in E. coli TMAO-reductase results in a decreased preference for trimethylamine-N- oxide relative to dimethyl sulfoxide. Mutation or deletion of the tyrosine in recombinant Rhodobacter DMSO-reductase results in a decreased specificity for S-oxides and an increased specificity for trimethylamine-N-oxide. Figure 4.8 shows an alignment of selected members of the DMSOR subfamily. The Rhodobacter DMSOR group and the DMSO-reductases of the γ-proteobacteria as well as the BSO-reductases contain a residue equivalent to this tyrosine. The TMAO-reductases lack this tyrosine or have a different amino acid at this position.
R. sphaeroides@Q57366 YGPTGTFGGSYG-WKSPGRLHNCQ- 168 R. sphaeroides@AAB94874 YGPTGTFGGSYG-WKNPGRLHNCQ- 143 Rhodobacter R. capsulatus@3318672 YGPQGVFGGSYG-WKSPGRLHNCT- 126 R. capsulatus@2981909 YGPSGVFGGSYG-WKSPGRLHNCT- 168 DMSO-reductases P. multocida@AAK03877 YGPSGLHAGQTG-WRATGQLHSST- 159 V. cholerae@G82168 YGPSGLHAGQTG-WRATGQLHSST- 159 TMAO S. putrefaciens@CAA06794 YGPTGTFGGSYG-WRSPGRLHNCQ- 159 S. massilia@O87948 REFLEKGVNA-D-RSTRGNGDFVR- 150 reductases E. coli@P33225 HGPSALLTAS-G-WQSTGMFHNAS- 170 V. cholerae@H82137 YGPASIFAGSYG-WRSNGVLHKAS- 136 E. coli@AAA23522 HGPSALLTAS-GGWQSTGMFHNAS- 109 E. coli@AAG58700 YGPASIFAGSYG-WRSNGVLHKAS- 89 C. jejuni@G81444 NGPSAIFAGSYG-WRSSGVLHKAQ- 162 BSO H. influenzae@P44798 HGSTGIFAGSYG-WFSCGSLHASR- 166 reductases R. sphaeroides@P54934 HGSTGIFAGSYG-WFSCGSLHASR- 111 H. pylori@G64570 KGASAIFGGSYG-WKSSGNMQNSR- 152 H. pylori@H71865 N----IFNASYGGWGHAGSLHRCN- 148 M. tuberculosis@E70916 -PDRLKYPMKRVGKRGEGKFERIS- 103 P. multocida@AAK03838 DPDRLKYPMKRVGKRGEGKFERIS- 200 H. influenzae@P45004 YGNEAVHVL-YGTGVDGGNITNSN- 166 Y. pestis@AAD37319 YGNEAVHVL-YGTGVDGGNITNSN- 181 E. coli@F64914 YGNEAVHVL-YGTGVDGGNITNSN- 176 DMSO-reductases E. coli@AAG56575 YGNESIYLN-YGTGTLGGTMTRSW- 176 of the E. coli@BAB34402 YGNEAVYIQ-YSSGIVGGNITRSSP 143 γ-proteobacteria E. coli@P77374 YGNESIYLN-YGTGTLGGTMTRSWP 174 E. coli@E64914 YGNEAVYIQ-YSSGIVGGNMTRSSP 173 E. coli@P18775 YGNESIYLN-YGTGTLGGTMTRSWP 181 E. coli@AAG57632 YGNESIYLN-YGTGTLGGTMTKSWP 113 P. acidigallici@CAB50913 YGPSAILSTPSSHHMWGNVGYRHS- 155 Transhydroxylase Figure 4.8 Partial sequence alignment of the members of the DMSO-reductase subfamily The highlighted Tyr (Tyr-156 in the R. sphaeroides alignment) is equivalent to the Tyr-114 of Johnson and Rajagopalan (2001) because the DMSOR-precursor protein was used in this alignment.
In transhydroxylase an interesting tyrosine at position 560 in the α-subunit could be observed. The side chain of Tyr A560 adopts two different conformations and locks the active site if it is in the right conformation. Without substrate or inhibitor, the channel to the active site could be locked by Tyr A560 so that access to the molybdenum is denied (Figure 4.9 A). The structure of 90 Discussion
transhydroxylase soaked with pyrogallol (2.2Å; Supplementary Table 1) is showing the tyrosine in a switched away position that access to the molybdenum is free (Figure 4.9 B).
Figure 4.9 Switching of the tyrosine A560 in the α-subunit green: tyrosine blocking the active site, red: switched tyrosine (A) native transhydroxylase, (B) transhydroxylase complex with pyrogallol
This tyrosine seems not to be involved in the catalytic mechanism but it could serve as a “protector” of the active site. It could block the access to the catalytic center for non specific compounds so that this active site is always free for true substrates. In case of a substrate entering the channel to the active site a recognition, process could be initialised, the tyrosine will switch away and reopen the access to the molybdenum. Another interesting tyrosine, Tyr A404 is located near the catalytic center and will be described in the next chapter.
4.4.5 Towards the reaction mechanism of transhydroxylase
Understanding the transhydroxylase reaction, converting pyrogallol to phloroglucinol, represents a chemical challenge. It was shown that 1,2,3,5-tetrahydroxybenzene was required in cell-free extracts at stoichiometric amounts to make the reaction run (Brune and Schink, 1990). When partially enriched transhydroxylase enzyme fractions were incubated with pyrogallol in the absence of tetrahydroxybenzene, neither conversion of pyrogallol to phloroglucinol nor formation of tetrahydroxybenzene was detected in the standard assay of Brune and Schink (1990). Reichenbecher and Schink (1999) showed a complete conversion of pyrogallol to phloroglucinol when the amount of enzyme was drastically increased and the pyrogallol concentration was decreased in the assay. Note that formation of tetrahydroxybenzene was not observed during the experiment. In one experiment of Reichenbecher and Schink (1999) as little Discussion 91
as 0.7 µM (≈ 0.18 nmol) pyrogallol were incubated with 0.13 mg (≈ 1 nmol transhydroxylase) enriched enzyme preparation in a final volume of 0.25 ml. This was the only experiment in which formation of tetrahydroxybenzene was observed. Reichenbecher and Schink (1999) 18 showed in an experiment using OH2 that the pyrogallol-phloroglucinol transhydroxylase of P. acidigallici transfers hydroxyl substituents only between organic carriers. There was no exchange with free water. With this, the transhydroxylase differs from all other moco containing enzymes that catalyze hydroxylations by oxygen transfer between water and an organic substrate (Schultz et al., 1995; Kisker et al., 1997)
4.4.5.1 Former proposed rection mechanisms
Over the years, various proposals for the catalytic mechanism of transhydroxylase have been made. In the first mechanism a Mo(VI)-oxo group reacts by electrophilic substitution with the position 5 of pyrogallol, forming a tetrahydroxybenzene-Mo(VI) ether (Figure 4.10). The oxygen-Mo bond is cleaved and one molecule tetrahydroxybenzene is formed. The tetrahydroxybenzene undergoes a 180° turn and an oxidation to a para-quinone whereas the Mo(VI) center is reduced to a Mo(IV) center. In the following steps the Mo(IV) binds to the keto-group at position 2 of the ortho-quinone and a Mo(VI)-oxo is cleaved whereas the product phloroglucinol is formed.
92 Discussion
OH H+ O HO OH HO OH
OH H+ O H O HO OH OH (VI) OH Mo Mo(VI) HO OH HO OH H+ OH Mo(VI) - O -2e
Mo(VI) H+ O +2e-
H+ O HO OH H O O
(IV) OH Mo HO OH H O HO OH H+ Mo(IV) O HO OH (IV) Mo
Figure 4.10 Putative mechanism for the transhydroxylase reaction via ‘Umpolung’ of the hydroxyl group by the molybdenum cofactor (Hille et al., 1999).
In the second mechanism the substrate pyrogallol is oxidized to an ortho-quinone that is attacked by a nucleophilic Mo(IV)-OH center (Figure 4.11). One molecule tetrahydroxybenzene is formed that undergoes a 180° turn at the active site and reacts with the molybdenum center in a reverse way as described in the first half of the cycle (Hille et al., 1999).
Discussion 93
OH H+ O HO OH HO OH
OH + H OH OH + HO OH OH (IV) O Mo HO OH Mo(IV) HO O H+ OH Mo(IV)
OH
Mo(IV) OH
O
(VI) Mo H+ O HO OH
OH OH
(IV) OH Mo HO OH H O HO OH H+ Mo(IV) +OH HO OH (IV) Mo
Figure 4.11 Putative mechanism for the transhydroxylase reaction via ‘Umpolung’ of the substrate by oxidation to an ortho-quinone and nucleophilic hydroxylation by a molybdenum-coordinated OH group (Hille et al., 1999).
Although tetrahydroxybenzene is a potential intermediate in the last two mechanisms, it is not obvious why it is needed to start the reaction. One explanation is that the turning of tetrahydroxybenzene is only possible by a dissociation/reassociation process. After release from the enzyme it may compete with pyrogallol or phloroglucinol for the active site (Hille et al., 1999). Consequently, a low intracellular pool of tetrahydroxybenzene originates that is hard to detect. This leads to inactive transhydroxylase, which has to be reactivated. About 10% of the cell protein is transhydroxylase (Reichenbecher and Schink, 1999). This appears reasonable if (i) the transhydroxylating process is the rate-limiting step in the degradation of gallic acid, or (ii) transhydroxylase is inactivated by tetrahydroxybenzene formed in the reaction. The extreme substrate inhibition of transhydroxylase (Reichenbecher and Schink, 1999) can be a consequence of the in-activation of transhydroxylase by tetrahydroxybenzene being formed during the reaction. The specific activity under in vitro conditions can only be maximized by addition of 94 Discussion
equal amounts of pyrogallol and tetrahydroxybenzene (Brune and Schink, 1990) and running the reaction in a cycle with reactivation of the inhibited transhydroxylase by dehydroxylation of tetrahydroxybenzene. Reichenbecher and Schink (1999) demonstrated the formation of tetrahydroxybenzene when small amounts of pyrogallol and transhydroxylase were incubated without addition of tetrahydroxybenzene. In this experiment the molar transhydroxylase : pyrogallol ratio was about five. Brune (1990) observed dimethyl-sulfide production during incubation of transhydroxylase with pyrogallol and DMSO. He postulated that the effect of DMSO originates from an oxidative production of tetrahydroxy-benzene from pyrogallol. Because transhydroxylase belongs to the group of the DMSO-reductases it is possible that transhydroxylase has a DMSO-reductase activity. The proposed mechanism of R. sphaeroides DMSOR (Garton et al., 1997) suggests a Mo(IV) site that binds DMSO and that is oxidized to a Mo(VI)-oxo site during catalytic turnover. According to this mechanism transhydroxylase in the inactivated Mo(IV) state converts DMSO to dimethylsulfide and reactivates itself to the Mo(VI)-oxo species that is able to hydroxylate pyrogallol. Therefore, tetrahydroxybenzene is not formed by a direct reaction of DMSO and pyrogallol but can be formed after reactivation of transhydroxylase by DMSO. If this interpretation is correct, then there is no need for an anaplerotic reaction to synthesize tetrahydroxybenzene de novo during growth as postulated by Hille et al. (1999). This hypothesis can be checked if transhydroxylase is incubated with pyrogallol and 18O-marked DMSO. The 18O should be found in the product phloroglucinol and in released tetrahydroxy-benzene. A control experiment incubating pyrogallol and DMSO under the same conditions will show that there is no oxidation of pyrogallol by DMSO (Abt, 2001).
4.4.5.2 New reaction mechanism
In the two reaction mechanisms described above, the molybdenum is involved in catalysis. Soaking experiments of transhydroxylase crystals with the substrate (pyrogallol) and inhibitor (1,2,4-trihydroxybenzene) showed that the molybdenum is binding its substrates in a specific way. Figure 3.24 shows the pyrogallol binding specifically with its O1 oxygen to the molybdenum replacing the acetate or the hydroxyl or water group in the acetate-free enzyme. The structure of the bound inhibitor is showing a formation of a new O-binding at position 5 (Figure 3.25). 1,2,4-trihydroxybenzene could be also transformed into 1,2,3,5-trihydroxybenzene (personal communication, Dr. Andreas Brune Universität Konstanz). This emphasizes the hypothesis that the molybdenum has the function of coordinating and positioning the substrate (pyrogallol) and is not involved in catalysis. The newly proposed reaction mechanism is similar Discussion 95
to the proposed mechanism suggested by Hille et al., 1999 where a cyclic oxidation/reduction mechanism with the formation of two hexahydroxydiphenylethers as intermediates is shown. The 3D structure of transhydroxylase reveals that a mechanism with co-catalyst is operating in transhydroxylase. Figure 4.12 shows a schematic view of the active site with bound pyrogallol as found in the relevant complex structure and manually docked co-catalyst. It adopts the position of the phenyl ring of Tyr A560 in the closed conformation and can form several hydrogen bonds with the protein and pyrogallol indicated by dashed lines in Figure 4.12. O2, the hydroxyl to be transferred to C5 of pyrogallol, is in close proximity to this atom. Tyr A404 is the most probable candidate to act as general base in this transfer reaction because its OH-group lies in appropriate distances to both atoms (3.4 Å to O2 and 3.3 Å to C5) but also Cys A557 could play this role (respective distances, 3.0 Å and 4.0 Å). This mechanism is in line with the experimental findings that 1,2,3,5-tetrahydroxybenzene is needed to start the reaction and thatthe transferred hydroxyl does not come from the solvent.
Figure 4.12 Schematic view of the active site with bound pyrogallol molecule and manually docked co-catalyst 1,2,3,5-tetrahydroxy-benzene both coloured blue.
Based on these data the proposed mechanism (Figure 4.13) involves Asp A174, His A144, Tyr A404 and the Mo as catalytic residues. Pyrogallol enters the active site first (a) and is bound (b) as in the transhydroxylase pyrogallol complex structure, being stabilized by the protonated Asp A174. His A144 abstracts the proton from O1, thus promoting its binding to the Mo and inducing the binding to the co-substrate (c). Tyr A404 is in the deprotonated state and abstracts a proton from O2 (d). Subsequently, it attacks the C5 of pyrogallol in a nucleophilic manner. A 96 Discussion
bridging bond from O2 to C5 of the pyrogallol is formed causing the flip of one double bond in the ring system (e). Now Tyr A404 abstracts the proton from C5, which leads to the formation of a diphenylether (f). A nucleophilic attack of the protonated tyrosine side chain on the bridging oxygen causes the bond break between this atom and the co-substrate, which has been transformed to the phloroglucinol product (g) which is released. Asp A174 and His A144 attack O2 and O1 of the quinoid structure in a nucleophilic manner, thus transforming it into the aromatic co-substrate, which is released (h). a) b)
D174 D174 NH NH O O O OH H144 H144 OH N O N H
HO OH HO O Mo(VI) +H+ Mo(VI) H
c) d) e)
D174 D174 D174 NH NH NH O OH O OH O OH H144 H144 H144 O N O N O N H H H
HO O HO O HO O Mo(VI) Mo(VI) Mo(VI) H H
-H+ H OH O O HO O O Y404 Y404 Y404 HO OH HO OH HO OH
OH OH OH
f) g) h)
D174 D174 D174 NH NH NH O OH O OH O O H144 H144 H144 O N O N OH N H H
HO O HO O HO OH Mo(VI) Mo(VI) Mo(VI)
-H+ O HO OH O OH O Y404 Y404 Y404 HO OH HO OH 1,2,3,5-tetrahydroxy-benzene
OH OH released
phloroglucinol product released Figure 4.13 Proposed reaction scheme for transhydroxylase based on the 3D structures and mechanistic studies. Discussion 97
4.4.5.3 Is the reaction molybdenum transhydroxylase efficient?
As shown in Figure 3.20 the active site is only accessible through a narrow. This channel is very narrow and the substrate has to pass the channel to get coordinated by the molybdenum. The pyrogallol must enter this channel in the right direction to bind with its O1 group to the molybdenum, because a torsion of this molecule inside the channel is not possible (Figure 4.14).
Figure 4.14 Stereo drawing of the molecular surface representation of transhydroxylase in complex with pyrogallol coordinated by the molybdenum in the narrow channel.
After this coordination the cosubstrate (1,2,3,5-tetrahydroxybenzene) has to enter the channel in a straight direction so that its OH2 group can be transferred to the C5 of the pyrogallol (Figure 4.15). After transferring the OH2 group to the C5 group of the pyrogallol, the cosubstrate is converted to phloroglucinol and has leave the channel like the new produced cosubstrate and another catalytic cycle could start.
98 Discussion
Figure 4.15 Stereo drawing of the molecular surface representation of transhydroxylase. The access channel with pyrogallol and the modelled cosubstrate are coordinated at the active site.
In this case it seems difficult to envision this as an efficient reaction mechanism. However, kinetic studies at transhydroxylase by Reichenbecher et al. (1999) describing a high turnover rate and also high specific activity of 3.8 U mg-1 was measured in this work. In addition, transhydroxylase is occurring in high amounts in P. acidigallici. Nearly 10% of the enzyme mass of this organism in consisting of transhydroxylase (Brune et al.; 1992). This fact is also shown in the high yield of transhydroxylase after purification. Because of large amounts of transhydroxylase in P. acidigallici there could be always a free accessible active center for performing this reaction. A similar case was reported at the adenylylsulfate reductase (APSR) from the hyperthermophilic Archaeoglobus fulgidus by Fritz et al. (2002). APS reductase converts adenosine phosphosulfate to sulfite and adenosine monophosphate. The active site of APS reductase is buried deeply in the protein interior and is accessible only from the outside through a 17-Å-long channel with a diameter of around 10 Å.
Discussion 99
4.4.5.4 The role of the β-subunit and its [4Fe-4S]-clusters
The role of the [4Fe-4S] clusters in the ß-subunit is unclear. Their closest Fe-Fe distances are 10.1 Å and 9.2 Å (Figure 4. 16), which would be well suited for an efficient electron transfer. But the closest Fe-Mo and Fe-MGD-group distances are 23.4 Å and 12.7 Å.
Figure 4.16 The active site of transhydroxylase with the distances between the cofactors.
The first Fe-Mo distance with 23.4 Å is too long for an effective electron transfer from the nearest [4Fe-4S] cluster to the Mo-ion, and the same applies to the distance to the MGD molecule. However, a closest distance of 12.4 Å from a [4Fe-4S] cluster to the methyl group C8M of FAD has been found in adenylysulfate reductase from Archaeoglobus fulgidus (Fritz et al.; 1999). Effective electron transfer can take place here due to a strictly conserved tryptophan located between the two cofactors and in van der Waals contact to both centres. No such aromatic residue between the [4Fe-4S] cluster and the MGD group is found in transhydroxylase which renders this pathway of an efficient electron transfer rather improbable. The catalysed reaction of transhydroxylase is a net non-redox reaction and does not need redox equivalents from outside. Therefore, it lacks the [4Fe-4S] cluster in the α-subunit, that would allow an effective electron transfer between the Mo redox center and the [4Fe-4S] clusters of the ß-subunit as in tungsten-containing formate dehydrogenase (FDH-T), formate dehydrogenase N (FDH-N) and nitrate reductase A (NarGH). It may be suggested that transhydroxylase evolved 100 Discussion
from such enzymes and contains the ß-subunit as a relict without catalytic function in the observed reaction. A similar effect is described by Jormakka et al. (2002) in formate dehydrogenase N. This enzyme is containing 5 [4Fe-4S] clusters (FeS-0 to FeS-4). Electron transport between the MGD and the FeS-1 without the need of FeS-0 is suggested. Also the existence of an FeS-0 cluster in the α-subunit of Escherichia coli nitrite reductase A has been questioned because one of the four cysteine residues for the potential cluster binding is replaced by a histidine residue and the cluster has not been detected by EPR (Rothery et al.; 1998).
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110 Appendix
6. Appendix
6.1 Abbreviations
Å Ångstrøm; 1 Å = 10-10 m ADH Alcohol dehydrogenase AH Acetylene hydratase of Pelobacter acetylenicus APSR Adenylylsulfate reductase BSA Bovine serum albumin BSO/BSOR Biotin sulfoxide/Biotin sulfoxide reductase Da Dalton; 1 Da = 1 g mol-1 DMSO/DMSOR Dimethyl sulfoxide/ Dimethyl sulfoxide reductase DNA Deoxyribonucleic acid EDTA Ethylenediaminetetraacetic acid Tetrasodium salt dihydrate EPR Electron paramagnetic resonance FDH-H Formate dehydrogenase H FDH-N Formate dehydrogenase N FDH-T Tungsten-containing formate dehydrogenase Fe Iron FPLC Fast protein liquid chromatography HEPES N-[2-Hydroxy-ethyl]piperazine-N’-[ethanesulfonic acid] HPLC High performance liquid chromatography ICP-MS Inductively coupled plasma mass spectrometry KPi Potassium phosphate MGD Molybdopterine-guanine-dinucleo NADH Nicotinamide adenine dinucleotide PAGE Polyacrylamide gel electrophoresis PEG Polyethylene glycol PVDF Polyvinylidene difluoride S Sulfur SDS Sodium dodecyl sulfate TEA Triethanolamine = 2,2',2''-Nitrilotriethanol = Tris(2- hydroxyethyl)amine
Appendix 111
TMAO/TMAOR Trimethylamine-N-oxide/Trimethylamine-N-oxide reductase Tris Tris-(hydroxymethyl)-aminomethane UV/Vis Ultraviolet/Visible light v/v Volume per volume w/v Weight per volume Y-ADH Yeast alcohol dehydrogenase
6.2 Amino acids
A Ala alanine M Met methionine B Asx Asn or Asp N Asn asparagine C Cys cysteine P Pro proline D Asp aspartate Q Gln glutamine E Glu glutamate R Arg arginine F Phe phenylalanine S Ser serine G Gly glycine T Thr threonine H His histidine V Val valine I Ile isoleucine W Trp tryptophan K Lys lysine Y Tyr tyrosine L Leu leucine Z Glx Gln or Glu
Adapted from Eur. J. Biochem. (1985), 150, 1 – 5
112 Appendix
6.3 Nucleic acid bases
A adenine S G/C strong G guanine W A/T weak C cytosine B G/C/T not A T thymine D A/G/T not C R A/G purine H A/C/T not G Y C/T pyrimidine V A/G/C not T K G/T keto group N A/G/C/T any M A/C amino group I inosine "wobble" to U/C/A/T
Adapted from Eur. J. Biochem. (1985), 150, 1 – 5
6.4 International System of Units (SI)
In this work the International System of Units SI, shown in the standard DIN 1301 is used. Conversions to non-legal, historical units were done according to the Physikalisch Technische Bundesanstalt, Braunschweig. An overview on SI base units, SI prefixes, SI units, and non-legal units including conversion rates is available at http://www.ptb.de/english/misc/sie.pdf (English version) or http://www.ptb.de/deutsch/onlnpub/sid/sid.pdf (German version). Appendix 113
6.5 Figures and Tables
Figure 1.1 The families of mononuclear molybdenum enzymes Figure 1.2 Molybdenum enzymes. Figure 1.3 Cofactors of molybdenum and tungsten enzymes. Figure 1.4 Structures of the most common types of iron-sulfur centers. Figure 1.5 Comparison of EPR properties of different types of iron-sulfur centers. Figure 1.6 Acetylene degradation pathway of P. acetylenicus (Rosner 1994, Schink, 1985). Figure 1.7 Redox properties of acetylene hydratase (Meckenstock et al., 1999). Figure 1.8 Transhydroxylase reaction in the pathway of gallic acid degradation in Pelobacter acidigallici (Brune and Schink, 1992). Figure 1.9 EPR spectra of dithionite-reduced transhydroxylase from P. acidigallici. Figure 2.1 The 50 l batch culture system. Figure 2.2 Reciprocal lattice planes and Bragg’s law. Figure 2.3 The Ewald construction. Figure 2.4 An Argand diagram of the Gaussian plane. A complex wave function can be described by a rotating vector with a length A (amplitude) and an angle a (phase) with the real axis. B) The difference between MIR and MAD. Figure 2.5 A Harker diagram for the deduction of protein phase angles by anomalous scattering (after Drenth, 1994). Figure 2.6 Schematic view of a sitting drop crystallization unit. Figure 2.7 The main functionalities of the humidity apparatus. Figure 3.1 Growth of P. acetylenicus Figure 3.2 Purification scheme of P. acetylenicus acetylene hydratase Figure 3.3 SDS-PAGE (12,5%) of acetylenehydratase purification from tungstate grown P. acetylenicus. Figure 3.4 Diffraction image of the acetylene hydratase crystal which was used for the SAD experiment. Figure 3.5 Model of acetylenehydratase Figure 3.6 Molybdate cultivation of P. acidigallici in a 50 l batch culture with gallic acid as energy and carbon source in saltwater medium. Figure 3.7 Tungstate cultivation of P. acidigallici in a 50 l batch culture with gallic acid as energy and carbon source in saltwater medium. Figure 3.8 Purification scheme of P. acidigallici transhydroxylase. Figure 3.9 SDS-PAGE (12,5%) of the purification of P. acidigallici transhydroxylase. Figure 3.10 SDS-PAGE (12,5%) of the purification of tungstate P. acidigallici transhydroxylase. Figure 3.11 Crystals of transhydroxylase Figure 3.12 Unit cells of P1 and R3 (P1=green; R3=orange) of transhydroxylase. Figure 3.13 Transformation of crystals in a capillary (Kiefersauer et al, 2000). Figure 3.14 Transformation of crystals in a loop (Kiefersauer et al, 2000). Figure 3.15 Choice of wavelengths for the MAD experiment. Figure 3.16 Polar plot of a self-rotation function for κ = 180 ° (twofoldcorrelation). Figure 3.17 Overall structure of TH. Figure 3.18 Structure based amino acid sequence alignment for a selection of members of the DMSO reductase family Figure 3.19 Topology diagram for α-subunit Figure 3.20 Molecular surface representation of TH displaying the access channel for substrate and co-substrate. Figure 3.21 Stereo drawing of the active site in TH plus 2FO-FC electron density contoured at 1 σ for selected residues of native TH with bound acetate molecule 114 Appendix
Figure 3.22 Structure based amino acid sequence alignment for a selection of members of the DMSO reductase family of transhydroxylase ß-subunit. Figure 3.23 Topology diagram for ß-subunit. Figure 3.24 Stereo drawing of the active site in TH plus 2FO-FC electron density contoured at 1 σ for selected residues; TH in complex with substrate pyrogallol, labelled as PG. Figure 3.25 Stereo drawing of the active site in TH plus 2FO-FC electron density contoured at 1 σ for selected residues Figure 4.1 Active site of acetylene hydratase model. VI Figure 4.2 Reduction of [Et4N]2[W O2(mnt)2]. Figure 4.3 Coordination of the Molybdopterin cofactor at the preliminary acetylene hydratase model. Figure 4.4 Possible reaction mechanisms of acetylene hydratase. 2+ Figure 4.5 Structure of Ta6Br12 . Figure 4.6 Changes of unit cell parameters (axes length and angles) while the MAD experiment. Figure 4.7 Comparison of transhydroxylase to DMSO-reductase from R. capsulatus. Figure 4.8 Partial sequence alignment of the members of the DMSO-reductase subfamily. Figure 4.9 Switching of the tyrosine A560 in the α-subunit. Figure 4.10 Putative mechanism for the transhydroxylase reaction via ‘Umpolung’ of the hydroxyl group by the molybdenum cofactor. Figure 4.11 Putative mechanism for the transhydroxylase reaction via ‘Umpolung’ of the substrate by oxidation to an ortho-quinone and nucleophilic hydroxylation by a molybdenum-coordinated OH group. Figure 4.12 Schematic view of the active site with bound pyrogallol molecule and manually docked co-catalyst 1,2,3,5-tetrahydroxy-benzene both coloured blue. Figure 4.13 Proposed reaction scheme for transhydroxylase based on the 3D structures and mechanistic studies. Figure 4.14 Stereo drawing of the molecular surface representation of transhydroxylase in complex with pyrogallol coordinated by molybdenum in the narrow channel. Figure 4.15 Stereo drawing of the molecular surface representation of transhydroxylase. Figure 4.16 The active site of transhydroxylase with the distances between the cofactors.
Table 1.1 Physical and chemical properties of molybdenum and tungsten. Table 1.2 Stoichiometric formulations for substrate reactions of selected molybdenum and tungsten enzymes. Table 1.3 Molybdenum and tungsten enzymes (Stiefel, 1997). Table 2.1 Freshwater medium (Schink, 1985). Table 2.2 Modified trace element solution SL10 (Widdel and Pfennig, 1981). Table 2.3 Vitamin solution (Widdel and Pfennig, 1981). Table 2.4 Special trace element solutions (Widdel, 1980; modified). Table 2.5 P. acetylenicus tungstate cultivation; additional ingredients. Table 2.6 Saltwater medium (Brune and Schink, 1990). Table 2.7 P. acidigallici molybdate cultivation; additional ingredients. Table 2.8 P. acidigallici tungstate cultivation; additional ingredients. Table 3.1 Purification of acetylene hydratase from tungstate grown P. acetylenicus. Table 3.2 Scaled dataset of acetylene hydratase X-ray experiments Table 3.3 Purification of molybdate cultivated P. acidigallici transhydroxylase. Table 3.4 Purification of tungstate cultivated P. acidigallici transhydroxylase. Table 3.5 Metal content of molybdenum transhydroxylase and tungsten transhydroxylase Appendix 115
6.6 Supplementary Table
Native Native/SAD PG-complex1 INH-complex2 Space group P1 P1 P1 P1 Unit cell a (Å) 174.02 172.96 172.47 172.57 b (Å) 179.64 178.58 178.14 178.44 c (Å) 181.22 179.91 179.05 179.67 α (º) 63.38 63.95 64.11 63.83 ß (º) 63.98 64.32 64.45 64.40 γ (º) 64.90 65.03 65.10 65.04 Wavelength (Å) 1.7426 1.73647 1.05 0.9393 Resolution range 38.63-2.35 25-2.8 (2.95- 25-2.2 (2.32- 25-2.0 (2.11- (Å)3 (2.39-2.35) 2.80) 2.20) 2.00) Measurements 1,733,679 2,895,987 2,639,960 3,432,221 Unique reflections 635,960 378,146 814,421 1,084767 Completeness3 90.1 (87.2) 92.4 (68.7) 96.9 (96.1) 96.6 (92.0) Multiplicity3 2.3 (2.2) 7.6 (6.9) 3.2 (3.2) 3.2 (2.7) I/σ (I)3 7.0 (1.2) 4.9 (2.1) 5.1 (1.4) 4.0 (1.9) 3,4 Rmerge (%) 13.4 (60.0) 12.7 (28.8) 14.0 (51.5) 12.0 (34.0) 3,5 Ranom (%) 5.7 (12.9) Phasing power6 1.12 / 0.578 Mean FOM7 0.22 / 0.56 8 Rcryst / Rfree (%) 19.9 / 25.4 17.9 / 22.4 17.2 / 20.2 Number of atoms Non-hydrogen 110,412 110,172 110,172 protein (altconf) (240) PGD 1,128 1,128 1,128 Mo ions 12 12 12 Fe ions 144 144 144 Ca ions 24 24 24 Inorganic sulfur 144 144 144 Ligand 489 108 120 Water molecules 10,157 10,089 10,091 Average B-factor 33.3 21.1 18.8 (all atoms, Å2) R.m.s deviation Bond lengths (Å) 0.08 0.008 0.008 Angles (º) 1.9 1.8 1.8 1 Pyrogallol-TH-complex 2 1,2,4-trihydroxy-TH-complex 3 Values in parentheses are for the highest resolution shell. 4 Rmerge (I) = ΣΣ|I(h)I −
116 Appendix
6 Phasing power = rms (F(+h) − F(-h)) / rms residual, where (F(+h) − F(-h)) is either the observed anomalous difference, used for the calculation of the first figure, or the calculated anomalous difference, used for the calculation of the second figure. The rms residual is an estimate of the remaining heavy atom structure factor based on the anomalous differences and errors in the measurement. 7 FOM − figure of merit, first figure obtained from SOLVE, second figure after RESOLVE. 8 R-factors were calculated using data F > 0 σ, R-factor = Σhkl||Fo| − |Fc|| / Σhkl|Fo|, where |Fo| and |Fc| are the observed and calculated structure factor amplitudes for reflection hkl, applied to the work (Rcryst) and test (Rfree) (10 % omitted from refinement) sets, respectively. 9Acetate Appendix 117
6.7 Acknowledgements
Danksagung
Mein besonderer Dank geht an Prof. Dr. Peter M.H. Kroneck für die Betreuung und Unter- stützung der vorliegenden Arbeit, für die Förderung der Zusammenarbeit mit anderen Labors sowie für die Ermöglichung des Besuches von (inter)nationalen Tagungen und Kursen.
Prof. Dr. Albrecht Messerschmidt danke ich für die Übernahme des Coreferates dieser Arbeit, die interessanten Diskussion, die große Hilfe bei der Strukturanalyse und die sehr interessanten Fahrten zum ESRF nach Grenoble.
Prof. Dr. Robert Huber danke ich für die Möglichkeit, in der Abteilung Strukturforschung am Max-Planck-Institut für Biochemie in Martinsried Kristallisationsexperimente durchführen zu können.
Ein Dank geht an Dr. Martin Augustin der mir bei meinen Aufenthalten in Martinsried mit Rat und Tat beiseite stand und viel dazu beigetragen hat meine Experiment erfolgreich durchzuführen.
Ein Dank geht an Dr. Oliver Einsle, der mir die ersten Schritte im Handling mit Kristallen gezeigt hat und für die Einführung in die Grundlagen der Strukturforschung.
Prof. Dr. Bernhard Schink danke ich für anregende Gespräche.
PD. Dr. Matthias Boll danke ich für nette gesellige Abende bei diversen Veranstaltungen und die anregenden Gespräche im Schlosskeller Rauischholzhausen.
Dr. Rainer Kiefersauer danke ich für die ausgiebige Hilfe bei der Transformation von Kristallen.
Diese Arbeit wurde durch das Schwerpunktprogramm „Radikale in der enzymatischen Katalyse“ der Deutschen Forschungsgemeinschaft (DFG) unterstützt. 118 Appendix
Außerdem möchte ich mich bedanken bei …
meinen Eltern, ohne deren Unterstützung ich nie soweit gekommen wäre.
allen, die mir bei der Erstellung der Doktorarbeit mit Rat und Tat geholfen haben, insbesondere bei den aktuellen und ehemaligen Mitgliedern der Arbeitsgruppe Kroneck: Dietmar Abt, Günther Fritz, Thorsten Kehl, Michael Koch, Marc Rudolf, Thorsten Ostendorp, Alexander Schiffer, Alma Steinbach, Klaus Sulger.
bei den ehemaligen und aktuellen Mitarbeiten der Arbeitsgruppe Huber: Gust, Schnitte, Lutti, Olli, Werner, Arne, Steffi, Susi, Berta, Tim, Klaus, Otto, Rainer, Rasso, Holger, Irena, Jo, Fritzi, Rupsi und Spuki und Stoni.für eine tolle Zeit in Martinsried, vielen interessanten Disskussionen, den Stadionbesuchen, Waldheimaufenthalten, Bürger`s sessions und verrückten Abenden im Happy Billard.
meinen Vertiefungskursstudenten Karin Daub, Michael Koch und Andreas Schnur für ihre Hilfe und Mitarbeit bei zahlreichen Versuchen.
meinen Hiwis Bastian Thaa, Emina Savarese und Martina Reis für ihre Mitarbeit bei zahlreichen Experimenten.
meinen Freunden aus den verschiedenen Fakultäten der Uni Konstanz für viele Feten, Fußballabende und Kneipenaufenthalten.
und zuletzt meiner lieben Frau Alexandra, die immer zu mir hielt, mich in allen Belangen unterstüzt und mir das Leben erleichtert hat.
Appendix 119
6.8 Curriculum vitae
Personal Data:
Name: Holger Nießen Born: 26.02.1974 (Lindau/B, Germany) Family status: married
Education:
Since October 2000 Ph. D. Thesis in the group of Prof. P.M.H. Kroneck, Universität Konstanz: Structural aspects of molybdenum-transhydroxylase from Pelobacter acidigallici and tungsten-acetylene hydratase from Pelobacter acetylenicus May 2002 FEBS/EU- Advanced practical training course “The role of metals in biology, medicine and the environment”, Louvain-la-Neuve, Belgium 29.08.2000 Diploma in Biology 1999 - 2000 Diploma Thesis in the group of Prof. P.M.H. Kroneck, Universität Konstanz: Isolation an characterisation of ATP-Sulfurylase from Desulfovibrio desulfuricans Essex 6 1995 - 2000 Student of Biology, Universität Konstanz, Germany 1994 - 1995 Student of Process Ingeniering Technische Universität Stuttgart, Germany May 1993 Matriculation, Bodenseegymnasium Lindau/B, Germany 1984 - 1993 Bodenseegymnasium Lindau/B, Germany 1982 - 1984 Primary School Schachen, Lindau/B, Germany 1980 - 1982 Primary School Bodolz, Lindau/B, Germany
120 Appendix
6.9 Publications
Niessen, H. Abt, D. J., Einsle O., Schink B, Kroneck P. M. H. and Messerschmidt A. (2004). Crystallization and preliminary X-ray analysis of the tungsten-dependant acetylene hydratase from Pelobacter acetylenicus. in preperation
Messerschmidt A., Niessen, H. Abt, D. J., Einsle O., Schink B and Kroneck P. M. H. (2004). Structure-derived mechanism of transhydroxylase, a molybdenum enzyme with no apparent valence change at the metal centre during catalysis. PNAS, submitted.
Abt D.J., Einsle O., Nießen H., Krieger R., Messerschmidt A., Schink B. and Kroneck P.M.H. (2001). Crystallization and preliminary X-ray analysis of the molybdenum dependent pyrogallol-phloroglucinol transhydroxylase of Pelobacter acidigallici. Acta Cryst. D 58(2), 343- 5.
Appendix 121
6.10 Conference abstracts
Nießen H., Abt D.J., Schink B., Messerschmidt A., Huber R. and Kroneck P.M.H. (2003). The 3-dimensional structure of the pyrogallol-phloroglucinol transhydroxylase from Pelobacter acidigallici. VAAM 2003, Berlin
Nießen H., Abt D.J., Brinkmann H., Meyer A., Messerschmidt A., Huber R., Schink B and Kroneck P.M.H (2002). Novel bacterial molybdopterin enzymes: Mo-transhydroxylase and W- acetylene hydratase. FEBS/EU Advanced practical training course “The role of Metals in Biology, Medicine and the Environment.” Louvain-la-Neuve, Belgium
Nießen H., Abt D.J., Schink B., Messerschmidt A., Huber R. and Kroneck P.M.H. (2002). New structural aspects of the pyrogallol-phloroglucinol transhydroxylase of Pelobacter acidigallici. VAAM 2002, Göttingen
Nießen H., Abt D.J., Schink B., Messerschmidt A., Huber R. and Kroneck P.M.H. (2002). Novel bacterial molybdopterin enzymes: Mo-Transhydroxylase and W-Acetylenehydratase. DFG-Priority Programme: Radicals in enzymatic catalysis, Schloß Rauischholzhausen.
Abt D.J., Nießen H., Schink B. and Kroneck P.M.H. (2001). Novel bacterial molybdopterin enzymes: Mo-transhydroxylase and W-acetylene hydratase. 10th International Conference on Bioinorganic Chemistry (ICBIC 10), Florence, Italy
Nießen H. and Kroneck P.M.H. (2000). Isolation an characterisation of ATP-Sulfurylase from Desulfovibrio desulfuricans Essex 6 . VAAM 2000, München