Exploring nicotinamide cofactor promiscuity in NAD(P)H-dependent flavin containing monooxygenases (FMOs) using natural variation within the phosphate binding loop. Structure and activity of FMOs from Cellvibrio sp. BR and Pseudomonas stutzeri NF13

Article abstract of DOI:10.1016/j.molcatb.2014.08.019

Flavin-containing monooxygenases (FMOs) catalyse asymmetric oxidation reactions that have potential for preparative organic synthesis, but most use the more expensive, phosphorylated nicotinamide cofactor NADPH to reduce FAD to FADH₂ prior to f

Full text of DOI:10.1016/j.molcatb.2014.08.019

Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Exploring nicotinamide cofactor promiscuity in NAD(P)H-dependent
flavin containing monooxygenases (FMOs) using natural variation
within the phosphate binding loop. Structure and activity of FMOs
from Cellvibrio sp. BR and Pseudomonas stutzeri NF13
Chantel N. Jensena, Sohail T. Alib, Michael J. Allenb, Gideon Grogana,∗
a York Structural Biology Laboratory, Department of Chemistry, University of York, Heslington, York YO10 5DD, UK
b Plymouth Marine Laboratory, Prospect Place, Plymouth PL1 3DH, UK
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 22 July 2014
Received in revised form 18 August 2014
Accepted 30 August 2014
Available online 11 September 2014
Flavin-containing monooxygenases (FMOs) catalyse asymmetric oxidation reactions that have poten-
tial for preparative organic synthesis, but most use the more expensive, phosphorylated nicotinamide
cofactor NADPH to reduce FAD to FADH2 prior to formation of the (hydro)peroxy intermediate required
for substrate oxygenation. A comparison of the structures of NADPH-dependent FMO from Methylophaga
aminisulfidivorans (mFMO) and SMFMO from Stenotrophomonas maltophilia, which is able to use both
NADPH and NADH, suggested that the promiscuity of the latter enzyme may be due in part to the
substitution of an Arg–Thr couple in the NADPH phosphate recognition site in mFMO, for a Gln–His
couple in SMFMO (Jensen et al., 2012, Chembiochem, 13, 872–878). Natural variation within the phos-
phate binding region, and its influence on nicotinamide cofactor promiscuity, was explored through the
cloning, expression, characterisation and structural studies of FMOs from Cellvibrio sp. BR (CFMO) and
Pseudomonas stutzeri NF13 (PSFMO), which possess Thr–Ser and Gln–Glu in the putative phosphate recog-
nition positions, respectively. CFMO and PSFMO displayed 5- and 1.5-fold greater activity, respectively,
than SMFMO for the reduction of FAD with NADH, and were also cofactor promiscuous, displaying a
ratio of activity with NADH:NADPH of 1.7:1 and 1:1.3, respectively. The structures of CFMO and PSFMO
revealed the context of the phosphate binding loop in each case, and also clarified the structure of the
mobile helix–loop–helix motif that appears to shield the FAD-binding pocket from bulk solvent in this
class of FMOs, a feature that was absent from the structure of SMFMO.
Keywords:
Flavoprotein
Monooxygenase
FAD
Biotransformations
Sulfide
© 2014 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND
license (http://creativecommons.org/licenses/by-nc-nd/3.0/).
1. Introduction
employ the phosphorylated nicotinamide cofactor NADPH to
reduce a molecule of FAD within the enzyme, which then reacts
Flavin-containing monooxygenases (FMOs) [1] catalyse the
oxygenation of heteroatoms, such as nitrogen and sulfur, in various
organic substrates, and have been studied both for their role in
metabolism in higher eukaryotes, including humans [2,3], and
also for their contributions to microbial metabolism, in which
they are able to catalyse the oxidation of amines [4] and amino
acids such as ornithine [5]. In the case of microbial enzymes, the
identification of interesting FMO activity has led to the exploitation
of those enzymes for biotechnological applications [6,7], in which
the enzymatic characteristics of high turnover rates and chemo-,
regio- and enantioselectivity, are very valuable. FMOs typically
with molecular oxygen to form a (hydro)peroxy flavin species that
is the catalytic oxidant in reaction [1]. A well-studied example
of a bacterial FMO is that from Methylophaga aminisulfidivorans
(Uniprot Q83XK4, mFMO), an enzyme of monomer molecular
weight 46 kDa that was identified on the basis of its ability to
form the pigment indigo through oxidative transformation of
indole [8]. mFMO has also been shown to catalyse the asymmetric
sulfoxidation of a series of prochiral thioethers, when employed
as part of a fusion enzyme with phosphite dehydrogenase for
the recycling of the nicotinamide cofactor [9]. Studies of mFMO
have shown
a dependence for the phosphorylated cofactor
NADPH, but the lower cost of the non-phosphorylated analogue,
NADH, has meant that recent studies of FMOs have been directed
towards enzymes that might employ that cofactor for flavin
reduction.
∗
Corresponding author. Tel.: +44 1904 328266; fax: +44 1904 328266.
E-mail address: gideon.grogan@york.ac.uk (G. Grogan).
http://dx.doi.org/10.1016/j.molcatb.2014.08.019
1381-1177/© 2014 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/3.0/).

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C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
expected size were isolated using a PCR Cleanup kit® (Qiagen). Tar-
get genes were then sub-cloned into the pET-YSBL-LIC-3C vector
following a previously published procedure [20]. The recombinant
plasmids were then used to transform cells of E. coli XL1-Blue
(Novagen), which, after transformation and overnight growth on LB
agar containing 30 ␮g mL−1 kanamycin as antibiotic marker, were
subjected to miniprep procedures that resulted in plasmids suit-
able for DNA sequencing. Once the sequence of the genes had been
confirmed, gene expression was conducted by transforming cells
of E. coli BL21 (DE3) with the recombinant plasmids. 5 mL of LB
medium containing 30 ␮g mL−1 kanamycin was inoculated with a
single colony of the relevant strain. This starter culture was grown
at 37 ◦C overnight with shaking at 180 r.p.m. Each 5 mL culture was
then used to incolulate 500 mL LB broth containing 30 ␮g mL−1
kanamycin in a 2 L Erlenmeyer flask. These larger cultures were
grown with shaking at 37 ◦C until the optical density, as deter-
mined by measurement at 600 nm, had reached 0.8. The cultures
were then induced through the addition of 1 mM isopropyl ␤-
D-1-thiogalactopyranoside (IPTG), and growth continued at 18 ◦C
overnight. Cells were then harvested by centrifugation for 15 min
at 4225 g using a Sorvall GS3 rotor in a Sorvall RC5B Plus centrifuge.
Following centrifugation, the resultant cell pellets were resus-
pended in 25 mL 50 mM Tris/HCl buffer pH 7.5, containing 300 mM
sodium chloride (‘buffer’) per L of cell growth. These suspensions
were then subjected to cell disruption using an ultrasonicator for
3 × 30 s periods at 4 ◦C with intervals of 1 min. The soluble fraction
after sonication was recovered by centrifuging the suspension for
30 min at 26,892 g in a Sorvall SS34 rotor. Supernatants were then
filtered using a 2 ␮m Amicon filter, and then subjected t nickel affin-
ity chromatography using a 5 mL His-TrapTM Chelating HP column.
After loading the filtered protein solution, the column was washed
with five column volumes of buffer containing imidazole (30 mM).
The FMOs were then eluted from the column using a 30–500 mM
imidazole gradient over twenty column volumes. Column fractions
containing FMOs were identified using SDS-PAGE and combined.
Pooled fractions were concentrated, typically, to a volume of 4 mL
using a Centricon® filter membrane (10 kDa cut-off) and 2 mL of
this solution then loaded onto an S75 SuperdexTM 16/60 size exclu-
sion column that had been pre-equilibrated with buffer. FMOs were
eluted with buffer at a flow rate of 1 mL min−1. Fractions were ana-
lysed by SDS-PAGE and those that contained pure FMOs pooled and
stored at 4 ◦C for crystallisation, enzyme assays or biotransforma-
tions. For the purposes of crystallisation, the histidine tags of CFMO
or PSFMO were cleaved using 3 C protease and using a procedure
described previously [20]. Typical CFMO and PSFMO preparations
yielded 20 mg and 7.5 mg pure protein per litre of cells, respectively.
We recently reported the cloning, expression and structural
characterisation of another FMO, named SMFMO, from the marine
bacterium Stenotrophomonas maltophilia [10]. This target was inter-
esting as it displayed the ability to use either NADPH or NADH as
the cofactor for reduction of the flavin. SMFMO was hence able
to use NADH, along with a formate dehydrogenase/sodium for-
mate based recycling system, to catalyse the asymmetric oxidation
of thioethers, and also the Baeyer–Villiger oxidation of a strained,
fused cyclobutanone substrate [10]. Related activities have recently
been described by the group of Fraaije, who have characterised
other FMOs, notably FMO-D, from Rhodococcus jostii RHA1, related
to SMFMO, and which are similarly able to employ NADPH or
NADH [11]. The structure of SMFMO was determined [10], and
analysis of the nicotinamide cofactor binding loop revealed dif-
ferences between NADPH-dependent mFMO [12–14] and SMFMO
that might be significant in the recognition of the NADPH 2ꢀ ribose
phosphate that distinguishes NADPH and NADH [10]. In this region
in mFMO, Arg234 and Thr235 project towards the phosphate and
interact directly with the phosphate oxygen atoms, whereas in
SMFMO Gln193 and His194 are found in equivalent positions.
A double mutant of SMFMO that was designed to mimic the
phosphate binding loop of mFMO, changed the preference of the
enzyme for NADPH to NADH from a ratio of 1.5:1 to 1:3.5 [15].
These mutations were not successful in removing activity with
NADH, however. Multiple studies on the wider group of NAD(P)H-
dependent flavoprotein monooxygenases (FPMOs) have shown
that, whilst positively charged basic residues are often involved in
the specific recognition of negatively-charged phosphate [16–18],
NADH-dependent activity can be engineered through the mutation
of the cofactor binding loop to include a negatively charged car-
boxylate side chain enzymes that excludes phosphate, presumably
through charge repulsion [19]. Engineering a glutamate residue
into the cofactor-binding loop of SMFMO, in an attempt to gen-
erate a more NADH-specific variant, resulted in a mutant that was
not produced in the soluble fraction of the Escherichia coli strain
used for gene expression, however [15]. In this report, we describe
the cloning, expression, and characterisation of two homologs of
SMFMO, CFMO from Cellvibrio sp. BR (Uniprot code I3IEE4) and
PSFMO from Pseudomonas stutzeri NF13 (M2V3J0). These homologs
display natural variation in the cofactor-binding loop, Thr–Ser in
CFMO and Gln–Glu in PSFMO, which suggested there may be altered
cofactor preference compared to either mFMO or SMFMO. The
enzyme activity with NADH and NADPH and a range of prochi-
ral sulfides is assessed, and the structures of the enzymes, which
reveal the context of the substituted amino acids within the puta-
tive cofactor binding loop, are presented.
2. Experimental
2.3. Enzyme assays
2.1. Chemicals
Steady-state kinetic constants for the NADH and NADPH-
dependent reduction of FAD by the FMOs were determined
using the method employed previously [10,21]. In a 1 mL quartz
cuvette containing Tris–HCl buffer pH 7.5 (50 ␮mol) the decrease
in absorbance at 340 nm was monitored for concentrations of
NAD(P)H (10–100 ␮M) after the addition of enzyme (CFMO or
PSFMO, 3.9 nmol). All data points represented the average of three
separate runs. Kinetic constants (KM and kcat) were calculated using
a value for ε of 6220 mol dm−3 cm−1 using GrafitTM (Erithacus
Sofware).
Chemicals, including media and buffer components, sulfide sub-
strates and cofactors were purchased from Sigma-Aldrich (Poole
U.K.).
2.2. Gene synthesis, cloning, expression and protein purification
The genes encoding CFMO and PSFMO were synthesised
by GeneArt (Invitrogen), with sequences optimised for expres-
sion in E. coli using the GeneArt server program. Genes were
then amplified by PCR from the commercial genes using the
following primers: For CFMO: Forward: CCAGGGACCAGCAATG-
GATACACCGGTTATGG; Reverse: GAGGAGAAGGCGCGTTAGGCGC-
TATCCAGATACTG; For PSFMO: Forward: CCAGGGACCAGCAATGC-
CTCCGATTCTGG; Reverse: GAGGAGAAGGCGCGTTACGGACGACG-
GCTCGG. PCRs were analysed on agarose gels, and bands of the
2.4. Biotransformations
Biotransformations using isolated enzymes with cofactor
recycling were performed using the method previously described
for SMFMO [10]. For NADH-dependent biotransformations: To

C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
193
Table 1
Data collection and resolution statistics for CFMO and PSFMO. Values for the highest resolution shells are given in parentheses.
CFMO
PSFMO
Beamline
Diamond i24
Diamond i24
Wavelength (Å)
Resolution (Å)
Space group
Unit cell
0.96862
22.44–2.39 (2.45–2.39)
C2
0.96862
35.94–1.83 (1.88–1.83)
P3221
a = b = 63.56; c = 189.82
a = 115.41; b = 95.09; c = 92.37
˛ = ˇ = 90.0; ꢀ = 126.3
˛ = ˇ = 90.0; ꢀ = 120.0
No. of molecules in the asymmetric unit
Unique reflections
Completeness (%)
Rmerge (%)
Rp. i. m.
Multiplicity
2
1
31344 (2329)
98.1 (98.4)
0.09 (0.50)
0.09 (0.50)
3.2 (2.9)
9.0 (2.1)
0.99 (0.76)
25
24.6/28.7
0.014
1.66
38
40
41893 (3176)
100 (99.9)
0.112 (0.62)
0.054 (0.30)
9.8 (9.9)
14.9 (3.8)
1.00 (0.90)
13
16.2/20.4
0.02
2.26
19
22
ꢁI/ꢁ(I)ꢂ
CC1/2
Overall B factor from Wilson plot (Å2)
Rcryst/Rfree (%)
r.m.s.d. 1–2 bonds (Å)
r.m.s.d. 1–3 bonds (◦)
Avge main chain B (Å2)
Avge side chain B (Å2)
Avge water B (Å2)
31
24
a 10 mL round bottomed flask containing Tris–HCl buffer pH
7.5 (5 mL) were added substrate(s) (1–6 in 100 ␮L ethanol to
a final concentration of 5 mM), NADH (5 mg, a final concentra-
tion of 0.7 mM), formate dehydrogenase (5 mg), sodium formate
(6.8 mg, 0.1 mmol) and CFMO or PSFMO (1 mL of a 5 mg mL−1
solution, 0.13 ␮mol). The reactions were then stirred for 24 h
at room temperature. Aliquots (500 ␮L) were taken at intervals
and extracted with ethyl acetate (500 ␮L). The organic layer was
transferred to a GC vial and analysed by GC as described previ-
ously [10]. For NADPH-dependent biotransformations: To a 10 mL
round bottomed flask containing Tris–HCl buffer pH 7.5 (5 mL)
were added substrate(s) (1–6 in 100 ␮L ethanol to a final con-
centration of 5 mM), NADPH (5.7 mg, a final concentration of
0.7 mM), glucose-6-phosphate-dehydrogenase (0.14 mg), glucose-
6-phosphate (5.2 mg, 0.02 mmol) and CFMO or PSFMO (1 mL of a
5 mg mL−1 solution, 0.13 ␮mol). The reactions were then stirred
for 24 h at room temperature and organic extracts of 500 ␮L
aliquots analysed as previously [10]. Chiral analysis of sulfoxide
products was carried out using BGB 173 and BGB-175 columns
(30 m × 0.25 mm × 0.25 ␮m; each from BGB-Analytik) according to
procedures described previously [10].
Rigaku Micromax-007HF generator fitted with Osmic multilayer
optics and a MARRESEARCH MAR345 imaging plate detector. Crys-
tals that diffracted to a resolution of greater than 3 A were retained
˚
for full dataset collection at the synchrotron.
2.6. Data collection, structure solution, model building and
refinement of CFMO and PSFMO
Datasets described herein were collected at the Diamond Light
Source, Didcot, Oxfordshire, U.K. Data for CFMO and PSFMO were
each collected on beamline I24. Data were processed and inte-
grated using XDS [23] and scaled using SCALA [24] included in the
Xia2 processing system [25]. Data collection statistics are given in
Table 1. The crystals of CFMO were in space group C2. The struc-
ture of CFMO was solved using MOLREP [26], using a monomer
model of SMFMO (PDB code 4a9w) [10]. The solution contained
two molecules in the asymmetric unit, representing one dimer,
and the solvent content was 55%. The crystals of PSFMO were in
space group P3221. The structure of PSFMO was again solved using
SMFMO as a model, but in this case, the solution contained only one
molecule in the asymmetric unit. The solvent content in this case
was 57%. The structures of CFMO and PSFMO were built and refined
using iterative cycles using Coot [27] and REFMAC [28], employing
local NCS restraints in refinement. The final structures exhibited
Rcryst and Rfree values of 24.6 and 28.7 (CFMO) and 16.2 and 20.4%
(PSFMO), respectively. Each structure was validated prior to depo-
sition using PROCHECK [29]. Refinement statistics for all structures
are presented in Table 1. The Ramachandran plot for CFMO showed
93.4% of residues to be situated in the most favoured regions, 6.2% in
additional allowed and 0.5% residues in outlier regions. For PSFMO,
the corresponding values were 92.8%, 4.6% and 2.6%, respectively.
The coordinates and structure factors for CFMO and PSFMO have
been deposited in the Protein Data Bank with the accession codes
4usq and 4usr, respectively.
2.5. Protein crystallisation
Pure PSFMO and CFMO was subjected to crystallisation trials
using a range of commercially available screens in 96-well plates
employing 300 nL drops at a range of protein concentrations (3,
10 and 20 mg mL−1). The best crystals for PSFMO were obtained
using the Clear Strategy Screen (CSS) [22] conditions containing
35% (w/v) tacsimate pH 7.0 and His6-tag cleaved protein at a con-
centration of 20 mg mL−1. The best crystals for CFMO were obtained
using the CSS conditions containing 1.5 M ammonium sulfate and
non-cleaved protein at 20 mg mL−1. Larger crystals for diffrac-
tion analysis using optimised conditions were prepared using the
hanging-drop vapour diffusion method in 24-well plate Linbro
dishes and using crystallisation drops of 2 ␮L, comprised of 1 ␮L of
reservoir solution and 1 ␮L of protein solution at 20 mg mL−1. For
PSFMO, the best crystals were again obtained in crystal drops con-
taining 35% (w/v) tacsimate at pH 7.0 with no further additions. For
CFMO, the best crystals were obtained in crystal drops containing
1.5 M ammonium sulfate and 1% propan-2-ol (v/v) at pH 7.0. Crys-
tals were flash-cooled in a cryogenic solution containing the mother
liquor with 10% (v/v) glycerol, and tested for diffraction using a
3. Results and discussion
3.1. Target selection
Following the failure to incorporate a glutamate residue within
the cofactor binding loop of SMFMO that might engender discrimi-
natory binding of NADH over NADP in that enzyme, it was decided
to select targets from the genomic databanks representative of

194
C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
Fig. 1. Sequence alignment of SMFMO, CFMO and PSFMO. The characteristic Rossman motifs are highlighted in blue; the FMO motif [30] in green, and the two residues
proposed to be the closest to the 2ꢀ ribose hydroxyl phosphate in red. (For interpretation of the references to color in this figure legend, the reader is referred to the web
version of this article.)
natural variation within that loop, particularly corresponding to
the Arg–Thr and Gln–His couples of mFMO of SMFMO, respectively.
It was hoped that these variants might be sufficiently different
within the loop to display a shift in preference for one nicotin-
amide cofactor, preferably NADH, over NADPH. CFMO (Uniprot
code I3IEE4) and PSFMO (M2V3J0) were selected on this basis. Each
is a putative FMO of a similar size to SMFMO (361 and 358 amino
acids, respectively, versus 357 for SMFMO) and contains two Ross-
man domains and the FXGXXXHXXXY FMO motif [30]. A sequence
alignment of these targets and SMFMO (Fig. 1) revealed 58% and
61% sequence identity between CFMO and SMFMO and PSFMO
and SMFMO, respectively. In place of the Arg234Thr235 couple in
mFMO or Gln193His194 couple in SMFMO, CFMO possessed Ser202
and Thr203, and PSFMO, Gln194 and Glu195, respectively. The lat-
ter was particularly interesting in being one of the only homologs
identified from database searches as having either a glutamate
or aspartate residue within the putative phosphate recognition
region. No homolog was identified yet that possessed a Glu or Asp
residue in place of Gln193 in SMFMO. Genes encoding CFMO and
PSFMO were synthesised, subcloned and expressed in the soluble
fractions of transformed strains of E. coli BL21 (DE3). The pro-
teins were readily purified using nickel affinity and size exclusion
chromatography as described in Section 2, each yielding protein
solutions of a bright yellow colour, indicative of the presence of
bound FAD.
shown in Table 2, with data for SMFMO [10] reproduced for
comparison.
The first observation was that both CFMO and PSFMO were both
able to accept NADH or NADPH for the reduction of FAD. CFMO was
observed to bind both NADH and NADPH with increased affinity
compared to SMFMO, as illustrated by Km values of 3 and 5 ␮M com-
pared to values in the range of 24–27 ␮M for NADH and NADPH,
respectively. Although kcat values were lower for CFMO than the
other two enzymes, the low Km contributed to CFMO having a cat-
alytic activity, as determined by values of kcat/KM, of 5.3 and 5.4
fold greater than SMFMO for NADH and NADPH, respectively. These
values also reveal a slight preference for NADH by a factor of 1.7,
compared with SMFMO, for which the value was 1.5. For PSFMO,
the KM was similar with either NADH (16 ␮M) or NADPH (17 ␮M)
which were each lower than for SMFMO. kcat/KM was slightly
lower with NADH (1880 M−1 s−1) than NADPH (2350 M−1 s−1),
indicating
a preference for NADPH in this case of 1.3, but
Table 2
Kinetic constants for CFMO and PSFMO in the reduction of flavin by either NADH or
NADPH. Values for SMFMO, reproduced from Ref. [10] are also given for comparison.
Enzyme/cofactor
KM (␮M)
kcat (s−1
)
kcat/KM (M−1 s−1
)
SMFMO*
NADH
SMFMO
NADPH
CFMO
24
27
3
9
5
0.03
1250
0.02
0.02
0.02
0.03
0.04
740
1
1
6670
4000
1880
2350
NADH
CFMO
3.2. Cofactor promiscuity in the reduction of FAD
5
NADPH
PSFMO
NADH
PSFMO
NADPH
The ability of CFMO and PSFMO to use either NADH or NADPH
to reduce FAD was assessed using standard UV spectrophoto-
metry assays [10,21], in which the oxidation of NAD(P)H was
monitored at 340 nm. Kinetic parameters (Km and kcat) were deter-
mined for each enzyme and each cofactor and the results are
16
2
17 2.0
*
Data taken from Ref. [10].

C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
195
NADPH, respectively. Overall, PSFMO catalysed sulfoxidation reac-
tions gave higher conversions than either SMFMO or CFMO, but
with poorer e.e.s overall, save for substrate 6.
3.4. Structures of CFMO and PSFMO
In order to shed light on the nature of the cofactor binding loops
in CFMO and PSFMO, the structures of each in complex with the
˚
˚
flavin FAD were determined to resolutions of 2.39 A and 1.83 A,
respectively. Crystals of CFMO grew in the C2 space group, with
two molecules ‘A’ and ‘B’, representing one dimer in the asymmet-
ric unit. The CFMO dimer was made up of two monomers (Fig. 3b),
2
˚
sharing an interfacial area of approximately 1207 A . Analysis of
the CFMO structure using PISA [31] found that the interactions
that stabilise the dimer included six hydrogen bonds, including
those between the backbone nitrogen of Trp108(A) and the side
chain oxygen of Glu124(B) and the side chain N H of Gln320(A)
with the backbone carbonyl of Ile145(B). The calculated dissoca-
tion energy of the dimer interface (ꢂiG) was −4.4 kcal mol−1. The
monomer of CFMO superposed with the structure of the SMFMO
Fig. 2. Sulfide substrates screened in this study against CFMO and PSFMO.
each value was again higher than that for SMFMO with either
cofactor.
˚
3.3. Biotransformations of prochiral sulfides using CFMO and
PSFMO using either NADH or NADPH as cofactor
monomer with an r.m.s.d. of 0.77 A over 317 C␣ atoms. The sec-
ondary elements of the structure are summarised in Fig. 3a. Each
CFMO monomer consists of two domains, an FAD binding domain
and the putative substrate binding domain. Electron density was
visible for the majority of the backbone in each monomer, from
residue Ser13 to Ala361, with a stretch of missing density corre-
sponding to six amino acids between positions Gln236 and Asp243
(PVGGLG) in subunit A and fifteen between Ala227 and Asp243
(AQEGREIEQPVGGLG) in subunit B that could not be modelled. Side
chain density in the region of residues 224–236 in subunit A was
also missing. It appears that the residues for which there is poor
electron density are part of a flexible helix–loop–helix structure,
incorporating helix ␣8, which is found over the FAD binding pocket,
perhaps shielding the active site from bulk solvent. An equiva-
lent region of density was missing in structures of both wild-type
SMFMO (4A9W) [10] and the Gln193Arg/His194Thr mutant (4C5O)
[15]. There was substantial residual density in the omit map in the
putative active site following building and refinement of the pro-
tein atoms of CFMO. This was successfully modelled and refined as
FAD in the flat, oxidised form.
Both CFMO and PSFMO were then challenged with a range of
prochiral sulfide substrates 1–6 (Fig. 2), with the addition of either
catalytic NADH or NADPH as cofactor, and in the presence of a
suitable cofactor recycling system, as had been used for SMFMO
[10]. The results of all biotransformations by CFMO and PSFMO,
compared to those obtained with SMFMO are shown in Table 3.
CFMO oxidised sulfides 1 and 3–6 to, for the most part, their corre-
sponding (R)-sulfoxides, with either cofactor. However, substrate
1 was transformed to the (S)-sulfoxide in the presence of NADPH.
The conversions were overall higher with NADH, however, the
enantiomeric excess of sulfoxide products was slightly higher with
NADPH in general. Substrate 2 was not converted by CFMO in
the presence of either cofactor. The highest conversion was seen
for substrate 3, methyl tolyl sulfide, with 65% for NADH (22% e.e.
(R)-sulfoxide product) and 32% for NADPH (32% e.e.). The most
enantioselective transformation was that of substrate 6 with 66%
-(R) and 77% -(R) for achieved with NADH or NADPH, respectively.
CFMO was overall less active than SMFMO with sulfide substrates
and NADH, although both the conversion and product enantiomeric
excess in the oxidation of phenyl methyl sulfide 4 were superior.
Interestingly, when NADPH was employed as nicotinamide cofactor
CFMO gave higher conversions compared to SMFMO, except for
substrate 3, possibly indicative of its greater activity as shown by
the kinetic constants. For PSFMO, the highest conversions were
observed for substrate 3 and 4, but enantioselectivity was poor in
each case. PSFMO again gave (R)-sulfoxide products for the most
part. The most enantioselective reaction was observed for substrate
6, yielding (R)-sulfoxide product of 85% and 57% with NADH and
Crystals of PSFMO grew in the P3221 space group, with one
molecule in the asymmetric unit, although both size-exclusion
studies (not shown), and the relationship of the monomer to its
closest crystallographic symmetry partner, are strongly suggestive
of an active dimer for PSFMO as with SMFMO and CFMO. Anal-
ysis of a dimer pair by PISA [31] revealed an interfacial area of
2
˚
955 A , with six H-bonds and 8 salt bridges formed between the
monomers. The calculated dissociation energy of the dimer inter-
face (ꢂiG) was −7.1 kcal mol−1. The secondary structural elements
are summarised in Fig. 4a and the monomer structure is shown in
Fig. 4b. In the case of PSFMO, there was electron density for each
Table 3
Results of biotransformations of prochiral sulfide substrates 1 and 3–6 by CFMO or PSFMO using either NADH or NADPH as nicotinamide cofactor. Values for SMFMO,
reproduced from Ref. [10] are also given for comparison.
Sulfide
NADH
NADPH
Conversion (%); absolute configuration; e.e.
Conversion (%); absolute configuration; e.e. (%)
SMFMO*
CFMO
PSFMO
SMFMO*
CFMO
PSFMO
1
3
4
5
6
27, (R)-, 71% e.e.
90, (R)-, 25% e.e.
8, (R)-, 21% e.e.
6, (S)-, 15% e.e.
40, (R)-, 80% e.e.
17, (R)-, 54% e.e.
65, (R)-, 22% e.e.
64, (R)-, 58% e.e.
No conversion
14, (R)-, 66% e.e.
61, (R)-, 30% e.e.
97, (R)-, 47% e.e.
99, (R)-, 14% e.e.
13, (S)-, 10% e.e.
50, (R)-, 85% e.e.
2, (R)-, 57% e.e.
33, (R)-, 44% e.e.
1, n.d.
<1, n.d.
9, (R)-, 82% e.e.
6, (S)-, 43% e.e.
32, (R)-, 32% e.e.
38, (R)-, 64% e.e.
11, (R)-, 3% e.e.
47, (R)-, 77% e.e.
28, (R)-, 24% e.e.
78, (R)-, 32% e.e.
73, (S)-, 4% e.e.
No conversion
22, (R)-, 57% e.e.
*
Data taken from Ref. [10].
n.d. = not determined.

196
C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
Fig. 3. (a) Primary structure of CFMO, with secondary structure assignment, created using DSSP [32,33] and represented using ALINE [34]. (b) Structure of dimer of CFMO, in
ribbon format, with subunit A in green and B in coral. One FAD molecule per monomer is shown in cylinder format with carbon atoms in grey. Helix ␣8, which forms part of
the flexible helix–loop–helix structure that may shield the active site from bulk solvent, is also highlighted. (For interpretation of the references to color in this figure legend,
the reader is referred to the web version of this article.)
amino acid in the monomer from residue Met1 to Pro354, which
allowed for the first time the observation of the entire backbone
of the flexible helix–loop–helix structure, represented by residues
207–234 in PSFMO, and incorporating helices ␣9 and ␣10, over FAD
that had been missing from SMFMO structures and only partially
present in CFMO, although side-chain density for residues 222–227
was poor. Electron density was also present for residues Gly-2, Pro-
1 and Ala 0 at the N-terminus, which represent the first part of the
linker to the His-tag in the protein produced using the YSBLIC-3C
construct. The monomer of PSFMO superposed with the structure
again clearly visible in the omit map after building and refinement
of the protein atoms, and was also modelled and refined success-
fully in the oxidised form, the density revealing no puckering of the
tricyclic ring system that might be indicative of the reduced flavin.
3.5. The FAD binding region in CFMO and PSFMO
In both CFMO and PSFMO, the tricyclic isoalloxazine ring of FAD
was bound at the domain interface in a cavity beneath the flexible
helix–loop–helix region of each enzyme (Fig. 5a and b). The oxygen
atoms of the pyrimidinedione ring form H-bonds with the backbone
˚
of the SMFMO with an r.m.s.d. of 0.90 A over 327 C␣ atoms. FAD was
Fig. 4. (a) Primary structure of PSFMO, with secondary structure assignment, created using DSSP [32,33] and represented using ALINE [34]. (b) Structure of PSFMO monomer,
in ribbon format. One FAD molecule per monomer is shown in cylinder format with carbon atoms in grey. Helix ␣9, which with helix ␣10 forms part of the flexible
helix–loop–helix structure over the active site, and for which electron density is continuous in the PSFMO structure, is also highlighted.

C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
197
Fig. 5. FAD environment within the active sites of (a) CFMO and (b) PSFMO. The peptide backbones are shown in with transparency levels of 30%. Amino acid side chains
within the active site are shown in cylinder format in coral for CFMO and green for PSFMO. The FAD molecules are shown in cylinder format with carbon atoms in grey,
surrounded by the Fo − Fc omit map in blue displayed at a level of 3ꢁ in each case, which was obtained through refinement in the absence of FAD. The maps illustrate the
˚
˚
superior quality of the resolution (1.83 A vs. 2.39 A) in the PSFMO structure.
N
Hs of CFMO Phe62 (PSFMO Phe53) and Leu340 (Leu322). In
revealed that the phosphate binding sites for NADPH, including
the 2ꢀ hydroxyl ribose phosphate that distinguishes NADPH from
NADH, were occupied by sulfate ions that had resulted from the
crystallisation conditions in lithium sulfate (Fig. 6a). This obser-
vation led to the hypothesis that substitution of arginine and
threonine residues in mFMO for glutamine and histidine in SMFMO
were one of the factors that determined cofactor promiscuity in
the latter enzyme. Having acquired the structure of CFMO and
PSFMO, these were now superimposed with the structure of mFMO
in order to gain insight into the changes in the cofactor binding
loops that may occur as a result of having threonine/serine and
glutamine/glutamate in these positions. In the case of CFMO the
side-chain oxygen atoms of Thr202 and Ser203 are both orientated
towards the putative phosphate binding site (Fig. 6b). The struc-
ture(s) of mFMO had previously demonstrated that threonine is
able to make an effective contact with one of the phosphate oxygen
atoms of NADPH [12–14].
The nucleotide binding site of PSFMO featured a glycerol
molecule, from the cryoprotectant, occupying approximately
the same position as the adenine ring of NADPH in mFMO, and
with the hydroxyl groups forming hydrogen bonds with the
side chain of Tyr140, the backbone NH of Gln194 and a water
molecule. Superimposition of the putative phosphate binding sites
of SMFMO and PSFMO revealed that the backbone C␣ atoms of
common with SMFMO, the aromatic side-chain of a phenylalanine
residue CFMO Phe 62 (PSFMO Phe53) is found beneath the pyrim-
idinedione ring in the place where, in mFMO, an asparagine residue
is found, which is thought to stabilise the oxygenating species in
catalysis. Mutation of this Asn to a serine residue in mFMO led to
removal of oxygenase activity [13], but in SMFMO, mutation of the
Phe residue to a valine resulted in a mutant of inverted enantiose-
lectivity for the transformation of para-tolyl methyl sulfide [15].
The similarity in the FAD environment in all three homologs is
reflected in the broadly similar substrate ranges and enantioselec-
tivities of the cofactor-promiscuous enzymes with (R)-selectivity
conserved for the most part towards substrates 1, 3, 4 and 6 with
NADH. The poorer performance of NADPH-dependent biotransfor-
mations by SMFMO, CFMO and PSFMO appears at variance with
their comparable abilities to reduce FAD, but may be due to more
effective participation of the NADH cofactor in stabilisation of the
active site during oxygenation, as has been observed for mFMO [12].
3.6. Cofactor binding loops in CFMO and PSFMO
While the structures of SMFMO [10,15] have not yet been deter-
mined in complex with a nicotinamide cofactor, superimposition
with the structure(s) of NADPH-complexes of mFMO (e.g. 2XLT)
Fig. 6. The putative NADP+ ribose 2ꢀ phosphate recognition site in (a) SMFMO, (b) CFMO and (c) PSFMO, each superimposed with the corresponding site in mFMO, derived
from PDB accession code 2XLT. The backbone and side chain and NADP+ carbon atoms for mFMO are shown in grey in each case. Glc = glycerol in (c). Equivalent features for
SMFMO, CFMO and PSFMO are shown in yellow, coral and green in (a), (b) and (c) respectively. (For interpretation of the references to color in this figure legend, the reader
is referred to the web version of this article.)

198
C.N. Jensen et al. / Journal of Molecular Catalysis B: Enzymatic 109 (2014) 191–198
residues thought to be responsible for cofactor promiscuity in
SMFMO, Gln193 and His194, are in the same position as those
in PSFMO, Gln194 and Glu195 (Fig. 6c). However, the side chains
of both Gln194 and Glu195 are observed to point away from the
putative phosphate binding site into the bulk solvent.
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No data on the activity with mFMO and NADH have been
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of the NADH ribose oxygen atoms in the presence of that cofactor,
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CFMO and PSFMO provide new examples of enzymes within the
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of enzymes, and also suggesting new avenues for the engineering
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Acknowledgements
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We are grateful to PML Applications Ltd. and to the Biotech-
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studentship award to C.N.J.
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