The anaerobic linalool metabolism in the
betaproteobacteria Castellaniella defragrans 65Phen
and Thauera linaloolentis 47Lol
Dissertation
Zur Erlangung des Doktorgrades der Naturwissenschaften
- Dr. rer. nat. -
Dem Fachbereich Biologie/Chemie der
Universität Bremen
vorgelegt von
Robert Marmulla
Bremen, November 2015
Die vorliegende Arbeit wurde in der Zeit von April 2012 bis Dezember 2015 in der
Mikrobiologie Abteilung des Max-Planck-Instituts für Marine Mikrobiologie in Bremen im
Rahmen des Graduiertenprogramms „International Max Planck Research School of Marine
Microbiology“ angefertigt.
1. Gutachter: Prof. Dr. Jens Harder
2. Gutachter: PD Dr. Ulrich Ermler
3. Gutachter: Prof. Dr. Uwe Nehls
4. Gutachter: Prof. Dr. Matthias Ullrich
Tag des Promotionskolloquiums: 11.12.2015
Table of contents
Summary
1
Zusammenfassung
3
Chapter I
Introduction
5
Isoprenoids and monoterpenes
5
Castellaniella defragrans 65Phen and linalool dehydratase/isomerase
10
Thauera linaloolentis 47Lol
12
Chapter II
Microbial monoterpene transformations - a review
18
Aim of the work
54
Chapter III
A novel purification protocol for the linalool dehydratase/isomerase
58
Chapter IV
X-ray structure of linalool dehydratase/isomerase reveals enzymatic alkene synthesis
79
Chapter V
A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
99
Chapter VI
Linalool isomerase, a membrane-anchored enzyme in the anaerobic monoterpene
degradation in Thauera linaloolentis 47Lol
112
Chapter VII
The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
141
Chapter VIII
General discussion and outlook
Linalool dehydratase/isomerase
167
167
Biotechnological application for the linalool dehydratase/isomerase
and linalool isomerase
169
Linalool isomerase
171
Second metabolic pathway for linalool in C. defragrans 65Phen
182
Acknowledgements
Eidesstattliche Erklärung
List of abbreviations
AFW
Artificial fresh water medium
ATP
Adenosine triphosphate
Atu
Acyclic terpene utilization
Blastp
Basic local alignments sequence tool for protein sequences
CV
column volume
DMAPP
Dimethylallyl diphosphate
(g)DNA
(genomic) Deoxyribonucleic acid
DTT
Dithiothreitol
EDTA
Ethylenediaminetetraacetic acid
GeoA
Geraniol dehydrogenase
GeoB
Geranial dehydrogenase
GPP
Geranyl diphosphate
HIC
Hydrophobic interaction chromatography
HMN
2,2,4,4,6,8,8-Heptamethylnonane
IPP
Isopentenyl diphosphate
IPTG
Isopropyl-E-D-1-thioglactopyranoside
Ldi
Linalool dehydratase/isomerase
Lis
Linalool isomerase
Liu
Leucine / isovalerate utilization
LPP
Linalyl diphosphate
MALDI-ToF MS
Matrix assisted laser desorption/ionization time of flight mass
spectrometry
NCBI
National Center for Biotechnology Information
NPP
Neryl diphosphate
ORF
Open reading frame
PAGE
Polyacrylamide gel electrophoreses
PCR
Polymerase chain reaction
RAST
Rapid Annotation using Subsystem Technology
SDS
Sodium dodecylsulfate
SEC
Size-exclusion chromatography
Tris-HCl
Tris(hydroxymethyl)-aminomethane - hydrochloric acid
Triton X100
Polyethylene glycol p-(1,1,3,3-tetramethylbutyl)-phenyl ether
Tween20
Polyoxyethylene (20) sorbitan monolaurate
Summary
Summary
The linalool dehydratase/isomerase enables Castellaniella defragrans 65Phen to grow on the
acyclic monoterpenes E-myrcene and geraniol. This enzyme catalyzes the stereospecific
hydration of E-myrcene to (S)-linalool and the isomerization of (S)-linalool to geraniol and
has been generally characterized. However, the catalytic mechanism was still unknown. I
developed an improved purification protocol to yield high amounts of pure protein, suitable
for structure determination by X-ray crystallography. A protein structure was successfully
determined and revealed a homopentameric assembly for the native enzyme. An active center
was identified within an (D,D)6-barrel of a subunit, partially covered by a extending loop from
a neighboring subunit. Co-crystallization with (R,S)-linalool confirmed the location of the
active center but did not provide detailed information to deduce a catalytic mechanism.
However, an acid-base catalysis was suggested.
Even though, the linalool dehydratase/isomerase accepts only the (S)-isomer of (R,S)-linalool,
C. defragrans 65Phen metabolized both isomers. Indications for a complex linalool
metabolism in C. defragrans 65Phen were investigated by physiological and biochemical
experiments. The formation of the monocyclic monoterpenes D-terpinene and terpinolene
from (R,S)-linalool as substrate was observed in enzyme assays. Early studies, have only
shown the formation of geranic acid as oxidation product from geraniol in cultures, but not its
consumption. Geranic acid did not support growth of C. defragrans 65Phen under denitrifying
conditions. Physiological and biochemical experiments lead to the hypothesis of a central
monoterpene metabolism via a monocyclic intermediate, regardless of the initial structure of
the monoterpene (acyclic, mono- or bicyclic).
Thauera linaloolentis 47Lol grows on linalool as sole carbon and energy source under
denitrifying conditions. A 3,1-hydroxyl-'1-'2-mutase (linalool isomerase) has been proposed
as initial enzyme for the isomerization of the tertiary alcohol to the primary alcohol. This
enzyme activity was successfully enriched six-fold by subcellular fractionation and sucrosegradient centrifugation. A putative gene, showing considerable sequence similarity to the
linalool dehydratase/isomerase of C. defragrans 65Phen, was identified in a draft genome.
Bioinformatic analysis suggested a membrane association by four transmembrane helices and
a cytosolic C-terminal domain. Purification to homogeneity was hindered by this membrane
association and sensitivity towards detergents. However, the enriched linalool isomerase
activity was generally characterized with respect to pH and temperature optimum, oxygensensitivity and its kinetic properties.
-1-
Summary
The overall linalool metabolism in T. linaloolentis 47Lol was investigated by physiological,
proteomic and genomic approaches. Transposon mutagenesis revealed cellular adaptations as
defense against the toxic effects of the monoterpene alcohol linalool. Strong evidence was
found for the active expression of the acyclic terpene utilization (Atu) pathway. Homologous
genes, encoding the enzymes of this pathway, were identified in the draft genome. Proteins
identified in linalool-grown cells matched the predicted genes. Enzyme assay experiments
confirmed the presence of a NAD+-dependent geraniol dehydrogenase and geranial
dehydrogenase. To my knowledge, this is the first experimentally-supported description of an
active Atu pathway outside of the genus Pseudomonas.
-2-
Zusammenfassung
Zusammenfassung
Die Linalool dehydratase/isomerase ermöglicht es Castellaniella defragrans 65Phen auf den
azyklischen Monoterpenen E-Myrcene und Geraniol zu wachsen. Dieses Enzym katalysiert
die
stereospezifische
Hydratisierung
von
E-Myrcene
zu
(S)-Linalool,
sowie
die
Isomerisierung von (S)-Linalool zu Geraniol und wurde grundlegend charakterisiert. Der
katalytische Mechanismus war jedoch unbekannt. Ein verbessertes Reinigungsprotokoll
wurde entwickelt, um hohe Ausbeuten an reinem Protein zu erhalten, welches die
Voraussetzungen für die Strukturaufklärung mit Röntgenstrukturanalyse erfüllte. Die
Proteinstruktur konnte erfolgreich aufgeklärt werde und zeigte eine Homopentamerstruktur
des nativen Enzymes. Ein aktives Zentrum wurde in einer „(D,D)6-barrel“ der einzelnen
Untereinheiten identifiziert, welches partiell durch eine Schleife der benachbarten
Untereinheit gedeckelt wird. Co-Kristallisation mit (R,S)-linalool bestätigten die Lage des
aktiven Zentrums. Allerdings war die Qualität der Daten der Co-Kristallisation nicht
ausreichend, um einen detaillierten Mechanismus ableiten zu können. Ein Säure-BaseMechanismus wurde vorgeschlagen.
Obwohl die Linalool dehydratase/isomerase nur das (S)-Isomer von (R,S)-Linalool akzeptiert,
metabolisiert C. defragrans 65Phen beide Isomere. Hinweise auf einen komplexen LinaloolStoffwechsel in C. defragrans 65Phen wurden mittels physiologischer und biochemischer
Experimente untersucht. Die Bildung der monozyklischen Monoterpene D-Terpinen und
Terpinolen ausgehend von (R,S)-Linalool als Substrat wurde in Enzymassays beobachtet.
Frühe Studien zeigten nur die Bildung von Geraniumsäure als Oxidationsprodukt von
Geraniol in Kulturen, aber nicht dessen Verbrauch. C. defragrans 65Phen wuchs nicht auf
Geraniumsäure unter denitrifizierenden Bedingungen. Physiologische und biochemische
Experimente führten zur Hypothese, dass der zentrale Monoterpen-Stoffwechsel, unabhängig
der Ausgangsstruktur der Monoterpene (azyklisch, mono- oder bizyklisch), über ein
monozyklisches Zwischenprodukt verläuft.
In ersten Studien konnte gezeigt werden, dass Thauera linaloolentis 47Lol Linalool und
Geraniol als alleinige Kohlenstoff- und Energiequelle unter denitrifizierenden Bedingungen
nutzen kann. Eine 3,1-Hydroxyl-'1-'2-Mutase (Linalool isomerase) wurde als beteiligtes
Enzym für die initiale Isomerisierung des tertiären Alkohols zum primären Alkohol
-3-
Zusammenfassung
vorgeschlagen. Diese Enzymaktivität wurde in der vorliegenden Arbeit mittels subzellulärer
Fraktionierung und Sucrose-Dichtegradienten-Zentrifugation sechsfach angereichert. Ein
Kandidatengen mit wesentlicher Sequenzähnlichkeit zur Linalool dehydratase/isomerase aus
C. defragrans 65Phen wurde im Genom identifiziert. Bioinformatische Analysen ergaben eine
Membranassoziation des Enzyms mit vier Transmembranhelices und einer cytosolischen Cterminalen
Domäne.
Eine
Reinigung
bis
zur
Homogenität
war
aufgrund
der
Membranassoziation und der Sensitivität gegenüber Detergenzien nicht möglich. Die
angereicherte Linalool isomerase Aktivität wurde grundlegend in Bezug auf das pH- und
Temperaturoptimum, die Sauerstoffsensitivität und der kinetischen Parameter charakterisiert.
Der
allgemeine
Linalool-Stoffwechsel
in
T.
linaloolentis
47Lol
wurde
mittels
physiologischen, proteomischen und genomischen Ansätzen untersucht. Die Analyse von
Transposon-Insertation-Mutanten zeigte zellulare Adaptionen als Verteidigung auf die
toxischen Effekte des Monoterpenalkohols Linalool. Deutliche Beweise für eine aktive
Expression des „acyclic terpene utilization (Atu)“ Stoffwechselwegs wurden gefunden.
Homologe Gene, welche die Enzyme dieses Stoffwechselwegs kodieren, wurden im Genom
identifiziert. In Linalool-gewachsenen Zellen wurden Proteine identifiziert, welche mit den im
Genom gefundenen Genen übereinstimmten. Enzymassays bestätigten das Vorhandensein
einer NAD+-abhängigen Geraniol Dehydrogenase und einer Geranial Dehydrogenase. Meiner
Kenntnis nach ist dies die erste experimentell-bestätigte Beschreibung des AtuStoffwechselwegs außerhalb des Genus Pseudomonas.
-4-
Chapter I - Introduction
Chapter I
Introduction
Isoprenoids and monoterpenes
Isoprenoids, also known as terpenoids, are naturally occurring hydrocarbons. The name
“isoprenoid” is derived from the isoprene-rule formulated by Otto Wallach in 1914 (Wallach,
1914). This concept was further developed and resulted in the “biogenic isoprene rule”,
stating that all terpenoids consist of repetitive isoprene units (2-methyl-buta-1,3-diene, C5H8)
(Ruzicka, 1953). Based on this rule, terpenoids can be divided into hemi- (C5), mono- (C10),
sesqui- (C15), di- (C20), tri- (C30), tetra- (40) or poly (>40) -terpenes (Breitmaier, 2006) and
express a vast structural variety. To date, more than 55000 different structures are known
(Ajikumar et al., 2008). Mainly produced as secondary plant metabolites, they are the main
constituent of essential oils. However, terpenoids occur in all domains of life and their roles in
nature are quite diverse. They serve as buildings blocks in the biosynthesis of biological
molecules like plant hormones (gibberellins), pigments (carotenoids), electron carriers
(quinones) and membrane components (sterols) (Kesselmeier and Staudt, 1999, Davis and
Croteau, 2000). Besides their contribution to biological structures, volatile terpenoids are used
as messaging molecules in plant reproduction and development but also as attractants for
pollinators or as repellents for herbivores (Gershenzon and Dudareva, 2007, Dudareva et al.,
2013, Muhlemann et al., 2014).
Terpenoid synthesis was intensively investigated and two pathways have been described: the
mevalonate pathway (MVA) and the 1-deoxy-D-xylose-5-phosphate (DXP) or 2-C-methylerythritol 4-phosphate (MEP) pathway. Both pathways are actively expressed in plants but
located in different cell compartments. The MVA pathway is expressed in the cytosol, while
the DXP/MEP pathway is found in the plastids. Eukaryotes and archaea use the MVA
pathway only. The DXP pathway is mostly used by bacteria. The common products of both
pathways are the universal precursor isopentenyl diphosphate (IPP) and dimethylallyl
diphosphate (DMAPP). In the MVA pathway, mevalonate is synthesized via 3-hydroxy-3methyl-glutaryl-CoA (HMG) from acetyl-CoA building blocks. A key enzyme is the HMGreductase. Mevalonate is transformed into IPP by the expenditure of three ATP and a
-5-
Chapter I - Introduction
decarboxylation. The interconversion between IPP and DMAPP is catalyzed by the
isopentenyl diphosphate isomerase. The DXP pathway is initiated by fusion of pyruvate with
glyceraldehyde-3-phosphate and a decarboxylation to yield 1-deoxy-D-xylose-5-phosphate
(DXP). DXP is subsequently transformed in an enzymatic cascade into IPP and DMAPP (Fig.
1). Both C5 precursor molecules are linked in a head-to-tail condensation by prenyl
transferases to form geranyl diphosphate (GPP) and farnesyl diphosphate (FPP) for
monoterpene and sesquiterpene synthesis, respectively. Longer terpenoids are synthesized by
linking GPP and FPP molecules (Dewick, 2002, Kirby and Keasling, 2009, Dudareva et al.,
2013).
-6-
Chapter I - Introduction
Figure 1: Biosynthetic pathway for isopentenyl diphosphate and dimethylallyl diphosphate, adapted
from Kirby et al. (Kirby and Keasling, 2009). Shown are the main steps of the 1-deoxy-D-xylose-5phosphate (DXP) or 2-C-methyl-erythritol 4-phosphate (MEP) and mevalonate (MVA) pathway. In
the DXP pathway, the central metabolite 1-deoxy-D-xylose-5-phosphate is derived from
glyceraldehyde-3-phosphate and pyruvate and further converted to 2-C-methyl-erythritol-4-phosphate
by a specific synthase (DXS) and reductase (DXR), respectively. 2-C-methyl-erythritol-4-phosphate is
converted to isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP). 3-hydroxy-3methylglutaryl-CoA is derived from acetyl-CoA in the MVA pathway. A 3-hydroxy-3-methylglutarylCoA reductase catalysis the conversion to mevalonate which is further transformed to IPP. IPP and
DMAPP interconverted by a specific isomerase.
-7-
Chapter I - Introduction
Monoterpenes (C10H16) are synthesized as acyclic, mono- and bicyclic molecules starting
from geranyl- or neryl diphosphate (GPP and NPP). After subtraction of the diphosphate
moiety, monoterpenes (e.g. myrcene) are formed by removal of a proton, while monoterpene
alcohols (e.g. linalool) are formed by addition of a hydroxyl group. The linalyl cation,
resulting from the removal of the diphosphate moiety from GPP or NPP and an intramolecular
shift of the positive charge to the tertiary position, is the universal precursor for acyclic
monoterpenoids. A ring-closure in the linalyl or the neryl cation leads to the D-terpinyl cation,
which is the common precursor of all mono- and bicyclic monoterpenes (e.g. limonene,
phellandrene) (Croteau, 1986, Croteau, 1987, Davis and Croteau, 2000, Cheng et al., 2007,
Bohlmann and Gershenzon, 2009).
Due to their volatility, isoprene and monoterpenes have a significant impact on the
atmospheric carbon budget. Emission rates of non-methane volatile organic compounds
(VOC) - comprising VOCs of biogenic (BVOC) and anthropogenic sources - into the
atmosphere are estimated to be about 1150 Tg C yr-1 (Atkinson and Arey, 2003). The largest
fraction of emitted BVOCs is represented by isoprene and monoterpenes with emission rates
of 500 Tg C yr-1 and 127 Tg C yr-1, respectively (Günther et al., 1995). Overall BVOC
emission is predicted to increase in the future and to have a larger impact on global changes
(Peñuelas and Staudt, 2010). Lifetimes of isoprene and monoterpenes within the atmosphere
range from minutes to hours. They are prone to chemical reactions and photooxidation due to
their reactive carbon double bonds (Atkinson and Arey, 2003, Kesselmeier and Staudt, 1999).
Soils are loaded with monoterpenes through plant roots, by leaf fall and litter decomposition.
Roots of Pinus species were shown to contain substantial amounts of monoterpenes (415 μg
g-1 fresh wt) with emission rates between 9 and 119 μg g-1 dry wt h-1. Limonene and D-/Epinene were the main components of emitted monoterpenes (Lin et al., 2007). D-/E-pinene
concentrations (2 – 90 mg m-3) were highest in soils around spruce and pine trees (Smolander
et al., 2006). Studies on conifers showed monoterpene concentrations up to 1500 μg g-1 dry
wt, mainly represented by E-pinene (Ludley et al., 2009). Conifer litter covered soils emitted
about 35 μg monoterpenes m-2 h-1 (Hayward et al., 2001, Ludley et al., 2009). Keeping these
numbers in mind, monoterpenes can represent a substantial fraction of biologically available
carbon in soils with strong vegetation. Besides this contribution to the biological available
carbon pool in soil environments (Misra et al., 1996, Owen et al., 2007), monoterpenes have a
-8-
Chapter I - Introduction
strong influence on nitrogen cycling. When used as carbon source, monoterpenes can
stimulate ammonium uptake and its incorporation into biomass. It was also shown that they
have inhibitory effects on nitrification. Studies showed potential effects on all other levels of
the nitrogen cycling as well (White, 1994, Smolander et al., 2006). The adverse effect on
nitrification was further investigated by several groups. Inhibition of nitrification up to 90 %
was observed in the presence of monoterpene mixtures in forest soils. Denitrification, in
contrast, was not affected (Paavolainen et al., 1998). Further studies showed that selected
monoterpenes had an inhibitory effect on aerobic methane oxidation, while denitrification was
again not affected. Amaral et al. tested the general effect of monoterpenes on the metabolism
of heterotrophic bacteria. A reduced carbon dioxide formation in the presence of
monoterpenes was only observed in S. aureus and B. subtilis but not in E. coli or P.
aeruginosa (Amaral et al., 1998). The difference in sensibility might be attributed to the
different cellular features of Gram-positive and Gram-negative bacteria. Below toxic
concentration, essential oils and monoterpenes support microbial growth under aerobic and
anaerobic conditions (Harder and Probian, 1995, Misra et al., 1996, Misra and Pavlostathis,
1997, Harder et al., 2000, Ramirez et al., 2010). Monoterpene transformations were described
for bacterial enrichment cultures and pure cultures. Fungi were described to transform
monoterpenes mainly by use of mono- and di-oxygenases (Farooq et al., 2004). However,
only a limited number of bacteria with standing nomenclature and their enzymatic machinery
for monoterpene transformations are described. These are presented in the review article
following this introduction (Marmulla and Harder, 2014). More recently, Pseudomonas sp.
M1 was studied intensively for its myrcene metabolism. Early studies proposed the initial
formation of myrcene-8-ol and a subsequent oxidation of the alcohol to the corresponding
acid. 2-methyl-6-methylen-2,7-octadienoic acid is activated by formation of the CoA thioester
2-methyl-6-methylen-2,7-octadienoyl-CoA and further degraded in E-oxidation-like reactions
(Iurescia et al., 1999). Now, it was reported that Pseudomonas sp. M1 responded with drastic
changes on the transcriptomic level to myrcene-exposure and showed an induced expression
of myrcene-degradation specific genes within a 28 kb genomic island. The proposed
degradation via myrcene-8-ol was supported and expanded by identification of genes
potentially encoding the involved enzymes: an alcohol and an aldehyde dehydrogenase (myrB
and myrA), a CoA-ligase (myrC) and an enoyl-CoA hydratase (myrD). An epoxide hydrolase
-9-
Chapter I - Introduction
was found to be expressed in myrcene grown cells and indicates an epoxidation of myrcene as
the initial activation reaction (Soares-Castro and Santos, 2015).
Castellaniella defragrans 65Phen and linalool dehydratase/isomerase
The betaproteobacterium C. defragrans 65Phen (ex. Alcaligenes defragrans) was isolated
under denitrifying conditions with the monocyclic monoterpene D-phellandrene as sole
carbon and energy source. It was described as mesophile, rod-shaped and motile with a
strictly respiratory metabolism. Molecular oxygen, nitrate, nitrite, nitric oxide and nitrous
oxide served as terminal electron acceptor. Fatty acids, selected amino acids and
monoterpenes were used as carbon sources. Sugars did not support growth. Among
monoterpenes, C. defragrans 65Phen used acyclic, monocyclic and bicyclic monoterpenes
and monoterpene alcohols as growth substrates under denitrifying conditions. A sp2hybridized C1-atom in cyclic monoterpenes is essential for the complete mineralization to
carbon dioxide (Foss et al., 1998, Heyen and Harder, 1998, Kämpfer et al., 2006).
The mineralization of the pure hydrocarbon monoterpene E-myrcene is initiated by the
linalool dehydratase/isomerase (Ldi). This bifunctional enzyme catalyzes the reversible
hydration of E-myrcene to (S)-linalool and its isomerization to geraniol (Brodkorb et al.,
2010, Lüddeke and Harder, 2011). Linalool dehydratase/isomerase was purified to
homogeneity and generally characterized. It is expressed as a precursor protein, possessing a
signal peptide for periplasmatic localization. The signal peptide is absent in the native
enzyme, which was proposed to assemble into a homo-tetrameric complex with an apparent
molecular weight of 160 kDa. The enzyme showed a pH-optimum at slightly alkaline pH and
a high oxygen-sensitivity. It was completely inactivated in the presence of > 1 % O2 (v/v), but
activity was restored by applying anoxic conditions and treatment with a reducing agent (e.g.
dithiothreitol). Ldi was shown to refold into an active state after treatment with 6 M urea. A
ldi gene was identified on a 50 kb fosmid by Edman degradation and reverse translation. The
identity of the gene was confirmed by heterologous expression in E. coli BL21(DE3)
(Brodkorb et al., 2010). Geraniol, resulting from the hydration of E-myrcene and
isomerization is subsequently oxidized to geranic acid. Geranic acid formation was shown in
cell suspensions and cell-free protein extracts of 65Phen with E-myrcene or D-phellandrene as
- 10 -
Chapter I - Introduction
substrate. Geraniol and geranial were suggested as possible intermediates (Heyen and Harder,
2000). A geraniol and a geranial dehydrogenase were identified, purified and heterologously
expressed in E. coli BL21(DE3). Both enzymes were generally characterized. The geraniol
dehydrogenase was described as a homodimer and affiliated with the benzyl alcohol
dehydrogenases within the medium-chain dehydrogenase/reductases (MDR) and possessed a
zinc-binding domain. The putative aldehyde dehydrogenase gene was expressed in E.coli
BL21(DE3) and catalyzed the oxidation of geranial to geranic acid in vitro. This geranial
dehydrogenase affiliated with the aldehyde dehydrogenase superfamily and showed the
presence of a partial Rossmann fold. The genes, encoding enzymes involved in the initial Emyrcene degradation, are not organized in an operon-like structure (Lüddeke et al., 2012b).
Recently, the genome of C. defragrans 65Phen became available (NZ_HG916765). The
genome comprises 3452 protein coding sequences and has an overall GC-content of 68.9 %.
A novel pathway for anaerobic limonene mineralization was elucidated by transposon
mutagenesis and proteomics. Initially, limonene is oxidized to perillyl alcohol at the primary
methyl group. This reaction is catalyzed by a limonene dehydrogenase (ctmAB). Perillyl
alcohol is further oxidized via perillyl aldehyde to perillic acid by the geraniol and geranial
dehydrogenases, respectively. Perillic acid is subsequently activated by the addition of
coenzyme A and the formation of an energy-rich thioester bond. The sp2-hydridyzed C1 atom
enables an oxidative ring cleavage to yield 3-isopropenyl-pimelyl-CoA and the complete
mineralization in the central metabolism. The enzymes involved in the functionalization and
oxidation of limonene are encoded in the cyclic terpene metabolism cluster (ctmABCDEFG),
while enzymes involved in the oxidative ring cleavage are encoded in the monoterpene ring
cleavage cluster (mrcABCDEFGH) (Petasch et al., 2014).
- 11 -
Chapter I - Introduction
Thauera linaloolentis 47Lol
T. linaloolentis 47Lol, a betaproteobacterium within the family Rhodocyclaceae, was isolated
on (R,S)-linalool as sole carbon and energy source under denitrifying conditions. The
facultative anaerobic bacterium can use the monoterpene alcohols linalool and geraniol as
sole carbon and energy sources. Both are completely oxidized to carbon dioxide coupled to
complete denitrification. In addition, only short chain fatty acids and ethanol supported
growth of 47Lol but not sugars. In contrast to C. defragrans 65Phen the substrate range for
monoterpenes is rather narrow. Non-functionalized and cyclic monoterpenes did not support
growth (Foss and Harder, 1998). The metabolism of linalool was studied in electron acceptor
limited cultures. Geraniol and geranial were observed in linalool-grown cultures. Nerol, the
cis-isomer of geraniol, was not formed. However, geraniol and nerol were oxidized to their
corresponding aldehydes by 47Lol but only geranial was further metabolized. Geraniol fed
cultures showed formation of linalool and geranial. A 3,1-hydroxyl-'1-'2-mutase was
proposed as novel enzyme, catalyzing the isomerization of the tertiary monoterpene alcohol
linalool to the primary alcohol geraniol but the overall monoterpene metabolism was not
completely elucidated (Foss and Harder, 1997). Recently, two datasets from whole genome
sequencing approaches became available on NCBI (ASM31020v1, ASM62130v1) and may
give an insight into the genomic potential of this organism.
- 12 -
Chapter I - Introduction
References
AJIKUMAR, P. K., TYO, K., CARLSEN, S., MUCHA, O., PHON, T. H. & STEPHANOPOULOS,
G. 2008. Terpenoids: Opportunities for biosynthesis of natural product drugs using engineered
microorganisms. Molecular Pharmaceutics, 5, 167-190.
AMARAL, J. A., EKINS, A., RICHARDS, S. R. & KNOWLES, R. 1998. Effect of selected
monoterpenes on methane oxidation, denitrification, and aerobic metabolism by bacteria in
pure culture. Applied and Environmental Microbiology, 64, 520-525.
ATKINSON, R. & AREY, J. 2003. Atmospheric degradation of volatile organic compounds.
Chemical Reviews, 103, 4605-4638.
BOHLMANN, J. & GERSHENZON, J. 2009. Old substrates for new enzymes of terpenoid
biosynthesis. Proceedings of the National Academy of Sciences, 106, 10402-10403.
BREITMAIER, E. 2006. Terpenes; Flavors, Fragrances, Pharmaca, Pheromones, Wiley-VCH.
BRODKORB, D., GOTTSCHALL, M., MARMULLA, R., LÜDDEKE, F. & HARDER, J. 2010.
Linalool dehydratase-isomerase, a bifunctional enzyme in the anaerobic degradation of
monoterpenes. Journal of Biological Chemistry, 285, 30436-30442.
CHENG, A. X., LOU, Y. G., MAO, Y. B., LU, S., WANG, L. J. & CHEN, X. Y. 2007. Plant
terpenoids: Biosynthesis and ecological functions. Journal of Integrative Plant Biology, 49,
179-186.
CROTEAU, R. 1986. Biosynthesis of Cyclic Monoterpenes. Acs Symposium Series, 317, 134-156.
CROTEAU, R. 1987. Biosynthesis and catabolism of monoterpenes. Chemical Reviews, 87, 929-954.
DAVIS, E. M. & CROTEAU, R. 2000. Cyclization enzymes in the biosynthesis of monoterpenes,
sesquiterpenes, and diterpenes. In: LEEPER, F. J. & VEDERAS, J. C. (eds.) Biosynthesis.
Springer Berlin Heidelberg.
DEWICK, P. M. 2002. The biosynthesis of C5-C25 terpenoid compounds. Natural Product Reports,
19, 181-222.
DUDAREVA, N., KLEMPIEN, A., MUHLEMANN, J. K. & KAPLAN, I. 2013. Biosynthesis,
function and metabolic engineering of plant volatile organic compounds. New Phytologist,
198, 16-32.
FAROOQ, A., ATTA UR, R. & CHOUDHARY, M. I. 2004. Fungal transformation of monoterpenes.
Current Organic Chemistry, 8, 353-366.
FÖRSTER-FROMME, K. & JENDROSSEK, D. 2010. Catabolism of citronellol and related acyclic
terpenoids in pseudomonads. Applied Microbiology and Biotechnology, 87, 859-869.
- 13 -
Chapter I - Introduction
FOSS, S. & HARDER, J. 1997. Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. Fems Microbiology Letters, 149,
71-75.
FOSS, S. & HARDER, J. 1998. Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov.,
isolated on oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) and nitrate.
Systematic and Applied Microbiology, 21, 365-373.
FOSS, S., HEYEN, U. & HARDER, J. 1998. Alcaligenes defragrans sp. nov., description of four
strains isolated on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alphaphellandrene) and nitrate. Systematic and Applied Microbiology, 21, 237-244.
GERSHENZON, J. & DUDAREVA, N. 2007. The function of terpene natural products in the natural
world. Nature Chemical Biology, 3, 408-414.
GÜNTHER, A., HEWITT, C. N., ERICKSON, D., FALL, R., GERON, C., GRAEDEL, T.,
HARLEY, P., KLINGER, L., LERDAU, M., MCKAY, W. A., PIERCE, T., SCHOLES, B.,
STEINBRECHER, R., TALLAMRAJU, R., TAYLOR, J. & ZIMMERMAN, P. 1995. A
global model of natural volatile organic compound emission. Journal of Geophysical
Research-Atmospheres, 100, 8873-8892.
HANSEN, R. E. & WINTHER, J. R. 2009. An introduction to methods for analyzing thiols and
disulfides: Reactions, reagents, and practical considerations. Analytical Biochemistry, 394,
147-158.
HARDER, J., HEYEN, U., PROBIAN, C. & FOSS, S. 2000. Anaerobic utilization of essential oils by
denitrifying bacteria. Biodegradation, 11, 55-63.
HARDER, J. & PROBIAN, C. 1995. Microbial degradation of monoterpenes in the absence of
molecular oxygen. Applied and Environmental Microbiology, 61, 3804-3808.
HAYWARD, S., MUNCEY, R. J., JAMES, A. E., HALSALL, C. J. & HEWITT, C. N. 2001.
Monoterpene emissions from soil in a Sitka spruce forest. Atmospheric Environment, 35,
4081-4087.
HEYEN, U. & HARDER, J. 1998. Cometabolic isoterpinolene formation from isolimonene by
denitrifying Alcaligenes defragrans. Fems Microbiology Letters, 169, 67-71.
HEYEN, U. & HARDER, J. 2000. Geranic acid formation, an initial reaction of anaerobic
monoterpene metabolism in denitrifying Alcaligenes defragrans. Applied and Environmental
Microbiology, 66, 3004-3009.
HOLLIDAY, G. L., MITCHELL, J. B. O. & THORNTON, J. M. 2009. Understanding the functional
roles of amino acid residues in enzyme catalysis. Journal of Molecular Biology, 390, 560-577.
- 14 -
Chapter I - Introduction
IURESCIA, S., MARCONI, A. M., TOFANI, D., GAMBACORTA, A., PATERNO, A.,
DEVIRGILIIS, C., VAN DER WERF, M. J. & ZENNARO, E. 1999. Identification and
sequencing of beta-myrcene catabolism genes from Pseudomonas sp strain M1. Applied and
Environmental Microbiology, 65, 2871-2876.
KÄMPFER, P., DENGER, K., COOK, A. M., LEE, S. T., JÄCKEL, U., DENNER, E. B. M. &
BUSSE, H. J. 2006. Castellaniella gen. nov., to accommodate the phylogenetic lineage of
Alcaligenes defragrans, and proposal of Castellaniella defragrans gen. nov., comb. nov and
Castellaniella denitrificans sp nov. International Journal of Systematic and Evolutionary
Microbiology, 56, 815-819.
KESSELMEIER, J. & STAUDT, M. 1999. Biogenic volatile organic compounds (VOC): An
overview on emission, physiology and ecology. Journal of Atmospheric Chemistry, 33, 23-88.
KIRBY, J. & KEASLING, J. D. 2009. Biosynthesis of Plant Isoprenoids: Perspectives for Microbial
Engineering. Annual Review of Plant Biology. Palo Alto: Annual Reviews.
LIN, C., OWEN, S. M. & PEÑUELAS, J. 2007. Volatile organic compounds in the roots and
rhizosphere of Pinus spp. Soil Biology and Biochemistry, 39, 951-960.
LIU, B., FROSTEGARD, A. & SHAPLEIGH, J. P. 2013. Draft genome sequences of five strains in
the genus Thauera. Genome announcements, 1.
LÜDDEKE, F., DIKFIDAN, A. & HARDER, J. 2012a. Physiology of deletion mutants in the
anaerobic
beta-myrcene
degradation
pathway
in
Castellaniella
defragrans.
BMC
Microbiology, 12.
LÜDDEKE, F. & HARDER, J. 2011. Enantiospecific (S)-(+)-linalool formation from beta-myrcene
by linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences, 66, 409-412.
LÜDDEKE, F., WÜLFING, A., TIMKE, M., GERMER, F., WEBER, J., DIKFIDAN, A.,
RAHNFELD, T., LINDER, D., MEYERDIERKS, A. & HARDER, J. 2012b. Geraniol and
geranial dehydrogenases induced in anaerobic monoterpene degradation by Castellaniella
defragrans. Applied and Environmental Microbiology.
LUDLEY, K. E., JICKELLS, S. M., CHAMBERLAIN, P. M., WHITAKER, J. & ROBINSON, C. H.
2009. Distribution of monoterpenes between organic resources in upper soil horizons under
monocultures of Picea abies, Picea sitchensis and Pinus sylvestris. Soil Biology and
Biochemistry, 41, 1050-1059.
MARINO, S. M. & GLADYSHEV, V. N. 2010. Cysteine function governs its conservation and
degeneration and restricts Its utilization on protein surfaces. Journal of Molecular Biology,
404, 902-916.
- 15 -
Chapter I - Introduction
MARINO, S. M. & GLADYSHEV, V. N. 2012. Analysis and functional prediction of reactive
cysteine residues. Journal of Biological Chemistry, 287, 4419-4425.
MISRA, G. & PAVLOSTATHIS, S. G. 1997. Biodegradation kinetics of monoterpenes in liquid and
soil-slurry systems. Applied Microbiology and Biotechnology, 47, 572-577.
MISRA, G., PAVLOSTATHIS, S. G., PERDUE, E. M. & ARAUJO, R. 1996. Aerobic biodegradation
of selected monoterpenes. Applied Microbiology and Biotechnology, 45, 831-838.
MUHLEMANN, J. K., KLEMPIEN, A. & DUDAREVA, N. 2014. Floral volatiles: from biosynthesis
to function. Plant Cell and Environment, 37, 1936-1949.
OWEN, S. M., CLARK, S., POMPE, M. & SEMPLE, K. T. 2007. Biogenic volatile organic
compounds as potential carbon sources for microbial communities in soil from the rhizosphere
of Populus tremula. Fems Microbiology Letters, 268, 34-39.
PAAVOLAINEN, L., KITUNEN, V. & SMOLANDER, A. 1998. Inhibition of nitrification in forest
soil by monoterpenes. Plant and Soil, 205, 147-154.
PEÑUELAS, J. & STAUDT, M. 2010. BVOCs and global change. Trends in Plant Science, 15, 133144.
PETASCH, J., DISCH, E.-M., MARKERT, S., BECHER, D., SCHWEDER, T., HÜTTEL, B.,
REINHARDT, R. & HARDER, J. 2014. The oxygen-independent metabolism of cyclic
monoterpenes in Castellaniella defragrans 65Phen. BMC Microbiology, 14, 164.
RAMIREZ, K. S., LAUBER, C. L. & FIERER, N. 2010. Microbial consumption and production of
volatile organic compounds at the soil-litter interface. Biogeochemistry, 99, 97-107.
RUZICKA, L. 1953. The isoprene rule and the biogenesis of terpenic compounds. Experientia, 9, 357367.
SMOLANDER, A., KETOLA, R. A., KOTIAHO, T., KANERVA, S., SUOMINEN, K. & KITUNEN,
V. 2006. Volatile monoterpenes in soil atmosphere under birch and conifers: Effects on soil N
transformations. Soil Biology & Biochemistry, 38, 3436-3442.
SOARES-CASTRO, P. & SANTOS, P. M. 2015. Deciphering the genome repertoire of Pseudomonas
sp. M1 toward β-myrcene biotransformation. Genome Biology and Evolution, 7, 1-17.
WALLACH, O. 1914. Terpene und Campher: Zusammenfassung eigener Untersuchungen auf dem
Gebiet der alicyclischen Kohlenstoffverbdinungen, Leipzig, Veit.
WHITE, C. S. 1994. Monoterpenes: Their effects on ecosystem nutrient cycling. Journal of Chemical
Ecology, 20, 1381-1406.
- 16 -
Chapter I - Introduction
- 17 -
Chapter II - Microbial monoterpene transformations - a review
Chapter II
Microbial monoterpene transformations - a review
Robert Marmulla and Jens Harder*
Department of Microbiology, Max Planck Institute for Marine Microbiology, Bremen,
Germany
* Jens Harder, Max Planck Institute for Marine Microbiology, Celsiustr. 1, Bremen 28359,
Germany, jharder@mpi-bremen.de
Edited by:
Collin Murrell, University of East Anglia, UK
Reviewed by: Terry John McGenity, University of Essex, UK
Andrew Crombie, University of East Anglia, UK
Review article in Frontiers in Microbiology
published 15 July 2014
dio:10.3389/fmicb.2014.00346
- 18 -
Chapter II - Microbial monoterpene transformations - a review
Abstract
Isoprene and monoterpenes constitute a significant fraction of new plant biomass. Emission
rates into the atmosphere alone are estimated to be over 500 Tg per year. These natural
hydrocarbons are mineralized annually in similar quantities. In the atmosphere, abiotic
photochemical processes cause lifetimes of minutes to hours. Microorganisms encounter
isoprene, monoterpenes, and other volatiles of plant origin while living in and on plants, in the
soil and in aquatic habitats. Below toxic concentrations, the compounds can serve as carbon
and energy source for aerobic and anaerobic microorganisms. Besides these catabolic
reactions, transformations may occur as part of detoxification processes. Initial
transformations of monoterpenes involve the introduction of functional groups, oxidation
reactions, and molecular rearrangements catalyzed by various enzymes. Pseudomonas and
Rhodococcus strains and members of the genera Castellaniella and Thauera have become
model organisms for the elucidation of biochemical pathways. We review here the enzymes
and their genes together with microorganisms known for a monoterpene metabolism, with a
strong focus on microorganisms that are taxonomically validly described and currently
available from culture collections. Metagenomes of microbiomes with a monoterpene-rich
diet confirmed the ecological relevance of monoterpene metabolism and raised concerns on
the quality of our insights based on the limited biochemical knowledge.
Introduction
Annually about 120 Pg of carbon dioxide are assimilated by plants. A part is transformed into
chemically complex molecules and released into the environment by emission or excretion
(Ghirardo et al., 2011). Volatile organic compounds (VOCs) comprise a large number of
molecules, including various hydrocarbons, single carbon compounds (e.g. methane),
isoprene and terpenes (e.g. mono- and sesquiterpenes). The atmosphere is loaded with an
estimated VOC emission rate of about 1150 Tg C yr-1 (Stotzky and Schenck, 1976, Guenther
et al., 1995, Atkinson and Arey, 2003). These estimates included only non-methane VOCs of
biogenic origin (BVOCs); a second source are anthropogenic VOCs. Among the BVOCs,
isoprene and monoterpenes dominate with estimated emission rates of about 500 Tg C yr−1
and 127 Tg C yr−1, respectively (Guenther et al., 1995). Monoterpenes (C10H16) consist of two
- 19 -
Chapter II - Microbial monoterpene transformations - a review
linked isoprene (C5H8) units and include in the strict sense only hydrocarbons. Often the term
monoterpene is applied including monoterpenoids which are characterized by oxygencontaining functional groups. Structural isomers - acyclic, mono- and bicyclic
monoterpenes -, stereoisomers as well as a variety of substitutions result in a large diversity of
molecules. Today, more than 55000 different isoprenoids are known (Ajikumar et al., 2008).
Monoterpenes are not only emitted as cooling substances (Sharkey et al., 2008), but can also
be stored intracellularly serving mainly as deterrent or infochemical (Dudareva et al., 2013).
Wood plants mainly accumulate pinene and other pure hydrocarbon monoterpenes as
constituents of their resins, whereas citrus plants are the major source of limonene. Flowers,
however, produce and emit a variety of oxygenated monoterpenes (e.g. linalool) (Kesselmeier
and Staudt, 1999 and references therein, Sharkey and Yeh, 2001, Bicas et al., 2009).
In the atmosphere, monoterpenes are transformed in purely chemical reactions within hours.
Photolysis and reactions with molecular oxygen, ozone, hydroxyl radicals, NOx species, and
chlorine atoms result in carbonyls, alcohols, esters, halogenated hydrocarbons, and
peroxynitrates. These products condense and lead to the formation of secondary aerosols.
Rain or precipitation transports them to soils (Atkinson and Arey, 2003, Fu et al., 2009,
Ziemann and Atkinson, 2012). Monoterpenes reach the surface layers of soils by leaf fall and
excreted resins. Also roots emit monoterpenes into the rhizosphere (Wilt et al., 1993,
Kainulainen and Holopainen, 2002). Deeper soil layers do contain significant less
monoterpenes
than
the
surface
soil
layer.
Emission
into
the
atmosphere
and
biotransformations in the surface layer mainly by microorganisms are the major sinks. An
alternative, abiotic photoreactions like in the atmosphere, is limited by light availability in soil
(Kainulainen and Holopainen, 2002, Insam and Seewald, 2010).
Bacteria encountering monoterpenes have to deal with their toxic effects (reviewed by
Bakkali et al., 2008). In order to prevent the accumulation of monoterpenes in the cell and
cytoplasmatic membrane, bacteria modify their membrane lipids, transform monoterpenes
and use active transport by efflux pumps (Papadopoulos et al., 2008, Martinez et al., 2009).
Below toxic concentrations monoterpenes are used by microorganisms as sole carbon and
energy source. The mineralization of the hydrocarbons requires the introduction of functional
groups to access beta-oxidation like fragmentation reactions yielding central metabolites, e.g.
- 20 -
Chapter II - Microbial monoterpene transformations - a review
acetyl-CoA. In many aerobic microorganisms molecular oxygen serves as reactive agent to
functionalize the monoterpenes (Fig. 1).
Figure 1: Selected monoterpene transformations. (a) (+)-camphor [1] hydroxylation to 5hydroxycamphor [2]; (b) 1,8-cineole [10] hydroxylation to hydroxyl-1,8-cineole [11]; (c) D-pinene [3]
epoxidation to D-pinene oxide [5]; (d) (R)-limonene [6] hydroxylation to perillyl alcohol [15]; (e)
myrcene [25] hydration to (S)-(+)-linalool [17] and isomerization to geraniol [24].
Strains of Pseudomonas and Rhodococcus have become model organisms for the elucidation
of pathways in aerobic bacteria. Nearly 40 years after the first reports on aerobic
mineralization (Seubert, 1960, Seubert and Fass, 1964, Dhavalikar and Bhattacharyya, 1966,
Dhavalikar et al., 1966), the mineralization of monoterpenes in denitrifying bacteria and
methanogenic communities was discovered (Harder and Probian, 1995, Harder and Foss,
1999). Betaproteobacterial strains of the genera Castellaniella and Thauera are the study
objects for the elucidation of anaerobic pathways. All these bacteria were obtained in singlefed batch enrichments with high substrate concentrations (mmol
concentrations in nature (μmol
‫כ‬
‫כ‬
L−1), in contrast to low
L−1). Consequently, in batch enrichments isolated strains
exhibit often a solvent tolerance; they grow in the presence of a pure monoterpene phase.
- 21 -
Chapter II - Microbial monoterpene transformations - a review
Cultivation was rarely attempted by physical separation followed by single - fed batch
cultivations. Such dilution-to-extinction series performed in replicates - also known as mostprobable-number (MPN) method - revealed a frequent presence of the degradative capacities
in natural populations: denitrifying communities in sewage sludge and forest soil yielded 106
to 107 monoterpene-utilizing cells mL−1, representing 0.7 to 100 % of the total cultivable
nitrate-reducing microorganisms (Harder et al., 2000). MPN cultivations for aerobic bacteria
have not been reported so far, and for both cases the highly abundant bacteria with the
capacity to grow on monoterpenes have not been identified.
Over the last 50 years, many monoterpene transformations have been reported for microbial
cultures, but the biochemical pathways were rarely disclosed. More important for the
maintenance of our knowledge, only a small portion of the investigated strains were deposited
in culture collections. Without detailed knowledge of genes or the availability of strains, the
observations of biotransformation experiments are of limited value for future studies.
Therefore, this review on the transformation of monoterpenes focusses on enzymes for which
the gene and protein sequences are available in public databases as well as on microorganisms
that at least have been deposited in a public culture collection and ideally are validly
described (Table 1). A broad overview on microbial biotransformations is also provided by a
number of older review articles (Trudgill, 1990, Trudgill, 1994, van der Werf et al., 1997,
Hylemon and Harder, 1998, Duetz et al., 2003, Ishida, 2005, Li et al., 2006, Bicas et al., 2009,
Li and Lan, 2011, Schewe et al., 2011, Tong, 2013). KEGG and MetaCyc, two widely used
reference datasets of metabolic pathways, include degradation pathways of limonene, pinene,
geraniol and citronellol. Single reactions of p-cymene and p-cumate degradation are covered.
MetaCyc additionally covers the metabolism of myrcene, camphor, eucalyptol, and carveol.
Bicyclic monoterpenes
(+)-Camphor [1, Fig. 2] (C10H16O) is the substrate of one of the first and best described
monoterpene transforming enzymes, a specific cytochrome P450 monooxygenase (camABC,
P450cam, EC 1.14.15.1) from Pseudomonas putida (ATCC 17453). Initially, (+)-camphor is
hydroxylated. The resulting 5-exo-hydroxycamphor [2] is oxidized by a NAD-reducing
dehydrogenase (EC 1.1.1.327) which gene camD is part of the operon camDCAB. The
- 22 -
Chapter II - Microbial monoterpene transformations - a review
diketone is oxidized in a Baeyer–Villiger like oxidation to a lactone, either by a 2,5diketocamphane
1,2-monooxygenase
or
a
3,6-diketocamphane
1,6-monooxygenase
(camE25−1E25−2 or camE36, EC 1.14.13.162). The lactone spontaneously hydrolyses to 2-oxo- '3-4,5,5-trimethylcyclopentenyl-acetic acid which is activated as coenzyme A thioester by a
specific synthase (camF1,2, EC 6.2.1.38). This CoA-ester serves as substrate for another
specific monooxygenase (camG, EC 1.14.13.160), which initiates the cleavage of the second
ring by formation of a lactone. After hydrolysis of the lactone, the linear product is oxidized
to isobutanoyl-CoA and three acetyl-CoA. All corresponding genes (camABCDEFG) have
been identified on a linear plasmid (Ougham et al., 1983, Taylor and Trudgill, 1986, Aramaki
et al., 1993, Kadow et al., 2012, Leisch et al., 2012, Iwaki et al., 2013).
Figure 2: (+)-camphor [1]; D-pinene [3]; E-pinene [4].
The most abundant bicyclic monoterpene is pinene with the isomers α-pinene [3] and βpinene [4] (C10H16), a main constituent of wood resins (e.g. conifers). Pseudomonas rhodesiae
(CIP 107491) and P. fluorescens (NCIMB 11671) grew on α-pinene as sole carbon source. αpinene is oxidized to α-pinene oxide [5] by a NADH-dependent α-pinene oxygenase (EC
1.14.12.155) and undergoes ring cleavage by action of a specific α-pinene oxide lyase (EC
5.5.1.10), forming apparently isonovalal as first product which is isomerized to novalal (Best
et al., 1987, Bicas et al., 2008, Linares et al., 2009). The cleavage reaction of α-pinene oxide
was also described for a Nocardia sp. strain P18.3 (Griffiths et al., 1987, Trudgill, 1990,
Trudgill, 1994).
An alternative route for pinene degradation via a mono-cyclic p-menthene derivate has been
described for Pseudomonas sp. strain PIN (Yoo and Day, 2002).
Bacillus pallidus BR425
degrades α- and β-pinene apparently via limonene [6] and pinocarveol. While α-pinene is
transformed into limonene and pinocarveol, β-pinene yields pinocarveol only. Both
intermediates may be further transformed into carveol [7] and carvone. The activity of a
specific monooxygenases has been suggested, but experimental evidence is lacking (Savithiry
- 23 -
Chapter II - Microbial monoterpene transformations - a review
et al., 1998). Serratia marcescens uses α-pinene as sole carbon source. Trans-verbenol [8]
was a detectable metabolite. In glucose and nitrogen supplemented medium, this strain
formed α-terpineol [9]. The two oxidation products were considered to be dead-end products
as they accumulated in cultures (Wright et al., 1986). A general precaution has to be
mentioned here for many biotransformation studies: monoterpenes contain often impurities
and oxidation products which may be utilized as substrates resulting in traces of monoterpene
and monoterpenoid transformation products that are not further metabolized. Stoichiometric
experiments have to show that the amount of metabolite is larger than the amount of impurity
in the substrate. Only such careful stoichiometric experiments, mutants in functional genes or
the characterization of enzymes in vitro can provide a proof of the presence of a
biotransformation.
Eucalyptol, the bicyclic monoterpene 1,8-cineole [10] (C10H18O), is transformed in several
pathways. Novosphingobium subterranea converts 1,8-cineole initially into 2-endohydroxycineole, 2,2-oxo-cineole and 2-exo-hydroxycineole. Acidic products from ring
cleavages have been identified in situ (Rasmussen et al., 2005). Hydroxy-cineole formation
occurred in 1,8-cineole-grown cultures of Pseudomonas flava (Carman et al., 1986). A
cytochrome P450 monooxygenase from Bacillus cereus UI-1477 catalyzes the hydroxylation
of 1,8-cineole, yielding either 2R-endo- or 2R-exo-hydroxy-1,8-cineole [11] (Liu and
Rosazza, 1990, Liu and Rosazza, 1993). Another 1,8-cineole-specific P450 monooxygenase
(EC 1.14.13.156) has been purified and characterized from Citrobacter braakii, which yielded
2-endo-hydroxy-1,8-cineole only. Further oxidation and lactonization were followed by a
spontaneous lactone ring hydrolysis (Hawkes et al., 2002). Biotransformation in Rhodococcus
sp. C1 involves an initial hydroxylation to 6-endo-hydroxycineol [12] and further oxidation to
6-oxocineole by a 6-endo-hydroxycineol dehydrogenase (EC 1.1.1.241). A 6-oxocineole
monooxygenase (EC 1.14.13.51) converts the ketone into an unstable lactone. Spontaneous
decomposition results in (R)-5,5-dimethyl-4-(3-oxobutyl)-4,5-dihydrofuran-2(3H)-one. An
initial monooxygenase activity has not been detected in cell-free systems, while the
dehydrogenase and oxygenase activities have been measured in crude cell extracts (Williams
et al., 1989).
- 24 -
Chapter II - Microbial monoterpene transformations - a review
Monocyclic monoterpenes
Limonene [6, Fig. 3] (C10H16) is the most abundant mono-cyclic monoterpene, besides
toluene the second most abundant VOC indoors (Brown et al., 1994). It represents the main
component of essential oils from citrus plants, e.g. lemon and orange.
Figure 3: (R)-limonene [6]; carveol [7]; trans-verbenol [8].
Rhodococcus erythropolis DCL14 transforms (R,S)-limonene via limonene-1,2-epoxide into
limonene-1,2-diol [13, Fig. 5], applying a limonene-1,2 monooxygenase (EC 1.14.13.107)
and a limonene-1,2-epoxide hydrolase (EC 3.3.2.8), respectively. A specific dehydrogenase
(EC 1.1.1.297) forms the ketone, 1-hydroxy-2-oxolimonene, which is oxidized to a lactone by
a 1-hydroxy-2-oxolimonene 1,2-monooxygenase (EC 1.14.13.105). Enzyme activities were
only detected in limonene-induced cells, suggesting a tight regulation of the limonene
degradation. R. erythropolis DCL14 harbors a second pathway for limonene degradation.
Initially, (R)-limonene is hydroxylated by a NADPH-dependent limonene 6-monooxygenase
(EC 1.14.13.48) to trans-carveol [7]. Subsequently, trans-carveol is oxidized to carvone and
dihydrocarvone by a carveol dehydrogenase (EC 1.1.1.243) and carvone reductase (EC
1.3.99.25), respectively. A monocyclic monoterpene ketone monooxygenase (EC
1.14.13.105) inserts an oxygen atom, forming isopropenyl-7-methyl-2-oxo-oxepanone [14,
Fig. 6]. This lactone is cleaved by a specific ε-lactone hydrolase (EC 3.1.1.83) yielding
hydroxyl-3-isopropenyl-heptanoate. Oxidation and activation as coenzyme A thioester enable
a further degradation in accordance to the beta-oxidation (van der Werf et al., 1999b, van der
Werf and Boot, 2000). R. opacus PWD4 uses (R)-limonene on the same pathway. Biomass
from a glucose-toluene chemostat culture transformed limonene into trans-carveol, which was
further oxidized to carvone by a trans-carveol dehydrogenase (EC 1.1.1.275) (Duetz et al.,
2001).
- 25 -
Chapter II - Microbial monoterpene transformations - a review
Figure 4: D-terpineol [9]; 1,8-cineole [10]; 6-hydroxycineol [12].
Figure 5: limonene-1,2-diol [13]; perillyl alcohol [15]; cryptone [16].
Figure 6: isopropenyl-7-methyl-2-oxo-oxepanone [14]; J-terpineol [18]; p-menth-1-ene-6,8-diol [19].
Studies on the limonene metabolism in P. gladioli identified α-terpineol [9, Fig. 4] and
perillyl alcohol [15] as major metabolites. However, none of the involved enzymes has been
purified or further characterized (Cadwallader et al., 1989). A α-terpineol dehydratase from P.
gladioli was isolated and partially purified. A D-terpineol dehydratase from P. gladioli was
isolated and partially purified. The hydration reaction to the isopropenyl double bond of (4R)(+)-limonene resulted in (4R)-(+)-α-terpineol as only product (Cadwallader et al., 1992).
Geobacillus stearothermophilus (ex Bacillus) showed growth on limonene as sole carbon
source. The main limonene transformation product was perillyl alcohol, while α-terpineol and
perillyl aldehyde were found in minor concentrations. After heterologous expression of a
- 26 -
Chapter II - Microbial monoterpene transformations - a review
putative limonene degradation pathway in E. coli, α-terpineol was identified as major product
of the biotransformation. Other studies reported a limonene hydroxylation on the methyl
group yielding perillyl alcohol, which underwent further oxidation to perillic acid (Chang and
Oriel, 1994, Chang et al., 1995). Additional studies on the recombinant limonene hydroxylase
confirmed the production of perillyl alcohol from limonene but revealed in addition the
formation of carveol. The limonene hydroxylase showed dependency on molecular oxygen
and NADH as cofactors and was suggested to belong to the (S)-limonene 7-monooxygenase
family (EC 1.14.13.49) (Cheong and Oriel, 2000).
Enterobacter agglomerans 6L and Kosakonia cowanii 6L (ex Enterobacter cowanii)
transformed (R)-limonene [6]. The main metabolites detected in ether extracts of E.
agglomerans 6L cultures were γ-valerolactone and cryptone [16]. In assays using four
recombinant expressed limonene-transforming enzymes from K. cowanii 6L, linalool [17, Fig.
8] was identified as main product besides smaller amounts of dihydrolinalool. It was proposed that the potential limonene hydroxylase converts limonene into linalool, perillyl
alcohol, α-terpineol and γ-terpineol [18] (Park et al., 2003, Yang et al., 2007).
Pseudomonas putida (MTCC 1072) converts limonene to p-menth-1-ene-6,8-diol [19] and
perillyl alcohol (Chatterjee and Bhattacharyya, 2001). No sequence information was found in
public databases. Two other strains of Pseudomonas putida (F1 and GS1) have been found to
convert (+)-limonene to perillic acid in co-substrate fed-batch cultures (Speelmans et al.,
1998). Experimental results indicated the participation of the p-cymene pathway (CYM)
(Mars et al., 2001). Castellaniella defragrans grows anaerobically on cyclic monoterpenes as
sole carbon and energy source under denitrifying conditions (Foss et al., 1998). Recent
experiments suggested an oxygen-independent hydroxylation on the methyl group of
limonene to perillyl alcohol as the initial activation step, followed by subsequent oxidation to
perillic acid (Petasch et al., 2014).
- 27 -
Chapter II - Microbial monoterpene transformations - a review
Figure 7: p-cymene [20]; p-cumate [21]; menthol [22].
Figure 8: (S)-linalool [17]; citronellol [23]; geraniol [24]; E-myrcene [25].
P-cymene [20, Fig. 7] (C10H14) is an aromatic monoterpene (p-isopropyl-toluene).
Pseudomonas putida F1 (ATCC 700007) degrades p-cymene to p-cumate [21] via the CYMpathway (cymBCAaAbDE). A two-component p-cymene monooxygenase (cymAaAb, EC
1.14.13.-) introduces a hydroxyl group on the methyl group of p-cymene. The resulting pcumic alcohol is oxidized to the corresponding carboxylic acid by an alcohol and an aldehyde
dehydrogenase (cymB and cymC, EC 1.1.1.- and EC 1.2.1.-). The genes cymD and cymE
encode for a putative outer membrane protein and an acetyl coenzyme A synthetase,
respectively. However, their role in the pathway remains unclear (Eaton, 1997). Upstream of
the cym-operon, the genes for the further degradation of p-cumate are located. They are
organized in another operon and comprise eight genes (cmtABCDEFGH). P. putida F1 has
been shown to use p-cumate as sole carbon source. It is hydroxylated by a ferredoxin
dependent p-cumate 2,3-dioxygenase. The genes cmtAaAd encode a ferredoxin reductase and
a ferredoxin, and cmtAbAc encode the large and the small subunits of the dioxygenase (EC
1.14.12.-). The resulting cis-2,3-dihydroxy-2,3-dihydro-p-cumate is oxidized and ring
cleavage occurs by introduction of another oxygen molecule. The responsible enzymes are a
- 28 -
Chapter II - Microbial monoterpene transformations - a review
specific dehydrogenase (cmtB, EC 1.3.1.58) and a 2,3-dihydroxy-p-cumate dioxygenase
(cmtC, EC 1.13.11.-), respectively. Further degradation is accomplished by a decarboxylation
and elimination of an isobutyrate molecule, catalyzed by a 2-hydroxy-3-carboxy-6-oxo-7methylocta-2,4-dienoate decarboxylase (cmtD, EC 4.1.1.-) and a 2-hydroxy-6-oxo-7methylocta-2,4-dienoate hydrolase (cmtE, EC 3.7.1.-). The product, 2-hydroxypenta-2,4dienoate, undergoes a water addition by a specific hydratase (cmtF, EC 4.2.1.80). Then, a
carbon-carbon lyase reaction yields pyruvate and acetaldehyde, catalyzed by 2-oxo-4hydroxyvalerate aldolase (cmtG, EC 4.1.3.39). Acetaldehyde is oxidized and enters as acetylCoA the citrate cycle (Eaton, 1996).
Thauera terpenica 21Mol utilizes menthol [22] as sole carbon source. The proposed
degradation mechanism involves two initial oxidation reactions leading to menth-2-enone,
followed by a hydration and an additional oxidation step. Finally, ring cleavage may occur
and the molecule is attached to coenzyme A to yield 3,7-dimethyl-5-oxo-octyl-CoA (Foss and
Harder, 1998, Hylemon and Harder, 1998).
Acyclic monoterpenes
First studies on acyclic monoterpenoids in the early sixties by Seubert and colleagues
described the degradation of citronellol [23], geraniol [24], and nerol via an oxidation of the
alcohol to an acid, followed by the formation of a CoA-thioester and subsequent betaoxidation in Pseudomonas citronellolis (ATCC 13674) (Seubert, 1960, Seubert et al., 1963,
Seubert and Remberger, 1963, Seubert and Fass, 1964). This knowledge has been extended
towards other Pseudomonas strains (Cantwell et al., 1978). The complete degradation
pathway has been classified as the acyclic terpene utilization and leucine utilization
(ATU/LIU) pathway involving the genes atuABCDEFGH and liuRABCDE. After the initial
formation of cis-geranyl-CoA, a geranyl-coenzyme-A carboxylase (atuCF, EC 6.4.1.5)
elongates the methylgroup. A hydroxyl group is introduced by an isohexenyl-glutaconyl-CoA
hydratase (atuE, EC 4.2.1.57), followed by a water addition and elimination of an acetate
molecule catalyzed by a 3-hydroxy-3-isohexenylglutaryl-CoA lyase (liuE, EC 4.1.3.26). The
resulting 7-methyl-3-oxooct-6-enoyl-CoA is further degraded via two beta-oxidation like
reactions to yield 3-methylcrotonyl-CoA, which enters the leucine degradation pathway
- 29 -
Chapter II - Microbial monoterpene transformations - a review
(liuRABCDE) (Höschle et al., 2005, Aguilar et al., 2006, Förster-Fromme et al., 2006, FörsterFromme and Jendrossek, 2010, Chávez-Avilés et al., 2010). Citronellol degradation is
reported for many Pseudomonas strains, including P. aeruginosa PAO1 (ATCC 15692), P.
mendocina (ATCC 25411), and P. delhiensis (DSM 18900) (Cantwell et al., 1978, Prakash et
al., 2007, Förster-Fromme and Jendrossek, 2010). Among the few reactions described in
detail is a molybdenum dependent dehydrogenase responsible for the geranial oxidation to
geranylate in P. aeruginosa PAO1 (Höschle and Jendrossek, 2005).
The acyclic monoterpene β-myrcene [25] (C10H16) is transformed by Pseudomonas
aeruginosa (PTCC 1074) into dihydrolinalool, 2,6-dimethyloctane and α-terpineol. Limonene
has been proposed as possible intermediate in α-terpineol formation but was not detected in
the culture broth (Esmaeili and Hashemi, 2011). Pseudomonas sp. M1 accomplishes
degradation by hydroxylation on the C8 position to myrcene-8-ol, which is further oxidized,
linked to coenzyme A and metabolized in a beta-oxidation like manner (Iurescia et al., 1999).
The formation of geraniol from β-myrcene has been observed with resting cells of
Rhodococcus erythropolis MLT1, regardless of the presence of a cytochrome P450 inhibitor.
The reaction was dependent on aerobic conditions, however it remains unclear if a
monooxygenase or lyase system is involved (Thompson et al., 2010).
The tertiary alcohol linalool is also transformed at the C8 position. A linalool monooxygenase
(EC 1.14.13.151) has been described in P. putida PpG777 and Novosphingobium
aromaticivo-rans (ATCC 700278D-5) (Ullah et al., 1990, Bell et al., 2010). In the absence of
molecular oxygen, Castellaniella defragrans 65Phen has an unique enzyme for the linalool
transformation, the linalool dehydratase-isomerase (Brodkorb et al., 2010). Castellaniella and
Thauera strains were the first anaerobic microorganisms shown to anaerobically degrade and
mineralize monoterpenes (Harder and Probian, 1995, Harder et al., 2000). The linalool
dehydratase-isomerase (EC 4.2.1.127 and 5.4.4.4) of C. defragrans 65Phen catalyzes a regioand stereo-specific hydration of β-myrcene yielding the tertiary alcohol (S)-(+)-linalool [17]
and the isomerization to the primary alcohol geraniol (Brodkorb et al., 2010, Lueddeke and
Harder, 2011). Geraniol and geranial dehydrogenases formed geranic acid (Heyen and
Harder, 2000, Lueddeke et al., 2012). T. linaloolentis 47Lol grows on linalool as sole carbon
and energy source. A similar isomerization of linalool to geraniol with subsequent oxidation
of geraniol to geranial has been observed in cultures (Foss and Harder, 1997).
- 30 -
Chapter II - Microbial monoterpene transformations - a review
Monoterpene transformation by fungi
Fungi excrete laccases which are copper-containing oxidases. Utilizing molecular oxygen as a
cosubstrate, an unspecific oxidation of organic molecules is initiated by these enzymes.
Additionally, fungi express a variety of cytochrome P450 mono-and di-oxygenases. Thus,
several fungi were described to transform monoterpenes during growth in rich medium
(reviewed by Farooq et al., 2004). Species with a reported capacity to transform
monoterpenes are Aspergillus niger, Botrytis cinerea, Diplodia gossypina, Mucor
circinelloides, Penicillium italicum, Penicillium digitatum, Corynespora cassiicola, and
Glomerella cingulata. For a long time, no species have been described to use monoterpenes as
sole carbon and energy source for growth (Trudgill, 1994 and references therein). Recently,
Grosmannia clavigera, a bark beetle-associated fungal pathogen of pine trees, was shown to
grow on a mono- and diterpene mixture, containing α/β-pinene and 3-carene (DiGuistini et
al., 2011). ABC efflux transporter and cytochrome P450 enzymes confer a monoterpene
resistance to the blue-stain fungi (Lah et al., 2013, Wang et al., 2013).
Monoterpenes in the carbon cycle
Habitats with a dense vegetation of wood and flowers are expected to contain larger
populations of monoterpene transforming microorganisms. Whereas coniferous forests emit
up to 6.7 g carbon ‫ כ‬m−2 ‫ כ‬yr−1, broadleaf evergreen forest and grassland emit only 3.5 and 2.5
g carbon ‫ כ‬m−2 ‫ כ‬yr−1, respectively (Tanaka et al., 2012). Monoterpene emission rates between
0.3 and 7 g carbon ‫ כ‬m−2 ‫ כ‬yr−1 for the United States - mainly α- and β-pinene, limonene and βmyrcene (Geron et al., 2000) - can support the aerobic growth of 0.15–3.5 g bacteria ‫ כ‬m−2
‫כ‬
yr−1, assuming 50 % of carbon incorporated into biomass. This is a significant potential,
considering the presence of around 10 g microbial biomass in the top centimeter of soil per
square meter.
In marine systems, isoprene and monoterpenes (mainly α-pinene) are produced by
phytoplankton and algae and partially emitted into the atmosphere (reviewed by Yassaa et al.,
2008, Shaw et al., 2010). Isoprene emission was estimated to 0.2–1.2 Tg carbon ‫ כ‬yr−1 (Palmer
and Shaw, 2005, Gantt et al., 2009, Shaw et al., 2010). For the ocean surface area this results
in an emission rate of 0.0025 g carbon
‫כ‬
m−2
‫כ‬
- 31 -
yr−1. Current uncertainties in the size of
Chapter II - Microbial monoterpene transformations - a review
emission based on shipborne measurements in comparison to satellite data (Luo and Yu,
2010) may be resolved by incorporating an export from the continental atmosphere to the
oceanic atmosphere (Hu et al., 2013). Isoprene-amended samples from marine habitats were
enriched in bacteria affiliating with Actinobacteria, Alphaproteobacteria, and Bacteroidetes
and first strains were shown to degrade isoprene and aliphatic hydrocarbons (Acuña Alvarez
et al., 2009).
In summary, these findings indicate a higher abundance of monoterpene transforming and
mineralizing bacteria in soils than in the ocean. Indeed, most monoterpene trans-forming
bacteria have been enriched or isolated from soil and freshwater samples in habitats with
monoterpene emitting vegetation.
Databases for pathway analysis and a look at metagenomes
Databases are nowadays available for the analysis of enzymatic reactions and metabolic
pathways in metagenomic and genomic sequence datasets. The most relevant are the Kyoto
Encyclopedia of Genes and Genomes (KEGG), MetaCyc and the Biocatalysis/Biodegradation
database of the University of Minnesota.
First studies used KEGG to identify monoterpene-related genes in metagenomes of
microbiomes in insects and nematodes feeding on a monoterpene-rich diet. Pine beetles
encounter the high terpenoid concentrations of conifers and may take advantage of
detoxification processes catalyzed by their symbionts/microbiomes (Adams et al., 2013). The
KEGG pathway for limonene and pinene degradation (ko00903) was used to identify genes
encoding enzymes putatively involved in monoterpene degradation. Five enzymes were
present and more abundant in the metagenomes than in a combined metagenomic set of plant
biomass-degrading communities. These enzymes were an aldehyde dehydrogenase, an
oxidoreductase, an enoyl-CoA hydratase and two hydratases/epimerases. Whether these genes
are truly involved in monoterpene metabolism or the degradation of cyclic compounds, e.g.
related aromatic lignin monomers, is an open question. Taxonomically, these genes affiliated
with the genera Pseudomonas, Rahnella, Serratia, and Stenotrophomonas.
The pinewood nematode Bursaphlenchus xylophilus transcribes cytochrome P450 genes as
main metabolic pathway for xenobiotics detoxification, but not all enzymes needed for
- 32 -
Chapter II - Microbial monoterpene transformations - a review
terpenoid metabolism were detected by transcriptomic analysis. Metagenomic data of
nematode bacterial symbionts included the complete α-pinene degradation pathway (Cheng et
al., 2013). Annotation based on KEGG revealed that the degradation pathways for limonene
and pinene (map00903) and for geraniol (map00281) accounted for 2.5 % of mapped
metagenes. The majority of these genes affiliated to Pseudomonas, Achromobacter, and
Agrobacterium. Strains isolated from the nematode and capable of growth on α-pinene
affiliated to Pseudomonas, Achromobacter, Agrobacterium, Cytophaga, Herbaspirillum, and
Stenotrophomonas.
Conclusion
The synthesis and transformation of BVOCs, especially terpenoids, by plants is well studied
(Kesselmeier and Staudt, 1999). Corresponding pathways have been elucidated and a variety
of corresponding enzymes have been isolated and characterized (Mahmoud and Croteau,
2002, Yu and Utsumi, 2009). In contrast, the exploration of the microbial transformation and
mineralization of monoterpenes has accumulated a small coverage of the field. Simply, over
the last 50 years, research on bacterial monoterpene metabolism had only found the interest of
very few principal investigators. Now, large sequence datasets of organisms and biological
communities provide an unprecedented insight into the diversity of pathways and provide us
with challenging hypotheses. However, the basis for the annotation is the biochemical
characterization of enzymes which is only available for few monoterpenes. Only three
pathways are completely known on the genetic and enzymatic level: the ones for camphor
(CAM), p-cymene (CYM/CMT), and citronellol/geraniol (ATU/LIU). For pinene, the gene
for a key enzyme, the α-pinene oxide lyase (EC 5.5.1.10), is still unknown. The lack of such a
key enzyme sequence for a KEGG pathway (map00903) illustrates our uncertainty in the
interpretation of metagenomic and genomic datasets. Progress in proteomic and metabolomic
analyses in the last years support now biochemical and genetic experiments which will swiftly
reveal the desired identification of key enzymes in the monoterpene metabolism.
- 33 -
Pseudomonas
fluorescens
NCIMB 11671
Pseudomonas
fluorescens
NCIMB 11671
D-pinene
monooxygenase
D-pinene oxide lyase
1,8-cineole 2-endomonooxygenase
Monocyclic
monoterpene ketone
monooxygenase
Limonene 1,2-diol
dehydrogenase
Limonene 1,2monooxygenase
1.14.13.155
5.5.1.10
1.14.13.156
1.14.13.105
1.1.1.297
1.14.13.107
Rhodococcus
erythropolis DCL
14
Rhodococcus
erythropolis DCL
14
Rhodococcus
erythropolis DCL
14
Citrobacter
braakii
Pseudomonas
rhodesiae PF1 (CIP
107491)
Organism
Enzyme name
EC
number
- 34 -
(R)-limonene
Limonene 1,2diol
(-)-menthone
1,8-cineole
1,2-epoxymenth-8-ene
1-hydroxy-pmenth-8-en-2one
NAD+
Oxygen,
NAD(P)H
(4R,7S)-4methyl-7(propan-2yl)oxepan-2-one
2-endo-hydroxy1,8-cineole
Water,
NAD(P)+
NADH
Water,
NADP+
Water,
NADP+
Water,
NAD+
D-pinene oxide
(E)-2,6dimethyl-5methylidenehept-2-enal (isonovalal)
Co-product
Product
Oxygen,
NADPH
Oxygen,
NADPH
Oxygen,
NADH
D-pinene
D-pinene oxide
Co-substrate
Substrate
Table 1: Summary table of monoterpene transforming enzymes in validly described species of bacteria.
(van der
Werf et al.,
1999b)
(van der
Werf et al.,
1999b)
(van der
Werf et al.,
1999b)
(Hawkes et
al., 2002)
(Fontanille
et al.,
2002)
(Best et al.,
1987)
(Best et al.,
1987)
Reference
Chapter II - Microbial monoterpene transformations - a review
Rhodococcus
opacus PWD4
(DSM 44313)
Rhodococcus
erythropolis DCL
14
Carvone reductase
Trans-carveol
dehydrogenase
Monoterpene H-lactone
hydrolase
(4R)-limonene-1,2epoxide hydrolase
(1S,2S,4R)-limonene1,2-diol dehydrogenase
1.3.99.25
1.1.1.275
3.1.1.83
3.3.2.8
1.1.1.297
Rhodococcus
erythropolis DCL
14
Rhodococcus
erythropolis DCL
14
Rhodococcus
erythropolis DCL
14
Rhodococcus
erythropolis DCL
14
Carveol dehydrogenase
1.1.1.243
Rhodococcus
erythropolis DCL
14
Organism
(S)-limonene 6monooxygenase
Enzyme name
1.14.13.48
EC
number
- 35 -
Menth-8-ene1,2-diol
1,2-epoxy-pmenth-8-ene
(4S,7R)-7methyl-4-prop1-en-2yloxepan-2-one
(+)-transcarveol
(+)dihydrocarvone
(-)-trans-carveol
(S)-limonene
Substrate
Menth-8-ene1,2-diol
1-hydroxy-pmenth-8-en-2one
Water
NAD+
6-hydroxy-3prop-1-en-2ylheptanoate
(+)-carvone
NAD+
Water
(-)-carvone
(-)-carvone
NADP+
Oxidized
electron
acceptor
(-)-trans-carveol
Product
Oxygen,
NADPH
Co-substrate
NADH
NADH
Reduced
electron
acceptor
NADPH
Water,
NADP+
Co-product
(van der
Werf et al.,
1999a)
(van der
Werf et al.,
1999a)
(van der
Werf et al.,
1999a)
(Duetz et
al., 2001)
(van der
Werf et al.,
1999b)
(van der
Werf et al.,
1999b)
(van der
Werf et al.,
1999b)
Reference
Chapter II - Microbial monoterpene transformations - a review
Linalool dehydratase
(-isomerase)
Linalool (dehydratase)
-isomerase
Geraniol
dehydrogenase
Geranial dehydrogenase Castellaniella
defragrans 65Phen
(DSM 12143)
4.2.1.127
5.4.4.4
1.1.1.347
1.2.1.86
Castellaniella
defragrans 65Phen
(DSM 12143)
Castellaniella
defragrans 65Phen
(DSM 12143)
Castellaniella
defragrans 65Phen
(DSM 12143)
Pseudomonas
putida PpG777
Novosphingobium
aromaticivorans
ATCC 700278D-5
Linalool 8monooxgenase
1.14.13.151
Geobacillus
stearothermophilus
(ex Bacillus strain
BR388)
Organism
(S)-limonene 7monooxygenase
Enzyme name
1.14.13.49
EC
number
- 36 -
Geranial
Geraniol
(S)-(+)-linalool
E-myrcene
Linalool
(S)-limonene
Substrate
Water,
NAD+
NAD+
Water
2 oxygen,
2 NADH
Oxygen,
NADPH
Co-substrate
Geranylate
Geranial
Geraniol
(S)-(+)-linalool
(6E)-8oxolinalool
(-)-perillyl
alcohol
Product
NADH
NADH
3 Water,
2 NAD+
Water,
NADP+
Co-product
(Lueddeke
et al.,
2012)
(Lueddeke
et al.,
2012)
(Brodkorb
et al.,
2010)
(Brodkorb
et al.,
2010)
(Ullah et
al., 1990)
(Bell et al.,
2010)
(Cheong
and Oriel,
2000)
Reference
Chapter II - Microbial monoterpene transformations - a review
p-cymene
monooxygenase,
hydroxylase
subunit(CymAa)
p-cymene
monooxygenase,
reductase subunit
(CymAb)
p-cumic alcohol
dehydrogenase (CymB)
p-cumic aldehyde
dehydrogenase(CymC)
Putative outer
membrane protein,
unknown
function(CymD)
Acetyl-CoA synthetase
(CymE)
1.14.13.-
1.1.1.-
1.2.1.-
-.-.-.-
6.2.1.1
Enzyme name
1.14.13.-
Cym
pathway
EC
number
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Organism
- 37 -
Acetate
p-cumic
aldehyde
p-cumic alcohol
p-cymene
p-cymene
Substrate
CoA, ATP
Acetyl-CoA
p-cumic acid
p-cumic
aldehyde
NAD+
Water,
NAD+
p-cumic alcohol
p-cumic alcohol
Product
Oxygen,
NADH
Oxygen,
NADH
Co-substrate
Diphosphate,
AMP
NADH
NADH
Water,
NAD+
Water,
NAD+
Co-product
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1997)
(Eaton,
1997)
Reference
Chapter II - Microbial monoterpene transformations - a review
p-cumate 2,3dioxygenase
(CmtAaAbAcAd)
2,3-dihydroxy-2,3dihydro-p-cumate
dehydrogenase (CmtB)
2,3-dihydroxy-pcumate dioxygenase
(CmtC)
2-hydroxy-3-carboxy6-oxo-7-methylocta2,4-dienoate
decarboxylase (CmtD)
2-hydroxy-6-oxo-7methylocta-2,4dienoate hydrolase
(CmtE)
2-hydroxypenta-2,4dienoate hydratase
(CmtF)
1.3.1.58
1.13.11.-
4.1.1.-
3.7.1.-
4.2.1.80
Enzyme name
1.14.12.-
pathway
Cmt
EC
number
Pseudomonas
putida MT-2
(ATCC 33015)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Pseudomonas
putida F1 (ATCC
700007)
Organism
- 38 -
2-hydroxypenta-2,4dienoate
2-hydroxy-6oxo-7methylocta-2,4dienoate
2-hydroxy-3carboxy-6-oxo7-methylocta2,4-dienoate
2,3-dihydroxyp-cumate
Cis-2,3dihydroxy-2,3dihydro-pcumate
p-cumate
Substrate
Water
Water
2-oxo-4hydroxypentanoate
2-hydroxypenta2,4-dienoate
2-hydroxy-6oxo-7methylocta-2,4dienoate
2-hydroxy-3carboxy-6-oxo7-methylocta2,4-dienoate
2,3-dihydroxy-pcumate
NAD+
Oxygen
Cis-2,3dihydroxy-2,3dihydro-pcumate
Product
Oxygen,
NADH
Co-substrate
Isobutyrate
Carbon
dioxide
NADH
NAD+
Co-product
(Harayama
et al.,
1989)
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1996)
(Eaton,
1996)
Reference
Chapter II - Microbial monoterpene transformations - a review
2-oxo-4hydroxyvalerate
aldolase (CmtG)
Acetaldehyde
dehydrogenase (CmtH)
1.2.1.10
Enzyme name
4.1.3.39
EC
number
Pseudomonas
putida PG (DSM
8368)
Pseudomonas
putida PG (DSM
8368)
Organism
- 39 -
Acetaldehyde
2-oxo-4hydroxypentanoate
Substrate
NAD+,
CoA
Co-substrate
Acetyl-CoA
Acetaldehyde
Product
NADH
Pyruvate
Co-product
(Platt et al.,
1995)
(Platt et al.,
1995)
Reference
Chapter II - Microbial monoterpene transformations - a review
Pseudomonas
citronellolis
(ATCC 13674)
Pseudomonas
mendocina (ATCC
25411)
Putative citronellylCoA desaturase (AtuD)
Geranyl-CoA
carboxylase,
carboxylase alphasubunit (AtuF)
Geranyl-CoA
carboxylase,
carboxylase betasubunit (AtuC)
AtuC, AtuF
1.3.99.-
6.4.1.5
6.4.1.5
Pseudomonas
citronellolis
(ATCC 13674)
Pseudomonas
citronellolis
(ATCC 13674)
Pseudomonas
citronellolis
(ATCC 13674)
Putative citronellylCoA synthetase (AtuH)
6.2.1.-
Pseudomonas
citronellolis
(ATCC 13674)
Organism
Citronellol / citronellal
dehydrogenase (AtuB;
AtuG)
Enzyme name
1.1.99.- /
1.2.99.-
Atu
pathway
EC
number
- 40 -
Cis-geranylCoA
Cis-geranylCoA
Citronellyl-CoA
Citronellate
Citronellol /
citronellal
Substrate
Bicarbonate,
ATP
Bicarbonate,
ATP
Oxidized
electron
acceptor
CoA, ATP
Water,
oxidized
electron
acceptor
Co-substrate
Isohexenylglutaconyl-CoA
Isohexenylglutaconyl-CoA
Cis-geranyl-CoA
Citronellyl-CoA
Citronellal /
citronellate
Product
ADP,
phosphate
ADP,
phosphate
Reduced
electron
acceptor
Diphosphate,
AMP
Reduced
electron
acceptor
Co-product
(Cantwell
et al.,
1978)
(Fall and
Hector,
1977,
FörsterFromme et
al., 2006)
(FörsterFromme et
al., 2006)
(FörsterFromme et
al., 2006)
(FörsterFromme et
al., 2006)
(FörsterFromme et
al., 2006)
Reference
Chapter II - Microbial monoterpene transformations - a review
4.1.3.26
4.2.1.57
EC
number
Pseudomonas
aerugenosa PAO1
(ATCC 15692)
LiuE
Pseudomonas
mendocina (ATCC
25411)
AtuE
Pseudomonas
citronellolis
(ATCC 13674)
Pseudomonas
aeruginosa PAO1
(ATCC 15692)
AtuE
3-hydroxy-3-isohexenyl-glutarylCoA:acetate lyase
(LiuE)
Pseudomonas
citronellolis
(ATCC 13674)
Pseudomonas
aeruginosa PAO1
(ATCC 15692)
AtuC, AtuF
Isohexenyl-glutaconylCoA hydratase (AtuE)
Organism
Enzyme name
- 41 -
3-hydroxy-3isohexenylglutaryl-CoA
Isohexenylglutaconyl-CoA
Substrate
Water
Co-substrate
7-methyl-3-oxo6-octenoyl-CoA
3-hydroxy-3isohexenylglutaryl-CoA
Product
Acetate
Co-product
(ChávezAvilés et
al., 2010)
(FörsterFromme et
al., 2006,
ChávezAvilés et
al., 2010)
(Cantwell
et al.,
1978)
(Díaz-Pérez
et al., 2004,
Höschle et
al., 2005)
(FörsterFromme et
al., 2006)
(Díaz-Pérez
et al., 2004,
Höschle et
al., 2005)
Reference
Chapter II - Microbial monoterpene transformations - a review
Pseudomonas
putida (ATCC
17453)
- 42 -
3,6diketocamphane
Oxygen,
NADH
Oxygen,
NADH
(-)-5-oxo-1,2campholide
(+)-5-oxo-1,2campholide
2,5diketocamphane
/ 3,6diketocamphane
5-oxo-hydroxycamphor
Water,
NAD+
Water,
NAD+
NADH
Water,
oxidized
putidaredoxin
(Taylor and
Trudgill,
1986)
(Taylor and
Trudgill,
1986)
(Aramaki
et al.,
1993)
(Bell et al.,
2010)
3,6-diketocamphane
1,6-monooxygenase
(CamE36)
2,5diketocamphane
NAD+
Oxygen,
reduced
putidaredoxin
1.14.13.162
Pseudomonas
putida (ATCC
17453)
5-oxo-hydroxycamphor
(+)(-)-camphor
2,5-diketocamphane
1,2-monooxygenase
(CamE25-1, CamE25-2,
CamE36)
Pseudomonas
putida (ATCC
17453)
Novosphingobium
aromaticivorans
(ATCC 700278D5)
Pseudomonas
putida (ATCC
29607)
1.14.13.162
Reference
5-exo-hydroxycamphor
dehydrogenase (CamD)
Co-product
1.1.1.327
Product
(Poulos et
al., 1985)
Co-substrate
Camphor 5monooxygenase
(CamABC)
Substrate
1.14.15.1
Organism
(Iwaki et
al., 2013
and
references
therein)
Enzyme name
Cam
pathway
EC
number
Chapter II - Microbial monoterpene transformations - a review
Pseudomonas
putida (ATCC
17453)
Pseudomonas
putida (ATCC
17453)
2-oxo-Δ3-4,5,5trimethylcyclopentenyl
acetyl-CoA 1,2monooxygenase
(CamG)
1.14.13.160
Organism
(2,2,3-trimethyl-5oxocyclopent-3-enyl)
acetyl-CoA synthase
(CamF1, CamF2)
Enzyme name
6.2.1.38
EC
number
- 43 -
[(1R)-2,2,3trimethyl-5oxocyclopent-3enyl] acetylCoA
[(1R)-2,2,3trimethyl-5oxocyclopent-3enyl] acetate
Substrate
Oxygen,
NADPH
ATP, CoA
Co-substrate
[(2R)-3,3,4trimethyl-6-oxo3,6-dihydro-1Hpyran-2-yl]
acetyl-CoA
[(1R)-2,2,3trimethyl-5oxocyclopent-3enyl] acetyl-CoA
Product
Water,
NADP+
Diphosphate,
AMP
Co-product
(Ougham et
al., 1983,
Leisch et
al., 2012)
(Ougham et
al., 1983)
Reference
Chapter II - Microbial monoterpene transformations - a review
Chapter II - Microbial monoterpene transformations - a review
References
ACUÑA ALVAREZ, L., EXTON, D. A., TIMMIS, K. N., SUGGETT, D. J. & MCGENITY, T. J. 2009.
Characterization of marine isoprene-degrading communities. Environmental Microbiology, 11, 32803291.
ADAMS, A. S., AYLWARD, F. O., ADAMS, S. M., ERBILGIN, N., AUKEMA, B. H., CURRIE, C. R.,
SUEN, G. & RAFFA, K. F. 2013. Mountain pine beetles colonizing historical and native host trees are
associated with a bacterial community highly enriched in genes contributing to terpene metabolism.
Applied and Environmental Microbiology, 79, 3468-3475.
AGUILAR, J. A., ZAVALA, A. N., DÍAZ-PÉREZ, C., CERVANTES, C., DÍAZ-PÉREZ, A. L. &
CAMPOS-GARCÍA, J. 2006. The atu and liu clusters are involved in the catabolic pathways for
acyclic monoterpenes and leucine in Pseudomonas aeruginosa. Applied and Environmental
Microbiology, 72, 2070-2079.
AJIKUMAR, P. K., TYO, K., CARLSEN, S., MUCHA, O., PHON, T. H. & STEPHANOPOULOS, G.
2008. Terpenoids: Opportunities for biosynthesis of natural product drugs using engineered
microorganisms. Molecular Pharmaceutics, 5, 167-190.
ARAMAKI, H., KOGA, H., SAGARA, Y., HOSOI, M. & HORIUCHI, T. 1993. Complete nucleotidesequence of the 5-exo-hydroxycamphor dehydrogenase gene on the CAM plasmid of Pseudomonas
putida (ATCC-17453). Biochimica et Biophysica Acta, 1174, 91-94.
ATKINSON, R. & AREY, J. 2003. Atmospheric degradation of volatile organic compounds. Chemical
Reviews, 103, 4605-4638.
BAKKALI, F., AVERBECK, S., AVERBECK, D. & WAOMAR, M. 2008. Biological effects of essential
oils - A review. Food and Chemical Toxicology, 46, 446-475.
BELL, S. G., DALE, A., REES, N. H. & WONG, L. L. 2010. A cytochrome P450 class I electron transfer
system from Novosphingobium aromaticivorans. Applied Microbiology and Biotechnology, 86, 163175.
BEST, D. J., FLOYD, N. C., MAGALHAES, A., BURFIELD, A. & RHODES, P. M. 1987. Initial
enzymatic steps in the degradation of alpha-pinene by Pseudomonas fluorescens Ncimb 11671.
Biocatalysis and Biotransformation, 1, 147-159.
BICAS, J. L., DIONISIO, A. P. & PASTORE, G. M. 2009. Bio-oxidation of terpenes: An approach for
the flavor industry. Chemical Reviews, 109, 4518-4531.
BICAS, J. L., FONTANILLE, P., PASTORE, G. M. & LARROCHE, C. 2008. Characterization of
monoterpene biotransformation in two pseudomonads. Journal of Applied Microbiology, 105, 19912001.
- 44 -
Chapter II - Microbial monoterpene transformations - a review
BRODKORB, D., GOTTSCHALL, M., MARMULLA, R., LUEDDEKE, F. & HARDER, J. 2010.
Linalool dehydratase-isomerase, a bifunctional enzyme in the anaerobic degradation of monoterpenes.
Journal of Biological Chemistry, 285, 30436-30442.
BROWN, S. K., SIM, M. R., ABRAMSON, M. J. & GRAY, C. N. 1994. Concentrations of volatile
organic compounds in indoor air - a review. Indoor Air - International Journal of Indoor Air Quality
and Climate, 4, 123-134.
CADWALLADER, K. R., BRADDOCK, R. J. & PARISH, M. E. 1992. Isolation of alpha-terpineol
dehydratase from Pseudomonas gladioli. Journal of Food Science, 57, 241-&.
CADWALLADER, K. R., BRADDOCK, R. J., PARISH, M. E. & HIGGINS, D. P. 1989. Bioconversion
of (+)-limonene by Pseudomonas gladioli. Journal of Food Science, 54, 1241-1245.
CANTWELL, S. G., LAU, E. P., WATT, D. S. & FALL, R. R. 1978. Biodegradation of acyclic
isoprenoids by Pseudomonas species. Journal of Bacteriology, 135, 324-333.
CARMAN, R. M., MACRAE, I. C. & PERKINS, M. V. 1986. The oxidation of 1,8-cineole by
Pseudomonas flava. Australian Journal of Chemistry, 39, 1739-1746.
CHANG, H. C., GAGE, D. A. & ORIEL, P. J. 1995. Cloning and expression of a limonene degradation
pathway from Bacillus stearothermophilus in Escherichia coli. Journal of Food Science, 60, 551-553.
CHANG, H. C. & ORIEL, P. 1994. Bioproduction of perillyl alcohol and related monoterpenes by isolates
of Bacillus stearothermophilus. Journal of Food Science, 59, 660-&.
CHATTERJEE, T. & BHATTACHARYYA, D. K. 2001. Biotransformation of limonene by Pseudomonas
putida. Applied Microbiology and Biotechnology, 55, 541-546.
CHÁVEZ-AVILÉS, M., DÍAZ-PÉREZ, A. L. & CAMPOS-GARCÍA, J. 2010. The bifunctional role of
LiuE from Pseudomonas aeruginosa, displays additionally HIHG-CoA lyase enzymatic activity.
Molecular Biology Reports, 37, 1787-1791.
CHENG, X. Y., TIAN, X. L., WANG, Y. S., LIN, R. M., MAO, Z. C., CHEN, N. S. & XIE, B. Y. 2013.
Metagenomic analysis of the pinewood nematode microbiome reveals a symbiotic relationship critical
for xenobiotics degradation. Scientific Reports, 3.
CHEONG, T. K. & ORIEL, P. J. 2000. Cloning and expression of the limonene hydroxylase of Bacillus
stearothermophilus BR388 and utilization in two-phase limonene conversions. Applied Biochemistry
and Biotechnology, 84-6, 903-915.
DHAVALIKAR, R. & BHATTACHARYYA, P. 1966. Microbiological transformation of terpenes. 8.
Fermentation of limonene by a soil Pseudomonad. Indian Journal of Biochemistry, 3, 144-157.
DHAVALIKAR, R., RANGACHARI, P. & BHATTACHARYYA, P. 1966. Microbiological
transformations of terpenes. 9. Pathways of degradation of limonene in a soil Pseudomonad. Indian
Journal of Biochemistry, 3, 158-164.
- 45 -
Chapter II - Microbial monoterpene transformations - a review
DÍAZ-PÉREZ, A. L., ZAVALA-HERNANDEZ, A. N., CERVANTES, C. & CAMPOS-GARCIA, J.
2004. The gnyRDBHAL cluster is involved in acyclic isoprenoid degradation in Pseudomonas
aeruginosa. Applied and Environmental Microbiology, 70, 5102-5110.
DIGUISTINI, S., WANG, Y., LIAO, N. Y., TAYLOR, G., TANGUAY, P., FEAU, N., HENRISSAT, B.,
CHAN, S. K., HESSE-ORCE, U., ALAMOUTI, S. M., TSUI, C. K. M., DOCKING, R. T.,
LEVASSEUR, A., HARIDAS, S., ROBERTSON, G., BIROL, I., HOLT, R. A., MARRA, M. A.,
HAMELIN, R. C., HIRST, M., JONES, S. J. M., BOHLMANN, J. & BREUIL, C. 2011. Genome and
transcriptome analyses of the mountain pine beetle-fungal symbiont Grosmannia clavigera, a
lodgepole pine pathogen. Proceedings of the National Academy of Sciences of the United States of
America, 108, 2504-2509.
DUDAREVA, N., KLEMPIEN, A., MUHLEMANN, J. K. & KAPLAN, I. 2013. Biosynthesis, function
and metabolic engineering of plant volatile organic compounds. New Phytologist, 198, 16-32.
DUETZ, W. A., BOUWMEESTER, H., VAN BEILEN, J. B. & WITHOLT, B. 2003. Biotransformation
of limonene by bacteria, fungi, yeasts, and plants. Applied Microbiology and Biotechnology, 61, 269277.
DUETZ, W. A., FJALLMAN, A. H. M., REN, S. Y., JOURDAT, C. & WITHOLT, B. 2001.
Biotransformation of D-limonene to (+) trans-carveol by toluene-grown Rhodococcus opacus PWD4
cells. Applied and Environmental Microbiology, 67, 2829-2832.
EATON, R. W. 1996. p-Cumate catabolic pathway in Pseudomonas putida F1: Cloning and
characterization of DNA carrying the cmt operon. Journal of Bacteriology, 178, 1351-1362.
EATON, R. W. 1997. p-Cymene catabolic pathway in Pseudomonas putida F1: Cloning and
characterization of DNA encoding conversion of p-cymene to p-cumate. Journal of Bacteriology, 179,
3171-3180.
ESMAEILI, A. & HASHEMI, E. 2011. Biotransformation of myrcene by Pseudomonas aeruginosa.
Chemistry Central Journal, 5.
FALL, R. R. & HECTOR, M. L. 1977. Acyl-coenzyme-a carboxylases. Homologous 3-methylcrotonylcoA and geranyl-coA carboxylases from Pseudomonas citronellolis. Biochemistry, 16, 4000-4005.
FAROOQ, A., ATTA-UR-RAHMAN & CHOUDHARY, A. I. 2004. Fungal transformation of
monoterpenes. Current Organic Chemistry, 8, 353-367.
FONTANILLE, P., LE FLÈCHE, A. & LARROCHE, C. 2002. Pseudomonas rhodesiae PF1: A new and
efficient biocatalyst for production of isonovalal from D-pinene oxide. Biocatalysis and
Biotransformation, 20, 413-421.
FÖRSTER-FROMME, K., HÖSCHLE, B., MACK, C., BOTT, M., ARMBRUSTER, W. &
JENDROSSEK, D. 2006. Identification of genes and proteins necessary for catabolism of acyclic
- 46 -
Chapter II - Microbial monoterpene transformations - a review
terpenes and leucine/isovalerate in Pseudomonas aeruginosa. Applied and Environmental
Microbiology, 72, 4819-4828.
FÖRSTER-FROMME, K. & JENDROSSEK, D. 2010. Catabolism of citronellol and related acyclic
terpenoids in pseudomonads. Applied Microbiology and Biotechnology, 87, 859-869.
FOSS, S. & HARDER, J. 1997. Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. FEMS Microbiology Letters, 149, 71-75.
FOSS, S. & HARDER, J. 1998. Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on
oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) and nitrate. Systematic and
Applied Microbiology, 21, 365-373.
FOSS, S., HEYEN, U. & HARDER, J. 1998. Alcaligenes defragrans sp. nov., description of four strains
isolated on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alpha-phellandrene) and
nitrate. Systematic and Applied Microbiology, 21, 237-244.
FU, P. Q., KAWAMURA, K., CHEN, J. & BARRIE, L. A. 2009. Isoprene, monoterpene, and
sesquiterpene oxidation products in the high arctic aerosols during late winter to early summer.
Environmental Science & Technology, 43, 4022-4028.
GANTT, B., MESKHIDZE, N. & KAMYKOWSKI, D. 2009. A new physically-based quantification of
marine isoprene and primary organic aerosol emissions. Atmospheric Chemistry and Physics, 9, 49154927.
GERON, C., RASMUSSEN, R., ARNTS, R. R. & GUENTHER, A. 2000. A review and synthesis of
monoterpene speciation from forests in the United States. Atmospheric Environment, 34, 1761-1781.
GHIRARDO, A., GUTKNECHT, J., ZIMMER, I., BRUGGEMANN, N. & SCHNITZLER, J. P. 2011.
Biogenic volatile organic compound and respiratory CO2 emissions after 13C-labeling: Online tracing
of C translocation dynamics in poplar plants. PLoS ONE, 6.
GRIFFITHS, E. T., HARRIES, P. C., JEFFCOAT, R. & TRUDGILL, P. W. 1987. Purification and
properties of alpha-pinene oxide lyase from Nocardia sp. strain P18.3. Journal of Bacteriology, 169,
4980-4983.
GUENTHER, A., HEWITT, C. N., ERICKSON, D., FALL, R., GERON, C., GRAEDEL, T., HARLEY,
P., KLINGER, L., LERDAU, M., MCKAY, W. A., PIERCE, T., SCHOLES, B., STEINBRECHER,
R., TALLAMRAJU, R., TAYLOR, J. & ZIMMERMAN, P. 1995. A global model of natural volatile
organic compound emissions. Journal of Geophysical Research-Atmospheres, 100, 8873-8892.
HARAYAMA, S., REKIK, M., NGAI, K. L. & ORNSTON, L. N. 1989. Physically associated enzymes
produce and metabolize 2-hydroxy-2,4-dienoate, a chemically unstable intermediate formed in catechol
metabolism via meta-cleavage in Pseudomonas putida. Journal of Bacteriology, 171, 6251-6258.
HARDER, J. & FOSS, S. 1999. Anaerobic formation of the aromatic hydrocarbon p-cymene from
monoterpenes by methanogenic enrichment cultures. Geomicrobiology Journal, 16, 295-305.
- 47 -
Chapter II - Microbial monoterpene transformations - a review
HARDER, J., HEYEN, U., PROBIAN, C. & FOSS, S. 2000. Anaerobic utilization of essential oils by
denitrifying bacteria. Biodegradation, 11, 55-63.
HARDER, J. & PROBIAN, C. 1995. Microbial degradation of monoterpenes in the absence of molecular
oxygen. Applied and Environmental Microbiology, 61, 3804-3808.
HAWKES, D. B., ADAMS, G. W., BURLINGAME, A. L., DE MONTELLANO, P. R. O. & DE VOSS,
J. J. 2002. Cytochrome P450cin (CYP176A), isolation, expression, and characterization. Journal of
Biological Chemistry, 277, 27725-27732.
HEYEN, U. & HARDER, J. 2000. Geranic acid formation, an initial reaction of anaerobic monoterpene
metabolism in denitrifying Alcaligenes defragrans. Applied and Environmental Microbiology, 66,
3004-3009.
HÖSCHLE, B., GNAU, V. & JENDROSSEK, D. 2005. Methylcrotonyl-CoA and geranyl-CoA
carboxylases are involved in leucine/isovalerate utilization (Liu) and acyclic terpene utilization (Atu),
and are encoded by liuB/liuD and atuC/atuF, in Pseudomonas aeruginosa. Microbiology-Sgm, 151,
3649-3656.
HÖSCHLE, B. & JENDROSSEK, D. 2005. Utilization of geraniol is dependent on molybdenum in
Pseudomonas aeruginosa: evidence for different metabolic routes for oxidation of geraniol and
citronellol. Microbiology-Sgm, 151, 2277-2283.
HU, Q. H., XIE, Z. Q., WANG, X. M., KANG, H., HE, Q. F. & ZHANG, P. F. 2013. Secondary organic
aerosols over oceans via oxidation of isoprene and monoterpenes from Arctic to Antarctic. Scientific
Reports, 3.
HYLEMON, P. B. & HARDER, J. 1998. Biotransformation of monoterpenes, bile acids, and other
isoprenoids in anaerobic ecosystems. FEMS Microbiology Reviews, 22, 475-488.
INSAM, H. & SEEWALD, M. S. A. 2010. Volatile organic compounds (VOCs) in soils. Biology and
Fertility of Soils, 46, 199-213.
ISHIDA, T. 2005. Biotransformation of terpenoids by mammals, microorganisms, and plant-cultured
cells. Chemistry & Biodiversity, 2, 569-590.
IURESCIA, S., MARCONI, A. M., TOFANI, D., GAMBACORTA, A., PATERNO, A., DEVIRGILIIS,
C., VAN DER WERF, M. J. & ZENNARO, E. 1999. Identification and sequencing of beta-myrcene
catabolism genes from Pseudomonas sp. strain M1. Applied and Environmental Microbiology, 65,
2871-2876.
IWAKI, H., GROSSE, S., BERGERON, H., LEISCH, H., MORLEY, K., HASEGAWA, Y. & LAU, P. C.
K. 2013. Camphor pathway redux: Functional recombinant expression of 2,5- and 3,6-diketocamphane
monooxygenases of Pseudomonas putida ATCC 17453 with their cognate flavin reductase catalyzing
Baeyer-Villiger reactions. Applied and Environmental Microbiology, 79, 3282-3293.
- 48 -
Chapter II - Microbial monoterpene transformations - a review
KADOW, M., LOSCHINSKI, K., SASS, S., SCHMIDT, M. & BORNSCHEUER, U. T. 2012.
Completing the series of BVMOs involved in camphor metabolism of Pseudomonas putida NCIMB
10007 by identification of the two missing genes, their functional expression in E. coli, and
biochemical characterization. Applied Microbiology and Biotechnology, 96, 419-429.
KAINULAINEN, P. & HOLOPAINEN, J. K. 2002. Concentrations of secondary compounds in Scots
pine needles at different stages of decomposition. Soil Biology & Biochemistry, 34, 37-42.
KESSELMEIER, J. & STAUDT, M. 1999. Biogenic volatile organic compounds (VOC): An overview on
emission, physiology and ecology. Journal of Atmospheric Chemistry, 33, 23-88.
LAH, L., HARIDAS, S., BOHLMANN, J. & BREUIL, C. 2013. The cytochromes P450 of Grosmannia
clavigera: Genome organization, phylogeny, and expression in response to pine host chemicals. Fungal
Genetics and Biology, 50, 72-81.
LEISCH, H., SHI, R., GROSSE, S., MORLEY, K., BERGERON, H., CYGLER, M., IWAKI, H.,
HASEGAWA, Y. & LAU, P. C. K. 2012. Cloning, Baeyer-Villiger biooxidations, and structures of the
camphor pathway 2-oxo-'3-4,5,5-trimethylcyclopentenylacetyl-coenzyme A monooxygenase of
Pseudomonas putida ATCC 17453. Applied and Environmental Microbiology, 78, 2200-2212.
LI, H. J. & LAN, W. J. 2011. Biotransformation of limonene by microorganisms. Progress in Chemistry,
23, 2318-2325.
LI, H. J., LAN, W. J., CAI, C. H., ZHOU, Y. P. & LIN, Y. C. 2006. Biotransformation of limonene by
marine bacteria. Chinese Journal of Analytical Chemistry, 34, 946-950.
LINARES, D., FONTANILLE, P. & LARROCHE, C. 2009. Exploration of D-pinene degradation
pathway of Pseudomonas rhodesiae CIP 107491. Application to novalic acid production in a
bioreactor. Food Research International, 42, 461-469.
LIU, W. G. & ROSAZZA, J. P. N. 1990. Stereospecific hydroxylation of 1,8-cineole using a microbial
biocatalyst. Tetrahedron Letters, 31, 2833-2836.
LIU, W. G. & ROSAZZA, J. P. N. 1993. A soluble Bacillus cereus cytochrome-P-450cin system catalyzes
1,4-cineole hydroxylations. Applied and Environmental Microbiology, 59, 3889-3893.
LUEDDEKE, F. & HARDER, J. 2011. Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift Fur Naturforschung Section C-a Journal of Biosciences, 66,
409-412.
LUEDDEKE, F., WUELFING, A., TIMKE, M., GERMER, F., WEBER, J., DIKFIDAN, A.,
RAHNFELD, T., LINDER, D., MEYERDIERKS, A. & HARDER, J. 2012. Geraniol and geranial
dehydrogenases induced in anaerobic monoterpene degradation by Castellaniella defragrans. Applied
and Environmental Microbiology, 78, 2128-2136.
LUO, G. & YU, F. 2010. A numerical evaluation of global oceanic emissions of D-pinene and isoprene.
Atmospheric Chemistry and Physics, 10, 2007-2015.
- 49 -
Chapter II - Microbial monoterpene transformations - a review
MAHMOUD, S. S. & CROTEAU, R. B. 2002. Strategies for transgenic manipulation of monoterpene
biosynthesis in plants. Trends in Plant Science, 7, 366-373.
MARS, A. E., GORISSEN, J. P. L., VAN DEN BELD, I. & EGGINK, G. 2001. Bioconversion of
limonene to increased concentrations of perillic acid by Pseudomonas putida GS1 in a fed-batch
reactor. Applied Microbiology and Biotechnology, 56, 101-107.
MARTINEZ, J. L., SÁNCHEZ, M. B., MARTÍNEZ-SOLANO, L., HERNANDEZ, A., GARMENDIA,
L., FAJARDO, A. & ALVAREZ-ORTEGA, C. 2009. Functional role of bacterial multidrug efflux
pumps in microbial natural ecosystems. FEMS Microbiology Reviews, 33, 430-449.
OUGHAM, H. J., TAYLOR, D. G. & TRUDGILL, P. W. 1983. Camphor revisited - Involvement of a
unique monooxygenase in metabolism of 2-oxo-'3-4,5,5-trimethylcyclopentenylacetic acid by
Pseudomonas putida. Journal of Bacteriology, 153, 140-152.
PALMER, P. I. & SHAW, S. L. 2005. Quantifying global marine isoprene fluxes using MODIS
chlorophyll observations. Geophysical Research Letters, 32.
PAPADOPOULOS, C. J., CARSON, C. F., CHANG, B. J. & RILEY, T. V. 2008. Role of the MexABOprM efflux pump of Pseudomonas aeruginosa in tolerance to tea tree (Melaleuca altenifolia) oil and
its monoterpene components terpinen-4-ol, 1,8-cineole, and alpha-terpineol.
Applied and
Environmental Microbiology, 74, 1932-1935.
PARK, Y. J., KIM, I. C. & CHANG, H. C. 2003. Microbial conversion of (+)-limonene by an
Enterobacter agglomerans isolate. Journal of Microbiology and Biotechnology, 13, 636-639.
PETASCH, J., DISCH, E.-M., MARKERT, S., BECHER, D., SCHWEDER, T., HÜTTEL, B.,
REINHARDT, R. & HARDER, J. 2014. The oxygen-independent metabolism of cyclic monoterpenes
in Castellaniella defragrans 65Phen. BMC Microbiology, 14, 164.
PLATT, A., SHINGLER, V., TAYLOR, S. C. & WILLIAMS, P. A. 1995. The 4-hydroxy-2-oxovalerate
aldolase and acetaldehyde dehydrogenase (acylating) encoded by the nahM and nahO genes of the
naphthalene catabolic plasmid pWW60-22 provide further evidence of conservation of meta-cleavage
pathway gene sequences. Microbiology-Uk, 141, 2223-2233.
POULOS, T. L., FINZEL, B. C., GUNSALUS, I. C., WAGNER, G. C. & KRAUT, J. 1985. The 2.6-A
crystal structure of Pseudomonas putida cytochrome-P-450. Journal of Biological Chemistry, 260,
6122-6130.
PRAKASH, O., KUMARI, K. & LAL, R. 2007. Pseudomonas delhiensis sp. nov., from a fly ash dumping
site of a thermal power plant. International Journal of Systematic and Evolutionary Microbiology, 57,
527-531.
RASMUSSEN, J. A. M., HENDERSON, K. A., STRAFFON, M. J., DUMSDAY, G. J., COULTON, J. &
ZACHARIOU, M. 2005. Two new biocatalysts for improved biological oxidation of 1,8-cineole.
Australian Journal of Chemistry, 58, 912-916.
- 50 -
Chapter II - Microbial monoterpene transformations - a review
SAVITHIRY, N., GAGE, D., FU, W. J. & ORIEL, P. 1998. Degradation of pinene by Bacillus pallidus
BR425. Biodegradation, 9, 337-341.
SCHEWE, H., MIRATA, M. A., HOLTMANN, D. & SCHRADER, J. 2011. Biooxidation of
monoterpenes with bacterial monooxygenases. Process Biochemistry, 46, 1885-1899.
SEUBERT, W. 1960. Degradation of isoprenoid compounds by microorganisms. I. Isolation and
characterization of an isoprenoid-degrading bacterium, Pseudomonas citronellolis n. sp. Journal of
Bacteriology, 79, 426-434.
SEUBERT, W. & FASS, E. 1964. Untersuchungen über den bakteriellen Abbau von Isoprenoiden. 5. Der
Mechanismus des Isoprenabbaus. Biochemische Zeitschrift, 341, 35-44.
SEUBERT, W., FASS, E. & REMBERGER.U 1963. Untersuchungen über den bakteriellen Abbau von
Isoprenoiden. 3. Reinigung und Eigenschaften der Geranylcarboxylase. Biochemische Zeitschrift, 338,
265-275.
SEUBERT, W. & REMBERGER, U. 1963. Untersuchungen über den bakteriellen Abbau von
Isoprenoiden. 2. Die Rolle der Kohlensäure. Biochemische Zeitschrift, 338, 245-264.
SHARKEY, T. D., WIBERLEY, A. E. & DONOHUE, A. R. 2008. Isoprene emission from plants: Why
and how. Annals of Botany, 101, 5-18.
SHARKEY, T. D. & YEH, S. S. 2001. Isoprene emission from plants. Annual Review of Plant Physiology
and Plant Molecular Biology, 52, 407-436.
SHAW, S. L., GANTT, B. & MESKHIDZE, N. 2010. Production and Emissions of Marine Isoprene and
Monoterpenes: A Review. Advances in Meteorology.
SPEELMANS, G., BIJLSMA, A. & EGGINK, G. 1998. Limonene bioconversion to high concentrations
of a single and stable product, perillic acid, by a solvent-resistant Pseudomonas putida strain. Applied
Microbiology and Biotechnology, 50, 538-544.
STOTZKY, G. & SCHENCK, S. 1976. Volatile organic compounds and microorganisms. Critical
Reviews in Microbiology, 4, 333-382.
TANAKA, K., KIM, H. J., SAITO, K., TAKAHASHI, H. G., WATANABE, M., YOKOHATA, T.,
KIMOTO, M., TAKATA, K. & YASUNARI, T. 2012. How have both cultivation and warming
influenced annual global isoprene and monoterpene emissions since the preindustrial era? Atmospheric
Chemistry and Physics, 12, 9703-9718.
TAYLOR, D. G. & TRUDGILL, P. W. 1986. Camphor revisited: studies of 2,5-diketocamphane 1,2monooxygenase from Pseudomonas putida ATCC 17453. Journal of Bacteriology, 165, 489-497.
THOMPSON, M. L., MARRIOTT, R., DOWLE, A. & GROGAN, G. 2010. Biotransformation of betamyrcene to geraniol by a strain of Rhodococcus erythropolis isolated by selective enrichment from hop
plants. Applied Microbiology and Biotechnology, 85, 721-730.
- 51 -
Chapter II - Microbial monoterpene transformations - a review
TONG, W.-Y. 2013. Biotransformation of Terpenoids and Steroids. In: RAMAWAT, K. G. &
MÉRILLON, J.-M. (eds.) Natural Products. Springer Berlin Heidelberg.
TRUDGILL, P. W. 1990. Microbial metabolism of monoterpenes - recent developments. Biodegradation,
1, 93-105.
TRUDGILL, P. W. 1994. Microbial metabolism and transformation of selected monoterpenes, Kluwer
Academic Publishers, PO Box 989, 3300 AZ Dordrecht, Netherlands 101 Phillip Drive, Norwell,
Massachusetts 02061, USA.
ULLAH, A. J. H., MURRAY, R. I., BHATTACHARYYA, P. K., WAGNER, G. C. & GUNSALUS, I. C.
1990. Protein components of a cytochrome P-450 linalool 8-methyl hydroxylase. Journal of Biological
Chemistry, 265, 1345-1351.
VAN DER WERF, M., DE BONT, J. A. M. & LEAK, D. J. 1997. Opportunities in microbial
biotransformation of monoterpenes. In: BERGER, R. G. (ed.) Advances in Biochemical Engineering
Biotechnology; Biotechnology of aroma compounds. Springer-Verlag, Heidelberger Platz 3, D-1000
Berlin, Germany 175 Fifth Avenue, New York, New York 10010, USA.
VAN DER WERF, M. J. & BOOT, A. M. 2000. Metabolism of carveol and dihydrocarveol in
Rhodococcus erythropolis DCL14. Microbiology-Uk, 146, 1129-1141.
VAN DER WERF, M. J., ORRU, R. V. A., OVERKAMP, K. M., SWARTS, H. J., OSPRIAN, I.,
STEINREIBER, A., DE BONT, J. A. M. & FABER, K. 1999a. Substrate specificity and
stereospecificity of limonene-1,2-epoxide hydrolase from Rhodococcus erythropolis DCL14; an
enzyme showing sequential and enantioconvergent substrate conversion. Applied Microbiology and
Biotechnology, 52, 380-385.
VAN DER WERF, M. J., SWARTS, H. J. & DE BONT, J. A. M. 1999b. Rhodococcus erythropolis
DCL14 contains a novel degradation pathway for limonene. Applied and Environmental Microbiology,
65, 2092-2102.
WANG, Y., LIM, L., DIGUISTINI, S., ROBERTSON, G., BOHLMANN, J. & BREUIL, C. 2013. A
specialized ABC efflux transporter GcABC-G1 confers monoterpene resistance to Grosmannia
clavigera, a bark beetle-associated fungal pathogen of pine trees. New Phytologist, 197, 886-898.
WILLIAMS, D. R., TRUDGILL, P. W. & TAYLOR, D. G. 1989. Metabolism of 1,8-cineole by a
Rhodococcus species: Ring cleavage reactions. Journal of General Microbiology, 135, 1957-1967.
WILT, F. M., MILLER, G. C., EVERETT, R. L. & HACKETT, M. 1993. Monoterpene concentrations in
fresh, senescent, and decaying foliage of single-leaf pinyon (Pinus monophylla Torr and Frem:
Pinaceae) from the Western Great-Basin. Journal of Chemical Ecology, 19, 185-194.
WRIGHT, S. J., CAUNT, P., CARTER, D. & BAKER, P. B. 1986. Microbial oxidation of D-pinene by
Serratia marcescens. Applied Microbiology and Biotechnology, 23, 224-227.
- 52 -
Chapter II - Microbial monoterpene transformations - a review
YANG, J. E., PARK, Y. J. & CHANG, H. C. 2007. Cloning of four genes involved in limonene
hydroxylation from Enterobacter cowanii 6L. Journal of Microbiology and Biotechnology, 17, 11691176.
YASSAA, N., PEEKEN, I., ZÖLLNER, E., BLUHM, K., ARNOLD, S., SPRACKLEN, D. &
WILLIAMS, J. 2008. Evidence for marine production of monoterpenes. Environmental Chemistry, 5,
391-401.
YOO, S. K. & DAY, D. F. 2002. Bacterial metabolism of alpha- and beta-pinene and related
monoterpenes by Pseudomonas sp. strain PIN. Process Biochemistry, 37, 739-745.
YU, F. N. A. & UTSUMI, R. 2009. Diversity, regulation, and genetic manipulation of plant mono- and
sesquiterpenoid biosynthesis. Cellular and Molecular Life Sciences, 66, 3043-3052.
ZIEMANN, P. J. & ATKINSON, R. 2012. Kinetics, products, and mechanisms of secondary organic
aerosol formation. Chemical Society Reviews, 41, 6582-6605.
- 53 -
Aim of the work
Aim of the work
Castellaniella defragrans 65Phen initiates the degradation of E-myrcene by hydration to the
tertiary alcohol (S)-linalool, which is further isomerized to the primary alcohol geraniol by the
bifunctional enzyme linalool dehydratase/isomerase (Ldi). No similar enzymes are reported in
databases and the catalyzed reactions are rather unusual. An aim of the presented work was the
structural analysis of the linalool dehydratase/isomerase by X-ray crystallography and the
elucidation of the catalytic mechanism of this enzyme. In order to produce sufficient amounts of
protein, a purification protocol based on immobilized-metal affinity chromatography and a polyhistidine-tag was attempted. As recovery of native and active Ldi by this purification approach
cannot be assured, an alternative purification protocol for the wildtype enzyme based on classical
column chromatography with a focus on high yields was also desired.
In another approach, the function of selected amino acids in the Ldi was investigated by amino
acid exchange experiments. Cysteine residues play an important role in proteins. They contribute
to the tertiary structure by formation of disulfide bonds or have catalytical properties when
present as free thiols (Hansen and Winther, 2009, Marino and Gladyshev, 2010, Marino and
Gladyshev, 2012). Therefore, the cysteine residues were selected as first targets. Other amino
acids, possibly involved in enzyme catalysis of the Ldi, were selected based on an “educatedguess approach”. The absence of cofactors in the Ldi suggests acid-base catalysis. Amino acids
known for their capacity to donate or to subtract a proton (e.g. histidine) were selected (Holliday
et al., 2009). Additionally, hydrophobic amino acids were targeted as they may be involved in
stabilization and coordination of the hydrophobic substrates E-myrcene, (S)-linalool and geraniol.
A knock-out mutant of C. defragrans 65Phen, deficient in the ldi gene, was unable to grow on Emyrcene and geraniol but still grew on mono- and bicyclic monoterpenes and on (R,S)-linalool
(Lüddeke et al., 2012a). Recently, a degradation pathway of limonene has been elucidated
(Petasch et al., 2014) and might apply also to other monocyclic monoterpenes. However, the
metabolic fate of linalool in C. defragrans 65Phen 'ldi remained unresolved and was
investigated by physiological and enzymatic approaches. The results will be discussed in the
presented work.
- 54 -
Aim of the work
The formation of geraniol from linalool has been observed in cultures of Thauera linaloolentis
47Lol and a 3,1-hydroxyl-'1-'2-mutase was suggested as novel enzyme activity. Additionally,
the oxidation of geraniol to geranial has been reported (Foss and Harder, 1998, Foss and Harder,
1997). The isolation and characterization of the 3,1-hydroxyl-'1-'2-mutase (linalool isomerase)
activity was attempted during this project. The oxidation of geraniol to geranial suggested a
further degradation in analogy to the Atu pathway, known from Pseudomonas spp. (FörsterFromme and Jendrossek, 2010). Two draft genomes of T. linaloolentis 47Lol became publicly
available during the initial phase of the project. A second aim of this project was to investigate
the fate of geraniol by cultivation based experiment as well as on proteomic and genomic level.
- 55 -
Aim of the work
References
FÖRSTER-FROMME, K. & JENDROSSEK, D. 2010. Catabolism of citronellol and related acyclic
terpenoids in pseudomonads. Applied Microbiology and Biotechnology, 87, 859-869.
FOSS, S. & HARDER, J. 1997. Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. Fems Microbiology Letters, 149, 7175.
FOSS, S. & HARDER, J. 1998. Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on
oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) and nitrate. Systematic and
Applied Microbiology, 21, 365-373.
HANSEN, R. E. & WINTHER, J. R. 2009. An introduction to methods for analyzing thiols and disulfides:
Reactions, reagents, and practical considerations. Analytical Biochemistry, 394, 147-158.
HOLLIDAY, G. L., MITCHELL, J. B. O. & THORNTON, J. M. 2009. Understanding the functional roles
of amino acid residues in enzyme catalysis. Journal of Molecular Biology, 390, 560-577.
LÜDDEKE, F., DIKFIDAN, A. & HARDER, J. 2012. Physiology of deletion mutants in the anaerobic
beta-myrcene degradation pathway in Castellaniella defragrans. BMC Microbiology, 12.
MARINO, S. M. & GLADYSHEV, V. N. 2010. Cysteine function governs its conservation and
degeneration and restricts Its utilization on protein surfaces. Journal of Molecular Biology, 404,
902-916.
MARINO, S. M. & GLADYSHEV, V. N. 2012. Analysis and functional prediction of reactive cysteine
residues. Journal of Biological Chemistry, 287, 4419-4425.
PETASCH, J., DISCH, E.-M., MARKERT, S., BECHER, D., SCHWEDER, T., HÜTTEL, B.,
REINHARDT, R. & HARDER, J. 2014. The oxygen-independent metabolism of cyclic
monoterpenes in Castellaniella defragrans 65Phen. BMC Microbiology, 14, 164.
- 56 -
Aim of the work
- 57 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Chapter III
A novel purification protocol for the linalool dehydratase/isomerase
Robert Marmulla1 and Jens Harder1*
1
Dept. of Microbiology, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359
Bremen
*
corresponding author: jharder@mpi-bremen.de
Manuscript in preparation
- 58 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Abstract
The linalool dehydratase/isomerase (Ldi) is the initial enzyme in the degradation of acyclic
monoterpenes in Castellaniella defragrans 65Phen. The bifunctional enzyme catalyzes the
reversible hydration of the acyclic monoterpene E-myrcene to (S)-linalool and its isomerization to
geraniol. The ldi wildtype gene encodes a periplasmic enzyme with a signal sequence for export.
In this study, we attempted the overexpression and purification of the mature protein in E. coli. In
one attempt, a His6-tag together with a small ubiquitin-like modifier (Sumo) domain was fused to
the N-terminus of the Ldi (his6-Sumo-tag; Sumo-Ldi) and successfully expressed in E. coli
BL21(DE3). Purification was hampered by an extremely strong binding to the nickel affinity
column and the purified enzyme showed only poor or no activity after removal of the his6-Sumopeptide. Alternatively, we developed a novel purification protocol for the wildtype Ldi
heterologously expressed in E. coli BL21(DE3). The purification of the wildtype protein was
simpler than the purification of the His6-tagged protein.
Keywords: Linalool dehydratase/isomerase, linalool, geraniol, myrcene, Castellaniella
defragrans 65Phen, his6-Sumo
Introduction
The linalool dehydratase/isomerase (Ldi, EC 4.2.1.127 and EC 5.4.4.4) is a bifunctional enzyme
which catalyzes the reversible hydration of E-myrcene to (S)-linalool and its further isomerization
to geraniol (Fig. 1). The enzyme, which had been identified in the betaproteobacterium
Castellaniella defragrans 65Phen, enables growth on the monoterpene hydrocarbon E-myrcene
(Foss et al., 1998, Heyen and Harder, 2000). Linalool dehydratase/isomerase had been purified to
homogeneity by chromatography and its gene sequence was disclosed (Brodkorb et al., 2010).
Based on SDS-PAGE, the mature enzyme had a molecular weight of around 40 kDa. A
homotetrameric complex was predicted as the native state from size-exclusion chromatography
experiments. The ldi gene has been cloned into the overexpression vector pET42a(+) and the
enzyme successfully expressed in E. coli BL21(DE3)Star (Brodkorb et al., 2010). The expressed
Ldi features a N-terminal signal peptide for periplasmatic localization via the Sec-pathway,
- 59 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
which is expected to be cleaved off during maturation after the transport over the inner
membrane. The mature enzyme consists of 371 amino acids with a molecular weight of 41.8 kDa,
a calculated isoelectric point at pH 6.02 and an overall hydrophobicity of -0.198 (GRAVY, Kyte
and Doolittle, 1982). However, the enzymatic mechanism is stereoselective for the (S)-linalool,
but unknown in its details (Lüddeke and Harder, 2011).
Figure 1: Reaction sequence catalyzed by the linalool dehydratase/isomerase from C. defragrans 65Phen.
E-myrcene is hydrated to (S)-linalool and further isomerized to geraniol.
The first reported purification protocol for the wildtype enzyme included an anion exchange
chromatography (SourceQ), followed by hydrophobic interaction chromatography on a butylsepharose FF column, an ammonium sulfate precipitation, size-exclusion chromatography and a
second anion exchanger (ResourceQ) (Brodkorb et al., 2010). We intended to improve this
procedure by applying a faster purification method with an N-terminal addition of a purification
tag to the protein. Over the last decades, novel purification systems were developed based on
specific protein interactions to a solid support, called affinity chromatography. Specific binding is
achieved by addition of so called purification tags to the protein of interest, either N- or Cterminally fused. The simplest tag is a multiple repetition of histidine residues. Other tags
resemble substrate binding domains of enzymes (e.g. MBP - maltose binding protein, GST glutathione S-transferase binding site, CBP - calmodulin binding protein) (Walls and Loughran,
2011, Young et al., 2012). Histidine residues are known to form stable complexes with divalent
metal ions (e.g. Ni2+, Co2+, Cu2+) on the surface of the column material; a method known as
immobilized-metal affinity chromatography (IMAC) (Porath et al., 1975, Charlton and
Zachariou, 2008). Bound protein is eluted from the column by the replacement mechanism.
Increasing concentrations of histidine or imidazole are used in IMAC, while for substrate binding
domains increasing concentrations of their natural substrate or of analogues are used. Purification
tags may interfere with downstream processing or enzyme activity and removal of the added
domain is often desired. Protease cleavage sites are introduced between tag and target protein for
- 60 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
this purpose (Arnau et al., 2006, Charlton and Zachariou, 2011). A rather novel purification tag
consisting of a His6-tag and the small ubiquitin-like modifier (Sumo) was developed and
successfully applied to different proteins from eukaryotes and prokaryotes like the SARS virus
protein (Zuo et al., 2005b), human epidermal growth factor (Su et al., 2006), bone morphogenetic
protein 14 (Li et al., 2011) and others (Peroutka et al., 2011). Sumo peptide was shown to
enhance solubility and decrease proteolytic degradation of the heterologously expressed protein
(Butt et al., 2005 and references therein). Even short peptides, which are under normal conditions
easily prone to degradation, were efficiently expressed in E. coli by using a His6-Sumo-tag fusion
system (Malakhov et al., 2004, Marblestone et al., 2006, Satakarni and Curtis, 2011). The Sumo
domain is removed by specific Sumo proteases (Li and Hochstrasser, 1999). In contrast to other
proteases, Sumo proteases recognize not only their consensus sequence but also the tertiary
structure of the Sumo peptide. Cleavage occurs after a conserved Gly-Gly pair without leaving
extra non-native amino acids attached on the target protein (Li and Hochstrasser, 1999, Hay,
2007, Mukhopadhyay and Dasso, 2007). The Sumo protease (Ulp1) from yeast has been studied
with respect on compatibility towards temperature and chemicals. It has been shown to be active
over a wide temperature range (4 - 22°C) as well as under reduced and oxidized conditions and in
the presence of urea or ethanol (Malakhov et al., 2004). The beneficial characteristics of the
Sumo-system caused its selection for a high-yield purification of the Ldi for future studies. The
Sumo peptide may also increase the solubility and stability of the heterologously expressed
fusion protein, while the His6-tag was necessary for facilitating Ldi purification by the IMAC
principle.
- 61 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Methods
Construction of expression vector
The ldi gene was amplified by PCR from the expression vector pET42-Ldi (Brodkorb et al.,
2010), using the Phusion Polymerase (Life Technologies, Thermo Fisher Scientific, Waltham,
USA) with a newly designed primer pair (FW: GCGGAACTGCCGCCGGGG and RV:
TTATTTCCCTGCGAGCTTGGCC, annealing temperature 68°C). The primer pair targeted the
coding sequence of the mature Ldi, lacking the leader peptide for periplasmatic localization. The
amplicon was cloned into the pET-Sumo vector and the plasmid was transformed into E. coli
BL21(DE3) according to manufacturer guidelines (Champion™ pET Sumo Expression System,
Life Technologies, Thermo Fisher Scientific). In a second approach, an overexpression plasmid
for the mature Ldi without the leader sequence was constructed using a new primer pair, targeting
position 79-1194 of the ldi gene, introducing an artificial start codon and the restriction enzyme
sites NdeI and BglII (FW: TGTGACATATGATGGCGGAACTGCCG and RV: CGCGAGATCT
TTATTTCCCTGCGA, annealing temperature 60°C). The amplicon was cloned into the pET-42a
vector by restriction digest with NdeI and BglII (Invitrogen, Thermo Fisher Scientific) according
to manufacturer guidelines and transformed into E. coli BL21(DE3).
Cultivation of E. coli BL21(DE3) and expression of Ldi
E. coli BL21(DE3) was cultivated in rich medium (yeast extract 12 g L-1, peptone 14 g L-1, NaCl
5 g L-1, MgSO4 x 7 H2O 0.25 g L-1, glycerol 2 g L-1, kanamycin 50 μg mL-1) at 37°C in 500 mL
batches in 1000 mL Erlenmeyer flasks at 165 rpm. Ldi expression was induced by 1 mM
Isopropyl β-D-1-thiogalactopyranoside at an optical density (600 nm) around 1. Cultures were
further cultivated for 4 to 5 hours at 32°C to an OD (600 nm) of approx. 3. Biomass was
harvested by centrifugation (14000 x g, 20 min, 4°C), frozen in liquid nitrogen and stored
at -80°C.
Preparation of soluble protein extract
One volume E. coli BL21(DE3) biomass was suspended in three volumes appropriate buffer
(Sumo-Ldi: 80 mM Tris-Cl pH 8.0 with 500 mM NaCl and 50 mM imidazole; wildtype Ldi: 80
mM Tris-Cl pH 8.0) and thawed quickly at room temperature. Cell disintegration was performed
- 62 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
by mechanical sheering using a French Press pressure cell at 8.26 MPa two times. The crude
extract was clarified by ultracentrifugation (150000 x g, 30 min, 4°C) and the supernatant
(soluble protein extract) was used in further experiments.
Activity assay
Enzymatic activity was determined by end-point measurement. Subfractions obtained during
purification were dialyzed against Tris-Cl buffer (80 mM, pH 8.0) in a 1:500 sample:buffer ratio
for at least 12 hours. The assay was performed in 4 mL glass vials. 300-500 μL of sample were
reduced with 5 mM dithiothreitol (DTT). Vials were closed with a butyl rubber septum and
flushed with N2 gas for 3 minutes via needles. After a pre-incubation of 20 min at room
temperature, 200 μL substrate (200 mM (R,S)-linalool or geraniol in 2,2,4,4,6,8,8heptamethylnonane, HMN) was added via a needle. Incubation took place at 37°C and shaking at
70 rpm (Sartorius Certomat R incubator, Sartorius Stedim Biotech GmbH, Goettingen, Germany)
for 11 to 12 hours. Product formation was determined by gas chromatography with flame
ionization detection (PerkinElmer Auto System XL, Überlingen, Germany). 1 μL sample of
HMN phase was injected onto an Optima-5 column (30 m x 0.32 mm, 0.25μm film thickness;
Macherey-Nagel, Germany) with hydrogen as carrier gas and the following temperature program:
injection port 250°C, detection port 350°C, initial column temperature 40°C for 2 min, increasing
to 100°C at a rate of 4°C min-1, keeping 100°C for 0.1 min, followed by an increase to 320°C at
45°C min-1 and hold for 3 min. The split ratio was set to 1:9.
Sumo protease activity and cleavage conditions
The Sumo protease cleavage of Sumo-Ldi into Sumo-tag and Ldi was investigated in soluble
protein extracts of E. coli BL21(DE3) pET-Sumo-Ldi (9.7 mg mL-1 protein) under oxidized and
reduced conditions, without and with 20 mM DTT, in 80 mM Tris-Cl (pH 8.0).
Protein purification
All purification steps were performed on an ÄKTA purifier system (GE Healthcare Europe
GmbH, Freiburg, Germany) at 6°C. All buffers were prepared and pH values adjusted at room
temperature (22°C) and filtered (0.2 μm) prior use. Protein was detected by absorption
measurement at 280 nm. SDS-PAGE was used to analyze purification progress. Protein
- 63 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
concentrations were determined according to the method described by Bradford using bovine
serum albumin as a calibration standard (Bradford, 1976).
Purification of Sumo-Ldi
Soluble protein extract was prepared from cell pellets of E. coli BL21(DE3) pET-Sumo-Ldi in
binding buffer (80 mM Tris-Cl, 500 mM NaCl, 50 mM imidazole, pH 8.0). A Ni2+-IDA column
(dimensions: 11 x 80 mm; CV 30.5 mL; ProBond Nickel chelating resin, Novex by Life
Technologies, Thermo Fisher Scientific) was equilibrated with the same buffer. Soluble protein
fraction (500 mg at 25 mg mL-1) was applied onto the column at a flow rate of 0.5 mL/min.
Unbound protein was removed with the flow through. Elution of bound protein was achieved by
applying 500 mM imidazole in binding buffer. In a second step, 1 M imidazole in buffer was
applied to assure the elution of all protein. Eluted protein was pooled according to the absorption
peaks, concentrated on an AMICON filter unit (15 kDa cut-off; Merck Millipore, Billerica, MA,
USA) and loaded onto a desalting Sephadex G25 column (dimensions: 24 x 100 mm; CV 45.2
mL) equilibrated with Tris-Cl buffer (80 mM, 100 mM NaCl, 50 mM imidazole, pH 8.0) at 0.5
mL/min to lower the imidazole concentration. The Sumo-Ldi containing sample was treated with
Sumo protease (5 μg recombinant protein Unit protease-1 h-1) and concentrated prior the second
chromatography separation to remove Sumo protease and the Sumo-tag. The Ni2+-IDA column
was equilibrated with Tris-Cl buffer (80 mM, 100 mM NaCl, 20 mM imidazole, 0.1 % w/v
Tween 20, pH 8.0). Flow through was collected and bound protein was eluted in a single step
with 500 mM imidazole. The Ldi containing flow through was concentrated on an AMICON
filter unit and applied to size-exclusion chromatography (40 mM Tris-Cl, pH 8.0; HiLoad 16/60,
Superdex200, dimensions: 16 x 600 mm). Calibration was performed with thyroglobulin (670
kDa), bovine gamma-globulin (158 kDa), chicken ovalalbumin (44 kDa), equine myoglobin (17
kDa) and vitamin B12 (1.35 kDa).
An alternative purification under denaturating conditions was performed in the presence of 6 M
urea in the purification buffers. Other conditions remained the same as described above. Urea
was removed from the Sumo-Ldi containing eluate prior protease treatment by dialysis against
Tris-Cl buffer (80 mM, 100 mM NaCl, 50 mM imidazole, pH 8.0). Subsequent separation of
Sumo-tag and Ldi was performed as described above.
- 64 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Purification of wildtype Ldi
The wildtype Ldi was purified in a three step chromatography approach, including hydrophobic
interaction, anion exchange and size-exclusion chromatography. Soluble protein extract from E.
coli BL21(DE3) pET42-Ldi was prepared in Tris-Cl buffer (80 mM, pH 8.0). The sample was
adjusted to 5 % ammoniumsulfate (v/v) with a saturated ammoniumsulfate solution to aid binding
onto a Phenyl Sepharose column (dimensions: 8 x 100 mm; CV 5 mL). The column was
equilibrated with Tris-Cl buffer (80 mM, pH 8.0; 5 % v/v (NH4)2SO4). After extensive washing
with binding buffer, a gradual elution of bound protein was performed by applying buffers with
decreasing hydrophobicity: Tris-Cl buffer (80 mM, pH 8.0), Tris-Cl buffer (20 mM, pH 8.0, 0.1%
Tween20 v/v), and pure water. The change from binding buffer to the first elution buffer was
performed in a linear gradient (0-100 % over 20 mL), while other buffer changes were performed
in a one-step gradient. Pooled fractions, eluted with 20 mM Tris-Cl (pH 8.0; 0.1 % Tween20),
were concentrated on an AMICON filter unit (10 kDa cut-off) and dialyzed against
NaH2PO4/Na2HPO4 buffer (50 mM, pH 7.0) for at least 12 hours. Dialyzed fraction was loaded
onto a ResourceQ anion exchange column (CV 1 mL), equilibrated with NaH2PO4/Na2HPO4
buffer (50 mM, pH 7.0). Bound protein was eluted with NaH2PO4/Na2HPO4 buffer (50 mM, pH
7.0) containing 1 M NaCl. The flow through was concentrated on an AMICON filter unit prior to
the application onto a size-exclusion column (HiLoad 16/60, Superdex200, dimensions: 16 x 600
mm), which was equilibrated with Tris-Cl (20 mM, pH 8.0), at a flow rate of 0.6 mL/min.
- 65 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Results
Purification of LDI via a Sumo-tag
The Champion™ pET Sumo Expression System was used to obtain mature Ldi without additional
non-native amino acids after purification. A His6-affinity tag and the Sumo peptide were fused inframe to the 5´-end of the ldi gene. The signal peptide for periplasmatic localization was removed
to prevent premature cleavage of the tag. The Sumo-Ldi expressed matched the in silico predicted
size of 55.2 kDa (Fig. 2).
Figure 2: Expression of the Sumo-Ldi fusion protein in E. coli BL21(DE3) pET42-Sumo-Ldi. (1) noninduced culture, (2) culture induced with 1 mM IPTG.
The fusion protein was successfully expressed in E. coli BL21(DE3) without formation of
inclusion bodies. Sumo protease cleavage resulted in protein bands of 42 kDa and 13 kDa. Based
on 20 % Sumo-Ldi in the soluble protein extract (Fig. 3), the Sumo protease had an activity of 5
μg protein Unit-1 protease h-1 at 12°C in Tris-Cl buffer (40 mM, pH 8.0) during a 15 h incubation.
- 66 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Figure 3: Test of Sumo protease activity under oxidized and reduced conditions with soluble protein
extract from E. coli BL21(DE3) pET-Sumo-Ldi: (1) non-cleaved protein extract, (2) non-cleaved protein
extract with 20 mM DTT, (3) protein extract after cleavage under oxidative conditions, (4) protein extract
after cleavage under with 20 mM DTT.
Although the purification of proteins based on the His6-Sumo-tag was successfully applied in
various cases (Marblestone et al., 2006, Peroutka et al., 2011, Zuo et al., 2005a), we encountered
multiple problems during the purification of the Sumo-Ldi fusion protein. Unusual high
imidazole concentrations were required to achieve selective binding (50 mM imidazole) and
elution (500 mM imidazole) of the fusion protein. Lower imidazole concentrations
(manufacturers manual: 10 mM, 20 mM and 250 mM imidazole in 50 mM NaH2PO4/Na2HPO4
buffer, pH 8.0 for binding, washing and elution) resulted in large quantities of undesired protein
bound to the column (data not shown). Sumo-Ldi, applied in the soluble protein extract (500 mg
at 25 mg mL-1), bound tightly to the column and was not detected in the flow through (Fig. 4,
lane 2). About 75 % of the applied protein passed the column without binding. Bound protein
was eluted by applying 500 mM imidazole and contained the Sumo-Ldi (Fig. 4, lane 3). The
protein eluted over a volume of approximately 50 mL with three not well separated peaks. Higher
imidazole concentrations caused an increase in absorption at 280 nm but no further peaks were
detected with 1 M imidazole in the buffer. The eluate was pooled and subjected to desalting
chromatography. The successful Sumo-protease treatment was confirmed by a shift of the 55 kDa
band to 42 kDa and the formation of a band at 13 kDa in SDS-PAGE. Sumo-Ldi was cleaved
almost completely (Fig. 4, lane 4).
- 67 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Figure 4: Purification of the Sumo-Ldi fusion protein from soluble protein extract of E. coli BL21(DE3)
pET-Sumo-Ldi. (1) soluble extract, (2) flow through from the first run, (3) eluate with 500 mM imidazole
from the first run, (4) eluate after Sumo protease treatment, (5) flow through from the second run, (6)
eluate from the second run, (7) peak shoulder of gelfiltration, (8) major peak of size-exclusion
chromatography.
After the concentrating step 80 mg (5.8 mg mL-1) of protein were obtained from approx. 8 to 10 g
wet biomass. Removal of the His6-Sumo-tag and uncleaved Sumo-Ldi on an Ni2+-IDA column
under the same conditions yielded half of the applied protein in the flow through, while another
15 % of the applied protein was eluted with imidazole. In total, only 70 % of the applied protein
was recovered from the column. The native Ldi complex was predicted to have a retention
volume of about 62 mL on a Superdex200 column (Brodkorb et al., 2010). Protein purified via
the His6-Sumo-tag eluted with a retention volume of 40 to 80 mL, with the main peak between 65
and 80 mL. This corresponded to a mixture of monomeric (80.3 mL), dimeric (73.2 mL), trimeric
(68.9 mL) and tetrameric (66 mL) Ldi complexes (Fig. 5). The main peak was pooled and yielded
11.4 mg of protein (0.76 mg mL-1). This accounted for an overall yield of about 2 % of the
initially applied soluble protein.
Purification of the Sumo-Ldi under denaturative conditions resulted in a similar peak distribution
in SEC as the native Sumo-Ldi and an inactive enzyme after removal of urea. Therefore, this
attempt was not considered as an alternative.
- 68 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Figure 5: Chromatograms from size-exclusion chromatography of Ldi purification: purified as Sumo-Ldi
(dashed line, primary y-axes) and purified as untagged Ldi (solid line, secondary y-axes).
Purification of wildtype Ldi
Ldi was overexpressed in E. coli as wildtype enzyme and as the mature enzyme without signal
peptide for export. The mature Ldi showed a reduced activity compared to the wildtype Ldi in
soluble protein extracts: 43 % and 17 % for dehydratase and isomerase activity, respectively. We
decided to purify the overexpressed Ldi wildtype enzyme.
The linalool dehydratase/isomerase showed an overall hydrophobic character. For this reason, a
hydrophobic interaction chromatography (HIC) in combination with a rather low initial
ammonium sulfate concentration in the binding buffer was chosen as a first purification step.
Soluble extract loaded on the column contained about 550 mg total protein (28 mg mL-1).
Approximately 80 % of the applied protein did not bind to the column. After a washing step with
a buffer containing no ammonium sulfate, the main Ldi fraction eluted in a low molarity Tris-Cl
buffer in the presence of 0.1 % (v/v) Tween20. This fraction contained 3 % of the initially
applied protein (18.5 mg, 0.75 mg mL-1). Another portion of Ldi and highly hydrophobic protein
was eluted with pure water, but not used for further purification steps. Anion exchange
chromatography was performed as a second purification step after the sample was concentrated
and dialyzed. Only 75 % of protein was recovered after the concentrating step. Due to an in silico
predicted isoelectric point of 6.02 the Ldi should bind to the anion exchanger under the given
- 69 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
conditions (pH 7.0). 25 % of the applied protein was recovered in the flow through (3.5 mg, 0.3
mg mL-1), containing also the Ldi activity. This sample was concentrated and applied to sizeexclusion chromatography. The main peak had a retention volume of 64 mL (Fig. 5) and
contained 1.9 mg protein (62.5 % of applied protein; 0.17 mg mL-1). This retention volume
indicated a pentameric enzyme complex, according to our calibration (63.7 mL). Final yields in
pure Ldi (> 98 % purity, Fig. 6) were about 0.33 % of the initially applied total protein.
Figure 6: Purification of the untagged Ldi from soluble protein extract of E. coli BL21(DE3) pET42-Ldi.
(1) soluble extract, (2) HIC flow through, (3) HIC flow through, continued, (4) HIC washing step with 80
mM Tris-Cl, (5) HIC elution step with 20 mM Tris-Cl + 0.1 % v/v Tween20, (6) HIC elution step with
water, first peak, (7) HIC elution step with water, second peak, (8) ResourceQ flow through, (9)
ResourceQ eluate, (10) Size-exculsion chromatography.
- 70 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Table 1: Purification of linalool dehydratase/isomerase from E. coli BL21(DE3) pET42-Ldi. Upper set
shows linalool dehydratase activity. Lower set shows geraniol isomerase activity.
Purification step
Protein [mg]
Acitivty
[pkat]
Specific activity
[pkat mg-1]
Relative specific
activity
Protein
yield [%]
Soluble extract
557.9
30412
55
1
100.0
HIC
18.0
23202
1288
24
3.2
ResourceQ
4.2
11372
2728
50
0.7
SEC
2.0
6049
3011
55
0.4
Soluble extract
557.9
30105
54
1
100.0
HIC
18.0
4333
241
4
3.2
ResourceQ
4.2
1998
479
9
0.7
SEC
2.0
764
380
7
0.4
The specific activities of the wildtype Ldi were successfully enriched by a factor of 55 and 7 for
the dehydratase and isomerase activity, respectively. Total activities were reduced to 20 % and 3
% for the dehydratase and isomerase, respectively (Table 1). For the dehydratase function a
constant increase in specific activity was detected, whereas the isomerase activity dropped in the
last purification step (SEC). The variable increase of activities during purification indicates that
both activities respond individually to changes in physicochemical parameters.
Enzyme activity
Even though the Sumo fusion system was described to be superior over other purification tags for
difficult-to-express proteins (Butt et al., 2005), it yielded for the Ldi fusion a pure protein without
enzymatic activity. Ldi dehydratase and isomerase activities of cleaved Sumo-Ldi were 4.3 pkat
mg protein-1 and 0.6 pkat mg protein-1 in soluble protein extracts, respectively. Samples obtained
during purification showed quite inconsistent results with respect to activity. In most cases
activity was not observed. Wildtype Ldi activities in soluble protein fractions of E. coli
BL21(DE3) pET42-Ldi were more than 10-fold higher (54 pkat mg protein-1 for both activities).
Reasons for that difference can only be speculated as the knowledge on the enzyme mechanism is
- 71 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
still scarce. A possible effect of the signal peptide for periplasmatic localization on the activity of
the Ldi cannot be excluded.
Discussion and conclusion
The Ldi, purified as Sumo-Ldi, was enriched to over 95 % according to SDS-PAGE with the
presented purification protocol. Reported literature values are in the range of 90 to 96 % purity
(Su et al., 2006, Zuo et al., 2005b, Wang et al., 2010, Li et al., 2011). We observed a tight
binding of the Sumo-Ldi to the Ni2+-IDA column. The Ldi has an overall hydrophobic character
and contains multiple endogenous histidine (12/371) and cysteine residues (4/371). These amino
acids are known to complex metals ions (Porath et al., 1975) and surface patches of histidines and
other amino acids cause unspecific binding of proteins to IMAC-columns. Unspecific binding of
hydrophobic proteins to IMAC-columns has also been reported (Arnold, 1991, Bolanos-Garcia
and Davies, 2006). Application of the Ldi, purified as Sumo-Ldi, onto size-exclusion
chromatography resulted in a broad peak between 65 and 80 mL, indicating a complex mixture of
the Ldi subunits assembled into a mono-, di-, tri- or tetrameric complexes. The Ldi was reported
to refold into an active complex after urea treatment (Brodkorb et al., 2010). Self-assembly of the
Ldi into its native, functional state after SEC was assumed. However, no activity was recovered.
Purification attempts under denaturative conditions in the presence of 6 M urea with subsequent
removal of urea by dialysis were not successful either (data not shown). Size-exclusion
chromatography of the purified wildtype enzyme resulted in a single peak. This difference in the
SEC separations might be attributed to a conformational change of the Ldi during purification as
Sumo-Ldi. Likely reasons are the absence of signal peptide, the presence of Sumo peptide and the
high imidazole concentrations during purification. Removal of the signal peptide resulted in a
cytoplasmic expression. The Ldi contains four cysteine residues, likely contributing to the tertiary
structure by disulfide bond formation. Correct formation of disulfide bonds occurs in the DsbADsbB and DsbC-DsbD systems in the periplasm (Denoncin and Collet, 2012). Consequently, a
cytoplasmic expression of the Ldi would lead to misfolding and an inactive enzyme. The low
activity of the N-terminally truncated Ldi mutant supports this hypothesis. The initial presence of
the Sumo-peptide might have promoted a misfolding additionally.
- 72 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
The overall hydrophobicity of Ldi led to the development of a new purification protocol for the
heterologously expressed wildtype Ldi. In the first purification protocol, the Ldi was bound to a
Butyl-Sepharose column in the presence of 15 % (v/v) of a saturated ammonium solution
(Brodkorb et al., 2010). Judged by the specific activity, this purification step was the most
effective. In order to increase efficiency of the purification, we used a Phenyl-Sepharose as initial
step and lowered the ammonium concentration. This lead to the removal of more than 80 %
undesired protein. In contrast to the old protocol, Ldi was not eluted with urea but with a low
molarity Tris-Cl buffer (20 mM) containing Tween20 to increase the hydrophobicity of the buffer
and to stabilize the protein. A subfraction of Ldi was also eluted with pure water but did not lead
to better overall yields. In the purification of the wildytpe enzyme from C. defragrans 65Phen,
Ldi was bound to a ResourceQ column at pH 7.0 and eluted with 150 mM NaCl (Brodkorb et al.,
2010). In the presented protocol we applied the same column and pH parameter. However, the
Ldi did not bind to the column. The binding was likely affected by the presence of the detergent
Tween20, which may be not fully removed by dialysis. The phenomenon of detergents shielding
the possible interaction sites of protein when bound to them is known in literature (Arnold and
Linke, 2008). We encountered additional protein loss during the concentrating steps using the
AMICON filter units. Even though it is not described in literature, it is known in the scientific
community that proteins tend to stick to the membrane of those filter units, especially at higher
concentrations. It is suggested by the manufacturer to block unspecific binding sites by addition
of bovine serum albumin or polymeric additives (e.g. polyethylene glycol). Final polishing by
size-exclusion chromatography resulted in a single peak at the retention volume of a homopentameric Ldi complex rather than the homo-tetrameric complex as previously described
(Brodkorb et al., 2010). The difference between the predicted retention volumes of a
homopentamer and a homotetramer are for the column 2.3 mL, which indicates a suboptimal
resolution of the column for this size determination. The retention volume of the pentameric Ldi
complex was 64 mL.
- 73 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
Conclusion
A Ldi mutant was successfully expressed with a N-terminally purification tag as His6-Sumo-Ldi
in E. coli BL21(DE3). Even though the purification yielded a single band in the SDS-PAGE, the
enzymatic activity was not preserved. Alterations on the N-terminus of the Ldi seem to have an
irreversible negative effect on conformation and activity. Alternatively, the heterologously
expressed wildtype enzyme was successfully purified in its active state by a newly developed
purification scheme to quantities sufficient for structural and mechanistic studies.
Competing interests
The authors declare that they have no competing interests.
Authors´ contribution
RM designed and performed the experiments. RM and JH drafted the manuscript. All authors
read and approved the final manuscript.
Acknowledgments
This study was financed by the Max Planck Society.
- 74 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
References
ARNAU, J., LAURITZEN, C., PETERSEN, G. E. & PEDERSEN, J. 2006. Current strategies for the use
of affinity tags and tag removal for the purification of recombinant proteins. Protein Expression
and Purification, 48, 1-13.
ARNOLD, F. H. 1991. Metal-affinity separations: a new dimension in protein processing. BioTechnology, 9, 151-156.
ARNOLD, T. & LINKE, D. 2008. The use of detergents to purify membrane proteins. Curr Protoc
Protein Sci, Chapter 4, Unit 4 8 1-4 8 30.
BOLANOS-GARCIA, V. M. & DAVIES, O. R. 2006. Structural analysis and classification of native
proteins from E. coli commonly co-purified by immobilised metal affinity chromatography.
Biochimica et Biophysica Acta (BBA) - General Subjects, 1760, 1304-1313.
BRADFORD, M. M. 1976. Rapid and sensitive method for quantitation of microgram quantities of
protein utilizing principle of protein-dye binding. Analytical Biochemistry, 72, 248-254.
BRODKORB, D., GOTTSCHALL, M., MARMULLA, R., LÜDDEKE, F. & HARDER, J. 2010. Linalool
dehydratase-isomerase, a bifunctional enzyme in the anaerobic degradation of monoterpenes.
Journal of Biological Chemistry, 285, 30436-30442.
BUTT, T. R., EDAVETTAL, S. C., HALL, J. P. & MATTERN, M. R. 2005. SUMO fusion technology
for difficult-to-express proteins. Protein Expression and Purification, 43, 1-9.
CHARLTON, A. & ZACHARIOU, M. 2008. Immobilized metal ion affinity chromatography of histidinetagged fusion proteins. Methods in Molecular Biology. Humana Press Inc, 999 Riverview Dr, Ste
208, Totowa, Nj 07512-1165 USA.
CHARLTON, A. & ZACHARIOU, M. 2011. Tag Removal by Site-Specific Cleavage of Recombinant
Fusion Proteins. Protein Chromatography: Methods and Protocols.
DENONCIN, K. & COLLET, J.-F. 2012. Disulfide bond formation in the bacterial periplasm: major
achievements and challenges ahead. Antioxidants & Redox Signaling, 19, 63-71.
FOSS, S., HEYEN, U. & HARDER, J. 1998. Alcaligenes defragrans sp. nov., description of four strains
isolated on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alphaphellandrene) and nitrate. Systematic and Applied Microbiology, 21, 237-244.
HAY, R. T. 2007. SUMO-specific proteases: a twist in the tail. Trends in Cell Biology, 17, 370-376.
HEYEN, U. & HARDER, J. 2000. Geranic acid formation, an initial reaction of anaerobic monoterpene
metabolism in denitrifying Alcaligenes defragrans. Applied and Environmental Microbiology, 66,
3004-3009.
KYTE, J. & DOOLITTLE, R. F. 1982. A simple method for displaying the hydropathic character of a
protein. Journal of Molecular Biology, 157, 105-132.
- 75 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
LI, J., CUI, X., JI, H., QIU, T., JI, X., DU, M., WU, H., XU, X. & ZHANG, S. 2011. High efficient
expression of bioactive human BMP-14 in E. coli using SUMO fusion partner. The Protein
Journal, 30, 592-597.
LI, S. J. & HOCHSTRASSER, M. 1999. A new protease required for cell-cycle progression in yeast.
Nature, 398, 246-251.
LÜDDEKE, F. & HARDER, J. 2011. Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences, 66, 409-412.
MALAKHOV, M. P., MATTERN, M. R., MALAKHOVA, O. A., DRINKER, M., WEEKS, S. D. &
BUTT, T. R. 2004. SUMO fusions and SUMO-specific protease for efficient expression and
purification of proteins. Journal of Structural and Functional Genomics, 5, 75-86.
MARBLESTONE, J. G., EDAVETTAL, S. C., LIM, Y., LIM, P., ZUO, X. & BUTT, T. R. 2006.
Comparison of SUMO fusion technology with traditional gene fusion systems: Enhanced
expression and solubility with SUMO. Protein Science, 15, 182-189.
MUKHOPADHYAY, D. & DASSO, M. 2007. Modification in reverse: the SUMO proteases. Trends in
Biochemical Sciences, 32, 286-295.
PEROUTKA, R. J., III, ORCUTT, S. J., STRICKLER, J. E. & BUTT, T. R. 2011. SUMO Fusion
Technology for Enhanced Protein Expression and Purification in Prokaryotes and Eukaryotes.
Heterologous Gene Expression in E coli: Methods and Protocols.
PORATH, J., CARLSSON, J., OLSSON, I. & BELFRAGE, G. 1975. Metal chelate affinity
chromatography, a new approach to protein fractionation. Nature, 258, 598-599.
SATAKARNI, M. & CURTIS, R. 2011. Production of recombinant peptides as fusions with SUMO.
Protein Expression and Purification, 78, 113-119.
SU, Z. J., HUANG, Y. D., ZHOU, Q. N., WU, Z. L., WU, X. P., ZHENG, Q., DING, C. C. & LI, X. K.
2006. High-level expression and purification of human epidermal growth factor with SUMO
fusion in Escherichia coli. Protein and Peptide Letters, 13, 785-792.
WALLS, D. & LOUGHRAN, S. T. 2011. Tagging recombinant proteins to enhance solubility and aid
purification. Protein Chromatography: Methods and Protocols.
WANG, H. Y., XIAO, Y. C., FU, L. J., ZHAO, H. X., ZHANG, Y. F., WAN, X. S., QIN, Y. X., HUANG,
Y. D., GAO, H. C. & LI, X. K. 2010. High-level expression and purification of soluble
recombinant FGF21 protein by SUMO fusion in Escherichia coli. BMC Biotechnology, 10.
YOUNG, C. L., BRITTON, Z. T. & ROBINSON, A. S. 2012. Recombinant protein expression and
purification: A comprehensive review of affinity tags and microbial applications. Biotechnology
Journal, 7, 620-634.
- 76 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
ZUO, X., LIE, S., HALL, J., MATTERN, M. R., TRAN, H., SHOO, J., TAN, R., WEISS, S. R. & BUTT,
T. R. 2005a. Enhanced expression and purification of membrane proteins by SUMO fusion in
Escherichia coli. Journal of Structural and Functional Genomics, 6, 103-111.
ZUO, X., MATTERN, M. R., TAN, R., LI, S., HALL, J., STERNER, D. E., SHOO, J., TRAN, H., LIM,
P., SARAFIANOS, S. G., KAZI, L., NAVAS-MARTIN, S., WEISS, S. R. & BUTT, T. R. 2005b.
Expression and purification of SARS coronavirus proteins using SUMO-fusions. Protein
Expression and Purification, 42, 100-110.
- 77 -
Chapter III - A novel purification protocol for the linalool dehydratase/isomerase
- 78 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Chapter IV
X-ray structure of linalool dehydratase/isomerase reveals enzymatic alkene synthesis
Sina Weidenweber1', Robert Marmulla2', Ulrich Ermler1 and Jens Harder2
1
Dept. of Molecular Membrane Biology, Max Planck Institute of Biophysics, D-60348 Frankfurt
am Main, Germany, and 2Dept. of Microbiology, Max Planck Institute for Marine Microbiology,
D-28359 Bremen, Germany
'
These authors contributed equally to this work.
*corresponding authors:
Ulrich Ermler,1Dept. of Molecular Membrane Biology, Max Planck Institute of Biophysics, D60348 Frankfurt am Main, Germany, Email: ulrich.ermler@biophys.mpg.de, Phone:
++496910546303, Fax:++496910541002
Jens Harder, 2Max Planck Institute for Marine Microbiology, Celsiusstrasse 1
28359 Bremen, Germany, Email: jharder@mpi-bremen.de, Phone: ++49 421 2028 750, Fax:
++49 421 2028 590
Manuscript in preparation
- 79 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Abstract
Hydrocarbon-degrading microorganisms have evolved enzymes suitable for the sustainable
biotechnological production of intermediates for fuel and polymer synthesis. Linalool
dehydratase/isomerase (Ldi) has been documented in several patents and patent applications as
enzyme for the biological production of butadiene and isoprene from alkenols. In biology, this
enzyme is involved in the anaerobic monoterpene biotransformation. It reversibly isomerizes the
primary monoterpene alcohol geraniol into the tertiary alcohol (S)-linalool and dehydrates (S)linalool to the alkene E-myrcene without using a cofactor. In this study, we report on the crystal
structures of Ldi from Castellaniella defragrans 65Phen and its complex with a terpene substrate
at 1.9 and 2.45 Å resolution, respectively. Architecturally, Ldi is organized as a homopentameric
protein complex with each monomer being built up of an (α,α)6 barrel fold. The terpene binding
site is located inside a 15 Å long predominantly hydrophobic channel at the monomer-monomer
interface which is rather unusual for (D,D)6 barrel fold proteins and dehydratases. Dependent on
the channel considered, the electron density for the terpene was either interpreted as β-myrcene or
geraniol. Three amino acid clusters pointing from different directions to the functional groups of
the terpene contain the key residues for catalyzing the hydroxy mutase and dehydratase reactions
by acid/base catalysis. The first cluster includes His128, Cys179, Tyr72 and a H2O molecule, the
second Asp38´, Tyr239 and Gln178 and the third Tyr44´, Cys170 and Glu171. The detailed
information on the active site and the terpene binding mode will guide future developments of
Ldi for biotechnological applications.
- 80 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Main text
Essential oils produced by plants mainly consist of monoterpenes; volatile hydrocarbons with the
composition C10H16 [1]. More than 127 Tg C yr-1 monoterpenes are thus emitted into the
atmosphere and an unknown amount enters soils, lakes and rivers [2]. Up to one liter
monoterpenes m-2 are in forest soils and monoterpenes are among the most frequent indoor
volatile hydrocarbons [3, 4]. Microorganisms use this abundant carbon source, but little is known
about degradation pathways and the enzymes involved in comparison to those of e.g. alkanes and
aromatic compounds [5, 6]. A novel enzyme, the linalool dehydratase/isomerase (LDI), was
found in the betaproteobacterium C. defragrans 65Phen that mineralizes monoterpenes coupled
to denitrification in the absence of molecular oxygen [7]. This periplasmic enzyme involved in
terpene degradation reversibly catalyzes the isomerisation from the primary alkenol geraniol into
the tertiary alkenol (S)-linalool and its dehydration to E-myrcene [8, 9] (Fig. 1). Dehydratase and
isomerase activities of the bifunctional Ldi were only detectable in the presence of a reductant
(DTT or dithionite) and the absence of O2 [8, 9]. The dehydration reaction of Ldi or genetically
engineered variants thereof can be used for transforming smaller substrate molecules into
butadiene and isoprene according to patents and patent claims [10-15]. The market for these two
compounds, precursors for many polymers (e.g. nylon, polyester, polyisoprene), amounts to
billions of dollars annually and has seen recently a supply shortage due to a transition in the
chemical industry from oil to gas as natural resource [16].
To optimize Ldi for biotechnological purposes by a more rational approach, detailed structural
data of the enzyme are indispensable, in particular, because Ldi does not share any sequence
relationships to other protein of sequence data banks. Therefore, Ldi was overexpressed as
wildtype-protein in E. coli and purified in an improved procedure as mature protein lacking the
N-terminal signal peptide for periplasmatic localization. Upon crystallization its X-ray structure
was determined with the single isomorphous replacement anomalous scattering method (SIRAS)
using thimerosal as heavy metal compound and refined to R/Rfree values of 19.0/22.5 % at 1.9 Å
resolution (Table 2).
The crystal structure of Ldi revealed a homopentameric protein complex (Fig. 2A) in agreement
with size-exclusion chromatographic data in solution. The pentamer resembles a planar rosette
with a central hole of ca. 30 Å in diameter and is held together by an extended monomermonomer interface. Architecturally, each monomer is built up of a classical (α,α)6 barrel fold
- 81 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
composed of six inner helices (63-80, 124-139, 179-193, 238-251, 294-306 and 341-351) flanked
by six lateral helices (25-35, 85-100, 146-163, 198-212, 255-266 and 309-322) oriented in an
antiparallel fashion (Fig. 2B). The bottom of the barrel is locked by short loops connecting the
inner and lateral helices and by the N (1-25)- and C (352-366)-terminal segments while its
entrance is formed by variable segments linking the lateral and the next inner helices (Fig. 2B). A
protein data bank research using the Dali server [17] revealed a large number of proteins with an
(α,α)6 barrel fold; the highest structural similarities were computed between Ldi and
rhamnogalacturonyl hydrolase from Bacillus subtilis (2d8l) [18] as well as cellobiose-2epimerase from Ruminococcus albus (3vw5) [19], with sequence identities of 16.5 and 15.8 %,
and with rmsds of 3.3 and 3.4 Å, respectively, using 77 % and 75 % of the Cα atoms for
calculation. Interestingly, (α,α)6 barrel folds are frequently found in isomerases and in enzymes
of the terpenoid metabolism, i.e. terpenoid cyclases and protein prenyl transferases [20], but in
contrast to Ldi these enzymes operate as a monomer.
The substrate binding site in (α,α)6 barrel enzymes is embedded inside a cavity at the barrel
entrance whereas the substrate cavity in Ldi is partly capped by the irregular segment 36’-52’ of
the neighbor monomer (marked by an apostrophe) of the homopentamer. Notably, this segment is
fixed to the (α,α)6 barrel core by an intra-subunit disulfide bond between Cys48’ and Cys101’
(Fig. 2B). Site-specific exchange of one of these cysteines completely inactivated the enzyme.
Despite the attached neighboring subunit the bottom of the cavity is still accessible from bulk
solvent via a narrow channel, ca. 15 Å long and 7 Å wide, which is placed at the interface
between the two subunits (Fig. 3A). The entrance of the channel is positioned at the inner side of
the rosette and lined up by several nonpolar residues including Ile37’, Ile40’, Pro59, Val62,
Val292 and Leu340 which may attract the hydrophobic terpenes. This type of active site
architecture is new in (α,α)6 barrel proteins and therefore unpredictable without structural data.
An alternative strategy of active site encapsulation has been reported for monomeric squalene
cyclases [21] that use an additional domain for this purpose. In contrast, rhamnogalacturonyl
hydrolase [18] and cellobiose-2-epimerase [19], structurally most related to Ldi, are endowed
with a more polar and exposed substrate binding site only partly shielded by the N- and Cterminal arms and an expanded loop.
- 82 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
To learn more about substrate binding and the catalytic mechanism, Ldi crystals were soaked
with (R,S)-linalool and dithiothreitol under enzyme activity conditions. Subsequently, the crystals
were frozen and structurally characterized at 2.45 Å resolution. Significant extra electron density
was clearly visible at the inferred substrate binding site at the bottom of the five channels in the
asymmetric unit (Fig. 3, 4). Although the terpene binding sites were incompletely and partially
inhomogeneous occupied, the best fit was realized with β-myrcene in two channels and geraniol
in the three remaining channels (Fig. 3B, 4B). This finding implicates a dehydratase or isomerase
reaction in the Ldi crystal which is plausible because the crystals were soaked at pH 9, the pH
optimum for enzymatic activity. In the free form thermodynamically most stable is β-myrcene,
followed by (S)-linalool and geraniol [8, 22]. In the enzyme complex under the conditions of
crystallization, the interaction with the enzyme seems to prefer geraniol over (S)-linalool. The
monoterpenes are embedded into the channel such that the unpolar methylpentene moiety is
directed towards the channel entrance and surrounded by various hydrophobic residues including
Val62, Tyr65, Phe69, Tyr239, Trp243, Leu294, Phe298, Leu341, Asp38’ and Phe39‘ (Fig. 3).
The functionally relevant buta-1,3-diene or but-2-en-1-ol moieties of β-myrcene and geraniol are
located at the channel bottom and are flanked by Tyr65, Met124, Cys170, Phe176, Gln178,
Cys179, Tyr239, Asp38’, Phe39’ and Tyr 44’ (Fig. 3B).
The described structural basis for substrate binding implicates that Ldi performs the dehydratase
and hydroxyl mutase reactions at the same position at the bottom of a narrow channel. As no
organic cofactor or metal ions were found, both reactions apparently proceed by acid/base
catalysis using protonable amino acid side chains. The functional groups of the monoterpenes
comprising the hydroxy groups of (S)-linalool and geraniol and the C3-C3’ double bond of βmyrcene are contacted by several polar side chains subdivided into three catalytic clusters (Fig.
4). Cluster I includes Cys179-SH, a firmly bound H2O, His128-NE2 and Tyr72-OH arranged in a
line hydrogen-bonded to each other. Notably, Cys179 is located at the N-terminus of helix 178192 which might stabilize a deprotonated thiolate as described for diaminopimelate epimerase
[23]. From the opposite side of the channel points cluster II toward the terpene centered at
Asp38’ of the neighboring subunit. Its carboxylate group interacts with Tyr44’-OH, Tyr239-OH
and Gln178-NE2. Moreover, the terpene-free Ldi structure, determined at higher resolution,
reveals additional visible H2O in hydrogen-bond distance with Asp38’ that are linked to a nearby
solvent-filled cavity. One of these H2O, if also present in the substrate bound complex, is ca. 5 Å
- 83 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
apart from C3 and C3’ and can move without steric hindrance toward the C3 atom. In this
modeled position it interacts with Asp38’, Tyr44’ and Tyr239 and is termined as 1. catalytic H2O
(Fig. 4A). Both clusters contact the C3-C3’ group of the terpenes and this ensures the
regioselectivity of the hydration reaction. Cluster III at the apex of the channel is primarily
composed of Cys170, Glu171 and Tyr44’. These residues might contact and activate one H2O
modeled into an empty space in front of the β-myrcene C1 (and equivalently of the (S)-linalool
C1) (Fig. 4A). This 2. catalytic H2O, which partly overlaps with the C1 hydroxy group of
geraniol of the superimposed Ldi-geraniol complex, is involved in the hydroxyl mutase reaction
(Fig. 4B).
All
three
catalytic
clusters
protonation/deprotonation
have,
in
principle,
the
competence
and/or hydroxylation/dehydroxylation steps
to
perform
required
for
the
the
dehydratase and isomerase reaction. The catalytic mechanism of Ldi outlined in Fig. 4 is based
on the orientation of the monoterpene relative to the three catalytic clusters. For finding the most
optimal solution, the Ldi-terpene complex structure was refined in several orientations of βmyrcene. We choose the structure with the minimal residual electron density in the substrate
binding site and the most uniform conformation of the terpenes in the five channels. Accordingly,
hydration of E-myrcene to (S)-linalool might proceed by protonating C3’ by Cys179 of cluster I
and hydroxylating C3 by the 1. catalytic H2O of cluster II in an anti-selective manner (Fig. 4A,
C). Both reactions have to be synchronized to achieve the previously observed stereospecificity
of (S)-linalool formation [9]. An alternative scenario involves a nucleophilic attack of Cys179thiolate onto C3, C3’- protonation i.e. by Asp38’ and a nucleophilic attack of the 1. catalytic H2O
onto the Cys179-terpene thioether intermediate. Hydroxylation of C3 by the water sandwiched
between His128 and Cys179 at C3 would lead to (R)-linalool in the fitted orientation of βmyrcene and was therefore discarded (Fig. 4). The hydroxyl mutase reaction from geraniol to
(S)-linalool might proceed by a nucleophilic attack of the catalytic H2O on C3, an allylic
rearrangement and a release of the C1-hydroxyl group (Fig. 4B). Interestingly, Tyr44’ could act
both as acid and base in the isomerization reaction as it sits between the two functional groups
and can switch to a protonated/deprotonated form at the pH optimum of 9.
Hydration of an isolated electron-rich carbon double bond with the poor nucleophile water is a
chemically challenging reaction [24] and therefore nature normally prefers a conjugated electronpoor double bond using the Michael reaction for alcohol formation using enzymes like
- 84 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
aspartases/fumarases, enoyl-CoA hydratases and aconitases [25-27]. Isomerization between (S)linalool and geraniol is a basic chemical reaction, which was also found in a few other
intramolecular hydroxyl transferases as i.e. isochorismate synthase [28].
Hydratases and
isomerases use frequently histidines and aspartates or glutamates but rarely cysteines as
(de)protonable groups. In the postulated mechanism of Ldi a central role is attributed to Cys170
and Cys179 which was experimentally verified by site-directed mutagenesis studies. Each
exchange of the individual cysteines to serine resulted in an inactive enzyme. This is supported
by the inactivity of Ldi under aerobic conditions and in the absence of DTT which is likely due to
their oxidation either to cysteine sulfoxide or to Cys170-Cys179 disulfide (distance 4.1 Å). The
use of cysteines in Ldi for acid/base catalysis is not without precedents. In glutamate racemase
two cysteines act as acid/base [29] and in malate isomerase one cysteine acts as nucleophile
forming a covalent thioether bond and one as proton donor [30].
A sustainable world requires the transformation of renewable resources into intermediates of the
fine chemical industry. Low cost-production of butane-1,4-diol from feedstock by engineered
microorganisms has already been developed [31]. For butadiene and isoprene, several patents
used linalool dehydratases or derivatives thereof for the final step in the synthesis of the olefins
[10-15]. The presented structural data of Ldi revealed the terpene binding site between two
subunits inside a hydrophobic channel, unprecedentedly for (α,α)6 barrel proteins, and the
geometry of the bound E-myrcene and geraniol relative to specifically positioned amino acids
involved in binding and catalysis (Fig. 2-4). This knowledge provides a rational basis for
expanding the substrate specificity of the enzyme and for increasing its turnover rate. From the
new platform more targeted biotechnological avenues can be explored.
- 85 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Authors´ contribution
SW, RM, UE and JH designed the experiments. RM performed the protein expression, protein
purification and generation and analysis of mutants. SW performed the protein crystallography
and data evaluation. SW, RM, UE and JH drafted the manuscript. All authors read and approved
the final manuscript.
Acknowledgements
This work was supported by the German Research Foundation (DFG) in the SPP1319 and the
Max-Planck Society. We thank Hartmut Michel and Friedrich Widdel for continuous support and
the staff of PXII at the Swiss Light Source (Villigen) for help during data collection.
- 86 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
References
1.
Dudareva N, Klempien A, Muhlemann JK, Kaplan I: Biosynthesis, function and metabolic
engineering of plant volatile organic compounds. New Phytol 2013, 198:16-32.
2.
Günther A, Hewitt CN, Erickson D, Fall R, Geron C, Graedel T, Harley P, Klinger L, Lerdau M,
McKay WA et al: A global model of natural volatile organic compound emission. Journal of
Geophysical Research-Atmospheres 1995, 100:8873-8892.
3.
Ziemann PJ, Atkinson R: Kinetics, products, and mechanisms of secondary organic aerosol
formation. Chemical Society Reviews 2012, 41:6582-6605.
4.
Wilt FM, Miller GC, Everett RL, Hackett M: Monoterpene concentrations in fresh, senescent,
and decaying foliage of single-leaf pinyon (Pinus monophylla Torr and Frem: Pinaceae)
from the Western Great-Basin. Journal of Chemical Ecology 1993, 19:185-194.
5.
Marmulla R, Harder J: Microbial monoterpene transformations – A review. Frontiers in
Microbiology 2014, 5.
6.
Heider J, Schühle K: Anaerobic biodegradation of hydrocarbons including methane. In: The
Prokaryotes. Edited by Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F: Springer
Berlin Heidelberg; 2013: 605-634.
7.
Foss S, Heyen U, Harder J: Alcaligenes defragrans sp. nov., description of four strains isolated
on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alpha-phellandrene)
and nitrate. Systematic and Applied Microbiology 1998, 21:237-244.
8.
Brodkorb D, Gottschall M, Marmulla R, Lüddeke F, Harder J: Linalool dehydratase-isomerase,
a bifunctional enzyme in the anaerobic degradation of monoterpenes. Journal of Biological
Chemistry 2010, 285:30436-30442.
9.
Lüddeke F, Harder J: Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences 2011, 66:409-412.
10.
Botes AL, van Eck Conradie A. Methods for biosynthesis of isoprene. 2015.
PCT/US2014/049786.
11.
Botes AL, van Eck Conradie A. Methods for biosynthesizing 1,3-butadiene. 2015.
US20150079654.
12.
Campbell P, Bredow S, Zhou H, Doneske S, Monticello DJ. Microorganisms and processes for
the production of isoprene. 2014. US20140349362.
13.
Coelho PS, Farrow MF, Smith MA. De novo metabolic pathways for isoprene biosynthesis.
2014. PCT/US2013/067079.
14.
Koch DJ, Lopes MSG, Gouvea IE, Zeidler AFB, Dumaresq ASR, Rodrigues MEP. Modified
microorganisms and methods of using same for anaerobic coproduction of isoprene and
acetic acid. 2015. WO2015002977.
- 87 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
15.
Marlière P. Production of volatile dienes by enzymatic dehydration of light alkenols. 2015.
US 14/195,738.
16.
Bailey MP: The future of butadiene. Chemical Engineering 2014, 121:19-24.
17.
Holm L, Rosenström P: Dali server: conservation mapping in 3D. Nucleic Acids Research
2010, 38:W545-W549.
18.
Itoh T, Ochiai A, Mikami B, Hashimoto W, Murata K: Structure of unsaturated
rhamnogalacturonyl hydrolase complexed with substrate. Biochemical and Biophysical
Research Communications 2006, 347:1021-1029.
19.
Fujiwara T, Saburi W, Inoue S, Mori H, Matsui H, Tanaka I, Yao M: Crystal structure of
Ruminococcus albus cellobiose 2-epimerase: Structural insights into epimerization of
unmodified sugar. FEBS Letters 2013, 587:840-846.
20.
Wendt KU, Schulz GE: Isoprenoid biosynthesis: manifold chemistry catalyzed by similar
enzymes. Structure 1998, 6:127-133.
21.
Wendt KU, Poralla K, Schulz GE: Structure and function of a squalene cyclase. Science 1997,
277:1811-1815.
22.
Foss S, Harder J: Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. FEMS Microbiol Letters 1997,
149:71-75.
23.
Pillai B, Cherney MM, Diaper CM, Sutherland A, Blanchard JS, Vederas JC, James MNG:
Structural insights into stereochemical inversion by diaminopimelate epimerase: An
antibacterial drug target. Proceedings of the National Academy of Sciences 2006, 103:86688673.
24.
Resch V, Hanefeld U: The selective addition of water. Catal Sci Technol 2015, 5:1385-1399.
25.
Agnihotri G, Liu H-w: Enoyl-CoA hydratase: Reaction, mechanism, and inhibition.
Bioorganic & Medicinal Chemistry 2003, 11:9-20.
26.
Lauble H, Kennedy MC, Beinert H, Stout CD: Crystal structures of aconitase with transaconitate and nitrocitrate bound. Journal of Molecular Biology 1994, 237:437-451.
27.
Puthan Veetil V, Fibriansah G, Raj H, Thunnissen A-MWH, Poelarends GJ: Aspartase/fumarase
superfamily: A common catalytic strategy involving general base-catalyzed formation of a
highly stabilized aci-carboxylate intermediate. Biochemistry 2012, 51:4237-4243.
28.
Sridharan S, Howard N, Kerbarh O, Błaszczyk M, Abell C, Blundell TL: Crystal structure of
Escherichia coli enterobactin-specific isochorismate synthase (EntC) bound to its reaction
product isochorismate: Implications for the enzyme mechanism and differential activity of
chorismate-utilizing enzymes. Journal of Molecular Biology 2010, 397:290-300.
29.
Glavas S, Tanner ME: Active site residues of glutamate racemase. Biochemistry 2001, 40:61996204.
- 88 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
30.
Fisch F, Fleites CM, Delenne M, Baudendistel N, Hauer B, Turkenburg JP, Hart S, Bruce NC,
Grogan G: A covalent succinylcysteine-like intermediate in the enzyme-catalyzed
transformation of maleate to fumarate by maleate isomerase. Journal of the American
Chemical Society 2010, 132:11455-11457.
31.
Yim H, Haselbeck R, Niu W, Pujol-Baxley C, Burgard A, Boldt J, Khandurina J, Trawick JD,
Osterhout RE, Stephen R et al: Metabolic engineering of Escherichia coli for direct production
of 1,4-butanediol. Nature Chemical Biology 2011, 7:445-452.
32.
Bradford MM: Rapid and sensitive method for quantitation of microgram quantities of
protein utilizing principle of protein-dye binding. Analytical Biochemistry 1976, 72:248-254.
33.
Kabsch W: XDS. Acta Crystallographica Section D: Biological Crystallography 2010, 66:125132.
34.
Schneider TR, Sheldrick GM: Substructure solution with SHELXD. Acta Crystallographica
Section D 2002, 58:1772-1779.
35.
de La Fortelle E, Bricogne G: SHARP: A maximum-likelihood heavy-atom parameter
refinement program for the MIR and MAD methods. Methods in Enzymology 1997, 276:472494.
36.
Jones TA, Zou J-Y, Cowan SW, Kjeldgaard M: Improved methods for building protein models
in electron density maps and the location of errors in these models. Acta Crystallographica
Section A 1991, 47:110-119.
37.
Cowtan KD: Joint CCP4 and ESF-EACBM newsletter on protein crystallography. 1994:3438.
38.
Winn MD, Ballard CC, Cowtan KD, Dodson EJ, Emsley P, Evans PR, Keegan RM, Krissinel EB,
Leslie AGW, McCoy A et al: Overview of the CCP4 suite and current developments. Acta
Crystallographica Section D 2011, 67:235-242.
39.
Emsley P, Cowtan K: Coot: model-building tools for molecular graphics. Acta
Crystallographica Section D 2004, 60:2126-2132.
40.
Murshudov GN, Vagin AA, Dodson EJ: Refinement of macromolecular structures by the
maximum-likelihood method. Acta Crystallographica Section D 1997, 53:240-255.
41.
Adams PD, Afonine PV, Bunkoczi G, Chen VB, Davis IW, Echols N, Headd JJ, Hung L-W,
Kapral GJ, Grosse-Kunstleve RW et al: PHENIX: a comprehensive Python-based system for
macromolecular structure solution. Acta Crystallographica Section D 2010, 66:213-221.
42.
Schrodinger L: The PyMOL Molecular Graphics System, Version 1.4.1. In.; 2010.
- 89 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Figures
Figure 1: Reaction of Ldi. Ldi catalyzes the hydration of myrcene to (S)-linalool and the hydroxyl transfer
from (S)-linalool to geraniol. The presence of molecular oxygen reversibly inactivates Ldi [8, 9]. The
three terpenes are composed of a methylpentene tail (left of green line) and butadi-1,3-en (E-myrcene),
but-1-en-3-ol ((S)-linalool) and but-2-en-1-ol (geraniol) heads.
Figure 2: Structure of Ldi. A) The homopentamer. The terpene (orange), shown as stick model, is
accessible by a channel whose entrance points toward the interior of the pentameric ring (black arrow). B)
The Ldi dimer. The Ldi monomer is built up of an (D,D)6 barrel (inner/outer helices in light blue/blue,
helix linkers and N- and C-terminus in blue). The cavity at the barrel entrance has a depth of ca. 7 Å and a
width of ca. 10 Å and is capped by a loop (green) of the neighboring subunit (gray). Thus, the dimer
represents in the functional unit.
- 90 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Figure 3: Substrate binding. A) Channel architecture. The substrate binding site is embedded inside of 15
Å long and narrow L-shaped channel (shown as red surface) accommodated between two subunits (green,
blue) of the pentamer. B) The Ldi-myrcene structure. Myrcene (orange) is bound at the end of the
channel. Its electron density is contoured at 0.9 σ. The functional buta-1,3-diene group points to the apex,
the methylpentene moiety to the entrance. Tyr65 stacks against the entire terpene molecule but appears not
to be involved in acid/base catalysis due to its hydrophobic surrounding.
- 91 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Figure 4: Catalytic reaction of Ldi. A) The dehydration reaction. The active site is subdivided into three
catalytic clusters termed cluster I (yellow), II (blue), and III (green) grouped around myrcene (orange).
The 1. catalytic H2O (grey) was modelled into a site between myrcene, Tyr44’, Asp38’ and Gln178 and
the 2. catalytic H2O between Tyr44’, Cys160 and Glu171. B) The hydroxyl mutase reaction. The channel
with the highest occupied geraniol and the three catalytic cluster were shown. The electron density of
geraniol has a contour level of σ = 0.8. C) Postulated mechanism. Schematic representation of the catalytic
process based on the fitted orientation of the terpenes based on a 2.5 Å electron density map. In the
hydroxyl transfer reaction from geraniol to (S)-linalool Asp38’ and/or Tyr44’ abstract a proton from the 1.
catalytic H2O and Glu171, Tyr44’ or, in particular, Cys170 protonate the geraniol C1 hydroxy group.
- 92 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Methods
Cultivation of E.coli BL21(DE3) and expression of ldi
Linalool dehydratase/isomerase was heterologous expressed in E. coli BL21(DE3) pET42-Ldi [8]
in 500 ml rich medium (yeast extract 12 g L-1, peptone 14 g L-1, NaCl 5 g L-1, MgSO4 x 7 H2O
0.25 g L-1, glycerol 2 g L-1, kanamycin 50 μg mL-1) at 37°C in 1 L baffled flasks at 165 rpm. Ldi
expression was induced by 1 mM Isopropyl-E-D-1-thiogalactopyranoside at an optical density
(600 nm) around 1. Cultures were further cultivated for 4 to 5 hours at 32°C to an OD (600 nm)
of approx. 3. Biomass was harvested by centrifugation (14000 x g, 20 min, 4°C), frozen in liquid
nitrogen and stored at -80°C.
Activity assay
Enzymatic activity was determined by end-point measurement. Subfractions obtained during
purification were dialyzed against Tris-Cl buffer (80 mM, pH 8.0). The assay was performed in 4
mL glass vials. 300-500 μL of sample were treated with 5 mM DTT. Vials were closed with a
butyl septum and flushed with N2 gas for 3 minutes via needles. After a pre-incubation of 20
minutes at room temperature, substrate (200 mM (R,S)-linalool or geraniol in 2,2,4,4,6,8,8heptamethylnonane, HMN) was added via a needle. Incubation took place at 37°C under mild
shaking. Product formation was determined by gas chromatography with flame ionization
detection (PerkinElmer Auto System XL, Überlingen, Germany). 1 μL sample was injected onto
an Optima-5 column (30 m x 0.32 mm, 0.25 μm film thickness; Macherey-Nagel, Germany) with
hydrogen as carrier gas and the following temperature program: injection port 250°C, detection
port 350°C, initial column temperature 40°C for 2 min, increasing to 100°C at a rate of 4°C min1
, keeping 100°C for 0.1 min, followed by an increase to 320°C at 45°C min-1 and hold for 3 min.
The split ratio was set to 1:9.
Protein purification
E. coli BL21(DE3) pET42-Ldi biomass was suspended in Tris-Cl buffer (80 mM, pH 8.0) and
thawed quickly at room temperature. Cell disintegration was performed by mechanical sheering
using a French Press pressure cell at 8.26 MPa two times. The crude extract was clarified by
ultracentrifugation (150000 x g, 30 min, 4°C) and the supernatant (soluble protein extract) was
- 93 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
further used. All purification steps were performed on an ÄKTA purifier system (GE Healthcare)
at 6°C. All buffers were prepared and pH values adjusted at room temperature (22°C) and filtered
(0.2 μm) prior use. Protein was detected by absorption measurement at 280 nm. SDS-PAGE was
used to analyze purification samples and protein concentrations were determined according to the
method described by Bradford using bovine serum albumin as a calibration standard [32].
Ldi was purified in a three step chromatography approach, including hydrophobic interaction,
anion exchange and size-exclusion chromatography. Soluble protein extract from E. coli
BL21(DE3) pET42-Ldi was prepared in Tris-Cl buffer (80 mM, pH 8.0). The sample was
adjusted to 5 % ammoniumsulfate (v/v) with a saturated ammoniumsulfate solution to aid binding
onto a phenyl sepharose column (dimensions: 8 x 100 mm; CV 5 mL). The column was
equilibrated with Tris-Cl buffer (80 mM, pH 8.0; 5 % (NH4)2SO4). After extensive washing with
binding buffer, a gradual elution of bound protein was performed by applying buffers with
decreasing hydrophobicity: Tris-Cl buffer (80 mM, pH 8.0) - Tris-Cl buffer (20 mM, pH 8.0, 0.1
% Tween20 v/v) - pure water. The change from binding buffer to the first elution buffer was
performed in a linear gradient (0-100 % over 20 mL), while the other buffer changes were
performed in an one-step gradient. Pooled fractions from the peak eluted with 20 mM Tris-Cl
(pH 8.0; 0.1 % Tween20) were concentrated with an AMICON filter unit (10 kDa cut-off,
Merck-Millipore) and dialyzed against NaH2PO4/Na2HPO4 buffer (50 mM, pH 7.0) for at least 12
hours. This fraction was loaded onto a ResourceQ anion exchange column (CV 1 mL),
equilibrated with NaH2PO4/Na2HPO4 buffer (50 mM, pH 7.0). After washing off unbound protein
with binding buffer, bound protein was eluted with NaH2PO4/Na2HPO4 buffer (50 mM, pH 7.0)
containg 1 M NaCl. The flow through was concentrated with an AMICON filter unit prior the
application onto a size-exclusion chromatography column (HiLoad 16/60, Superdex200,
dimensions: 16 x 600 mm), equilibrated with Tris-Cl buffer (20 mM, pH 8.0), at a flow rate of
0.6 mL min-1. Final samples, matching the purity standards were dialyzed against 10 mM Tris-Cl
buffer (pH 8.0) and subsequently concentrated to 10 - 15 mg ml-1 protein for further processing in
crystallography.
- 94 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Crystallization, X-ray data collection, phase determination and refinement
Ldi was crystallized at 4°C under oxic as well as anoxic (97 % N2 : 3 % H2 v/v) conditions using
the sitting / hanging drop vapor diffusion method. Diffraction data were collected at the PX II
beamline at the Swiss Light Source (SLS, Villingen, Switzerland) and processed with XDS [33].
The multi-wavelength anomalous diffraction (MAD) data set was measured at the Hg (L-III)
absorption edge. The mercury positions were identified with SHELXC/D [34], the phases were
calculated with SHARP [35] and improved by solvent flattening. The quality of the resulting
electron density was sufficient for identifying five monomers of the Ldi pentamer but not for
chain tracing. Non-crystallographic symmetry operators and the protein envelope were calculated
with O [36]. For final structure determination we included a native data set into SHARP used the
single isomorphous replacement with anomalous scattering (SIRAS) method and five-fold
averaged the resulting electron density with DM [37] of the CCP4 package [38]. Automated
model building was performed using Buccaneer of the CCP4 package [38]. The model was
manually completed and refined using Coot [39], Refmac5 [40] and Refine in Phenix [41]
including the non-crystallographic symmetry (NCS) and the translation/liberation/screw motion
(TLS) options. All crystals of Ldi were isomorphous and new data had to be refined against the
model. A binding site for an elongated molecule present in all five subunits was assigned as
partly attached precipitant PEG550 MME. Data collection and refinement statistics are
summarized in Table 2. Crystal structures were superimposed using DaliLite [17]. All Figures
were prepared using PyMOL [42]. Atomic coordinates and structure factors of Ldi and Ldi with
substrate are available in the Protein Data Bank (PDB) under the accession numbers xxx and yyy,
respectively.
Construction of amino-exchange mutants
Amino acid-exchange was performed by site-directed mutagenesis, using primers with defined
mismatches (Table 1), the pET42-Ldi plasmid as a template and the Phusion Polymerase
according to the manufacturer manual (Life Technologies, Thermo Scientific Bio). The complete
plasmid was amplified by overlapping primers, ligated and transformed into E. coli. Mutants
were verified by sequencing of the plasmid with standard T7-primers using the BigDye
Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Life Technologies Corporation,
- 95 -
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Carlsbad, USA) with the following program: 95°C for 5 min, 99 cycles at 96°C for 30 sec, 55°C
for 20 sec and 60°C for 4 min. Fragments were analyzed on an ABI Prism 3130xl Genetic
Analyzer (Applied Biosystems Life Technologies Corporation, Carlsbad, USA). Verified Ldi
mutants were grown in 100 ml LB medium in 250 ml flasks and protein expression was induced
by addition of 1 mM IPTG. Soluble protein extract was obtained as described above and used for
evaluation of enzyme activity.
Table 1: Primer for amino acid exchange in the linalool dehydratase/isomerase.
Primer
Sequence
Mutation
pC220_fw
GGGCAGCTCCTTCGAGG
C48S
pC220_rw
CCTCGAAGGAGCTGCCC
pC379_fw
GATGAAGAGCAAGCGGGTC
pC379_rw
GACCCGCTTGCTCTTCATC
pC586_fw
CGGCATCGTCAGCGAGCC
pC586_rw
GGCTCGCTGACGATGCCG
pC613_fw
TTGTCCAGAGCAATTCGGTC
pC613_rw
GACCGAATTGCTCTGGACAA
pH460_fw
GTACAAGGGCTACTGAACC
pH460_rw
GGTTCAGTAGCCCTTGTAC
pTyr65Phe_FW
CATCAAGTTTTCGATCG
pTyr65Phe_RW
CGATCGAAAACTTGATG
pPhe69Tyr_FW
CGATCGCCTACTATGCG
pPhe69Tyr_RW
CGCATAGTAGGCGATCG
- 96 -
C101S
C170S
C179S
H128Y
Y65F
F69Y
Chapter IV - X-ray structure of linalool dehydratase/isomerase
Table 2: Crystallographic data.
Crystal
Data collection
Space group
Cell dimensions
a, b, c (Å)
DEJ (º)
ldi3
P212121
90.0, 105.5,
220.3
90, 90, 90
Wavelength
Resolution (Å)
0.99996
50 - 2.2
(2.3 - 2.2)
Rsym
10.9 (81.3)
I/VI
10.4 (1.9)
Completeness (%)
99.6 (99.6)
Redundancy
3.5 (3.4)
Refinement
Resolution (Å)
No. reflections
Rwork/ Rfree
No. atoms
Protein
Ligand/ion
Water
B-factors
Protein
Ligand/ion
Water
R.m.s deviations
Bond lengths
(Å)
Bond angles (º)
Ramachandran
favored/
disallowed (%)
ldi13
ldi31
ldi42
P212121
P212121
99.9, 106.7,
223.1
90, 90, 90
99.3, 106.3,
221.3
90, 90, 90
1.00002
0.97928
50 – 1.9
(2.0 – 1.9)
50 – 2.5
(2.6 – 2.5)
9.8 (71.0)
14.0 (131.4)
10.5 (2.9)
13.7 (2.4)
99.9 (99.8)
99.8 (99.2)
5.9 (5.9)
11.7 (11.7)
48.3 – 2.2
(2.23 –
2.20)
117597
17.5 (28.3)
20.9 (32.5)
49.5 – 1.91
(1.93 – 1.91)
49.1 – 2.5
(2.52 – 2.50)
183305
19.0 (27.5)
22.5 (32.7)
81564
17.6 (36.4)
22.5 (40.8)
14590
371
14602
580
14602
239
121
38.4
36.6
35.7
35.0
60.3
107.1
50.1
0.003
0.011
0.005
0.80
97.4 / 0.1
1.17
97.7 / 0.2
0.83
96.8 / 0.2
P212121
99.9, 106.1, 220.4
90, 90, 90
Peak
Inflection
1.00724 1.00939
50 - 2.4
50 - 2.7
(2.5 (2.8 - 2.7)
2.4)
13.1
12.3 (54.0)
(68.7)
7.9
8.2 (2.5)
(2.7)
99.8
98.7 (98.8)
(99.8)
4.6
3.3 (3.2)
(4.7)
- 97 -
Remote
0.99999
50 - 3.0
(3.1 3.0)
14.9
(53.9)
7.3 (2.7)
98.9
(98.5)
3.3 (3.4)
Chapter IV - X-ray structure of linalool dehydratase/isomerase
- 98 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Chapter V
A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Robert Marmulla1, Frauke Lüddeke1, Manuel Liebeke2 and Jens Harder1
1
Dept. of Microbiology, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359
Bremen
*
corresponding author: jharder@mpi-bremen.de
2
Dept. of Symbiosis, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359
Bremen
Manuscript in preparation
- 99 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Abstract
The betaproteobacterium Castellaniella defragrans 65Phen grows on various a-, mono- and
bicyclic monoterpenes as sole carbon and energy source under denitrifying conditions.
Mineralization of the acyclic monoterpene E-myrcene is initiated via hydration to (S)-linalool and
isomerization to geraniol by the linalool dehydratase/isomerase (Ldi). In this study, we report
growth on (R,S)-linalool by a deletion mutant ('ldi) of C. defragrans 65Phen that does not grow
on E-myrcene or geraniol. The metabolic fate of (R,S)-linalool in this mutant strain was observed
in cell-free protein extracts. The formation of the monocyclic monoterpenes D-terpinene and
terpinolene coincided with a decrease in (R,S)-linalool. This biotransformation required the
presence of ATP and divalent metal ions (Mg2+, Mn2+). We suggest an ATP-dependent activation
of (R,S)-linalool, followed by a subsequent intramolecular ring-closure to yield a monocyclic
monoterpene. Together with the absence of the acyclic terpene utilization pathway in the C.
defragrans 65Phen genome, our observations suggest a linalool degradation via the monocyclic
monoterpene degradation pathway.
Introduction
Castellaniella defragrans 65Phen was isolated on D-phellandrene under denitrifying conditions
(Foss, et al. 1998, Kämpfer, et al. 2006). Due to its capacity to use a broad range of acyclic and
cyclic monoterpenes under denitrifying conditions it became a model organism to investigate
anaerobic monoterpene transformations in bacteria (Marmulla and Harder 2014, Petasch, et al.
2014). The bifunctional enzyme linalool dehydratase/isomerase (Ldi) is the first enzyme in a Emyrcene biotransformation pathway. This enzyme hydrates E-myrcene stereospecifically to (S)linalool and further isomerizes (S)-linalool to geraniol (Brodkorb, et al. 2010, Lüddeke and
Harder 2011). The oxidation of geraniol to geranic acid was first detected in cultures (Heyen and
Harder 2000) and the responsible enzymes were characterized as geraniol dehydrogenase and
geranial dehydrogenase (Lüddeke, et al. 2012b). Degradation of geranic acid derived from the
oxidation of geraniol or citronellol had been investigated in various Pseudomonas species to
proceed via the acyclic terpene utilization pathway (Atu) (Förster-Fromme and Jendrossek 2010).
- 100 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
None of the genes coding for the enzymes of the Atu pathway was identified in the genome of C.
defragrans 65Phen (Petasch, et al. 2014). The deletion mutant strain C. defragrans 65Phen 'ldi
that lacked the linalool dehydratase/isomerase gene (ldi) was not able to grow on E-myrcene or
geraniol, but grew on monocyclic monoterpenes (e.g. D-phellandrene and limonene), indicating a
vital role of Ldi in the metabolism of acyclic monoterpenes (Lüddeke, et al. 2012a). We now
observed growth of the strains C. defragrans 65Phen and 65Phen 'ldi on the acyclic
monoterpene alcohol (R,S)-linalool as sole carbon and energy source under denitrifying
conditions. To characterize this novel linalool metabolism, enzyme activity measurements and
metabolite identification by GC and GC-MS were performed which supported the presence of a
terpene cyclase reaction.
Material and Methods
Bacterial strains and cultivation conditions
C. defragrans 65Phen wildtype and 'ldi strains were cultivated under anaerobic, denitrifying
conditions in artificial fresh water (AFW) medium. Medium was prepared as described (Foss, et
al. 1998) with replacement of the carbonate buffer by 10 mM Na2HPO4/NaH2PO4 and omission
of vitamins. The headspace contained nitrogen gas only. Monoterpenes were directly applied to
the water phase below toxic concentrations. Alternatively, they were provided in 2,2,4,4,6,8,8heptamethylnonane as carrier phase. Growth experiments were performed in 10 mL medium that
was shaken (90 rpm) at 28°C. Growth was determined by measurement of the optical density at
600 nm. Biomass for enzyme assays was obtained from 2 L cultures grown on 1.5 mM (R,S)linalool and 10 mM nitrate to the late exponential phase. The harvest occurred by centrifugation
at 16000 x g for 25 min at 4°C. For storage, biomass was frozen in liquid nitrogen and stored at 80°C.
Enzyme assay and product analysis
Monoterpenes were purchased from Sigma-Aldrich (Germany) and were of 95 to 97 % purity.
Enzyme assays were performed in 4 mL glass vials closed with a butyl-rubber septum. The setup
was prepared under aerobic conditions by providing either 1 mL cell lysate or soluble, protein
extract (2.1 ± 0.7 mg mL-1 protein in individual setups) and subsequent addition of supplements.
- 101 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
The followed supplements were tested (concentration are given as final concentrations in the
assays): DTT (1-2 mM), ATP (5-20 mM), MgSO4/MnSO4 (5 mM each) and EDTA (20 mM).
(R,S)-linalool and (R)-linalool were used as substrates. Individual assays received either 10 mM
or 58 mM substrate. Final assay volume was 1.2 mL. The vials were closed and flushed with
nitrogen gas for 2 minutes through a needle. Cell lysate was prepared by resuspending biomass in
Na2HPO4/NaH2PO4 buffer (50 mM, pH 7.0) and one passage through a One-Shot cell disruptor
(Constant Systems Ltd., UK) at 1.7 GPa. Soluble protein extract was obtained by
ultracentrifugation (150000 x g for 30 min at 4°C). The assay was incubated at 28°C under mild
shaking for 12 to 14 hours. Samples were transferred to 1.5 mL plastic tubes and mixed with 200
μL n-hexane by vortexing for 30 seconds. After 10 minutes incubation, the samples were mixed
again and centrifuged (13000 x g, 10 min) to achieve phase separation. The clear hexane phase
was recovered and transferred into a GC-vial. A 1 μL-sample was analyzed by gas
chromatography with flame ionization detection (Shimadzu GC-14A, Shimadzu Corporation) on
a Hydrodex-E-6TBDM column (25 m x 0.25 mm, Macherey-Nagel, Germany) with the following
temperature program: injection port 200°C, detection port 250°C, initial column temperature
60°C for 1 min, increasing to 130°C at a rate of 5°C min-1, keeping 130°C for 0.5 min, followed
by an increase to 230°C at 20°C min-1 and hold for 4 min. Quantification was performed from a
calibration with authentic standards. Samples showing product formation were further
characterized by GC-MS analyses. The analysis was performed on an Agilent 7890B gas
chromatograph connected to an Agilent 5977A MSD. Samples were injected in splitless mode
with an Agilent 7693 autosampler injector into deactivated splitless liners. Separation was
performed on an Agilent 30 m DB5-MS column with a 10 m DuraGuard column, using the
following temperature program: 40°C for 1 min, increasing to 160°C at a rate of 4°C min-1,
followed by 30°C min-1 to 280°C and a hold for 1 min. Metabolites were assigned using the NIST
2.0 Library and verified by injection of pure compounds under same conditions.
- 102 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Results and Discussion
C. defragrans 65Phen cells pregrown on D-phellandrene required initially weeks of incubation to
initiate growth on (R,S)-linalool. These adapted cells demonstrated in transfer cultures the
presence of two phenotypes. While growing on 800 μM (R,S)-linalool, one phenotype favored the
formation of E-myrcene: (S)-linalool was consumed and 511 μM E-myrcene accumulated.
Biomass increase and nitrate reduction were small (Table 1). The formation of E-myrcene, likely
by the Ldi, can be interpreted as a defense reaction of the organism against the toxic alcohols, as
they have a higher solubility than monoterpenes and are more toxic. The Ldi converts only (S)linalool (Lüddeke and Harder 2011), thus (R)-linalool is still found in these cultures. The second
phenotype showed growth and consumed nearly all nitrate (9.7 mM) and both linalool isomers
completely. Only minor amounts of E-myrcene were formed (Table 1). We interpret the
consumption of both linalool isomers as indication for a novel pathway.
Table 1: Physiological activities of C. defragrans incubated with 800 μM (R,S)-linalool (8 mM in HMN)
and 10 mM nitrate for 480 hours at 28°C. The inoculum was 1 % (v/v) of a linalool-adapted culture. All
monoterpene concentrations are referred to the aqueous phase. Variability of OD determination was ±
0.02.
Inoculum
increase
[g/L]
Nitrite
formation
[mM]
Nitrate
consumpti
on [mM]
(R)linalool
[μM]
(S)linalool
[μM]
Emyrcene
[μM]
0.001
0.00
0.00
0.0
400
400
0
0.132
0.12
0.00
9.7
0
0
76
0.038
0.03
0.06
2.7
323
0.8
511
Growth
Biomass
['ODmax]
--65Phen
Next, we tested whether the deletion strain 65Phen 'ldi showed growth on (R,S)-linalool. After
growth was established, transfer cultures grew on (R,S)-linalool, but not on E-myrcene, geraniol
or geranic acid (Fig. 1). The growth on linalool and the disappearance of both isomers confirmed
the presence of a second metabolic pathway for linalool-usage. The biomass yield is reduced in
comparison to biomass yields during growth on acetate or D-phellandrene and nitrate. Cultures
- 103 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
grown on (R,S)-linalool or D-phellandrene reached maximal optical densities of 0.13 or 0.35,
respectively (Lüddeke, et al. 2012a). This may be caused by a lower energy conservation
efficiency due to membrane damage by linalool or energy consumption in either a defense
reaction (export) or an activation of linalool.
Figure 1: Growth curve of C. defragrans 65Phen 'ldi on 750 μM monoterpene (myrcene ♦, linalool ■,
geraniol ▲, geranic acid ●) and 10 mM nitrate. Inoculum was 4 % (v/v) of a linalool-adapted culture.
To identify possible intermediates in the linalool degradation, we conducted various enzyme
assay experiments with cell lysates and cytosolic protein extracts together with (R,S)-linalool and
subsequent gas chromatography analysis. Two abundant substances were formed and identified
as D-terpinene and terpinolene by retention time comparison to authentic standards via GC and
GC-MS analyses (Fig. 2).
- 104 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Figure 2: Chromatograms of hexane extracts analyzed by gas chromatography (A) and mass spectra of
identified peaks (B, C). Assays differed by the absence (dashed line, control) and presence of 5 mM ATP
and 1 mM DTT (solid lines). Retention times for D-terpinene (1) and terpinolene (2) were 3.6 and 4.75
minutes, respectively. Red spectrum indicates measurement and blue spectrum indicates reference
spectrum in NIST library.
The formation of both monocyclic monoterpenes was stimulated by addition of divalent metal
ions and adenosintriphosphate (ATP). In initial experiments, 5 mM ATP and 1 mM dithiothreitol
(DTT) enabled the cell-free protein extract to form 3.1 mM D-terpinene and 0.35 mM terpinolene
- 105 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
(Fig. 2). Dialyzed membrane-free soluble extracts required ATP for the product formation, with a
shift in substrate formation: more terpinolene than D-terpinene was formed. Addition of DTT had
a stimulating effect on the reaction of the protein extract. The chelating agent EDTA (20 mM)
suppressed the formation completely. ATP concentrations of 10 and 20 mM caused a
monoterpene formation of 0.16 and 0.08 mM D-terpinene and 1.71 and 3.23 mM terpinolene,
respectively. Both (R)- and (R,S)-linalool were transformed under those conditions.
These results reveal an ATP-dependent activation in the biotransformation of (R,S)-linalool to a
cyclic monoterpene intermediate in C. defragrans 65Phen (Fig. 3). To our knowledge, an ATPdependent pyrophosphorylation of (R,S)-linalool has not been reported. We suggest an activation
to linalyl diphosphate, a common intermediate in monoterpene biosynthesis pathways (Davis and
Croteau 2000).
Figure 3: Proposed reaction sequence of linalool activation and cyclization to terpinolene. Shown is the
formation of linalyl diphosphate from linalool and ATP and subsequent intramolecular rearrangements
known from monoterpene biosynthesis.
In monoterpene biosynthesis in plants, the phosphate moiety of linalyl diphosphate is leaving,
resulting in a linalyl cation intermediate that can undergo cyclization to the terpinyl cation, the
precursor molecule of all cyclic monoterpenes. The cyclization is catalyzed by terpene cyclases
which are divided into class I and class II enzymes. While class I cyclases initiate carbocation
formation by the release of the pyrophosphate moiety, class II cyclases protonate a carbon-carbon
double bond in the initial step. Monoterpene cyclases belong to the first class, having a common
DDXXD-motif and rely on Mg2+ and/or Mn2+ as a divalent metal cofactor (Gao, et al. 2012,
Wendt and Schulz 1998). Even though these enzymes share common motif sequences and a Dhelical bundle as tertiary structure, they are quite dissimilar with respect to their overall sequence
(Yamada, et al. 2015). Homology search in the genome of C. defragrans 65Phen
- 106 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
(NZ_HG916765) did not reveal a potential monoterpene cyclase. As monoterpene synthases
accept only activated substrates (diphosphates of geraniol, linalool or nerol), it is likely that the
activation and further cyclization of linalool are catalyzed by two independent enzymes in C.
defragrans 65Phen; namely a linalool-specific diphosphate transferase and a monoterpene
cyclase.
The formation of more than one product is well known for monoterpene cyclases. Depending on
the amino acid sequence in the active center, terpene synthases can be specific for a product or
rather promiscuous (Christianson 2008). A change of a single amino acid can lead to a drastic
change in monoterpene composition (Köllner, et al. 2004). Also, in abiotic catalysis D-terpinene
and terpinolene together with eucalyptol were reported to be major products of the non-enzymatic
cyclization of linalool in a self-assembled organic cavity (Zhang and Tiefenbacher 2015). This
indicates a favored formation of D-terpinene and terpinolene from linalool as general principle.
The microbial degradation of the formed monocyclic monoterpenes may proceed via an oxygenindependent limonene dehydrogenase (ctmABCDEFG) and a suite of ring opening reactions
(mrcABCDEFGH) (Petasch, et al. 2014).
Conclusion
This study revealed the presence of a pathway for the synthesis of monocyclic monoterpenes
from the acyclic (R,S)-linalool in the monoterpene metabolism of C. defragrans 65Phen. In an
ATP-dependent reaction, (R,S)-linalool was transformed to D-terpinene and terpinolene. This
pathway may involve so far unknown donor:linalool diphosphate transferase and monoterpene
cyclase activities.
- 107 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Competing interests
The authors declare that they have no competing interests.
Authors´ contribution
FL performed initial experiments. RM performed experiments and GC measurements. ML
performed the GC-MS analysis. RM, ML and JH drafted the manuscript. All authors read and
approved the final manuscript.
Acknowledgments
This study was financed by the Max Planck Society.
- 108 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
References
Brodkorb D, Gottschall M, Marmulla R et al. Linalool dehydratase-isomerase, a bifunctional enzyme in
the anaerobic degradation of monoterpenes. Journal of Biological Chemistry 2010;285: 3043642.
Christianson DW. Unearthing the roots of the terpenome. Current Opinion in Chemical Biology 2008;12:
141-50.
Davis EM, Croteau R. Cyclization enzymes in the biosynthesis of monoterpenes, sesquiterpenes, and
diterpenes. In: Leeper FJ, Vederas JC (eds.) Biosynthesis volume 209: Springer Berlin Heidelberg,
2000, 53-95.
Förster-Fromme K, Jendrossek D. Catabolism of citronellol and related acyclic terpenoids in
pseudomonads. Applied Microbiology and Biotechnology 2010;87: 859-69.
Foss S, Heyen U, Harder J. Alcaligenes defragrans sp. nov., description of four strains isolated on
alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alpha-phellandrene) and
nitrate. Systematic and Applied Microbiology 1998;21: 237-44.
Gao Y, Honzatko RB, Peters RJ. Terpenoid synthase structures: a so far incomplete view of complex
catalysis. Natural Product Reports 2012;29: 1153-75.
Heyen U, Harder J. Geranic acid formation, an initial reaction of anaerobic monoterpene metabolism in
denitrifying Alcaligenes defragrans. Applied and Environmental Microbiology 2000;66: 3004-9.
Kämpfer P, Denger K, Cook AM et al. Castellaniella gen. nov., to accommodate the phylogenetic lineage
of Alcaligenes defragrans, and proposal of Castellaniella defragrans gen. nov., comb. nov and
Castellaniella denitrificans sp nov. International Journal of Systematic and Evolutionary
Microbiology 2006;56: 815-9.
Köllner TG, Schnee C, Gershenzon J et al. The variability of sesquiterpenes emitted from two Zea mays
cultivars is controlled by allelic variation of two terpene synthase genes encoding stereoselective
multiple product Enzymes. The Plant Cell 2004;16: 1115-31.
Lüddeke F, Dikfidan A, Harder J. Physiology of deletion mutants in the anaerobic beta-myrcene
degradation pathway in Castellaniella defragrans. BMC Microbiology 2012a;12.
Lüddeke F, Harder J. Enantiospecific (S)-(+)-linalool formation from beta-myrcene by linalool
dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of Biosciences
2011;66: 409-12.
Lüddeke F, Wülfing A, Timke M et al. Geraniol and geranial dehydrogenases induced in anaerobic
monoterpene degradation by Castellaniella defragrans. Applied and Environmental Microbiology
2012b;78: 2128-2136.
- 109 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
Marmulla R, Harder J. Microbial monoterpene transformations – A review. Frontiers in Microbiology
2014;5.
Petasch J, Disch E-M, Markert S et al. The oxygen-independent metabolism of cyclic monoterpenes in
Castellaniella defragrans 65Phen. BMC Microbiology 2014;14: 164.
Wendt KU, Schulz GE. Isoprenoid biosynthesis: manifold chemistry catalyzed by similar enzymes.
Structure 1998;6: 127-33.
Yamada Y, Kuzuyama T, Komatsu M et al. Terpene synthases are widely distributed in bacteria.
Proceedings of the National Academy of Science USA 2015;112: 857-62.
Zhang Q, Tiefenbacher K. Terpene cyclization catalysed inside a self-assembled cavity. Nature Chemistry
2015;7: 197-202.
- 110 -
Chapter V - A novel metabolic pathway for linalool in Castellaniella defragrans 65Phen
- 111 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Chapter VI
Linalool isomerase, a membrane-anchored enzyme in the anaerobic monoterpene
degradation in Thauera linaloolentis 47Lol
Robert Marmulla1, Barbara Šafarić1, Stephanie Markert2, Thomas Schweder2, Jens Harder1*
1
Dept. of Microbiology, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359
Bremen
*
corresponding author: jharder@mpi-bremen.de
2
Institute for Pharmacy, Dept. of Pharmaceutical Biotechnology, University of Greifswald,
Felix-Hausdorff-Str. 3, D-17487 Greifswald
Research article submitted to BioMed Central (BMC) Biochemistry
- 112 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Abstract
Background: Thauera linaloolentis 47Lol uses the tertiary monoterpene alcohol (R,S)-linalool as
sole carbon and energy source under denitrifying conditions. The conversion of linalool to
geraniol had been observed in carbon-excess cultures, suggesting the presence of a 3,1-hydroxyl'1-'2-mutase (linalool isomerase) as responsible enzyme. To date, only a single enzyme
catalyzing such a reaction is described: the linalool dehydratase/isomerase (Ldi) from
Castellaniella defragrans 65Phen acting only on (S)-linalool.
Results: The linalool isomerase activity was located in the inner membrane. It was enriched by
subcellular fractionation and sucrose gradient centrifugation. MALDI-ToF MS analysis of the
enriched protein identified the corresponding gene named lis that codes for the protein in the
strain with the highest similarity to the Ldi. Linalool isomerase is predicted to have four
transmembrane helices at the N-terminal domain and a cytosolic domain. Enzyme activity
required a reductant for activation. A specific activity of 3.42 ± 0.28 nkat mg * protein-1 and a kM
value of 455 ± 124 μM were determined for the thermodynamically favored isomerization of
geraniol to both linalool isomers at optimal conditions of pH 8 and 35°C.
Conclusion: The linalool isomerase from T. linaloolentis 47Lol represents a second member of
the enzyme class 5.4.4.4, next to the linalool dehydratase/isomerase from C. defragrans 65Phen.
Besides considerable amino acid sequence similarity both enzymes share common characteristics
with respect to substrate affinity, pH and temperature optima, but differ in the turnover of linalool
isomers.
Keywords: Linalool, geraniol, Thauera, isomerase, allyl alcohol, monoterpene
- 113 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Introduction
Monoterpenes (C10H16), naturally occurring hydrocarbons, are the main constituents of essential
oils and belong to the diverse group of terpenoids, of which more than 50000 structures are
known to date [1, 2]. They are produced as secondary plant metabolites and serve diverse
functions like signaling, attraction/repellence of pollinators and insects, thermotolerance and are
involved in allelopathy [3, 4]. Atmospheric monoterpene emission from plants was estimated to
be 127 Tg C yr-1 [5], with half-lives of minutes to hours in the atmosphere due to their proneness
to chemical and photooxidative reactions. Monoterpenes enter soils by precipitation from the
atmosphere, excretion from roots and by leaf fall [6-9]. The hydrophobic character of
monoterpenes causes cell toxicity, mainly by accumulation into and destabilization of the cell
membranes [10]. Below toxic concentrations, microorganisms can use monoterpenes as carbon
and energy source for growth. Several bacteria have been described to transform monoterpenes in
the presence of oxygen as a cosubstrate applying mono- and dioxygenases [11], but also other
biotransformations are described [12]. Linalool, a tertiary monoterpene alcohol (Fig. 1), is the
main component of essential oils in lavender and coriander. Its chemical structure prevents a
direct oxidation of the hydroxyl group. In order to enable its biological degradation, an additional
functionalization or isomerization is needed. Linalool is oxidized to 8-hydroxylinalool under
aerobic conditions [13, 14]. Other strategies are used in the absence of oxygen. Foß and Harder
reported the formation of the primary alcohol geraniol in linalool-grown cultures of Thauera
linaloolentis 47Lol (Fig. 1). This betaproteobacterium was isolated on linalool as sole carbon end
energy source under denitrifying conditions. A 3,1-hydroxyl-'1-'2-mutase was proposed as novel
enzymatic function initializing the mineralization of linalool [15, 16].
Figure 1: Linalool isomerase catalyzes the reversible reaction from linalool to geraniol.
- 114 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
A similar reaction was described for the bifunctional enzyme linalool dehydratase/isomerase
from Castellaniella defragrans 65Phen. The enzyme catalyzes the reversible hydration of Emyrcene to (S)-linalool and its isomerization to geraniol. It is an oxygen-sensitive, periplasmatic
protein of 43 kDa including a signal peptide for export [17, 18]. The tertiary alcohol 2-methyl-3buten-2-ol (232-MB) may also be transformed by enzyme-catalyzed isomerization reactions. 232MB is a metabolite in bacterial degradation of fuel oxygenates. Its mineralization via an initial
isomerization to 3-methyl-2-buten-1-ol (prenol) in Aquincola tertiaricarbonis L108 and
Methylibium petroleiphilum PM1 was proposed [19]. Malone et al. proposed a 2-methyl-3-buten2-ol isomerase to be involved in Pseudomonas putida MB-1, isolated on 232-MB as sole carbon
source. Prenol was suggested as product which is further oxidized and metabolized [20].
However, the corresponding enzymes were so far not characterized. For intramolecular hydroxylgroup transfer (EC 5.4.4.x), only seven different enzyme activities are described: (hydroxyamino)
benzene mutase (EC 5.4.4.1), isochorismate synthase (EC 5.4.4.2), 3-(hydroxyamino) phenol
mutase (EC 5.4.4.3), geraniol isomerase (EC 5.4.4.4), 9,12-octadecadienoate 8-hydroperoxide
8R-isomerase (EC 5.4.4.5), 9,12-octadecadienoate 8-hydroperoxide 8S-isomerase (EC 5.4.4.6)
and hydroxyperoxy icosatetraenoate isomerase (EC 5.4.4.7).
We report the enrichment of the linalool isomerase activity in protein fractions of T. linaloolentis
47Lol and the kinetic properties of the enzyme. A corresponding gene was identified in the draft
genome. We suggest to place the linalool isomerase of T. linaloolentis 47Lol as a new member in
the enzyme family of intramolecular hydroxyl group transferases (EC 5.4.4.x) with the EC
number 5.4.4.4 next to the geraniol isomerase function of the linalool dehydratase/isomerase
from C. defragrans 65Phen.
- 115 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Results and Discussion
Identification of a candidate protein for linalool isomerase
The linalool dehydratase/isomerase (Ldi, NCBI:CBW30776) was used in similarity searches to
identify a putative linalool isomerase protein in a draft genome of T. linaloolentis 47Lol. One
protein showed a considerable similarity with an overall identity of 20 % (positives 33 %, Evalue 3E-10, NCBI:ENO87364). The corresponding gene is isolated from the adjacent genes (>
150 bp) and encodes a protein of 644 amino acids with a calculated molecular weight of 71.8
kDa, an isoelectric point of 6.06 and a hydrophobicity of -0.115 (GRAVY) [21]. No signal
peptide was predicted by the SignalP software. For the N-terminus, four transmembrane domains
within the first 139 amino acids were predicted together with a localization in the inner
membrane with the C-terminal protein fold in the cytoplasm. Conserved domains as described in
Pfam were not present. The specific hydrophobicity values (GRAVY) were 0.94 and -0.406 for
the N- and C-terminal parts of the protein (amino acids 1 – 139 and 140 – 644, respectively). The
similarity to the Ldi was restricted to the C-terminal domain.
Enrichment of the linalool isomerase activity (Lis)
The Lis enzyme activity was detected in crude cell-free protein extracts containing also
membrane fragments after application of high pressure cell disruption. Cell disintegration by
osmosis or ultra sonification retained the activity in the membrane fraction. Supported by the
large cytoplasmatic domain of the candidate protein, we attempted the isolation as soluble
enzyme from dialyzed crude extracts. The pH was increased to 9.5 to eventually increase the
anionic character of the enzyme. The enzyme activity did not bind to a DEAE column. After
ammonium sulfate addition (10 % v/v of a saturated solution), the enzyme activity was retained
on a Phenyl-Sepharose column and eluted with pure water. A strong binding to hydrophobic
columns has also been observed for the Ldi [17] and other monoterpene-transforming enzymes
[22]. A size determination of the eluted fraction yielded a molecular weight of over 600 kDa, as
native protein in phosphate buffer as well as in the presence of urea as denaturant. Visualization
of the proteins on SDS-PAGE revealed the presence of many proteins, including abundant
proteins between 60 and 70 kDa and around 33 kDa (data not shown).
The large size together with the predicted membrane association of the Lis candidate gene
- 116 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
suggested as alternative purification approach a subcellular fractionation including a sucrosegradient centrifugation for the identification of membrane-associated proteins. First, the outer
membrane and periplasmatic proteins were removed by spheroplast formation. After
disintegration by high pressure release, the inner membrane (IM) fraction was separated from the
cytosolic soluble fraction by ultracentrifugation (158 Svedberg). Total Lis activity was five times
higher in the membrane fraction than in the soluble fraction (15.8 and 3.3 nkat, respectively) (Fig.
2 and Tab. 1).
Figure 2: SDS-PAGE of different fractions during the inner membrane preparation from spheroplasts.
(F1) spheroplasts, (F2) spheroplasts after cell disintegration, (F3) outer membrane and periplasmatic
protein fraction, (F4) crude extract after spheroplast disintegration, (F5) unbroken spheroplast and cell
debris, (F6) cytoplasmatic, soluble protein fraction, (F7) inner membrane fraction.
- 117 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Table 1: Enrichment of Lis activity from spheroplasts of T. linaloolentis 47Lol. (F1) spheroplasts, (F2)
lysed spheroplasts, (F3) outer membrane and periplasmatic proteins, (F4) crude extract from spheroplasts,
(F5) unbroken spheroplast and cell debris, (F6) cytoplasmatic, soluble proteins, (F7) inner membrane
fraction. *F4 and F5 are derived from F2. F6 and F7 are derived from F4.
Sample
Protein
[mg]
Activity [pkat]
Specific
activity
[pkat mg-1]
Relative
specific
activity
Protein yield
[%]*
F1
71.9
7295
101
F2
79.0
7800
99
1.0
100.0
F3
67.6
3060
45
0.5
F4
51.8
24984
482
4.9
65.6
F5
29.1
513
18
0.2
36.8
F6
38.0
3283
86
0.9
73.3
F7
21.4
15848
741
7.5
41.3
Both fractions were separated on sucrose gradients. The Lis activity of the inner membrane
fraction (Fig. 3 and Tab. 2) was concentrated in two fractions with a sucrose density between
1.17 and 1.22 g mL-1 (IM4 and IM5).
- 118 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure 3: SDS-PAGE of sucrose gradient fractions after separation of the inner membrane (IM) fraction
from spheroplast disintegration. (F7) inner membrane fraction from spheroplast disintegration, (IM 1) (IM 7) fractions 1-7 (top-to-bottom) of sucrose gradient.
Table 2: Enrichment of Lis activity by sucrose gradient centrifugation from inner membrane pellets (F7)
obtained from spheroplasts of T. linaloolentis 47Lol. 1 mL fractions from top to bottom are shown.
Activity [pkat]
Specific
activity
[pkat mg-1]
Relative
specific
activity
Protein yield
[%]
6.74
5925
879
1
100.0
IM 1
0.20
139
697
0.8
3.0
IM 2
1.06
238
225
0.3
15.7
IM 3
1.44
870
604
0.7
21.4
IM 4
1.92
3085
1607
1.8
28.5
IM 5
3.28
3513
1071
1.2
48.7
IM 6
1.44
1050
729
0.8
21.4
IM 7
0.02
0
0
0.0
0.3
Sample
Protein
[mg]
Applied sample
(F7)
- 119 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
For the soluble protein fraction (SP) (Fig. 4 and Tab. 3), the Lis activity was also concentrated in
the fractions with this sucrose content (SP4 and SP5). Protein gels revealed an enrichment of
proteins with a molecular size between 60 to 70 kDa and around 32 kDa for the active fractions
from IM and SP.
Figure 4: SDS-PAGE of sucrose gradient fractions after separation of the cytoplasmatic, soluble protein
fraction from spheroplast disintegration. (F6) Soluble protein fraction from spheroplast disintegration,
(SP 1) - (SP 7) fractions 1-7 (top-to-bottom) of sucrose gradient.
- 120 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Table 3: Enrichment of Lis activity by sucrose gradient centrifugation from cytoplasmatic soluble protein
fraction (F6) obtained from spheroplasts of T. linaloolentis 47Lol. 1 mL fractions from top to bottom are
shown.
Sample
Protein
[mg]
Activity [pkat]
Specific
activity
[pkat mg-1]
Relative
specific
activity
Protein yield
[%]
Applied sample
(F6)
9.2
1863
202
1.0
100.0
SP 1
0.6
14
24
0.1
6.1
SP 2
1.5
16
11
0.1
16.7
SP 3
3.1
165
54
0.3
33.2
SP 4
2.7
273
100
0.5
29.5
SP 5
1.2
141
114
0.6
13.3
SP 6
0.3
13
47
0.2
3.0
SP 7
0.3
0
0
0.0
2.8
The Lis activity was pelleted from the active fractions (SP4 and SP5) by decrease of the sucrose
concentration and ultracentrifugation. SDS-PAGE characterized the dissolved pellet as
containing an approx. 60 kDa protein as most abundant protein (Fig. 5 and Tab. 4). This
purification stage and was used for enzyme kinetics.
- 121 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure 5: SDS-PAGE of Lis activity enrichment by spheroplast disintegration and sucrose gradient
centrifugation. (F1) spheroplasts, (F2) spheroplasts after cell disintegration, (F4) crude extract after
spheroplast disintegration, (F5) cytoplasmatic soluble protein fraction, (SP 4/5) fractions 4 and 5 from
sucrose gradient centrifugation, (Sucrose supernatant) supernatant of second ultracentrifugation, (Sucrose
pellet) protein pellet of second ultracentrifugation.
Table 4: Enrichment of Lis activity from spheroplasts of T. linaloolentis 47Lol by sucrose gradient
centrifugation. (F1) spheroplasts, (F2) lysed spheroplasts, (F4) crude extract, (F6) cytoplasmatic, soluble
protein fraction, (SP 4/5) fractions 4 and 5 from sucrose gradient, (Sucrose supernatant) supernatant after
second ultracentrifugation, (Sucrose pellet) protein pellet after second ultracentrifugation.
Sample
Protein
[mg]
Activity [pkat]
Specific
activity
[pkat mg-1]
Relative
specific
activity
Protein yield
[%]
F1
40.8
1408
35
F2
111.7
3130
28
1
100.0
F4
94.7
13572
143
5
84.8
F6
58.8
3440
59
2
52.6
SP 4/5
3.7
745
200
7
3.3
Sucrose
supernatant
-
-
-
-
-
Sucrose pellet
1.9
323
171
6
1.7
- 122 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
The applied ultracentrifugation pelleted particles from the soluble fraction with a Svedberg
constant of above 158 S. High pressure cell disintegration produces a number of small membrane
fragments and vesicles. Small vesicles (microsomes) require a sedimentation coefficient between
100-10000 S [23] to be pelleted. We conclude from our observations that the Lis activity is
located on a protein associated with the inner membrane. Only a minor fraction of the total
enzyme activity, present as small membrane-protein aggregates, was enriched in the fraction that
is traditionally considered to contain only cytosolic proteins.
The protein gels showed consistently protein(s) between 60 and 70 kDa in fractions with Lis
activity. MALDI-ToF mass spectroscopy identified the protein band at approx. 60 kDa from
several gels as NCBI:ENO87364, the protein that was predicted to be a candidate for the linalool
isomerase. The discrepancy between the calculated molecular weight of 71.8 kDa and the
observed weight of between 60 and 70 kDa is also likely a result of the hydrophobic nature of the
protein: hydrophobic, including membrane proteins tend to bind more SDS and show a faster
migration in denaturing gel electrophoreses [24].
Further purification of the Lis activity was not successful, as a detergent-mediated release of the
protein from the membrane was impaired by irreversible inhibitory effects on the enzyme
activity. Detergents aid solubility of membrane proteins but also have an adverse effect on
stability and functionality of proteins [25, 26]. Terpenoid synthases were shown to have a rather
hydrophobic core shielding the catalytic center from surrounding bulk solvent [27]. Also the Lis
enzyme may have a hydrophobic protein domain sensitive to detergents that change the
conformation into a non-active state.
Heterologous gene expression
Expression of the native lis gene in E. coli BL21(DE3) yielded an induced protein band around
60 kDa (Fig. 6). No inclusion bodies were observed. The expressed protein was located in the
membrane fraction, but Lis activity was not observed in crude cell extracts or in cell-free protein
fractions. Expression of a N-terminally truncated version of the linalool isomerase, representing
the cytosolic part of the enzyme only, yielded a soluble protein that also did not show activity.
The lis gene did not have rare codons. We assume that the folding in E. coli was incorrect.
- 123 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure 6: SDS-PAGE after heterologous expression of the lis gene in E. coli BL21(DE3) pET42-Lis. (1)
Cells from induced culture, (2) crude protein extract after cell disintegration and (3) soluble protein
fraction after ultracentrifugation.
Characterization of linalool isomerase activity
The Lis activity was enriched in the absence of a reductant and enzymatically inactive. Activity
was restored by addition of a reducing agent and under anoxic conditions in the enzyme assay.
The activation could be initiated by cationic (ferrous iron), neutral (dithiothreitol) or anionic
(dithionite) compounds. However, maximum activity was measured with 4 mM dithionite (268
pkat * mg protein-1). A large excess of reductant caused a decrease in activity, e.g. 10 mM
dithionite or 8 mM DTT were less suitable for activation (Tab. 5).
- 124 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Table 5: Influence of different reducing agents on linalool isomerase activity.
Reducing agent
Dithionite
Dithiothreitol
Cysteine
Ferrous iron
Reducing potential
Concentration
Specific activity
[mV]
[mM]
[pkat * mg protein-1]
- 660
2
89
4
268
10
210
2
199
8
145
16
83
5
23
10
7
5
112
- 330
- 220
2+
- 236 (Fe(OH)3/Fe )
Lis activity was determined in the thermodynamically favorable direction from geraniol to
linalool [15] and was observed between pH 7 and 9.5, with an optimum around pH 8. The
temperature optimum was 35°C and the activation energy was 80.4 ± 6.9 kJ mol-1. For
comparison, the pH and temperature optimum of the Ldi enzyme were at pH 9 and 35°C,
respectively, and the activation energy was 68.6 kJ mol-1 [17]. Geraniol formation from linalool
was detected within 4 hours of incubation (Fig. 7).
- 125 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure 7: Formation of geraniol by linalool isomerase. The most enriched protein sample after sucrose
gradient centrifugation and precipitation was used: 200 μL sample (1.04 mg mL-1 protein), 5 mM
dithionite, 1 μL (R,S)-linalool (≈ 30 mM). Incubation was performed at 28°C and individual samples were
extracted with 200 μL n-hexane after 0, 2 and 4 hours (from bottom to top) and measured by gas
chromatography (Shimadzu GC 14A, see methods). Retention time of geraniol was at 12.55 min.
Kinetic parameters were determined with the most enriched sample in duplicates for the geraniol
isomerization to linalool. The enzyme followed Michaelis-Menten-kinetics with an apparent kMvalue of 455 ± 124 μM and a Vmax of 3.42 ± 0.28 nkat * mg protein-1 (Fig. 8). A similar substrate
affinity was determined for the linalool dehydratase/isomerase (kM 500 μM). Maximal velocity
was higher for the Ldi (Vmax 410 nkat * mg protein-1) than for the Lis, however, the purification
level for Lis was lower, and we do not know whether part of the enzyme is in an inactive state.
- 126 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure 8: Michaelis-Menten plot of geraniol isomerization by linalool isomerase. An apparent k M-value
and Vmax value of 455 ± 124 μM and 3.42 ± 0.28 nkat * (mg protein) -1 were determined for the
isomerization of geraniol to linalool.
Lis did not show a stereospecificity towards linalool isomers: both (R) and (S)-linalool were
formed (Fig. 9). In contrast, the Ldi accepts only (S)-linalool as a substrate [17].
Figure 9: Gas chromatogram demonstration of the absence of stereoselectivity of the linalool isomerase.
Soluble protein extract (1.4 mg protein; solid line) and inner membrane-enriched fraction (1.6 mg protein;
dashed line) from spheroplast disintegration were incubated with geraniol, subsequently extracted with nhexane and analyzed by gas chromatography. Retention times: (R)- and (S)-linalool 7.85 and 8.05 min,
geraniol 12.45 min
- 127 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Earlier studies already showed the regiospecific formation of geraniol from linalool without nerol
formation [15]. To confirm the regioselectivity, Lis activity was tested with nerol or citronellol
and in combination with geraniol. No activity was measured with nerol or citronellol alone.
Linalool isomerase activity dropped to approx. 50 % in the presence of nerol. Activity in the
presence of citronellol and geraniol was barely detectable. Thus, the enzyme is regioselective and
seems to bind nerol with a similar affinity as geraniol, whereas citronellol, which lacks the C2-C3
double bond, is stronger bound than geraniol.
UV-VIS spectroscopy in a range of 200 to 800 nm did not provide evidence for the presence of
cofactors. Cofactor-independent enzymes are known for allylic rearrangements that are catalyzed
by acid-base mechanisms [28]. The isomerization of geraniol to linalool requires a protonation of
the hydroxyl group to leave as water, and a shift in electron density leading to a tertiary
carbocation intermediate which may be attacked by water or a hydroxyl ion, resulting in the
formation of linalool.
Conclusion
We identified a linalool isomerase in T. linaloolentis 47Lol by partial protein purification. The
gene encodes a two-domain protein with a N-terminal anchor in the inner membrane that is
characterized by four transmembrane helices and a C-terminal cytosolic domain which showed
considerable similarity to the linalool dehydratase/isomerase from C. defragrans 65Phen. The
enzyme is active in the reduced state and sensitive towards detergents. It expands the enzyme
class of intramolecular hydroxyl group transferases as a new member, linalool isomerase
(5.4.4.4).
- 128 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Figure S1: Alignment of the linalool dehydratase/isomerase (NCBI:CBW30776) from C. defragrans
65Phen and the linalool isomerase from T. linaloolentis 47Lol (NCBI:ENO87364). Color indicates
similarity.
- 129 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Material and Methods
Bacterial strains, cultivation conditions and biomass harvest
T. linaloolentis 47Lol was cultivated under anaerobic, denitrifying conditions in artificial fresh
water (AFW) medium. Medium was prepared as described by Foss et al. [29] with modifications:
carbonate buffer was replaced by 10 mM Na2HPO4/NaH2PO4 and vitamins were omitted. The
headspace contained nitrogen gas only. 1-2 mM (R,S)-linalool (>97 % purity; Sigma-Aldrich,
Germany) were directly applied without carrier phase as sole carbon and energy source. Cultures
were incubated at 28°C under mild shaking (60 rpm). Alternatively, they were stirred. Bacterial
biomass for protein purification was obtained from 2 L cultures grown on 2 mM linalool and 20
mM nitrate by centrifugation (16000 x g, 25 min, 4°C). If not used directly, biomass was frozen
in liquid nitrogen and stored at -80°C.
Purification attempts by chromatography
First attempts on the purification of the linalool isomerase were performed by classical column
chromatography using anion exchange and hydrophobic interaction columns (DEAE; binding
buffer: 80 mM Tris-Cl, pH 9.5, elution buffer: 80 mM Tris-Cl, pH 9.5 with 1M NaCl; Phenyl
Sepharose; binding buffer: 80 mM Tris-Cl, pH 8.0 with 10 % v/v of a saturated ammonium
sulfate solution, elution buffers: 80 mM Tris-Cl, pH 8.0 and water). Size-exclusion
chromatography was performed on a HiLoad 16/60 Superdex200 column (GE healthcare,
dimensions: 16 x 600 mm; 20 mM KH2PO4/K2HPO4, pH 8.0 with or without 6 M urea).
Calibration was performed with thyroglobulin (670 kDa), bovine gamma-globulin (158 kDa),
chicken ovalalbumin (44 kDa), equine myoglobin (17 kDa) and vitamin B12 (1.35 kDa).
Soluble protein extract was prepared by resuspending biomass in Tris-Cl buffer (80 mM, pH 9.5
for DEAE and pH 8.0 for HIC) and fast thawing at room temperature. Cell disintegration was
performed by mechanical sheering using a One-Shot cell disruptor (Constant Systems Ltd.,
Daventry, UK) at 1.7 GPa two times. The crude extract was clarified by ultracentrifugation
(150000 x g, 30 min, 4°C).
All purification steps were performed on ice or at 5°C. SDS-PAGE was used to characterize
purification samples and protein concentrations were determined according to the method
described by Bradford using bovine serum albumin as a calibration standard [30].
- 130 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Spheroplast preparation
Spheroplasts were prepared according to the protocol for subcellular fractionation described by
Koßmehl et al. [31]. Cells were washed with 1.5 M NaCl solution, collected by centrifugation
(14200 x g, 20 min, 4°C) and resuspended in 20 % (w/v) sucrose, 30 mM Tris-Cl (pH 8.0) and 2
mM EDTA for osmotic shock treatment. The cell suspension was incubated for 20 min at 30°C.
A spheroplast enriched pellet was formed by centrifugation (14200 x g, 20 min, 4°C). The pellet
was resuspended in 5 mL of ice-cold 40 mM Tris-Cl (pH 8.0) and disintegrated by a One Shot
Cell Disruptor (Constants Systems Ltd., UK) in two passages at 1.7 GPa. After removal of larger
cell debris and unbroken cells by centrifugation (14200 x g, 20 min), the supernatant was further
clarified by ultracentrifugation (104000 x g, 1 h, 4°C). The resulting pellet and supernatant
corresponded to an inner membrane fraction (IM, F7, Fig. 1) and a soluble protein fraction (SP,
F6, Fig. 1), respectively.
Sucrose density gradient centrifugation
A linear sucrose gradient was created by overlaying a 20 % (w/v) over a 70 % (w/v) sucrose
solution, 3 mL each. The tubes were incubated horizontally for 90 min to allow mixing. A 1 mLprotein sample was loaded carefully on top of the gradient and centrifugation was performed in a
L-70 ultracentrifuge (70.1Ti rotor, Beckmann Coulter) at 260000 x g, for 4 h at 4°C, with slow
acceleration and deceleration. The linearity of the sucrose gradient was confirmed by
gravimetrical measurement. 1 mL fractions covering the gradient were analyzed for enzyme
activity and for protein content on SDS-PAGE. The two most active fractions were pooled,
diluted with 40 mM Tris-Cl (pH 8.0) to a final volume of 8 mL and a second ultracentrifugation
step (260000 x g, 4°C, 1.5 h ≡ 42 Svedberg) was performed to remove sucrose. This resulted in
the formation of a protein pellet enriched in Lis activity, which was resuspended in 1 mL buffer
and used for enzyme kinetic characterization.
Proteomics by MALDI-ToF MS
Protein samples, obtained from individual purifications, were analyzed by SDS-PAGE coupled
with matrix-assisted laser desorption/ionization time of flight (MALDI-ToF) mass spectrometry.
Protein bands in gels were excised manually, and the Ettan Spot Handling Workstation (GE
Healthcare) was used for trypsin digestion and embedding of the resulting peptide solutions in an
- 131 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
α-cyano-4-hydroxycinnamic acid matrix for spotting onto MALDI targets. MALDI-ToF MS
analysis was performed on an AB SCIEX TOF/TOFTM 5800 Analyzer (AB Sciex / MDS
Analytical Technologies [32]. Spectra in a mass range from 900 to 3700 Da (focus 1700 Da)
were recorded and analyzed by GPS Explorer™ Software Version 3.6 (build 332, Applied
Biosystems) and the Mascot search engine version 2.4.0 (Matrix Science Ltd, London, UK) using
the RAST draft genome as reference.
Heterologous gene expression
The predicted Lis gene was isolated from genomic DNA of T. linaloolentis 47Lol by means of
PCR, using the Phusion Polymerase according to the manufacturer manual (Life Technologies,
Thermo
Fisher
Scientific,
Waltham,
USA)
with
(TCGTACATATGATGAGCAATATGGAATCG)
and
the
primer
pair
pLI_BglII_RV
pLI_NdeI_FW
(CGATAGATCT
TCAGTGGCCCGGCTTG, annealing temperature 59°C). Additionally, a N-terminal truncated
version of the gene was constructed with the primer pair pLI_NdeI_FW-truncated-N
(TCGTACATATGATGCGCGGCGCCAAGC) and pLI_BglII_RV (annealing temperature
68°C), which had an artificial start codon (ATG) and covered the amino acids 141-644 of the Lis.
The genes were cloned into the pET42a overexpression plasmid and transformed into E. coli
BL21(DE3). E. coli BL21(DE3) pET42-Lis or pET42-Lis-'N were grown in liquid LB medium
at 37°C and protein expression was induced by addition of 1 mM IPTG at an optical density (600
nm) of around 1. Cultures were further incubated at 30°C for 4-5 hours. Biomass was harvested
by centrifugation (16000 x g, 25 min, 4°C). Protein extracts were prepared as described above.
Geraniol isomerase activity
Enzymatic activity was determined by end-point analysis for the thermodynamically favored
reaction from geraniol to linlaool. Subfractions obtained during purification were adjusted to
Tris-Cl buffer (40 mM, pH 8.0). Individual assays were prepared in 4 mL glass vials in an
anaerobic chamber with N2 headspace, containing 300-500 μL of sample and 5 mM dithionite as
reducing agent. Samples were incubated for 20 minutes prior addition of 200 μL organic phase
(200 mM geraniol in 2,2,4,4,6,8,8-heptamethylnonane, HMN). Vials were air-tight closed with
butyl rubber stoppers and incubated for 14-16 hours at 28°C under mild shaking. Product
formation was determined by gas chromatography with flame ionization detection (PerkinElmer
- 132 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
Auto System XL, Überlingen, Germany). 1 μL of the HMN phase was injected onto an Optima-5
column (30 m x 0.32 mm, 0.25 μm film thickness; Macherey-Nagel, Germany) with hydrogen as
carrier gas and the following temperature program: injection port 250°C, detection port 350°C,
initial column temperature 40°C for 2 min, increasing to 100°C at a rate of 4°C min-1, keeping
100°C for 0.1 min, followed by an increase to 320°C at 45°C min-1 and hold for 3 min. The split
ratio was set to 1:9.
The effect of detergents on Lis activity was tested for Triton X100 and Tween20 (0.5 %, 1 %
w/v), CHAPS (0.1 % w/v) and n-octyl-D-D-glucoside (0.1 %, 0.5 %, 1 % w/v). Detergents were
added to the soluble spheroplast fraction (protein concentration 0.1 mg mL-1) and standard
enzyme assays were performed.
Reducing agents were tested with a dialyzed (Visking dialysis tubing 12-14 kDa cut-off, Serva)
soluble protein extract (4 mg mL-1 protein). The following reducing agents were used: dithionite
(2, 4 and 10 mM), dithiothreitol (2, 8 and 16 mM), cysteine (5 and 10 mM), ferrous iron (5 mM).
The activity assay was conducted as described aforementioned.
Temperature dependency on Lis activity was determined between 12°C and 50°C. Samples (300
μL, 6.8 mg mL-1 protein) were prepared like in the standard activity assay and pre-incubated for
20 minutes at the individual temperatures prior substrate addition. The assay was terminated after
8 hours and analyzed by gas chromatography. Activation energy was calculated from the
Arrhenius plot (y = -9664.3 x + 36.2; R2 = 0.914).
The pH-optimum was determined by incubating crude cell lysate (20 mg mL-1) in a pH-range
from 7 to 9.5 in Tris-Cl (40 mM). The enzyme assay was performed according to the standard
enzyme assay.
Kinetic parameter (kM and Vmax) were determined for the most enriched enzyme fraction in
biological duplicates (68 and 80 μg mL-1 protein; 10 to 20 μg total protein in final assay).
Samples were incubated with geraniol concentration from 0.125 to 4 mM, directly applied
without carrier phase. Both, substrate and enzyme were prepared separately with 7 mM dithionite
and pre-incubated. Reactions were started by injecting an equal volume (200 μL) of enzyme to
the substrate solution. Samples were incubated at 28°C for 90 minutes and terminated by addition
of 100 mM NaOH (final concentration). 1 μL of sample was directly subjected to GC analysis.
Kinetic parameters were calculated from primary data plotted in a Michaelis-Menten-graph.
Substrate specificity of the Lis was tested with geraniol, nerol and cironellol. A 400 μL-sample
(active fraction after sucrose gradient, SP 4/5) was incubated with 200 μL of 200 mM geraniol,
- 133 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
nerol and citronellol in HMN as well as with 200 μL of geraniol-nerol and geraniol-citronellol
mixtures in HMN (100 mM each). Assays were prepared as described for the standard activity
assay. All substrates were prepared in HMN.
Stereoselectivity was tested with soluble protein extract (1.4 mg protein) and inner membraneenriched fraction (1.6 mg protein) from spheroplast disintegration. Samples were treated with 5
mM dithionite and incubated with 10 mM geraniol under anaerobic conditions at 28°C for 14
hours and subsequently extracted with 200 μL n-hexane. Monoterpene analysis was performed
by gas chromatography with flame ionization detector (Shimadzu GC-14A, Shimadzu
Corporation) on a Hydrodex-ß-6TBDM column (25 m x 0.25 mm, Macherey-Nagel, Germany)
with the following temperature program: injection port 200°C, detection port 250°C, initial
column temperature 60°C for 1 min, increasing to 130°C at a rate of 5°C min-1, keeping 130°C
for 0.5 min, followed by an increase to 230°C at 20° min-1 and hold for 4 min.
Linalool isomerase activity
The forward reaction of the linalool isomerase - linalool to geraniol - was tested in a separate
assay. 200 μL of enriched fraction after sucrose gradient centrifugation were treated with 5 mM
dithionite and incubated with 1 μL (R,S)-linalool under anaerobic conditions at 28°C for 0, 2 and
4 hours. Samples were extracted with 200 μL n-hexane and analyzed by GC (Shimadzu GC14A).
UV-VIS spectrum for cofactors
The most enriched, active protein sample (0.95 mg mL-1 protein) was analyzed by UV-VIS
spectroscopy (Beckman DU-640 spectrophotometer) in the range of 200 to 800 nm to detect
cofactors.
Gene identification and bioinformatic analysis
A draft genome for T. linaloolentis 47Lol was obtained by merging data from two public
available sequencing datasets: ASM31020 (4.199 Mbp on 220 contigs) published 2012 and
ASM62130 (4.214 Mbp on 46 contigs) published 2014 on NCBI. Contigs were automatically
merged using Sequencher 4.6 with a minimum match percentage of 95 % and a minimum overlap
of 50 bp. The resulting draft genome had 4.4 Mbp on 23 contigs and was uploaded to RAST for
- 134 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
further analysis [33, 34]. Identification of a putative gene, coding for a linalool isomerase (lis),
was performed by homology search using the linalool dehydratase/isomerase sequence from
Castellaniella degragrans 65Phen. The identified gene was analyzed by various bioinformatic
tools: SignalP 4.1 for prediction of signal peptides [35], TMHMM, SOSUI and Philius for
prediction of transmembrane helices [36-39] and the Pfam database to search for motifs and
domain patterns [40].
Competing interests
The authors declare that they have no competing interests.
Authors´ contribution
RM and JH designed the study. RM and BS performed the purification and the experiments for
enzyme characterization. SM and TS conducted mass-spectrometry based protein identifications
and analyzed data. RM and JH drafted the manuscript. All authors read and approved the final
manuscript.
Acknowledgments
We thank Dirk Albrecht for MALDI-ToF measurements. This study was financed by the Max
Planck Society.
- 135 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
References
1.
Breitmaier E: Terpenes; Flavors, Fragrances, Pharmaca, Pheromones, vol. 1st edition: WileyVCH; 2006.
2.
Ajikumar PK, Tyo K, Carlsen S, Mucha O, Phon TH, Stephanopoulos G: Terpenoids:
Opportunities for biosynthesis of natural product drugs using engineered microorganisms.
Molecular Pharmaceutics 2008, 5:167-190.
3.
Dudareva N, Klempien A, Muhlemann JK, Kaplan I: Biosynthesis, function and metabolic
engineering of plant volatile organic compounds. New Phytol 2013, 198:16-32.
4.
Sharkey TD, Wiberley AE, Donohue AR: Isoprene emission from plants: why and how. Annals
of Botany 2008, 101:5-18.
5.
Günther A, Hewitt CN, Erickson D, Fall R, Geron C, Graedel T, Harley P, Klinger L, Lerdau M,
McKay WA et al: A global model of natural volatile organic compound emission. Journal of
Geophysical Research-Atmosphere 1995, 100:8873-8892.
6.
Atkinson R, Arey J: Atmospheric degradation of volatile organic compounds. Chemical
Reviews 2003, 103:4605-4638.
7.
Kainulainen P, Holopainen JK: Concentrations of secondary compounds in Scots pine needles
at different stages of decomposition. Soil Biology & Biochemistry 2002, 34:37-42.
8.
Wilt FM, Miller GC, Everett RL, Hackett M: Monoterpene concentrations in fresh, senescent,
and decaying foliage of single-leaf pinyon (Pinus monophylla Torr. and Frem.: Pinaceae)
from the western great basin. Journal of Chemical Ecology 1993, 19:185-194.
9.
Ziemann PJ, Atkinson R: Kinetics, products, and mechanisms of secondary organic aerosol
formation. Chemical Society Reviews 2012, 41:6582-6605.
10.
Bakkali F, Averbeck S, Averbeck D, Waomar M: Biological effects of essential oils - A review.
Food and Chemical Toxicology 2008, 46:446-475.
11.
Schewe H, Mirata MA, Holtmann D, Schrader J: Biooxidation of monoterpenes with bacterial
monooxygenases. Process Biochemistry 2011, 46:1885-1899.
12.
Marmulla R, Harder J: Microbial monoterpene transformations – A review. Frontiers in
Microbiology 2014, 5.
13.
Bell SG, Dale A, Rees NH, Wong LL: A cytochrome P450 class I electron transfer system
from Novosphingobium aromaticivorans. Applied Microbiology and Biotechnology 2010,
86:163-175.
- 136 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
14.
Ullah AJH, Murray RI, Bhattacharyya PK, Wagner GC, Gunsalus IC: Protein components of a
cytochrome P-450 linalool 8-methyl hydroxylase. Journal of Biological Chemistry 1990,
265:1345-1351.
15.
Foss S, Harder J: Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. FEMS Microbiol Letters 1997,
149:71-75.
16.
Foss S, Harder J: Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on
oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) and nitrate. Systematic
and Applied Microbiology 1998, 21:365-373.
17.
Brodkorb D, Gottschall M, Marmulla R, Lüddeke F, Harder J: Linalool dehydratase-isomerase,
a bifunctional enzyme in the anaerobic degradation of monoterpenes. Journal of Biological
Chemistry 2010, 285:30436-30442.
18.
Lüddeke F, Harder J: Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences 2011, 66:409-412.
19.
Schuster J, Schäfer F, Hübler N, Brandt A, Rosell M, Härtig C, Harms H, Müller RH, Rohwerder
T: Bacterial degradation of tert-amyl alcohol proceeds via hemiterpene 2-methyl-3-buten-2ol by employing the tertiary alcohol desaturase function of the Rieske nonheme
mononuclear iron oxygenase MdpJ. Journal of Bacteriology 2012, 194:972-981.
20.
Malone VF, Chastain AJ, Ohlsson JT, Poneleit LS, Nemecek-Marshall M, Fall R:
Characterization of a Pseudomonas putida allylic alcohol dehydrogenase induced by growth
on 2-methyl-3-buten-2-ol. Applied and Environmental Microbiology 1999, 65:2622-2630.
21.
Kyte J, Doolittle RF: A simple method for displaying the hydropathic character of a protein.
Journal of Molecular Biology 1982, 157:105-132.
22.
Lüddeke F, Wülfing A, Timke M, Germer F, Weber J, Dikfidan A, Rahnfeld T, Linder D,
Meyerdierks A, Harder J: Geraniol and geranial dehydrogenases induced in anaerobic
monoterpene degradation by Castellaniella defragrans. Applied and Environmental
Microbiology 2012, 78:2128-2136.
23.
Kleinsmith LJ, Kish VM: Principles of cell and molecular biology, 2 edn: HarperCollins; 1995.
24.
Rath A, Glibowicka M, Nadeau VG, Chen G, Deber CM: Detergent binding explains
anomalous SDS-PAGE migration of membrane proteins. Proceedings of the National
Academy of Sciences 2009, 106:1760-1765.
25.
Gohon Y, Popot JL: Membrane protein-surfactant complexes. Current Opinion in Colloid &
Interface Science 2003, 8:15-22.
- 137 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
26.
Seddon AM, Curnow P, Booth PJ: Membrane proteins, lipids and detergents: not just a soap
opera. Biochimica Et Biophysica Acta-Biomembranes 2004, 1666:105-117.
27.
Christianson DW: Structural biology and chemistry of the terpenoid cyclases. Chemical
Reviews 2006, 106:3412-3442.
28.
Schwab JM, Henderson BS: Enzyme-catalyzed allylic rearrangements. Chemical Reviews
1990, 90:1203-1245.
29.
Foss S, Heyen U, Harder J: Alcaligenes defragrans sp. nov., description of four strains isolated
on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alpha-phellandrene)
and nitrate. Systematic and Applied Microbiology 1998, 21:237-244.
30.
Bradford MM: Rapid and sensitive method for quantitation of microgram quantities of
protein utilizing principle of protein-dye binding. Analytical Biochemistry 1976, 72:248-254.
31.
Koßmehl S, Wöhlbrand L, Drüppel K, Feenders C, Blasius B, Rabus R: Subcellular protein
localization (cell envelope) in Phaeobacter inhibens DSM 17395. Proteomics 2013, 13:27432760.
32.
Wolf C, Hochgräfe F, Kusch H, Albrecht D, Hecker M, Engelmann S: Proteomic analysis of
antioxidant strategies of Staphylococcus aureus: Diverse responses to different oxidants.
Proteomics 2008, 8:3139-3153.
33.
Aziz RK, Bartels D, Best AA, DeJongh M, Disz T, Edwards RA, Formsma K, Gerdes S, Glass
EM, Kubal M et al: The RAST server: Rapid annotations using subsystems technology. BMC
Genomics 2008, 9.
34.
Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA, Gerdes S, Parrello B,
Shukla M et al: The SEED and the Rapid Annotation of microbial genomes using Subsystems
Technology (RAST). Nucleic Acids Research 2014, 42:D206-D214.
35.
Petersen TN, Brunak S, von Heijne G, Nielsen H: SignalP 4.0: discriminating signal peptides
from transmembrane regions. Nature Methods 2011, 8:785-786.
36.
Sonnhammer EL, von Heijne G, Krogh A: A hidden Markov model for predicting
transmembrane helices in protein sequences. Proceedings / International Conference on
Intelligent Systems for Molecular Biology ; ISMB International Conference on Intelligent Systems
for Molecular Biology 1998, 6:175-182.
37.
Hirokawa T, Boon-Chieng S, Mitaku S: SOSUI: classification and secondary structure
prediction system for membrane proteins. Bioinformatics 1998, 14:378-379.
38.
Imai K, Asakawa N, Tsuji T, Akazawa F, Ino A, Sonoyama M, Mitaku S: SOSUI-GramN: high
performance prediction for sub-cellular localization of proteins in Gram-negative bacteria.
Bioinformation 2008, 2:417-421.
- 138 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
39.
Reynolds SM, Käll L, Riffle ME, Bilmes JA, Noble WS: Transmembrane topology and signal
peptide prediction using dynamic bayesian networks. PLoS Computational Biology 2008,
4:e1000213.
40.
Finn RD, Bateman A, Clements J, Coggill P, Eberhardt RY, Eddy SR, Heger A, Hetherington K,
Holm L, Mistry J et al: Pfam: the protein families database. Nucleic Acids Research 2014, 42:
222-230.
- 139 -
Chapter VI - Linalool isomerase from Thauera linaloolentis 47Lol
- 140 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Chapter VII
The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Robert Marmulla1, Edinson Puentes Cala1, Stephanie Markert2, Thomas Schweder2, Jens
Harder1*
1
Dept. of Microbiology, Max Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359
Bremen
*
corresponding author: jharder@mpi-bremen.de
2
Institute for Pharmacy, Dept. of Pharmaceutical Biotechnology, University of Greifswald,
Felix-Hausdorff-Str. 3, D-17487 Greifswald
Research article submitted to BioMed Central (BMC) Microbiology
- 141 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Abstract
Background: The betaproteobacterium Thauera linaloolentis 47Lol was isolated on the tertiary
monoterpene alcohol (R,S)-linalool as sole carbon and energy source under denitrifying
conditions. Growth experiments indicated the formation of geraniol and geranial. Thus, a 3,1hydroxyl-'1-'2-mutase (linalool isomerase) activity may initiate the degradation, followed by
enzymes of the acyclic terpene utilization (Atu) and leucine/isovalerate utilization (Liu) pathways
that were extensively studied in Pseudomonas spp. growing on citronellol or geraniol.
Results: A transposon mutagenesis yielded 39 transconjugants that could not grow anaerobically
on linalool and nitrate in liquid medium. The deficiencies were apparently based on gene
functions required to overcome the toxicity of linalool, but not due to inactivation of genes in the
degradation pathway. Growing cultures formed geraniol and geranial transiently, but also geranic
acid. Analysis of expressed proteins detected several enzymes of the Atu and Liu pathways. The
draft genome of T. linaloolentis 47Lol had atu and liu genes with homology to those of
Pseudomonas spp..
Conclusion: The in comparison to monoterpenes larger toxicity of monoterpene alcohols is
defeated by several modifications of the cellular structure and metabolism in T. linaloolentis
47Lol. The acyclic terpene utilization pathway is used in T. linaloolentis 47Lol during growth on
(R,S)-linalool and nitrate under anoxic conditions. This is the first experimental verification of an
active Atu pathway outside of the genus Pseudomonas.
Keywords: Monoterpene, Linalool, Geraniol, acyclic terpene utilization
- 142 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Introduction
Linalool (C10H18O, 3,7-dimethylocta-1,6-dien-3-ol), a tertiary monoterpene alcohol, is the main
constituent in essential oils of lavender and coriander. It is also a fragrance of flowers [1, 2]. As
tertiary alcohols cannot be directly oxidized to ketones, microorganisms initiate their metabolism
with oxidation reactions by oxygenases at other parts of the molecule [3] or with isomerizations
without molecular oxygen. The latter case has been described for the betaproteobacterium
Castellaniella defragrans 65Phen. A linalool dehydratase/isomerase metabolizes (S)-linalool to
geraniol [4, 5]. The further degradation pathway to geranic acid is catalyzed by a NAD+dependent geraniol and a geranial dehydrogenase (GeoA, GeoB) [6]. However, a pathway for
geranic acid degradation was not found in the genome of C. defragrans 65Phen [7]. Expected
were genes and operons as described for the degradation of the monoterpene alcohols geraniol
and citronellol in Pseudomonas aeruginosa PAO1. In this strain, oxidation of the alcohols to their
corresponding carboxylic acids and the oxidation of citronellic acid to geranic acid is followed by
activation as CoA thioester (Fig. 1). The tertiary carbon atom is transformed into a secondary
carbon atom by carboxylation of the beta-methyl group as initial reaction. Hydration on the C3
atom yields 3-hydroxy-3-isohexenylglutaryl-CoA followed by acetate cleavage which removes
the methyl group initially present at the tertiary carbon atom. 7-methyl-3-oxo-6-octenoyl-CoA is
the product of the pathway that was named acyclic terpene utilization (Atu). For several
Pseudomonas species the enzymes were identified: citronellol/citronellal dehydrogenase (AtuB
and AtuG), citronellyl-CoA dehydrogenase (AtuD), long-chain acyl-CoA synthase (AtuH),
geranyl-CoA carboxylase (AtuC and AtuF), isohexenyl-glutaconyl-CoA hydratase (AtuE) and 3hydroxy-3-isohexenylglutaryl-CoA:acetate lyase (AtuA). The corresponding genes are arranged
in an operon like structure (atuABCDEFGH) under the control of a transcriptional regulator
(AtuR) [8-10]. 7-methyl-3-oxo-6-octenoyl-CoA undergoes two rounds of E-oxidation to two
acetyl-CoA and 3-methyl-crotonyl-CoA. The latter enters the leucine/isovalerate utilization
pathway (Liu) for complete mineralization [11].
- 143 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Figure 1: Proposed pathway for the linalool metabolism in T. linaloolentis 47Lol, based on the acyclic
terpene utilization pathway (Atu) in Pseudomonas spp. [adapted from 9]. Linalool isomerase (Lis),
geraniol dehydrogenase (GeoA), geranial dehydrogenase (GeoB).
- 144 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
The absence of the Atu pathway in the denitrifying C. defragrans 65Phen motivated us to search
for the pathway in another denitrifying betaproteobacterium T. linaloolentis 47Lol. It was
isolated on linalool as sole carbon and energy source under denitrifying conditions and uses
linalool and geraniol as growth substrates. It has a strict respiratory metabolism, using molecular
oxygen, nitrate, nitrite or nitrous oxide as terminal electron acceptor [12]. Growth studies
revealed the regioselective formation of geraniol from linalool in the stationary phase, while
linalool and geranial were detected in geraniol-grown cultures. These findings suggested the
presence of a linalool isomerase, a 3,1-hydroxyl-'1-'2-mutase [13]. In this study, we attempted
the identification of genes and proteins essential for the linalool degradation by transposon
mutagenesis and by mass spectroscopy of proteins.
Results and Discussion
Transposon insertion mutagenesis
5916 transconjugants were screened on solid medium for denitrifying growth on acetate or
linalool as carbon source. 92 mutants had a growth deficiency on linalool, thus a mutation
frequency of 0.0155 was observed. Transfer cultures in liquid medium revealed for 53 of these
transposon mutants a biomass yield equal to the wildtype, only 39 transconjugants lacked growth
on linalool. The transposon insertion sites in the genome were identified by sequencing and
comparison to the genome sequence. Surprisingly, genes of the Atu pathway were not inactivated
by transposon insertions. The majority of the inactivated genes was affiliated to the following
functional classifications (Supplement table S1): DNA modification/processing (10 mutants),
cellular transport systems (7 mutants), membrane integrity (4 mutants), miscellaneous (7
mutants) and unclassified (11 mutants). In general, these genes, involved in biosynthesis and
repair, have to be seen as reflection of the cell toxicity of monoterpenes rather than the
involvement in the biodegradation. Monoterpene alcohols exhibit usually a higher toxicity than
the pure hydrocarbons. Once these compounds have passed the polar part of the LPS, they can
integrate into the outer and inner membranes as well as the hydrophobic core of proteins. The
amphiphilic monoterpene alcohols may act as surfactants. Destabilization of the membrane
system and cellular structures causes ion leakages and a collapse of the proton motive force.
- 145 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Effects on proteins and on enzyme activities have been reported [14-16]. As defense against the
penetration and accumulation of lipophillic substances, Gram-negative bacteria alter the
composition and structure of their outer membrane (OM) and the lipopolysaccharide layer (LPS)
[17, 18], e.g., a change in membrane composition from saturated fatty acids to cyclopropanecontaining fatty acids has been observed from acetate-grown to monoterpene-grown cells of C.
defragrans 65Phen [19]. Among the transconjugants were three mutants affected in LPS core and
O-antigen synthesis or modification, while another mutant was affected in a penicillin-binding
protein (PDB 2) involved in peptidoglycan synthesis during cell growth and division [20, 21].
Bacteria counteract the toxic effects of monoterpenes by active export with energy-driven
transporters. Well known examples for such transport systems are multidrug resistance (MDR) or
resistance-nodulation-cell division (RND) efflux pumps, however the substrate range of the
exporters is often not known [22-24]. We obtained linalool-growth deficient transconjugants with
insertion into an ABC-type multidrug transporter or branched-chain amino acid transporters
(three mutants). Ten insertions were in genes involved in DNA modification and transcriptional
control. Two transconjugants had inactivated a transcriptional repressor of the TetR family
(ENO85695). The repressor has 38 % identity on amino acid level (blastp) to AtuR, a specific
transcriptional regulator for the acyclic terpene utilization in Pseudomonas spp.. Insertion
mutants of P. aeruginosa showed a consecutive basal expression of Atu proteins [25]. However,
the atu genes in 47Lol are not arranged in the operon structure of Pseudomonas and are distant to
the repressor gene. Thus the transcriptional role of this putative repressor remains unclear.
Another transconjugant had an insertion in a transcriptional regulator of the ModE family, known
to be involved in molybdenum uptake and incorporation [26]. The oxidation of geraniol to
geranic acid in P. aeruginosa PAO1 is dependent on molybdenum, but other Pseudomonas
species grow on geraniol and express the Atu pathway without evidence for a molybdenum
requirement [10].
The high frequency of transconjugants with a deficiency in growth on linalool coincides with
related studies that revealed distinct changes in the transcriptome and proteome in cells switching
the growth substrate from non-toxic aliphatic short chain acids to monoterpenes. Pseudomonas
sp. M1 changed the expression of nearly 30 % of the genome in response to a change from lactate
to myrcene, including genes involved in the Atu pathway, citric acid cycle and E-oxidation, genes
for a restructuring of the LPS layer and membranes and an up-regulation of nitrate-respiration
gene clusters [27]. The toxicity of monoterpenes was also reflected in the proteomes of C.
- 146 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
defragrans 65Phen cells grown on acetate and D-phellandrene: among the 107 identified induced
proteins were several transporters and membrane biosynthesis proteins [7]. The failure to identify
transconjugants in the degradation pathway for linalool may be attributed to the increased toxicity
of the monoterpenoid. A similar transposon mutagenesis in C defragrans 65Phen identified the
central degradation pathway for monoterpenes [7], but these alkenes are less toxic than
monoterpene alcohols [6].
Characterization of growth on linalool
T. linaloolentis 47Lol was grown on linalool under denitrifying conditions. Besides the
previously demonstrated formation of geraniol and geranial as metabolites [13] we established
the detection of geranic acid (Fig. 2 a,b). Geraniol and geranial accumulated transiently to 7 and
10 μM, while geranic acid accumulated up to 200 μM. Low cellular concentrations of the toxic
geraniol and geranial are due to high affinities of the corresponding enzymes, which is part of a
cellular defense [28].
Figure 2: Growth of T. linaloolentis 47Lol on (R,S)-linalool under denitrifying conditions. (A) Optical
density at 600 nm as a measure for growth and linalool concentration in mM, (B) Optical density at 600
nm as a measure for growth and geraniol, geranial and geranic acid concentration in μM. Experiment was
performed in duplicates. Initial increase in linalool concentration is due to time-dependent dissolution of
linalool in medium. Linalool decreased accompanied with an increase in optical density. Geraniol and
geranial accumulated only transiently, while geranic acid accumulated up to 200 μM.
- 147 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Activities of the initial enzymes - linalool isomerase, geraniol dehydrogenase (GeoA) and
geranial dehydrogenase (GeoB) - were measured in cell-free protein extracts of linalool-grown
cultures. Linalool isomerase was measured for the thermodynamically favored backward reaction
from geraniol to linalool and showed an activity of 260 pkat * mg protein-1. NAD+-reductase
activity for geraniol and citral (mixture of geranial and neral) were 269 ± 51 pkat * mg protein-1
and 247 ± 107 pkat * mg protein-1, respectively. Abundant proteins in the cell-free protein extract
were identified by SDS-PAGE and MALDI-ToF-based mass spectrometric analysis (Fig. S1).
Proteins annotated as linalool isomerase, geraniol dehydrogenase, the two subunits of geranylCoA carboxylase as well as a 3-hydroxy-3-isohexenylglutaryl-CoA:acetate lyase were identified
together with the two subunits of the methylcrotonyl-CoA carboxylase and the isovaleryl-CoA
dehydrogenase of the leucine degradation pathway (liu operon).
Genome annotation
Two publicly available draft genomes of T. linaloolentis 47Lol were assembled and yielded a
draft genome with 23 contigs of 4,402,076 bp, an overall G+C content of 66.6 %, 4084 open
reading frames, 46 transfer RNA genes and 1 ribosomal RNA operon. The genome completely
covers central metabolic pathways, e.g. citric acid cycle, E-oxidation of fatty acids, oxygen
respiration and denitrification. Genetic information from P. citronellolis, P. aeruginosa PAO1
(Atu and Liu pathway) and C. defragrans 65Phen (linalool isomerization and geraniol/geranial
oxidation) were used to identify homologous genes in T. linaloolentis 47Lol (Table 1).
- 148 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Table 1: Genes involved in the acyclic terpene utilization (Atu) and leucin isovalerate utilization (Liu)
pathway. Genes were identified by amino acids alignments using C. defragrans 65Phen1 and P.
aeruginosa PAO12 sequences as references. Proteins identified in the proteomic approach were analyzed
by SDS-PAGE coupled to MALDI-ToF MS.
Enzyme
NCBI
accession
Length
[AA]
AA
similarity
[%]
E-value
Protein
identification
by MS
Linalool isomerase1
ENO87364
644
20
3E-10
Yes
Geraniol dehydrogenase1
(GeoA)
ENO84122
366
46
7E-96
Yes
Geranial dehydrogenase1
(GeoB)
ENO84123
456
31
7E-39
No
Acyl-CoA synthase2
(AtuH)
ENO87356
545
24
1E-15
No
545
54
0
Yes
Geranyl-CoA
carboxylase2 beta-subunit ENO87361
(AtuC)
Geranyl-CoA
carboxylase2 alpha
subunit (AtuF)
ENO87362
705
52
0
Yes
Isohexenyl-glutaconylCoA hydratase2 (AtuE)
ENO87363
258
30
7E-27
No
3-Hydroxy-3isohexenylglutarylCoA:acetate lyase2
(AtuA)
ENO84124
615
52
0
Yes
535
72
0
Yes
3-Methylcrotonyl-CoA
carboxylase2 beta-subunit ENO88226
(LiuB)
3-Methylcrotonyl-CoA
carboxylase2 alpha
subunit (LiuD)
ENO88223
668
53
0
Yes
3-Methylglutaconyl-CoA
hydratase2 (LiuC)
ENO88225
265
44
3E-73
No
HydroxymethylglutarylCoA lyase2 (LiuE)
ENO88221
312
63
2E-129
No
- 149 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Annotations were fully supported by the aforementioned protein identification. One result of the
reassembly was the formation of two large contigs that contained all genes of the Atu and Liu
pathways over a region of 24 and 9.1 kb, respectively. The atu genes were organized with the
putative linalool isomerase gene lis downstream of atuCFE and the genes for the geraniol
oxidation geoAB upstream of atuA (Fig. 3). The genes in between were annotated as hypothetical
proteins. Gene homologs for atuB and atuG, the putative citronellol and citronellal
dehydrogenases, were identified in the draft genome but did not show significant similarity in a
reciprocal best match analysis.
Figure 3: Gene arrangement of acyclic terpene utilization (atu) and leucine isovalerate utilization (liu)
pathway genes in the draft genome of T. linaloolentis 47Lol. Annotation and NCBI accession numbers are
indicated. Gaps size between atuH and atuC is 3488 bp and hap size between lis and geoA is 7617 bp. The
atu genes are located on a 388748 bp large contig, while the liu genes are located on a 205711 bp contig.
- 150 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Conclusion
A transposon mutagenesis targeting the degradation pathway for linalool yielded mutants
defective in the defense against the toxic monoterpene alcohol. Enzyme activities of a linalool
isomerase, geraniol and geranial dehydrogenases together with the formation of geranic acid
demonstrated the initial degradation pathway. Mass spectrometry identified the carboxylases of
Atu and Liu pathways which indicated the utilization of these pathways for geranic acid
degradation. The draft genome contained the atu genes together with candidate genes for the
initial degradation pathway. To our knowledge, this is the first description of an active Atu
pathway outside of the genus Pseudomonas and under anoxic conditions.
- 151 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Methods
Bacterial strains and cultivation conditions
T. linaloolentis 47Lol was cultivated under anaerobic, denitrifying conditions in artificial fresh
water (AFW) medium. Medium was prepared as described by Foss et al. [19] with modifications.
Carbonate buffer was replaced by 10 mM Na2HPO4/NaH2PO4 and vitamins were omitted. The
headspace contained nitrogen gas only. Acetate or (R,S)-linalool were used as growth substrates.
Growth experiments were performed in 10 mL cultures, incubated at 28°C under mild shaking
(90 rpm). Growth was estimated by measuring the optical density at 600 nm. Monoterpenes were
purchased from Sigma-Aldrich (Germany) and were of 95 to 97 % purity. Geranial was provided
as citral; the mixture of geranial and neral.
Solid AFW medium (0.5 g L-1 KH2PO4, 0.5 g L-1 NH4Cl, 0.5 g L-1 MgSO4 x 7 H2O, 0.1 g L-1
CaCl2 x 2 H2O, 0.85 g L-1 NaNO3, 11.9 g L-1 HEPES, 15 g L-1 agar, pH 7.2) was mixed,
autoclaved and supplemented with trace-element and selenite-tungstate stock solutions as
described [19]. Plates contained either 2 mM (R,S)-linalool or 50 mM acetate as carbon source.
Liquid brain heart infusion (BHI) medium (12 g L-1 brain heart infusion, 10 g L-1 peptone, 4 g L-1
hydrolyzed casein, 5 g L-1 NaCl, 2 g L-1 glucose, 2.5 g L-1 Na2HPO4, 0.2 g L-1 NaNO3, pH 7.4)
was used to yield higher biomass for genomic DNA extraction. Antibiotics were provided from
stock solutions in the following working concentrations: 25-50 μg mL-1 kanamycin, 50 μg mL-1
rifampicin, 1 μg mL-1 ofloxacin.
Transposon insertion mutagenesis
Transposon insertion mutagenesis was performed by using the mini-Tn5 plasmid and biparental
conjugation as described by Larsen et al. [29]. Two spontaneous T. linaloolentis 47Lol mutants
resistant to the antibiotics ofloxacin (47Lol-OF) or rifampicin (47Lol-RIF) were obtained by
repeated culturing of the wildtype in anoxic medium on 20 mM acetate and 10 mM nitrate in the
presence of various concentrations of the antibiotics (20 – 200 μg mL-1 rifampicin, 0.05 – 0.15 μg
mL-1 ofloxacin). Escherichia coli BW20767 [30] harboring the pRL27 plasmid with the mini-Tn5
transposon was grown overnight in lysogeny broth with 50 μg mL-1 kanamycin at 37°C. Cells
were collected by centrifugation (8000 x g, 5 min) and washed twice with fresh medium without
antibiotic. The cells were resuspended in medium, diluted to an optical density (600 nm) of 1 and
mixed in a donor:recipient ratio of 1:3. 200 μL of this mixture were incubated on AFW-plates (50
- 152 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
mM acetate, no antibiotics) at 28°C for 24 hours aerobically. Cells were resuspended in 1 mL
AFW and diluted twofold and tenfold and plated on AFW-plates containing 50 mM acetate,
kanamycin and rifampicin or ofloxacin. The plates were incubated for up to 5 days aerobically at
28°C. Colonies were randomly screened for transposon-integration in the genome by PCR with
the primer pair pRL27 Tn5_F (CGTTACATCCCTGGCTTGTT) and pRL27 Tn5_R
(TGAAGAAGGTGTTGCTGA) [7]. Transconjugants were replica-plated on AFW-plates
containing either 20 mM acetate or 2 mM (R,S)-linalool and nitrate. Colonies showing deficiency
for anaerobic growth on linalool were selected and transferred into liquid culture and a second
anoxic screening on acetate and linalool in the presence of nitrate was performed. Cultures
deficient in growth on linalool were selected for further analysis. Genomic DNA was prepared to
determine the location of transposon insertion and to identify the affected gene by direct
sequencing. Mutants were grown in 10 mL BHI medium overnight. Biomass was collected by
centrifugation (4500 x g, 10 min), resuspended in 150 μL buffer (Tris-Cl 50 mM, 10 mM EDTA,
pH 8, 50 units RNase A) and treated with 150 μL lysis buffer (200 mM NaOH, 1 % w/v SDS).
After mixing and incubation at 96°C for 20 minutes the solution was neutralized by addition of
225 μL neutralization buffer (3 M sodium acetate, pH 5.5). Precipitates were removed by
centrifugation (20000 x g, 10 min). Genomic DNA was precipitated from the supernatant by
addition of ice-cold isopropanol to a content of 50 % (v/v), incubation at -20°C for 60 min and
centrifugation (20000 x g, 20 min, 4°C). The DNA pellet was washed with 70 % (v/v) ice-cold
ethanol, centrifuged and dissolved in 40 μL water. Sequencing reactions were performed with the
BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems Life Technologies
Corporation, USA), gDNA (1 – 3 μg) as template and the primer pair tnpRL 17_1
(AACAAGCCAGGGATGTAA) and tnpRL 13_2 (CAGCAACACCTTCTTCACGA) [29] using
the following program: 96°C for 20 sec, 99 cycles of 96°C for 10 sec, 56°C for 5 sec, 60°C for 4
min. Amplicons were analyzed with an ABI Prism 3130xl Genetic Analyzer (Applied Biosystems
Life Technologies Corporation, USA).
Metabolite analysis
T. linaloolentis 47Lol was cultivated on 1.5 mM (R,S)-linalool and 10 mM nitrate in duplicate
cultures. Inoculum was 2 % (v/v) of a linalool-adapted culture. Two cultures with similar optical
densities were sampled for metabolite extraction, covering growth over the lag-phase,
exponential phase to the entry into the stationary phase. 10 mL culture liquid was transferred to a
- 153 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
15 mL plastic tube, 3 g of NaCl and 250 μL of n-hexane were added. The sample was mixed by
vortexing for 30 seconds and 10 minutes incubation on a tilting shaker. Phase separation was
achieved by 10 minutes centrifugation (3500 x g, 10 min, 5°C). The clear hexane phase was
recovered into a GC-vial. 1 μL sample was analyzed by gas chromatography with flame
ionization detection (Shimadzu GC-14A, Shimadzu Corporation) on a Hydrodex-E-6TBDM
column (25 m x 0.25 mm, Macherey-Nagel, Germany) with the following temperature program:
injection port 200°C, detection port 250°C, initial column temperature 60°C for 1 min, increasing
to 130°C at a rate of 5°C min-1, keeping 130°C for 0.5 min, followed by an increase to 230°C at
20°C min-1 and hold for 4 min. To determine the geranic acid concentration, 50 μL of the hexane
phase were mixed with 50 μL 1 mM NaOH and the hexane was allowed to evaporate. 5 μL of 20
mM phosphoric acid were added and the sample was analyzed by an Acquity UPLC H-class
system (Waters Corporation, USA). 1 μL sample was separated on a reverse phase (BEH C18,
1.7 μm, 2.1 x 50 mm) column with 1 mM H3PO4 at 0.6 mL min-1 in a water-acetonitrile gradient
from 10 to 70 % acetonitrile (v/v) at 30°C. UV detection was performed at 221 nm.
Geraniol-, Geranial dehydrogenase and Linalool isomerase assays
Soluble protein extract of T. linaloolentis 47Lol was prepared from biomass suspension in TrisCl buffer (40 mM, pH 8.0) by passing through an One-Shot cell disruptor (Constant Systems
Ltd., UK) at 1.7 GPa two times, followed by ultracentrifugation (150000 x g, 30 min, 4°C). The
clarified supernatant was used for further experiments. Geraniol- and Geranial dehydrogenase
activities were determined by absorption measurement of NADH formation at 340 nm on a
Beckman DU640 Spectrophotometer (Beckman coulter, USA) as described previously [6].
Soluble protein extract was dialyzed against Tris-Cl buffer (VISKING dialysis tubing, 14 kDa
cutoff, SERVA). The final assay (1 mL) was performed in a quartz cuvette at 22°C and contained
1 mM geraniol or 1.5 mM citral, 1 mM NAD+ (final concentrations) and various amounts of
protein. Reactions were started by addition of NAD+. Rates were calculated based on a molar
extinction coefficient of 6220 M-1 cm-1.
Linalool isomerase activity was determined in soluble protein extracts. Glass vials (4 mL) with
300 to 500 μL sample were reduced with 5 mM dithionite, closed with butyl rubber stoppers and
flushed with nitrogen gas for 3 minutes, to provide anoxic conditions. Samples were incubated
for 20 minutes at room temperature. The reaction was started by addition of 200 μL geraniol (200
mM) in 2,2,4,4,6,8,8-heptamethylnonane (HMN) through a needle and incubated for 14 to 16
- 154 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
hours at 28°C under mild shaking. Product formation was determined in samples from the HMN
phase by gas chromatography with flame ionization detection (PerkinElmer Auto System XL,
Überlingen, Germany) on an Optima-5 column (30 m x 0.32 mm, 0.25 μm film thickness;
Macherey-Nagel, Germany) with the following temperature program: injection port 250°C,
detection port 350°C, initial column temperature 40°C for 2 min, increasing to 100°C at a rate of
4°C min-1, keeping 100°C for 0.1 min, followed by an increase to 320°C at 45°C min-1 and hold
for 3 min. The split ratio was set to 1:9.
Proteomics by MALDI-ToF MS
T. linaloolentis 47Lol cultures were grown on (R,S)-linalool to the late exponential phase and
harvested by centrifugation. Cells were suspended in Tris-Cl buffer (40 mM, pH 8.0) and soluble
protein extract was prepared as described above. Protein samples, obtained from individual
purifications,
were
analyzed
by
SDS-PAGE
coupled
with
matrix-assisted
laser
desorption/ionization time of flight (MALDI-ToF) mass spectrometry (MS). Protein bands in gels
were excised manually, and the Ettan Spot Handling Workstation (GE Healthcare) was used for
trypsin digestion and embedding of the resulting peptide solutions in an α-cyano-4hydroxycinnamic acid matrix for spotting onto MALDI targets. MALDI-ToF MS analysis was
performed on an AB SCIEX TOF/TOFTM 5800 Analyzer (AB Sciex / MDS Analytical
Technologies [31]. Spectra in a mass range from 900 to 3700 Da (focus 1700 Da) were recorded
and analyzed by GPS Explorer™ Software Version 3.6 (build 332, Applied Biosystems) and the
Mascot search engine version 2.4.0 (Matrix Science Ltd, London, UK) using the RAST draft
genome as reference.
Draft genome and gene analysis
Two publicly available sequencing datasets, ASM31020 (4.199 Mbp on 220 contigs, available
since November 2012) and ASM62130 (4.214 Mbp on 46 contigs, available since April 2014),
were merged using Sequencher 4.6 with a minimum match percentage of 95 % and a minimum
overlap of 50. The resulting draft genome was uploaded to RAST for further analysis [32, 33].
Known genes from Castellaniella defragrans 65Phen, Pseudomonas aeruginosa PAO1 and P.
citronellolis, encoding enzymes involved in the linalool and geraniol metabolism, were used to
- 155 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
identify homologs in T. linaloolentis 47Lol. Overall similarity of the identified genes to their
homologs was determined by blastp [34].
Competing interests
The authors declare that they have no competing interests.
Authors‘ contribution
RM, EPC and JH participated in the design of the study. RM carried out growth experiments,
enzyme measurements and bioinformatic work. EPC performed the transposon mutagenesis
experiment. SM and TS conducted mass-spectrometry based protein identifications and analyzed
data. RM and JH drafted the manuscript. All authors read and approved the final manuscript.
Acknowledgments
This study was financed by the Max Planck Society. We thank Dirk Albrecht for MALDI-ToF
measurements.
- 156 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
References
1.
Kesselmeier J, Staudt M: Biogenic volatile organic compounds (VOC): An overview on
emission, physiology and ecology. Journal of Atmospheric Chemistry 1999, 33:23-88.
2.
Sharkey TD, Yeh S: Isoprene emission from plants. Annual Review of Plant Physiology and
Plant Molecular Biology 2001, 52:407-436.
3.
Marmulla R, Harder J: Microbial monoterpene transformations – A review. Frontiers in
Microbiology 2014, 5.
4.
Brodkorb D, Gottschall M, Marmulla R, Lüddeke F, Harder J: Linalool dehydratase-isomerase,
a bifunctional enzyme in the anaerobic degradation of monoterpenes. Journal of Biological
Chemistry 2010, 285:30436-30442.
5.
Lüddeke F, Harder J: Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences 2011, 66:409-412.
6.
Lüddeke F, Wülfing A, Timke M, Germer F, Weber J, Dikfidan A, Rahnfeld T, Linder D,
Meyerdierks A, Harder J: Geraniol and geranial dehydrogenases induced in anaerobic
monoterpene degradation by Castellaniella defragrans. Applied and Environmental
Microbiology 2012, 78:2128-2136.
7.
Petasch J, Disch E-M, Markert S, Becher D, Schweder T, Hüttel B, Reinhardt R, Harder J: The
oxygen-independent metabolism of cyclic monoterpenes in Castellaniella defragrans 65Phen.
BMC Microbiology 2014, 14:164.
8.
Förster-Fromme K, Jendrossek D: Identification and characterization of the acyclic terpene
utilization gene cluster of Pseudomonas citronellolis. FEMS Microbiology Letters 2006,
264:220-225.
9.
Förster-Fromme K, Jendrossek D: Catabolism of citronellol and related acyclic terpenoids in
pseudomonads. Applied Microbiology and Biotechnology 2010, 87:859-869.
10.
Höschle B, Jendrossek D: Utilization of geraniol is dependent on molybdenum in
Pseudomonas aeruginosa: evidence for different metabolic routes for oxidation of geraniol
and citronellol. Microbiology 2005, 151:2277-2283.
11.
Förster-Fromme K, Höschle B, Mack C, Bott M, Armbruster W, Jendrossek D: Identification of
genes and proteins necessary for catabolism of acyclic terpenes and leucine/isovalerate in
Pseudomonas aeruginosa. Applied and Environmental Microbiology 2006, 72:4819-4828.
12.
Foss S, Harder J: Thauera linaloolentis sp. nov. and Thauera terpenica sp. nov., isolated on
oxygen-containing monoterpenes (linalool, menthol, and eucalyptol) and nitrate. Systematic
and Applied Microbiology 1998, 21:365-373.
- 157 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
13.
Foss S, Harder J: Microbial transformation of a tertiary allylalcohol: Regioselective
isomerisation of linalool to geraniol without nerol formation. FEMS Microbiology Letters
1997, 149:71-75.
14.
Nazzaro F, Fratianni F, De Martino L, Coppola R, De Feo V: Effect of essential oils on
pathogenic bacteria. Pharmaceuticals 2013, 6:1451-1474.
15.
Bakkali F, Averbeck S, Averbeck D, Waomar M: Biological effects of essential oils - A review.
Food and Chemical Toxicology 2008, 46:446-475.
16.
Sikkema J, Debont JAM, Poolman B: Mechanisms of membrane toxicity of hydrocarbons.
Microbiological Reviews 1995, 59:201-222.
17.
Nikaido H: Molecular basis of bacterial outer membrane permeability revisited.
Microbiology and Molecular Biology Reviews 2003, 67:593-656.
18.
van den Berg B: Going forward laterally: Transmembrane passage of hydrophobic molecules
through protein channel walls. Chembiochem 2010, 11:1339-1343.
19.
Foss S, Heyen U, Harder J: Alcaligenes defragrans sp. nov., description of four strains isolated
on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alpha-phellandrene)
and nitrate. Systematic and Applied Microbiology 1998, 21:237-244.
20.
Macheboeuf P, Contreras-Martel C, Job V, Dideberg O, Dessen A: Penicillin binding proteins:
key players in bacterial cell cycle and drug resistance processes. FEMS Microbiology Review
2006, 30: 673-691.
21.
Sauvage E, Kerff F, Terrak M, Ayala JA, Charlier P: The penicillin-binding proteins: structure
and role in peptidoglycan biosynthesis. FEMS Microbiology Review 2008, 32: 234-258.
22.
Lubelski J, Konings WN, Driessen AJM: Distribution and physiology of ABC-type
transporters contributing to multidrug resistance in bacteria. Microbiology and Molecular
Biology Reviews : MMBR 2007, 71:463-476.
23.
Martinez JL, Sánchez MB, Martínez-Solano L, Hernandez A, Garmendia L, Fajardo A, AlvarezOrtega C: Functional role of bacterial multidrug efflux pumps in microbial natural
ecosystems. FEMS Microbiology Review 2009, 33: 430-449.
24.
Ramos JL, Duque E, Gallegos M-T, Godoy P, Ramos-González MI, Rojas A, Terán W, Segura A:
Mechanisms of solvent tolerance in Gram-negative bacteria. Annual Review of Microbiology
2002, 56:743-768.
25.
Förster-Fromme K, Jendrossek D: AtuR is a repressor of acyclic terpene utilization (Atu) gene
cluster expression and specifically binds to two 13 bp inverted repeat sequences of the atuAatuR intergenic region. FEMS Microbiology Letters 2010, 308: 166-174.
- 158 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
26.
Grunden AM, Self WT, Villain M, Blalock JE, Shanmugam KT: An analysis of the binding of
repressor protein ModE to modABCD (Molybdate transport) operator/promoter DNA of
Escherichia coli. Journal of Biological Chemistry 1999, 274:24308-24315.
27.
Soares-Castro P, Santos PM: Deciphering the genome repertoire of Pseudomonas sp. M1
toward β-myrcene biotransformation. Genome Biology and Evolution 2015, 7:1-17.
28.
Kim JM, Marshall MR, Cornell JA, Iii JFP, Wei CI: Antibacterial activity of carvacrol, citral,
and geraniol against Salmonella typhimurium in culture medium and on fish cubes. Journal
of Food Science 1995, 60:1364-1368.
29.
Larsen R, Wilson M, Guss A, Metcalf W: Genetic analysis of pigment biosynthesis in
Xanthobacter autotrophicus Py2 using a new, highly efficient transposon mutagenesis system
that is functional in a wide variety of bacteria. Archices of Microbiology 2002, 178:193-201.
30.
Metcalf WW, Jiang W, Daniels LL, Kim S-K, Haldimann A, Wanner BL: Conditionally
replicative and conjugative plasmids carrying lacZα for cloning, mutagenesis, and allele
replacement in bacteria. Plasmid 1996, 35:1-13.
31.
Wolf C, Hochgräfe F, Kusch H, Albrecht D, Hecker M, Engelmann S: Proteomic analysis of
antioxidant strategies of Staphylococcus aureus: Diverse responses to different oxidants.
Proteomics 2008, 8:3139-3153.
32.
Aziz RK, Bartels D, Best AA, DeJongh M, Disz T, Edwards RA, Formsma K, Gerdes S, Glass
EM, Kubal M et al: The RAST server: Rapid annotations using subsystems technology. BMC
Genomics 2008, 9.
33.
Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA, Gerdes S, Parrello B,
Shukla M et al: The SEED and the Rapid Annotation of microbial genomes using Subsystems
Technology (RAST). Nucleic Acids Research 2014, 42:D206-D214.
34.
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ: Basic local alignment search tool.
Journal of Molecular Biology 1990, 215:403-410.
- 159 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Figure S1: SDS-PAGEs (A, B) of cell-free protein extract of T. linaloolentis 47Lol grown on 1 mM
(R,S)-linalool and 10 mM nitrate to the late exponential phase. Indicated bands were extracted and
analyzed by MALDI-ToF MS. Marker is given in kDa. (1) AtuF - geranyl-CoA carboxylase alpha subunit,
(2) LiuD - methylcrotonyl-CoA carboxylase biotin-containing subunit, (3) AtuA - 3-hydroxy-3isohexenylglutaryl-CoA:acetate lyase, (4) AtuC - geranyl-CoA carboxylase beta subunit, (5) LiuA isovaleryl-CoA dehydrogenase, (6) GeoA - geraniol dehydrogenase, (7) Lis - linalool isomerase, (8) LiuB
- methylcrotonyl-CoA carboxylase carboxyl transfer subunit.
- 160 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Table S1: Transposon insertion mutants.
Mutant
Length
[bp]
NCBI
Protein
accession
annotation
Probable function
DNA modification /
transcriptional control
Integration host factor,
D-subunit
Transcription regulation
DNA-repair protein;
DNA repair
Type III restriction enzyme
mechanism
ENO89557
DNA-binding domain of
ModE
Transcription regulation
(molybdenum metabolism)
675
ENO85695
Trancriptional regulator,
TetR family
Transcription regulation
63
294
ENO87982
XRE family transcriptional
regulator
Transcription regulation
(response to xenobiotics)
64
675
ENO85695
Trancriptional regulator,
TetR family
Transcription regulation
83
1074
ENO88415
DNA sulfur modification
protein DndB
DNA binding and
recognition of modification
sites
88
2568
ENO89640
PAS/PAC sensor hybrid
histidine kinase
Signaling and transcription
regulation
91
1542
ENO89296
ATP-dependent
DNA repair mechanism
endonuclease family protein
93
903
ENO82884
Integrase core domain
(Transposase)
18
309
ENO90120
36
2124
ENO88423
41
813
47
- 161 -
DNA mobility
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Mutant
Length
[bp]
NCBI
Protein
accession
annotation
Probable function
Transport
37
2433
ENO85793
TonB-dependent
Transport
siderophore receptor (FpvA)
38
2430
ENO90450
TonB-dependent
siderophore receptor
Transport
60
1332
ENO85236
Branched-chain amino acid
transport system permease
protein LivM
Branched-chain amino acid
transport
ENO88656
Membrane protein
belonging to the AzlC
superfamily
Branched-chain amino acid
transport
Branched-chain amino acid
transport
69
705
73
705
ENO88656
Membrane protein
belonging to the AzlC
superfamily
80
1140
ENO86020
ABC-type multidrug
transprort system
Transport
81
2475
ENO86288
TonB-dependent
siderophore receptor
Transport
- 162 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Mutant
Length
[bp]
NCBI
Protein
accession
annotation
Probable function
Membrane integrity
21
1971
ENO84346
Penicillin-binding protein 2
Peptidoglycan synthesis
(cell elongation)
Lipooligosaccharide core
biosynthesis
52
963
ENO84489
ADP-L-glycero-D-mannoheptose-6-epimerase
(EC 5.1.3.20)
68
1386
ENO89893
UDP-phosphate galactose
phosphotransferase
Lipopolysachharide Oantigen synthesis
85
1386
ENO89893
UDP-phosphate galactose
phosphotransferase
Lipopolysachharide Oantigen synthesis
Miscellaneous
54
1548
ENO84316
ATP-dependent RNA
helicase
RNA metabolism
58
264
ENO89266
Lysophospholipase
(EC 3.1.1.5)
Phospholipid metabolism
61
16533
ENO89831
Putative large exoprotein
Similarity to
involved in heme utilization
hemagglutinin-like protein;
or adhesion of
involved in cell adhesion
ShlA/HecA/FhaA family
62
1041
ENO84082
Peptidase Gluzincin family
(MA2)
Protein metabolism
ENO84083
Fic (Filamentation induced
by cAMP) family protein
Regulatory mechanism of
cell division, folate
metabolism
66
1146
- 163 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Mutant
67
87
Length
[bp]
504
504
NCBI
Protein
accession
annotation
ENO85121
ENO85121
Probable function
Putative phasin protein
Surface proteins of
intracellular storage
granules
Putative phasin protein
Surface proteins of
intracellular storage
granules
Unclassified
28
7296
ENO88318*
Hypothetical protein
unknown
43
1812
ENO90436*
Putative ATP and DNAbinding domain
DNA metabolism
46
1209
ENO90599
Membrane protein,
Acyltransferase
unknown
1524;
Reverse transcriptase;
Unknown;
927
ENO87300;
ENO87299
Threonine dehydratase
Amino acid metabolism
1104
ENO86348
Hypothetical protein
unknown
ENO88413
Hypothetical protein
Motif for dnd systemassociated protein 4 within
DNA sulfur modification
system
ENO87344
von Willebrand factor A
containing CoxE domainlike protein
unknown
Similarity to labA-like
proteins (NYN domain)
Unknown (might be
involved in RNA core
metabolism; RNA
processome)
53
55
70
71
525
1179
76 #
77
819
ENO89891
- 164 -
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
Mutant
Length
[bp]
NCBI
Protein
accession
annotation
79
1170
ENO89611
Hypothetical protein
Unknown (DUF2863)
84
1104
ENO86348
Hypothetical protein
Unknown
* DNA sequence identical but translation is incorrect in NCBI
#
no sequence was obtained from the sequencing for mutant 76
- 165 -
Probable function
Chapter VII - The anaerobic linalool metabolism in Thauera linaloolentis 47Lol
- 166 -
Chapter VIII - General discussion and outlook
Chapter VIII
General discussion and outlook
Linalool dehydratase/isomerase
The linalool dehydratase/isomerase was successfully expressed in E. coli BL21(DE3) and
purified to homogeneity in quantities satisfactory for crystallography. Overall yields of about 0.3
% of the soluble protein were still low and improvement is desirable for future experiments or a
biotechnological application. An increase in yield is unlikely to be achieved by the current
expression system and purification scheme without large up-scaling. Purification by an Nterminal tag resulted in a non-active enzyme. Due to the absence of cofactors and
posttranslational modifications, I assume the absence of the signal peptide and the presence of the
Sumo peptide in Sumo-Ldi expression as major cause for the loss of activity.
The Ldi structure showed the formation of a disulfide bond between C48 and C101, contributing
to the formation of a loop which points towards the active cavity of the adjacent subunit in the
pentameric assembly. Located within this loop are amino acids likely to be involved in catalysis.
Formation of disulfide bonds is a spontaneous process under oxidative conditions and needs to be
controlled and regulated to assure correct folding and functionality of proteins. Oxidative protein
folding in the periplasm of Gram-negative bacteria involves four disulfide bond proteins (Dsb),
acting as the DsbA-DsbB system, which aids disulfide bond formation, and the DsbC-DsbD
system, which catalyzes disulfide bond isomerization from a non-native to native state (Kojima et
al., 2001, Denoncin and Collet, 2012). In contrast to the periplasm, the cytoplasm is kept in a
reduced state and disulfide bond formation does not occur. The removal of the signal peptide in
the Sumo-Ldi resulted in a cytoplasmic expression, thereby preventing the formation of disulfide
bonds and thus resulted in a misfolded protein. I explain the low residual activity of the Ldi by
the “accidentally” correct formation of the disulfide bond between cysteine 48 and cysteine 101
after cell disintegration and exposure to molecular oxygen. The lower activity of the N-terminally
truncated Ldi (without signal peptide) compared to the wildtype enzyme supports this hypothesis.
The initial presence of the Sumo peptide might have had an additional negative effect on the
folding.
- 167 -
Chapter VIII - General discussion and outlook
Nevertheless, overall purification might be improved by use of a C-terminal tag. The determined
Ldi structure showed a solvent accessible C-terminus (chapter IV), which was not buried inside
the structure and therefore potentially accessible for a purification tag without having a negative
effect on the folding. Most purification tags are available for N- as well as C-terminal protein
expression (Arnau et al., 2006, Young et al., 2012). Ldi expression with a C-terminal purification
tag and the natural signal peptide on the N-terminus would promote the expression into the
periplasm and most likely result in correct formation of the disulfide bond (C48, C101) and a
correct overall folding of the Ldi.
A homo-pentameric structure was elucidated for the Ldi and a mechanism was proposed.
However, the detailed mechanism and the direct amino acid-substrate interactions remained
undisclosed. To study the enzyme in action, a snapshot of the substrate bound in the active site is
desired. Such a snapshot is achieved by using substrate and intermediate analogs which interact
with active amino acid residues, are not turned over, but are covalently linked to them instead.
The mechanism of monoterpene synthases was studied by use of fluorinated geranyl- or linalyl
diphosphate, homolinalyl diphosphate, 6,7-dihydrogeranyl diphosphate and 3-aza-geranyl
diphosphate (Schwab et al., 2001, Whittington et al., 2002, Karp et al., 2007). No suitable
analogs for E-myrcene, linalool or geraniol were available and we conducted initial experiments
with linalool soaked Ldi crystals. Only refraction data for a few crystals with a bound
monoterpene was obtained. It was questionable if the bound mass was still linalool or if it was
converted to the thermodynamically more stable E-myrcene or to geraniol. However, the initial
hypothesis of a shared active site between two subunits was confirmed as the mass of the bound
substrate was located accordingly. The available structure also gives indication to the oxygensensitivity of the enzyme. While C48 and C101 form a disulfide bond and support the tertiary
structure, C170 and C179 are present in the free thiol state and contribute to the catalysis.
Oxidation of all cysteine residues results consequently in a non-active enzyme.
- 168 -
Chapter VIII - General discussion and outlook
Biotechnological application for the linalool dehydratase/isomerase and the linalool isomerase
Isoprene and butadiene are large volume intermediates in industry, as they are used as starting
material in many synthetic processes. Annual butadiene and isoprene productions of over 800000
tons were reported (Lee et al., 2011). These values relate to a market value of over 20 billion and
4 billion USD for butadiene and isoprene, respectively. There is a growing interest in the
production of these chemicals from sugars as biotechnological fermentation products, e.g.
butandiol. Information on enzymes forming dienes by dehydration from alcohols is still scarce. A
well-described enzyme catalyzing such a reaction is the linalool dehydratase/isomerase
(Brodkorb et al., 2010). The importance of the enzyme is reflected by the number of pending and
accepted patents. Linalool dehydratase/isomerase was described in multiple patents as an
example for alkenol dehydration. Industrial relevant reactions are the dehydration of 3-methyl-2buten-1-ol (prenol), 2-methyl-3-buten-2-ol or 3-buten-2-ol to isoprene or 1,3-butadiene,
respectively (Fig. 1) (Garcez et al., 2013, Coelho et al., 2014, Lopes et al., 2014, Pearlman et al.,
2014, Botes and van Eck Conradie, 2015a, Botes and van Eck Conradie, 2015b, Koch et al.,
2015a, Koch et al., 2015b, Marlière et al., 2015, Osterhout et al., 2015). A few patents claim to
make already use of the Ldi or a derivative thereof heterologously expressed in E. coli. The
inventors state that enzymes, derived from the Ldi, catalyze the dehydration of alkenol substrates
to the corresponding hydrocarbons, with the general formula CnH2nO and CnH2n-2 (3 < n < 7),
respectively. The formation of isoprene from 2-methyl-3-buten-2-ol and prenol or the formation
of 1,3-butadiene from 3-buten-2-ol were described. The natural substrates of the linalool
dehydratase/isomerase (E-myrcene, (S)-linalool, geraniol) were only poorly accepted by these
engineered enzymes (Bredow et al., 2014, Campbell et al., 2014, Marlière, 2014, Marlière et al.,
2014, Marlière, 2015, Marlière et al., 2015). The conversion of 2-methyl-3-buten-2-ol or 3methyl-2-buten-1-ol to isoprene resembles the conversion of linalool or geraniol to myrcene,
respectively, having a C5 instead of a C10 substrate. While the aforementioned molecules share
structural features, 1,3-butadiene and 3-buten-2-ol lack a methyl group. It is doubtful, if these
molecules are accepted by Ldi-derived enyzmes without protein engineering by amino acid
exchange. However, the use of the Ldi for biotechnological processes is advised, as the enzyme
expresses a natural affinity and tolerance towards the hydrophobic substrates and products. With
knowledge of the protein structure, mutations can be introduced by a targeted approach to
achieve high-yield conversion rates for hemiterpenes and the production of isoprene but also to
broaden the substrate and product spectrum to yield the desired 1,3-butadien.
- 169 -
Chapter VIII - General discussion and outlook
Figure 1: Dehydration reactions of alkenols to alkenes. The upper panel shows the isomerization of 3methyl-2-buten-1-ol (prenol) to 2-methyl-3-buten-2-ol and its dehydration to 2-methyl-1,3-butadien
(isoprene). The lower panel shows the dehydration of 3-buten-2-ol to 1,3-butadiene.
Besides the production of bulk chemicals like isoprene and 1,3-butadiene, there is also a growing
demand on the biosynthesis of fine chemicals like (optically active) tertiary alcohols. Their
synthesis is challenging. While chemical synthesis needs to be tightly controlled and resulting
racemic mixtures need to be separated, biocatalysts can be used to produce enantiopure products.
A repertoire of enzymes has been applied for this purpose, e.g. esterases and lipases (Kourist and
Bornscheuer, 2011). Studies on the degradation of fuel-oxygenates indicated the presence of
isomerases of microbial origin, converting 3-methyl-2-buten-1-ol (prenol) to 2-methyl-3-buten-2ol (Malone et al., 1999, Schuster et al., 2012). The linalool dehydratase/isomerase and the
linalool isomerase catalyze the reversible isomerization of a primary to a tertiary alcohol.
However, both enzymes as well as other biocatalysts have drawbacks with respect to their
biotechnological application. Protein engineering is used to optimize biocatalysts to broaden the
substrate spectrum, to narrow the product spectrum and to increase the turnover and stability
(Reetz, 2011, Reetz, 2013). The Ldi, as a bifunctional enzyme, converts the product of the first
reaction, the tertiary alcohol (S)-linalool, into the hydrocarbon E-myrcene. The linalool isomerase
catalyzes the isomerization reaction only, but lacks stereospecificity. These enzymes can
potentially be manipulated to yield optically pure linalool or even broaden the substrate and
product spectrum to other desired alcohols.
- 170 -
Chapter VIII - General discussion and outlook
Linalool isomerase
The linalool isomerase was identified as a membrane-associated protein. The N-terminus
contains four transmembrane helices, while the remaining protein is localized in the cytoplasm.
This prediction from a candidate gene was supported by subcellular fractionation experiments.
The tight association towards the inner membrane hindered the purification to homogeneity.
Release of the active protein from the membrane was impaired due to the high sensitivity of the
enzyme towards commonly used detergents (Triton X-100, Tween20 or n-octyl-E-Dglucopyranoside). Membrane proteins constitute for about one third of total cell protein. Those
proteins are less well studied than soluble proteins and structural information is rather scarce
(Wallin and von Heijne, 1998) because their isolation is more challenging. Membrane-spanning
parts of such proteins need to be shielded from bulk solvent in order to preserve the native
conformation and structure. This is achieved by the use of detergents. A broad variety of
detergents was successfully applied in membrane protein purification and characterization,
ranging from simple amphiphilic molecules (e.g. Tween, Triton) to aliphatic sugars (e.g. n-octylE-D-glucopyranoside, n-dodecyl-E-D-maltopyranoside) (Privé, 2007). Even though, some
membrane proteins are still unstable in the presence of a detergent or undergo structural changes
often accompanied with a loss of functionality. Therefore, more complex amphiphilic molecules
were investigated for membrane protein research (Tribet et al., 1996, Popot, 2010). Lately,
amphiphilic polymers (amphipols) received more attention and were shown to be superior over
detergents (Chae et al., 2010b). In contrast to detergents, which assemble into monolayermicelles, amphipols form a bilayer, mimicking a cellular membrane and retaining the natural
protein fold. However, these amphipols are usually not strong enough to release the protein from
the membrane and need to be applied in combination with other detergents (Chae et al., 2010a,
Popot, 2010). Other nonconventional surfactants (e.g. Nanodiscs, amphipols or fluorinated
surfactants) were also successfully applied in membrane protein research (reviewed by Popot,
2010). I suggest testing those nonconventional surfactants alone and in combination with
detergents for future purification attempts. Heterologous expression of the full-length protein and
a N-terminally truncated version - representing the cytosolic domain only - in E. coli BL21(DE3)
were successful but the resulting proteins were inactive. No rare codons were identified in the lis
gene which leads to the assumption that the inactivity did not result from mis-translation.
However, the employed E. coli BL21(DE3) strain is widely used, novel derivatives designed for
expression of membrane proteins are available (reviewed by Rosano and Ceccarelli, 2014). The
- 171 -
Chapter VIII - General discussion and outlook
E. coli strains C41(DE3), CD43(DE3) and Lemo21(DE3) were shown to be superior over
BL21(DE3) with respect to expression of membrane and globular proteins (Miroux and Walker,
1996, Wagner et al., 2008) and should be tested for expression of the linalool isomerase.
Alternatively, membrane proteins can be intentionally expressed in inclusion bodies followed by
denaturing and careful refolding of the protein (Grisshammer and Tate, 1995, Rosano and
Ceccarelli, 2014).
The linalool isomerase is the second member in the enzyme class 5.4.4.4 within intramolecular
hydroxyl group transferases (EC 5.4.4.x). Linalool isomerase and the linalool dehydratase/
isomerase of C. defragrans 65Phen share most of their characteristics, like pH and temperature
optima, oxygen-sensitivity, cofactor-independency and the affinity for their substrate geraniol.
An amino acid based alignment revealed an identity of 20 % (Fig. S1 and S2). The mature
linalool dehydratase/isomerase consists of 371 amino acids and is predicted to be in the
periplasm, while the linalool isomerase consists of 644 amino acids and is predicted to be
membrane-associated with its C-terminal domain located in the cytoplasm. A secondary structure
prediction was performed for the mature Ldi (Fig. 2) and the Lis (Fig. 3) using the JPred tool
(Drozdetskiy et al., 2015).
Figure 2: Secondary structure prediction of the mature linalool dehydratase/isomerase, without signal
peptide. Helices are indicated in red and sheets are indicated in green.
- 172 -
Chapter VIII - General discussion and outlook
Figure 3: Secondary structure prediction of the linalool isomerase. Helices are indicated in red and sheets
are indicated in green.
- 173 -
Chapter VIII - General discussion and outlook
The predicted positions of helices for the Ldi matched the helices we observed in the crystal
structure (Table 1). Secondary structure prediction of Lis showed a high number of helices
similar to the Ldi. I therefore performed a sequence alignment of the mature Ldi and the Lis with
the Clustal Omega-tool (Sievers et al., 2011) in combination with a secondary structure
prediction (Fig. 4). Sequence similarity of Lis to Ldi was restricted to the C-terminal domain of
the enzyme. The secondary structure of the aligned sequences was in agreement with the
individual datasets and the conserved amino acids.
Table 1: Structural helices of the mature linalool dehydratase/isomerase compared to helices in the model
of the linalool isomerase.
Inner helices
Outer helices
C-terminal helix
Amino acid
(Ldi)
Length (AA)
Amino acid
(Lis)
Length (AA)
63 - 80
17
217 - 234
17
124 - 139
15
287 - 301
14
179 - 193
14
257 - 372
15
238 - 251
13
418 - 427
9
294 - 306
12
482 - 493
11
341 - 351
10
530 - 538
8
25 - 35
10
184 - 194
10
85 - 100
15
239 - 255
16
146 - 163
17
327 - 340
13
198 - 212
14
377 - 390
13
255 - 266
11
431 - 442
11
309 - 322
13
498 - 510
12
355 - 360
6
544 - 548
5
- 174 -
Chapter VIII - General discussion and outlook
Figure 4: Alignment and secondary structure prediction of the mature linalool dehydratase/isomerase and
linalool isomerase. Color indicates similarity. Helices are indicated in red and sheets are indicated in
green.
- 175 -
Chapter VIII - General discussion and outlook
Based on this similarity, I attempted to model the cytosolic domain of the linalool isomerase
(amino acid 139-644) with the I-Tasser (Iterative threading assembly refinement, V 4.3) tool,
using the structural data of the Ldi (single subunit) as a template (Yang et al., 2015). The
experimentally determined Ldi structure and the Lis model showed significant structural
similarity. Data evaluation revealed a C-score of -2.05 and a TM-score of 0.47 ± 0.15. C-score
range is between -5 and 2. Higher values indicate a higher confidence in the predicted structure.
A TM-score higher than 0.5 indicates a highly similar topology, while values lower than 0.17
indicate a random topology (Zhang and Skolnick, 2005). Even though, the C-score of the
obtained linalool isomerase structure is low, the TM-score suggests a similar topology to the
linalool dehydratase/isomerase. The C-terminal end of the linalool isomerase (last 100 amino
acids) did not seem to be properly modeled as the Ldi-template is shorter than the Lis domain
(Ldi: 371 amino acids, Lis domain: 506 amino acids) and did therefore not provide sufficient
template information. This may have lowered the C-score artificially. In the model, the Nterminus of the Lis model stretched out of the structure and would potentially continue in the four
transmembrane helices, by which the enzyme is anchored in the inner membrane. Ramachandran
plot analysis of the obtained structure showed 82.7 % of the amino acids in the favored regions
and additional 11.7 % in the allowed region. The superimposed structures of Ldi and Lis showed
the same (D,D)6 barrel conformation (Fig. 5) and a similar helix pattern was observed (Table 1).
The hydrophobic cavity formed by the inner six helices was conserved. Within this cavity, four
conserved amino acids (Ldi: Y65, M124, C170, E171, C179; Lis: Y220, M287, C349, E350,
C358) and three similar amino acids (Ldi: F69, H128, F176; Lis: H224, W246, Y355) matched in
the superimposed structures and in the sequence alignment (Fig. 7 A,B). From structural data of
the Ldi, we deduced amino acids potentially involved in the isomerization of geraniol to (S)linalool: Tyr65, Met124, Cys170, Glu171, Phe176, Cys179 and Tyr239. I have identified the
same or similar amino acids in the Lis model: Tyr220, Met287, Cys349, Glu350, Tyr355, Cys358
and Lys480 (Table 2, Fig. 7 and 8).
- 176 -
Chapter VIII - General discussion and outlook
Figure 5: Comparison the linalool dehydratase/isomerase structure (Ldi, yellow) and the linalool
isomerase model (Lis, blue). (A) left to right: front view of Ldi, Lis and superimposed. (B) left to
right: sideview of Ldi, Lis and superimposed. (C) left to right: rear view of Ldi, Lis and
superimposed.
- 177 -
Chapter VIII - General discussion and outlook
Table 2: Amino acids shared by the mature linalool dehydratase/isomerase and linalool isomerase in a
sequence alignment and in the superimposed structures.
Amino acid (Ldi)
Amino acid (Lis)*
Conserved in alignment
Conserved in structure*
Pro 59
Gln 214
No
Yes
Val 62
Ala 217
Yes
Yes
Tyr 65
Tyr 220
Yes
Yes
Phe 69
His 224
Yes
Yes
Tyr 72
Phe 227
Yes
Yes
Met 124
Met 287
Yes
Yes
His 128
Trp 291
Yes
Yes
Cys 170
Cys 349
Yes
Yes
Glu 171
Glu 350
Yes
Yes
Phe 176
Tyr 355
Yes
Yes
Gln 178
Val 357
Yes
Yes
Cys 179
Cys 358
Yes
Yes
Tyr 239
Lys 480
No
#
No #
Trp 243
Gly 420
No
Yes
#
Yes #
Val 292
Lys 480
Yes
Leu 294
Phe 482
Yes
Yes
Phe 298
Ala 486
Yes
Yes
Leu 340
Leu 528
No
Yes
Leu 341
Ala 529
No
Yes
*Structure of Lis cytoplasmic domain (AA 139 - 644) was modeled with the I-Tasser tool using the
structural data of the Ldi as template. Amino acid counts are given for the full length protein.
# Lys480 in Lis is not conserved in the alignment. However, the location of the side chain in the
superimposed structures is in accordance with Tyr239 of Ldi.
- 178 -
Chapter VIII - General discussion and outlook
Figure 6: Superimposed structures of the linalool dehydratase/isomerase (Ldi, yellow) and the
linalool isomerase model (Lis, blue). (A) Close up of the active centers. (B) Close up of active
center in Lis model with selected amino acids Tyr220, Met287, Cys349, Glu350, Tyr355, Cys358
and Lys480, which were identified as similar to the Ldi by sequence alignment and from the
superimposed structures.
Figure 7: (A) Superimposed structures with selected amino acids of Lis and Ldi. (B)
Superimposed structures with selected amino acids of Lis (labeled) and Ldi (unlabeled).
- 179 -
Chapter VIII - General discussion and outlook
Figure 8: (A) Superimposed structures with selected amino acids of Lis (unlabeled), Ldi
(labeled: Tyr65, Met124, Cys170, Glu171, Phe176, Cys179, Tyr239) and linalool fitted into the
cavity. (B) Close up of active center in Lis model with selected amino acids (Tyr220, His224,
Phe227, Met287, Trp291, Cys349, Glu350, Trp353, Tyr355, Cys358, Phe361, Phe424, Lys480,
Phe482), which were identified based on similarity to the Ldi and which cover the pocket,
pointing towards the substrate
The identity and similarity of the amino acids in the Ldi structure and the Lis model indicate a
similar reaction mechanism. A vital role of Cys170 and Cys179 in the Ldi was supported by
amino acid exchange experiments. Exchange of the individual residues to serine resulted in a
non-active enzyme. Cysteine 349 and C358 are the equivalents in the linalool isomerase model.
As the detailed hydroxyl mutase mechanism of the Ldi is still not disclosed, a mechanism for Lis
can only be speculated. Common amino acids in the active center may play a crucial role in
catalysis: two cysteines (Ldi: C170, C179; Lis: C349, C358), a glutamate (Ldi: E171 ; Lis: E350)
and a tyrosine (Ldi: Y65 ; Lis: Y220).
A mechanism for the geraniol isomerization to linalool may involve protonation of the geraniol
hydroxy group by Cys349. Release of a water molecule from the terpene results in a carbocation
with a positive charge on the C1 atom. A intramolecular shift of the positive charge would result
in the more stable tertiary linalyl cation. Nucleophilic attack of a solvent water molecule followed
- 180 -
Chapter VIII - General discussion and outlook
by a deprotonation leads to linalool. The reaction from linalool to geraniol could involve a
protonation of the linalool hydroxy group by Cys358 or Tyr355. Release of the resulting water
molecule, a concerted attack of a water molecule, coordinated by Cys349 and Glu350, onto the
C1 atom of the terpene and a deprotonation would lead to geraniol. The latter reaction would be
similar to this proposed for the linalool dehydratase/isomerase (chapter IV).
In contrast to the Ldi, which transforms (S)-linalool only, the Lis accepts both linalool isomers.
We have attributed the stereochemistry in Ldi to the loop of the neighboring subunit covering the
active site. This loop carries amino acids (Asp38, Phe39, Tyr44), which coordinate a catalytic
water molecule and thereby ensure stereospecific formation of (S)-linalool (Chapter IV). The
absence of this loop in the Lis model would result in an unguided nucleophilic attack of water to
C3 of the terpene and results in the formation of (R,S)-linalool.
- 181 -
Chapter VIII - General discussion and outlook
Second metabolic pathway for linalool in C. defragrans 65Phen
Castellaniella defragrans 65Phen is able to use a broad variety of monoterpenes as growth
substrate and became a model organism for anaerobic monoterpene degradation. However, the
underlying metabolism is still not yet fully disclosed. Besides acyclic monoterpenes, C.
defragrans 65Phen metabolizes mono- and bi-cyclic monoterpenes. Initial studies have shown
that a sp2-hybridized C1 atom is vital for the degradation of monocyclic monoterpenes (Heyen
and Harder, 1998). Recently, a pathway for limonene mineralization has been proposed. Initially,
limonene is oxidized to perillyl alcohol and further oxidized to perillic acid. The sp2-hybridized
C1 atom enables an oxidative ring-opening and further degradation by E-oxidation-like reactions
(Petasch et al., 2014). Similar ring-opening reactions have been proposed for cyclohexane
carboxylic acid degradation (Elshahed et al., 2001, Kung et al., 2014). Degradation of the acyclic
monoterpene E-myrcene involves the hydration to (S)-linalool and an isomerization to geraniol,
catalyzed by the linalool dehydratase/isomerase (Ldi). Geraniol is further oxidized to geranic acid
but the ultimate fate has not been investigated (Heyen and Harder, 2000, Brodkorb et al., 2010,
Lüddeke and Harder, 2011). The knock-out mutant C. defragrans 65Phen 'ldi did not grow Emyrcene or geraniol but grew on monocyclic monoterpenes (e.g. limonene, D-phellandrene)
(Lüddeke et al., 2012a) and on (R,S)-linalool. This suggested a novel metabolic pathway for
(R,S)-linalool.
In the presented work, I report a novel enzyme activity in C. defragrans 65Phen transforming
(R,S)-linalool into the monocyclic monoterpenes D-terpinene and terpinolene in an ATPdependent reaction. C. defragrans 65Phen has been shown to grow on these monoterpenes (Foss
et al., 1998). Potentially they can be metabolized in analogy to limonene. Similar cyclization
reactions have been described for monoterpene synthases. These enzymes are well known in
plants and received recently also attention in microbial cells. The underlying catalytic
mechanisms are well studied and topic of many research articles and reviews. Monoterpene
synthases possess a conserved DDXXD and NSA/DTE motif (Chen et al., 2011, Gao et al., 2012)
and share overall structural similarities of an D-helical bundle in which the active site is located
(Whittington et al., 2002, Hyatt et al., 2007, Kampranis et al., 2007). Besides these similarities,
monoterpene synthases are rather dissimilar in their primary structure (Yamada et al., 2015). Due
to this aspect it was not possible to identify a putative monoterpene synthase in the genome of C.
defragrans 65Phen based on sequence homology search. Monoterpene synthases accept either
- 182 -
Chapter VIII - General discussion and outlook
geranyl-, neryl- or linalyl diphosphate (Bohlmann and Gershenzon, 2009) as substrate and it can
be speculated that the formation of D-terpinene and terpinolene from linalool is catalyzed by one
novel enzyme or that linalool is first activated by a phosphotransferase and the resulting linalyl
diphosphate is used as substrate for a monoterpene synthase.
As the fate of geranic acid is not completely elucidated in C. defragrans 65Phen and genes
involved in the acyclic terpene utilization pathway have net been found in the genome (Heyen
and Harder, 2000, Petasch et al., 2014), it is questionable if there is a degradation pathway for
geranic acid. The transformation of (R,S)-linalool into a monocyclic monoterpene intermediate
enables a further degradation in analogy to limonene. From this perspective, the linalool
dehydratase/isomerase would serve rather in a process of detoxification by converting geraniol
and linalool into the less soluble myrcene. A first ring-cleavage of bicyclic monoterpenes (e.g.
pinene, carene) results also in monocyclic intermediates and I suggest a central metabolism for all
monoterpenes in C. defragrans 65Phen via a monocyclic monoterpene intermediate (Fig. 9).
- 183 -
Chapter VIII - General discussion and outlook
Figure 9: Proposed monoterpene degradation in Castellaniella defragrans 65Phen. E-myrcene (1) is
hydrated to (S)-linalool (2) and isomerized to geraniol (3) by the bifunctional linalool
dehydratase/isomerase (Ldi). Geraniol is further oxidized to geranial and geranic acid (4) by a geranioland geranial dehydrogenase (GeoA and GeoB), respectively (Lüddeke et al., 2012b). (R,S)-linalool is
transformed into D-terpinene (6) and terpinolene (7) in an ATP-dependent reaction sequence. These
monocyclic monoterpenes are designated to further degradation in analogy to limonene (8), which is
functionalized by the limonene dehydrogenase (ctmAB). The so formed perillyl alcohol is oxidized to
perllyl aldehyde (10) and perillic acid (11). This pathway and the subsequent degradation of perillic acid
was recently described (Petasch et al., 2014). Additionally, a first ring-cleavage of bicyclic monoterpenes
(e.g. carene (12) and pinene (13)) results in a monocyclic monoterpene and can be degraded like
limonene.
- 184 -
Chapter VIII - General discussion and outlook
References
ARNAU, J., LAURITZEN, C., PETERSEN, G. E. & PEDERSEN, J. 2006. Current strategies for the use
of affinity tags and tag removal for the purification of recombinant proteins. Protein Expression
and Purification, 48, 1-13.
BOHLMANN, J. & GERSHENZON, J. 2009. Old substrates for new enzymes of terpenoid biosynthesis.
Proceedings of the National Academy of Sciences, 106, 10402-10403.
BOTES, A. L. & VAN ECK CONRADIE, A. 2015a. Methods for biosynthesis of isoprene.
PCT/US2014/049786.
BOTES, A. L. & VAN ECK CONRADIE, A. 2015b. Methods for biosynthesizing 1,3-butadiene.
US20150079654.
BREDOW, S., DONESKE, S., LI, M., ZHOU, H., MONTICELLO, D. J. & CAMPBELL, P. 2014.
Microorganisms and processes for the conversion of glycerol to isoprene. WO2014015210.
BRODKORB, D., GOTTSCHALL, M., MARMULLA, R., LÜDDEKE, F. & HARDER, J. 2010. Linalool
dehydratase-isomerase, a bifunctional enzyme in the anaerobic degradation of monoterpenes.
Journal of Biological Chemistry, 285, 30436-30442.
CAMPBELL, P., BREDOW, S., ZHOU, H., DONESKE, S. & MONTICELLO, D. J. 2014.
Microorganisms and processes for the production of isoprene. US20140349362.
CHAE, P. S., GOTFRYD, K., PACYNA, J., MIERCKE, L. J. W., RASMUSSEN, S. G. F., ROBBINS, R.
A., RANA, R. R., LOLAND, C. J., KOBILKA, B., STROUD, R., BYRNE, B., GETHER, U. &
GELLMAN, S. H. 2010a. Tandem facial amphiphiles for membrane protein stabilization. Journal
of the American Chemical Society, 132, 16750-16752.
CHAE, P. S., RASMUSSEN, S. G. F., RANA, R. R., GOTFRYD, K., CHANDRA, R., GOREN, M. A.,
KRUSE, A. C., NURVA, S., LOLAND, C. J., PIERRE, Y., DREW, D., POPOT, J.-L., PICOT,
D., FOX, B. G., GUAN, L., GETHER, U., BYRNE, B., KOBILKA, B. & GELLMAN, S. H.
2010b. Maltose-neopentyl glycol (MNG) amphiphiles for solubilization, stabilization and
crystallization of membrane proteins. Nature Methods, 7, 1003-1008.
CHEN, F., THOLL, D., BOHLMANN, J. & PICHERSKY, E. 2011. The family of terpene synthases in
plants: a mid-size family of genes for specialized metabolism that is highly diversified throughout
the kingdom. The Plant Journal, 66, 212-229.
COELHO, P. S., FARROW, M. F. & SMITH, M. A. 2014. De novo metabolic pathways for isoprene
biosynthesis. PCT/US2013/067079.
DENONCIN, K. & COLLET, J.-F. 2012. Disulfide bond formation in the bacterial periplasm: major
achievements and challenges ahead. Antioxidants & Redox Signaling, 19, 63-71.
- 185 -
Chapter VIII - General discussion and outlook
DROZDETSKIY, A., COLE, C., PROCTER, J. & BARTON, G. J. 2015. JPred4: a protein secondary
structure prediction server. Nucleic Acids Research, 43, W389-W394.
ELSHAHED, M. S., BHUPATHIRAJU, V. K., WOFFORD, N. Q., NANNY, M. A. & MCINERNEY, M.
J. 2001. Metabolism of benzoate, cyclohex-1-ene carboxylate, and cyclohexane carboxylate by
“Syntrophus aciditrophicus” strain SB in syntrophic association with H2-using microorganisms.
Applied and Environmental Microbiology, 67, 1728-1738.
FOSS, S., HEYEN, U. & HARDER, J. 1998. Alcaligenes defragrans sp. nov., description of four strains
isolated on alkenoic monoterpenes ((+)-menthene, alpha-pinene, 2-carene, and alphaphellandrene) and nitrate. Systematic and Applied Microbiology, 21, 237-244.
GAO, Y., HONZATKO, R. B. & PETERS, R. J. 2012. Terpenoid synthase structures: a so far incomplete
view of complex catalysis. Natural Product Reports, 29, 1153-1175.
GARCEZ, L. M. S., SLOVIC, A. M., DE ANDRADE COUTINHO, P. L. & DE CASTRO
MORSCHBACKER, A. L. R. 2013. Modified microorganisms and methods of co-producing
butadiene with 1-propanol and/or 1,2-propanediol. WO2013192183
GRISSHAMMER, R. & TATE, C. G. 1995. Overexpression of Integral Membrane-Proteins for Structural
Studies. Quarterly Reviews of Biophysics, 28, 315-422.
HEYEN, U. & HARDER, J. 1998. Cometabolic isoterpinolene formation from isolimonene by
denitrifying Alcaligenes defragrans. Fems Microbiology Letters, 169, 67-71.
HEYEN, U. & HARDER, J. 2000. Geranic acid formation, an initial reaction of anaerobic monoterpene
metabolism in denitrifying Alcaligenes defragrans. Applied and Environmental Microbiology, 66,
3004-3009.
HYATT, D. C., YOUN, B., ZHAO, Y., SANTHAMMA, B., COATES, R. M., CROTEAU, R. B. &
KANG, C. 2007. Structure of limonene synthase, a simple model for terpenoid cyclase catalysis.
Proceedings of the National Academy of Sciences, 104, 5360-5365.
KAMPRANIS, S. C., IOANNIDIS, D., PURVIS, A., MAHREZ, W., NINGA, E., KATERELOS, N. A.,
ANSSOUR, S., DUNWELL, J. M., DEGENHARDT, J., MAKRIS, A. M., GOODENOUGH, P.
W. & JOHNSON, C. B. 2007. Rational conversion of substrate and product specificity in a Salvia
monoterpene synthase: Structural insights into the evolution of terpene synthase function. Plant
Cell, 19, 1994-2005.
KARP, F., ZHAO, Y., SANTHAMMA, B., ASSINK, B., COATES, R. M. & CROTEAU, R. B. 2007.
Inhibition of monoterpene cyclases by inert analogues of geranyl diphosphate and linalyl
diphosphate. Archives of Biochemistry and Biophysics, 468, 140-146.
KOCH, D. J., LOPES, M. S. G., GOUVEA, I. E., ZEIDLER, A. F. B., DUMARESQ, A. S. R. &
RODRIGUES, M. E. P. 2015a. Modified microorganisms and methods of using same for
anaerobic coproduction of isoprene and acetic acid. PCT/US2014/045097.
- 186 -
Chapter VIII - General discussion and outlook
KOCH, D. J., LOPES, M. S. G., GOUVEA, I. E., ZEIDLER, A. F. B., DUMARESQ, A. S. R. &
RODRIGUES, M. E. P. 2015b. Modified microorganisms and methods of using same for
anaerobic coproduction of isoprene and acetic acid. WO2015002977.
KOJIMA, R., OKUMURA, M. & INABA, K. 2001. Structural basis of disulfide bond formation in the
bacterial periplasm and mammalian ER. eLS. John Wiley & Sons, Ltd.
KOURIST, R. & BORNSCHEUER, U. T. 2011. Biocatalytic synthesis of optically active tertiary
alcohols. Applied Microbiology and Biotechnology, 91, 505-517.
KUNG, J. W., MEIER, A.-K., MERGELSBERG, M. & BOLL, M. 2014. Enzymes involved in a novel
anaerobic cyclohexane carboxylic acid degradation pathway. Journal of Bacteriology, 196, 36673674.
LEE, J. W., KIM, H. U., CHOI, S., YI, J. & LEE, S. Y. 2011. Microbial production of building block
chemicals and polymers. Current Opinion in Biotechnology, 22, 758-767.
LOPES, M. S. G., SLOVIC, A. M., GOUVEA, I. E., PEREZ, J. R. & PARIZZI, L. P. 2014. Modified
microorganisms and methods of making butadiene using same. PCT/US2012/070161.
LÜDDEKE, F., DIKFIDAN, A. & HARDER, J. 2012a. Physiology of deletion mutants in the anaerobic
beta-myrcene degradation pathway in Castellaniella defragrans. BMC Microbiology, 12.
LÜDDEKE, F. & HARDER, J. 2011. Enantiospecific (S)-(+)-linalool formation from beta-myrcene by
linalool dehydratase-isomerase. Zeitschrift für Naturforschung Section C - a Journal of
Biosciences, 66, 409-412.
LÜDDEKE, F., WÜLFING, A., TIMKE, M., GERMER, F., WEBER, J., DIKFIDAN, A., RAHNFELD,
T., LINDER, D., MEYERDIERKS, A. & HARDER, J. 2012b. Geraniol and geranial
dehydrogenases induced in anaerobic monoterpene degradation by Castellaniella defragrans.
Applied and Environmental Microbiology.
MALONE, V. F., CHASTAIN, A. J., OHLSSON, J. T., PONELEIT, L. S., NEMECEK-MARSHALL, M.
& FALL, R. 1999. Characterization of a Pseudomonas putida allylic alcohol dehydrogenase
induced by growth on 2-methyl-3-buten-2-ol. Applied and Environmental Microbiology, 65,
2622-2630.
MARLIÈRE, P. 2014. Production of volatile dienes by enzymatic dehydration of light alkenols.
PCT/EP2013/067727.
MARLIÈRE, P. 2015. Production of volatile dienes by enzymatic dehydration of light alkenols. US
14/195,738.
MARLIÈRE, P., ANISSIMOVA, M. & ALLARD, M. 2015. Method for the enzymatic production of 3buten-2-one. PCT/EP2014/064102.
MARLIÈRE, P., DELCOURT, M. & MAZALEYRAT, S. 2014. Alkenol dehydratase variants.
PCT/EP2014/060085.
- 187 -
Chapter VIII - General discussion and outlook
MIROUX, B. & WALKER, J. E. 1996. Over-production of proteins in Escherichia coli: Mutant hosts that
allow synthesis of some membrane proteins and globular proteins at high levels. Journal of
Molecular Biology, 260, 289-298.
OSTERHOUT, R. E., BURGARD, A. P., PHARKYA, P. & ANDRAE, S. 2015. Microorganisms and
methods for improving product yields on methanol using acetyl-CoA synthesis. WO2015084633.
PEARLMAN, P. S., CHEN, C., BOTES, A. & VAN ECK CONRADIE, A. 2014. Methods for
biosynthesizing 1,3-butadiene. WO2013188546.
PETASCH, J., DISCH, E.-M., MARKERT, S., BECHER, D., SCHWEDER, T., HÜTTEL, B.,
REINHARDT, R. & HARDER, J. 2014. The oxygen-independent metabolism of cyclic
monoterpenes in Castellaniella defragrans 65Phen. BMC Microbiology, 14, 164.
POPOT, J.-L. 2010. Amphipols, nanodiscs, and fluorinated surfactants: Three nonconventional
approaches to studying membrane proteins in aqueous solutions. Annual Review of Biochemistry,
79, 737-775.
PRIVÉ, G. G. 2007. Detergents for the stabilization and crystallization of membrane proteins. Methods,
41, 388-397.
REETZ, M. T. 2011. Laboratory evolution of stereoselective enzymes: A prolific source of catalysts for
asymmetric reactions. Angewandte Chemie International Edition, 50, 138-174.
REETZ, M. T. 2013. Biocatalysis in organic chemistry and biotechnology: Past, present, and future.
Journal of the American Chemical Society, 135, 12480-12496.
ROSANO, G. L. & CECCARELLI, E. A. 2014. Recombinant protein expression in Escherichia coli:
advances and challenges. Frontiers in Microbiology, 5, 172.
SCHUSTER, J., SCHÄFER, F., HÜBLER, N., BRANDT, A., ROSELL, M., HÄRTIG, C., HARMS, H.,
MÜLLER, R. H. & ROHWERDER, T. 2012. Bacterial degradation of tert-amyl alcohol proceeds
via hemiterpene 2-methyl-3-buten-2-ol by employing the tertiary alcohol desaturase function of
the Rieske nonheme mononuclear iron oxygenase MdpJ. Journal of Bacteriology, 194, 972-981.
SCHWAB, W., WILLIAMS, D. C., DAVIS, E. M. & CROTEAU, R. 2001. Mechanism of monoterpene
cyclization: Stereochemical aspects of the zransformation of noncyclizable substrate analogs by
recombinant (−)-limonene synthase, (+)-bornyl diphosphate synthase, and (−)-pinene synthase.
Archives of Biochemistry and Biophysics, 392, 123-136.
SIEVERS, F., WILM, A., DINEEN, D., GIBSON, T. J., KARPLUS, K., LI, W., LOPEZ, R.,
MCWILLIAM, H., REMMERT, M., SÖDING, J., THOMPSON, J. D. & HIGGINS, D. G. 2011.
Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal
Omega. Molecular Systems Biology, 7.
TRIBET, C., AUDEBERT, R. & POPOT, J.-L. 1996. Amphipols: Polymers that keep membrane proteins
soluble in aqueous solutions. Proceedings of the National Academy of Sciences, 93, 15047-15050.
- 188 -
Chapter VIII - General discussion and outlook
WAGNER, S., KLEPSCH, M. M., SCHLEGEL, S., APPEL, A., DRAHEIM, R., TARRY, M.,
HOGBOM, M., VAN WIJK, K. J., SLOTBOOM, D. J., PERSSON, J. O. & DE GIER, J. W.
2008. Tuning Escherichia coli for membrane protein overexpression. Proceedings of the National
Academy of Sciences of the United States of America, 105, 14371-14376.
WALLIN, E. & VON HEIJNE, G. 1998. Genome-wide analysis of integral membrane proteins from
eubacterial, archaean, and eukaryotic organisms. Protein Science, 7, 1029-1038.
WHITTINGTON, D. A., WISE, M. L., URBANSKY, M., COATES, R. M., CROTEAU, R. B. &
CHRISTIANSON, D. W. 2002. Bornyl diphosphate synthase: Structure and strategy for
carbocation manipulation by a terpenoid cyclase. Proceedings of the National Academy of
Sciences, 99, 15375-15380.
YAMADA, Y., KUZUYAMA, T., KOMATSU, M., SHIN-YA, K., OMURA, S., CANE, D. E. &
IKEDA, H. 2015. Terpene synthases are widely distributed in bacteria. Proceedings of the
National Academy of Sciences of the United States of America, 112, 857-62.
YANG, J., YAN, R., ROY, A., XU, D., POISSON, J. & ZHANG, Y. 2015. The I-TASSER Suite: protein
structure and function prediction. Nature Methods, 12, 7-8.
YOUNG, C. L., BRITTON, Z. T. & ROBINSON, A. S. 2012. Recombinant protein expression and
purification: A comprehensive review of affinity tags and microbial applications. Biotechnology
Journal, 7, 620-634.
ZHANG, Y. & SKOLNICK, J. 2005. TM-align: a protein structure alignment algorithm based on the TMscore. Nucleic Acids Research, 33, 2302-2309.
- 189 -
Chapter VIII - General discussion and outlook
Figure S1: Alignment of the linalool dehydratase/isomerase preprotein, including the signal peptide for
export, from C. defragrans 65Phen and the linalool isomerase from T. linaloolentis 47Lol. Color indicates
similarity.
Figure S2: Alignment of the mature linalool dehydratase/isomerase, including the signal peptide for
export, from C. defragrans 65Phen and the linalool isomerase from T. linaloolentis 47Lol. Color indicates
similarity.
- 190 -
Chapter VIII - General discussion and outlook
- 191 -
Acknowledgements
First, I would like to thank Prof. Dr. Jens Harder for the opportunity to perform my PhD on such
an interesting topic as the monoterpenes as well as for being a reviewer for my thesis. I really
appreciate your constant support. You were always there if I needed advice or for discussions.
Even though, I started already in 2008 with an internship on the monoterpene topic, it was never
boring as there was always a new feature to work on, and I´ve learned a lot of things over the
years. It was really an experience.
PD Dr. Ulrich Ermler, I am thankful for your help on the structure determination of the linalool
dehydratase/isomerase, an enzyme I became somehow addicted to over the years. I also thank
you for your time and effort to review my thesis and for being part of my thesis committee.
Prof. Dr. Uwe Nehls, thank you for constructive discussions throughout the committee meetings
and for your time and effort in being part of my thesis committee.
I thank Prof. Dr. Matthias Ullrich that he took the time to read my thesis and to join my thesis
committee.
I also thank Greta Giljan and Edinson Puentes for reading my thesis and being part of my thesis
committee.
I thank Prof. Dr. Friedrich Widdel for giving me the opportunity to perform my PhD in the
microbiology department of the MPI. Even though I barely took advantage of it, his door was
always open.
I would like to thank Dennis, Artur and Vladi for the pleasant atmosphere in the office during my
master thesis and the first month of my PhD. I really missed our discussions about scientific and
non-scientific topics as well as the office parties.
- 192 -
Of course, I also thank my “intermediate” office mates Iines and Deni. Besides the discussion on
your master thesis topics we laughed a lot about all different things and I really enjoyed your
company.
Last but not least, I thank my “current” office mates Edinson, Jana and Tanja for sharing an
exciting time with me. I especially like to thank Tanja for our discussion on recent topics in the
news and for sharing one or the other rumor, and Edinson for our discussion and conversations
about everything and nothing ;) I wish all of you the best while finishing your PhD and for the
future.
During my time in the microbiology department I have met a lot of different people and
unfortunately had to say goodbye to them as well. However, I would like to thank all of them,
creating a pleasant and warm atmosphere. There was always somebody to talk to, to discuss
certain issues but also to just make a joke or to get distracted when the workload became too
much. Special thanks go to Marina, Gao, Jan, Johannes, Richard and Stefano.
Without the help of our lovely technicians (Christina, Daniela, Ingrid, Ramona) a lot of things
wouldn´t have worked as well as they did. As a person, being in the institute rather early in the
morning, they were always the first faces I have seen and their smile gave me a good start into
the day. Special thanks go to Christina, who became also a friend over the years. Unfortunately, I
don´t remember the first coffee, we had together but I do remember a lot of “interesting” talks
and discussion during “tea kitchen meetings” ;)
I also thank you for showing me all the
techniques and always being a help in the lab.
Over the years, there have been a number of students who helped me a lot. Sometimes it was
quite frustrating but I believe all of you have learned something for the future, experienced that
lab-life is not always straight forward, but also I have learned some of things from you. I enjoyed
your company in the lab and the discussions we had. Without your contribution I would probably
not have come this far. I would like to thank all of you; Andreas, Barbara, Edinson, Irene and
Tina.
- 193 -
Thank you, Sina Weidenweber, to join the Linalool dehydratase/isomerase project. Without your
effort the project would have not been such a success. I know we were lucky to get nice crystals
more or less right away and that we were able to gain the information we needed to solve the
structure of this enzyme. But all luck and good wishes won´t help, if there is nobody to do the
work in a proper way. I wish you all the best for your future career.
I´d like to thank Manual Liebeke from the Symbiosis department for measuring one or the other
sample on the GC-MS and his help to interpret the data.
I want to thank all the participants of the MarMic program, especially Christiane and Karl-Heinz
as coordinators. I thank all lecturers and tutors as well as my MarMic class for this great
experience in my life, exciting lecture topics, fascinating lab courses and lovely moments inside
and outside of the office :)
Even though, I never met you personally, I thank Stephanie Markert, Dörte Becher and Thomas
Schweder from the University of Greifswald for their help with measuring my protein samples on
the MALDI-ToF MS.
Last but not least, I would like to thank all my friends for their constant support and for being
there when I had to get upset about something, to share my thoughts or just to distract me from
the everyday work life. Especially, Princess Linda for becoming a really a friend and nice as well
as entertaining evenings, talks and messages; and Greta basically for just being there, talking
about everything, cheering me up and sharing pleasant and unpleasant moments.
Of course, I also thank all other students and coworkers, I have met during my time in the
institute.
Selbstverständlich möchte ich mich auch ganz herzlich bei meiner Familie, speziell meiner Mutti
und meinen Großeltern, bedanken. Ohne euch wäre mein bisheriger Lebensweg wohl wesentlich
steiniger gewesen. Ich danke euch von ganzem Herzen für all eure Unterstützung, euren
Zuspruch und das Vertrauen in mich und meine Fähigkeiten.
- 194 -
Erklärung der selbstständigen Erarbeitung
Erklärung gemäß §6 Abs. 5 der Promotionsordnung (Stand 14.03.2007) der
Universität Bremen im Fachbereich 2 für Biologie/Chemie.
Hiermit versichere ich, dass ich die vorliegende Dissertation mit dem Titel „The
anaerobic linalool metabolism in the betaproteobacteria Castellaniella defragrans 65Phen
and Thauera linaloolentis 47Lol“
1. ohne unerlaubte fremde Hilfe selbstständig angefertigt habe.
2. keine anderen als die von mir angegebenen Quellen und Hilfsmittel benutzt habe.
3. die den benutzten Werken wörtlich oder inhaltlich entnommenen Stellen als solche
kenntlich gemacht habe.
Bremen,
(Robert Marmulla)
- 195 -