Tài liệu Báo cáo khoa học: Structural and mechanistic aspects of flavoproteins: photosynthetic electron transfer from photosystem I to NADP+ doc

MINIREVIEW
Structural and mechanistic aspects of flavoproteins:
photosynthetic electron transfer from photosystem I
to NADP
+
Milagros Medina
Departamento de Bioquı
´
mica y Biologı
´
a Molecular y Celular and BFIF, Universidad de Zaragoza, Spain
Introduction
Many electron-transfer reactions in biological systems
depend on redox chains that involve flavoproteins [1].
In these chains, questions remain regarding not only
the mechanisms of electron transfer and hydride
transfer, but also the role that flavins might play in
these events. The primary function of photosystem I
(PSI) is to reduce NADP
+
to NADPH, which is then
used in the assimilation of CO
2
[2,3]. In plants, this
occurs via reduction of the soluble [2Fe–2S] ferre-
doxin (Fd) by PSI. Subsequent reduction of NADP
+
by Fd
rd
is catalysed by FAD-containing ferredoxin–
NADP

+
reductase (FNR) [4]. In most cyanobacteria,
and some algae under low iron conditions, flavo-
doxin (Fld) (an FMN flavoprotein), in particular
Keywords
electron transfer; ferredoxin; ferredoxin–
NADP
+
reductase; flavodoxin; hydride
transfer; NAD(P)
+
⁄ H; photosystem I;
protein–flavin complexes; protein–protein
and protein–ligand interaction; redox
potential regulation
Correspondence
M. Medina, Departamento de Bioquı
´
mica y
Biologı
´
a Molecular y Celular, Facultad de
Ciencias, Pedro Cerbuna 12, Universidad de
Zaragoza, 50009-Zaragoza, Spain
Fax: +34 976 762123
Tel: +34 976 762476
E-mail:
(Received 28 January 2009, revised 22
April 2009, accepted 4 May 2009)
doi:10.1111/j.1742-4658.2009.07122.x

This minireview covers the research carried out in recent years into differ-
ent aspects of the function of the flavoproteins involved in cyanobacterial
photosynthetic electron transfer from photosystem I to NADP
+
, flavodox-
in and ferredoxin–NADP
+
reductase. Interactions that stabilize protein–
flavin complexes and tailor the midpoint potentials in these proteins, as
well as many details of the binding and electron transfer to protein and
ligand partners, have been revealed. In addition to their role in photosyn-
thesis, flavodoxin and ferredoxin–NADP
+
reductase are ubiquitous fla-
voenzymes that deliver NAD(P)H or low midpoint potential one-electron
donors to redox-based metabolisms in plastids, mitochondria and bacteria.
They are also the basic prototypes for a large family of diflavin electron
transferases with common functional and structural properties. Under-
standing their mechanisms should enable greater comprehension of the
many physiological roles played by flavodoxin and ferredoxin–NADP
+
reductase, either free or as modules in multidomain proteins. Many aspects
of their biochemistry have been extensively characterized using a combina-
tion of site-directed mutagenesis, steady-state and transient kinetics, spec-
troscopy and X-ray crystallography. Despite these considerable advances,
various key features of the structural–function relationship are yet to be
explained in molecular terms. Better knowledge of these systems and their
particular properties may allow us to envisage several interesting applica-
tions of these proteins beyond their physiological functions.
Abbreviations

2¢P-AMP, 2¢-phospho-AMP portion of NADP
+
⁄ H; CTC, charge-transfer complex; Fd, ferredoxin; Fd
ox,
oxidized ferredoxin; Fd
rd,
reduced
ferredoxin; Fld, flavodoxin; Fld
hq,
hydroquinone flavodoxin; Fld
ox,
oxidized flavodoxin; Fld
sq,
semiquinone flavodoxin; FNR, ferredoxin–NADP
+
reductase; FNR
ox,
oxidized ferredoxin–NADP
+
reductase; FNR
sq,
semiquinone ferredoxin–NADP
+
reductase; NMN, nicotinamide
mononucleotide portion of NAD(P)
+
⁄ H; PSI, photosystem I.
3942 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
Fld
sq

⁄ Fld
hq
, substitutes for the Fd
ox
⁄ Fd
rd
pair in this
reaction [5,6].
PSI
rd
þ Fld
sq
! PSI + Fld
hq
NADP
þ
þ 2Fld
hq
¡
FNR
NADPH + 2Fld
sq
Two Fld
sq
molecules transfer two electrons from
two PSI molecules to one FNR. FNR becomes fully
reduced through formation of the intermediate,
FNR
sq
, and later transfers both electrons simulta-

neously to NADP
+
[7,8]. During this process, the Fld
molecule must move between its docking site in PSI
and the docking site in FNR, and the formation of
short-life transient complexes is required. Mutational
and structural studies that characterize such interac-
tions are the subject of this minireview. Studies on
proteins from the cyanobacterium Anabaena are pref-
erentially considered because it is the most thoroughly
investigated system containing both Fld and FNR
(hereafter, AnFld and AnFNR) [5,8].
PSI architecture
Cyanobacterial PSI exists as either a monomer or tri-
mer embedded in the thylacoid membrane. It contains
12 subunits, 96 chlorophylls, 22 carotenoids, 2 phyllo-
quinones, 4 lipids and 3 [4Fe–4S] clusters per mono-
mer [9,10]. Protein subunits with more relevant
functions are conserved between plant and cyanobacte-
ria [11]. When light strikes one of the antenna chloro-
phylls and the exciton is transferred to the pair of
chlorophylls in the PSI reaction centre, charge separa-
tion occurs. Low-potential electrons are transferred
across the membrane by a chain that ends in three
[4Fe–4S] clusters, F
X
,F
A
and F
B

.F
X
is coordinated
by cysteines located in both of the large PSI subunits,
PsaA and PsaB, via a loop that also plays a role in the
attachment of PsaC [12]. PsaC, PsaD and PsaE are
located at the cytosolic site (Fig. 1A) [2,7,13–16]. PsaC
carries the terminal F
A
and F
B
clusters. After binding
of the protein carrier to this PSI site, the electron is
A
F
B
F
A
R39
(PsaE)
K106
(PsaD)
K34
(PsaC)
I59
W57
N58
Y94
D146
D90

C
K2, K3 T56, W57, N58
Y94
E61, D65
E67
E16
E20
T12
D150
D144, E145
I59
I92
D96, N97
B
R264
K75
L76
L78
E301
K72
R16
Y303
Fig. 1. (A) Molecular surface with the electrostatic potential of the putative Fd ⁄ Fld-binding site of Synechococcus elongatus PSI (PDB code
1jb0) [10]. The surface is transparent to show the internal position of the F
A
and F
B
centres (represented as spheres) in the PsaC subunit of
PSI (S. elongatus numbering is used). (B) Molecular surface with the electrostatic potential of Anabaena FNR at the Fld-docking site (PDB
code 1que) [62]. The FAD group is drawn in CPK with carbons shown in orange. PSI and FNR positions for the interaction with the protein

carrier are indicated. (C) Molecular surface with electrostatic potential of Anabaena Fld (PDB code 1flv) [28]. Detail of residues in the close
FMN environment in the oxidized Fld O-down conformation is shown on the right. FMN is drawn in CPK (balls or sticks), with carbons
shown in orange. Figure 1(A,B) is reproduced from the supplementary material in Gon˜i et al. [48] .
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3943
transferred from F
B
to Fld (or Fd), which subse-
quently leaves the PSI site bringing the electron to
FNR.
Flavins: key cofactors in protein
electron transfer reactions
Free flavins stabilize very little of their one-electron
reduced form, the semiquinone, because the midpoint
potential for reduction of the oxidized state to the
semiquinone, E
ox ⁄ sq
, is more negative than that for
reduction of the semiquinone to the hydroquinone,
E
sq ⁄ hq
[17]. Binding of FAD or FMN to the apopro-
tein usually displaces E
ox ⁄ sq
to a less negative value,
whereas E
sq ⁄ hq
shifts to a more negative value, stabiliz-
ing the semiquinone [1,18,19]. This allows flavoproteins
to function as key intermediates at the interface

between one- and two-electron transfers [20,21] and
allows flavoproteins to participate in many biological
processes [19,21,22].
Flavodoxin
The redox activity of Fld derives from its FMN cofac-
tor. The Fld semiquinone is exceptionally stable and its
midpoint potentials are quite negative. This is a direct
consequence of the differing stability of the oxidized,
semiquinone and reduced ApoFld:FMN complexes
[23–25]. In AnFld, the values are E
ox ⁄ sq
= )266 mV
and E
sq ⁄ hq
= )439 mV at pH 8.0 and 25 °C, with a
maximum stabilization of the semiquinone of  96%
[25,26]. Therefore, Fld is proposed to replace Fd
(E
ox ⁄ rd
= )384 mV) by exchanging electrons between
its semiquinone and hydroquinone states [5,8].
Oxidized Flds fold into a five-stranded parallel
b sheet sandwiched between five a helices [27–31], with
the FMN group located at the edge of the globular
protein and its two isoalloxazine methyls solvent acces-
sible (Fig. 1C). The polypeptide chain in the loops sur-
rounding amino acid residue 50 and amino acid
residue 90 (50’s and 90’s loops) make close contact
with the isoalloxazine and modulate its reduction
properties [24–26,32–37]. The H-bond network

observed in the AnFld FMN environment is conserved
in Flds across species [38], but specific interactions
around the flavin vary [39–41]. The 90’s loop usually
provides a Tyr stacked against the FMN si-face (Y94
in AnFld) which makes a large contribution to the
midpoint potential [23–25]. The residue from the 50’s
loop that stacks at the re-inner face is commonly a
Trp (W57 in AnFld) [20,28,29,39,42,43], but nonaro-
matic residues (L, H, M or A) have also been found
[29,40,44,45]. Both residues ensure that the flavin is in
an electronegative environment which allows tight
FMN
hq
binding while making formation of its anion
thermodynamically unfavourable [24,25].
In several Flds, rearrangement of the peptide bond
equivalent to 58–59 in AnFld allows a main chain car-
bonyl to flip from an ‘O-down’ conformation to an
‘O-up’ conformation. In the ‘O-up’ conformation, an
H-bond occurs between this carbonyl and N(5)H from
the neutral semiquinone [46,47]. In Anacystis nidulans
Fld, the flip involves breaking a weak H-bond present
in the oxidized state between the FMN N(5) and the
NH of V59, in favour of a stronger H-bond between
the carbonyl of N58 (‘O-up’ conformation) and FMN
N(5)H [41]. The semiquinone states of A. nidulans and
Anabaena Flds are less stable than those from other
species because the semiquinone H-bond with the CO
of Asn is weaker than the bond formed with the sma-
ller Gly, and because of the presence in the oxidized

state of a N(5)–HN59 H-bond that is absent in other
Flds. Replacements at T56, W57, N58, I59 and E61
in AnFld regulate the ability of the N58–I59 peptide
to H-bond with the N(5) or N(5)H, and modulate the
energy of its conformational change [25,26,48]. There-
fore, the backbone rearrangements of N58–I59
provide a versatile device for modulating the strength
of FMN binding and E
ox ⁄ sq
and E
sq ⁄ hq
in AnFld
[25,26,48].
Fld has a large excess of acidic residues which pro-
duce a strong dipole that orients its negative end
towards the FMN isoalloxazine. The importance of
electrostatic repulsion in the control of E
ox ⁄ sq
, and
particularly of E
sq ⁄ hq
, has also been demonstrated with
several Flds [25,37,41,48–53]. Electron nuclear double
resonance and 1D and 2D electron spin echo envelope
modulation spectroscopies applied to AnFld
sq
also led
to assignment of the interaction parameters of N(1),
N(3), H(5), H(6), CH3(8) and N(10) with the electron
spin [54,55]. Analysis of mutants indicated that the

stacking of a bulky residue at the re-face of the flavin
decreases the electron-spin density in the benzene ring,
whereas an aromatic residue at the si-face increases the
spin density at N(5) and C(6) [56].
Ferredoxin–NADP
+
reductase
The first structure obtained for a photosynthetic
FNR was from spinach (spFNR). spFNR folds in
two domains, one of which presents a noncovalently
bound FAD molecule and the other binds NADP
+
[57,58]. Structures from other species have also been
reported [59–62]. The FAD-binding domain in
AnFNR includes residues 1–138 and is made up of
six antiparallel b strands arranged in two perpendicu-
Flavoproteins in photosynthetic electron transfer M. Medina
3944 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
lar b sheets, with a short a helix at the bottom and
another a helix and a long loop that is maintained by
a small two-stranded antiparallel b sheet at the top
(Fig. 1B). The NADP
+
-binding domain includes resi-
dues 139–303 and consists of a core of five parallel
b strands surrounded by seven a helices [62]. FAD is
bound outside the antiparallel b barrel, and its isoal-
loxazine lies between two tyrosines, Y79 and the C-
terminal Y303 in AnFNR.
The two one-electron midpoint potentials of the fla-

vin in FNRs are close to each other, therefore these
proteins stabilize only 10–20% of the maximal amount
of semiquinone [63,64]. An E
ox ⁄ hq
value of )325 mV
has been reported for recombinant AnFNR at pH 8.0
and 10 °C [63,65]. Despite differences in buffer, tem-
perature and pH, the reported values for AnFNR are
in good agreement, however, slightly more negative
midpoint potentials are reported for other species
[64,66,67]. Replacements for Y79 have been reported
for the pea and spinach enzymes, Y89 and Y95,
respectively, suggesting that the aromaticity of this res-
idue is essential for FAD binding and that H-bonding
through the Tyr-OH is involved in the correct posi-
tioning of the NADP
+
substrate for efficient catalysis
[61,68]. Replacement of Y303 with Ser and of E301
(situated at the active site) with Ala shift the flavin
midpoint potential to considerably less negative values,
although semiquinone stabilization is severely ham-
pered, introducing constraints into one-electron trans-
fer processes [26,63]. In addition, L76, L78 and
particularly K75 at the FAD-domain modulate E
ox ⁄ hq
[63,65]. FNR:Fd complexation correlates with the
AnFd midpoint potential becoming 15 mV more nega-
tive and that of AnFNR becoming 27–40 mV less neg-
ative [69]. As seen with the spinach proteins,

complexation makes electron transfer thermodynami-
cally more favourable [70]. Similarly, the midpoint
potential for reduction of NADP
+
in complex with
FNR is 40 mV less negative than that of the free
NADP
+
⁄ NADPH pair [66,71]. Assignment of hyper-
fine couplings to nuclei of the isoalloxazine semi-
quinone have also been reported for AnFNR
sq
and
pFNR
sq
[72]. These studies indicated that the net effect
of the C-terminal Tyr is withdrawal of electron density
from the benzene ring towards the pyrazine ring, plac-
ing the accepted electron nearer to a site where it can
best be neutralized by protonation – the N5 position.
Electron transfer from PSI to
flavodoxin
Fd and Fld differ in size and in the chemical nature of
their redox cofactors. There is no sequence homology
between them, but structural alignment based on their
surface electrostatic potentials shows cofactor superpo-
sition in the region where both proteins accumulate
the negative end of their molecular dipole moments
[73]. Their biding site on PSI was analysed by studying
the kinetic behaviour of site-directed mutants and by

electron microscopy on cross-linked complexes
[12,15,16,74,75]. The cytosolic subunits of Synechococ-
cus elongatus (Sy) PSI, PsaC, PsaD and PsaE, and the
extrinsic loop of PsaA, present a positively charged
surface potential (Fig. 1A) and are proposed to partici-
pate in electrostatic docking of the negatively charged
Fd or Fld (Fig. 1C) [13,16]. The PsaC subunit cannot
be deleted without loss of PSI activity because it car-
ries the F
B
donor [14]. PsaD contributes to the electro-
static steering of Fd toward its binding site [76,77],
whereas several roles are proposed for PsaE [78,79].
K35 from the PsaC subunit of Chlamydomonas rein-
hardtii is critical for the interaction and, therefore, effi-
cient electron transfer [15,80]. The residues of PSI and
Fd facing each other have not yet been identified, with
the exception of K106 in SyPsaD, which interacts with
E93 in SyFd (E95 in AnFd) [77,81]. The scarce data
about the interaction of Fld and PSI suggest similar
functions for PsaC, PsaD and PsaE, but there are
insufficient data to propose a Fld docking site
[13,16,82,83]. Analysis of different AnFd and SyFd
mutants revealed that E31, R42, T48, D67, D68, D69,
E94, and particularly D59, D62 and E95 (AnFd num-
bering) influence electron transfer and are involved in
either the binding process or electron transfer itself
[84–86].
Although the Fld
sq

⁄ Fld
hq
pair is involved in shut-
tling electrons between PSI and FNR, a physiological
role for the Fld
ox
⁄ Fld
sq
pair cannot be precluded
[7,14]. Reduction of AnFld
ox
to the semiquinone state
by PSI has been a useful model with which to ana-
lyse the interaction forces and electron transfer
parameters involved in the physiological reaction [48].
Wild-type AnFld forms a transient PSI:Fld
ox
complex
prior to electron transfer [87]. Site-directed mutagene-
sis has been used to find the role of specific AnFld
side chains in the interaction and electron transfer
with PSI [53,84,88–90]. Many of these Fld mutants
(T12V, E16Q, T56G, W57 replaced by K, R, F, L, A
and Y, I59 replaced by A and K, Y94 replaced by A
and F, N97K, I59A ⁄ I92A and I59E ⁄ I92E) accept
electrons from PSI following transient complex for-
mation [53,87,90,91]. For some (T12V, W57Y and
Y94F), k
et
was lowered considerably, suggesting that

the complex is not optimal for electron transfer.
Changes in the midpoint potential are proposed to be
responsible for the W57Y Fld behaviour [90], but
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3945
changes in the electrostatics within the FMN environ-
ment which favour a complex less efficient for elec-
tron transfer explain the results for the other mutants
[88]. By contrast, k
et
was enhanced considerably for
most of the remaining mutants. More or less negative
values of E
ox ⁄ sq
for these Flds compared with wild-
type Fld might influence this kinetic behaviour by
decreasing or increasing, respectively, the driving
force of the reaction. In addition, analysis of multiple
charge reversal Fld mutants has recently indicated
that changes in the orientation and magnitude of the
molecular dipole moment have a critical effect on
electron transfer [48].
However, for some Fld mutants (E20K, T56S,
W57E, N58C, N58K, I59E, E61A, E61K, I92 replaced
by A, E and K and D96N and also some multiple
charge reversal mutants), electron transfer from PSI to
Fld
ox
takes place via a collisional-type mechanism
[48,53,91]. Noticeably, some mutants show rates higher

than those obtained for the wild-type in all the
PSI ⁄ Fld ratios assayed [88]. These effects are not
related to a change in Fld midpoint potentials. They
might be interpreted as being caused by a modification
in the accessibility of the flavin, but more generally
can be explained by a conformational change in the
orientation of the interacting PSI and Fld surfaces,
leading to a smaller edge-to-edge distance between F
B
and the flavin ring [48]. However, for other mutations,
rates at a given concentration are lower than the corre-
sponding rate for wild-type Fld. Because, in general,
the introduced mutations are not in the direct isoall-
oxazine coordination, this is unlikely to be because of
differences in the structural FMN environment, but
rather because the orientation between the protein
dipoles is not optimal for electron transfer or because
of a change in the electrostatic potential of the protein
[48].
In conclusion, subtle changes in the isoalloxazine
environment influence Fld binding ability and modu-
late the electron-exchange process by producing differ-
ent orientations and distances between redox centres.
Observations indicate that these side chains contribute
to the orientation of AnFld on the PSI, producing a
wild-type complex that is not the most optimal for
electron transfer. Mutation of these residues changes
Fld surface topology, and the module and orientation
of the molecular dipole, contributing to their altered
behaviour. Mutational studies on I59 and I92 AnFld

indicate that their hydrophobicity is far from critical,
suggesting that either hydrophobic interactions do not
play a crucial role or that the hydrophobic surface of
Fld must be provided by the solvent-exposed portion
of FMN.
Electron flow from flavodoxin to
NADP
+
mediated by FNR
Interaction and electron transfer between Fld
and FNR
Crystal structures of Fd:FNR complexes have been
reported for Anabaena [92] and maize leaf [93]. Despite
the two structures exhibiting a different orientation for
Fd [94], the [2Fe–2S] cluster lies close to the FAD of
FNR in both. Mutants of the two partners have also
contributed to the identification of residues essential
for complex formation and electron transfer [5,65,95–
102]. Both electrostatic and hydrophobic interactions
play an important role in the association and dissocia-
tion processes in these complexes [5,8,92]. K72 (K88 in
spFNR), K75, L76, L78 and V136 in AnFNR are key
for the interaction with Fd [5,97,100,103]. Similarly,
residues on the AnFd surface have a moderate effect
on complex stability and electron transfer with the
reductase, with E94, F65 and S47 being crucial
[69,104–106]. Despite proposals that FNR interacts
with Fld and Fd using the same region [100,101], in
general, replacing some FNR residues had more dras-
tic effects in processes involving Fld, suggesting that

the individual residues do not contribute equally to
complex formation with both partners. This was
the case for R16, K72, and particularly K75
[65,89,100,107]. In addition, K138 and R264 in the
NADP
+
-binding domain of AnFNR are more impor-
tant in establishing interactions with AnFld than with
AnFd [100,108]. Moreover, although removal of the
E139 AnFNR negative charge has a deleterious effect
on electron transfer reactions with AnFd, it appears to
enhance electron transfer with AnFld [109]. Electron
transfer with Fld is severely diminished upon the intro-
duction of negatively charged side chains at L76, L78
and V136 in AnFNR [89]. Therefore, these nonpolar
residues participate in the establishment of interactions
with both AnFld and AnFd. With this in mind, it was
expected that one or more negatively charged or
hydrophobic residues on the Fld surface would interact
with some of the above specified residues on FNR.
A number of AnFld variants containing replace-
ments, either at the putative interaction surface with
FNR or in the FMN environment, have been analy-
sed. None of the E16, E20, T56, I59, E61, D65, I92,
Y94, D96 and N97 positions is key, but they do con-
tribute cooperatively to the orientation and strengthen-
ing of the FNR:Fld complexes [53,88]. Simultaneous
replacement of I59 and I92 indicated that they are not
involved in crucial specific interactions [53,89]. T12,
W57 and N58 seem to be more important in the inter-

Flavoproteins in photosynthetic electron transfer M. Medina
3946 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
action [53,88–90] and, in addition, FMN might be crit-
ical [53]. It is somehow unclear whether the residues
stacking the FMN ring, W57 and Y94, truly affect
protein binding, or if the altered electron transfer
properties in mutations be explained in terms of
altered flavin accessibility and ⁄ or thermodynamic
parameters [24,25,90]. Therefore, electron transfer pro-
cesses between FNR and Fld resulted in the modula-
tion (either negatively or positively) of some mutants,
but in no case was electron transfer prevented. Consid-
ering also that the crystallographic structure of the
AnFNR:AnFld interaction appears highly elusive, these
observations suggest that the interaction of Fld with
FNR is less specific than that of Fd.
Theoretical models of this interaction, one based on
the rat NADPH–cytochrome P450 reductase (CPR)
structure [95] and the other obtained by docking [110],
confirm that a mutual orientation between FNR and
Fld, similar to the corresponding binding domains in
CPR, is highly probable and places the redox centres
closer than observed in the FNR:Fd structures
[92,93,95]. The docking model fits well with the experi-
mental data, showing that all Fld residues important
for the interaction with FNR are in contact with the
FAD cofactor. Different interface propensities for the
same FNR residues, with either Fd or Fld, are consis-
tent with experimental observations indicating that,
although FNR uses the same site for interaction with

Fd and Fld, each individual residue does not partici-
pate to the same extent in interactions with each of the
partners [100]. This is in agreement with the fact that,
although multiple chemical modifications produced
Flds less suitable for electron transfer [111], site-direc-
ted mutagenesis has not revealed any residues critical
for the interaction with FNR [53,88–90]. Replacement
of the few Fld positions, T12, W57, N58 and Y94,
with a high interface propensity produced opposing
effects: some Fld:FNR complexes can be either weaker
or stronger and less optimal for electron transfer than
those with wild-type Fld, but others can appear more
optimal for a particular electron transfer process.
Docking suggests that wild-type Fld could adopt
different orientations on the FNR surface without sig-
nificantly altering the distance between the methyl
groups of FAD and FMN (Fig. 2A). This might
explain why subtle changes in the Fld still produce
functional complexes. Moreover, the enhanced or
hindered reactivity can also be explained if there is a
single orientation of Fld in the complex that is
retained and changes either the overall interaction or
the electron transfer parameters. Recent analysis of
multiple charge-reversal mutations on the Fld surface
concluded that interactions do not rely on a precise
complementary surface in the reacting molecules. In
E301
L263
R264
Y79

S80
BA
Fig. 2. Proposed interactions in the Anabaena FNR active site leading to electron transfer ⁄ hydride transfer. (A) Model of a ternary
Fld:FNR:NADP
+
complex. FNR is shown as a grey surface with the atoms of Y303 CPK coloured, with C shown in violet. The position of
NADP
+
on the FNR surface corresponds to the X-ray FNR:NADP
+
complex (PDB code 1gjr), in which Y303 prevents stacking of the flavin
and nicotinamide rings. The figure also shows several positions determined by docking of Fld onto FNR (Fld in green, light orange and pink
correspond to docking solutions ranked 1, 3 and 5 respectively). (B) Detail of the proposed FNR active site centre in a model of ternary com-
plexes competent for hydride transfer. FNR active site residues are given as sticks and CPK coloured, with C in grey. The nicotinamide por-
tion of the coenzyme presents the position derived from the structure in complex with Y303S FNR (PDB code 2bsa) [117]. The dotted
surface around the nicotinamide indicates the position of Y303 in wild-type FNR. For both structures, FAD in FNR, FMN in Fld and NADP
+
are show as sticks and are CPK coloured, with carbons in yellow, orange and pink, respectively.
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3947
fact, analysis indicates that the initial orientation, dri-
ven by alignment of the Fld molecule dipole moment
with that of FNR, contributes to the formation of sev-
eral alternative binding modes competent for the effi-
cient electron transfer reaction [48]. Similar behaviours
have been reported in other electron transfer systems,
where dynamic ensembles, as opposed to single confor-
mations, contribute to the electron transfer process
[112,113].
Electrostatic nonspecific interactions as major

determinants of the efficient interaction between
Fld and its counterparts
Some mutations appear to favour single orientations
which improve the association and electron transfer
with a particular partner and native Fld complexes are
not the most optimal for electron transfer. Such obser-
vation, in agreement with docking analysis [110], sug-
gests that the flavin atoms might be mainly involved in
the interaction and be solely responsible for electron
transfer. Therefore, subtle changes in the isoalloxazine
environment not only influence Fld-binding abilities,
but also modulate the electron transfer process by pro-
ducing different orientations and distances between the
redox centres. This further confirms that Fld interacts
with different structural partners through nonspecific
interactions, which in turn decrease the potential effi-
ciency that could be achieved if unique and more
favourable orientations were produced with a reduced
number of partners. During Fld-dependent photosyn-
thetic electron transfer, the Fld molecule must move
from its docking site in PSI to that in FNR. In vivo,
the formation of transient complexes of Fld with PSI
and FNR is useful, but not critical, during this process
to promote electron transfer and avoid the reduction
of oxygen by the donor centres [7,8,14,48,53]. Thus,
electrostatic alignment appears to be one of the major
determinants of the orientation of Fld on the partner
surface. The fact that simultaneous replacement on the
Fld surface did not hinder or enhance processes with
PSI and FNR also suggests a different interaction

mode with each partner [48].
Interaction of FNR with the NADP
+
coenzyme
and the hydride transfer event
Once the FAD cofactor of FNR has accepted two
electrons, they have to be transferred to NADP
+
. The
FNR protein portion has a dual role in this process
by: (a) modulating the FAD midpoint potential to a
value that makes the hydride transfer reversible, and
(b) providing the environment for an efficient encoun-
ter between the N5 of the flavin and the C4 of the nic-
otinamide. FNR is highly specific for NADP
+
⁄ H
versus NAD
+
⁄ H and different studies have established
a role for several FNR residues in determining coen-
zyme binding, specificity and enzymatic efficiency
[114–120]. Three FNR regions appear to be mainly
responsible for the interaction: 2¢-phospho-AMP (2¢P-
AMP) and pyrophosphate of the NADP
+
⁄ H binding
sites, and the position occupied by the C-terminal resi-
due where the nicotinamide portion of NADP
+

(NMN) is proposed to bind for hydride transfer [114–
117,119]. S223 and Y235 at the AnFNR 2¢P-AMP site
are critical in determining the specificity and efficient
coenzyme orientation [99,115,121]. The 155–160 and
261–268 loops, which accommodate the coenzyme
pyrophosphate portion, also confer specificity and the
volume of residues in the latter loop fine-tunes FNR
catalytic efficiency [114,116,119,120]. R100 (K166 in
spFNR), situated at the FAD-binding domain, allows
its guanidinium group to H-bond to the NADP
+
pyrophosphate, providing the necessary flexibility to
address the NMN moiety of NADP
+
towards the
active site [99,108,114]. Finally, the Tyr at the si -face
contributes to the correct positioning of the substrate
NADP
+
[68], whereas the C-terminal Tyr at the
re-face is surely critical for modulating NADP
+
⁄ H
biding affinity and selectivity [117,118,122–126].
Structural studies have allowed us to postulate a
stepwise mechanism in which the nucleotide must bind
to a bipartite site [59,62,114,117,127]. The first stage is
recognition of the 2¢P-AMP moiety [62]. The interme-
diate state represents a narrowing of the cavity to
match the adenine and the pyrophosphate, whereas the

nicotinamide is placed in a pocket near the FAD [114].
However, in this arrangement, the C-terminal Tyr pre-
vents interaction of nicotinamide with the isoalloxazine
and its energetically unfavourable displacement is then
expected if the hydride transfer optimal interaction is
to be achieved (Fig. 2A) [59,114,118,127]. Only FNR
variants in which the C-terminal Tyr has been replaced
produced structures with a rearrangement between fla-
vin and nicotinamide that was compatible with hydride
transfer, for example Y303S in AnFNR, and Y308W
or Y308S in pFNR (Fig. 2B) [59,117]. These FNRs
improved the affinity for NADP
+
and produced a
close interaction between flavin and nicotinamide
[117,118]. However, because of this strong binding,
they show low catalytic efficiency [59,117,118]. All the
data indicate a fine-tuning of the FNR efficiency pro-
duced by minor structural changes in the regions
involved in coenzyme binding [117,119].
Reduction of NADP
+
by FNR
hq
occurs by a
formal hydride transfer from the flavin anionic hydro-
Flavoproteins in photosynthetic electron transfer M. Medina
3948 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
quinone to the nicotinamide. In vitro, the reaction
takes place via a two-step mechanism, in which the

first observed process is related to formation of the
FNR
hq
–NADP
+
charge-transfer complex (CTC-2) via
an intermediate Michaelis–Menten complex (MC-2),
followed by hydride transfer to produce an equilibrium
mixture of the CTC-2 and FNR
ox
–NADPH (CTC-1)
CTCs. Both CTCs are also detected for the reverse
reaction, although the mechanism has some differ-
ences. Spectroscopic properties for these CTCs and
their hydride transfer rates for interconversion have
been estimated [66,121]. In AnFNR, CTC-2 accumu-
lates during the reaction and at equilibrium, whereas
CTC-1 evolves rapidly into other FNR states. Forma-
tion of these CTCs appears necessary for efficient
hydride transfer and the relative conformation and ori-
entation of FNR and NADP
+
⁄ H during the interac-
tion are critical [121]. Hydride transfer in systems
involving flavins and pyridine nucleotides is highly
dependent on the approach and colinear orientation
of the N5 of the flavin, the hydride to be transferred,
and C4 of the nicotinamide. In FNR, displacement
of the C-terminal Tyr appears to be required for the
interaction to occur [117,118]. The data reported to

date suggest good agreement between CTC formation
and hydride transfer rates [96,121]. However, this
hypothesis is based on a limited data set and preli-
minary characterization of the process for Y303S
AnFNR suggests that, at least for this mutant, there
might not be a direct correlation [128]. Therefore, it
may be that for wild-type FNR the most favourable
orientation between the nicotinamide and the flavin
might present considerably less overlap of the rings
than in the mutant structure (Fig. 2B), and other
relative orientations that maintain C4, H and N5
colinearity might account for the efficient hydride
transfer. Further work is needed to clarify these
points.
Most data indicate a similar interaction in higher
plant and cyanobacterial FNRs, but NADP
+
disor-
ders their spectra differently [71]. The different spec-
tra of higher plant FNRs show a peak at  510 nm
indicative of a stacking interaction between the nico-
tinamide and the isoalloxazine, which is not seen in
cyanobacterial FNRs [71]. Spectra obtained upon
addition of NADP
+
to C-terminal Tyr mutants pro-
duce prominent peaks in AnFNR and pFNR, sug-
gesting greater nicotinamide occupancy of the active
site [117]. Thus, although the AnFNR UV spectra
and electron spin density distribution of AnFNR

sq
are perturbed by NADP
+
[55,72], lower nicotinamide
occupancy of the active site is expected in AnFNR
relative to higher plant FNRs. Therefore, differences
in nicotinamide binding to the active sites cannot be
discounted.
FNR catalytic site
The structure of the catalytically competent
FNR:NADP
+
conformation indicates that in AnFNR,
S80, C261, E301 and Y303 constitute the FNR cata-
lytic site [59,117]. Y303 plays distinct and complemen-
tary roles during the catalytic cycle by lowering the
affinity for NADP
+
⁄ H to levels compatible with turn-
over, by stabilizing the flavin semiquinone required for
electron splitting and by modulating the electron trans-
fer rates [59,117,118]. Moreover, a role in providing
adequate orientation between the reacting rings might
be envisaged [128]. S80 and C261 contribute to the
efficient flavin:nicotinamide interaction through the
production of CTCs during hydride transfer (in
spFNR S96 and C272) [96,129]. This Ser also contrib-
utes to semiquinone stabilization, and the volume of
the Cys residue modulates the enzyme catalytic effi-
ciency [119]. E301 has been studied in AnFNR and

spFNR (E312) [98,130]. Structural and functional dif-
ferences were found when the same mutants were pro-
duced in both species, again suggesting differences in
their mechanisms [131]. E301 was more critical for sta-
bilization of the semiquinone and midpoint potential
in AnFNR [63,98,130]. Studies in spFNR concluded
that E301 does not act as a proton donor [98], but
whether it transfers protons in AnFNR could not be
determined [130]. In fact, in the AnFd:AnFNR and
AnFld:AnFNR dockings, the carboxylic group of E301
is no longer exposed to solvent and it is one of the res-
idues with highest propensity for being at the interface
[110]. Similar observations have been extracted from
the Fd:FNR crystal structure [92]. This suggests a pos-
sible pathway for proton transfer between the external
medium and the AnFNR isoalloxazine N5 via S80
[58,62]. In addition, in both enzymes, this residue is
critical for proper binding of the nicotinamide to the
active centre, CTC stabilization and efficient flavin
reduction by NADPH [98,130].
FNR catalytic cycle: the ternary complex
The ability of FNR, Fd and NADP
+
to form a ter-
nary complex is fully accepted, indicating that
NADP
+
is able to occupy a site on FNR without dis-
placing Fd [70,71,114,127], and similar behaviour also
applies for Fld [132]. During catalysis, the order in

which substrates are added is not important, although
Fd and Fld lower the affinity for NADP
+
and occupa-
tion of the NADP
+
-binding site weakens the Fd:FNR
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3949
and Fld:FNR complexes [70,132]. The two binding
sites are not completely independent, and the overall
reaction is proposed to work in an ordered two-sub-
strate process, with the pyridine nucleotide binding
first [70,133]. Complex formation between Fd
rd
and
FNR:NADP
+
was found to increase the rate of elec-
tron transfer by facilitating the rate-limiting step of the
process – dissociation of the product (Fd
ox
) [127].
Thus, in the system involving Fd, negative cooperativi-
ty in the ternary interaction is translated into positive
cooperativity at the kinetic level. However, some key
features of the process remain to be explained in
molecular terms because the expected molecular move-
ments are not apparent and, although reversibile, dif-
ferent mechanisms seem to apply in each direction

[127]. Binding equilibrium and steady-state studies in
wild-type and mutant proteins envisage similar mecha-
nisms for Fld [5,90,130,132]. Fld, reduced to Fld
hq
by
PSI, will cycle between Fld
hq
and Fld
sq
upon passing a
single electron to FNR [5]. Fast kinetic methods have
been used to analyse binding and electron transfer
between FNR and Fld, but to date the reactions have
been followed at single wavelengths and have not
involved NADP
+
⁄ H [90]. Therefore, it remains to the
future to better evaluate the intermediate and final spe-
cies of the equilibrium mixture in the electron transfer
process in the binary Fld:FNR and ternary
Fld:FNR:NADP
+
electron transfer systems.
Flds and FNRs in nonphotosynthetic
organisms and as building blocks for
more complex proteins
Flds are electron transfer proteins involved in a variety
of photosynthetic and nonphotosynthetic reactions in
bacteria [134]. In eukaryotes, only a few Flds have
been reported [135–137], but a descendant of the Fld

gene helps to build multidomain proteins [134,138]. A
photosynthetic function was first proposed for FNR,
but flavoproteins with FNR activity have been
described in chloroplasts, phototropic and heterotro-
phic bacteria, apicoplasts, and animal and yeast mito-
chondria [139]. Two unrelated families of proteins can
be found in these enzymes: the plant type and the glu-
tathione reductase type [126]. Based on their structural
and functional properties, plant-type FNRs are classi-
fied as plastidic type and bacterial type. Plastidic
FNRs efficiently catalyse electron transfer and hydride
transfer between low-potential one-electron carriers
and NADP
+
⁄ H, usually participating in the produc-
tion of NADPH. Bacterial FNRs generally exhibit
considerably slower turnover, provide the cell with
reduced electron carriers and are examples of novel
methods of FAD and NADP
+
⁄ H binding. However,
their structures, the particular residues involved in
FAD binding and the residues at the catalytic centre
are well conserved [91]. In addition, all plant-type
FNRs may share a similar catalytic mechanism [140].
The general fold found in FNR is also present in
other enzymes. Many of these enzymes are multidomain
proteins that, in addition to the FNR-like domain, also
contain Fd- or Fld-like domains. These proteins contain
FAD (or FMN) and a FMN or Fe–S protein, and shut-

tle electrons from NAD(P)H to the metal centres via
their FNR and Fd ⁄ Fld domains [138,141–144]. The Fld
and FNR domains in diflavin reductases appear to have
evolved independently [141,142,144]. Despite most
charged residues in Anabaena proteins being conserved
in these domains, the dipole moment orientations
between the FNR and Fld domains are far from colin-
ear [48]. Long-range electrostatic forces to attract their
interaction surfaces have been decreased. However, resi-
dues on the Fld- and FNR-domain interaction surfaces
may have been conserved to orientate the Fld domain
when pivoting between the FNR domain and the
electron acceptor [144].
The FNR family also contains NAD
+
⁄ H- and
NADP
+
⁄ H-dependent members. Some NAD
+
⁄
H-dependent members do not present the C-terminal
aromatic residue stacking the flavin and a cavity
appears open at its re-face [138,145,146], but the
NMN moiety is usually not observed in the structures
of their complexes and, when observed an interaction
between the flavin and the nicotinamide compatible
with hydride transfer is not present [114,138,145,147].
Catalytic differences between NADP
+

-dependent
members are related to the different energies required
to produce stacking of the nicotinamide at the re-face
to FAD [119,138]. These observations are compatible
with a mechanism in which the initial interactions
between the enzyme and 2¢P-AMP must evolve
towards the production of alternative structures for
each protein. The fine-tuning of the enzyme catalytic
efficiency is governed by the distance between and
mutual orientation of the N5 of FAD and the nicotin-
amide C4. Therefore, it is reasonable to suppose that
ancestral FNR adapted its NAD(P)
+
⁄ H-binding site,
modulating unique orientations to adapt its efficiency
to the coenzyme oxidation or reduction rates required
in each particular electron transfer chain.
Applications of current knowledge
about Flds and FNRs
The NADPH-producing electron transfer chain has
been used to explore the possibility of redesigning
Flavoproteins in photosynthetic electron transfer M. Medina
3950 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
existing electron transfer systems so that they can per-
form functions other than that for which they were
synthesized [148,149]. Strategies to engineer stress tol-
erance in plants based on the typical stress response of
photosynthetic micro-organisms are underway [150–
152]. The change in the enzyme specificity with respect
to its coenzyme is another example of redesign. FNR

regions involved in coenzyme binding have been mod-
elled to mimic the site in NAD
+
⁄ H-dependent
enzymes [115–118,120], but further work is needed to
improve their catalytic efficiency. Finally, Flds and
FNRs are essential for the survival of some human
pathogens and so may be important in the field of
drug design [126,153–155].
Acknowledgements
This work has been supported by Ministerio de Educa-
cio
´
n y Ciencia, Spain (Grant BIO2007-65890-C02-01).
I would like to thank Drs J. Sancho and C. Go
´
mez-
Moreno who initially introduced me in the Flavopro-
teins and Photosynthesis. In particular, I appreciate all
the support and collaboration received from Dr
Go
´
mez-Moreno over more than 20 years. I also thank
to Drs M. L. Peleato, M. F. Fillat and T. Bes, as well
as all those collaborators that over the years have
helped us to obtain X-ray structures and kinetic data:
Drs J. A. Hermoso, G. Tollin, J. K. Hurley, M. A.
Rosa, M. Herva
´
s and J. A. Navarro. Finally, I must

thank to my current and former PhD students, Dr M.
Martı
´
nez-Ju´ lvez, Dr J. Tejero, Dr I. Nogue
´
s, Dr S.
Frago, J. R. Peregrina, G. Gon
˜
i, A. Serrano, B. Her-
guedas and I. Lans for their collaboration and interest
to better understand the FNR system and for all they
teach me everyday.
References
1Mu
¨
ller F (1990) Chemistry and Biochemistry of Flavo-
enzymes. CRC Press, Boca Raton, FL.
2 Golbeck JH (2006) Photosystem I. The Light-driven
Platocyanin:Ferredoxin Oxidoreductase. Springer,
Dordrecht.
3 Vishniac W & Ochoa S (1952) Fixation of carbon
dioxide coupled to photochemical reduction of pyridine
nucleotides by chloroplast preparations. J Biol Chem
195, 75–93.
4 Arakaki AK, Ceccarelli EA & Carrillo N (1997) Plant-
type ferredoxin–NADP
+
reductases: a basal structural
framework and a multiplicity of functions. FASEB J
11, 133–140.

5 Medina M & Go
´
mez-Moreno C (2004) Interaction of
ferredoxin–NADP
+
reductase with its substrates: opti-
mal interaction for efficient electron transfer. Photosyn-
th Res 79, 113–131.
6 Bottin H & Lagoutte B (1992) Ferredoxin and flavo-
doxin from the cyanobacterium Synechocystis sp
PCC 6803. Biochim Biophys Acta 1101, 48–56.
7Se
`
tif P (2006) Electron transfer from the bound iron–
sulfur clusters to ferredoxin ⁄ flavodoxin: kinetic and
structural properties of ferredoxin ⁄ flavodoxin reduc-
tion by photosystem I. In Photosystem I. The Light-dri-
ven Platocyanin:Ferredoxin Oxidoreductase (Golbeck
JH, ed.), pp. 439–454. Springer, Dordrecht.
8 Hurley JK, Tollin G, Medina M & Go
´
mez-Moreno C
(2006) Electron transfer from ferredoxin and flavodox-
in to ferredoxin-NADP
+
reductase. In Photosystem I.
The Light-driven Platocyanin:Ferredoxin Oxido-
reductase (Golbeck JH, ed.), pp. 455–476. Springer,
Dordrecht.
9 Fromme P, Jordan P & Kranb N (2001) Structure of

photosystem I. Biochim Biophys Acta 1507, 5–31.
10 Jordan P, Fromme P, Witt hydride transfer, Klukas O,
Saenger W & Kranb N (2001) Three-dimensional
structure of cyanobacterial photosystem I at 2.5 A
˚
resolution. Nature 411, 909–917.
11 Ben-Shem A, Frolow F & Nelson N (2003) Crystal
structure of plant photosystem I. Nature 426, 630–635.
12 Antonkine ML, Jordan P, Fromme P, Kranb N,
Golbeck JH & Stehlik D (2003) Assembly of protein
subunits within the stromal ridge of photosystem I.
Structural changes between unbound and sequentially
PS I-bound polypeptides and correlated changes of the
magnetic properties of the terminal iron sulfur clusters.
J Mol Biol 327, 671–697.
13 Mu
¨
hlenhoff U, Kruip J, Bryant DA, Rogner M, Se
`
tif
P & Boekema E (1996) Characterization of a redox-
active cross-linked complex between cyanobacterial
photosystem I and its physiological acceptor flavodox-
in. EMBO J 15, 488–497.
14 Se
`
tif P (2001) Ferredoxin and flavodoxin reduction by
photosystem I. Biochim Biophys Acta 1507, 161–179.
15 Se
`

tif P, Fischer N, Lagoutte B, Bottin H & Rochaix
JD (2002) The ferredoxin docking site of photosys-
tem I. Biochim Biophys Acta 1555, 204–209.
16 Lelong C, Boekema EJ, Kruip J, Bottin H, Rogner
M&Se
`
tif P (1996) Characterization of a redox
active cross-linked complex between cyanobacterial
photosystem I and soluble ferredoxin. EMBO J 15,
2160–2168.
17 Draper RD & Ingraham LL (1968) A potentiometric
study of the flavin semiquinone equilibrium. Arch Bio-
chem Biophys 125, 802–808.
18 Mayhew SG (1999) The effects of pH and semiquinone
formation on the oxidation–reduction potentials of fla-
vin mononucleotide. A reappraisal. Eur J Biochem 265,
698–702.
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3951
19 Mayhew SG, Foust GP & Massey V (1969) Oxida-
tion–reduction properties of flavodoxin from Pepto-
streptococcus elsdenii. J Biol Chem 244, 803–810.
20 Massey V (2000) The chemical and biological versatil-
ity of riboflavin. Biochem Soc Trans 28, 283–296.
21 Mayhew SG & Ludwig ML (1975) Flavodoxins and
electron-transferring flavoproteins. Enzymes 12, 57–118.
22 Mayhew SG & Tollin G (1992) General properties of
flavodoxins. In Chemistry and Biochemistry of Flavoen-
zymes (Mu
¨

ller F, ed.), pp. 389–426. CRC Press, Boca
Raton, FL.
23 Swenson RP & Krey GD (1994) Site-directed mutagen-
esis of tyrosine-98 in the flavodoxin from Desulfovib-
rio vulgaris (Hildenborough): regulation of oxidation–
reduction properties of the bound FMN cofactor by
aromatic, solvent, and electrostatic interactions.
Biochemistry 33, 8505–8514.
24 Lostao A, Go
´
mez-Moreno C, Mayhew SG & Sancho J
(1997) Differential stabilization of the three FMN
redox forms by tyrosine 94 and tryptophan 57 in flavo-
doxin from Anabaena and its influence on the redox
potentials. Biochemistry 36, 14334–14344.
25 Frago S, Gon
˜
i G, Herguedas B, Peregrina JR, Serrano
A, Pe
´
rez-Dorado I, Molina R, Go
´
mez-Moreno C,
Hermoso JA, Martı
´
nez-Ju´ lvez M et al. (2007) Tuning
of the FMN binding and oxido-reduction properties by
neighboring side chains in Anabaena flavodoxin. Arch
Biochem Biophys 467, 206–217.
26 Nogue

´
s I, Campos LA, Sancho J, Go
´
mez-Moreno C,
Mayhew SG & Medina M (2004) Role of neighboring
FMN side chains in the modulation of flavin reduction
potentials and in the energetics of the FMN:apoprotein
interaction in Anabaena flavodoxin. Biochemistry 43,
15111–15121.
27 Fukuyama K, Wakabayashi S, Matsubara H & Rogers
LJ (1990) Tertiary structure of oxidized flavodoxin
from an eukaryotic red alga Chondrus crispus at 2.35 A
˚
resolution. Localization of charged residues and impli-
cation for interaction with electron transfer partners.
J Biol Chem 265, 15804–15812.
28 Rao ST, Shaffie F, Yu C, Satyshur KA, Stockman BJ,
Markley JL & Sundarlingam M (1992) Structure of the
oxidized long-chain flavodoxin from Anabaena 7120 at
2A
˚
resolution. Protein Sci 1, 1413–1427.
29 Freigang J, Diederichs K, Schafer KP, Welte W &
Paul R (2002) Crystal structure of oxidized flavodoxin,
an essential protein in Helicobacter pylori. Protein Sci
11, 253–261.
30 van Mierlo CP, van der Sanden BP, van Woensel P,
Muller F & Vervoort J (1990) A two-dimensional
1
H-NMR study on Megasphaera elsdenii flavodoxin in

the oxidized state and some comparisons with the two-
electron-reduced state. Eur J Biochem 194, 199–216.
31 Knauf MA, Lohr F, Blumel M, Mayhew SG &
Ruterjans H (1996) NMR investigation of the solution
conformation of oxidized flavodoxin from Desulfo-
vibrio vulgaris. Determination of the tertiary structure
and detection of protein-bound water molecules. Eur J
Biochem 238, 423–434.
32 Bradley LH & Swenson RP (1999) Role of glutamate-
59 hydrogen bonded to N(3)H of the flavin mono-
nucleotide cofactor in the modulation of the redox
potentials of the Clostridium beijerinckii flavodoxin.
Glutamate-59 is not responsible for the pH dependency
but contributes to the stabilization of the flavin semiq-
uinone. Biochemistry 38, 12377–12386.
33 Bradley LH & Swenson RP (2001) Role of hydrogen
bonding interactions to N(3)H of the flavin mono-
nucleotide cofactor in the modulation of the redox
potentials of the Clostridium beijerinckii flavodoxin.
Biochemistry 40, 8686–8695.
34 Chang F, Bradley LH & Swenson RP (2001) Evalua-
tion of the hydrogen bonding interactions and their
effects on the oxidation–reduction potentials for the
riboflavin complex of the Desulfovibrio vulgaris flavo-
doxin. Biochim Biophys Acta 1504, 319–328.
35 Chang FC & Swenson RP (1997) Regulation of oxida-
tion–reduction potentials through redox-linked ioniza-
tion in the Y98H mutant of the Desulfovibrio vulgaris
[Hildenborough] flavodoxin: direct proton nuclear
magnetic resonance spectroscopic evidence for the

redox-dependent shift in the pKa of histidine-98.
Biochemistry 36, 9013–9021.
36 Druhan LJ & Swenson RP (1998) Role of methionine
56 in the control of the oxidation–reduction potentials
of the Clostridium beijerinckii flavodoxin: effects of
substitutions by aliphatic amino acids and evidence for
a role of sulfur-flavin interactions. Biochemistry 37,
9668–9678.
37 McCarthy AA, Walsh MA, Verma CS, O’Connell DP,
Reinhold M, Yalloway GN, D’Arcy D, Higgins TM,
Voordouw G & Mayhew SG (2002) Crystallographic
investigation of the role of aspartate 95 in the modula-
tion of the redox potentials of Desulfovibrio vulgaris
flavodoxin. Biochemistry 41, 10950–10962.
38 Romero A, Caldeira J, Legall J, Moura I, Moura JJ &
Romao MJ (1996) Crystal structure of flavodoxin from
Desulfovibrio desulfuricans ATCC 27774 in two oxida-
tion states. Eur J Biochem 239, 190–196.
39 Watt W, Tulinsky A, Swenson RP & Watenpaugh KD
(1991) Comparison of the crystal structures of a flavo-
doxin in its three oxidation states at cryogenic temper-
atures. J Mol Biol 218, 195–208.
40 Ludwig ML, Pattridge KA, Metzger AL, Dixon MM,
Eren M, Feng Y & Swenson RP (1997) Control of
oxidation–reduction potentials in flavodoxin from
Clostridium beijerinckii: the role of conformation
changes. Biochemistry 36, 1259–1280.
41 Hoover DM, Drennan CL, Metzger AL, Osborne C,
Weber CH, Pattridge KA & Ludwig ML (1999)
Flavoproteins in photosynthetic electron transfer M. Medina

3952 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
Comparisons of wild-type and mutant flavodoxins
from Anacystis nidulans. Structural determinants
of the redox potentials. J Mol Biol 294,
725–743.
42 Massey V & Palmer G (1966) On the existence of spec-
trally distinct classes of flavoprotein semiquinones. A
new method for the quantitative production of flavo-
protein semiquinones. Biochemistry 5, 3181–3189.
43 Hoover DM & Ludwig ML (1997) A flavodoxin that
is required for enzyme activation: the structure of oxi-
dized flavodoxin from Escherichia coli at 1.8 A
˚
resolu-
tion. Protein Sci 6, 2525–2537.
44 Alagaratnam S, van Pouderoyen G, Pijning T, Dijkstra
BW, Cavazzini D, Rossi GL, Van Dongen WM, van
Mierlo CP, van Berkel WJ & Canters GW (2005) A
crystallographic study of Cys69Ala flavodoxin II from
Azotobacter vinelandii : structural determinants of redox
potential. Protein Sci 14, 2284–2295.
45 Hu Y, Li Y, Zhang X, Guo X, Xia B & Jin C (2006)
Solution structures and backbone dynamics of a flavo-
doxin mioC from Escherichia coli in both apo- and
holo- forms: implications for cofactor binding and
electron transfer. J Biol Chem 281, 35454–35466.
46 Kasim M & Swenson RP (2001) Alanine-scanning of
the 50’s loop in the Clostridium beijerinckii flavodoxin:
evaluation of additivity and the importance of interac-
tions provided by the main chain in the modulation of

the oxidation-reduction potentials. Biochemistry 40,
13548–13555.
47 Chang FC & Swenson RP (1999) The midpoint poten-
tials for the oxidized-semiquinone couple for Gly57
mutants of the Clostridium beijerinckii flavodoxin cor-
relate with changes in the hydrogen-bonding interac-
tion with the proton on N(5) of the reduced flavin
mononucleotide cofactor as measured by NMR chemi-
cal shift temperature dependencies. Biochemistry 38,
7168–7176.
48 Gon
˜
i G, Herguedas B, Herva
´
s M, Peregrina JR, De la
Rosa MA, Go
´
mez-Moreno C, Navarro JA, Hermoso
JA, Martı
´
nez-Ju´ lvez M & Medina M (2009) Flavodox-
in: a compromise between efficiency and versatility in
the electron transfer from photosystem I to ferredoxin-
NADP
+
reductase. Biochem Biophys Acta Bioenerget-
ics 1787, 144–154.
49 Geoghegan SM, Mayhew SG, Yalloway GN & Butler
G (2000) Cloning, sequencing and expression of the
gene for flavodoxin from Megasphaera elsdenii and the

effects of removing the protein negative charge that is
closest to N(1) of the bound FMN. Eur J Biochem
267, 4434–4444.
50 Zhou Z & Swenson RP (1995) Electrostatic effects of
surface acidic amino acid residues on the oxidation-
reduction potentials of the flavodoxin from Desulfovib-
rio vulgaris (Hildenborough). Biochemistry 34,
3183–3192.
51 Zhou Z & Swenson RP (1996) The cumulative electro-
static effect of aromatic stacking interactions and the
negative electrostatic environment of the flavin mono-
nucleotide binding site is a major determinant of the
reduction potential for the flavodoxin from Desulfovib-
rio vulgaris [Hildenborough]. Biochemistry 35, 15980–
15988.
52 Feng Y & Swenson RP (1997) Evaluation of the role
of specific acidic amino acid residues in electron trans-
fer between the flavodoxin and cytochrome c
3
from
Desulfovibrio vulgaris. Biochemistry 36, 13617–13628.
53 Gon
˜
i G, Serrano A, Frago S, Herva
´
s M, Peregrina JR,
De la Rosa MA, Go
´
mez-Moreno C, Navarro JA &
Medina M (2008) Flavodoxin-mediated electron trans-

fer from photosystem I to ferredoxin–NADP
+
reduc-
tase in Anabaena: role of flavodoxin hydrophobic
residues in protein–protein interactions. Biochemistry
47, 1207–1217.
54 Martı
´
nez JI, Alonso PJ, Go
´
mez-Moreno C &
Medina M (1997) One- and two-dimensional ESEEM
spectroscopy of flavoproteins. Biochemistry 36,
15526–15537.
55 Medina M, Go
´
mez-Moreno C & Cammack R (1995)
Electron spin resonance and electron nuclear double
resonance studies of flavoproteins involved in the pho-
tosynthetic electron transport in the cyanobacterium
Anabaena sp. PCC 7119. Eur J Biochem 227, 529–536.
56 Medina M, Lostao A, Sancho J, Go
´
mez-Moreno C,
Cammack R, Alonso PJ & Martı
´
nez JI (1999) Elec-
tron-nuclear double resonance and hyperfine sublevel
correlation spectroscopic studies of flavodoxin mutants
from Anabaena sp. PCC 7119. Biophys J 77, 1712–

1720.
57 Karplus PA & Bruns CM (1994) Structure–function
relations for ferredoxin reductase. J Bioenerg Bio-
membr 26, 89–99.
58 Bruns CM & Karplus PA (1995) Refined crystal struc-
ture of spinach ferredoxin reductase at 1.7 A
˚
resolu-
tion: oxidized, reduced and 2’-phospho-5’-AMP bound
states. J Mol Biol 247, 125–145.
59 Deng Z, Aliverti A, Zanetti G, Arakaki AK, Ottado J,
Orellano EG, Calcaterra NB, Ceccarelli EA, Carrillo
N & Karplus PA (1999) A productive NADP
+
binding
mode of ferredoxin–NADP
+
reductase revealed by
protein engineering and crystallographic studies. Nat
Struct Biol 6, 847–853.
60 Dorowski A, Hofmann A, Steegborn C, Boicu M &
Huber R (2001) Crystal structure of paprika ferre-
doxin–NADP
+
reductase. Implications for the electron
transfer pathway. J Biol Chem 276, 9253–9263.
61 Aliverti A, Faber R, Finnerty CM, Ferioli C, Pandini
V, Negri A, Karplus PA & Zanetti G (2001) Biochemi-
cal and crystallographic characterization of ferredoxin–
NADP

+
reductase from nonphotosynthetic tissues.
Biochemistry 40, 14501–14508.
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3953
62 Serre L, Vellieux FM, Medina M, Go
´
mez-Moreno C,
Fontecilla-Camps JC & Frey M (1996) X-Ray struc-
ture of the ferredoxin:NADP
+
reductase from the cya-
nobacterium Anabaena PCC 7119 at 1.8 A
˚
resolution,
and crystallographic studies of NADP
+
binding at
2.25 A
˚
resolution. J Mol Biol 263, 20–39.
63 Faro M, Go
´
mez-Moreno C, Stankovich M & Medina
M (2002) Role of critical charged residues in reduction
potential modulation of ferredoxin–NADP
+
reductase.
Eur J Biochem 269, 2656–2661.
64 Corrado ME, Aliverti A, Zanetti G & Mayhew SG

(1996) Analysis of the oxidation–reduction potentials
of recombinant ferredoxin–NADP
+
reductase from
spinach chloroplasts. Eur J Biochem 239, 662–667.
65 Martı
´
nez-Ju´ lvez M, Nogue
´
s I, Faro M, Hurley JK,
Brodie TB, Mayoral T, Sanz-Aparicio J, Hermoso JA,
Stankovich MT, Medina M et al. (2001) Role of a
cluster of hydrophobic residues near the FAD cofactor
in Anabaena PCC 7119 ferredoxin-NADP
+
reductase
for optimal complex formation and electron transfer to
ferredoxin. J Biol Chem 276, 27498–27510.
66 Batie CJ & Kamin H (1986) Association of ferredoxin–
NADP
+
reductase with NADP(H) specificity and
oxidation–reduction properties. J Biol Chem 261,
11214–11223.
67 Cassan N, Lagoutte B & Se
`
tif P (2005) Ferredoxin-
NADP
+
reductase. Kinetics of electron transfer, tran-

sient intermediates, and catalytic activities studied by
flash-absorption spectroscopy with isolated photosys-
tem I and ferredoxin. J Biol Chem 280, 25960–25972.
68 Arakaki AK, Orellano EG, Calcaterra NB, Ottado J &
Ceccarelli EA (2001) Involvement of the flavin si-face
tyrosine on the structure and function of ferredoxin–
NADP
+
reductases. J Biol Chem 276, 44419–44426.
69 Hurley JK, Weber-Main AM, Stankovich MT, Ben-
ning MM, Thoden JB, Vanhooke JL, Holden HM,
Chae YK, Xia B, Cheng H et al. (1997) Structure–
function relationships in Anabaena ferredoxin: correla-
tions between X-ray crystal structures, reduction
potentials, and rate constants of electron transfer to
ferredoxin:NADP
+
reductase for site-specific ferre-
doxin mutants. Biochemistry 36, 11100–11117.
70 Batie CJ & Kamin H (1984) Electron transfer by ferre-
doxin:NADP
+
reductase. Rapid-reaction evidence for
participation of a ternary complex. J Biol Chem 259,
11976–11985.
71 Sancho J & Go
´
mez-Moreno C (1991) Interaction of
ferredoxin-NADP
+

reductase from Anabaena with its
substrates. Arch Biochem Biophys 288, 231–238.
72 Medina M & Cammack R (2007) ENDOR and related
EMR methods applied to flavoprotein radicals. Appl
Magn Reson 31, 457–470.
73 Ullmann GM, Hauswald M, Jensen A & Knapp EW
(2000) Structural alignment of ferredoxin and flavo-
doxin based on electrostatic potentials: implications for
their interactions with photosystem I and ferredoxin-
NADP
+
reductase. Proteins 38, 301–309.
74 Fromme P, Bottin H, Kranb N&Se
`
tif P (2002) Crys-
tallization and electron paramagnetic resonance char-
acterization of the complex of photosystem I with its
natural electron acceptor ferredoxin. Biophys J 83,
1760–1773.
75 Mu
¨
hlenhoff U, Zhao J & Bryant DA (1996) Interac-
tion between photosystem I and flavodoxin from the
cyanobacterium Synechococcus sp. PCC 7002 as
revealed by chemical cross-linking. Eur J Biochem 235,
324–331.
76 Li N, Zhao JD, Warren PV, Warden JT, Bryant DA
& Golbeck JH (1991) PsaD is required for the stable
binding of PsaC to the photosystem I core protein of
Synechococcus sp. PCC 6301. Biochemistry 30, 7863–

7872.
77 Chitnis VP, Jungs YS, Albee L, Golbeck JH & Chitnis
PR (1996) Mutational analysis of photosystem I poly-
peptides. Role of PsaD and the lysyl 106 residue in the
reductase activity of the photosystem I. J Biol Chem
271, 11772–11780.
78 Barth P, Lagoutte B & Se
`
tif P (1998) Ferredoxin
reduction by photosystem I from Synechocystis sp.
PCC 6803: toward an understanding of the respective
roles of subunits PsaD and PsaE in ferredoxin binding.
Biochemistry 37, 16233–16241.
79 van Thor JJ, Geerlings TH, Matthijs HC & Helling-
werf KJ (1999) Kinetic evidence for the PsaE-depen-
dent transient ternary complex
photosystem I ⁄ ferredoxin ⁄
ferredoxin:NADP
+
reductase in a cyanobacterium.
Biochemistry 38, 12735–12746.
80 Fischer N, Hippler M, Se
`
tif P, Jacquot JP & Rochaix
JD (1998) The PsaC subunit of photosystem I provides
an essential lysine residue for fast electron transfer to
ferredoxin. EMBO J 17, 849–858.
81 Hanley J, Se
`
tif P, Bottin H & Lagoutte B (1996) Muta-

genesis of photosystem I in the region of the ferredoxin
cross-linking site: modifications of positively charged
amino acids. Biochemistry 35, 8563–8571.
82 Meimberg K, Fischer N, Rochaix JD & Mu
¨
hlenhoff U
(1999) Lys35 of PsaC is required for the efficient pho-
toreduction of flavodoxin by photosystem I from Chla-
mydomonas reinhardtii. Eur J Biochem 263, 137–144.
83 Meimberg K, Lagoutte B, Bottin H & Mu
¨
hlenhoff U
(1998) The PsaE subunit is required for complex for-
mation between photosystem I and flavodoxin from
the cyanobacterium Synechocystis sp. PCC 6803.
Biochemistry 37, 9759–9767.
84 Navarro JA, Herva
´
s M, Genzor CG, Cheddar G, Fil-
lat MF, De la Rosa MA, Go
´
mez-Moreno C, Cheng H,
Xia B, Chae YK et al. (1995) Site-specific mutagenesis
demonstrates that the structural requirements for effi-
cient electron transfer in Anabaena ferredoxin and
Flavoproteins in photosynthetic electron transfer M. Medina
3954 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
flavodoxin are highly dependent on the reaction part-
ner: kinetic studies with photosystem I, ferredoxin–
NADP

+
reductase, and cytochrome c. Arch Biochem
Biophys 321, 229–238.
85 Poncelet M, Cassier-Chauvat C, Leschelle X, Bottin H
& Chauvat F (1998) Targeted deletion and mutational
analysis of the essential (2Fe-2S) plant-like ferredoxin
in Synechocystis PCC 6803 by plasmid shuffling. Mol
Microbiol 28, 813–821.
86 Guillouard I, Lagoutte B, Moal G & Bottin H (2000)
Importance of the region including aspartates 57 and
60 of ferredoxin on the electron transfer complex with
photosystem I in the cyanobacterium Synechocystis sp.
PCC 6803. Biochem Biophys Res Commun 271, 647–
653.
87 Medina M, Herva
´
s M, Navarro JA, De la Rosa MA,
Go
´
mez-Moreno C & Tollin G (1992) A laser flash
absorption spectroscopy study of Anabaena sp.
PCC 7119 flavodoxin photoreduction by photosystem I
particles from spinach. FEBS Lett 313, 239–242.
88 Nogue
´
s I, Herva
´
s M, Peregrina JR, Navarro JA, De la
Rosa MA, Go
´

mez-Moreno C & Medina M (2005) An-
abaena flavodoxin as an electron carrier from photo-
system I to ferredoxin–NADP
+
reductase. Role of
flavodoxin residues in protein–protein interaction and
electron transfer. Biochemistry 44, 97–104.
89 Nogue
´
s I, Martı
´
nez-Ju´ lvez M, Navarro JA, Herva
´
sM,
Armenteros L, De la Rosa MA, Brodie TB, Hurley
JK, Tollin G, Go
´
mez-Moreno C et al. (2003) Role of
hydrophobic interactions in the flavodoxin mediated
electron transfer from photosystem I to ferredoxin-
NADP
+
reductase in Anabaena PCC. Biochemistry 42,
2036–2045.
90 Casaus JL, Navarro JA, Herva
´
s M, Lostao A, De la
Rosa MA, Go
´
mez-Moreno C, Sancho J & Medina M

(2002) Anabaena sp. PCC 7119 flavodoxin as electron
carrier from photosystem I to ferredoxin-NADP+
reductase. Role of Trp(57) and Tyr(94). J Biol Chem
277, 22338–22344.
91 Nogue
´
sI,Pe
´
rez-Dorado I, Frago S, Bittel C, Mayhew
SG, Go
´
mez-Moreno C, Hermoso JA, Medina M, Cor-
tez N & Carrillo N (2005) The ferredoxin-NADP(H)
reductase from Rhodobacter capsulatus: molecular
structure and catalytic mechanism. Biochemistry 44,
11730–11740.
92 Morales R, Charon MH, Kachalova G, Serre L,
Medina M, Go
´
mez-Moreno C & Frey M (2000) A
redox-dependent interaction between two electron-
transfer partners involved in photosynthesis. EMBO
Rep 1, 271–276.
93 Kurisu G, Kusunoki M, Katoh E, Yamazaki T,
Teshima K, Onda Y, Kimata-Ariga Y & Hase T
(2001) Structure of the electron transfer complex
between ferredoxin and ferredoxin-NADP
+
reductase.
Nat Struct Biol 8, 117–121.

94 Hanke GT, Kimata-Ariga Y, Taniguchi I & Hase T
(2004) A post genomic characterization of Arabidopsis
ferredoxins. Plant Physiol 134, 255–264.
95 Mayoral T, Martı
´
nez-Ju´ lvez M, Pe
´
rez-Dorado I, Sanz-
Aparicio J, Go
´
mez-Moreno C, Medina M & Hermoso
JA (2005) Structural analysis of interactions for com-
plex formation between ferredoxin-NADP
+
reductase
and its protein partners. Proteins 59, 592–602.
96 Aliverti A, Bruns CM, Pandini VE, Karplus PA,
Vanoni MA, Curti B & Zanetti G (1995) Involvement
of serine 96 in the catalytic mechanism of ferredoxin-
NADP
+
reductase: structure-function relationship
as studied by site-directed mutagenesis and X-ray
crystallography. Biochemistry 34, 8371–8379.
97 Aliverti A, Corrado ME & Zanetti G (1994) Involve-
ment of lysine-88 of spinach ferredoxin-NADP
+
reduc-
tase in the interaction with ferredoxin. FEBS Lett 343,
247–250.

98 Aliverti A, Deng Z, Ravasi D, Piubelli L, Karplus PA
& Zanetti G (1998) Probing the function of the invari-
ant glutamyl residue 312 in spinach ferredoxin-
NADP
+
reductase. J Biol Chem 273, 34008–34015.
99 Aliverti A, Lubberstedt T, Zanetti G, Herrmann RG &
Curti B (1991) Probing the role of lysine 116 and lysine
244 in the spinach ferredoxin-NADP
+
reductase by site-
directed mutagenesis. J Biol Chem 266, 17760–17763.
100 Martı
´
nez-Ju´ lvez M, Medina M & Go
´
mez-Moreno C
(1999) Ferredoxin-NADP
+
reductase uses the same
site for the interaction with ferredoxin and flavodoxin.
J Biol Inorg Chem 4, 568–578.
101 Hurley JK, Morales R, Martı
´
nez-Ju´ lvez M, Brodie TB,
Medina M, Go
´
mez-Moreno C & Tollin G (2002)
Structure-function relationships in Anabaena ferre-
doxin ⁄ ferredoxin-NADP

+
reductase electron transfer:
insights from site-directed mutagenesis, transient
absorption spectroscopy and X-ray crystallography.
Biochim Biophys Acta 1554, 5–21.
102 Aliverti A, Livraghi A, Piubelli L & Zanetti G (1997)
On the role of the acidic cluster Glu 92-94 of spinach
ferredoxin I. Biochim Biophys Acta 1342, 45–50.
103 Hurley JK, Hazzard JT, Martı
´
nez-Ju´ lvez M, Medina
M, Go
´
mez-Moreno C & Tollin G (1999) Electrostatic
forces involved in orienting Anabaena ferredoxin dur-
ing binding to Anabaena ferredoxin:NADP
+
reductase:
site-specific mutagenesis, transient kinetic measure-
ments, and electrostatic surface potentials. Protein Sci
8, 1614–1622.
104 Hurley JK, Salamon Z, Meyer TE, Fitch JC, Cusano-
vich MA, Markley JL, Cheng H, Xia B, Chae YK,
Medina M et al. (1993) Amino acid residues in Anaba-
ena ferredoxin crucial to interaction with ferredoxin-
NADP
+
reductase: site-directed mutagenesis and laser
flash photolysis. Biochemistry 32, 9346–9354.
105 Hurley JK, Cheng H, Xia B, Markley JL, Medina M,

Go
´
mez-Moreno C & Tollin G (1993) An aromatic
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3955
amino-acid is required at position-65 in Anabaena
ferredoxin for rapid electron transfer to ferredoxin-
NADP
+
reductase. J Am Chem Soc 115, 11698–11701.
106 Hurley JK, Medina M, Go
´
mez-Moreno C & Tollin G
(1994) Further characterization by site-directed muta-
genesis of the protein–protein interface in the ferre-
doxin ⁄ ferredoxin:NADP
+
reductase system from
Anabaena: requirement of a negative charge at position
94 in ferredoxin for rapid electron transfer. Arch Bio-
chem Biophys 312, 480–486.
107 Martı
´
nez-Ju´ lvez M, Medina M, Hurley JK, Hafezi R,
Brodie TB, Tollin G & Go
´
mez-Moreno C (1998)
Lys75 of Anabaena ferredoxin-NADP
+
reductase is a

critical residue for binding ferredoxin and flavodoxin
during electron transfer. Biochemistry 37, 13604–13613.
108 Martı
´
nez-Ju´ lvez M, Hermoso J, Hurley JK, Mayoral
T, Sanz-Aparicio J, Tollin G, Go
´
mez-Moreno C &
Medina M (1998) Role of Arg100 and Arg264 from
Anabaena PCC 7119 ferredoxin-NADP
+
reductase for
optimal NADP
+
binding and electron transfer. Bio-
chemistry 37, 17680–17691.
109 Faro M, Frago S, Mayoral T, Hermoso JA, Sanz-
Aparicio J, Go
´
mez-Moreno C & Medina M (2002)
Probing the role of glutamic acid 139 of Anabaena
ferredoxin–NADP
+
reductase in the interaction with
substrates. Eur J Biochem 269, 4938–4947.
110 Medina M, Abagyan R, Go
´
mez-Moreno C & Fernan-
dez-Recio J (2008) Docking analysis of transient com-
plexes: interaction of ferredoxin–NADP

+
reductase
with ferredoxin and flavodoxin. Proteins 72, 848–862.
111 Medina M, Go
´
mez-Moreno C & Tollin G (1992)
Effects of chemical modification of Anabaena flavodox-
in and ferredoxin-NADP
+
reductase on the kinetics of
interprotein electron transfer reactions. Eur J Biochem
210, 577–583.
112 Worrall JA, Reinle W, Bernhardt R & Ubbink M
(2003) Transient protein interactions studied by NMR
spectroscopy: the case of cytochrome c and adreno-
doxin. Biochemistry 42, 7068–7076.
113 Volkov AN, Ferrari D, Worrall JA, Bonvin AM &
Ubbink M (2005) The orientations of cytochrome c in
the highly dynamic complex with cytochrome b
5
visual-
ized by NMR and docking using HADDOCK. Protein
Sci 14, 799–811.
114 Hermoso JA, Mayoral T, Faro M, Go
´
mez-Moreno
C, Sanz-Aparicio J & Medina M (2002) Mechanism
of coenzyme recognition and binding revealed by
crystal structure analysis of ferredoxin-NADP
+

reduc-
tase complexed with NADP
+
. J Mol Biol 319, 1133–
1142.
115 Medina M, Luquita A, Tejero J, Hermoso J, Mayoral
T, Sanz-Aparicio J, Grever K & Go
´
mez-Moreno C
(2001) Probing the determinants of coenzyme specific-
ity in ferredoxin-NADP
+
reductase by site-directed
mutagenesis. J Biol Chem 276, 11902–11912.
116 Tejero J, Martı
´
nez-Ju´ lvez M, Mayoral T, Luquita A,
Sanz-Aparicio J, Hermoso JA, Hurley JK, Tollin G,
Go
´
mez-Moreno C & Medina M (2003) Involvement of
the pyrophosphate and the 2’-phosphate binding
regions of ferredoxin-NADP
+
reductase in coenzyme
specificity. J Biol Chem 278, 49203–49214.
117 Tejero J, Pe
´
rez-Dorado I, Maya C, Martı
´

nez-Ju´ lvez M,
Sanz-Aparicio J, Go
´
mez-Moreno C, Hermoso JA &
Medina M (2005) C-terminal tyrosine of ferredoxin-
NADP
+
reductase in hydride transfer processes with
NAD(P)
+
⁄ H. Biochemistry 44, 13477–13490.
118 Piubelli L, Aliverti A, Arakaki AK, Carrillo N, Cec-
carelli EA, Karplus PA & Zanetti G (2000) Competi-
tion between C-terminal tyrosine and nicotinamide
modulates pyridine nucleotide affinity and specificity in
plant ferredoxin-NADP
+
reductase. J Biol Chem 275,
10472–10476.
119 Musumeci MA, Arakaki AK, Rial DV, Catalano-
Dupuy DL & Ceccarelli EA (2008) Modulation of the
enzymatic efficiency of ferredoxin-NADP(H) reductase
by the amino acid volume around the catalytic site.
FEBS J 275, 1350–1366.
120 Peregrina JR, Herguedas B, Hermoso JA, Martı
´
nez-
Ju´ lvez M & Medina M (2009) Protein motifs involved
in coenzyme interaction and enzymatic efficiency in
Anabaena ferredoxin-NADP

+
reductase. Biochemistry
48, 3109–3119.
121 Tejero J, Peregrina JR, Martı
´
nez-Ju´ lvez M, Gutierrez
A, Go
´
mez-Moreno C, Scrutton NS & Medina M
(2007) Catalytic mechanism of hydride transfer
between NADP
+
⁄ H and ferredoxin-NADP
+
reductase
from Anabaena PCC 7119. Arch Biochem Biophys 459,
79–90.
122 Gutierrez A, Doehr O, Paine M, Wolf CR, Scrutton
NS & Roberts GC (2000) Trp-676 facilitates nicotin-
amide coenzyme exchange in the reductive half-reac-
tion of human cytochrome P450 reductase: properties
of the soluble W676H and W676A mutant reductases.
Biochemistry 39, 15990–15999.
123 Dohr O, Paine MJ, Friedberg T, Roberts GC & Wolf
CR (2001) Engineering of a functional human NADH-
dependent cytochrome P450 system. Proc Natl Acad
Sci USA 98, 81–86.
124 Elmore CL & Porter TD (2002) Modification of the
nucleotide cofactor-binding site of cytochrome P-450
reductase to enhance turnover with NADH in vivo.

J Biol Chem 277, 48960–48964.
125 Adak S, Sharma M, Meade AL & Stuehr DJ (2002) A
conserved flavin-shielding residue regulates NO syn-
thase electron transfer and nicotinamide coenzyme
specificity. Proc Natl Acad Sci USA 99, 13516–13521.
126 Aliverti A, Pandini V, Pennati A, de Rosa M &
Zanetti G (2008) Structural and functional diversity
of ferredoxin-NADP
+
reductases. Arch Biochem
Biophys 474, 283–291.
Flavoproteins in photosynthetic electron transfer M. Medina
3956 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS
127 Carrillo N & Ceccarelli EA (2003) Open questions in
ferredoxin-NADP
+
reductase catalytic mechanism. Eur
J Biochem 270, 1900–1915.
128 Peregrina JR, Lans I, Sa
´
nchez-Azqueta A, Frago S,
Gonza
´
lez-Lafont A, LLuch JM, Garcia-Viloca M &
Medina M (2008). Flavin role in protein-protein inter-
action and electron transfer: from photosystem I to
NADPH, a case study. Paper presented at the Flavins
and Flavoproteins, Jaca, Spain.
129 Aliverti A, Piubelli L, Zanetti G, Lubberstedt T, Herr-
mann RG & Curti B (1993) The role of cysteine residues

of spinach ferredoxin-NADP
+
reductase as assessed by
site-directed mutagenesis. Biochemistry 32, 6374–6380.
130 Medina M, Martı
´
nez-Ju´ lvez M, Hurley JK, Tollin G &
Go
´
mez-Moreno C (1998) Involvement of glutamic acid
301 in the catalytic mechanism of ferredoxin-NADP
+
reductase from Anabaena PCC 7119. Biochemistry 37,
2715–2728.
131 Mayoral T, Medina M, Sanz-Aparicio J, Go
´
mez-
Moreno C & Hermoso JA (2000) Structural basis of
the catalytic role of Glu301 in Anabaena PCC 7119
ferredoxin-NADP
+
reductase revealed by X-ray crys-
tallography. Proteins 38, 60–69.
132 Vela
´
zquez-Campoy A, Gon
˜
i G, Peregrina JR & Med-
ina M (2006) Exact analysis of heterotropic interac-
tions in proteins: characterization of cooperative ligand

binding by isothermal titration calorimetry. Biophys J
91, 1887–1904.
133 Batie CJ & Kamin H (1984) Ferredoxin:NADP
+
oxidoreductase. Equilibria in binary and ternary
complexes with NADP
+
and ferredoxin. J Biol Chem
259, 8832–8839.
134 Sancho J (2006) Flavodoxins: sequence, folding, bind-
ing, function and beyond. Cell Mol Life Sci 63, 855–864.
135 Peleato ML, Ayora S, Inda LA & Go
´
mez-Moreno C
(1994) Isolation and characterization of two different
flavodoxins from the eukaryote Chlorella fusca. Bio-
chem J 302 (Pt 3), 807–811.
136 Rogers LJ (1987) Ferredoxins, flavodoxins and related
proteins: structure, function and evolution. In The Cy-
anobacteria (Fay P & Van Baalen C, eds), pp. 35–67.
Elsevier, Amsterdam.
137 Price NT, Smith AJ & Rogers LJ (1991) Relationship
of the flavodoxin isoforms from Porphyra umbilicalis.
Phytochemistry 30, 2841–2843.
138 Wolthers KR & Scrutton NS (2007) Protein interac-
tions in the human methionine synthase–methionine
synthase reductase complex and implications for the
mechanism of enzyme reactivation. Biochemistry 46,
6696–6709.
139 Ceccarelli EA, Arakaki AK, Cortez N & Carrillo N

(2004) Functional plasticity and catalytic efficiency in
plant and bacterial ferredoxin-NADP(H) reductases.
Biochim Biophys Acta 1698, 155–165.
140 Bortolotti A, Pe
´
rez-Dorado I, Gon
˜
i G, Medina M,
Hermoso JA, Carrillo N & Cortez N (2009) Coenzyme
binding and hydride transfer in Rhodobacter capsulatus
ferredoxin ⁄ flavodoxin NADP(H) oxidoreductase. Bio-
chim Biophys Acta 1794, 199–210.
141 Panda K, Haque MM, Garcin-Hosfield ED, Durra D,
Getzoff ED & Stuehr DJ (2006) Surface charge inter-
actions of the FMN module govern catalysis by nitric-
oxide synthase. J Biol Chem 281, 36819–36827.
142 Wang M, Roberts DL, Paschke R, Shea TM, Masters
BS & Kim JJ (1997) Three-dimensional structure of
NADPH-cytochrome P450 reductase: prototype for
FMN- and FAD-containing enzymes. Proc Natl Acad
Sci USA 94, 8411–8416.
143 Correll CC, Ludwig ML, Bruns CM & Karplus PA
(1993) Structural prototypes for an extended family of
flavoprotein reductases: comparison of phthalate
dioxygenase reductase with ferredoxin reductase and
ferredoxin. Protein Sci 2, 2112–2133.
144 Wolthers KR, Lou X, Toogood HS, Leys D &
Scrutton NS (2007) Mechanism of coenzyme binding
to human methionine synthase reductase revealed
through the crystal structure of the FNR-like module

and isothermal titration calorimetry. Biochemistry 46,
11833–11844.
145 Bewley MC, Marohnic CC & Barber MJ (2001) The
structure and biochemistry of NADH-dependent cyto-
chrome b
5
reductase are now consistent. Biochemistry
40, 13574–13582.
146 Correll CC, Batie CJ, Ballou DP & Ludwig ML (1992)
Phthalate dioxygenase reductase: a modular structure
for electron transfer from pyridine nucleotides to
[2Fe–2S]. Science 258, 1604–1610.
147 Grunau A, Paine MJ, Ladbury JE & Gutierrez A
(2006) Global effects of the energetics of coenzyme
binding: NADPH controls the protein interaction
properties of human cytochrome P450 reductase.
Biochemistry 45, 1421–1434.
148 Faro M, Schiffler B, Heinz A, Nogue
´
s I, Medina M,
Bernhardt R & Go
´
mez-Moreno C (2003) Insights into
the design of a hybrid system between Anabaena ferre-
doxin–NADP
+
reductase and bovine adrenodoxin.
Eur J Biochem 270, 726–735.
149 Zollner A, Nogue
´

s I, Heinz A, Medina M, Go
´
mez-
Moreno C & Bernhardt R (2004) Analysis of the
interaction of a hybrid system consisting of bovine
adrenodoxin reductase and flavodoxin from the cyano-
bacterium Anabaena PCC7119. Bioelectrochemistry 63,
61–65.
150 Tognetti VB, Palatnik JF, Fillat MF, Melzer M,
Hajirezaei MR, Valle EM & Carrillo N (2006) Func-
tional replacement of ferredoxin by a cyanobacterial
flavodoxin in tobacco confers broad-range stress toler-
ance. Plant Cell 18 , 2035–2050.
M. Medina Flavoproteins in photosynthetic electron transfer
FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS 3957
151 Tognetti VB, Zurbriggen MD, Morandi EN, Fillat
MF, Valle EM, Hajirezaei MR & Carrillo N (2007)
Enhanced plant tolerance to iron starvation by func-
tional substitution of chloroplast ferredoxin with a
bacterial flavodoxin. Proc Natl Acad Sci USA 104,
11495–11500.
152 Zurbriggen MD, Tognetti VB, Fillat MF, Hajirezaei
MR, Valle EM & Carrillo N (2008) Combating stress
with flavodoxin: a promising route for crop improve-
ment. Trends Biotechnol 26, 531–537.
153 Kimata-Ariga Y, Kurisu G, Kusunoki M, Aoki S,
Sato D, Kobayashi T, Kita K, Horii T & Hase T
(2007) Cloning and characterization of ferredoxin and
ferredoxin–NADP
+

reductase from human malaria
parasite. J Biochem 141, 421–428.
154 Leadbeater C, McIver L, Campopiano DJ, Webster
SP, Baxter RL, Kelly SM, Price NC, Lysek DA, Noble
MA, Chapman SK et al. (2000) Probing the NADPH-
binding site of Escherichia coli flavodoxin oxidoreduc-
tase. Biochem J 352 (Pt 2), 257–266.
155 Cremades N, Bueno M, Toja M & Sancho J (2005)
Towards a new therapeutic target: Helicobacter pylori
flavodoxin. Biophys Chem 115, 267–276.
Flavoproteins in photosynthetic electron transfer M. Medina
3958 FEBS Journal 276 (2009) 3942–3958 ª 2009 The Author Journal compilation ª 2009 FEBS