Light-Driven Enzymatic Decarboxylation of Fatty Acids

Article abstract of DOI:10.1002/anie.201807119

The photoenzymatic decarboxylation of fatty acids to alkanes is proposed as an alternative approach for the synthesis of biodiesel. By using a recently discovered photodecarboxylase from Chlorella variabilis NC64A (CvFAP) we demonstrate the irreversible preparation of alkanes from fatty acids and triglycerides. Several fatty acids and their triglycerides are converted by CvFAP in near-quantitative yield and exclusive selectivity upon illumination with blue light. Very promising turnover numbers of up to 8000 were achieved in this proof-of-concept study.

Full text of DOI:10.1002/anie.201807119

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Accepted Article
Title: Light-driven enzymatic decarboxylation of fatty acids
Authors: Mieke Huijbers, Wuyuan Zhang, and Frank Hollmann
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Angewandte Chemie International Edition
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COMMUNICATION
Light-driven enzymatic decarboxylation of fatty acids
+
+
Mieke M.E. Huijbers , Wuyuan Zhang , Fabio Tonin, and Frank Hollmann*
Abstract: Photoenzymatic decarboxylation of fatty acids into
alkanes is proposed as alternative approach for biodiesel synthesis.
Using a recently discovered photodecarboxylase from Chlorella
variabilis NC64A (CvFAP) we demonstrate the irreversible
preparation of alkanes from fatty acids and triglycerides. Several
fatty acids and their triglycerides are converted by CvFAP upon
illumination with blue light in near-quantitative yield and exclusive
selectivity. In this proof-of-concept study very promising turnover
numbers of up to 8000 were achieved.
2
photochemical decarboxylation using Pd-doped TiO operating
under much milder reaction conditions.[3] However, one common
issue of all classical chemical processes for fatty acid
decarboxylation is their rather poor selectivity yielding complex
product mixtures due to Kolbe- and Hofer-Moest-type side
reactions.
Enzymatic counterparts so far are limited to the oxidative
[
4]
decarboxylation of fatty acids to terminal alkenes.
or to
activated carboxylic acids such as malonic acids[ or aromatic
5]
carboxylates.[6]
The conversion of (waste) fatty acids and oils into biofuels has
been in focus of research for decades.[1] The most widely used
approach is to transform fatty acid (esters) into the
corresponding methyl- and ethyl esters (FAMEs and FAEEs,
respectively). Especially hydrolase-catalysed transesterification
of oils and fats has been in focus due to its mild reaction
conditions. Equilibrium issues, however, still challenge the
practicability of this approach. Decarboxylation of fatty acids into
the corresponding C1-shortened alkanes may be an interesting
alternative to the (trans)esterification strategy (see Scheme 1).
Very recently Beisson and coworkers reported on an algal
fatty acid photodecarboxylase from Chlorella variabilis NC64A
7]
(
CvFAP).[ This flavoenzyme catalyses the light-dependent
decarboxylation of some long-chain fatty acids to the
corresponding C1-shortened alkanes. CvFAP requires
photoactivation by blue light (450 nm) indicating that only the
photoexcited FAD in the enzyme’s active site is capable of the
reaction. Examples of photoenzymes are quite rare: next to
CvFAP, only flavin-dependent DNA-repair enzymes[8] and
[9]
protochlorophyllide oxidoreductases, are known today.
We therefore set out to explore the preparative potential of
CvFAP for the production of alkanes from biobased fatty acids
and triglycerides.
Two variants of CvFAP were recombinantly produced in
Escherichia coli: one variant comprises the complete sequence
of CvFAP (residues 1-654, full-length, Figure S1), while the
second variant lacks a predicted chloroplast targeting sequence
and comprises residues 62-654 of CvFAP (short-length, Figure
S1). After production in Escherichia coli, the crude extract as
well as the purified enzyme were used for catalytic experiments.
While short-length CvFAP showed a good overproduction in E.
coli and was purified in one step to a purity of about 40%, full-
length CvFAP showed almost no overproduction in E. coli and
there was no clear band on SDS-PAGE after purification (Figure
S2). Though both enzyme preparations showed significant
decarboxylation activity (Figures S5 and S6) we continued our
investigations using the short-length CvFAP. Interestingly, crude
enzyme preparations exhibited a higher activity and robustness
of CvFAP as compared to purified preparations. The activity of
the purified enzyme was increased by a factor of 2 when
reconstituted with filtered E. coli cell extract. Pre-incubating the
enzyme in blue light causes a loss of activity, which is less
pronounced when pre-incubating the enzyme in filtered E. coli
cell extract (Table S1). At present time we no satisfying
explanation for stabilization effect and further experiments will
be necessary to shed more light on this. For all further
experiments crude cell free extracts were used. It is worth
mentioning that no background activity was found in the crude
extracts prepared from E. coli cells containing an empty vector.
Furthermore, performing experiments either in the absence of
blue light or in the absence of CvFAP (or using thermally
inactivated cell extract) gave no determinable conversion of e.g.
Scheme 1. Enzymatic transesterification (upper) and decarboxylation (lower)
reactions for synthesis of biofuels.
On the one hand, the specific heat of combustion of alkanes is
somewhat (ca. 9%) higher than that of corresponding FAMEs.[1]
On the other hand, the irreversible decarboxylation of fatty acids
should lead to simpler reaction schemes as compared to the
reversible (trans)esterification procedure where issues such as
water content (leading to saponification and catalyst inactivation)
and equilibrium (generally significant molar surpluses of
methanol or ethanol are required to achieve near-full
conversion) arise.
Established chemical methods for the decarboxylation of
fatty acids require rather harsh reaction conditions and rare-
metal catalysts, which will challenge the overall eco-efficiency of
the proposed alkane synthesis.[2] Recent advancements in
[
*]
Dr. M.M.E. Huijbers,[+] Dr. W. Zhang,[+] Dr. F. Tonin, Dr. F. Hollmann
Department of Biotechnology, Delft University of Technology
Van der Maasweg 9, 2629 HZ Delft, the Netherlands
E-mail: f.hollmann@tudelft.nl
[+
]
These authors contributed equally to this work.
Supporting information for this article is given via a link at the end of
the document.
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Angewandte Chemie International Edition
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COMMUNICATION
palmitic acid. Performing the photoenzymatic reaction under
differences of the substrate binding in CvFAP’s access channel
different atmospheric conditions (air, Ar,
N
2
or
H
2
)
gave
and positioning of the reactive group towards the cofactor.
essentially the same result (i.e. full conversion of palmitic acid
within 3h, Table S2). To increase the solubility of the
hydrophobic fatty acids such as palmitic acid, DMSO was
applied as a cosolvent. CvFAP tolerates up to 50% (v/v) DMSO
Table 1. Substrate scope of the photoenzymatic decarboxylation reaction.
(
Figure S7). In further studies, the reaction mixture contained
30% (v/v) of DMSO.
Under these conditions, we were pleased to observe a
Substrate
[Product]
mM]
Conversion
[%]
TON (CvFAP)[b]
500
smooth conversion of palmitic acid into pentadecane in the
presence of CvFAP and blue light (Figure 1). The reaction was
strictly light-dependent.
[a]
[
C
12
H
24
O
2
3.0
11
(Lauric acid)
C
14
H
28
O
2
6.9
25
1150
(
Myristic acid)
C
16
H
32
O
2
27.7
28.7
26.1
17.7
14.6[c]
25.7
96
4610
(Palmitic acid)
C
17
H
34
O
2
96
4780
(
Margaric acid)
C
18
H
36
O
2
92
4350
(Stearic acid)
C
18
H
34
O
2
(9)
65[c]
49[c]
90
2950
(Oleic acid)
C
18
H
32
O
2
(9, 12)
2600
(Linoleic acid)
C
20
H
40
O
2
4580
(Arachidic acid)
Figure 1. Photoenzymatic decarboxylation of palmitic acid into pentadecane.
Conditions: [CvFAP] = 6.0 M, [palmitic acid] = 13 mM, Tris-HCl (pH 8.5, 100
mM), pH 8.5, 30 % DMSO, blue light illumination.
Reaction conditions: [substrate] = 30 mM, [CvFAP] = 6.0 μM, Tris-HCl buffer
-
1
(pH 8.5, 100 mM) , 30 % DMSO, blue light illumination (intensity = 13.7 µE L
s ) for 14 hours. [a] Conversion: [product]final([product]final +[substrate]final)-1; [b]
-
1
-
1
TON: [product]final  [CvFAD] . [c ] Due to the lack of standard reference, the
conversion was calculated by using: conversion ([substrate]initial
=
-
-
1
[
substrate]final)  [substrate]initial .
Further characterisation experiments showed that the rate of
pentadecane production depended on the enzyme concentration,
the intensity of the light source and, to some minor extent, the
reaction temperature (Figures S5, S8 and S9).
A preparative-scale synthesis was performed under the
optimal conditions (see Supporting Information). 155 mg of
pentadecane was obtained, giving 79% of conversion (TON of
CvFAP 7916) and 61% of isolated yield (Figures S11 and S12).
Next, we explored the scope of the photoenzymatic
decarboxylation reaction somewhat further (Table 1). Generally
speaking, a broad range of different fatty acids was converted.
For some of them full conversion was observed also resulting in
quite favourable turnover numbers (TON, i.e. concentration of
product divided by concentration of the biocatalyst) for the
enzyme.
In view of the envisaged production of alkanes from (waste)
oils and fats we also investigated a bi-enzymatic cascade
comprising lipase-catalysed hydrolysis of triolein to the free oleic
acid and glycerol followed by the CvFAP-catalysed
photodecarboxylation reaction (Table 2).
The two-step cascade using homogeneously dissolved
triolein (20 mM) gave a satisfying overall yield of more than 80%
(Table 2, entry 5) and respectable turnover number for the
photodecarboxylase of 8280, which encouraged us to proceed
to a cosolvent-free two liquid phase system. A first experiment
using 890 U of lipase gave a relatively low product yield of 16
mM, which was attributed to a drop of pH in the aqueous phase
to 5.2 leading to a decreased CvFAP activity (Table 2, entry 1).
Indeed, either lowering the lipase activity or adjusting the pH
value after the hydrolysis reaction lead to significantly increased
product formation (Table 2, entries 2&3). Performing both steps
simultaneously (one-step) (Table 2, entry 4) was shown to be
practical though the product yield and consequently the TON of
In accordance with previous observations[7a] CvFAP showed
the highest activity with long-chain fatty acids (C>14).
Interestingly, the conversions of oleic acid and linoleic acid were
significantly lower than of the fully saturated counterpart (stearic
acid). We therefore investigated the relative activity of CvFAP
towards some isomers of oleic acid (cis/trans- and regioisomers,
Table S3) and docked these isomers into the crystal structure of
CvFAP (PDB: 5NCC, Figures S14-17).[7a] A good correlation
between the distance of the substrate carboxylate group to the
flavin cofactor and the initial rate of the decarboxylation rate was
found (Table S3). Hence, the differences in conversion and rate
observed for the different substrates may be assigned to
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COMMUNICATION
CvFAP were rather modest. Again, a pH drop in the aqueous
phase was observed. Optimisation of the relative activities of
lipase and photodecarboxylase will circumvent this limitation and
reveal the full potential of the proposed photobiocatalytic
synthesis of alkanes from triglycerides.
For production of the CvFAP in E. coli, two constructs were designed
based on a previously reported construct.[7a] The constructs both consist
of sequentially a 6x His-tag, thioredoxin (TrxA) tag, tobacco etch virus
(TEV) protease cleavage site and the gene coding for CvFAP (GenBank:
KY511411.1). The first construct comprises the full-length sequence
(residues 1-654) of CvFAP, while the second construct lacks the residues
Table 2. Photobiocatalytic cascade for the transformation of triolein into (Z)-
encoding for a predicted chloroplast targeting sequence and thereby
comprises residues 62-654 of CvFAP (Fig. S1). The sequence coding for
CvFAP was codon optimised for expression in E. coli. The construct was
synthesised by Baseclear (Leiden, the Netherlands) and cloned into a
pET28a vector using NdeI and HindIII restriction sites. Competent E. coli
BL21 (DE3) cells (NEB) were transformed with the plasmid for
recombinant enzyme production.
heptadec-8-ene.
Preparation of the cell free extract containing CvFAP
(full-length/short-length)
10 mL pre-cultures were inoculated with E. coli BL21 (DE3) cells
harboring the designed pET28a-His-TrxA-CvFAP plasmid. These
cultures were grown overnight in terrific broth (TB) medium, containing
50 μg/mL kanamycin. The pre-cultures were used to inoculated large
cultures (500 mL TB + 50 μg/mL kanamycin in 2 L shake flasks). Cells
were grown at 37 °C, 180 rpm, until an OD600 between 0.7-0.8 was
reached. Protein production was induced by the addition of 0.5 mM IPTG
and the cells were left at 17 °C, 180 rpm, for about 20 hours. Cells were
harvested by centrifugation (11000 g at 4 °C for 10 min), washed with
Tris-HCl buffer (50 mM, pH 8, containing 100 mM NaCl) and centrifuged
again. The cell pellet was resuspended in the same buffer, and 1 mM
PMSF was added. Cells were lysed by passing them passed twice
through a Multi Shot Cell Disruption System (Constant Systems Ltd,
Daventry, UK) at 1.5 kbar, followed by centrifugation of the cell lysate
Entry
conditions
[CrLip]
[Product]
[mM]
TON (CvFAP)
-
1
[U ml ]
1
2
3
two-step, pH 7.5
two-step, pH 7.5
890
16.4
27.7
34.5
1366
2308
2875
89
(38000 g at 4 °C for 1 h). After centrifugation, 5% glycerol (w/v) was
two-step,
adjustment of pH
after step 1
890
added to the soluble fraction, the cell extract was aliquoted, frozen in
liquid nitrogen and stored at -80 °C.
The total protein content of the cell extract was determined by a BCA
Assay (Interchim), using BSA as a standard. CvFAP production was
4
5
one-step, pH 7.5
890
19.8
1650
8280
analysed by SDS-PAGE (Figure S2) using
a Criterion™ Cell
homogeneous,
two-step, pH7.5
2500
49.7 (83%
conversion)
electrophoresis system (Bio-Rad). As molecular weight marker,
a
Precision Plus Protein Standard (Bio-Rad) was used. The gel was
analysed using a gel imaging system (GBox, Syngene, Cambridge, UK)
and the amount of CvFAP in the cell extract was estimated from the
relative intensity of the bands on gel.
General conditions two-step procedure: triolein : Tris-HCl buffer (pH 8.5,100
mM) phase ratio = 1:1 (v/v); CrLip: Lipase from Candida rugosa; T= 37 oC;
step 1: reaction time = 12h; followed by step 2: [CvFAP] = 6 M; irradiation
-
1
-1
with blue light (450 nm; intensity = 13.7 µE L s ); reaction time = 20h. TON:
-
1
[
product]final  [CvFAP] .
As a control, a cell free extract of E. coli BL21 (DE3) cells harboring an
empty pET28a vector was prepared according to the same protocol.
Overall, we have demonstrated the synthetic potential of the
novel photodecarboxylase from Chlorella variabilis NC64A. We
are convinced that further optimisations of the reaction setup
and of the biocatalyst(s) will yield a practical approach to
valorise non-edible triglycerides and acids into biofuels.
Photocatalytic setup
The photoenzymatic decarboxylation reactions using CvFAP were
performed at 37 °C in total volume of 1.0 mL Tris-HCl buffer (pH 8.5, 100
mM) containing 30% DMSO as cosolvent. Unless mentioned otherwise,
200 μL of DMSO containing 65.5 mM of palmitic acid, 100 μL of pure
Experimental Section
DMSO, 500 μL of Tris-HCl buffer (pH 8.5, 100 mM) and 200 μL of CvFAP
stock solution (30 μM cell extract in Tris-HCl buffer) were added to a
transparent glass vial (total volume 5.0 mL).The vial was sealed and
exposed to blue LED light under gentle magnetic stirring. The homemade
setup is shown in Fig. S3. The final conditions of this reaction were:
Cloning of CvFAP
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COMMUNICATION
[
palmitic acid] = 13.1 mM and [CvFAP] = 6 μM in Tris-HCl pH (8.5, 100
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substrates and products were extracted with ethyl acetate (containing 5
mM of 1-octanol as internal reference) in a 2:1 ratio (v/v). The remaining
organic phase was analysed using Gas Chromatography.
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mL). The hydrolysis of triolein was then performed at 37 °C a thermal
shaker (700 rpm). After 12 hours, 200 μL of CvFAP stock solution (30 μM
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analysed using Gas Chromatography.
[
5]
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described in two-step approach, except that after the addition of all
reaction components into the reaction vial, the mixture was illuminated by
blue LED light under gentle magnetic stirring at 37 °C.
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We thank the Netherlands Organisation for Scientific Research
for financial support through a VICI grant (no. 724.014.003). We
thank Dr. Jonathan Z. Bloh for the characterization of our
homemade light setup.
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Keywords: biocatalysis • biofuels • decarboxylation • fatty acids
•
photocatalysis •
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COMMUNICATION
Entry for the Table of Contents
COMMUNICATION
In the spotlight - a novel photoenzyme
enables quantitative and selective
decarboxylation of fatty acids to
alkanes.
Mieke M.E. Huijbers, Wuyuan Zhang,
Fabio Tonin and Frank Hollmann*
Page No. – Page No.
Light-driven enzymatic
decarboxylation of fatty acids
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