<html><head><meta name="color-scheme" content="light dark"></head><body><pre style="word-wrap: break-word; white-space: pre-wrap;">\makeatletter
\@ifundefined{HCode}
{\documentclass[screen,CRCHIM,Unicode,biblatex,published]{cedram}
\addbibresource{crchim20241041.bib}
\newenvironment{noXML}{}{}
\def\tsup#1{$^{{#1}}$}
\def\tsub#1{$_{{#1}}$}
\def\ndash{\text{--}}
\usepackage[T1]{fontenc}
\def\thead{\noalign{\relax}\hline}
\def\tbody{\noalign{\relax}\hline}
\def\endthead{\noalign{\relax}\hline}
\def\tabnote#1{\vskip4pt\parbox{.95\linewidth}{#1}}
\RequirePackage{etoolbox}
\def\jobid{crchim20241041}
%\graphicspath{{/tmp/\jobid_figs/web/}}
\graphicspath{{./figures/}}
\newcounter{runlevel}
\let\MakeYrStrItalic\relax
\csdef{Seqnsplit}{\\}
\def\refinput#1{}
\def\back#1{}
\def\botline{\\\hline}
\def\mn{\phantom{$-$}}
\def\0{\phantom{0}}
\usepackage{multirow} 
\def\nrow#1{\@tempcnta #1\relax%
\advance\@tempcnta by 1\relax%
\xdef\lenrow{\the\@tempcnta}}
\def\morerows#1#2{\nrow{#1}\multirow{\lenrow}{*}{#2}}
\usepackage{hyperref}
\makeatletter
\g@addto@macro{\UrlBreaks}{\UrlOrds}
\gappto{\UrlBreaks}{\UrlOrds}
\DOI{10.5802/crchim.407}
\datereceived{2024-12-13}
\daterevised{2025-03-22}
\dateaccepted{2025-05-23}
\ItHasTeXPublished
}
{\documentclass[crchim]{article}
\usepackage[T1]{fontenc}
\def\CDRdoi{10.5802/crchim.407}
\let\refinput\input
\makeatletter
\def\CDRsupplementaryTwotypes#1#2{}
}
\makeatother

\usepackage{upgreek}

\dateposted{2025-06-24}
\begin{document}

\begin{noXML}

\CDRsetmeta{articletype}{research-article}

\title{Regioselective hydration of geraniol by 
\textit{Escherichia coli} fumarases in whole-cell
biotransformations}

\alttitle{Hydratation r\'{e}gios\'{e}lective du g\'{e}raniol par les
fumarases d'\textit{Escherichia coli} dans des biotransformations en cellules
enti\`{e}res}

\author{\firstname{Natalie} \lastname{H\"{a}rterich}\IsEqualContrib}
\address{Institute of Biochemistry and Technical Biochemistry,
Department of Technical Biochemistry, University of Stuttgart, 70569
Stuttgart, Germany}

\author{\firstname{Philip} \lastname{Horz}\IsEqualContrib}
\addressSameAs{1}{Institute of Biochemistry and Technical Biochemistry,
Department of Technical Biochemistry, University of Stuttgart, 70569
Stuttgart, Germany}

\author{\firstname{Yingtong} \lastname{Fan}}
\addressSameAs{1}{Institute of Biochemistry and Technical Biochemistry,
Department of Technical Biochemistry, University of Stuttgart, 70569
Stuttgart, Germany}

\author{\firstname{Benjamin} \lastname{Aberle}\CDRorcid{0000-0002-1649-0425}}
\addressSameAs{1}{Institute of Biochemistry and Technical Biochemistry,
Department of Technical Biochemistry, University of Stuttgart, 70569
Stuttgart, Germany}
\address{Microbiology and Biotechnology at the Institute of Molecular
Physiology, Johannes Gutenberg University Mainz, 55128 Mainz, Germany}

\author{\firstname{Bernhard}\nobreakauthor\lastname{Hauer}\CDRorcid{0000-0001-6259-3348}\IsCorresp}
\addressSameAs{1}{Institute of Biochemistry and Technical Biochemistry,
Department of Technical Biochemistry, University of Stuttgart, 70569
Stuttgart, Germany}
\addressSameAs{2}{Microbiology and Biotechnology at the Institute of
Molecular Physiology, Johannes Gutenberg University Mainz, 55128 Mainz, Germany}
\email[B. Hauer]{hauerber@uni-mainz.de}

\keywords{\kwd{Fumarase}\kwd{Hydratase}\kwd{Water addition}\kwd{Geraniol}\kwd{Terpenes}\kwd{Alkenes}}

\altkeywords{\kwd{Fumarase}\kwd{Hydratase}\kwd{Addition d'eau}\kwd{G\'{e}raniol}\kwd{Terp\`{e}nes}\kwd{Alc\`{e}nes}}

\begin{abstract}
The regioselective hydration of carbon--carbon double bonds to generate
alcohols is a fundamental reaction in synthetic organic chemistry,
offering pathways to valuable secondary and tertiary alcohols.
Biocatalysis using hydratase enzymes, which add water to a double bond,
provides a selective and sustainable alternative to traditional
chemical methods. This study investigates the potential of
\textit{Escherichia coli} to hydrate the monoterpene geraniol in
whole-cell biotransformation systems. Through a targeted knockout
approach using the Keio collection, fumarases were identified as key
contributors to geraniol hydration. Overexpression studies further
revealed that FumA and FumB overexpression substantially enhanced
geraniol hydration activity at the terminal alkene, suggesting
promiscuity towards this non-native substrate. This result indicates an
expanded substrate scope of class I fumarases beyond their established
role in fumarate metabolism. By establishing a link between geraniol
hydration and specific genes, we aim to extend the enzymatic toolbox
for monoterpene transformations. Utilizing the inherent
regioselectivity and atom economy of fumarases, the potential of
fumarases as efficient biocatalysts in terpene modification could open
new avenues to advance applications in green chemistry and
biocatalysis.
\end{abstract}

\begin{altabstract}
L'hydratation r\'{e}gios\'{e}lective des doubles liaisons
carbone-carbone pour g\'{e}n\'{e}rer des alcools est une r\'{e}action
fondamentale en chimie organique synth\'{e}tique, qui permet d'obtenir
des alcools secondaires et tertiaires de grande valeur. La biocatalyse
utilisant des enzymes hydratases, qui ajoutent une mol\'{e}cule d'eau
\`{a} une double liaison, constitue une alternative s\'{e}lective et
durable aux m\'{e}thodes chimiques traditionnelles. Cette \'{e}tude
examine le potentiel d'\textit{Escherichia coli} \`{a} hydrater le g\'{e}raniol,
un monoterp\`{e}ne, dans des syst\`{e}mes de biotransformation \`{a}
cellules enti\`{e}res. Gr\^{a}ce \`{a} une approche de knock-out
cibl\'{e}e utilisant la collection Keio, les fumarases ont \'{e}t\'{e}
identifi\'{e}es comme des contributeurs cl\'{e}s \`{a} l'hydratation du
g\'{e}raniol. Des \'{e}tudes de surexpression ont en outre
r\'{e}v\'{e}l\'{e} que la surexpression de FumA et de FumB augmentait
significativement l'activit\'{e} d'hydratation du g\'{e}raniol au
niveau de l'alc\`{e}ne terminal, ce qui sugg\`{e}re une promiscuit\'{e}
avec ce substrat non natif. Ce r\'{e}sultat indique que les fumarases
de classe I ont un champ d'action \'{e}largi au-del\`{a} de leur
r\^{o}le \'{e}tabli dans le m\'{e}tabolisme du fumarate. En
\'{e}tablissant un lien entre l'hydratation du g\'{e}raniol et des
g\`{e}nes sp\'{e}cifiques, nous visons \`{a} \'{e}tendre la bo\^{i}te
\`{a} outils enzymatique pour les transformations des monoterp\`{e}nes.
En utilisant la r\'{e}gios\'{e}lectivit\'{e} et l'\'{e}conomie d'atomes
inh\'{e}rentes aux fumarases, le potentiel des fumarases en tant que
biocatalyseurs efficaces dans la modification des terp\`{e}nes pourrait
ouvrir de nouvelles voies pour faire progresser les applications en
chimie verte et en biocatalyse.
\end{altabstract}

\thanks{German Federal Ministry of Education and Research (BMBF) (grant
number 031B1343A)}

\thanks{\textbf{Note.} This article is submitted at the invitation of
the editorial committee (Juliette Martin)}

\shortrunauthors

\maketitle

%{\vspace*{12pt}}

\twocolumngrid

\end{noXML}

\section{Introduction} \label{sec1}

The chemical synthesis of alcohols remains a long-standing challenge.
In addition to the time-consuming and sometimes costly preparation of
the partially toxic transition metal catalysts, harmful by-products can
be produced, or the reaction is dependent on multiple reaction steps
(protective group chemistry), harsh reaction conditions and \mbox{often} does
not reach the desired selectivity for chiral products~\cite{1,2,3,4}.
These challenges evoke a sustained interest to explore additional or
new approaches. Nature provides an ideal template, offering not only
sustainable reaction conditions but also remarkable selectivity often
provided by enzymes. The atom-economical, biocatalytic addition of
water to alkenes offers an environmentally friendly alternative to the
harsh reaction conditions of organic chemistry. Hydratases belong to
the enzyme class of hydro-lyases (EC 4.2.1.X) and enable uncomplicated
access to secondary and tertiary alcohols, thus opening up new
possibilities for their synthesis~\cite{5}. The enzymes are categorized
into two groups based on their mechanism, the first group comprises
hydratases that hydrate conjugated C${=}$C double bonds in
${\upalpha}$,${\upbeta}$-unsaturated carbonyl compounds by a
nucleophilic Michael addition. The second group catalyzes the addition
of water to non-activated C${=}$C double bonds by an electrophilic
addition~\cite{6,7}.

The second group of enzymes is present, for example, in the metabolism
of terpenes, where hydratases are involved in the conversion of
terpenes by hydrating specific C${=}$C double bonds, thereby forming
tertiary alcohols with high selectivity. A prominent example is that of
carotenoid hydratases, which catalyze the hydration of terminal prenyl
units. Based on their function and their natural substrates, these
hydratases are divided into two distinct evolutionary groups, the CrtC
superfamily and the CruF family. Both acyclic (e.g., lycopene) and
monocyclic (e.g., ${\upgamma}$-carotene) carotenoid substrates can be
converted to their corresponding hydroxy compounds (Figure~S1). An
interesting representative example of this enzyme group is the
membrane-bound CrtC from \textit{Rubrivivax gelatinosus}, which can
convert the C\tsub{40} carotenoid lycopene in a regioselective manner
to hydroxylycopene and 1,1-dihydroxylycopene and can therefore catalyze
both single and double hydration~\cite{8,9,10}. In addition to the
hydration of carotenoids, hydratases such as kievitone hydratase and
phaseollidine hydratase have been demonstrated to convert
isoflavonoids, including kievitone, xanthohumol, and phaseollidine, to
their corresponding hydroxy derivatives
(Figure~\ref{fig1})~\cite{7,11,12}. In addition to very large terpenes,
short-chain monoterpenes such as limonene can also be modified by
hydratases. Hydrations of limonene have already been observed in yeasts
and bacteria, but mostly in whole cell preparations, which means that
isolated enzyme studies are rare. An exceptional case is the
membrane-bound ${\upalpha}$-terpineol dehydratase from
\textit{Pseudomonas gladioli}, which was isolated in 1992~\cite{13}.
Although the enzyme was classified as a dehydratase, its natural
function was shown to be the hydration of limonene to
${\upalpha}$-terpineol. The only published work on the heterologous
expression of a limonene hydratase is by Chang et~al.\ who expressed a
limonene hydratase (LIH) from \textit{Geobacillus stearothermophilus}
in \textit{E.~coli} and \mbox{converted} limonene to terpineol~\cite{14,15}.
Another interesting case is the bifunctional enzyme linalool
dehydratase isomerase (LinD). This enzyme catalyzes the hydration of
myrcene to (\textit{S})-linalool and its further isomerization to
geraniol, as well as the respective reverse reactions, with the
formation of myrcene from geraniol being the thermodynamically
preferred process~\cite{16}. In addition to natural substrates,
shortened and extended linalool derivatives could also be
converted~\cite{16,17,18,19,20}.

\begin{figure*}
{\vspace*{-2pt}}
\includegraphics{fig01}
{\vspace*{-3pt}}
\caption{\label{fig1}Overview of different hydratases adding water to
C${=}$C double bonds of different terpenes, terpenoids and fatty acids
following the Markovnikov-rule (electrophilic addition).}
{\vspace*{-3pt}}
\end{figure*}

One of the most frequently used enzymes for the conversion of
non-activated C${=}$C double bonds can be found in the
cofactor-dependent fatty acid hydratases (FAHs). They catalyze the
regioselective addition of water to C${=}$C double bonds of unsaturated
fatty acids to form the corresponding hydroxy fatty acids~\cite{10,19}.
FAHs have attracted a lot of attention mainly due to their wide
distribution in food-safe microorganisms such as
\textit{Lactobacillus}~\cite{21}. The oleate hydratase from
\textit{Elizabethkingia meningoseptica}, discovered in 1962, is the
most extensively studied FAH. This enzyme catalyzes the conversion of
oleic acid to 10-hydroxystearic acid.\ The enzyme requires flavin
adenine dinucleotide (FAD) as a cofactor, but its redox state does not
change during the reaction. Enzyme engineering has already opened up a
broad substrate spectrum with different FAHs. Fatty acids with
different chain lengths (C\tsub{11}--C\tsub{22}) or  numbers of double
bonds (up to~6) as well as oleic acid derivatives with different head
groups could be converted~\cite{21,22,23,24,25}. In addition, shorter,
non-activated alkenes (C\tsub{5}--C\tsub{10}) were also converted to
the corresponding secondary alcohol using a carboxylic acid dummy
substrate~\cite{26,27,28}. Furthermore, the acceptance of alkynes,
internal alkenes, substituted alkenes, and styrene derivatives was
demonstrated with this dummy substrate~\cite{26,27}.\looseness=-1

Fumarases, also known as fumarate hydratases, are enzymes catalyzing
the reversible hydration of fumarate to malate, a reaction central to
the citric acid cycle and ubiquitous across all kingdoms of life
(Figure~\ref{fig2}A)~\cite{29}. Prominent examples include the porcine
fumarase, the \textit{Saccharomyces cerevisiae} fumarase, and the three
fumarases from \textit{Escherichia coli} (FumA, FumB, and FumC).
Fumarases are categorized into two classes: class~I, dimeric enzymes
that contain a 4Fe--4S cluster, and class~II, comprising tetrameric,
thermostable, cofactor-free enzymes that are well studied due to their
robustness~\cite{30,31,32}. Fumarases are known for their exceptional
stereoselectivity, and a narrow substrate scope~\cite{33,34}. From an
early stage, halogenated derivatives of \mbox{fumarate,} such as
\mbox{fluorofumarate} and chlorofumarate, have been demonstrated to be
accepted by class~II fumarases, such as the porcine fumarase, which
served as an early model enzyme (Figure~\ref{fig2})~\cite{35,36}. In
the case of FumA from \textit{E.~coli}, a member of class~I fumarases,
reports from 1994 state that fluorofumarate is a promiscuous substrate
for hydration~\cite{37}. Moreover, it was demonstrated that a
C${\equiv}$C triple bond can be accepted by both fumarase classes in
the hydration of acetylene dicarboxylate to oxalacetate~\cite{35}. It
was also shown that the epoxide
\textsc{l}-\textit{trans}-2,3-epoxysuccinate can be transformed to a
diol~\cite{38}.

\begin{figure}
{\vspace*{6pt}}
\includegraphics{fig02}
\caption{\label{fig2}(A) Fumarases catalyze the reversible hydration of
fumarate to (\textit{S})-malate. (B) The substrate scope reported for
fumarases is limited to structures very close to fumarate and malate.}
\end{figure}

Among the fumarases of \textit{E.~coli}, FumA and FumB are class~I
enzymes that share 90\% sequence homology~\cite{39}. FumC is a class~II
fumarase, like the fumarases found in eukaryotic organisms, and is
therefore structurally very different from FumA and FumB~\cite{31}. The
expression of the three \mbox{fumarases} in \textit{E.~coli} varies with
oxygen availability, carbon source, and growth conditions~\cite{37,40}.
While FumA predominantly mediates fumarate hydration under most
conditions, FumB expression is elevated under anaerobic conditions,
possibly due to its high affinity for (\textit{S})-malate. FumC is
thought to compensate for the limitations of FumA and FumB during iron
limitation, superoxide radical accumulation, or \mbox{elevated}
temperatures~\cite{30,37,40}. Notably, FumA and FumB, despite being
class~I fumarases, demonstrate catalytic efficiencies that are
comparable to those of the class~II fumarases, which are typically
faster than class~I~\cite{37}.

Industrially, fumarases have been utilized since the 1970s for the
production of (\textit{S})-malate. Processes employing immobilized
\textit{Brevibacterium flavum} cells and whole-cell
\textit{Corynebacterium glutamicum} have demonstrated large-scale
production capabilities, reaching outputs of up to 2000 tons
annually~\cite{41}. 

For the hydration of small dicarboxylic acids, fumarases are
well-established enzymes valued for their stability and industrial
utility. On the other hand, large terpene substrates are converted by,
for example, carotenoid and kievitone hydratases. Internal alkenes in
fatty acids can be hydrated with high selectivity and efficiency by
FAHs. From a synthetic point of view, there is still a gap for a more
general enzyme platform for the isoprene moiety in smaller molecules
and for enzymes that can selectively add water to linear monoterpenes.
Although the hydration of sesquiterpenes and geraniol has been observed
in several studies utilizing fungal fermentations, these activities
have yet to be attributed to specific genes or enzyme
sequences~\cite{42,43,44,45}. This study revisits the catalytic
potential of fumarases, proposing greater versatility regarding the
substrate scope than was previously anticipated, and a potential to
address the current gap in the hydration of monoterpenes.\looseness=-1

\section{Results and discussion} \label{sec2}

The hydration of geraniol was observed in \textit{E.~coli} strains
ITB94, BL21(DE3), and BW25113 with and without a vector system,
indicating that endogenous \textit{E.~coli} enzymes catalyze the
reaction. The hydration does not occur spontaneously, as verified by
buffer control experiments. This observation initiated an investigation
to identify the specific gene responsible for this activity. To achieve
this, we utilized the Keio collection, a comprehensive library of
single-gene deletions in \textit{E.~coli} K-12 BW25113~\cite{46}. From
the library of 3985 strains, we selected strains lacking a gene
annotated as hydratase-encoding (according to ecocyc.org) and tested 42
individual knockout strains. These strains were screened using a
{96-deep-well} plate assay coupled with GC-FID analysis.  The main
product of the reaction was identified through a preparative-scale
reaction (see Section~\ref{ssec44}), followed by isolation and
characterization using NMR (Figures S4 and S5). By this approach, the
strain lacking the enzyme responsible for the hydration activity should
be identified.

Screening the selected hydratase candidates revealed varying levels of
the hydration product across the knockout strains, with the notable
exception of three strains: those lacking a gene for one of the three
fumarases ($\Delta$\textit{fumA}, $\Delta$\textit{fumB},
$\Delta$\textit{fumC}; Figure~\ref{fig3}). In these knockouts, the
hydration product was entirely absent, strongly suggesting that
fumarase enzymes are responsible for the observed hydration of
geraniol.

\begin{figure*}
\includegraphics{fig03}
\caption{\label{fig3}Screening of various knockout strains from the
\textit{E.~coli} Keio collection to identify hydratases responsible for
the conversion of geraniol. The hydration activity of different
\textit{E.~coli} strains, lacking genes annotated as hydratases, was
measured using 96-deep-well screening (1~mM substrate, 1\% DMSO (v/v),
30~\textdegree C, 24~h, 300~rpm, extraction with 500~$\upmu$L
cyclohexane/ethylacetate 1:1).}
\end{figure*}

Given the complete loss of product formation in $\Delta$\textit{fumA},
$\Delta$\textit{fumB} and $\Delta$\textit{fumC} strains, we further
investigated the activity of each fumarase gene individually by
overexpressing them in \textit{E.~coli} ITB94, the strain in which the
hydration activity was first discovered. For this purpose, three
pDHE1650 plasmids were constructed, each harboring a gene encoding for
one of the fumarases (FumA, FumB or FumC). The overexpression was
confirmed in whole cells (Figure~S3). Given that the strain ITB94 still
contains all three fumarase genes within its genome, a background
conversion was expected and observed using an empty vector control
(pDHE1650 without gene insert). To optimize the reaction conditions for
clear differentiation between background and overexpressed enzyme
activity, experiments were conducted with 50~mg/mL cell concentration,
10~mM substrate concentration, and a reaction time of 24~h. Under these
conditions, the influence of the enzymes on geraniol conversion was
clearly detectable, as background activity remained sufficiently low
while significant hydration activity was observed for the overexpressed
enzymes. The hydration of geraniol at the terminal alkene position was
considerably increased by the overexpression of FumA (17-fold) and FumB
(22-fold), in {comparison} to the empty vector control
(Figure~\ref{fig4}). In contrast, overexpression of FumC did not
noticeably alter geraniol hydration activity. Besides the terminal
hydration product, we also detected an internal hydration product.
Overexpression of FumA and FumB also enhanced the formation of the
internal hydration product by 2.4-fold and 4.5-fold, respectively
(Figure~\ref{fig4}). With FumC overexpression, no {internal} hydration
product was \mbox{detected.}

\begin{figure}
{\vspace*{4pt}}
\includegraphics{fig04}
\caption{\label{fig4}Formation of the terminal (grey) and internal
(green) hydration products of geraniol with \textit{E.~coli} whole
cells overexpressing one of the three fumarases (FumA, FumB, FumC)
analyzed by GC-FID (10~mM substrate, 1\% DMSO (v/v), 30~\textdegree C,
24~h, 300~rpm, extraction with 500~$\upmu$L cyclohexane/ethylacetate
1:1). The values shown are average values from triplicates, the error
bars show the standard deviation.}
\end{figure}

FumA and FumB catalyze the hydration of geraniol to a similar extent,
while FumC does not {demonstrate} promiscuity with geraniol under the
tested conditions. The similar behavior of FumA and FumB in geraniol
hydration aligns with previous findings that these enzymes catalyze the
fumarase reaction with nearly identical kinetic
parameters~\cite{37,47}. This is also reasonable in view of the fact
that they share 90\% sequence similarity~\cite{39}. Notably, FumA and
FumB exhibited regioselectivity for terminal hydration, with FumA
achieving an 11.3-fold higher formation of the terminal hydration
product and FumB a 7.8-fold higher formation of the terminal product.
Slight differences between FumA and FumB have been reported before for
certain substrates, such as D-tartrate, which may reflect their
physiological specialization, with FumA being more relevant under
aerobic and FumB under anaerobic conditions~\cite{40}. 

In addition to the hydration of geraniol, the reduction of geraniol to
citronellol was also observed during the experiments. Among the
overexpressed fumarases, cells overexpressing FumB showed an increase
in geraniol reduction, while overexpression of FumA had little effect,
and overexpression of FumC resulted in decreased citronellol formation
(Figure~S2). These results, especially those with FumC, suggest an
indirect link between fumarase overexpression and geraniol reduction,
emphasizing the interconnected nature of metabolic networks. Further
investigation into the specific enzymes responsible for this reduction,
similar to the study presented here, could elucidate the underlying
mechanisms and expand our understanding of the metabolic context
surrounding these reactions.

\section{Conclusion} \label{sec3}

Our findings demonstrate that \textit{E.~coli} fumarases might exhibit
a broader substrate promiscuity than previously recognized. The
hydration of the terpenoid geraniol, observed with overexpression of
FumA and FumB, suggests an extension of the substrate range for these
enzymes, which are considered highly specific for fumarate and only a
few derivatives~\cite{33,34}. While the hydration of geraniol has been
previously reported using the marine fungus \textit{Hypocrea} sp.\ 
MFAac46-2 in a three-day fermentation~\cite{42}, our study represents
the first direct linkage of this activity to specific genes and their
enzyme sequences. The unexpected substrate conversion by these class~I
fumarases, despite their more thermolabile and oxygen-sensitive nature
compared to class~II fumarases, may encourage a reconsideration of
class~I fumarases as valuable tools in biocatalysis. Furthermore, this
discovery suggests that fumarases may contribute to side reactions
involving terpenes and alkenes in fermentative processes. To the best
of our knowledge, fumarases have not been tested for activity towards
terpenes before. And since fumarases are very fast-acting enzymes on
their natural substrates ($k_{\mathrm{cat}}$ of 3100
s$^{-1}$)~\cite{37}, the assay times used were often in the range of
seconds and minutes, which may be a reason why much slower promiscuous
reactions are not detected. The possibility of using fumarases in
terpene hydration and related reactions highlights their potential as
highly efficient biocatalysts, which are particularly valued for their
100\% atom economy---a key attribute in sustainable chemical processes.
These results underscore the promise of hydratases and fumarases as
versatile tools and encourage deeper exploration of their applications
in biocatalysis and green chemistry.

\section{Experimental section} \label{sec4}
\subsection{Materials} \label{ssec41}

All chemicals and solvents were purchased from different suppliers
(Alfa Aesar, Carl Roth GmbH, \mbox{Enamine,} Macherey-Nagel, Merck,
Sigma-Aldrich, Thermo Fisher, VWR) without further purification.
Phusion High-Fidelity DNA Polymerase and DpnI were purchased from New
England Biolabs. 

\subsection{Screening of Keio collection (knockout strains)} \label{ssec42}
\subsubsection{Expression}\label{sssec421}

The investigation of the Keio Collection took place in 96-deep-well
plates (DWPs). Therefore, overnight cultures were prepared in 96-DWPs
by inoculating 1~mL of lysogeny broth (LB) medium containing
30~$\upmu$g/mL kanamycin (knockouts) or 150~$\upmu$g/mL ampicillin
(empty vector) per well, with glycerol stocks of the knockout variants
and empty vector controls. The plates were incubated at 37~\textdegree
C and 300~rpm for 16~h in an orbital shaker. The main cultures in
96-DWPs were set up using TB-media containing either 30~$\upmu$g/mL
kanamycin (knockouts) or 150~$\upmu$g/mL ampicillin (empty vector),
inoculated with 1\% overnight culture, and incubated for 24~h at 
37~\textdegree C and 300~rpm in the orbital shaker. After 24~h of
incubation, the plates were centrifuged at 4000~{g} for 20~min at
4~\textdegree C, and the harvested cells were immediately used for
biotransformation experiments. 

\subsubsection{Biotransformation (screening)}\label{sssec422}

For biotransformations, the fresh cells were resuspended in 
495~$\upmu$L KPi-buffer (50~mM, pH 7.0) and 5~$\upmu$L of geraniol
substrate stock (100~mM substrate in DMSO; final concentration 1~mM)
were added and incubated for 24~h at 30~\textdegree C and 300~rpm. The
reactions were stopped by adding 500~${\upmu}$L of
cyclohexane/ethylacetate 1:1. Extraction took place by shaking and
inverting with subsequent phase separation by centrifugation at
4000~{g} for 10~min at room temperature. Finally, 200~${\upmu}$L of the
organic phase was transferred into autosampler glass vials with inlets
for GC-FID analysis.

\subsection{Cloning and biotransformations of fumarases}\label{ssec43}
\subsubsection{Cloning of FumA, FumB and FumC in pDHE1650 vector}\label{sssec431}

The genes for the fumarase types FumA, FumB and FumC were successfully
integrated into the pDHE1650 vector using Gibson Assembly~\cite{48}.
For PCR, the standard protocol of Phusion$^{\text{\textregistered}}$\
High Fidelity Polymerase was used. The PCR products were digested using
\textit{Dpn}I (1~$\upmu$L \textit{Dpn}I for 25~$\upmu$L PCR product,
4~h at 37~\textdegree C), purified and transformed via heat shock into
\textit{E.~coli} XL1-Blue and after sequencing into ITB94 (derivate of
commercial strain TG1, with L-rhamnose-isomerase knockout and two
unspecific ADH---${\Delta}$\textit{yahK} and
${\Delta}$\textit{yjgB}---knockouts). 

\subsubsection{Expression}\label{sssec432}

Single colonies were inoculated in 5~mL overnight cultures (LB-medium
with ampicillin 150~$\upmu$g/mL) and incubated at 37~\textdegree C and
180~rpm. Main cultures were set up using terrific broth (TB) medium
containing 150~$\upmu$g/mL ampicillin and 0.5~g/L L-rhamnose (500~mL
medium in 2 L Erlenmeyer flasks), inoculated with 1\% overnight culture
and incubated at 30~\textdegree C and 180~rpm for 24~h. Cells were
harvested at 10,000~{g} for 30~min at 4~\textdegree C and immediately
used for biotransformation experiments.

\subsubsection{Biotransformation}\label{sssec433}

To investigate the functionality and differences in fumarase
biotransformation, experiments were conducted after expression of the
cells at a 500~${\upmu}$L scale in 2~mL glass vials. The experimental
setup included 495~${\upmu}$L of 50~mg/mL whole-cell suspension (FumA,
B, C, or empty vector, in 50~mM NaPi, pH 7.4) mixed with 5~$\upmu$L of
geraniol stock (1 M in DMSO, final substrate concentration: 10~mM). The
buffer control was prepared with 495~${\upmu}$L NaPi (50~mM, pH 7.4)
and 5~${\upmu}$L of geraniol stock (final concentration 10~mM, in
DMSO). Each reaction was performed in triplicates and incubated at
30~\textdegree C for 24~h with shaking at 300~rpm. The reaction
mixtures were extracted with 500~${\upmu}$L of cyclohexane/ethylacetate
1:1, vortexed, and centrifuged at 4000~{g} at rt for 5~min.
Subsequently, 200~${\upmu}$L of the organic phase was transferred into
autosampler glass vials with inlets for further GC analysis.

\subsection{Semi-preparative biotransformation}\label{ssec44}

Semi-preparative biotransformations took place in 100~mL shot flasks
with 100~mL of reaction volume. Therefore, cells with empty vector were
cultivated as described in Section~\ref{ssec43}. Fresh harvested cells
were resuspended in KPi-buffer (50~mM, pH 7.0) to a cell concentration
of 40~mg$_{\mathrm{cww}}$/mL and 10~mM substrate (geraniol) was added.
The biotransformation was incubated for 5d at 30~\textdegree C at
200~rpm in an orbital shaker. The reaction was stopped by extracting
three times with cyclohexane/ethylacetate 1:1 (in total 1~L solvent)
and the pooled organic phase was concentrated {in~vacuo}. The
crude product was then dissolved in CH\tsub{2}Cl\tsub{2} and purified
by column chromatography on silica gel 60M 0.04--0.063~mm and with
cyclohexane/ethylacetate 8:1 as the eluent. The determination of the
product structures was achieved by NMR (additional figures in the SI).

\subsubsection{Terminal hydrated OH-geraniol}\label{sssec441}

Isolated product: 97.4~mg of yellow clear oil (56\% isolated yield):

$^{1}$H NMR (500~MHz, CDCl\tsub{3}) $\delta=5.13$ 
(t, $^{3}J_{\mathrm{H},\mathrm{H}}=$\ 7.1~Hz, 1H), 4.12 
(d, $^{3}J_{\mathrm{H},\mathrm{H}}=$\ 7.1~Hz, 2H), 1.69 
(s, 3H), 1.25 (s, 6H) ppm.

$^{13}$C NMR (125~MHz, CDCl\tsub{3}) ${\delta}=132.1$, 
124.2, 74.0, 59.9, 42.4, 41.7, 29.7 (2C), 21.1, 17.7 ppm.

The NMR data obtained are consistent with previously reported
values~\cite{49}.

\subsubsection{Internal hydrated OH-geraniol}\label{sssec442}

Isolated product: 6.47~mg of yellow clear oil (3.7\% isolated yield):

$^{1}$H NMR (500~MHz, CDCl\tsub{3}) ${\delta} = 5.12$  (t,
$^{3}J_{\mathrm{H},\mathrm{H}}=7.0$~Hz, 1H), 3.96 (m, 2H), 2.05 (m,
2H), 1.83  (ddd, $^{3}J_{\mathrm{H},\mathrm{H}}=14.3$, 7.0, 5.0~Hz,
1H), 1.69 (s, 4H), 1.63 (s, 3H), 1.54 (m, 2H), 1.26 (s, 3H) ppm.

$^{13}$C NMR (125~MHz, CDCl\tsub{3}) ${\delta}=132.6$, 124.1, 71.3,
61.3, 42.2, 39.7, 26.9, 25.7, 22.7, 17.7 ppm.

The NMR data obtained are consistent with previously reported
values~\cite{50}.

\subsection{Gas chromatography}\label{ssec45}

To analyze the products from biotransformations, gas chromatography
(GC) was used. Analysis was performed on a Shimadzu GC2010 instrument
using a ZB-5 column (Zebron-Phenomenex, 30~m $\times$ 0.25~mm, 
0.25~$\upmu$m film) and hydrogen as carrier gas. The GC was equipped
with a flame ionization detector (FID) set to 335~\textdegree C.
Injections of 1~$\upmu$L injection volume were performed with an inlet
temperature of 260~\textdegree C in split mode (split 1:10). The oven
temperature started at 110~\textdegree C with a gradient of
10~\textdegree C per min up to 300~\textdegree C, holding the end
temperature for 1~min.

\section*{Declaration of interests}

The authors do not work for, advise, own shares in, or receive funds
from any organization that could benefit from this article, and have
declared no affiliations other than their research organizations.

\section*{Funding}

This project has received funding from the German Federal Ministry of
Education and Research (BMBF) - 031B1343A.

\section*{Acknowledgements}

We thank Andreas Schneider for assistance with the evaluation of NMR
spectra and Nicolas D. Travnicek for assistance with chromatographic
isolation of the products from preparative biotransformations.

\CDRGrant[BMBF]{031B1343A}

\section*{Supplementary data}

Supporting information for this article is available on the journal's
website under \printDOI\ or from the author.  

\CDRsupplementaryTwotypes{supplementary-material}{\cdrattach{crchim-407-suppl.pdf}}

\back{}

\printbibliography
\refinput{crchim20241041-reference.tex}

\end{document}
</pre></body></html>