Scoping the Enantioselective Desymmetrization of a Poorly Water-Soluble Diester by Recombinant Pig Liver Esterase

Article abstract of DOI:10.1021/acs.oprd.8b00277

Previously, the biocatalytic desymmetrization of dimethyl cyclohex-4-ene-cis-1,2-dicarboxylate to (1S,2R)-1-(methoxycarbonyl)cyclohex-4-ene-2-carboxylic acid, an important intermediate toward the synthesis of biologically active molecules, had been well-c

Full text of DOI:10.1021/acs.oprd.8b00277

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Full Paper
Scoping the Enantioselective Desymmetrization of a Poorly
Water-Soluble Diester by Recombinant Pig Liver Esterase
Murray P. Meissner, Philipp Süss, Henrike Brundiek, John M. Woodley, and Jan von Langermann
Org. Process Res. Dev., Just Accepted Manuscript • DOI: 10.1021/acs.oprd.8b00277 • Publication Date (Web): 16 Oct 2018
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Scoping the Enantioselective Desymmetrization of a
Poorly Water-Soluble Diester by Recombinant Pig
Liver Esterase
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Murray P. Meissner,θ,β Philipp Süss,§ Henrike Brundiek,§ John M. Woodley,*,θ Jan von
Langermann*,†
θTechnical University of Denmark (DTU), Department of Chemical and Biochemical
Engineering, Søltofts Plads, Building 229, 2800 Lyngby, Denmark
βOxford Biotrans Ltd., 127 Olympic Avenue, Milton Park, OX14 4SA, United Kingdom
§Enzymicals AG, Walther-Rathenau-Str. 49a, 17489 Greifswald, Germany
†University of Rostock, Institute of Chemistry, Biocatalytic Synthesis Group, Albert-Einstein-
Str. 3a, 18059 Rostock, Germany
Corresponding Authors
jw@kt.dtu.dk (John M. Woodley) and jan.langermann@uni-rostock.de (Jan von Langermann)
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ABSTRACT. Previously, the biocatalytic desymmetrization of dimethyl cyclohex-4-ene-cis-1,2-
dicarboxylate to (1S,2R)-1-(methoxycarbonyl)cyclohex-4-ene-2-carboxylic acid, an important
intermediate towards the synthesis of biologically active molecules, had been well-characterized
in terms of pH and temperature optima and several aspects of process performance. Eventually
this promising reaction could convert 200 mM (40 g·L-1) of substrate with > 99.5% e.e. using the
recombinant pig liver esterase, ECS-PLE06, at a scale of 8.8 L. However, the precise influence of
substrate concentration and the poorly water-soluble nature of the substrate (approximately 60 mM
in water at 25 °C for structurally similar dimethyl 1,4-cyclohexanedicarboxylate) remained
elusive. Therefore, this work focuses on using a recently published methodology based on reaction
trajectory analysis to identify mass transfer limitations in this reaction. With the constraints of
mass transfer on space-time yield considered, it was possible to evaluate and improve biocatalyst
yield (mass of product per mass of biocatalyst) through the use of higher substrate concentrations.
Ultimately the complete conversion of approximately 75 g·L-1 substrate was achieved in 3.65 h
yielding an excellent productivity of 20 g·L-1·h-1 with a biocatalyst yield of 4.36 g·gbiocat-1. This
work also highlights the simplicity of applying a reaction trajectory analysis methodology,
importance of scale during reaction characterizations and identifies future directions for reaction
improvement to address substrate supply and product inhibition/deactivation.
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KEYWORDS. Biocatalysis; recombinant Pig Liver Esterase (PLE); Reaction Scoping; Kinetic
Characterizations; Substrate Mass Transfer
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TOC-figure
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Introduction
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Biocatalysis has become an established branch of chemistry in its own right resulting from the
excellent stereo-, regio- and enantioselectivity of biocatalysts even under mild reaction
conditions.1,2 Consequentially, biocatalysts can realize sustainable manufacturing processes,
satisfying 10 of the 12 principles of green chemistry: waste prevention and high atom efficiency
(avoiding the formation of by-products) through their high selectivity, less hazardous syntheses,
safer solvents (most often carried out in water), and low energy demand due to their operation
under mild process conditions, their origin from renewable sugar feedstocks through fermentation,
avoidance of derivatization, their nature as catalysts and not stoichiometric reagents, the ability to
apply real-time process analytics to prevent waste, and most importantly, their inherent safety.3
Additionally, these traits frequently (although not always) infer better process economics. Finally,
biocatalysts can be further improved by protein engineering, offering an additional degree of
freedom towards process design. Such improvements are gained through tailoring the amino acid
sequences of enzymes to enhance stability, activity or selectivity.4 Indeed, numerous biocatalytic
processes have been successfully commissioned to date, using these approaches.5-7
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Conventional catalysis is most often performed in homogeneous gas-phase reactors or organic
solvents, where reaction scale-up is often dependent on adequate heat transfer and appropriate
reactor design. For biocatalysis, under mild conditions (ambient temperature, near neutral pH and
in aqueous-based media), this is less relevant. Instead, the conditions of the reaction become
important and biocatalytic reaction performance can be estimated (or “scoped”) using a standard
set of process performance measures (metrics).8 These metrics may be benchmarked against
industrial requirements, while the importance and value of each respective metric may differ
depending on the value of the product and target market. It is advantageous to be able to quantify
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these metrics at a scale as small as possible, in order to allow fast and efficient process evaluation
using the smallest possible amounts of (often expensive) materials. However, it turns out that in
many cases these metrics do not scale well. Understanding these limitations of scale-up for
biocatalysis helps address them during process development.
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A major reason for performance changes across scales is due to substrate mass transfer
limitations. It is well known that enzymes require at least some water to be present in their
environment to be active.9 Since organic substrates and products of synthetic interest are frequently
poorly water-soluble, this results in mass transfer limitations.10 This applies also to gas-liquid
systems, such as the important enzyme-catalyzed oxidations11, due to the particularly low water-
solubility of oxygen. Interestingly, the scale-dependence of gas-liquid mass transfer is different
from liquid-liquid mass transfer (see Table 1). Mixing is a function of power input, which can vary
significantly with scale.12 More specifically, it is easier to thoroughly mix a smaller volume to
achieve good dispersion (high power input per unit volume) than it is to mix a large volume. As
dispersion increases, so does the specific interfacial area (m-1) of the second phase component
(smaller gas bubbles or liquid droplets), which leads to improved mass transfer. However, when
considering gas-liquid mass transfer, an even greater influence is the increased gas hold-up at
larger scales, allowing an increase in overall volumetric mass transfer coefficient (kLa).13 In
contrast, liquid-liquid systems are not affected by hold-up because the volumetric component
remains in the reaction volume unlike gases, which pass through a reactor. Furthermore, liquid-
liquid systems have the ability to be emulsified by adding surfactants resulting in a homogeneous
medium, which being independent of mixing thereby makes them scale-independent. While solid-
liquid systems are not in focus in this work, similar parallels can be drawn, although they are less
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dependent on mixing to ensure a high interfacial area and are rather influenced more by initial
particle size.
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Table 1. Effect of scale on gas-liquid and liquid-liquid mass transfer.
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Gas-Liquid Mass Transfer
Liquid-Liquid Mass transfer
Dispersion (i.e.
Dispersion (i.e.
bubble size)
Hold-up a
droplet size /
Hold-up
coalescence) b
↑
↓
↓↓
↑↑
↑
↓
Small-scale
Large-scale
N/A
N/A
a The extent of effect of hold-up on mass transfer is more or less pronounced for plug-flow or
well-mixed regimes, respectively.12
b Stable emulsions create a homogeneous medium that will be independent of mixing.
It is therefore important to consider the scale at which reaction assessments are performed
because some mass transfer limited reactions (e.g. gas-liquid systems) will not proceed at the same
rate at a small-scale as at larger scales. The most important metrics to evaluate under these
circumstances are the biocatalyst yield and product concentration. These indicate if the catalyst is
sufficiently efficient and if enough product can be generated in a reasonable amount of time. Time-
course measurements are an effective means to assess reaction performance and gauge these
metrics. Recently, a method making use of comparing normalized time-course measurements (or
reaction trajectories) under different experimental conditions was used to distinguish mass transfer
limitations from other kinetic effects (e.g. enzyme stability, substrate affinity or intrinsic
activity).14 Additionally, the mechanism of enzyme deactivation or potential substrate/product
inhibition can also be identified using such time-course progressions.14-17 Time-course
progressions also give indication of the expected productivity of a reaction, and already at this
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stage, reactions that proceed too slowly may be earmarked for improvement. Naturally, only
metrics assessed at a larger scale can be considered true process metrics, although understanding
how to gauge a reflection of the metrics at smaller scales is invaluable to the effectiveness of
process development.
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In this work, a recently published procedure of reaction scoping for biocatalysis14 was applied
to the enantioselective desymmetrization of dimethyl cyclohex-4-ene-cis-1,2-dicarboxylate (1) to
(1S,2R)-1-(methoxycarbonyl)cyclohex-4-ene-2-carboxylic acid (2) catalyzed by recombinant pig
liver esterase (rPLE; ECS-PLE06; Scheme 1).18 The monoester product of this reaction is of value
as a pharmaceutical intermediate towards the synthesis of biologically active molecules19 (some
examples are shown in Scheme 1)20, the antibiotic plantencin21 as well as some active
pharmaceutical ingredients in treatments towards HIV22 and cytomegalovirus (Herpes)23.
Previously, this reaction and enzyme were characterized in terms of pH and temperature optima
as well as reaction performance.18,24,25 A simple method of pH control was implemented that made
use of inexpensive sodium bicarbonate degradation in the presence of the carboxylic acid product
2, thereafter forming CO2 and passing from the reaction medium. This was found to be more
effective than conventional titration of base to control the pH. In addition, the pH remains stable
within non-saturated bicarbonate solutions. Eventually this promising reaction showed adequate
performance to convert substrate concentrations of 200 mM (40 g·L-1) with an enantiomeric excess
of > 99.5%.18 However, this reaction performance came from a series of improvised experiments
and the bottleneck towards large-scale production was not determined. On this basis, we argued
that it would be valuable to perform a bottleneck identification using reaction trajectory analysis14
to determine the major process limitation. Using time-course measurements in this way we argued
it would be possible to distinguish the dominating bottleneck to the required performance of ECS-
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PLE06 under given conditions: either enzyme kinetics, enzyme stability or, more likely, substrate
mass transfer. With this limitation identified, the reaction could be further up-scaled and its
suitability for process implementation assessed.
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COOMe
COOH
ECS-PLE06
+
+ H2O
CH3OH
NaHCO3 (aq)
COOMe
COOMe
1
2
O
O
H
NH
COO
NH3
O
H
anticapsin
(+)-aucantene cis-carbapenems
Scheme 1. Desymmetrization of diester 1 to (1S,2R)-monoester 2 catalyzed by recombinant pig
liver esterase (ECS-PLE06).
Results and Discussion
Understanding the influence of the poorly water-soluble substrate on this system initially
required a series of time-course experiments with varying enzyme concentrations (expressed as
dosed activities) to identify potential mass transfer limitations. With this limitation on reaction rate
considered, process performance metrics were assessed. Further scale-up was then considered with
the goal of maximizing product concentration.
The first step in this systematic procedure was to elucidate the major reaction limitation by
varying enzyme loadings at a fixed, moderate substrate dose. Figure 1a shows representative time-
course progressions at different enzyme doses at a fixed substrate concentration of 80 mM.
Normalizing the time axis by the varied parameter (enzyme activity) reveals that the reaction
becomes substrate mass transfer limited at enzyme doses above ca. 6 U·mL-1, since the reaction
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trajectories diverge (Figure 1b). Interestingly, the product was found to have a beneficial effect by
better dispersing the substrate in the reaction medium since it acts as an emulsifier. Likewise,
methanol was also formed as a by-product, which would (at higher concentrations) help improve
substrate solubility in the medium (and thereby reduce mass transfer limitations). The slight
improvement in reaction rate due to these factors can be observed after initial product formation,
especially at the lowest enzyme doses.
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Figure 1. (a) Conversion as a function of time for the ECS-PLE06 catalyzed reaction of 80 mM
diester 1 to monoester 2 using 40 mL saturated NaHCO3 as a buffer (pH 8.15) at 40 °C in a stirred
round bottom flask (left). (b) Data replotted with the x-axis normalized by enzyme concentration
to reveal the major process limitation (right). ECS-PLE06 concentrations, [E0], were varied as
follows: ● – 1.54 U·mL-1, ■ – 2.99 U·mL-1, + – 4.50 U·mL-1, * – 5.94 U·mL-1, ♦ – 8.72 U·mL-1,
▲ – 11.37 U·mL-1.
Figure 2 is a plot of space-time yield and biocatalyst yield as a function of enzyme activity.
According to Figure 2, the upper limit of productivity due to substrate mass transfer limitations is
21 g·L-1·h-1. Clearly, the expected linear response of space-time yield to enzyme activity no longer
holds beyond this point, indicating a non-enzyme related limitation. On the other hand, biocatalyst
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yield appears to improve at lower enzyme concentrations because the same amount of substrate is
still converted, even though the reactions take longer. Another way of improving the biocatalyst
yield is to increase the substrate loading thereby converting more substrate with the same amount
of enzyme. In this way, a better reflection of biocatalyst yield is obtained because, in reality,
industrial processes should operate with the highest possible space-time yield in order to afford
cost-effective processes (shorter batches and smaller reaction volumes). However, the most
representative biocatalyst yield should still be evaluated under conditions where the reaction is not
kinetically limited. This is especially important to consider in cases such as this, which are limited
by liquid-liquid mass transfer.
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Figure 2. Space-time yield (STY; blue ●) and biocatalyst yield (red ▲) as a function of rPLE dose
(presented as activity) after the full conversion of 80 mM substrate. The dashed line indicates
linearity through the origin. Data in yellow are those obtained after the full conversion of 400 mM
substrate.
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Hence, the substrate concentration was increased 5-fold (400 mM) and an enzyme dose of
6 U·mL-1 was maintained because mass transfer limitations only occurred at higher enzyme
concentrations. Full conversion and no kinetic limitations were observed at this higher substrate
concentration, which was confirmed by the overlap of trajectories on a normalized time axis plot
(Figure 3). The resultant 4.9-fold improvement in biocatalyst yield from 0.89 to 4.36 g·gbiocat-1 at
an almost identical maximum space-time yield is shown in Figure 2.
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Figure 3. Reaction with 80 mM substrate (●) and 400 mM substrate (▲) in 40 mL saturated
NaHCO3 buffer (pH 8.15) at 40 °C both with an enzyme dose of 6 U mL-1 on a normalized time-
axis plot.
In order to investigate the potential of further increasing substrate concentration, it was necessary
to marginally increase the scale of reaction by operating in a 250 mL stirred tank reactor. At this
scale, better control and measurement of the reaction environment was made possible through
defined stirring, temperature and pH control. Substrate concentrations greater than 500 mM caused
the enzyme to agglomerate and denature. Therefore, three 400 mM batch doses of 1 were supplied
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at 3 h intervals in a fed-batch manner to mimic the previous reaction profiles observed at smaller
scale. Nevertheless, full conversion was not reached even after 24 h (data not shown). In order to
rule out potential substrate inhibition caused by too high single substrate concentration doses, a
constant feed of substrate at 5.81 g·h-1 was instead supplied to the reactor by use of a syringe pump.
This would hypothetically result in a calculated 1.19 M (225 g·L-1) product and a space-time yield
of 22.0 g·L-1·h-1 in a 9 h reaction, assuming the reaction is not limited. However, during a fed-
batch reaction, product formation ceased after 7 h, after which substrate accumulated at a rate equal
to that of the pump feed rate (Figure 4a). Since the mass balance of substrate and product closed
throughout the reaction, this result indicated that indeed the reaction had stopped after 400 mM
product formation (Figure 4b). We postulated therefore that either enzyme stability or product
inhibition could be the cause. In order to confirm this, the end-point reaction medium was dosed
with a fresh 6 U·mL-1 of ECS-PLE06 and left to incubate at 40 °C for 48 h. Conversion was only
marginally improved by approximately 10% (data not shown) indicating that product inhibition or
product induced deactivation was the most likely cause of activity loss.
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Figure 4. (a) Fed-batch conversion of 1.2 M diester 1 (●) to monoester 2 (▲) in a 250 mL (end
volume) stirred tank reactor. pH was maintained with saturated NaHCO3 buffer (pH 8.15) at 40 °C
and included an enzyme dose of 6 U·mL-1 (above). (b) Mass balance corresponding to the sum of
substrate and product in the reaction medium (●) the actual fed-batch rate of substrate supply to
the reactor by a syringe pump (below). Error bars reflect 95% confidence intervals (n = 3).
In order to test this further, the time-dependent stability of ECS-PLE06 was evaluated with
respect to temperature (40 °C), high buffer strength solutions (saturated NaHCO3), and by-product
methanol concentrations of up to 5% (v/v) (corresponding to a high molar concentration of
ca. 1.3 M; Figure 5). ECS-PLE06 showed a negligible activity loss after 24 h in the presence of all
environmental factors. Furthermore, the enzyme gained activity whilst being incubated in NaHCO3
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buffer and methanol, which might point towards an affinity of ECS-PLE06 for strongly buffered
environments containing methanol. This confirmed that the remaining limitation to be addressed
was product inhibition or product induced deactivation.
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Figure 5. Stability of ECS-PLE06 as a function of time at room temperature in 50 mM phosphate
buffer at pH 7.5 (control, solid bars), at 40 °C in 50 mM phosphate buffer (pH 7.5; diagonal striped
bars), at 40 °C in saturated NaHCO3 buffer (1.14 M, pH 8.15; opaque bars), and with 5% (v/v)
methanol (~1.3 M) at 40 °C in saturated NaHCO3 buffer (horizontal striped bars). Error bars reflect
95% confidence intervals (n = 4).
In situ product removal (ISPR) techniques could potentially be applied to address product
inhibition/deactivation. The method of reactive crystallization would be particularly interesting
because it could be implemented through the selection of an appropriate metal bicarbonate salt. In
this way, the charged product anion could potentially precipitate from solution when bound to the
metal cation without negatively affecting enzyme performance. Furthermore, the bicarbonate ion
would also help buffer the pH during reaction presenting a potent ‘self-healing’ system. This
method of reactive crystallization ISPR was recently demonstrated with a transaminase reaction
to improve the thermodynamic equilibrium to favor product formation.26 Ultimately, product in
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the form of solid precipitates resulted in a product concentration in excess of 1 M. For the case of
the rPLE reaction, two-liquid phase ISPR approaches would not be feasible because the substrate,
being hydrophobic, would partition more strongly than the product into an organic phase. Instead,
further focus could be made towards more expensive ISPR methods such as selective ion exchange
resins27,28 or charged membranes29. Another possibility could be to remove the product in an
external loop through pH swing (precipitation) or cooling crystallization, although this is less
likely to be feasible since the excess bicarbonate would need to be removed and such a system
would also require a means to retain the biocatalyst from the harsh conditions in the external
loop.30,31 The reaction itself might be also further improved by the use of specific ionic liquids as
additives.32
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Conclusions
Through the application of a standardized methodology relying on reaction trajectory analysis,
it was concluded that the reaction was limited by mass transfer. This was due to the low solubility
of substrate in the water-based medium. ECS-PLE06 is a robust enzyme that is highly thermostable
and tolerant towards solvents25, therefore mass transfer limits could still potentially be reduced by
increasing temperature or adding water-miscible co-solvents to this system to improve substrate
solubility. This specific system also showcases the elegance of ‘self-healing’ systems where pH
can easily be maintained by the reaction medium itself through an oversaturated, benign carbonate
buffer reaction with product cation species.
In this work improved performance of ECS-PLE06 was achieved with final process metrics
summarized in Table 2. These results point towards a promising biocatalytic reaction of up to
0.4 M (75 g·L-1) batches of product with an expected biocatalyst yield and productivity of
-1
4.36 g·gbiocat and 20.2 g·L-1·h-1, respectively. Beyond this relatively high concentration limit,
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product inhibition or product induced inactivation will prevent further conversion of substrate. If
higher product concentrations are required, then alternative ISPR techniques must be considered.
Furthermore, substrate must be dosed in a fed-batch manner to avoid enzyme agglomeration and
activity loss above 500 mM substrate.
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Table 2. Process performance metrics for the ECS-PLE06-catalyzed desymmetrization of 1 to 2.
Reaction conversion
Product concentration
Space-time yield
100%
74 g·L-1
20.2 g·L-1·h-1
1.20 g·gbiocat-1·h-1
Specific productivity
Biocatalyst yield
-1
4.36 g·gbiocat
Materials and Methods
REAGENTS. All reagents were of analytical grade and supplied by Sigma-Aldrich (St. Louis,
MO). Dimethyl cyclohex-4-ene-cis-1,2-dicarboxylate (1), (1S,2R)-1-(methoxycarbonyl)cyclohex-
4-ene-2-carboxylic acid (2) and lyophilized rPLE (ECS-PLE06; EC 3.1.1.1; ≥ 0.3 U·mg-1) were
kindly supplied by Enzymicals AG (Greifswald, Germany).
ACTIVITY ASSAY. The activity of pig liver esterase was quantified through
spectrophotometric measurements of the degradation of p-nitrophenyl acetate (pNPA) to p-
nitrophenolate. Reaction mixtures contained 850 μL of phosphate buffer (pH 7.5, 50 mM), 50 μL
of enzyme stock solution and 100 μL of pNPA in DMSO (10 mM). Initial rate measurements were
taken at 400 nm (ε30 °C = 17759 L·mol-1·cm-1) over 100 s at 30 °C. One unit (U) of esterase activity
was defined as the generation of 1 μmol of p-nitrophenolate per minute under the standard activity
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assay conditions. Measurements were made using a UV-vis spectrophotometer (Specord 50,
Analytik Jena AG, Jena, Germany). Reference samples contained distilled H2O and negative
controls were performed in the absence of enzyme.
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SMALL-SCALE APPARATUS. Small-scale reactions were performed in a 100 mL round
bottom flask using a magnetic stirrer for mixing and an external water bath for temperature control
at 40 °C. 40 mL of saturated NaHCO3 buffer (pH 8.15) was used to control the pH. ECS-PLE06
(≥ 0.3 U·mg-1) was dosed to this medium, and the reaction was started by the addition of 0.6 g of
1 (80 mM). Samples (500 μL) were taken periodically and concentrations of 1 and 2 were
determined using the procedure detailed in the analytical methods section. Reaction progress could
be indirectly followed by use of a CO2 bubble counter fixed to the top of the apparatus.
FED-BATCH 250 mL STIRRED TANK REACTOR. Fed-batch experiments were performed
in a 300 mL glass jacketed stirred tank reactor (STR). 42 g of NaHCO3 (corresponding to 2 M
final concentration, which translates to a saturated solution with excess solid NaHCO3) was added
with 1500 U of ECS-PLE06 (6 U·mL-1 final concentration) to 190 mL distilled H2O. Temperature
was maintained at 40 °C by use of an external CC-K6 cryostat (Huber, Offenburg, Germany) and
mixing ensured by a pitched 4-blade turbine with overhead stirring motor at 300 rpm. pH was
measured by an InLab Semi-Micro electrolyte sensor (Mettler Toledo, Colombus, OH) and was
found to be 8.15 ± 0.38 (mean ± SD) over all experiments using the sodium hydrogen carbonate
buffered system. 60 mL of substrate 1 (corresponding to a final concentration of 1.2 M) was fed
by a R99-E syringe pump (Razel Scientific Instruments, Fairfax, VT) over the first 10 hours to
make up a final reaction volume of 250 mL. Triplicate samples (500 μL) were taken periodically
over 24 h and concentrations of substrate and product were determined using the procedure
detailed in the analytical methods section.
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ANALYTICAL METHODS. Concentrations of 1 and 2 were quantified by GC-FID (TRACE
1310 Series, Thermo Fisher Scientific, Waltham, MA). Triplicate samples of 500 μL were taken
from reaction media and acidified with 500 μL of concentrated HCl (1:1 volumes) to render 2
insoluble in the aqueous environment and to denature residual protein activity. Samples were then
diluted and extracted with methyl tert-butyl ether (MTBE) containing n-decane (25 mM) as an
internal standard by vortexing on the highest setting and phase separation followed by
centrifugation. 1 μL of the organic phase fraction was injected into a TR-1701 column (30 m x
0.25 mm x 0.25 μm; Thermo Fisher Scientific, Waltham, MA) where the oven temperature was
ramped from 130 °C to 200 °C at a rate of 5 °C min-1 and thereafter held at an isotherm of 200 °C
for 1 min. Injector and detector temperatures were maintained at 250 °C throughout. The carrier
gas flow rate was 1.2 mL min-1 with a split ratio of 35:1. Experimental substrate and product
concentrations were determined using a standard curve generated by subjecting samples of known
concentrations to the same acidification/solvent extraction procedure.
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AUTHOR INFORMATION
Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval
to the final version of the manuscript.
Funding Sources
Funding by the German Federal Ministry of Education and Research (BMBF – Bundesministerium
für Bildung und Forschung; project number: 031A123) and the German Research Foundation
(DFG – Deutsche Forschungsgemeinschaft; LA 4183/1-1) is gratefully acknowledged. The
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authors thank the Technical University of Denmark (DTU) for financial support for one of us
(MPM).
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Notes
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The authors declare no competing financial interest.
ACKNOWLEDGMENT
The authors thank Prof. Dr. Udo Kragl (University of Rostock, Germany) and Prof. Dr. Uwe
Bornscheuer (University of Greifswald, Germany) for their ongoing support and fruitful
discussions.
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TOC-figure
61x45mm (300 x 300 DPI)
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