1. A single amino acid residue controls ROS production in the
respiratory Complex I from Escherichia coli
Juho Knuuti,1
Galina Belevich,1
Vivek Sharma,2
Dmitry A. Bloch1
* and Marina Verkhovskaya1
**
1
Helsinki Bioenergetics Group, Institute of
Biotechnology, University of Helsinki, PO Box 65
(Viikinkaari 1), FIN-00014, Helsinki, Finland.
2
Department of Physics, Tampere University of
Technology, P. O. Box 692, FIN-33101, Tampere,
Finland.
Summary
Reactive oxygen species (ROS) production by respira-
tory Complex I from Escherichia coli was studied in
bacterial membrane fragments and in the isolated and
purified enzyme, either solubilized or incorporated in
proteoliposomes. We found that the replacement of a
single amino acid residue in close proximity to the
nicotinamide adenine dinucleotide (NADH)-binding
catalytic site (E95 in the NuoF subunit) dramatically
increases the reactivity of Complex I towards dioxygen
(O2). In the E95Q variant short-chain ubiquinones
exhibit strong artificial one-electron reduction at the
catalytic site, also leading to a stronger increase in
ROS production. Two mechanisms can contribute to
the observed kinetic effects: (a) a change in the reac-
tivity of flavin mononucleotide (FMN) towards di-
oxygen at the catalytic site, and (b) a change in the
population of the ROS-generating state. We propose
the existence of two (closed and open) states of the
NAD+
-bound enzyme as one feature of the substrate-
binding site of Complex I. The analysis of the kinetic
model of ROS production allowed us to propose that
the population of Complex I with reduced FMN is
always low in the wild-type enzyme even at low
ambient redox potentials, minimizing the rate of reac-
tion with O2 in contrast to E95Q variant.
Introduction
Complex I [nicotinamide adenine dinucleotide (NADH):qui-
none oxidoreductase, type I] catalyses electron input to the
respiratory chain. It uses NADH as a two-electron donor
and transfers these electrons to quinone (most often ubi-
quinone) via flavin mono-nucleotide (FMN) and eight iron-
sulphur (FeS) clusters located in the hydrophilic domain of
the enzyme. Quinone reduction is coupled to translocation
of three to four protons across the inner mitochondrial
or bacterial cytoplasmic membrane (Wikström, 1984;
Hinkle et al., 1991; Galkin et al., 1999; 2006; Wikström and
Hummer, 2012). Complex I has also been suggested to be
one of the major sites of formation of superoxide by mito-
chondria (Turrens and Boveris, 1980; Galkin and Brandt,
2005; Kussmaul and Hirst, 2006), and has been therefore
linked to some neurodegenerative diseases and ageing
(Lin and Beal, 2006; Fukui and Moraes, 2008; Rhein et al.,
2009). Indeed, it was reported that Complex I is the major
site of superoxide formation in brain mitochondria under
normal conditions (Barja and Herrero, 1998), and that it
produces most of the reactive oxygen species (ROS) in
Parkinson’s disease and ageing (for a review see Turrens,
2003), although the relevance of superoxide formation
by Complex I under physiological conditions has been
questioned (Zoccarato et al., 2004; Grivennikova and
Vinogradov, 2006; Drechsel and Patel, 2010; Kareyeva
et al., 2012). The site of this ‘electron leak’ from Complex I
to O2 has been studied in different systems, ranging from
intact mitochondria (Treberg et al., 2011) to the isolated
enzyme (Kussmaul and Hirst, 2006; Ohnishi et al., 2010),
there are no definite conclusions as yet. Here, we have
studied ROS production by isolated and native membrane-
bound Complex I from Escherichia coli, with the aid of
site-specific mutations to alter vital electron transfer prop-
erties. Complex I contains non-covalently bound FMN,
which resides on the bottom of a cavity of the catalytic
subunit and is, therefore, accessible to oxygen from the
surroundings. FMN has a low midpoint redox potential
(Sled et al., 1993; Euro et al., 2008b), and has been pro-
posed to be the main site of O2
•−
generation in mitochon-
drial Complex I (Galkin and Brandt, 2005; Grivennikova
and Vinogradov, 2006; Kussmaul and Hirst, 2006).
However, the rates of ROS production by purified mito-
chondrial [16–22 nmol mg−1
min−1
(Galkin and Brandt,
2005; Kussmaul and Hirst, 2006; Kareyeva et al., 2012)],
and bacterial [25–50 nmol mg−1
min−1
(Esterhazy et al.,
2008; this article)] Complex I, are very slow compared
to the NADH oxidation rates. Furthermore, some other
NADH-dependent flavin-containing enzymes, such as
Accepted 3 October, 2013. For correspondence. *E-mail dmitry
.bloch@helsinki.fi; Tel. (+358) 9 191 59754; Fax (+358) 9 191 59920;
**E-mail marina.verkhovskaya@helsinki.fi; Tel. (+358) 9 191 59748;
Fax (+358) 9 191 59920.
Molecular Microbiology (2013) 90(6), 1190–1200 ■ doi:10.1111/mmi.12424
First published online 29 October 2013
© 2013 John Wiley & Sons Ltd

2. NADH/NADPH oxidases belonging to the Nox family (e.g.
MJ0649 from Methanocaldococcus jannaschii) produce
superoxide at much higher rates (60 μmol min−1
mg−1
),
which is c. 60% of the NADH oxidation rate (Case et al.,
2009). ROS production by mitochondria respiring on sub-
strates that reduce matrix NAD+
is also moderate (for a
review see Brand, 2010). This rate can be significantly
increased by blocking the electron output with a specific
Complex I inhibitor (e.g. Turrens and Boveris, 1980), but
such treatment raises the NADH/NAD+
ratio making it
possible that other NADH-dependent enzymes, such as
pyruvate or α-ketoglutarate dehydrogenase, could partici-
pate (Starkov et al., 2004). Under certain conditions mito-
chondrial dihydrolipoamide dehydrogenase was found to
be the major source of NADH-dependent ROS production
(Kareyeva et al., 2012).
In submitochondrial particles NADH-dependent O2
•−
generation was found to be only slightly increased
by rotenone (Vinogradov and Grivennikova, 2005;
Grivennikova and Vinogradov, 2006), and the isolated
enzyme showed no response (Galkin and Brandt, 2005;
Kussmaul and Hirst, 2006; Drose et al., 2009). The first
indication that reversed electron flow in mitochondria could
be a source of ROS was obtained by Hinkle et al. in 1967
(Hinkle et al., 1967). Later, it was reported that succinate or
glycerol-6-phosphate gives the highest rates of Complex
I-derived ROS production in mitochondria under condi-
tions of reversed electron transfer (see Brand, 2010 for
a review). This phenomenon has been explained by
the presence of two different O2
•−
-generating nucleotide
binding sites in Complex I, only one of which is active upon
forward electron transport (Vinogradov and Grivennikova,
2005), but this was not supported by the resolved structure
of Complex I (Sazanov and Hinchliffe, 2006). O2
•−
genera-
tion in the quinone binding site of Complex I was proposed
independently by Brand and Treberg (Lambert and Brand,
2004a,b; Treberg et al., 2011), but this suggestion was
disputed by Pryde and Hirst (Pryde and Hirst, 2011), who
concluded from experiments with submitochondrial parti-
cles that ROS production by Complex I is due to oxidation
of FMN by O2 during both forward and reversed electron
flow.
At any rate, it is well established that the rates of gen-
eration of O2
•−
by Complex I in mitochondria, submito-
chondrial particles, and purified Complex I are low relative
to the much faster rates of NADH oxidation by quinone. It
has been suggested that retaining NAD+
in the catalytic
site could diminish the accessibility of oxygen to FMN
(Kussmaul and Hirst, 2006), thus perhaps contributing to
the slow rates of ROS production. The resolved structure
of Complex I with the nucleotide bound [PBD entry 3IAS
(Berrisford and Sazanov, 2009)] indeed suggests that
NAD+
covers almost completely the part of the FMN
alloxazine ring that protrudes into the cavity. However, it is
difficult to predict to what extent such an effect might
prevent oxygen to react with reduced FMN.
The rate of ROS production by Complex I is expected to
be determined by the properties of the nucleotide binding
site and the bound FMN. The current study where a con-
served glutamate residue in the NuoF subunit (E95) is
mutated reveals a key element of the mechanism that
prevents massive electron leakage from the catalytic site
to dioxygen.
Results
Previously we have found that the replacement of the
invariant Glu95 with glutamine in the NADH- and FMN-
binding NuoF subunit results in moderate changes in
nucleotide affinity, but a significant upshift in the midpoint
redox potential of FMN (Euro et al., 2009a). The latter was
explained as an electrostatic effect on FMN due to replac-
ing the negatively charged carboxylate by the neutral
amide, since the side-chain of Glu95 resides in the nucleo-
tide binding cavity at a short distance from FMN. The
structure of the hydrophilic domain of Thermus thermophi-
lus Complex I (PDB entry 3IAM) reduced with NADH shows
that the latter is bound between E97, a counterpart of E95
in E. coli enzyme, and FMN. The distance between E97
and the nicotinamide ring of bound NADH is 3.2 Å. On the
other side E97 is located at the border of two subunits,
Nqo1 (NuoF) and Nqo2 (NuoE). The latter contains the
2Fe-2S cluster N1a, which lays 10 Å from E97 with Pro98
in between (Fig. 1).
An unexpected property of the E95Q variant of Complex
I is a strong increase of the rate of NADH-dependent
hydrogen peroxide production (Table 1). We compared
ROS production of wild-type enzyme, the E95Q variant,
and two other variants that lack quinone reductase activity.
In NuoCD R274A the electron transfer to ubiquinone was
blocked since the replacement of R274 by the neutral
alanine residue resulted in a strong decrease of the mid-
point redox potential of the last cluster in the intraprotein
FeS chain, N2 (Belevich et al., 2007), to such an extent that
it cannot be reduced by NADH. This cluster is the imme-
diate electron donor to ubiquinone. In the other variant,
NuoM E144A, the mutation was in the membrane subunit,
NuoM, quite distant from the quinone binding site (Euro
et al., 2008a). It is not yet known whether ubiquinone could
be reduced in this mutant, but the properties of the N2
cluster were unchanged. The rates of ROS production in
wild type and the NuoM E144A variant were rather similar,
and slightly lower in NuoCD R274A, in stark contrast to the
very high reactivity of the NuoF E95Q variant (Table 1). The
small decrease of ROS production rate in NuoCD R274A
can be explained by the lower content of Complex I in the
sample: normalization by hexaammineruthenium (III)
chloride (HAR)-reductase activity yields the same ratio of
ROS production by Complex I 1191
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

3. NADH-dependent HAR to oxygen reduction rate as for wild
type. This finding excludes any significant role of the N2
centre, or ubiquinone bound near that centre, in NADH-
dependent H2O2 generation by purified Complex I, since
both redox centres are reduced to a much smaller extent in
NuoCD R274A compared to wild type, yet with an insignifi-
cant change of ROS production velocity. No significant
difference in the rate of ROS production was observed
between solubilized Complex I and enzyme reconstituted
into liposomes (not shown).
ROS production by Complex I can be stimulated by
short-chain ubiquinones, insignificantly by decylubiqui-
none (DQ) and strongly by coenzyme Q1 (Q1) and mena-
dione (Table 2). This process is probably mediated by
formation of the respective semiquinone which interacts
with oxygen. It is known that short-chain ubiquinone ana-
logues, such as Q1, can be reduced by Complex I from an
artificial site resulting in O2
•−
generation (Cadenas et al.,
1977; Galkin and Brandt, 2005); the efficiency of this
process is increased with decrease of the quinone hydro-
phobicity (Cadenas et al., 1977). Most probably the site of
this artificial reaction is FMN (King et al., 2009). Menadi-
one is known to interact with flavin in many flavin-
containing enzymes that yields intensive ROS formation
(e.g. Matsuda et al., 2000). All in all, the data shown in
Table 2 indicates that FMN is the only significant site of
ROS production by Complex I under these conditions.
The dependence of the rate of ROS production on the
ambient redox potential calculated from the ratio [NADH]/
[NAD+
] is shown in Fig. 2. The experimental curve, which
is well reproducible and similar for wild type and the
NuoCD R274A and NuoM E144A variants, is less steep
and positively shifted (Fig. 2A) from a Nernstian n = 2
curve with Em = −350 mV, characteristic for FMN of E. coli
Complex I (Euro et al., 2008b). These data are in very
good agreement with that obtained on E. coli Complex I
by Esterhazy et al. (2008). In the mitochondrial enzyme
the rate of O2
•−
formation is half-maximal at −359 mV
(Kussmaul and Hirst, 2006). The titration of the rate of
ROS formation by the NuoF E95Q variant was steeper
Fig. 1. Disposition of E97 (counterpart of
E95 in E. coli Complex I) in the catalytic site
of Complex I from T. thermophilus (PDB
entry: 3IAM).
Table 1. ROS production by isolated Complex I does not correlate
with its NADH oxidase or ubiquinone reductase activities.
Variant
Complex I activity, μmol mg−1
min−1 Rate of NADH-
dependent
ROS production,
nmol mg−1
min−1
NADH:HAR
oxidoreductase
NADH:DQ
oxidoreductase
Wt 107.4 ± 7.0 26.9 ± 1.8 51.7 ± 3.5
NuoM E144A 109.7 ± 8.5 3.6 ± 0.5 55.6 ± 7.5
NuoCD R274A 72.4 ± 2.3 5.7 ± 0.3 44.1 ± 0.4
NuoF E95Q 24.4 ± 1.5 11.5 ± 1.5 749.4 ± 1.7
1192 J. Knuuti et al. ■
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

4. than that in the WT enzyme, and showed a clear positive
shift (Fig. 2B), which is in good agreement with the previ-
ously determined midpoint potentials of the two-electron
transition of FMN, −350 mV for wild type and −310 mV in
this mutant (Euro et al., 2009a). Assuming that both the
reduced and the half-reduced forms of FMN interact with
oxygen, it may be suggested that the deviation from a
two-electron Nernstian curve in the wild-type enzyme is
due to a somewhat higher semiflavin radical stability than
in the E95Q variant. Such an effect could explain the
change of shape of the titration curves, but not the strong
increase in the rate of ROS generation by the E95Q
variant.
The ubiquinone reductase activity of wild-type enzyme
is inhibited by rolliniastatin, the specific Complex I inhibitor
affecting the quinone binding site, by 90%, regardless of
whether DQ or Q1 are the acceptors, whereas the activity
of NuoF E95Q is inhibited by only 50% when the DQ is the
electron acceptor and almost not at all with Q1 (Fig. 3).
Since the NuoF E95 is quite distant (over 100 Å) from the
quinone binding site and can hardly affect its structure
directly, the observed relative insensitivity to rolliniastatin
indicates that one half of the DQ and almost all of the Q1
reductase activity are due to quinone reduction at another
artificial site, presumably FMN.
An increased capability of electron exchange in the
catalytic site of the NuoF E95Q variant is also revealed
by a very different reactivity to dithionite. The wild-type
enzyme is reduced slowly on a minutes timescale,
whereas the reduction of NuoF E95Q occurs much faster
(Fig. 4). The kinetics of optical changes at 450 nm upon
reduction, where the FMN contribution to the Complex I
spectrum is maximal, is shown in Fig. 4. The half times
are approximately 3 min and 10 s, for wild type and NuoF
E95Q respectively. Thus the acceleration of enzyme
reduction by the mutation is of the same order as the
acceleration of hydrogen peroxide production.
On the basis of our results we may conclude that the
catalytic site of Complex I becomes leaky with respect to
reactivity with O2 due to loss of the negative charge of
E95, although the structure of the cavity containing the
FMN and the nucleotide binding site was not significantly
perturbed, since the affinity for NADH was only slightly
changed (Euro et al., 2009a).
The fact that the rate of reduction of Complex I by
dithionite was considerably accelerated in the NuoF E95Q
Table 2. ROS production by isolated Complex I induced by short-chain ubiquinones.
Sample
ROS productiona
– DQ Q1 Menadione
Purified wt 1.00 ± 0.02 1.16 ± 0.01 9.28 ± 0.28 12.10 ± 0.31
Purified E95Q 15.98 ± 0.46 22.78 ± 0.04 95.46 ± 2.25 144.87 ± 1.17
Membranes wt 1.00 ± 0.14 1.07 ± 0.09 3.93 ± 0.02 5.55 ± 0.02
Membranes E95Qb
6.04 ± 0.90 5.98 ± 0.26 40.17 ± 1.47 59.07 ± 2.25
a. The initial rate of H2O2 production is normalized to that of wild type in the absence of added quinones, which are 51.7 ± 3.5 nmol mg−1
min−1
for purified enzyme and 2.49 ± 0.34 nmol mg−1
min−1
for native membrane-bound Complex I respectively.
b. Normalized to estimated 1.5 times lower content of mutated Complex I in native membranes. Estimation is based on the ratio of differences
between the specific quinone reductase activities of the membranes and purified proteins.
Fig. 2. Redox titration of the rate of H2O2
formation by purified Complex I. The
normalized rate is plotted against the
apparent ambient redox potential calculated
from the ratio [NADH]/[NAD+
].
A. Titration of wild-type Complex I, circles,
and mutants lacking ubiquinone reductase
activity, NuoCD R274A, squares, and NuoM
E144A, triangles.
B. Comparison of the redox titration of
wild-type Complex I, circles, and NuoF E95Q,
squares. Note that the absolute values for the
mutant are 15 times larger in comparison with
wild type.
ROS production by Complex I 1193
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

5. variant is consistent with the rate-limiting step being the
approach of the dithionite oxoanion to the active site. In
the wild-type enzyme this reaction is apparently limited by
the negative charge of the carboxylate group of E95.
However, this cannot explain the higher reactivity of the
E95Q variant with neutral short-chain ubiquinones.
Discussion
Here we report the ∼15-fold enhancement in the steady-
state rate of ROS production (vROS
) by the E95Q variant
of E. coli Complex I at low ambient redox potentials
(maintained by the NADH/NAD+
pair) as the result of the
increased electron leakage from the catalytic site. It comes
from the X-ray crystallography data that ROS formation is
very unlikely to occur at any of the FeS clusters due to their
shielding by the protein matrix; only FMN is partially
exposed to the solvent (PDB entry: 3IAS). We also assume
that the ubiquinone binding site does not participate in
ROS production, at least under the given experimental
conditions. Thus, our observations can only be accounted
for by changes in the properties of the active site caused by
the mutation.
In Complex I the flavin cofactor is partially buried at
the bottom of a deep cavity forming the catalytic site
(Sazanov and Hinchliffe, 2006). The side-chain of Glu95/
Glu97 (E. coli numbering, subunit NuoF, or T. thermophi-
lus numbering, subunit Nqo1) is located at the same
depth as flavin and around 6 Å (N5FMN-OE1E97) away from
the flavin π-system. The carboxylate and the flavin form a
binding pocket for the nucleotide, as shown in Fig. 1 (PDB
entry: 3IAM). It follows from the structure (see Supporting
Fig. 3. Relative insensitivity of ubiquinone
reductase activity of E95Q to rolliniastatin.
NADH oxidation was monitored in the
presence of DQ (A, B) and Q1 (C, D) by
purified wild-type (A, C) and E95Q (B, D)
Complexes I. NADH addition is indicated by
arrow. Rolliniastatin, 1 μM, was added at zero
time as indicated.
Fig. 4. Kinetics of wild-type and NuoF E95Q Complexes I reduction by dithionite. Set of redox spectra upon the reduction is shown for wild
type and E95Q variant. Kinetics of optical changes at 450 nm is characterized by τ1
2
≈ 3 min for wild type (circles) and τ1
2 ≈ 10 s for E95Q
(squares).
1194 J. Knuuti et al. ■
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

6. information, section 1) as well as from the effect of the
E95Q mutation on the Em of FMN (Euro et al., 2009a) that
the Glu95 side-chain is most probably in the anionic state.
In the nucleotide-bound state, the H-bonding between the
Glu95 and NAD+
is very unlikely; however, the electric
field of its charged carboxylate group is plausibly to affect
both redox potentials of the bound nucleotide and the
flavin, despite the polar nature of the solvent-exposed
nucleotide-binding cavity.
Here we do not consider oxygen interaction with the
FMN radical, despite the fact that the latter is known as a
powerful ROS generator in many chemical or enzymatic
systems (see, e.g. Massey, 1994; 1995; Massey and
Ghisla, 1974; Mattevi, 2006). NADH is a strict two-
electron donor so that any FMN radical would derive from
subsequent oxidation of the FMNH−
state. Thus the popu-
lation of the FMN radical state will be dependent on the
population of the FMNH−
state. Moreover, the participation
of the two-electron-reduced FMN in ROS production by
mitochondrial Complex I was shown experimentally
(Kussmaul and Hirst, 2006).
The mechanism of O2 interaction with the reduced flavin
is not known in Complex I. However, it is generally
accepted that the primary step in ROS generation in other
flavoproteins and chemical systems is the rate-limiting spin
conversion and charge transfer between FMNH−
and O2
resulting in the ‘caged’ radical pair formation followed by a
rebinding of O2
•−
at the C4a atom of the flavin with the
formation of a flavin hydroperoxide species, which can
further produce H2O2 (Massey, 1994; Mattevi, 2006). Pro-
duction of both O2
•−
and H2O2 was shown experimentally in
Complex I, although the ratio between these two species
varied and was also reported to be different in mitochon-
drial and bacterial enzymes (Esterhazy et al., 2008).
Hence, in this study, we do not discriminate between O2
•−
and H2O2 as both are considered ROS, the focus being
concentrated on the nature of the electron leakage from the
catalytic site.
Since ROS production is considered virtually irrevers-
ible, based on the very high quenching efficiency of the
detection system, its observed rate in Complex I at steady
state can be quantitatively expressed as
v kROS ROS
FMNH= × −
[ ] (1)
where kROS
is the rate constant of the reaction and
[FMNH−
] is the population of the reduced flavin. Given the
measured redox potentials of FMN in the nucleotide-free
WT and mutant enzymes [−350 mV and −310 mV respec-
tively (Euro et al., 2009a)], FMN should stay almost fully
reduced in both variants at Eh < −380 mV (ambient redox
potentials, typically provided by NADH). If it was indeed
the case, the E95Q mutant should have revealed much
higher reactivity of the reduced FMN towards dioxygen
(that is, a higher value of kROS
, e.g. due to the increased
accessibility of dioxygen to the reduced FMN) in order to
ensure the observed change in vROS
.
In Complex I direct interaction of O2 with FMN is blocked
at the flavin distal (opposite to the substrate-binding) side
due to the steric hindrance caused by the peptide back-
bone and side-chains of Gly183, Glu184 and Asn92
(T. thermophilus numbering, PDB entry: 3IAM); the flanks
of the flavin are also protected by the protein, except for its
redox-inactive dimethylbenzene ring and the N5 atom. The
latter was suggested as the site of O2
•−
formation in model
flavoproteins (McDonald et al., 2011). In Complex I it is
exposed to the cavity large enough to hold one or two H2O
molecules and possibly O2 or O2
•−
; however, the cavity itself
is isolated from the rest of the nucleotide-binding pocket by
Ser96, Glu97 and Tyr180 peptide backbone, Ser96 and
Asp94 side-chains (T. thermophilus numbering, PDB
entry: 3IAM), and by the C3 amide group of bound nicoti-
namide, preventing both O2 binding to, and O2
•−
escape
from the flavin N5 atom. Hence O2 can approach FMN only
from the proximal (NAD+
-binding) side, and it seems that
the presence of either NADH or NAD+
bound to FMN
prohibits the access of O2. Therefore, ROS production is
likely to occur either in the nucleotide-free enzyme (i.e.
after NAD+
dissociation) or in the situation, where NAD+
is
present in the binding pocket but does not form the π–π
stacked complex with FMNH−
(usually referred to as
charge–transfer complex, CTC, in many flavoproteins),
thereby allowing O2 to reach the proximal side of the flavin
plain. In the latter case, two conformational sub-states for
the NAD+
-bound enzyme could be proposed: a ‘closed’
state [(NAD+
-FR
)closed
] for the NAD+
-FR
CTC and an ‘open’
state [(NAD+
-FR
)open
], where such complex is not formed.
The latter possibility has indeed been discussed in the
literature for various flavoproteins (Gassner et al., 1996;
Palfey et al., 2001; Palfey and McDonald, 2010) and is
further supported by the observation that in the X-ray
crystal structure of Complex I (Berrisford and Sazanov,
2009) the planes of the flavin and nicotinamide rings are
not strictly parallel, indicating a possible mixture of several
sub-states. However, the E95Q replacement can hardly
affect the O2 accessibility to FMN in either case: in the
nucleotide-free catalytic site the distance between FMN
and Glu95 is large enough to accommodate an O2 mol-
ecule (> 5 Å); in (NAD+
-FR
)open
state the negatively charged
Glu95 can only attract NAD+
[in comparison with (NAD+
-
FR
)closed
state] providing more access of O2 to FMN and
more ROS production in WT.
The E95Q replacement could also affect the kROS
value
(Eq. 1) if NAD+
dissociation rate ( )koff
NAD+
became higher
for E95Q with respect to WT (in other words, NAD+
was
retained longer in the binding site of WT, attracted by
Glu95 and preventing the O2 access to FMN). The
decreased Km
NADH
, as well as the decreased Ki for NAD+
and ADP-ribose in the mutant with respect to wild type
ROS production by Complex I 1195
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

7. (Euro et al., 2009a) may argue in favour of this sugges-
tion. However, the changes in apparent affinities upon the
mutation [2 times for Km
NADH
, 10 μM in WT and 5 μM in
E95Q, and 2.4 times for Ki
NAD+
, 1 mM in WT and 0.39 mM
in E95Q respectively (Euro et al., 2009a)] are not large
enough to provide 15 times stronger binding of O2 in the
mutant.
Another possibility for the change in kROS
is the mutation-
specific change of the redox potential of the FMN-
bound dioxygen (see Supporting information, section 2).
However, the estimated effect ranging from 0 to −15 mV,
depending on the position of the bound O2, is too small to
account for the observed effect in full.
Therefore, we conclude that the change in reactivity of
the reduced flavin with respect to O2 can hardly explain
the observed mutation effects. Due to this we consider the
thermodynamic aspect of the reaction of Complex I with
oxygen. From the experimentally measured effect of the
E95Q mutation on the Em of FMN in the absence of
nucleotides [ΔEm = +40 mV (Euro et al., 2009a)], the esti-
mated value of the dielectric constant of the cavity (ε ≈ 53,
see Eq. S1), and the structural difference between the
nucleotide-free and nucleotide-bound enzymes, the inter-
action potential between Glu95 and FMNH−
was found,
δϕ1 = −35 meV (see Supporting information, section 2).
On the other hand, the interaction between the negatively
charged Glu95 and the positively charged, oxidized nico-
tinamide moiety of bound NAD+
is, by absolute value,
much stronger, δϕ2 = +59 meV. The sum δϕ1 + δϕ2 = +24
meV (+0.6 kcal mol−1
) indicates that in the WT enzyme the
carboxylate group of Glu95 stabilizes FMNH−
in the
nucleotide-bound state (NAD+
-FR
) relative to the same
state in the mutant. Similar conclusion was obtained from
the density functional theory (DFT) calculations using a
small model system comprising only of FMN and NAD+
or
NADH in the CTC configuration and the Glu95 residue
(see Supporting information, section 3, Tables S1 and S2,
and Fig. S1). Based on the electrostatic interaction ener-
gies calculated from the partial charges derived from DFT
calculations, the (NAD+
-FR
)closed
state is found more stable
in WT by 1.1 kcal mol−1
(at ε = 50). These results predict
the opposite effect of the mutation on ROS production.
Relative destabilization of the NAD+
-FR
state in the mutant
also means higher dissociation constant for NAD+
: a
change in 1.1 kcal mol−1
would raise KD
NAD+
in the mutant
for almost one order of magnitude. Keeping in mind the
∼ 40 meV stabilization of the nucleotide-free state FR
in
the mutant, the change in KD
NAD+
would be even larger,
which is very unlikely, given the Ki
NAD+
data (see above).
Therefore, the other explanation of the observed phenom-
enon should be suggested.
Finally, we analysed how the mutation affects the
steady-state probability, or life time, of transient enzyme
state(s) competent in ROS production ([FMNH−
] in Eq. 1).
The reaction scheme is shown in Fig. 5. After FMN reduc-
tion (step 3) there are two alternatives. In Cycle A, NAD+
dissociation (step 4) is followed by the O2 reaction with
state FR
(step 6). Note that Cycle A is realized only in the
situation when the FeS chain is reduced to a great extent,
so that the electron transfer from FMNH−
to the FeS chain
is no longer possible [step 5, kET2 0= , see for a review
(Verkhovskaya and Bloch, 2013)]. The other alternative is
Cycle B, where O2 reacts with the state NAD+
-FR
(step 7).
If NAD+
dissociation from FMNH−
is significantly slower
than ROS production ( )k koff
NAD ROS+
<< , then the contribu-
tion of Cycle A is negligible and Cycle B becomes pre-
dominant. Unlike Cycle A, Cycle B is independent of the
reduction level of the FeS chain. Although in various
NAD(P)H-dependent flavoenzymes NAD+
can leave the
binding site either before or after the oxidation of FMN to
Fig. 5. Kinetic scheme of the steady-state ROS production by Complex I. F and FR
stand for the nucleotide-free enzyme with oxidized and
two-electron-reduced FMN respectively; NAD+
-F and NAD+
-FR
are the enzyme with bound NAD+
; NADH-F is the enzyme with bound NADH.
Both ‘closed’ and ‘open’ states of NAD+
-FR
are shown. Redox state of the FeS cluster chain is not shown for simplicity. Highlighted are the
ROS-producing states (NAD+
-FR
)open
and FR
. Reaction steps are: NAD+
dissociation from oxidized enzyme (1), NADH binding to oxidized
enzyme (2), electron transfer from NADH to FMN (presumably through the hydride transfer within the charge–transfer complex, 3), NAD+
dissociation from the enzyme with reduced FMN (4), electron transfer from reduced FMN to the FeS cluster chain (5), interaction of reduced
FMN with dioxygen and ROS production (6, 7), respectively, as indicated by numbers. All reaction steps (1–5), except for the ROS production
(6, 7), are thermodynamically reversible; arrows show the preferred fluxes under typical conditions. Dotted line shows the pre-steady-state
reduction of the enzyme-bound FeS cluster chain by FR
. Dashed lines show the irreversible two-electron reduction of dioxygen.
1196 J. Knuuti et al. ■
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

8. a secondary substrate, there are examples in the litera-
ture where flavin remains reduced for a considerable time
(e.g. when the electron acceptors are not available) and
NAD+
stays attached to it as long as the flavin remains
reduced (Gassner et al., 1996), which should be due to
the electrostatic interactions between NAD+
and FMNH−
.
Seemingly contradictory with the latter proposal, the
experimentally measured, relatively high rate constant of
NAD+
dissociation [ koff
NAD
s
+
≈ −
103 1
(Verkhovskaya et al.,
2008)] was in fact determined under the conditions where
the FeS chain and FMN were not fully reduced; thus, two
different koff
NAD+
values should be considered, depending
on the reduction state of the flavin (Fig. 5, steps 1 and 4
respectively).
Noteworthy, Cycle B can be realized only when the
steady-state electron flux from NAD+
-FR
to O2 takes place.
In the absence of O2, the reaction will inevitably proceed
to the nucleotide-free, fully reduced state (FR
), despite its
accumulation is slow.
To quantitatively analyse the yield of the FMNH−
states,
the analytical solution was obtained for each cycle sepa-
rately (Cycle A, steps 2, 3, 4, and Cycle B, steps 1, 2, 3)
at equilibrium and without the electron leakage to O2
(kROS
= 0) (Fig. 6; see also: Supporting information,
section 4, Eqs S2–S4 and Figs S2 and S3). Cycle A does
not show any difference in the population of ROS-
production state (FR
) between WT and E95Q at low Eh
(< −380 mV). The equilibrium solution for Cycle B predicts
that the flavin can be reduced only to a certain, small
extent in the NADH-reduced enzyme even at low poten-
tials. Despite the enzyme is considered in redox equilib-
rium with NADH/NAD+
(given that the absolute ROS
production rate in Complex I, 0.5–1.0 s−1
in WT, is orders
of magnitude lower than the enzyme turnover rate,
kcat > 300 s−1
), in the absence of redox mediators the
sequence of the steps restricts the yield of FMNH−
: NADH
binding (Fig. 5, step 2) is followed by the electron redis-
tribution between NADH and FMN (step 3) according to
their Em; at the same time, the preceding NAD+
dissocia-
tion (step 1) does not contribute to repopulate FMNH−
.
When the electron transfer from NADH to FMN is uphill
( )/
ΔE E Em m
FMNH /FMN
m
NADH NAD
= − <
− +
0 , then [NAD+
-FR
] is low;
when it is downhill (ΔEm > 0), [NAD+
-FR
] is higher. Assum-
ing Em
NADH NAD
mV/ +
= −320 and the experimentally deter-
mined Em values for FMN, −350 (WT) and −310 (E95Q),
the former situation is realized in WT and the latter, in the
Fig. 6. Simulated Eh-dependences of the
equilibrium concentrations of intermediates in
Cycles A and B. The parameters used
are: Ntot
= [NADH] + [NAD+
] = 10−4
M,
KD
NADH
M= −
10 5
, KD
NAD
M
+
= −
10 5
,
Em
N
mV= −320 , Em
F
mV= −350 (WT),
Em
F
mV= −310 (E95Q). The ambient redox
potential is set by [NADH] and [NAD+
]:
E Eh m
N NADH
NAD
= − × +
30 lg
[ ]
[ ]
. All redox potentials
quoted, refer to the NHE scale. For the
respective kinetic schemes, see Fig. 5. For
further details, see Supporting information,
section 4. Note that for Cycle B, the
parameter set differs from that found by the
data fitting (see Fig. S5 and Table S3). The
parameter set used here is the one showing
the difference between the two cycles the
most (see Supporting information, section 4).
ROS production by Complex I 1197
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

9. mutant (Fig. 6, lower panel). By varying the KD values for
WT and E95Q, it was possible to obtain a good fit of the
vROS
data, given both experimental effects of the mutation:
the 15-fold increase in vROS
at low Eh and the ∼ 20 mV
upshift of the apparent midpoint potential (E 1
2) of the vROS
Eh-dependence (see Supporting information, sections 4
and 5, Eqs S5–S10, Figs S4 and S5, and Table S3).
As shown by the electrostatic and DFT calculations (see
above), the NAD+
-FR
state is relatively stable in WT as
compared with the mutant, which apparently contradicts
both the observations and the kinetic modelling. However,
the uncertainty in the position of nicotinamide in the ROS-
producing (NAD+
-FR
)open
state might affect the result of
such calculations. Since the nucleotide-binding cavity is
wide enough to accommodate NAD+
in a different position
as compared with the resolved structure (see Supporting
information, section 1), the electrostatic interaction
between Glu95, FMNH−
and NAD+
may vary strongly. The
lack of the expected mutation-caused change in the NAD+
affinity (Ki
NAD+
, see above) further supports this possibility.
We note that the equilibrium solution of Cycle B (Fig. 6,
lower panel) predicts a significant amount of the
nucleotide-bound enzyme state(s) at low Eh, which are
known to form CTC in many flavoproteins. However, CTC
can be difficult to detect due to strong variation in both
extinction coefficient and maximum wavelength; more-
over, both NADH-F and NAD+
-FR
CTC are often found to
be transient, short-lived states (see Massey and Ghisla,
1974; Gao and Ellis, 2007; Lin et al., 2012; and refer-
ences therein). No information about optical properties of
CTC in Complex I has been available yet. We were not
able to detect any signature of CTC in E. coli Complex I
under the ROS-producing conditions, either (data not
shown). Since in Complex I, the proposed ROS-producing
state (NAD+
-FR
)open
is not a CTC, it should not have an
absorption signature. On the other hand, the maximum
yield of [NAD+
-FR
] can be rather low in both WT and the
mutant in the NADH-non-saturating conditions and
depend on KD
NADH
(see Fig. S4 and Eq. S9).
An important implication of this analysis is the finding
that even low electron leakage from NAD+
-FR
results in
returning the enzyme to the non-ROS-producing state
(NADH-F), which accumulates. Therefore, the population
of the ROS producing states is always low in the WT
enzyme. We conclude that Complex I possesses an
inbuilt safety mechanism that prevents excessive reduc-
tion of FMN and consequently minimizes ROS production
at low ambient redox potentials regardless of the avail-
ability of the natural substrate, ubiquinone. This mecha-
nism is based on the stabilization of an intermediate state
(non-reactive with O2) of partially reduced enzyme, where
the FeS chain is either oxidized or reduced, but only to a
certain extent, and slow NAD+
release from the state with
reduced flavin.
Experimental procedures
Escherichia coli strains MWC215 (Δnhd) (Calhoun and
Gennis, 1993), GR70N (Green et al., 1984) (both have wild-
type Complex I) and variants NuoM E144A (Euro et al.,
2008a), NuoF E95Q (Euro et al., 2009a) and NuoCD R274A
(Belevich et al., 2007) were grown aerobically in LB medium
supplemented with appropriate antibiotics on demand at
37°C. Inverted cytoplasmic membrane vesicles were pre-
pared after cell wall removal by sonication as described pre-
viously (Belevich et al., 2007), and used the same day for
analysis. Enzyme extraction and further purification from iso-
lated membranes was done as previously described (Euro
et al., 2009b). The HAR and DQ reductase activities of the
membrane-bound and purified Complex I were measured as
in Belevich et al. (2011). Reconstitution of Complex I into
liposomes was performed as described in Verkhovskaya
et al. (2011). Protein concentration in the membrane prepa-
rations was determined by the BCA protein assay kit (Pierce)
using BSA as a standard. Absorbance at 280 nm was used to
quantify the purified protein (ε280 = 785160 M−1
cm−1
).
Complex I mediated NADH-dependent H2O2 production
was quantified by using 0.4 units ml−1
of horse-radish per-
oxidase (Sigma) catalysing oxidation of 10 μM Amplex
Red (Sigma) to resorufin (ε557–620 = 51.6 mM−1
cm−1
) at 30°C.
NADH was used for purified Complex I and nicotinamide
hypoxanthine dinucleotide (dNADH) for native membrane
vesicles of GR70N and its variants NuoF E95Q, NuoCD
R274A and NuoM E144A to avoid contribution of NADH:ubiq-
uinone reductase type II, which was completely removed
after purification. The MWC215 strain is a GR70N derivative
lacking the gene encoding this alternative NADH dehydroge-
nase; it was grown for high-yield wild-type enzyme purifica-
tion. H2O2 production was measured in 200 mM HEPES-KOH
pH 7.0 (RT) or 20 mM HEPES-KOH containing 360 mM man-
nitol pH 7.0 (RT). Initial ROS production rate was same in
both buffers, but the latter was chosen for the measurements
of Complex I proteoliposomes to preserve electrogenic prop-
erties of the enzyme. 0.005% n-dodecyl β-D-maltoside (DDM)
was included when solubilized enzyme was used, at con-
centrations of 0.6 μg ml-1
(1.0 nM) for E95Q variant and
6.1–7.8 μg ml−1
(11–15 nM) for other variants and WT respec-
tively. Media was supplemented with 1 μM rolliniastatin when
ROS production by membrane preparations was defined, at
total protein concentration of 45 μg ml−1
for E95Q variant and
90 μg ml-1
for other variants and WT respectively. H2O2 for-
mation by purified Complex I was titrated using the
NADH:NAD+
redox couple to set the ambient potential. The
nucleotides, NADH or dNADH and NAD+
were added to a
total concentration of 35 μM or (d)NADH concentration was
kept constant and NAD+
concentration was varied. The reac-
tion was initiated by Complex I addition and the rate, which
appeared non-linear, was quantified right after the enzyme
addition allowing short mixing and defining the maximal rate
as velocity without NAD+
addition. Background of H2O2 gen-
eration was determined in the presence of 525 units ml−1
catalase (Sigma).
Kinetic measurements were carried out using a high-
resolution CCD-array spectrometer (HR2000+, Ocean
Optics). The assay buffer comprised 50 mM HEPES-KOH,
pH 7.0, 15% glycerol and 0.005% DDM. Complex I was
added at a final concentration of 2.7–2.9 μM.
1198 J. Knuuti et al. ■
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200

10. Numerical simulations and the data fit were performed
using Matlab R2010 (MathWorks, Natick, MA). The data fit
was accomplished using a non-linear, constrained solver
‘fmincon’. A full description of the kinetic models is given in
Supporting information. The details of DFT calculations are
also described in Supporting information (section 3).
Acknowledgements
This work was supported by grants from Biocentrum Helsinki,
the Sigrid Jusélius Foundation and the Academy of Finland.
V.S. acknowledges postdoctoral researcher funding from the
Academy of Finland. We thank Prof. Mårten Wikström for
extensive discussion and his interest to this work. We thank
Eija Haasanen for purification of Complex I. We are also
thankful to the Center for Scientific Computing (CSC),
Finland, for providing computing time.
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Supporting information
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version of this article at the publisher’s web-site.
1200 J. Knuuti et al. ■
© 2013 John Wiley & Sons Ltd, Molecular Microbiology, 90, 1190–1200