Chemo-enzymatic synthesis of 11-hydroxyundecanoic acid and 1,11-undecanedioic acid from ricinoleic acid

Article abstract of DOI:10.1039/c5gc01017a

A practical chemoenzymatic synthetic method for 11-hydroxyundecanoic acid and 1,11-undecanedioic acid from ricinoleic acid (12-hydroxyoleic acid) was investigated. Biotransformation of ricinoleic acid into the ester (3) via 12-ketooleic acid (2) was driven by recombinant Escherichia coli cells expressing an alcohol dehydrogenase from Micrococcus luteus and the Baeyer-Villiger monooxygenase from Pseudomonas putida KT2440. The carbon-carbon double bond of the ester (3) was chemically reduced, and the ester bond was hydrolyzed to afford n-heptanoic acid (5) and 11-hydroxyundecanoic acid (7), which were converted into other related derivatives. For example, 11-hydroxyundecanoic acid was transformed into 1,11-undecanedioic acid (8) under fairly mild reaction conditions. Whole-cell biotransformation at high cell density (i.e., 20 g dry cells per L) allowed the final ester product concentration and volumetric productivity to reach 53 mM and 6.6 mM h^-1, respectively. The overall molar yield of 1,11-undecanedioic acid from ricinoleic acid was 55% based on the biotransformation and chemical transformation conversion yields of 84% and 65%, respectively.

Full text of DOI:10.1039/c5gc01017a

View Article Online
View Journal
Green
Chemistry
Accepted Manuscript
This article can be cited before page numbers have been issued, to do this please use: H. Jang, K.
Singha, H. Kim, Y. Kwon and J. B. Park, Green Chem., 2015, DOI: 10.1039/C5GC01017A.
This is an Accepted Manuscript, which has been through the
Royal Society of Chemistry peer review process and has been
accepted for publication.
Accepted Manuscripts are published online shortly after
acceptance, before technical editing, formatting and proof reading.
Using this free service, authors can make their results available
to the community, in citable form, before we publish the edited
article. We will replace this Accepted Manuscript with the edited
and formatted Advance Article as soon as it is available.
You can find more information about Accepted Manuscripts in the
Information for Authors.
Please note that technical editing may introduce minor changes
to the text and/or graphics, which may alter content. The journal’s
standard Terms & Conditions and the Ethical guidelines still
apply. In no event shall the Royal Society of Chemistry be held
responsible for any errors or omissions in this Accepted Manuscript
or any consequences arising from the use of any information it
contains.
www.rsc.org/greenchem

Page 1 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
1
2
Chemo-Enzymatic Synthesis of 11-Hydroxyundecanoic Acid and
1,11-Undecanedioic Acid from Ricinoleic Acid
3
4
5
HyunꢀYoung Jang1,†, Kaushik Singha2,†, HwanꢀHee Kim1, YongꢀUk Kwon2,*, and JinꢀByung
Park1,*
6
7
8
1Department of Food Science and Engineering, Ewha Womans University, Seoul 120ꢀ750,
9
Republic of Korea
10
11
2Department of Chemistry and Nano Science, Ewha Womans University, Seoul 120ꢀ750,
Republic of Korea
12
13
14
15
*To whom correspondence should be addressed. Eꢀmail: yukwon@ewha.ac.kr (Y.ꢀU. Kwon),
jbpark06@ewha.ac.kr (J.ꢀB. Park), Tel: +82ꢀ2ꢀ3277ꢀ6685/4509, Fax: +82ꢀ2ꢀ3277ꢀ4213
16
17
18
†H.ꢀY. Jang and K. Singha equally contributed to this work.
1

Green Chemistry
Page 2 of 25
View Article Online
DOI: 10.1039/C5GC01017A
19 Abstract
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
A practical chemoenzymatic synthetic method for 11ꢀhydroxyundecanoic acid and 1,11ꢀ
undecanedioic acid from ricinoleic acid (12ꢀhydroxyoleic acid) was investigated.
Biotransformation of ricinoleic acid into the ester (3) via 12ꢀketooleic acid (2) was driven by
recombinant Escherichia coli cells expressing an alcohol dehydrogenase from Micrococcus
luteus and the Baeyer–Villiger monooxygenase from Pseudomonas putida KT2440. The
carbonꢀcarbon double bond of the ester (3) was chemically reduced, and the ester bond was
hydrolyzed to afford nꢀheptanoic acid (5) and 11ꢀhydroxyundecanoic acid (7), which were
converted into other related derivatives. For example, 11ꢀhydroxyundecanoic acid was
transformed into 1,11ꢀundecanedioic acid (8) under fairly mild reaction conditions. Wholeꢀ
cell biotransformation at high cell density (i.e., 20 g dry cells/L) allowed the final ester
product concentration and volumetric productivity to reach 53 mM and 6.6 mM/h,
respectively. Overall molar yield of 1,11ꢀundecanedioic acid from ricinoleic acid was 55%
based on the biotransformation and chemical transformation conversion yields of 84% and
65%, respectively.
35
2

Page 3 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
36 Introduction
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
Medium chain (C7 to C13) α,ωꢀdicarboxylic acids and ωꢀaminocarboxylic acids are widely
used as building blocks and/or intermediates for producing plastics (e.g., polyamides and
polyesters), pharmaceuticals, plasticizers, lubricants, and hydraulic fluids 1ꢀ5. For example,
11ꢀaminoundecanoic acid is used to synthesize polyamide 11, which has outstanding
chemical, thermal, and impact resistance over a wide range of flexibilities compared to that of
other high performing and engineered plastics 2.
However, the medium chain α,ωꢀdicarboxylic acids and ωꢀaminocarboxylic acids are
usually produced chemically under harsh conditions requiring high temperature and pressure,
strong acids (e.g., H2SO4 and HNO3), and/or toxic oxidants (e.g., ozone) 1, 3. 11ꢀ
Aminoundecanoic acid is produced from ricinoleic acid methyl ester via cracking at high
temperature of 450 to 500oC and adding hydrogen bromide followed by nucleophilic
substitution with ammonia 2, 6
.
A variety of emerging ideas to tackle the problems in production of such chemicals have
been reported. Milder methods based on e.g., ruthenium catalysis in combination with
peracetic acid to replace ozonolysis, or the metathesis reaction of unsaturated fatty acids to
yield linear diacids have been investigated 7, 8. 11ꢀAminoundecanoic acid has been
synthesized from vernolic (cisꢀ12,13ꢀepoxyꢀcisꢀ9ꢀoctadecenoic) acid via a reaction sequence
that includes the formation of 12ꢀoxododecanoic acid oxime 2. Biotransformation routes using
Candida tropicalis have been also reported 1. Sebacic acid and azelaic acid were produced
from hydrocarbons via fermentation of C. tropicalis 5. However, these processes can be
constrained by the limitations in the availability of the substrates used and/or low product
yields.
3

Green Chemistry
Page 4 of 25
View Article Online
DOI: 10.1039/C5GC01017A
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
74
75
76
77
78
79
80
81
Microbial synthesis of medium chain (e.g., C8–C14) fatty acids, ωꢀhydroxycarboxylic
acids and α,ωꢀdicarboxylic acids from sugars and glycerol was also investigated 9ꢀ12. For
instance, the expression of the Pseudomonas putida alkane monooxygenase system, encoded
by alkBGT, in combination with all βꢀoxidation reversal enzymes in Escherichia coli allowed
to produce 6ꢀhydroxyhexanoic acid, 8ꢀhydroxyoctanoic acid, and 10ꢀhydroxydecanoic acid
from glycerol 9. The medium chain ωꢀhydroxycarboxylic acids were further oxidized into the
corresponding α,ωꢀdicarboxylic acids by coexpression of potential alcohol and aldehyde
dehydrogenases, chnD and chnE from Acinetobacter sp. strain SE19. However, the medium
chain carboxylic acids were produced to a mixture at a rather low concentration in the
medium.
Biotransformation of renewable fatty acids and/or plant oils into C9–C13 ωꢀ
hydroxycarboxylic acids, ωꢀaminocarboxylic acids, and α,ωꢀdicarboxylic acids has been
recently reported 13ꢀ16. For example, 11ꢀaminoundecanoic acid and 1,11ꢀundecanedioic acid
have been produced from ricinoleic acid; the C12ꢀhydroxyl group of ricinoleic acid was
oxidized to the ester (3) via 12ꢀketooleic acid (2) in a serial reaction with the alcohol
dehydrogenase (ADH) of Micrococcus luteus and the Baeyer–Villiger monooxygenase
(BVMO) of P. putida KT2440 (Scheme 1) 13, 14. The ester (3) was hydrolyzed by an esterase
to yield nꢀheptanoic acid and (Z)ꢀ11ꢀhydroxyundecꢀ9ꢀenoic acid. (Z)ꢀ11ꢀHydroxyundecꢀ9ꢀ
enoic acid was then further converted to 1,11ꢀundecanedioic acid by an ADH of P. putida
GPo1 and a reductase of E. coli. (Z)ꢀ11ꢀHydroxyundecꢀ9ꢀenoic acid was transformed into 11ꢀ
aminoundecanoic acid via oxidation followed by amination with the ADH of P. putida GPo1
and ωꢀtransaminase of Silicibacter pomeroyi. However, the final product concentration and
productivity of the biosynthetic routes remains very low because of high toxicity of the
4

Page 5 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
82
83
84
85
86
87
88
89
90
91
92
93
94
95
reaction intermediates and final products (e.g., nꢀheptanoic acid, (Z)ꢀ11ꢀhydroxyundecꢀ9ꢀ
enoic acid) to the microbial host cells (i.e., E. coli) 17 and low stability of the BVMOs 18ꢀ22
.
One of the approaches to bypass such product toxicity problem could be to use a chemoꢀ
enzymatic process 23, 24. Here, a chemoꢀenzymatic catalytic process to synthesize 11ꢀ
hydroxyundecanoic acid (7) and 1,11ꢀundecanedioic acid (8) from ricinoleic acid was
investigated. Ricinoleic acid was transformed into ester (3) by recombinant E. coli cells
expressing the ADH from M. luteus and the BVMO from P. putida KT2440 (Scheme 1).
Then, the carbonꢀcarbon double bond of ester (3) was chemically reduced and the ester bond
was hydrolyzed to nꢀheptanoic acid (5) and 11ꢀhydroxyundecanoic acid (7) (Scheme 2),
which can be converted into other related derivatives, including 11ꢀaminoundecanoic acid 14.
11ꢀHydroxyundecanoic acid was also oxidized to 1,11ꢀundecanedioic acid (8) under fairly
mild reaction conditions.
96 Materials and Methods
97
98
99
Microbial strains and culture media
Recombinant E. coli BL21(DE3) pACYCꢀADH and pJOEꢀBVMO were grown in Luriaꢀ
Bertani medium (5 g/L yeast extract, 10 g/L tryptone, and 10 g/L NaCl) supplemented with
chloramphenicol and ampicillin for seed cultivation as described in our previous study 13, 14
100
101
102
103
104
.
Riesenberg medium 25 supplemented with 10 g/L glucose and the appropriate antibiotics was
used for the main cultivation and biotransformation. The Riesenberg medium consisted of 4
g/L (NH4)2HPO4, 13.5 g/L KH2PO4, 1.7 g/L citric acid, 1.4 g/L MgSO4, and 10 mL/L trace
metal solution (10 g/L FeSO4, 2.25 g/L ZnSO4, 1.0 g/L CuSO4, 0.5 g/L MnSO4, 0.23 g/L
5

Green Chemistry
Page 6 of 25
View Article Online
DOI: 10.1039/C5GC01017A
105
106
107
Na2B4O7, 2.0 g/L CaCl2, and 0.1 g/L (NH4)6Mo7O24). Recombinant gene expression was
induced by adding 0.1 mM isopropylꢀβꢀDꢀthiogalactopyranoside (IPTG) and/or 2 g/L
rhamnose to the culture broth.
108
109
Biotransformation in a flask and bioreactor
110
111
112
113
114
115
116
117
118
Biotransformation was carried out as reported previously 17. In brief, biotransformation was
initiated at the stationary growth phase, usually 8 h after inducing gene expression with 0.1
mM IPTG and/or 2 g/L rhamnose. After changing the culture broth pH to 8.0 and increasing
temperature to 35°C, 15–60 mM ricinoleic acid and 0.5 g/L Tween80 were added to the
culture broth containing 3–20 g dry cells/L. Culture and biotransformation were performed in
a 250 mL flask (working volume: 20 mL) in a shaking incubator (200 rpm). The bioreactor
experiment was conducted in a 1 L scale reactor (Biotron, Bucheon, Korea). Agitation speed
and aeration rate were 400–1000 rpm and 1 vvm, respectively, to avoid oxygen limitation
during culture and biotransformation.
119
120
Product analysis by gas chromatography/mass spectrometry (GC/MS)
121
122
123
124
125
The concentrations of the remaining fatty acids and accumulating carboxylic acids in the
medium (e.g., ricinoleic acid (1), 12ꢀketooleic acid (2), and ester (3)) were determined as
described previously16, 17. The reaction medium was mixed with an equal volume of ethyl
acetate containing 0.1 or 0.5 g/L methyl palmitate as an internal standard. The organic phase
was harvested after vigorous vortexing and subjected to derivatization with NꢀmethylꢀNꢀ
6

Page 7 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
126
(trimethylsilyl) trifluoroacetamide. The trimethylsilyl derivatives were analyzed using a
Thermo Ultra Trace GC system connected to an ion trap mass detector (Thermo ITQ1100
GCꢀion Trap MS, Thermo Scientific, Indianapolis, IN, USA). The derivatives were separated
on a nonꢀpolar capillary column (30 m length, 0.25 ꢀm film thickness, HPꢀ5MS, Agilent
Technologies, Palo Alto, CA, USA). A linear temperature gradient was programmed as 90°C,
5°C/min to 280°C. Injection port temperature was 230°C. Mass spectra were obtained by
electron impact ionization at 70 eV. Scan spectra were obtained within the range of 100–600
m/z. Selected ion monitoring was used for detection and the fragmentation analysis of the
reaction products.
127
128
129
130
131
132
133
134
135
136
Isolation of ester intermediate (3) after biotransformation
137
138
139
140
141
The ester intermediate was isolated by simple extraction with ethyl acetate several times. The
combined organic layer was washed with saline, dried over sodium sulfate, filtered, and
evaporated in vacuo. The crude ester was purified by column chromatography (silica gel)
eluting with 20% ethyl acetateꢀhexane. The isolation/purification yield of ester (3) was >
80%, based on the observed concentration of the ester after biotransformation.
142
143
144
145
146
(Z)ꢀ11ꢀ(Heptanoyloxy)undecꢀ9ꢀenoic acid (3): 1H NMR (300 MHz, CDCl3) δ 5.67ꢀ5.46 (m,
2H), 4.60 (d, J = 6.0 Hz, 2H), 2.36ꢀ2.27 (m, 4H), 2.12ꢀ2.02 (m, 2H), 1.65ꢀ1.58 (m, 4H), 1.35ꢀ
1.21 (m, 14H), 0.86 (m, 3H): 13C NMR (75 MHz, CDCl3): δ 178.2, 172.0, 133.4, 121.6, 58.3,
32.5, 32.2, 29.5, 27.4, 27.2, 27.1, 27.0, 26.9, 25.6, 23.0, 22.7, 20.6, 12.1 (see the supporting
information, Fig. S1).
147
7

Green Chemistry
Page 8 of 25
View Article Online
DOI: 10.1039/C5GC01017A
148
149
Step-by-step chemical conversion of ester intermediate (3) to undecanedioic acid (8)
11ꢀ(Heptanoyloxy) undecanoic acid (6)
150
151
152
153
154
155
156
157
158
Ester (3) (1.0 g, 3.2 mmol) was taken in methanol (10 mL) and moist RaneyꢀNi (0.4 g) was
added after to this solution soaking it in filter paper. The heterogeneous mixture was
subjected to hydrogenation under hydrogen (1 atm) for 8 h. The reaction mixture was filtered
through Celite and washed several times with methanol. The filtrate was evaporated to afford
saturated ester (6) (0.95 g, 95%) as a yellowish semiꢀsolid, which determined to be
sufficiently pure (as per 1HꢀNMR). 1H NMR (300 MHz, CDCl3) δ 4.05 (t, J = 6.6 Hz, 2H),
2.31ꢀ2.26 (m, 4H), 1.61ꢀ1.59 (m, 6H), 1.28ꢀ1.25 (m, 18H), 0.87 (t, J = 6.0 Hz, 3H); 13C NMR
(75 MHz, CDCl3) δ 174.09 (2C), 64.38, 34.41, 31.45, 29.71, 29.40, 29.31, 29.18, 29.03,
28.81, 28.61, 25.89, 24.97, 24.65, 22.47, 14.01 (see the supporting information, Fig. S2).
159
11ꢀHydroxyundecanoic acid (7)
160
161
162
163
164
165
166
167
168
169
Saturated ester (6) (950 mg, 3.02 mmol) was treated with NaOH (1 N) in MeOHꢀH2O (4/1,
25 mL) at 60°C for 2.5 h. The reaction mixture was cooled, and pH was adjusted to 2 by
slowly adding aq. HCl (6 N) under iceꢀcold conditions. The reaction mixture was evaporated
to reduce the volume to about 5 mL (until water started to evaporate). The residual solvent
was saturated with NaCl and extracted with ethyl acetate (3 × 30 mL). The combined organic
extract was washed with saline (20 ml) and water (20 ml), dried over anhydrous Na2SO4,
filtered, and evaporated. The crude product was purified by silica gel column
chromatography using 50% ethyl acetateꢀhexane as the eluent to afford 11ꢀ
hydroxyundecanoic acid (7) (550 mg, 90%) as a white solid. 1H NMR (300 MHz, CDCl3) δ
3.63 (t, J = 6.6 Hz, 2H), 2.32 (t, J = 7.5 Hz, 2H), 1.64ꢀ1.51 (m, 4H), 1.35ꢀ1.25 (m, 12H); 13C
8

Page 9 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
170
NMR (75 MHz, CDCl3) δ 179.47, 62.96, 34.01, 32.60, 29.41, 29.28, 29.25, 29.12, 28.96,
25.63, 24.63 (see the supporting information, Fig. S3)
171
172
Undecanedioic acid (8)
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
A stock solution of H5IO6/CrO3 was prepared by dissolving periodic acid (H5IO6) (11.4 g, 50
mmol) and CrO3 (23 mg, 0.23 mmol) in wet acetonitrile (75% CH3CN/water) to a volume of
114 mL (complete dissolution typically required 1–2 h). The H5IO6/CrO3 stock solution (13
mL, 2.5 eq. of H5IO6, and 0.01 eq. of CrO3) was added to a solution of compound 7 (470 mg,
2.33 mmol) in wet acetonitrile (12 mL, 75% CH3CN/water) while maintaining the reaction
temperature at 0–5 °C for 30 min. The mixture was aged at 0°C for 45 min. The reaction was
quenched by adding an aqueous solution of Na2HPO4 (0.7 g in 7 ml H2O). Ethyl acetate (20
mL) was added and stirred. The organic layer was separated. The aqueous phase was
saturated with NaCl and extracted further with 10% MeOHꢀethyl acetate (4 × 20 ml). The
combined organic layer was washed with saline (15 ml), 5% NaHSO3 (15 mL), and saline (15
mL). The organic layer was dried over anhydrous Na2SO4 and concentrated to give the crude
dicarboxylic acid, which was further purified by recrystallization from ethyl acetateꢀhexane
to yield undecanedioic acid (8) (450 mg, 90%) as a white solid. 1H NMR (300 MHz, DMSOꢀ
d6) δ 2.17 (t, J = 6.6 Hz, 4H), 1.49ꢀ1.44 (m, 4H), 1.23 (m, 10H); 13C NMR (75 MHz, DMSOꢀ
d6) δ 174.90 (2C), 34.09 (2C), 29.23, 29.14 (2C), 28.98 (2C), 24.92 (2C) (see the supporting
information, Fig. S4).
189
190
191
Direct chemical conversion of ester intermediate (3) to undecanedioic acid (8)
The final product, undecanedioic acid (8) was synthesized directly without column
9

Green Chemistry
Page 10 of 25
View Article Online
DOI: 10.1039/C5GC01017A
192
193
194
195
196
197
198
chromatography through threeꢀstep chemical reactions of the crude ester. Crude ester (3)
(about 5.3 g, calculated amount of the ester: 4.0 g), which was obtained by centrifugation,
filtration, and extraction procedures using the reaction medium (volume: 300 mL, measured
concentration of the ester: 42.75 mM) after biotransformation, was subjected to
hydrogenation, hydrolysis, and oxidation as described above. Undecanedioic acid (8) (1.77 g,
65% yield based on the observed concentration of the crude ester) was isolated by
recrystallization at the final step.
199
200
201 Results and Discussion
202
Biocatalyst engineering for biotransformation of ricinoleic acid into ester 3
203
204
205
206
207
208
209
210
Wholeꢀcell biotransformation of ricinoleic acid (1) into 11ꢀhydroxyundecꢀ9ꢀenoic acid (4)
and nꢀheptanoic acid (5) (Scheme 1) was investigated in our previous study 17. As a result, the
biotransformation performance of the biocatalyst (i.e., recombinant E. coli BL21(DE3)
pACYCꢀADHꢀBVMO, pCOLAꢀPFEI expressing the ADH of M. luteus, the BVMO of P.
putida KT2440, and the esterase of P. fluorescens WI SIK) was limited by the toxic effect of
the final products on the host cells. Therefore, we examined biotransformation of ricinoleic
acid into ester 3 which is much less toxic to the cells (see the supporting information, Fig. S5),
which could be chemically converted into undecanedioic acid (Scheme 1 and 2).
211
212
Biotransformation productivity of the recombinant E. coli pACYCꢀADHꢀBVMO was
limited by low BVMO activity compared to ADH activity 17. Therefore, the BVMO was
10

Page 11 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
213
expressed with pJOEꢀBVMO, which allows for a higher copy number than that of the
pACYCꢀduet vector 21. Biotransformation with recombinant E. coli BL21(DE3) pACYCꢀ
ADH, pJOEꢀBVMO was initiated at the early stationary growth phase by adding 15 mM
ricinoleic acid and 0.5 g/L Tween80 to the culture broth. The starting material was converted
into ester 3 via 12ꢀketooctadecaꢀ9ꢀenoic acid (2) at a rate of 1.7 mM/h (0.43 g/L/h) at t < 6 h
(Fig. 1). The final product concentration and product yield was > 11 mM and 73%,
respectively. The biotransformation rate was ca. 50% greater compared to biotransformation
of ricinoleic acid to 11ꢀhydroxyundecꢀ9ꢀenoic acid and nꢀheptanoic acid 17. The high
biotransformation rate and low concentration of reaction intermediate 2 in the medium
suggested that the combination of pACYCꢀADH, pJOEꢀBVMO would be close to optimal for
expressing catalytic enzymes for wholeꢀcell biotransformation.
214
215
216
217
218
219
220
221
222
223
224
225
Biotransformation at high cell density
226
227
228
229
230
231
232
233
234
The next step was to increase final product concentration and volumetric productivity. This
was accomplished by enhancing the substrate and biocatalyst concentrations in the reaction
medium. The biotransformation was carried out after fedꢀbatch cultivation in a 1 L scale
bioreactor. When the recombinant E. coli BL21 pACYCꢀADH, pJOEꢀBVMO was grown to
10 g dry cells/L, ricinoleic acid was added to a final concentration of 30 mM in the culture
broth (Fig. 2A). The reaction dynamics were similar to those of the wholeꢀcell
biotransformation with 15 mM ricinoleic acid; the ester compound was stoichiometrically
produced from ricinoleic acid via 12ꢀketooctadecaꢀ9ꢀenoic acid (2). However, the final
product concentration and volumetric productivity increased to 25 mM and 6.2 mM/h,
11

Green Chemistry
Page 12 of 25
View Article Online
DOI: 10.1039/C5GC01017A
235
236
237
238
239
respectively, which were 2.3ꢀ and 3.6ꢀfold higher than the values obtained in the experiment
shown in Fig. 1 (Table 1). The volumetric productivity was ca. twoꢀfold greater than that of
recombinant E. coli pACYCꢀADHꢀ BVMO, pCOLAꢀPFEI during biotransformation of
ricinoleic acid into 11ꢀhydroxyundecꢀ9ꢀenoic acid and nꢀheptanoic acid under comparable
conditions 17.
240
241
242
243
244
Further increases in substrate and biocatalyst concentrations in the reaction medium to
63 mM and 20 g dry cells/L led to a final product concentration and volumetric productivity
of 53 mM (conversion yield, 84%) and 6.6 mM/h, respectively (Fig. 2B and Table 1). This
result indicates that the ester product was efficiently produced via wholeꢀcell biocatalysis at
high cell density.
245
246
Isolation of ester intermediate (3) after biotransformation
247
248
249
250
251
252
253
We used extraction, centrifugation, and filtration techniques to remove the cell mass and
isolate ester intermediate (3) after biotransformation. The ester intermediate was isolated by
simple extraction with ethyl acetate (2–3 times). The combined organic layer was washed
with saline, dried over sodium sulfate, evaporated, and chromatographed to afford the pure
ester intermediate. Ester (3) was recovered as a 90–95% crude mixture and was
purified/isolated to > 80% by column chromatography based on the observed concentration
of the ester intermediate after the biotransformation.
254
255
Chemical conversion of the ester intermediate to undecanedioic acid
12

Page 13 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
256
We developed a threeꢀstep chemical reaction including hydrogenation of the alkene,
hydrolysis of the ester, and oxidation of the alcohol to convert ester intermediate (3) to
undecanedioic acid (8) (Scheme 2). Stepꢀbyꢀstep reactions were examined initially to
optimize each step. At first, the alkene moiety was easily reduced using RaneyꢀNi under a
hydrogen atmosphere (1 atm) to afford the saturated ester 11ꢀ(heptanoyloxy)undecanoic acid
(6) as a semiꢀsolid at almost quantitative yield. Particularly, the RaneyꢀNi catalyst was very
effective and may be very economical for largeꢀscale reactions, compared to other metal
catalysts. Next, the ester was successfully hydrolyzed with sodium hydroxide in a coꢀsolvent
of methanol and water (4/1) at 60°C to provide 11ꢀhydroxyundecanoic acid (7) as a semiꢀ
solid with about a 90% yield. Oxidation of the terminal hydroxyl group to carboxylic acid
was the key step; thus, several oxidation conditions were examined. Among them, periodic
acid in the presence of a catalytic amount of chromium trioxide yielded the best conversion
efficiency, lowest cost, and mildness of the reaction conditions 26. 11ꢀHydroxyundecanoic
acid (7) was treated with a stock solution of periodic acid (2.5 eq.) and chromium trioxide
(0.01 eq.) in 75% aq. acetonitrile at 0°C to afford about 90% undecanedioic acid (8) after
recrystallization from ethyl acetateꢀhexane. The overall yield was about 80% over the three
stepꢀbyꢀstep syntheses. All intermediates were fully confirmed by their NMR spectra.
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
A threeꢀstep reaction will be needed for largeꢀscale preparations without any
purification procedures. Thus, we determined the optimal conditions without the column
purification steps. Ester intermediate (3) was prepared as a mixture of the ester and some
other fatty acids by simple centrifugation and filtration without column purification. After
hydrogenation of the mixture as described above, the methanol was partially evaporated.
Then, the esterꢀcontaining solution was treated with aq. sodium hydroxide (resulting
13

Green Chemistry
Page 14 of 25
View Article Online
DOI: 10.1039/C5GC01017A
279
280
281
282
283
284
285
286
287
288
289
290
291
292
concentration, 1 N) at 60°C for 2.5 h. After pH was adjusted to 2 by slow adding aq. HCl (6
N) at 0°C, the reaction mixture was evaporated to remove the lowꢀboiling point solvents and
extracted with ethyl acetate. Finally, the ωꢀhydroxy acidꢀcontaining mixture was treated with
periodic acid (2.5 eq.) and chromium trioxide (0.01 eq.) in 75% aq. acetonitrile at 0°C. After
simple extraction with ethyl acetate, the desired product, 1,11ꢀundecanedioic acid (8) was
purified by recrystallization from ethyl acetateꢀhexane. Based on the concentration of the
ester intermediate from the biotransformation, the overall yield of undecanedioic acid was >
65% after isolating the ester and the three chemical reactions, which was reasonable and
efficient after considering the purification yield of the ester and the stepꢀbyꢀstep chemical
yields over the three steps. Particularly, use of methanol was minimized and the timeꢀ
consuming, laborious and costꢀconsuming column purification steps were not needed. In
addition, the final product, 1,11ꢀundecanedioic acid, was obtained is a highly pure form by
recrystallization. We easily prepared 1–10 g of 1,11ꢀundecanedioic acid (8) in the laboratory
using these procedures, which could be applicable to future industrial preparations.
293
294
Feasibility of the ricinoleic acid transformation process
295
296
297
298
299
300
One of the key factors to influence feasibility of the ricinoleic acid transformation process for
industrial application may include final product concentration and volumetric productivity of
the biotransformation process, overall product yield, and cost of the reagents used for
bio/chemical transformations. Although cost of the reagents used for bio/chemical
transformations is difficult to estimate at the moment, the final product concentration and
volumetric productivity of the biotransformation can be compared between the complete
14

Page 15 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
301
biological transformation and the chemoꢀenzymatic transformation processes. As discussed
previously 17, the final product concentration and volumetric productivity of the complete
biological transformation process appeared to be limited in the range of 20 mM and 3 mM/h,
respectively, because of high toxicity of the reaction intermediates and final products (e.g., nꢀ
heptanoic acid, 11ꢀhydroxyundecꢀ9ꢀenoic acid) to the host cells (see the supporting
information, Fig. S5A). On the other hand, the final product concentration and volumetric
productivity of the chemoꢀenzymatic transformation process reached 53 mM and 6.6 mM/h,
respectively (Fig. 2B and Table 1). The final product concentration and volumetric
302
303
304
305
306
307
308
309
310
311
312
313
314
315
productivity of the chemoꢀenzymatic transformation process may further increase due to low
toxicity of the ester product (3) (see the supporting information, Fig. S5). Indeed, engineering
of the catalytic enzymes (e.g., BVMO) allowed the recombinant E. coli cells expressing the
ADH of M. luteus and the engineered BVMO of P. putida KT2440 to reach the final ester (3)
concentration and the volumetric productivity of over 100 mM in the reaction medium and 20
mM/h, respectively (unpublished data). This result indicates that cost effectiveness of the
chemoꢀenzymatic process could be larger than the complete biological transformation process.
316
317
Comparison to the complete chemical process
318
319
320
321
322
The key step and the most striking difference between the chemoꢀenzymatic and the complete
chemical processes for cracking ricinoleic acid (C18) into C7 and C11 products would be
cleavage method of the C11ꢀC12 bond of ricinoleic acid. This is driven by the BVMO
enzyme of P. putida KT2440 under physiological condition at the chemoꢀenzymatic process,
whereas conducted under high temperature over 450°C in case of the complete chemical
15

Green Chemistry
Page 16 of 25
View Article Online
DOI: 10.1039/C5GC01017A
323
324
325
326
327
328
process 6. The following step, oxidation of the cleavage products (i.e., 11ꢀhydroxyundecꢀ9ꢀ
enoic acid and methyl undecylenate at the chemoꢀenzymatic and complete chemical
processes, respectively) into 1,11ꢀundecanedioic acid would be comparable in terms of cost
effectiveness. Thereby, it was assumed that the chemoꢀenzymatic process could be less
energy consuming and more environmentꢀfriendly compared to the currently operating
chemical process.
329
330
331
332
333
334
335
Feasibility of the chemoꢀenzymatic transformation process could be also enhanced by
using castor oil instead of ricinoleic acid as the starting material. Ricinoleic acid accounts for
over 80% of the fatty acid constituents of castor oil, which is cheaper than ricinoleic acid and
commercially available. If a lipase is included into the reaction medium containing castor oil,
the plant oil will be hydrolyzed generating ricinoleic acid, oleic acid, and glycerol. Ricinoleic
acid will be converted into the ester product (3), whereas glycerol and oleic acid will be
consumed as energy sources by metabolic enzymes of E. coli.
336
337 Conclusion
338
339
340
341
342
343
344
A practical chemoꢀenzymatic synthesis of 11ꢀhydroxyundecanoic acid (7) and 1,11ꢀ
undecanedioic acid (8) from ricinoleic acid was examined. Biotransformation of ricinoleic
acid into ester (3) was driven by recombinant E. coli expressing the ADH from M. luteus and
the BVMO from P. putida KT2440. Ester (3) was chemically converted to 11ꢀ
hydroxyundecanoic acid (7) and then to 1,11ꢀundecanedioic acid (8) under fairly mild
reaction conditions. The wholeꢀcell biotransformation at high cell density (i.e., 20 g dry
cells/L) led to a final ester concentration and volumetric productivity of 53 mM and 6.6
16

Page 17 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
345
mM/h, respectively. The overall molar yield of 1,11ꢀundecanedioic acid from ricinoleic acid
was 55% based on the biotransformation and chemical transformation yields of 84% and 65%,
respectively. This method can be further applied to produce 9ꢀhydroxynonanoic acid and 1,9ꢀ
nonanedioic acid (azelaic acid) from oleic acid because 9ꢀ(nonanoyloxy)nonanoic acid can be
produced from oleic acid using recombinant E. coli expressing the C9 double bond hydratase
of Stenotrophomonas nitritireducens, the ADH from M. luteus, and the BVMO from P.
putida KT2440 14. This study will contribute to development of sustainable processes for the
production of medium chain fatty acids from renewable long chain fatty acids.
346
347
348
349
350
351
352
353
354 Acknowledgements
355
356
This study was supported by the R&D Program of MOTIE/KEIT [10044604 and 10044647],
Korea.
357
358
17

Green Chemistry
Page 18 of 25
View Article Online
DOI: 10.1039/C5GC01017A
359 References
360
1.
2.
U. Schorken and P. Kempers, Eur. J. Lipid Sci. Tech., 2009, 111, 627ꢀ645.
361
362
363
F. O. Ayorinde, E. Y. Nana, P. D. Nicely, A. S. Woods, E. O. Price and C. P.
Nwaonicha, J. Am. Oil Chem. Soc., 1997, 74, 531ꢀ538.
3.
4.
A. Kockritz and A. Martin, Eur. J. Lipid Sci. Tech., 2011, 113, 83ꢀ91.
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
388
M. A. R. Meier, J. O. Metzger and U. S. Schubert, Chem. Soc. Rev., 2007, 36, 1788ꢀ
1802.
5.
6.
7.
K. Kroha, Inform -Champaign-, 2004, 15, 568ꢀ571.
G. Das, R. K. Trivedi and A. K. Vasishtha, J. Am. Oil Chem. Soc., 1989, 66, 938ꢀ941.
U. Biermann, U. Bornscheuer, M. A. R. Meier, J. O. Metzger and H. J. Schafer,
Angew. Chem. Int. Ed., 2011, 50, 3854ꢀ3871.
8.
9.
J. O. Metzger, Eur. J. Lipid Sci. Technol., 2009, 111, 865ꢀ876.
J. M. Clomburg, M. D. Blankschien, J. E. Vick, A. Chou, S. Kim and R. Gonzalez,
Metab. Eng., 2015, 28, 202ꢀ212.
10.
E. J. Steen, Y. S. Kang, G. Bokinsky, Z. H. Hu, A. Schirmer, A. McClure, S. B. del
Cardayre and J. D. Keasling, Nature, 2010, 463, 559ꢀ562.
S. Sherkhanov, T. P. Korman and J. U. Bowie, Metab. Eng., 2014, 25, 1ꢀ7.
R. M. Lennen and B. F. Pfleger, Trends Biotechnol, 2012, 30, 659ꢀ667.
J. W. Song, E. Y. Jeon, D. H. Song, H. Y. Jang, U. T. Bornscheuer, D. K. Oh and J. B.
Park, Angew. Chem. Int. Ed., 2013, 52, 2534ꢀ2537.
11.
12.
13.
14.
15.
16.
17.
18.
J. W. Song, J. H. Lee, U. T. Bornscheuer and J. B. Park, Adv. Synth. Catal., 2014, 356,
1782ꢀ1788.
H. Y. Oh, S. U. Kim, J. W. Song, J. H. Lee, W. R. Kang, Y. S. Jo, K. R. Kim, U.
Bornscheuer, D. K. Oh and J. B. Park, Adv. Synth. Catal., 2015, 357, 408ꢀ416.
S. U. Kim, K. R. Kim, J. W. Kim, S. Kim, Y. U. Kwon, D. K. Oh and J. B. Park, J.
Agric. Food Chem., 2015, 63, 2773ꢀ2781.
H. Y. Jang, E. Y. Jeon, A. H. Baek, S. M. Lee and J. B. Park, Process Biochem., 2014,
49, 617ꢀ622.
A. H. Baek, E. Y. Jeon, S. M. Lee and J. B. Park, Biotechnol. Bioeng., 2015, 112,
889ꢀ895.
18

Page 19 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
389
19.
20.
21.
22.
23.
E. Y. Jeon, A. H. Baek, U. T. Bornscheuer and J. B. Park, Appl. Microbiol.
Biotechnol., 2015, 99, 6267ꢀ6275.
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
A. Kirschner, J. Altenbuchner and U. T. Bornscheuer, Appl. Microbiol. Biotechnol.,
2007, 73, 1065ꢀ1072.
J. Rehdorf, A. Kirschner and U. T. Bornscheuer, Biotechnol. Lett., 2007, 29, 1393ꢀ
1398.
H. L. van Beek, H. J. Wijma, L. Fromont, D. B. Janssen and M. W. Fraaije, FEBS
Open Bio, 2014, 4, 168ꢀ174.
J. Muschiol, C. Peters, N. Oberleitner, M. D. Mihovilovic, U. T. Bornscheuer and F.
Rudroff, Chem. Commun., 2015, 51, 5798ꢀ5811.
24.
25.
26.
S. Wallace and E. P. Balskus, Curr. Opin. Biotechnol., 2014, 30, 1ꢀ8.
D. Riesenberg, Curr. Opin. Biotechnol., 1991, 2, 380ꢀ384.
M. Zhao, J. Li, Z. Song, R. Desmond, D. M. Tschaen, E. J. J. Grabowski and P.
Reider, Tetrahedron Lett., 1998, 39, 5323ꢀ5326.
19

Green Chemistry
Page 20 of 25
View Article Online
DOI: 10.1039/C5GC01017A
406
407
Table 1. Biocatalytic performance of the recombinant Escherichia coliꢀbased biocatalyst
Reaction at
Reaction at fedꢀ
Reaction at fedꢀ
Reaction at fedꢀ
batch cultivation batch cultivation 1
batch cultivation 2 batch cultivation 3 c
Substrate concentration
(mM)
15
3
30
10
63
20
30
10
Biocatalyst concentration
(g dry cells/L)
Final product
11
25
53
21
concentration (mM)
Volumetric productivity
(mM/h) a
1.7
6.2
6.6
3.2
Product yield
(%)b
73
83
84
70
408
409
410
411
a Volumetric productivity was calculated based on the product concentration, which was
determined by gas chromatography/liquid chromatography (GC/MS), and biotransformation
time, which was measured when > 90% of the starting material was converted to the
products.
412
413
b Product yield was calculated based on substrate depletion and product concentration, which
were determined by GC/MS.
414
415
416
c Recombinant E. coli pACYCꢀADHꢀBVMO, pCOLAꢀPFEI was used to biotransformation
of ricinoleic acid to 11ꢀhydroxyundecꢀ9ꢀenoic acid. The result was adopted from our
previous study 17.
20

Page 21 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
417
Scheme Legends
418
419
420
421
Scheme 1. Designed biotransformation pathway. Ricinoleic acid (1) is enzymatically
converted into ester 3 (see the Results section for details), which can be hydrolyzed into nꢀ
heptanoic acid and (Z)ꢀ11ꢀhydroxyundecꢀ9ꢀenoic acid (4) 13. Adopted from our previous
study 13.
422
423
Scheme 2. Chemical conversion of ester intermediate (3) to undecanedioic acid (8)
424
Figure Legends
425
426
427
428
429
430
Figure 1. Time course of the biotransformation of ricinoleic acid (1) by recombinant
Escherichia coli BL21(DE3) pACYCꢀADH, pJOEꢀBVMO, expressing the alcohol
dehydrogenase (ADH) from Micrococcus luteus and the BVMO from Pseudomonas putida
KT2440. The biotransformation was initiated at the stationary growth phase (cell density: 3 g
dry cells/L) in Riesenberg medium by adding 15 mM ricinoleic acid and 0.5 g/L Tween80 to
the culture broth. Symbols indicate concentrations of ricinoleic acid (1) (△), 10ꢀketooleic acid
(2) (▽), and 3 (■).
431
432
433
434
435
436
Figure 2. Biotransformation of ricinoleic acid by Escherichia coli pACYCꢀADH, pJOEꢀ
BVMO at high cell density. Biotransformation was initiated by adding (A) 30 mM ricinoleic
acid and 0.5 g/L Tween80 to the culture broth after fedꢀbatch cultivation to a cell density of
10 g dry cells/L or (B) 63 mM ricinoleic acid and 0.5 g/L Tween80 to the culture broth after
fedꢀbatch cultivation to a cell density of 20 g dry cells/L. Symbols indicate concentrations of
ricinoleic acid (1) (△), 10ꢀketooleic acid (2) (▽), and 3 (■).
437
438
21

Green Chemistry
Page 22 of 25
View Article Online
DOI: 10.1039/C5GC01017A
439
440
Scheme 1
OH
O
OH
5
1
NAD+
Alcohol dehydrogenase
NADH
O
O
OH
5
2
NADPH
O2
Baeyer-Villiger monooxygenase
H2O
NADP+
O
O
OH
5
O
3
H2O
Esterase
O
OH
HO
OH
O
5
4
441
442
22

Page 23 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
443
444
445
Scheme 2
446
447
448
449
450
451
452
23

Green Chemistry
Page 24 of 25
View Article Online
DOI: 10.1039/C5GC01017A
453
454
455
Fig. 1
15
12
9
15
12
9
6
6
3
3
0
0
0
2
4
6
8
10
12
Reaction time (h)
456
457
458
24

Page 25 of 25
Green Chemistry
View Article Online
DOI: 10.1039/C5GC01017A
459
460
461
462
Fig. 2
A
30
24
18
12
6
30
24
18
12
6
0
0
0
2
4
6
8
Reaction time (h)
463
464
465
B
60
45
30
15
0
60
45
30
15
0
0
2
4
6
8
Reaction time (h)
466
25