Biocatalytic synthesis of non-vicinal aliphatic diols

Article abstract of DOI:10.1039/d0ob02086a

Biocatalysts are receiving increased attention in the field of selective oxyfunctionalization of C-H bonds, with cytochrome P450 monooxygenases (CYP450s), and the related peroxygenases, leading the field. Here we report on the substrate promiscuity of CYP505A30, previously characterized as a fatty acid hydroxylase. In addition to its regioselective oxyfunctionalization of saturated fatty acids (ω-1-ω-3 hydroxylation), primary fatty alcohols are also accepted with similar regioselectivities. Moreover, alkanes such as n-octane and n-decane are also readily accepted, allowing for the production of non-vicinal diols through sequential oxygenation. This journal is

Full text of DOI:10.1039/d0ob02086a

Organic &
Biomolecular Chemistry
View Article Online
View Journal | View Issue
PAPER
Biocatalytic synthesis of non-vicinal aliphatic
diols†
Cite this: Org. Biomol. Chem., 2021,
19, 439
Ana C. Ebrecht,a Jasmin C. Aschenbrenner, a,b Martha S. Smit a,b and
a
Diederik J. Opperman
*
Biocatalysts are receiving increased attention in the field of selective oxyfunctionalization of C–H bonds,
with cytochrome P450 monooxygenases (CYP450s), and the related peroxygenases, leading the field.
Here we report on the substrate promiscuity of CYP505A30, previously characterized as a fatty acid
hydroxylase. In addition to its regioselective oxyfunctionalization of saturated fatty acids (ω-1 –
ω-3 hydroxylation), primary fatty alcohols are also accepted with similar regioselectivities. Moreover,
alkanes such as n-octane and n-decane are also readily accepted, allowing for the production of non-
vicinal diols through sequential oxygenation.
Received 13th October 2020,
Accepted 3rd December 2020
DOI: 10.1039/d0ob02086a
rsc.li/obc
auxiliary redox partners has the advantage that only a single
biocatalyst has to be produced and the reaction mixture only
Introduction
Selective oxyfunctionalization of C–H bonds remains a signifi- requires the cofactor, molecular oxygen and substrate. In
cant challenge in chemical synthesis. This problem is further addition, self-sufficient CYP450s are generally considered
compounded for compounds without any directing functional more efficient and, in some cases, more stable than systems
groups such as n-alkanes. Biocatalysts such as cytochrome with multiple components.11
P450 monooxygenases (CYP450s),1 and more recently peroxy-
Members of the CYP102 family of CYP450s are considered
genases,2 have emerged as promising solutions to this fatty acid hydroxylases, catalysing, amongst others, the sub-
problem. With these heme-thiolate biocatalysts, the protein terminal (ω-1 to ω-3) hydroxylation of saturated fatty acids.12
structure provides a scaffold that restricts the binding orien- More in-chain hydroxylation activity towards long-chain fatty
tations, thus “selecting” the C–H groups accessible to the acids, albeit with low regioselectivity, has been observed for
heme-cofactor for oxyfunctionalization. A striking example of certain members of the CYP102 family.13 To date, the catalytic
this is the ability of some CYP450s (including CYP153s and scope and corresponding regio-, enantio- and stereoselectivity/
CYP52s) to perform terminal hydroxylation of alkanes and specificity of BM3 have been greatly enhanced through
fatty acids.3
directed evolution,14,15 rational design16 and the use of decoy
The CYP450, CYP102A1 (commonly referred to as BM3), molecules.17–19
has become the workhorse for selective oxyfunctionalization
CYP505, a family of self-sufficient fungal CYP450s, has also
reactions.4–6 Its prominence is due in part to its ease of pro- been classified as fatty acid hydroxylases, based in part on
duction and use and also an available X-ray crystal structure their remarkable sequence similarity to CYP102s, as well as
that was solved in the early 1990s,7,8 allowing catalytic and experimental evidence showing sub-terminal (ω-1–ω-3)
mechanistic insights.9,10 CYP102A1 is a self-sufficient CYP450 hydroxylation of C9–C18 fatty acids by CYP505A1 (P450 foxy).20
which does not require auxiliary proteins for activity. For cata- However, similar to CYP102s, the true physiological function
lytic activity, CYP450s require electrons that are donated, in and in vivo substrate for most of the members of this family
general, from reduced nicotinamide cofactors (NADH or remains unknown. Recently, the activity of CYP505D6 from
NADPH). The electrons are sequentially transferred from the Phanerochaete chrysosporium was investigated with respect to
reduced cofactors to the CYP450 heme center via specific its 1-dodecanol metabolism. Among the seven CYP505D P450s
redox partner proteins. Fusion between the CYP450 and these encoded by P. chrysosporium, two (CYP505D4 and D6) showed
higher expression profiles when induced with 1-dodecanol
compared to dodecanoic acid. Further investigation, however,
aDepartment of Biotechnology, University of the Free State, 205 Nelson Mandela
revealed CYP505D6 to show very little regioselectivity, produ-
Drive, Bloemfontein, 9300, South Africa. E-mail: opperdj@ufs.ac.za
bSouth African DST-NRF Centre of Excellence in Catalysis, c*change, South Africa
cing ω-1 to ω-7 hydroxylated products from 1-dodecanol.21
CYP505A30 has previously been described as a fatty acid
†Electronic supplementary information (ESI) available. See DOI: 10.1039/
hydroxylase able to hydroxylate saturated fatty acids (C12–C15)
d0ob02086a
This journal is © The Royal Society of Chemistry 2021
Org. Biomol. Chem., 2021, 19, 439–445 | 439

View Article Online
Paper
Organic & Biomolecular Chemistry
as well as arachidonic acid.22 The regioselectivity of that reported by Baker and co-workers.22 The remaining sub-
CYP505A30 mirrors that of BM3, where primarily sub-terminal strates were compared to the HS accumulation observed for
hydroxylation is observed, albeit in different product ratios. tetradecanoic acid (taken as 100%). The fatty alcohols only
Our research group recently reported on the in-chain (ω-7) induced approximately 60% HS in CYP505A30 and even lower
hydroxylation by the closely related CYP505E3 from Aspergillus (by ca. 10%) in the truncated HD. For the alkanes, the HS
terreus.23 Although hydroxylation of saturated fatty acids (C12– accumulation was between 30–60% in the full-length enzyme
C16) was observed, higher product formation (TTNs) was and 10–20% in the HD.
achieved with medium-chain alcohols (C10 and C12) and
The titration spectra were also used to estimate the dis-
even decane. Over-hydroxylation was also observed. This sociation constants (KD) (Table S1†). Longer chain fatty acids
sequential reactivity is not uncommon for CYP505s, as and fatty alcohols exhibited lower KD values, particularly for
CYP505B1 (FUM6) hydroxylates the branched-chain polyketide- 1-tetradecanol and 1-hexadecanol, with
amino acid precursor of fumonisin sequentially at the C14 and suggesting tighter binding to the enzyme. For alkanes, higher
C15 positions.24
KD values were estimated, with little variation between chain
a KD < 1 µM,
All this prompted us to explore the use of CYP505A30 as a lengths. This estimation, however, could be an artifact of the
fatty alcohol and alkane hydroxylase for the production of ali- low solubility of alkanes in water (less than 4 µM),29 which is
phatic diols. Whereas the synthesis of vicinal aliphatic diols lower than the calculated KD values. The limitation in the sub-
such as 1,2-diols from alkenes is well established, with the strate concentration may lead to an overestimation of the
regioselectivity directed by the CvC double bond, the syn- affinity for the compound. The same effect applies to 1-hexade-
thesis of non-vicinal aliphatic diols (apart from α,ω-diols) are canol since the KD value is higher than the solubility of the
less explored. These aliphatic diols are important building compound (∼0.17 µM).30
blocks for lubricants and new polymers.25,26
Kinetic characterization
Steady-state kinetics of CYP505A30 was performed by moni-
toring the oxidation of NADPH spectrophotometrically at
Results and discussion
340 nm. The fatty acids and primary fatty alcohols exhibited
Biocatalyst production
Michalis-Menten behavior (Fig. S9 and S10†). Contrary to
Similar to P450 BM3 (CYP102A1), CYP505A30 is a self-
sufficient P450, where the N-terminal CYP450 domain is
linked to a diflavin reductase domain (similar to the mamma-
lian cytochrome P450 reductase, CPR), requiring NADPH for
activity. CYP505A30 and a truncated version consisting of only
the heme-domain (CYP505A30HD) were recombinantly
expressed in E. coli, with both mostly peripherally associated
with the membranes. Expression analysis of the CYP505A30
also revealed truncations of the protein that ultimately co-puri-
fied with the N-terminally His6-tagged enzyme. A C-terminal
His6-tagged version was therefore created and could be puri-
fied to near homogeneity (Fig. S1 and 2†) with an estimated
yield of 2 mg L−1. The heme-domain was purified as a
CYP505A1 from Fusarium oxysporum, which exhibited sub-
strate inhibition for longer chain saturated fatty acids (C14–
C16),31 no substrate inhibition was observed with fatty acids
nor with the fatty alcohols tested. With alkanes, however, it
was not possible to perform kinetic characterization due to
the low solubility of the compounds. Only turnover frequen-
cies (kobs) are reported at substrate concentrations of 10 mM
(Table S1†).
Baker et al. reported pentadecanoic acid as the preferred
substrate of CYP505A30.22 With the exception of decanoic
acid, CYP505A30 exhibited higher kcat values for fatty acids
compared to the corresponding fatty alcohols (Table S1†).
Lower KM values for the corresponding fatty alcohols were,
however, generally observed. The catalytic efficiency (kcat/KM)
calculated was thus better with the fatty alcohols compared
to their fatty acid counterparts/equivalents (Fig. 1).
N-terminally His6-tagged protein with yields of up to 9 mg L−1
.
Spectral substrate binding
With CYP450s, the binding of the substrate typically induces Although 1-dodecanol yielded the best results, the kcat/KM
the displacement of the distal water molecule bound to the decreased drastically for longer chain fatty alcohols, with
catalytic heme-iron, resulting in a change of absorbance in the 1-tetradecanol and 1-hexadecanol displaying catalytic
spectrum of the enzyme. UV-Vis spectra of both the efficiency 100-fold lower than for tetradecanoic and hexade-
CYP505A30 and the truncated heme-domain, titrated with canoic acid, respectively. Hydrogen peroxide formation,
fatty acids, fatty alcohols and alkanes, were therefore com- which partially accounts for reducing equivalents lost to
pared. Both the full-length CYP505A30 and the heme-domain uncoupling, was also measured during the reactions with
bound a range of different substrates tightly, shifting the Soret selected fatty acids, alkanes and fatty alcohols, and found to
band from 418 nm to 388 nm, with an isosbestic point around constitute less than 16% (Table S1†) of the reducing equiva-
406 nm (Fig. S3–S8†), characteristic of type I substrate lents consumed.
binding.27,28 The extent of induced high-spin (HS) iron state
varied depending on the substrates (Table S1†). The highest
Biotransformations
absorbance difference observed was for the C12–C16 fatty Encouraged by the spectral and kinetic characterization, we
acids in the full-length enzyme and C14 for the HD, similar to performed biotransformations to determine the substrate
440 | Org. Biomol. Chem., 2021, 19, 439–445
This journal is © The Royal Society of Chemistry 2021

View Article Online
Organic & Biomolecular Chemistry
Paper
Fig. 1 Catalytic efficiency of CYP505A30 with different fatty acids and
primary fatty alcohols. Reaction conditions: 100 mM potassium phos-
phate buffer (pH 8), [CYP505A30] = 0.035–1 µM, [NADPH] = 0.3 mM, T
= 25 °C.
Fig. 2 Conversion of C8–C12 saturated fatty acids (FA) and primary
fatty alcohols (OH) by purified CYP505A30 (Purif) and CYP505A30
within E. coli cell free extracts (CFE). Reaction conditions: 200 mM pot-
assium phosphate buffer (pH 8), [CYP505A30] = 4 µM, [BmGDH] = 0.2 U
mL−1, [glucose] = 200 mM, [NADP+] = 0.3 mM, [FA or OH] = 10 mM, T =
30 °C, shaking = 200 rpm, t = 24 h (Purif), t = 2 h (CFE).
scope and regioselectivity of CYP505A30. Reactions (1 mL)
were performed with both purified biocatalyst as well as E. coli
cell free extract (CFE) containing CYP505A30 and allowed to
proceed for 0.5–24 h, where after the products were extracted
with an equal amount of ethyl acetate, derivatized and ana-
lysed by GC-MS. With CFE biotransformations, complete con-
version was observed within 2 h for both sets of fatty acids and
primary fatty alcohols tested (C8–C12), except for octanoic acid
that reached 90% conversion. Product yields from purified
enzymes were significantly lower, with even the 24 h reactions
not reaching the conversion levels seen with CFE biotrans-
formations after 2 h. This is consistent with our previous
results with alkane hydroxylation with CYP450s, where the
CFE contributes significantly to the stabilization of the
enzyme,32,33 allowing much higher TTNs. Additionally, the
CFE could increase the solubility of these substrate.34
(formation of triols) also observed for 1-decanol and 1-dodeca-
nol (Fig. S12†).
In addition to primary fatty alcohols, we also tested the
hydroxylation of different secondary fatty alcohols (2-, 3- and
4-octanol, 2-decanol, and 2-dodecanol). The enzyme also
accepted all these fatty alcohols as substrates, with near com-
plete conversion within 2 h, except for 3- and 4-octanol
(Table 1). The regioselectivities of CYP505A30 was mostly unal-
tered between the 1-, 2-, and 3-octanol, whereas 4-octanol gave
higher amounts of ω-1 (C7) or C2 hydroxylated products and
no ω-3 hydroxylation. The selectivity for ω-1 hydroxylation
decreased with 2-decanol and 2-dodecanol (Table 1). Reactions
again led to triol formation, a behavior seemingly character-
Comparable activities were observed between the fatty acids
and primary fatty alcohols, with TOF values ranging between
20 and 50 min−1 (Table S2†). The major products observed
were ω-1 and ω-2 hydroxylated fatty acids and fatty alcohols,
and ω-3 in a lower amount (<8%) (Fig. 2). Baker et al. showed
hydroxylation of dodecanoic and tetradecanoic acid in the
positions ω-1 to ω-3, with a high ratio of ω-1 hydroxylation.22
Our analysis of the product formation with fatty acids showed
the regioselectivity to be dependent on the chain-length. In
general, more than 70% hydroxylation in the ω-1 position was
observed. Upon complete conversion (after 2 h), overhydroxyla-
tion of dodecanoic acid occurred. For decanoic acid, overhy-
droxylated products were only observed at later time points
(Fig. S11†). CYP505A30 showed similar regioselectivities with
the corresponding primary fatty alcohols, with overoxidation
Table 1 Conversion of fatty alcohols by CYP505A30
Conv (2 h)
ω-1
ω-2
ω-3
Triols
23
1-Octanol
2-Octanol
3-Octanol
4-Octanol
1-Decanol
2-Decanol
1-Dodecanol
2-Dodecanol
>99%
>99%
83%
55
42
33
3
3
6
64
53a
24
71%
53a
18b
>99%
>99%
>99%
93%
44 [19]
51
27 [18]
43 [19]
27 [7]
46
15 [11]
28 [17]
6 [2]
3
58
22
Reaction conditions: 200 mM potassium phosphate buffer (pH 8),
0.8 mL CFE [CYP505A30] = 4 µM, [BmGDH] = 0.2 U mL−1, [glucose] =
100 mM, [NADP+] = 0.3 mM, [alcohol] = 10 mM, T = 30 °C, shaking =
200 rpm,
C2 hydroxylation. b Includes C3 hydroxylation.
t = 2 h. Values in brackets t =
30 min. a Includes
This journal is © The Royal Society of Chemistry 2021
Org. Biomol. Chem., 2021, 19, 439–445 | 441

View Article Online
Paper
Organic & Biomolecular Chemistry
istic of the CYP505 family. A. terreus CYP505E3, also exhibited
double-hydroxylation of long-chain fatty acids and alcohols,
with higher percentages in longer chain substrates.23
In addition, the CYP505 FUM6, is also involved in sequen-
tial hydroxylation steps of an already functionalized long-chain
alkyl substrate as part of its physiological function.24
CYP505A30 also catalysed the hydroxylation of n-octane and
n-decane, producing the 2- and 3-alcohols as the major pro-
ducts. Unlike CYP102A1 which only showed mono-hydroxy-
lation events with n-octane and n-decane,32,35 the alcohol pro-
ducts served as substrates for CYP505A30 in a second round of
hydroxylation to produce various diols.
Higher activity was observed with n-octane, with a TOF of
ca. 74 min−1 (Table S2†), yielding ∼36 mM total products after
24 h (TTN ca. 10700). The total product achieved with n-octane
was remarkable considering the yields reported in the litera-
ture. Optimized CYP102A1 mutants reached a maximum of
10 mM products, with a TTN of 4200 at best for the CFE reac-
tions with n-octane.32 Three major diol products were formed,
namely 2,7-, 2,6- and 3,6-octanediol (Fig. 3). The symmetrical
diols were formed from sequential ω-1 or ω-2 hydroxylations of
n-octane, whereas the 2,6-octanediol is a convergent product
from either ω-1 or ω-2 hydroxylation of 3-octanol or 2-octanol
respectively (Scheme 1 and S1, Fig. S15†). A similar trend was
observed with n-decane (Scheme S2, Fig. S15†) despite lower
TOF (ca. 9 min−1) and total products (ca. 8 mM, TTN 3100)
obtained within the same time period. However, diols consti-
Scheme 1 Sequential regioselective oxyfunctionalization of n-octane
in the production of non-vicinal diols.
tuted nearly 60% of the total products, possibly due to the low
solubility of n-decane and the decanol products formed being
the preferred substrate.
Similar product titers were reported with CYP102A1 and its
mutants32 and CYP505E323 (ca. 5–8 mM), further indicating
that substrate solubility could be limiting the reaction. The
scalability of the reactions was evaluated by performing a
10 mL reaction with n-octane. Despite a reduction in the TTNs
to ca. 9100, 23.4 mM total product, containing 4 mM diol, was
still observed after 24 h, translating to a space time yield (STY)
of 3.1 g L−1 day−1 in the aqueous phase.
Conclusions
CYP505A30, characterized initially as a sub-terminal fatty acid
hydroxylase, also accepts fatty alcohols and n-alkanes as sub-
strates. Similar, or even higher in the case of 1-decanol, cata-
lytic activities and conversions were observed for the shorter
chain fatty alcohols compared to the equivalent fatty acids.
The regioselectivity of CYP505A30 was unaffected by the
nature of the functionalized terminal group. Moreover, this
regioselectivity also remained unaffected regardless of the
position of the functional group (i.e. terminal alcohols vs. sub-
terminal alcohols). Due to the substrate promiscuity of
CYP505A30, n-alkanes could be sequentially converted via fatty
alcohol intermediates to non-vicinal aliphatic diols. We antici-
pate that similar to CYP102A1,14,36 the substrate scope and
regioselectivity of CYP505A30 is tuneable through directed
evolution, which would enable the production of single sym-
metrical non-vicinal diols. This unique activity opens up new
Fig. 3 GC-MS identification of silylated diols formed during conversion
of n-octane by CYP505A30. Reaction conditions: 200 mM potassium
phosphate buffer (pH 8), [CYP505A30] in CFE (0.8 mL)
= 4 µM,
[BmGDH] = 0.2 U mL−1, [glucose] = 200 mM, [NADP+] = 0.3 mM,
[octane] = 20% (v/v), T = 30 °C, shaking = 200 rpm, t = 24 h.
442 | Org. Biomol. Chem., 2021, 19, 439–445
This journal is © The Royal Society of Chemistry 2021

View Article Online
Organic & Biomolecular Chemistry
Paper
routes in polymer chemistry from secondary sub-terminal buffer pH 7.4, 40 mM imidazole, 500 mM NaCl) containing
diols, compared to their α,ω-diol counterparts.
1% (v/v) Triton X-100. After disruption of the cells and cen-
trifugation, the soluble fraction was loaded onto a HisTrap FF
column, and the protein was eluted in a linear gradient of
imidazole 40–500 mM. The last step consisted of a SEC using
buffer C containing 10% (v/v) glycerol. The enzyme eluted was
pooled and concentrated as described above.
Experimental
Biocatalyst production and purification
The open reading frame (ORF) coding for CYP505A30 from
Bacillus megaterium glucose dehydrogenase (BmGDH) was
Thermothelomyces thermophilus (Myceliophthora thermophila) expressed from pET28b(+) and purified as previously
ATCC 42464 (NCBI CP003004.1) was synthesized and cloned by described.38
GenScript into pET28b(+) via NdeI and HindIII to allow the pro-
Protein concentration was determined by Pierce BCA assay
duction of an N-terminal hexahistidine tagged enzyme. The (ThermoFisher Scientific) using bovine serum albumin as a
ORF was sub-cloned to pET22b(+) using the same restriction standard. CYP concentrations were determined using CO-
sites. To create a C-terminally hexahistidine tagged version, difference spectra.
the stop codon and a part of the plasmid backbone were
deleted using inverse PCR with primers CTH-F (5′-CAC CAC
CAC CAC CAC TGA GAT C-3′) and CTH-R (5′- GTC AAA GAC
ATC TGT GGC ATA CCG GTC-3′), as previously described.37 The
CPR domain was similarly removed from the pET28 construct
to create a N-terminally hexahistidine tagged variant of only
the heme domain (CYP505A30HD) using primers HD-F (5′-TAA
AAG CTT GCG GCC GCA CTC GAG CAC-3′) and HD-R (5′-GTC
TCG AAG AAT GGC CCG CAT GTA GAA GTC-3′).
Constructs were transformed into E. coli BL21-Gold(DE3)
(Stratagene) and selected for on LB-medium supplemented
with the appropriate antibiotic (100 µg mL−1 ampicillin or
30 µg mL−1 kanamycin). Heterologous expression of the CYPs
were performed in auto-induction media (ZYP-5052) sup-
plemented with 0.5 mM δ-aminolevulinic acid hydrochloride
and 50 μM FeCl3·6H2O (25 °C for 48 h).
For purification of CYP505A30, cells were harvested by cen-
trifugation (7000g, 10 min, 4 °C) and resuspended (0.2 g wet
weight mL−1) in buffer A (50 mM potassium phosphate buffer
pH 7.4) containing 1% (v/v) Triton X-100. Disruption of the
cells was carried out by single passage through a One-Shot Cell
disrupter System (Constant Systems Ltd) at 30 kPsi, followed
by centrifugation (30 000g, 30 min, 4 °C) and ultracentrifuga-
tion (100 000g, 90 min, 4 °C). The resulting soluble fraction
was loaded onto a 5 mL HisTrap FF column (GE Healthcare),
previously equilibrated with buffer A. The loaded column was
washed with 10 column volumes of buffer A containing 40 mM
Steady-state kinetics
Steady-state kinetics were performed by spectrophotometrically
monitoring the oxidation of NADPH at 340 nm (ε340 =
6.22 mM−1 cm−1), in 1 mL reactions containing 100 mM pot-
assium phosphate buffer pH 8, diluted CYP505A30
(0.035–1 μM). Reactions were initiated by the addition of
0.3 mM NADPH. Substrates were prepared in DMSO. DMSO
concentration did not exceed 1% of the total volume in the
reaction mixtures. Assays were carried out at 25 °C in a 1 cm
path-length cuvette using a Cary 300 Bio UV/Vis spectrophoto-
meter (Varian). Kinetic constants were determined by non-
linear regression to the Michaelis–Menten equation using the
Origin software (OriginLab).
Hydrogen peroxide uncoupling was measured as the per-
centage of NADPH that is used to produce hydrogen peroxide
during the reaction. CYP505A30 reactions were performed as
described above, containing 0.1 mM NADPH, 0.15 µM
CYP505A30 and excess substrate. Reactions were monitored
spectrophotometrically until complete consumption of
NADPH, where after hydrogen peroxide was quantified using
Ampliflu™ Red assay.39 Ampliflu solution consisted of 100 µM
Ampliflu Red, 0.2 U horseradish peroxidase and 2 U super-
oxide dismutase.
Substrate binding titrations
NaCl and 40 mM imidazole. Protein was eluted using a linear Spectral studies of CYP505A30(HD) were performed at 25 °C in
gradient of increasing NaCl and imidazole (40–500 mM). 100 mM potassium phosphate pH 8. Reactions contained
Fractions containing CYP505A30 were pooled, concentrated 2.5 μM enzyme in a final volume of 1 mL. Stock solutions of
using ultrafiltration (Amicon Ultra-30 kDa MWCO; Merck), and substrates were prepared in DMSO, and titrations were per-
desalted using PD-10 columns (GE Healthcare) equilibrated formed by the stepwise substrate addition of 0.5–2 μL, with the
with buffer C (50 mM potassium phosphate buffer pH 8.0). total volume added never exceeding 1% of the total sample
The desalted protein was loaded onto a 5 mL HiTrap Q HP volume. UV-Vis spectra (300–750 nm) were recorded for the
column (GE Healthcare), and the protein was eluted using a substrate-free enzyme, as well as after every titration until no
linear gradient of increasing NaCl (0–500 mM). Finally, size further shifts in the spectra were observed. The difference in
exclusion chromatography (SEC) was performed using absorbance along the spectrum was used for identification of
Sephacryl S200HR (XK 26 column, GE Healthcare) equilibrated minimum and maximum wavelengths. The difference in
with buffer C containing 100 mM NaCl and 10% (v/v) glycerol. absorbance between minimum and maximum were plotted
The eluted enzyme was pooled, concentrated, and stored at against substrate concentration to determine the dissociation
4 °C for up to 5 days. For CYP505A30HD purification, cells constant (KD). Data were fitted using Michaelis–Menten or
were resuspended in buffer B (50 mM potassium phosphate Morrison function using Origin software (OriginLab).
This journal is © The Royal Society of Chemistry 2021
Org. Biomol. Chem., 2021, 19, 439–445 | 443

View Article Online
Paper
Organic & Biomolecular Chemistry
Biotransformations
8 H. Li and T. L. Poulos, Nat. Struct. Biol., 1997, 4, 140–146.
9 M. A. Noble, C. S. Miles, S. K. Chapman, D. A. Lysek,
A. C. Mackay, G. A. Reid, R. P. Hanzlik and A. W. Munro,
Biochem. J., 1999, 339, 371–379.
10 L. A. Cowart, J. R. Falck and J. H. Capdevila, Arch. Biochem.
Biophys., 2001, 387, 117–124.
11 F. Hannemann, A. Bichet, K. M. Ewen and R. Bernhardt,
Biochim. Biophys. Acta, Gen. Subj., 2007, 1770, 330–344.
12 S. S. Boddupalli, R. W. Estabrook and J. A. Peterson, J. Biol.
Chem., 1990, 265, 4233–4239.
13 S. D. Munday, N. K. Maddigan, R. J. Young and S. G. Bell,
Biochim. Biophys. Acta, Gen. Subj., 2016, 1860, 1149–1162.
14 M. W. Peters, P. Meinhold, A. Glieder and F. H. Arnold,
J. Am. Chem. Soc., 2003, 125, 13442–13450.
15 G. D. Roiban and M. T. Reetz, Chem. Commun., 2015, 51,
2208–2224.
16 T. W. B. Ost, C. S. Miles, J. Murdoch, Y. F. Cheung,
G. A. Reid, S. K. Chapman and A. W. Munro, FEBS Lett.,
2000, 486, 173–177.
17 F. E. Zilly, J. P. Acevedo, W. Augustyniak, A. Deege and
M. T. Reetz, Angew. Chem., Int. Ed., 2011, 50, 2720–2724.
18 S. D. Munday, S. Dezvarei and S. G. Bell, ChemCatChem,
2016, 8, 2789–2796.
19 S. D. Munday, O. Shoji, Y. Watanabe, L. L. Wong and
S. G. Bell, Chem. Commun., 2016, 52, 1036–1039.
20 N. Nakayama, T. Asako and H. Shoun, J. Biochem., 1996,
119, 435–440.
Biotransformations were performed in 40 mL amber vials,
with a reaction mixture consisting of 200 mM potassium phos-
phate buffer pH 8, 200 mM glucose, 4 μM CYP505A30, 0.2 U
mL−1 BmGDH and 0.3 mM NADP+ in a final volume of 1 mL.
Reactions were started with the addition of substrate (10 mM
of fatty alcohols or fatty acid or 200 µL of alkane) and incu-
bated at 30 °C for 0.5, 2, and 24 hours with shaking (200 rpm).
For cell-free extract (CFE) biotransformations cells expressing
CYP505A30 were resuspended (0.2 g wet weight cells mL−1) in
buffer D (200 mM potassium phosphate buffer pH 8, 100 mM
glucose) and disrupted as stated above, followed by centrifu-
gation (20 000g, 30 min, 4 °C). CFE (800 µL) (4 µM) was added
to the reaction mixture.
Reactions were stopped and extracted by the addition of
150 μL HCl (5 M), followed by 1 mL ethyl acetate containing
2 mM internal standard (1-dodecanol or 1-undecanol). Samples
were analysed by GC-FID (Shimadzu GC-2010) and GC-MS
(Thermo Scientific TraceGC ultra – Trace DSQ) using the
columns and temperature programs described in Table S3.†
For GC-FID analysis, fatty acids samples were methylated by
mixing equal volumes of ethyl acetate extracts (100 µL) and tri-
methylsulfonium hydroxide (TMSH) preparation.40 For GC-MS
analysis, samples were silylated. For silylation, 100 µL of the
ethyl acetate extracts from the biotransformations were dried
under N2 at room temperature. Equal amounts of pyridine and
N,O-bis(trimethylsilyl)acetamide containing 2% (w/v) tri-
methylchlorosilane were added to the dried samples, mixed
and incubated at 70 °C for 1 h and analyzed by GC-MS.
21 K. Sakai, F. Matsuzaki, L. Wise, Y. Sakai, S. Jindou,
H. Ichinose, N. Takaya, M. Kato, H. Warilshi and
M. Shimizu, Appl. Environ. Microbiol., 2018, 84, 1–15.
22 G. J. Baker, H. M. Girvan, S. Matthews, K. J. McLean,
M. Golovanova, T. N. Waltham, S. E. J. Rigby, D. R. Nelson,
R. T. Blankley and A. W. Munro, ACS Omega, 2017, 2, 4705–
4724.
Conflicts of interest
There are no conflicts to declare.
23 M. J. Maseme, A. Pennec, J. van Marwijk, D. J. Opperman
and M. S. Smit, Angew. Chem., 2020, 59, 10359–10362.
24 S. Uhlig, M. Busman, D. S. Shane, H. Rønning, F. Rise and
R. Proctor, J. Agric. Food Chem., 2012, 60, 10293–10301.
25 S. Igari, S. Mori and Y. Takikawa, Wear, 2000, 244, 180–184.
26 S. P. Arnaud, L. Wu, M. A. Wong Chang, J. W. Comerford,
T. J. Farmer, M. Schmid, F. Chang, Z. Li and M. Mascal,
Faraday Discuss., 2017, 202, 61–77.
Acknowledgements
The authors would like to thank Mr Sarel Marais for GC-MS
analysis.
27 E. M. Isin and F. P. Guengerich, Anal. Bioanal. Chem., 2008,
392, 1019–1030.
Notes and references
1 P. R. Ortiz de Montellano, Chem. Rev., 2010, 110, 932–948.
2 Y. Wang, D. Lan, R. Durrani and F. Hollmann, Curr. Opin.
Chem. Biol., 2017, 37, 1–9.
3 L. Hammerer, C. K. Winkler and W. Kroutil, Catal. Lett.,
2018, 148, 787–812.
28 K. P. Conner, C. Woods and W. M. Atkins, Arch. Biochem.
Biophys., 2011, 507, 56–65.
29 J. A. Riddick, W. B. Bunger and T. K. Sakano, Organic sol-
vents: physical properties and methods of purification, Wiley,
4th edn, 1986.
4 S. Eiben, L. Kaysser, S. Maurer, K. Kühnel, V. B. Urlacher 30 S. H. Yalkowsky, Y. He and P. Jain, Handbook of Aqueous
and R. D. Schmid, J. Biotechnol., 2006, 124, 662–669.
5 F. H. Arnold and J. C. Lewis, Chimia, 2009, 63, 309–312.
6 C. J. C. Whitehouse, S. G. Bell and L. L. Wong, Chem. Soc.
Rev., 2012, 41, 1218–1260.
7 K. G. Ravichandran, S. S. Boddupalli, C. A. Hasemann,
J. A. Peterson and J. Deisenhofer, Science, 1993, 261, 731–736.
Solubility Data: An Extensive Compilation of Aqueous
Solubility Data for Organic Compounds, CRC Press LLC,
Boca Raton, FL, 2nd edn, 2003.
31 T. Kitazume, A. Tanaka, N. Takaya, A. Nakamura,
S. Matsuyama, T. Suzuki and H. Shoun, Eur. J. Biochem.,
2002, 269, 2075–2082.
444 | Org. Biomol. Chem., 2021, 19, 439–445
This journal is © The Royal Society of Chemistry 2021

View Article Online
Organic & Biomolecular Chemistry
Paper
32 A. Pennec, C. L. Jacobs, D. J. Opperman and M. S. Smit, 36 E. Weber, A. Seifert, M. Antonovici, C. Geinitz, J. Pleiss and
Adv. Synth. Catal., 2015, 357, 118–130. V. B. Urlacher, Chem. Commun., 2011, 47, 944–946.
33 S. Kochius, J. van Marwijk, A. Ebrecht, D. Opperman and 37 D. J. Opperman and M. T. Reetz, ChemBioChem, 2010, 11,
M. Smit, Catalysts, 2018, 8, 531. 2589–2596.
34 A. Mohammadzadeh-K, R. E. Feeney, R. B. Samuels and 38 F. M. Ferroni, C. Tolmie, M. S. Smit and D. J. Opperman,
L. M. Smith, Biochim. Biophys. Acta, 1967, 147, 583–
PLoS One, 2016, 11, 1–12.
589.
39 L. K. Morlock, D. Böttcher and U. T. Bornscheuer, Appl.
Microbiol. Biotechnol., 2018, 102, 985–994.
40 W. Butte, J. Chromatogr. A, 1983, 261, 142–145.
35 A. Glieder, E. T. Farinas and F. H. Arnold, Nat. Biotechnol.,
2002, 20, 1135–1139.
This journal is © The Royal Society of Chemistry 2021
Org. Biomol. Chem., 2021, 19, 439–445 | 445