Download Photo-catalytic oxidation of a di-nuclear manganese centre in an

Biochimica et Biophysica Acta 1787 (2009) 1112–1121
Contents lists available at ScienceDirect
Biochimica et Biophysica Acta
j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b b a b i o
Photo-catalytic oxidation of a di-nuclear manganese centre in an engineered
bacterioferritin ‘reaction centre’
Brendon Conlan a,⁎, Nicholas Cox a, Ji-Hu Su b, Warwick Hillier a, Johannes Messinger b,1, Wolfgang Lubitz b,
P. Leslie Dutton c, Tom Wydrzynski a,⁎
a
b
c
Australian National University, Canberra, Australia
Max Planck Institute for Bioinorganic Chemistry, Mülheim an der Ruhr, Germany
University of Pennsylvania, Philadelphia, USA
a r t i c l e
i n f o
Article history:
Received 13 February 2009
Received in revised form 16 April 2009
Accepted 21 April 2009
Available online 3 May 2009
Keywords:
Artificial photosynthesis
EPR
Manganese
Electron transfer
Protein engineering
Bacterioferritin
Zinc chlorin e6
a b s t r a c t
Photosynthesis involves the conversion of light into chemical energy through a series of electron transfer
reactions within membrane-bound pigment/protein complexes. The Photosystem II (PSII) complex in plants,
algae and cyanobacteria catalyse the oxidation of water to molecular O2. The complexity of PSII has thus far
limited attempts to chemically replicate its function. Here we introduce a reverse engineering approach to
build a simple, light-driven photo-catalyst based on the organization and function of the donor side of the
PSII reaction centre. We have used bacterioferritin (BFR) (cytochrome b1) from Escherichia coli as the protein
scaffold since it has several, inherently useful design features for engineering light-driven electron transport.
Among these are: (i.) a di-iron binding site; (ii.) a potentially redox-active tyrosine residue; and (iii.) the
ability to dimerise and form an inter-protein heme binding pocket within electron tunnelling distance of the
di-iron binding site. Upon replacing the heme with the photoactive zinc–chlorin e6 (ZnCe6) molecule and the
di-iron binding site with two manganese ions, we show that the two Mn ions bind as a weakly coupled dinuclear MnII,II
2 centre, and that ZnCe6 binds in stoichiometric amounts of 1:2 with respect to the dimeric form
of BFR. Upon illumination the bound ZnCe6 initiates electron transfer, followed by oxidation of the di-nuclear
Mn centre possibly via one of the inherent tyrosine residues in the vicinity of the Mn cluster. The light
dependent loss of the MnII EPR signals and the formation of low field parallel mode Mn EPR signals are
attributed to the formation of MnIII species. The formation of the MnIII is concomitant with consumption of
oxygen. Our model is the first artificial reaction centre developed for the photo-catalytic oxidation of a dimetal site within a protein matrix which potentially mimics water oxidation centre (WOC) photo-assembly.
© 2009 Elsevier B.V. All rights reserved.
1. Introduction
Oxygenic photosynthesis is the process whereby plants utilize
sunlight to catalyse the reduction of plastoquinone and the oxidation
of water into protons and molecular oxygen. This process is initiated in
the pigment/protein complex called Photosystem II (PSII). The
essential elements of PSII that catalyse the oxidation of water include:
(i.) the strongly oxidizing multi-chlorophyll complex termed P680;
(ii.) a redox-active tyrosine termed YZ; and (iii.) the oxygen evolving
centre (OEC) consisting of four μ-oxo bridged manganese ions and one
Abbreviations: BFR, wild type bacterioferritin; BFR1, bacterioferritin double
mutant H46R, H112R without cofactors; BFR1–M, BFR1 with two manganese ions
bound; BFR1–Z, BFR1 with ZnCe6 bound; BFR1–ZM, BFR1 with stoichiometric amounts
of ZnCe6 and manganese bound; EPR, electron paramagnetic resonance; ITC,
isothermal titration calorimetry; PSII, Photosystem II; ZnCe6, zincII chlorin e6
⁎ Corresponding authors.
E-mail addresses: [email protected] (B. Conlan),
[email protected] (T. Wydrzynski).
1
Current address: Department of Chemistry, Umeå University, Umeå, Sweden.
0005-2728/$ – see front matter © 2009 Elsevier B.V. All rights reserved.
doi:10.1016/j.bbabio.2009.04.011
calcium ion (Mn4OxCa) bound within the protein complex [1]. Upon
light activation of PSII, P680 forms the P680•+ radical cation
(E0 N1.2 V) by transferring an electron to a neighbouring pheophytin
molecule followed by rapid transfer of the electron to a bound
quinone molecule (QA). The charge separated state (P+/Q−
A ) undergoes hole transfer from P680•+ to YZ, and then in turn to the Mn4OxCa
cluster, which couples the one electron photochemistry of P680 to the
four electron chemistry of water oxidation via the sequential advance
of the S-state intermediates (S0–S4) [1].
In the design of artificial photoactive proteins, there are two
approaches that can be considered [2]. The first is the de novo design
followed by organic synthesis. A growing number of de novo proteins
are being synthesized with increasing functionality [3–6]; however,
their synthesis is currently limited to a polypeptide length of about
100 amino acid residues. The second approach, and the one adopted
for this work, involves the modification of natural protein scaffolds.
Target scaffolds are selected based on pre-existing structural features
that may be useful in re-engineering the protein. In addition, the
design of a light-activated redox catalyst has to take into consideration
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
the physiological limitations of intra-protein electron transfer.
Electron tunnelling between redox centres is reasonably accurately
described by a simple exponential decay with distance [7,8]. Packing
density and the secondary structure of the protein matrix play smaller
roles in mediating electron tunnelling rates [9–11]. As such natural
proteins with productive electron transfer reactions have been found
to have cofactors separated by not more than ∼ 14 Å [8,12,13].
The complexity of the PSII enzyme has thus far discouraged
attempts to reverse engineer a complete synthetic water oxidase [14].
As a result, previous engineering efforts have focussed mostly on
constructing biological motifs that mimic one particular aspect of the
water oxidase; for example, chlorin excitation/oxidation [15] and
metal binding/complex formation [16]. Hay et al. (2004) showed
light-activated electron transfer between a chlorin (ZnCe6) and a
quinone engineered into cytochrome b562. The quinone was covalently
attached approximately ∼ 10 Å away from the bound chlorin
producing surprisingly high efficiencies of electron transfer (∼ 20%).
Metal binding/complex formation was shown by Thielges et al. where
the purple non-sulphur bacterial reaction centre was modified to bind
a single MnII ion. Upon illumination, the bound Mn could reduce P870+
suggesting that the metal centre was oxidized [16]. Gray et al. have
shown electron transfer through ligation of a ruthenium complex to
the azurin protein. By differing the distance from a single copper ion,
light-induced oxidation of the copper ion and reduction of the
ruthenium cation could be achieved [17].
In order to mimic the reactions of the PSII donor side, we have
engineered a di-nuclear manganese centre within close proximity of a
light-active chlorin (ZnCe6). We have used a modified form of
bacterioferritin (BFR) from Escherichia coli as the protein scaffold to
bind these cofactors. BFR is a soluble ∼ 18.5 kDa oligomeric protein
which contains both iron and heme b binding sites [18–20]. The
1113
protein forms into a homodimer, binding a single heme group
symmetrically at the interface between the two protein monomeric
subunits via bis-methionine ligation, and each protein monomer
contains a binuclear metal binding site [21,22]. The crystal structure
for BFR was initially produced with two MnII ions bound in the metal
binding site [21,23] (Fig. 1a.). Either Mn or Fe metal ions can
selectively bind at the active enzymatic site. The BFR protein selfassembles to produce a large spherical-shaped shell, made up of 12
homodimeric units, that enclose a hollow cavity with a diameter of
∼8 nm [21,24,25]. The natural protein shell thus binds 12 heme groups
and 48 Fe ions [21] (Fig. 1b.). BFR presents an appealing starting point
for engineering a multi-step, light-activated protein as: (i.) it is a
highly stable protein (ii.) the heme can be extracted and replaced with
a photoactive chlorin [15,26]; (iii.) the binuclear metal site has ligands
similar to manganese catalase where the di-manganese site can reach
an oxidation state of Mn2III,IV [27–30]; and (iv.) the in silico estimated
cofactor separation within the BFR protein is similar in distance to that
found in PSII (Fig. 1c and d) [31].
Here we report the construction of a light-activated metallocatalyst, which can oxidize MnII to MnIII upon illumination with a
tyrosine radical formed during the process. The light-induced
oxidation state changes were characterized by low temperature
perpendicular and parallel mode X-band EPR spectroscopy.
2. Materials and methods
2.1. Cloning and mutagenesis
The bfr gene encoding bacterioferritin was amplified from E. coli
BL21 cells, using PCR primers 5′-GGTATTGAGGGTCGCATGAAAGGTGATACTAAAGTTATAA (Forward) and 5′-AGAGGAGAGTTAGAGCCTC-
Fig. 1. (a) A ribbon diagram of the E. coli bacterioferritin homodimer with two identical subunits each hosting di-nuclear metal centres and forming a heme binding pocket at the
interface of the subunits. Two manganese molecules bind per protein subunit (PDB 1BFR). (b) A diagram of the sphere structure formed by bacterioferritin. The sphere is composed of
twelve homodimers. (c) Schematic showing the estimated distance between the cofactors in the engineered bacterioferritin BFR1 complex. The distances are very similar to that
found in (d) which shows the distance between the cofactors in PSII.
1114
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
ATCAACCTTCTTCGCGGAT (Reverse). Ligation Independent Cloning
(LIC) regions in each primer (underlined) facilitated cloning into the
pET30 Xa/LIC vector (Novagen), for expression in E. coli BL21 cells. Two
external histidines (H46, H112) were changed to arginines using the
Stratagene Qickchange site-directed mutagenesis kit and the following
PCR primers. H46R 5′-AATGATGTGGAGTATCGCGAATCCATTGATGA
H112R 5′-GCCGATAGCGTTCGTGATTACGTCAGCC. This double mutant
(BFR1) was utilized for all measurements carried out in this study.
2.2. Expression, purification
BFR1 was expressed in E. coli cultures with induction using 1 mM
isopropyl-beta-D-thiogalactopyranoside (Sigma). Harvested cells
were pink in colour due to the high expression level of the heme
containing protein. Cells were lysed and the protein was purified from
the supernatant utilizing the His-tag and a nickel sepharose column
(GE Healthcare). The His-tag was cleaved using Factor Xa, and
removed again using a nickel sepharose column.
The purified protein was dark blood red in colour due to the bound
heme and was of single band purity on an overloaded SDS-PAGE gel.
Non-heme iron was removed using the modified method of
Bauminger et al. with exhaustive dialysis in the presence of
dithiothreitol and EDTA [32]. The heme was removed from the
heme binding pocket of the dimeric holoprotein using the method of
Teale et al. [26]. Protein concentrations for the apo-protein were
determined using ɛ280nm of 20340 M− 1 cm− 1. A 1 mM ZnCe6 stock
was prepared by the addition of zinc acetate to free base chlorin e6
dissolved in methanol, added in a 1:1 molar ratio with UV–Vis
confirmation of binding of the zinc to the chlorin [33]. UV–Vis
spectroscopy was carried out on a Cary 300 Spectrometer (Varian,
USA) and CD spectroscopy was carried out on a Jobin Yvon type III+
spectrometer (Horiba, Japan).
2.3. O2 measurements
All oxygen measurements were carried out at 25 °C on an
oxytherm Clarke type oxygen electrode (Hansatech, UK) modified to
contain a side illumination port. Oxygen measurements were carried
out on samples containing 30 μM BFR, 15 μM ZnCe6, 60 μM or 300 μM
MnCl2 or 300 µM FeSO4 in 50 mM MES pH 6.5. Samples were
illuminated for long periods of time until oxygen uptake plateaued.
2.4. Isothermal Titration Calorimetry (ITC)
ITC measurements were carried out on a Microcal calorimeter at
25 °C and the data was fit to a least squares model with the Microcal
Origin fitting software. The heat of dilution was subtracted from the
raw data prior to analysis. Manganese binding to BFR1 was measured
by injecting 11 µl aliquots of 900 μM MnCl2 into a 30 μM solution of
BFR1. BFR1 binding to ZnCe6 was measured by injecting 11 μl aliquots
of 300 μM BFR1 into a 15 μM solution of ZnCe6. All samples were
prepared in 25 mM tricine, 100 mM KCl pH 7.7.
2.5. EPR measurements
Low temperature (5 K) X-band EPR spectra were acquired on a
Bruker ESP 300E spectrometer using either a TM011 cavity or an
ER4116 dual mode cavity, perpendicular mode microwave frequency
9.6 GHz, parallel mode microwave frequency 9.3 GHz, modulation
frequency 100 kHz. The EPR spectrum of each sample was collected in
both parallel and perpendicular modes. To examine light-induced
changes in oxidation state of the metal centre in BFR1 complexes,
samples were prepared in the dark at room temperature then either
left for 1 h in the dark or illuminated for 1 h. The long illumination
time used coincided with the time taken for the oxygen uptake
reactions to go to completion (see previous section). Due to the
Fig. 2. Circular dichroism spectra of the apo forms of BFR and the double mutant BFR1
(H46R, H112R). The BFR and BFR1 had 59% and 60% α-helix respectively, with 7% βsheet and 33% random coil for both proteins.
concentrated nature of the samples they were stirred under pure
oxygen during illumination to prevent the samples becoming
anaerobic. Samples were then degassed and frozen to liquid nitrogen
and stored until use. All samples were measured within 24–48 h.
Samples were prepared in 50 mM MES pH 6.5 or in 25 mM tricine,
100 mM KCl pH 7.7 with 400 mM sucrose added as cryoprotectant.
2.6. EPR simulations/molecular modelling
Spectral simulations were solved numerically from a Hamiltonian
(36 × 36 matrix) using Scilab-4.4.1, an open source vector-based linear
algebra package (www.scilab.org). A least squares minimization
routine was employed to find the optimal solutions for the
parameters. A complete description of the EPR simulations can be
found in the supporting information.
Molecular modelling of mutations was carried out with the aid of
HyPERCHEM (Hypercube, USA) using energy minimization in a
periodic box with the AMBER94 force field. Raytraced images were
prepared using PyMOL v0.99 (Delano Scientific, USA).
3. Results
3.1. Cofactor assembly in bacterioferritin
3.1.1. ZnCe6 binding to BFR1
Previous work has shown that ZnCe6 will bind at the heme site of
cytochrome b562 via axial ligation to histidine [15]. BFR has two
surface exposed histidines at positions 46 and 112. To prevent nonspecific binding these two residues were replaced with arginines. The
secondary structure and binding of ZnCe6 to the double mutant (BFR1)
was identical to the wild type as probed by UV–Vis and Circular
Dichroism (CD) spectroscopy (Fig. 2). The wild type BFR and the BFR1
double mutant contain 59% and 60% α-helix respectively, with 7% βsheet and 33% random coil for both proteins. BFR1 was also found to
still form the spherical ∼ 444 kDa dodecamer of dimers as measured
by native gel electrophoresis. Thus for all the remaining experiments
BFR1 was used. The nomenclature employed throughout this manuscript is as follows: BFR1 (BFR1 devoid of all cofactors), BFR1–M (BFR1
with two Mn ions bound per protein unit, 4 per dimer), BFR1–Z (BFR1
with one ZnCe6 bound per dimer) or BFR1–ZM (BFR1 with four Mn and
one ZnCe6 bound per protein dimer).
BFR1 cofactor binding was monitored using UV–Visible spectroscopy (Figure S1) and isothermal titration calorimetry (ITC)2 (Fig. 3).
Figure S1 shows the spectra of ZnCe6 titrated into a solution BFR1.
2
ITC allows the accurate determination of binding constants (K), reaction
stoichiometries (n), enthalpies (ΔH°), and entropies (ΔS°), thus providing a complete
thermodynamic profile of the molecular interaction.
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
1115
with a binding constant of 2.54 (±0.45) × 106. All parameters are
reported in Table 1.
Fig. 3. Calorimetric titrations of cofactors with BFR1. The top of the figures is the raw ITC
output for the titration and the lower half of the figure is derived from the integrated
raw data. (a) Calorimetric titration of BFR1 binding to ZnCe6. The line represents the
best least squares fit to a model of a single binding site. (b) Calorimetric titration of MnII
binding to BFR1. The line represents the best least squares fit to a model of two
independent binding sites.
Binding ZnCe6 to BFR1 produces a shift in the absorbance maxima of
the QY band of the ZnCe6 from 632 to 639 nm. The association constant
of the ZnCe6 was measured by monitoring this QY shift upon its
titration into a solution of BFR1 (Ka ∼ 1.2 × 106). As a control a BFR
mutant in which the axial heme ligating M52 was converted to H52
was also measured. This mutant correctly assembled and was capable
of oxidizing iron. As expected, no absorbance shift of the chlorine QY
band was observed, suggesting that the ZnCe6 no-longer binds to the
BFR. This observation confirms the lack of heme binding activity of
BFR M52H previously reported by Andrews et al. [18].
Fig. 3a shows the injection heat for BFR1 titrated into ZnCe6 in
dilute solution (15 μM). The integrated heat (μCal) for each injection is
plotted over the molar ratio of protein to ZnCe6 in the bottom of Fig.
3a3. For this the heat of dilution of the protein was subtracted from the
raw data. It was found that 0.82 ZnCe6 molecules bind per BFR1 dimer
3.1.2. Mn binding to BFR1
MnII binding to BFR1 was measured using ITC. Fig. 3b shows the
injection heat for MnII titrated into a solution of BFR1. Analysis of the
integrated heat using the best fit of a least squares model allowing for
two independent binding sites revealed that BFR1 has both a high
affinity Mn site (Ka1 1.54 (±0.39) × 109 M) and a second lower affinity
site (Ka2 4.31 (±1.09) × 106 M) (Table 1). Titrations of Mn into BFR1
showed that binding of the first metal ion to site 1 is an exothermic
process, while binding to the lower affinity site 2 is an endothermic
process (Fig. 3b). According to the ITC fitting an average of 2.5
(±0.12) mol of Mn ions bind per mole of BFR1.
EPR measurements were undertaken to determine the oxidation
state of manganese ions bound to the bacterioferritin protein and their
coupling environment. We have used conventional perpendicular
mode EPR for the observation of half-integer spin systems (e.g. MnII,
and organic radicals) [34,35], and parallel polarization EPR for
observing EPR signals issued from integer spin systems, in particular
S = 2 (e.g. MnIII, MnIII,III
) [36–40].
2
EPR spectra of BFR1 titrated with two Mn ions per protein unit
revealed a broad signal (∼ 400 mT wide), dominated by a large
structured g = 2 derivative (Fig. 4a). The non-Curie temperature
dependence of this signal (Fig. 5a) suggested that it arises from an
even spin antiferromagnetically coupled system, most likely a MnII,II
2
dimer [41–44]. As the line shape of the MnII,II
dimer remained
2
constant for temperatures up to 50 K, the exchange coupling between
the two metal centres was expected to be small.
EPR simulations using the Spin Hamiltonian formalism of the line
shape and temperature dependence of the MnII,II
2 signal were undertaken to estimate the exchange/dipole couplings and zero-field
parameters for this system. These calculations are described in detail
in the supporting information. The exchange coupling was estimated
to be 1.3 K (0.9 cm− 1), the Mn–Mn distance ∼ 4 Å, commensurate with
crystallographic evidence [21]. Both D values for the Mn ions are small
(D b 0.2 K), typical for a MnII high spin (S = 5/2) centre [44–46].
The addition of the ZnCe6 pigment to the BFR1–M complex (i.e.
BFR1–ZM) led to a significant change in the dimer spectrum (Fig. 4b).
The main intensity of the spectrum shifted to g ∼ 2, and appeared less
structured. As before this new signal demonstrated non-Curie
temperature dependence (Fig. 5b) and its line shape was invariant
at all temperatures up to 50 K. It was noted that while the spectrum
about g ∼ 2 is altered, the wings (g ∼ 2.5 etc., now smaller by
approximately a factor of 2) matched the structure seen in the
original MnII dimer spectrum, i.e., without ZnCe6. This suggested that
the spectrum was heterogeneous, made up of both the original MnII
dimer spectrum and a new signal. The estimated contribution of the
original dimer spectrum to the total spectrum observed after the
addition of ZnCe6 was 59%. This suggested that the Zn–chlorin had an
effect on 41% of Mn clusters (i.e. 82% of reaction centres). This value is
Table 1
Parameters of best fit for ITC measurements of BFR1 cofactor binding.
Mn titrated into BFR1 (two independent binding site model)
Mn1
Mn2
Negative peaks correspond to an exothermic reaction and positive peaks
correspond to an endothermic reaction.
Ka (M− 1)
ΔHbind
(kcal mol− 1)
ΔSbind
(kcal mol− 1)
0.88 ± 0.01
1.64 ± 0.11
1.54 (±0.39) × 109
4.31 (±1.09) × 106
− 7380 ± 116
860 ± 87
17.3
33.2
BFR1 titrated into ZnCe6 (single binding site model)
BFR1
3
n/BFR1
n/ZnCe6
Ka (M− 1)
ΔHbind
(kcal mol− 1)
ΔSbind
(kcal mol− 1)
1.64 ± 0.02
2.54 (±0.45) × 106
− 4956 ± 89
12.7
Stoichiometry of the interaction (n), association constant (Ka), enthalpy of binding
(ΔHbind), and entropy of binding (ΔSbind).
1116
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
Fig. 4. CW x-band EPR spectra and simulated spectra for a.) BFR1–M (BFR1 with two Mn
ions bound per subunit) b.) BFR1–ZM (BFR1 complex with two manganese ions and one
ZnCe6 bound per subunit). Protein concentration 400 μM, Mn 800 μM, ZnCe6 200 μM.
Microwave frequency 9.44 GHz, microwave power 2 mW, modulation amplitude 2 mT,
temperature 5 K.
approximately the same as the estimate from the number of bound
Zn–chlorins per Mn cluster as estimated by ITC. The spectrum
generated by the subtraction of the original dimer spectrum appeared
as a ‘simple’ derivative about g ∼ 2. The spectra showed near linear
temperature dependence (Fig. 5c). Subsequent modelling of this
signal using the same formalism as described above suggested that it
could be explained by a substantial drop in the coupling between the
two metal centres, with the exchange component dropping to ∼0.01 K
and the Mn–Mn distance lengthening to ∼ 5 Å.
The binding of ZnCe6 also resulted in the formation of EPR signals
containing hyperfine structure centred at g = 4.5 and g = 9.8 (Fig. 4a).
These signals span from 50–185 mT, and were assigned to monomeric
MnII bound within a highly structured, weak ligand field. The
hyperfine splitting of this signal is approximately 9.5 mT, which is
consistent with MnII coordinated octahedrally with oxygen and
nitrogen ligands, as commonly seen in a number of manganese
enzymes [47–49]. Interestingly, BFR1–ZM samples prepared in the
dark exhibited an unusual multiline EPR signal (Figure S2) This signal
was super-imposed on the broad g = 2 homovalent MnII,II
spectral
2
components observed in the dark. This multiline has 5 mT splitting
and stretches from 290–410 mT, with up to 23 lines present. This
multiline signal is commonly attributed to MnII,II
2 dimer systems [50].
with the concomitant uptake of 0.25 O2 molecules per Fe atom when a
five fold excess of iron was present (Table 2). Both forms oxidize iron
with ratios of oxygen to Fe identical to the literature [51] but the rate
of iron oxidation for the mutant form is slightly higher than for the
wild type. The addition of ZnCe6 to the BFR1 protein did not change the
amount of oxygen taken up nor significantly affect the rate of iron
oxidation in the dark. Illumination of the BFR1–Z protein with iron
bound after iron oxidation had gone to completion produced further
oxygen uptake at a slower rate. Additional light-induced oxygen
uptake totalled approximately the same as dark iron oxygen uptake
(see Fig. 6a, and Table 2).
The BFR1–ZM complex, when measured under the same conditions, did not auto-oxidize Mn in the dark. Interestingly though,
subsequent illumination of this complex did lead to consumption of
oxygen (Fig. 6b–d). Control measurements demonstrated that only a
small amount of oxygen was consumed by the Mn free (BFR1–Z), and
for a protein free solution of ZnCe6 (Fig. 6e and f). Similarly, when
heme was bound to the protein in place of ZnCe6 and manganese
bound at the di-metal site, no oxygen uptake was observed in the dark
or upon illumination (Table 2). Buffered pH 6.5 Mn2+ solutions
showed very little oxygen uptake when illuminated (Fig. 6b). The
oxygen uptake rate of the BFR1–ZM complex was increased by the
addition of superoxide dismutase (SOD) and catalase but the total
amount of oxygen consumed was almost the same (Table 2). This
suggests that SOD and catalase may have a protective function as their
addition should slow ZnCe6 breakdown from reaction with superoxide
or peroxide. No oxygen was released upon dark adaptation. More
oxygen was consumed when a five fold excess of Mn was added
suggesting that more Mn was oxidized. The same amount of oxygen
uptake was found for BFR1–Z with excess iron. Both consumed a total
of ∼ 0.5 mol of oxygen per metal ion.
3.2.2. Light-induced chlorin oxidation/tyrosine oxidation as measured
by EPR
Light-induced oxidation of the bound ZnCe6 was readily observed
by EPR (Fig. 7a). Upon illumination of the BFR1–Z complex (i.e. no Mn
present) with short actinic light flashes at room temperature a
narrow 0.75 mT wide radical centred at g = 2.0022 was generated.
Under the same conditions the BFR1–ZM complex generated a
broader signal (2.5 mT radical) centred at g = 2.0058. This broader
radical resolved a 4–6 peak hyperfine structure with p–p spacing
∼1.5 mT characteristic of an oxidized tyrosine (Fig. 7b) [52–54]. A
corrected spin count for each species revealed that the tyrosine signal
area was approximately 6 times more intense than that of the ZnCe6
radical signal.
3.2. Light-induced oxidation state changes
3.2.1. O2 consumption of the di-metal site of BFR1
Bacterioferritins are members of a class of spherical shell-like iron
storage proteins that contain a ferroxidase site within the protein
which catalyses the oxidation and hydrolysis of iron. The oxidation
reaction involves the uptake of oxygen from solution and as a
consequence the reaction can be monitored using a Clarke type
oxygen electrode. Wild type BFR and BFR1 both oxidized FeII to FeIII
Fig. 5. The temperature dependence of the signals. (a) (red ◊) and b) (blue □) and the
subsequent linear temperature dependence of (a) minus (b) producing (c) (green Δ).
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
1117
Table 2
Oxygen uptake data.
Controls
Dark
Light
BFR1–Mn
Dark
Light
1
BFR –Fe
Dark
Light
Sample
Total oxygen
uptake (μM)
O2 uptake/
metal ion
kmax
(μMol/s)
Fe 300 μM (no protein)
BFR1–Z
ZnCe6 Mn 60 μM (no protein)
BFR1–ZM Mn 60 μM
BFR1 heme Mn 60 μM
BFR1–ZM Mn 60 μM
BFR1–ZM Mn 60 μM
Catalase & SOD
BFR1–ZM Mn 300 μM
BFR1–Z Fe 300 μM
wt-BFR Fe 300 μM
BFR1 Fe 300 μM
BFR1–Z Fe 300 μM
29
39
40
9
7
127
0.10
–
–
0.14
0.12
2.12
0.3
0.3
0.1
0.1
0.1
0.6
133
147
81
81
71
156a (75)b
2.22
0.49
0.27
0.27
0.24
0.52a (0.25)b
1.3
0.7
1.4
0.9
1.3
–a (0.2)b
Note. BFR 30 μM, ZnCe6 15 μM, samples with metal contained either stoichiometric amounts (60 μM) or a 5 fold excess (300 μM). aTotal O2 consumption light and dark cumulative.
b
Additional O2 consumption in light (as compared to dark). Rates are an average over 10 s.
3.2.3. Light-induced Mn2+ oxidation as measured by EPR
BFR1–ZM complexes were prepared containing either stoichiometric (2 Mn2+ per BFR unit) or 5-fold excess Mn2+ (10 Mn2+ per BFR
unit) in the dark. Excess Mn2+ (unbound) appeared as a narrow
100 mT wide 6 line EPR signal centred at g ∼ 2 characteristic of
Mn2+(H2O)6 [55]. This signal overlaid the structured MnII2 dimer
signal discussed above (see Mn Binding to BFR1). The MnII,II
dimer
2
signal was quantitatively the same for both samples.
Upon illumination of the sample containing stoichiometric Mn2+
the structured MnII,II
dimer signal decreased by ∼ 1/3. No new Mn
2
signal was observed in perpendicular or parallel mode EPR. Small
spectral features observed in parallel mode EPR were attributed to
background contaminant O2.
As with stoichiometric samples, illumination of samples that
contained excess Mn2+, led to a reduction of the structured MnII,II
2
dimer by ∼ 1/3. In these samples the additional Mn2+ six-line signal
present was also partially reduced by ∼25% (see Table 3). Concomitant
with the loss of unbound Mn2+ (six-line) was the appearance of broad
parallel polarized signal most likely arising from an even spin system
(Fig. 8). This new signal resolves a hyperfine structure with p–p
spacing of ∼ 15.6 mT. This spacing is larger than previously reported
for monomeric MnIII [37–39].
4. Discussion
Fig. 6. Oxygen uptake measurements carried out on a Clarke type oxygen electrode. The
light was turned on after 60 s for the measurement of all samples except (a) which was
measured in complete darkness and (f) where the light was turned on after oxygen
uptake in the dark had ceased. (a) BZM Mn 60 μM Dark. (b) ZnCe6 Mn 60 μM. (c) BZ. (d)
BZM Mn 60 μM. (e) BZM Mn 300 μM. (f) BZ Fe 300 μM. Protein concentration was
30 μM, ZnCe6 15 μM in 50 mM MES pH 6.5.
We have taken a bioengineering approach to study the lightactivated electron transport in the water oxidation reactions during
oxygenic photosynthesis. Bacterioferritin from E. coli was genetically
and biochemically modified to produce a minimalist model of the PSII
donor side in which analogues of PSII cofactors were bound. These
included a photo-oxidizable chlorin, a redox-active tyrosine, and a dinuclear manganese centre (Fig. 9). This model system demonstrates
both porphyrin excitation and metal complex oxidation.
4.1. Cofactor-protein assembly
Upon titration of the BFR1 into a solution of ZnCe6, ITC analysis
reveals that only 82% of the BFR1 binds ZnCe6 (Table 1). The small
fraction which does not bind may be due to partial aggregation of the
chlorin in solution, even at the low concentrations used, or it may
reflect heterogeneity in the sample; i.e. some of the binding pockets
Table 3
Relative metal ion EPR signal intensities.
Sample
Fig. 7. EPR spectra for the BFR1–Z complex after three flashes of light, without
manganese (a) and with manganese bound (b). Trace (a) reveals a narrow ZnCe6 radical
cation with a peak to peak width of 0.75 mT, centred at g = 2.0022. Trace (b) displays a
broad signal with a width of 2.5 mT, centred at g = 2.0058. Figure is arbitrarily scaled.
Protein concentration 150 μM, ZnCe6 75 μM, Mn 240 μM. Microwave frequency
9.44 GHz, microwave power 1 μW, modulation amplitude 0.3 mT, temperature 5 K.
BFR1ZM Mn 60 μM
BFR1ZM Mn 300 μM
Dark
Light
Structured g = 2
Six-line
Structured g = 2
Six-line
100%
100%
0
400%
66%
65%
0%
300%
Note. BFR 30 μM, ZnCe6 15 μM, metal concentration either stoichiometric amounts
(60 μM) or a 5 fold excess (300 μM) concentration per BFR1 monomer.
1118
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
Fig. 8. Parallel mode EPR spectra of the light minus dark signals of BFR1–ZM with a five
fold excess of Mn. The inset is a higher resolution scan of the multiline signal.
Microwave frequency 9.34 GHz, microwave power 10 mW, modulation amplitude 1 mT,
temperature 5 K. Protein concentration 250 μM, 125 μM ZnCe6, 2500 μM MnCl2 in
50 mM MES pH 6.5.
Fig. 9. Diagram of the amino acid ligands to the di-manganese centre with ZnCe6 bound
at the interface of the BFR dimeric complex. Residues from the A subunit are in blue and
the B subunit are in salmon. The three tyrosines have been labelled on either side of the
ZnCe6 group to show the orientation of the dimer subunits relative to each other. The
alpha helices are pictured as grey rods. The picture is modified from the PDB–1BFR.
are perturbed. The control mutant (M52H) did not bind any
porphyrins (chlorin or heme) possibly due to steric hindrance through
the introduction of two large histidine residues in the heme binding
pocket, preventing dimer formation. ITC analysis also revealed that
binding of ZnCe6 to the hydrophobic heme pocket has a large enthalpy
term (Table 1). This large and negative ΔHb value reflects strong noncovalent interactions (van der Waals and hydrogen bonds) between
ZnCe6 and BFR1 relative to their interaction with the solvent. The main
driving force for binding is most likely the hydrophobic interactions
between the porphyrin ring structure and the hydrophobic amino acid
residues lining the binding pocket.
As we showed earlier manganese ligates to the di-nuclear metal
binding site of bacterioferritin [2] via four glutamate residues (E18,
E51, E94, E127) one from each helix and two histidine residues (H54,
H130) [21]. E51 and E127 form di-μ-1,3-caboxylato bridges, where
each oxygen atom is coordinated to a different metal ion, while E18
and E94 form monodentate carboxylate ligands. H54 and H130
coordinate to the metal ions separately through the Nδ nitrogen (Fig.
9). ITC analysis of Mn binding to BFR1 revealed two distinct binding
sites in the dark with very different association constants. Site 1 bound
Mn very tightly whilst site 2 binding was weaker by almost three
orders of magnitude. Binding of the first Mn ion has a large and
negative ΔHb1 value, which may reflect structural changes in the
protein upon metal ligation [56]. The second Mn binding has a small
and positive ΔHb but a larger ΔSb than site 1 indicating that binding
Mn to site 2 is entropically driven. The most likely contributions to this
positive entropy are changes in the hydration of the protein and of the
metal ion upon binding to the protein [57]. These observations
coincide with previous crystal structure findings for BFR with
manganese bound, where site 2 was found to have a lower incidence
of occupation than site 1 [21].
In comparison, during the photo-assembly of the tetra-nuclear Mn
cluster in PSII, the first MnII ion binds tightly at a high affinity site in the
apo-complex [IM0 intermediate]. Light is then needed to start the
assembly process in which the bound MnII is oxidized to MnIII and a
proton is released [IM1]. The bound MnIII is unstable until a conformational change occurs in the dark and allows the formation of the more
stable MnIII–O–CaII [IM1⁎]. At this point the second MnII binds forming
the stable, spin-coupled cluster MnII–MnIII–O–CaII [IM2]. The formation
of the μ-oxo bridge thus helps stabilize the intermediates until the full
assembly of the tetra-nuclear MnCa cluster [58].
The ITC titrations also showed that an average of 2.51 mol of Mn
ions bound per mole of BFR1 monomer. This is higher than expected
but can be partly explained by the findings from the original crystal
structure where an extraneous Mn ion was shown to be bound at the
four-fold axis of the dodecamer of dimers by four Gln151 residues
internal to the structure [23]. This would account for a further 0.25
binding sites per mole of protein but still leaving 0.25 sites
unaccounted for. Nonspecific, low affinity binding sites have also
been detected in previous studies of the BFR protein. It is proposed
that these sites involve acidic residues located on the inner surface of
the protein shell [59]. Under the conditions of these experiments the
individual thermodynamic parameters of the weak binding sites
cannot be determined. This is due to the limiting form of the equation
for two classes of binding sites [60].
We characterized the oxidation states of the Mn centre by EPR
measurements. Simulation of dark Mn EPR signals strongly suggests
that manganese bound to the BFR1 protein remained in the 2+
oxidation state and was present as a weakly spin-coupled MnII,II
2
centre. The estimated Mn–Mn distance from EPR simulations was
∼4 Å, commensurate with the crystal structure [21]. The addition of
ZnCe6 appeared to alter approximately 50% of the MnII,II
complexes.
2
This behaviour may be rationalized as follows. Initially the homodimer
has two identical Mn clusters. Upon addition of ZnCe6 this symmetry
is lost. The BFR protein was engineered to bind ZnCe6 between the two
monomeric BFR1 subunits via a methionine residue. Only one
monomer of the dimer subunits provides this ligand (the 5th axial
coordinate of the ZnCe6). The methionine residue is one amino acid
from the crucial glutamic acid residue (E51) which ligates to both Mn
ions in the metal centre. It is possible that the binding of the ZnCe6
disrupts the coupling through movement of the methionine and thus
the glutamic acid residue producing a carboxylate shift (presumably
moving the two metal centres further apart).
4.2. O2 consumption
BFR functions in biology to bind iron and oxidize FeII2 to FeIII
2 . This
process involves the formation of a stabilizing μ-oxo bond between
the two metal ions. This reaction consumes dissolved O2 [61]. The
three phases to this process involve (i.) metal binding and deprotonation of amino acid ligands followed by (ii.) oxidation of bound metal
ions with the formation of a single μ-oxo bond between the metal ions
and uptake of one oxygen atom per centre. Once all ferroxidase iron
binding sites are filled, and all the 48 iron atoms bound per dodecamer
of dimers are oxidized, the complex then begins to oxidize excess FeII
within the ball and store it as ferric-oxy-hydroxide (iii.) [22,51,62].
+
i.) 2FeII + BFR → FeII,II
2 –BFR + 4H
III
II,II
ii.) Fe2 –BFR + ½O2 → Fe − O − FeIII–BFR
iii.) 4FeII + O2 + 6H20 → 4FeO(OH)(core) + 8H+
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
The oxygen assay of the BFR1 protein (Fig. 6) with iron present
revealed that 0.25 O2 molecules are taken up per iron oxidized. This
agrees with the literature values for the wild type and the ZnCe6
bound form, supporting the assumption that binding of the ZnCe6
does not change the basic function of the protein. Minimal oxygen
consumption is observed for BFR1–ZM complex suggesting it cannot
auto-oxidize MnII to MnIII in the dark.
Upon illumination the BFR1–ZM sample consumes ∼2 O2 per Mn.
This value is 4 times larger than for the BFR with Fe bound and may
suggest that the di-Mn site constructs multiple μ-oxo bridges as
compared to when Fe is bound in the wild type enzyme and only a
single μ-oxo bridge is formed. Control samples – samples that lack Mn
i.e. BFR1–Z – did not consume a significant amount of oxygen
confirming that O2 is reacting with the Mn. In samples with a five
fold excess of manganese, a significant increase in O2 consumption
was observed (∼ 20 μM). This likely indicates that ‘free’ Mn2+ is
oxidized within the ball structure forming MnIII-oxy-hydroxide. For
MnIII oxy-hydroxide formation to occur multiple oxidation events are
likely occurring at the manganese binding site. The addition of
catalase and SOD was found to increase the rate of oxygen
consumption but did not have any significant effect on the total
amount consumed4. We suggest that it acts to protect the chlorin,
removing super-oxides and other reactive oxygen species.
4.3. Tyrosine oxidation
In the absence of manganese, illumination of the BFR1–Z produces
a large ZnCe•6+ radical cation signal at g = 2.0022. In contrast, the
addition of Mn results in the loss of the ZnCe•6+ signal and the
appearance of a ‘tyrosine like’ signal. This new radical signal is unlikely
to be the ZnCe•6+ radical broadened due to through bond/through
space interactions with the Mn site as the distance between the Mn
centre and the ZnCe6 is large (∼14 Å). The intensity of the tyrosine like
signal is significantly in excess of the ZnCe6 (6 fold). This is
unsurprising as tyrosine radicals are generally much longer lived
than chlorin radicals [63,64]. Here it is suspected that in the time
between room temperature illumination and freezing the sample
down to 77 K (in the dark), a significant fraction of the chlorin radical
decays away. The most likely mechanism for tyrosine oxidation is
electron hole migration from the oxidized ZnCe6 •+ radical to a nearby
tyrosine residue. ZnCe6 has a high estimated oxidation potential of
∼ +1.1 V [15] which is sufficient to oxidize a tyrosine. It is unclear why
tyrosine oxidation does not occur when no manganese ions are bound.
When Mn (or any metal) binds to the BFR metal site deprotonation of
the metal ligands occurs along with structural changes to the ligands
themselves. These changes may alter the hydrogen bonding network
around the tyrosine allowing proton coupled electron transfer.
There are three tyrosine residues in BFR (Y25, Y58, and Y45) that
are close enough to the ZnCe6 (10.6 Å, 11.0 Å, and 4.2 Å respectively)
that they could potentially act as ‘efficient’ electron donors (Fig. 9). Of
these only two (Y25 and Y58) are found at the metal binding site and
both are involved in the hydrogen bonding network of the site.
Tyrosine 58 is not highly conserved and is commonly replaced by a
leucine in other bacterioferritins. Tyrosine 25 is hydrogen bonded to
glutamate 94 which is one of the metal ligands with the oxygen atom
of this tyrosine placed 4.3 Å from the nearest Mn ion. Ferritins, which
have very similar structure and function to bacterioferritin but do not
bind heme, have a redox-active tyrosine within the hydrogen bonding
network of the metal binding site. This residue is highly conserved in
ferritins and bacterioferritins [65]. As a consequence, we suggest that
tyrosine (Y25) is likely to be responsible for the tyrosine like radical in
our system. This is currently being tested in our lab.
4
Even though this is the case it is possible that oxygen acts as the electron acceptor
producing hydrogen peroxide.
1119
4.4. Mn2+ oxidation
The loss of MnII signals upon illumination of BFR1–ZM strongly
suggests that Mn oxidation occurs in this system. The broad structured
Mn2 II,II signal centred at g = 2 was found to decrease in BFR1–ZM
samples upon illumination. This structured Mn2 II,II signal decreased
on average by 34%.
No new Mn EPR signals were observed upon loss of the MnII signal
in BFR1–ZM. We would expect the one electron oxidation of an Mn2 II,II
to yield a mixed valence Mn2 II,III dimer. Strongly coupled mixed
valance dimers (J ≈ 10 cm− 1) display characteristic ‘multiline’ signals
seen in perpendicular mode EPR centred at g ∼ 2 with hyperfine
splittings of b90 G [30]. In contrast, the weakly coupled mixed valance
dimer should still have broader perpendicular signals and could
possibly resolve parallel polarization signals [30,66]. As neither of
these is seen, it is suggested the Mn cluster undergoes two electron
oxidation to an even spin MnIII,III
system. It is not clear as to whether
2
both proposed MnII oxidation events need to be photo-driven. As no
intermediate state (MnII,III
2 ) was observed, the second oxidation state
may not require light excitation of the chlorin. Alternatively the mixed
valance cluster may be more strongly oxidizing (electron donating) to
the oxidized tyrosine/chlorin than the MnII,II
2 precursor state. Oxygen
bridged MnIII,III
dimers with analogous ligand fields have been
2
observed at approximately zero-field (g = 20) [38]. We cannot
discount the presence of these signals in our illuminated BFR complex.
The BFR1–ZM sample with a five fold excess of Mn revealed the
same decrease in amplitude of the structured g = 2 MnII,II
2 signal upon
illumination as seen in samples with stoichiometric Mn. The six-line
signal also showed a significant decrease upon illumination. This
signal decreased by 25%, equivalent to 4 Mn oxidations per BFR dimer
(48 Mn ions per dodecamer of dimers). Parallel polarization EPR on
illuminated samples with excess Mn suggested the formation of a
manganese species with even spin character, upon illumination of the
BFR1–ZM complex; a broad signal centred at geff ∼ 4–5 with clear
hyperfine structure (Fig. 8). As this signal is not observed in
illuminated samples with stoichiometric Mn, it is suggested that this
signal comes from a new Mn species unlikely to be bound to BFR.
Comparing this to the native function of ferritins, it is tempting to
assign the species to MnIII-oxy-hydroxide (MnIIIOOH) which is
accumulated in the core of the sphere structure [67].
MnIII is a 3d4 (S = 2) transition ion [68]. There are potentially three
geometries that MnIII complexes in this protein ligand field can take: 6
coordinate octahedral, 5 coordinate square-pyramidal or trigonal–
bipyramidal [39]. The ground electronic state of the MnIII manifold
depending on the geometry of the ligand field can have two different
3d4 configurations corresponding to the electronic hole position. The
hole either occupies the dx2–y2 orbital (ground state 5B1) or the dz2
orbital (ground state 5A1). The hyperfine splitting for each of these
states is different due to the effect of spin–orbit coupling [39,69–71]. A
ground state of 5B1 yields a hyperfine splitting along AZ of ∼50 G
whereas a ground state of 5A1 yields a hyperfine splitting of ∼100 G
[39,70,72]. In this instance the sign of the Spin Hamiltonian parameter
D indicates the ground state of the manifold. A positive D value yields
an 5A1 ground state (as observed for the MnIII centre of the SOD
protein) whereas a negative D values yields a 5B1 ground state (as
observed for the MnIII photo-assembly intermediate of PS II)
[37,39,71,73].
Recently parallel polarization EPR signals have been reported for
MnIII peroxo complexes (i.e. MnIIIOOH) [66]. These signals were
generated by the oxidation of (L)MnII (where L = N-methyl-tris(2pyridylmethyl)ethan-1,2-diamine) using a 1000 fold excess of H2O2.
Two parallel polarization signals were observed: a structured signal at
g ∼ 8 with hyperfine splitting ∼ 50 G (analogous to MnIII signals
described above [39,40,72]) and a less structured signal at g ∼ 5 with
hyperfine splitting ∼ 95 G. It was noted that the addition of base
(triethyl-amine) led to the loss of the high field signal (g ∼ 5).
1120
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
Conversely the addition of acid (HClO4) leads to the loss of the low
field signal (g ∼ 8). As a consequence the low field signal (g ∼ 8) was
assigned to the monocationic species [LMnIII(OO)]+. Both UV–Vis and
ESI-MS supported this assignment. XSophie simulations estimated the
zero-field and hyperfine parameters of this complex to be: D =
−2.9 cm− 1, E/D = 0.075, AZ = 6.5 × 10− 3 cm− 1 [66]. Curiously, the
second signal (g ∼ 5) was not assigned to the corresponding protonated species [LMnIIIOOH]2+. It was noted that though the signal
could be modelled as arising from a MnIII signal (D = + 2.9 cm− 1,
E/D = 0.111), the resolved hyperfine structure (splittings of ∼95 G)
was inconsistent with assignment to a MnIII [66]. From the above
discussion this argument does not seem valid; Mn III can
potentially resolve hyperfine splitting of the order of ∼ 100 G.
We note that the D value changes sign for these two complexes,
which should reflect a re-ordering of electronic state manifold
(ground state interchanges from 5B1 to 5A1) as seen between MnIII
photo-assembly intermediate of PS II and Mn-SOD protein, leading
to an apparent increase in hyperfine splitting observed [37,39].
Our new Mn EPR signal observed in BFR bears a resemblance to the
g ∼ 5 signal seen in the above study [66] appearing at approximately
the same g position (g ∼ 4–5). It too has a large hyperfine spacing as
seen for the potential [LMnIIIOOH]2+ species. We do acknowledge
though that the hyperfine spacing seen (∼150 G) in the ‘new Mn EPR
signal’ is exceedingly large for a monomeric MnIII. The unusually large
hyperfine splitting may reflect the unique environment Mn centres
occupy in the core of the sphere protein structure (Mn–Mn
interactions etc.). Alternatively, the new EPR signal may represent a
di-Mn species, possibly a weakly coupled mixed valance dimer (i.e.
MnIIMnIII).
[20]
5. Conclusion
[21]
[6]
[7]
[8]
[9]
[10]
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
[19]
In this study we have shown that the modified BFR1–ZM complex
is capable of converting light energy into the stored higher oxidation
states of Mn. To our knowledge this work is the first example reported
in the literature of a bioengineered protein capable of photo-catalytic,
multi-electron oxidation of a di-manganese centre. This represents
one of the first steps in developing synthetic photo-catalytic, ‘green
enzymes’ that utilize light energy to catalyse oxygen evolution and
hydrogen production.
[22]
Acknowledgements
[27]
BC was supported by an ANU Postgraduate Scholarship. Funding
was supplied by ARC Grant DP0450421, by the Deutsche Forschungsgemeinschaft (Me1629/2-4), Solar-H2, BioH2, and the Max-PlanckGesellschaft which is gratefully acknowledged. PLD acknowledges
support from the US Department of Energy grant DE-FG0205ER46223. The authors would like to thank G.C. Dismukes for his
helpful discussions.
[23]
[24]
[25]
[26]
[28]
[29]
[30]
Appendix A. Supplementary data
[31]
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.bbabio.2009.04.011.
[32]
[33]
References
[34]
[1] T. Wydrzynski, K. Satoh, Photosystem II: The Light-Driven Water:Plastoquinone
Oxidoreductase, Springer, Dordrecht, 2005.
[2] B. Conlan, Designing Photosystem II: molecular engineering of photo-catalytic
proteins, Photosyn. Res. 98 (2008) 687–700.
[3] B.M. Discher, D. Noy, K.S. Reddy, S.X. Ye, J.D. Lear, C.C. Moser, P.L. Dutton, Selfassembly of membrane peptides and redox cofactors, Biophys. J. 84 (2003) 177A.
[4] B.R. Gibney, P.L. Dutton, Histidine placement in de novo-designed heme proteins,
Protein Sci. 8 (1999) 1888–1898.
[5] D.A. Moffet, M.A. Case, J.C. House, K. Vogel, R.D. Williams, T.G. Spiro, G.L.
McLendon, M.H. Hecht, Carbon monoxide binding by de novo heme proteins
[35]
[36]
[37]
derived from designed combinatorial libraries, J. Am. Chem. Soc. 123 (2001)
2109–2115.
W.F. DeGrado, Molecular design as an approach to understanding membrane
protein structure and function, FASEB J. 13 (1999) A1431.
C. Moser, J.M. Keske, K. Warncke, R.S. Farid, P.L. Dutton, Nature of biological
electron-transfer, Nature 355 (1992) 796–802.
C.C. Page, C.C. Moser, X.X. Chen, P.L. Dutton, Natural engineering principles of
electron tunnelling in biological oxidation–reduction, Nature 402 (1999) 47–52.
H.B. Gray, J.R. Winkler, Electron transfer in proteins, Annu. Rev. Biochem. 65
(1996) 537–561.
J.R. Winkler, A.J. Di Bilio, N.A. Farrow, J.H. Richards, H.B. Gray, Electron tunneling in
biological molecules, Pure. Appl. Chem. 71 (1999) 1753–1764.
J.R. Winkler, H.B. Gray, Electron tunneling in proteins: role of the intervening
medium, J. Biol. Inorg. Chem. 2 (1997) 399–404.
D. Noy, C.C. Moser, P.L. Dutton, Design and engineering of photosynthetic lightharvesting and electron transfer using length, time, and energy scales, Biochim.
Biophys. Acta 1757 (2006) 90–105.
C.C. Page, C.C. Moser, P.L. Dutton, Mechanism for electron transfer within and
between proteins, Curr. Opin. Chem. Biol. 7 (2003) 551–556.
T. Wydrzynski, W. Hillier, B. Conlan, Engineering model proteins for Photosystem
II function, Photosyn. Res. 94 (2007) 225–233.
S. Hay, B.B. Wallace, T.A. Smith, K.P. Ghiggino, T. Wydrzynski, Protein engineering
of cytochrome b562 for quinone binding and light-induced electrons transfer, Proc.
Natl. Acad. Sci. U. S. A. 101 (2004) 17675–17680.
M. Thielges, G. Uyeda, A. Camara-Artigas, L. Kalman, J.C. Williams, J.P. Allen,
Design of a redox-linked active metal site: manganese bound to bacterial
reaction centers at a site resembling that of Photosystem II, Biochemistry 44
(2005) 7389–7394.
H.B. Gray, J.R. Winkler, Electron tunneling through proteins, Q. Rev. Biophys. 36
(2003) 341–372.
S.C. Andrews, N.E. Lebrun, V. Barynin, A.J. Thomson, G.R. Moore, J.R. Guest, P.M.
Harrison, Site-directed replacement of the coaxial heme ligands of bacterioferritin
generates heme-free variants, J. Biol. Chem. 270 (1995) 23268–23274.
S.C. Andrews, J.M.A. Smith, C. Hawkins, J.M. Williams, P.M. Harrison, J.R. Guest,
Overproduction, purification and characterization of the bacterioferritin of
Escherichia coli and a C-terminally extended variant, Eur. J. Biochem. 213 (1993)
329–338.
G.D. Davis, C. Elisee, D.M. Newham, R.G. Harrison, New fusion protein systems
designed to give soluble expression in Escherichia coli, Biotechnol. Bioeng. 65
(1999) 382–388.
F. Frolow, A.J. Kalb, J. Yariv, Structure of a unique twofold symmetrical hemebinding site, Nat. Struct. Biol. 1 (1994) 453–460.
N.E. LeBrun, S.C. Andrews, J.R. Guest, P.M. Harrison, G.R. Moore, A.J. Thomson,
Identification of the ferroxidase center of Escherichia coli bacterioferritin,
Biochem. J. 312 (1995) 385–392.
A. Dautant, J.B. Meyer, J. Yariv, G. Precigoux, R.M. Sweet, A.J. Kalb, F. Frolow,
Structure of a monoclinic crystal form of cytochrome b1 (bacterioferritin) from E.
coli, Acta Crystallogr. D. 54 (1998) 16–24.
P.M. Harrison, P. Arosio, Ferritins: molecular properties, iron storage function and
cellular regulation, Biochim. Biophys. Acta 1275 (1996) 161–203.
G.R. Moore, Bacterial 4-alpha-helical bundle cytochromes, Biochim. Biophys. Acta
1058 (1991) 38–41.
F.W.J. Teale, Cleavage of the haem-protein link by acid methylethylketone,
Biochim. Biophys. Acta 35 (1959) 543.
V.V. Barynin, M.M. Whittaker, S.V. Antonyuk, V.S. Lamzin, P.M. Harrison, P.J.
Artymiuk, J.W. Whittaker, Crystal structure of manganese catalase from Lactobacillus plantarum, Structure 9 (2001) 725–738.
D.W. Yoder, J. Hwang, J.E. Penner-Hahn, Chapter 16: Manganese Catalases, in: H.S.
Sigel, A. (Eds.), Manganese and Its Role in Biological Processes, vol. 37, Marcell
Dekker, Inc., New York, 2000, pp. 527–557.
K.O. Schafer, R. Bittl, F. Lendzian, V. Barynin, T. Weyhermuller, K. Wieghardt, W.
Lubitz, MultifrequencyEPR , investigation of dimanganese catalase and related Mn
(III)Mn(IV) complexes, J. Phys. Chem., B. 107 (2003) 1242–1250.
C. Teutloff, K.O. Schafer, S. Sinnecker, V. Barynin, R. Bittl, K. Wieghardt, F. Lendzian,
W. Lubitz, High-field EPR investigations of Mn(III)Mn(IV) and Mn(II)Mn(III)
states of dimanganese catalase and related model systems, Magn. Reson. Chem.
43. Spec. no. (2005) S51–64.
B. Loll, J. Kern, W. Saenger, A. Zouni, J. Biesiadka, Towards complete cofactor
arrangement in the 3.0 angstrom resolution structure of Photosystem II, Nature
438 (2005) 1040–1044.
E.R. Bauminger, P.M. Harrison, D. Hechel, I. Nowik, A. Treffry, Mossbauer
spectroscopic investigation of structure–function relations in ferritins, Biochim.
Biophys. Acta 1118 (1991) 48–58.
A.R. Razeghifard, T. Wydrzynski, Binding of Zn–chlorin to a synthetic four-helix
bundle peptide through histidine ligation, Biochemistry 42 (2003) 1024–1030.
M.P. Hendrich, P.G. Debrunner, Electron-paramagnetic-res spectra of quintet
ferrous myoglobin and a model heme compound, J. Magn. Reson. 78 (1988)
133–141.
M.P. Hendrich, P.G. Debrunner, Integer-spin electron-paramagnetic resonance of
iron proteins, Biophys. J. 56 (1989) 489–506.
S.L. Dexheimer, J.W. Gohdes, M.K. Chan, K.S. Hagen, W.H. Armstrong, M.P. Klein,
Detection of EPR-spectra in S = 2 states of trivalent manganese complexes, J. Am.
Chem. Soc. 111 (1989) 8923–8925.
K.A. Campbell, E. Yikilmaz, C.V. Grant, W. Gregor, A.F. Miller, R.D. Britt, Parallel
polarizationEPR , characterization of the Mn(III) center of oxidized manganese
superoxide dismutase, J. Am. Chem. Soc. 121 (1999) 4714–4715.
B. Conlan et al. / Biochimica et Biophysica Acta 1787 (2009) 1112–1121
[38] M.M. Whittaker, V.V. Barynin, T. Igarashi, J.W. Whittaker, Outer sphere mutagenesis of Lactobacillus plantarum manganese catalase disrupts the cluster core —
mechanistic implications, Eur. J. Biochem. 270 (2003) 1102–1116.
[39] K.A. Campbell, D.A. Force, P.J. Nixon, F. Dole, B.A. Diner, R.D. Britt, Dual-mode EPR
detects the initial intermediate in photoassembly of the Photosystem II Mn
cluster: the influence of amino acid residue 170 of the D1 polypeptide on Mn
coordination, J. Am. Chem. Soc. 122 (2000) 3754–3761.
[40] K.A. Campbell, M.R. Lashley, J.K. Wyatt, M.H. Nantz, R.D. Britt, Dual-modeEPR ,
Study of Mn(III) salen and the Mn(III) salen-catalyzed epoxidation of cis-betamethylstyrene, J. Am. Chem. Soc. 123 (2001) 5710–5719.
[41] T. Howard, J. Telser, V.J. DeRose, An electron paramagnetic resonance study of Mn2
(H2O)(OAc)4(tmeda)2 (tmeda = N,N,N′,N′-Tetramethylethylenediamine): a
model for dinuclear manganese enzyme active sites, Inorg. Chem. 39 (2000)
3379–3385.
[42] S.V. Khangulov, V.V. Barynin, N.V. Voevodskaya, A.I. Grebenko, ESR spectroscopy
of the binuclear cluster of manganese ions in the active center of Mn-catalase from
Thermus thermophilus, Biochim. Biophys. Acta 1020 (1990) 305–310.
[43] P.J. Pessiki, S.V. Khangulov, D.M. Ho, G.C. Dismukes, Structural and functional
models of the dimanganese catalase enzymes. 2. Structure, electrochemical,
redox, and EPR properties, J. Am. Chem. Soc. 116 (1994) 891–897.
[44] B. Epel, K.O. Schafer, A. Quentmeier, C. Friedrich, W. Lubitz, Multifrequency EPR
analysis of the dimanganese cluster of the putative sulfate thiohydrolase SoxB of
Paracoccus pantotrophus, J. Biol. Inorg. Chem. 10 (2005) 636–642.
[45] G.C. Dismukes, Manganese enzymes with binuclear active sites, Chem. Rev. 96
(1996) 2909–2926.
[46] A.P. Golombek, M.P. Hendrich, Quantitative analysis of dinuclear manganese(II)
EPR spectra, J. Magn. Reson. 165 (2003) 33–48.
[47] W.C. McGregor, S.I. Swierczek, B. Bennett, R.C. Holz, Characterization of the
catalytically active Mn(II)-loaded argE-encoded N-acetyl-L-ornithine deacetylase
from Escherichia coli, J. Biol. Inorg. Chem. 12 (2007) 603–613.
[48] M.R. Schaab, B.M. Barney, W.A. Francisco, Kinetic and spectroscopic studies on the
quercetin 2,3-dioxygenase from Bacillus subtilis, Biochemistry 45 (2006)
1009–1016.
[49] A.K. Whiting, Y.R. Boldt, M.P. Hendrich, L.P. Wackett, L. Que, Manganese(II)dependent extradiol-cleaving catechol dioxygenase from Archrobacter globiformis
CM-2, Biochemistry 35 (1996) 160–170.
[50] A.J. Wu, J.E. Penner-Hahn, V.L. Pecoraro, Structural, spectroscopic, and reactivity
models for the manganese catalases, Chem. Rev. 104 (2004) 903–938.
[51] X. Yang, N.E. Le Brun, A.J. Thomson, G.R. Moore, N.D. Chasteen, The iron oxidation
and hydrolysis chemistry of Escherichia coli bacterioferritin, Biochemistry 39
(2000) 4915–4923.
[52] G. Bleifuss, M. Kolberg, S. Potsch, W. Hofbauer, R. Bittl, W. Lubitz, A. Graslund, G.
Lassmann, F. Lendzian, Tryptophan and tyrosine radicals in ribonucleotide
reductase: a comparative high-field EPR study at 94 GHz, Biochemistry 40
(2001) 15362–15368.
[53] W. Hofbauer, A. Zouni, R. Bittl, J. Kern, P. Orth, F. Lendzian, P. Fromme, H.T. Witt, W.
Lubitz, Photosystem II single crystals studied by EPR spectroscopy at 94 GHz: the
tyrosine radical Y–D(center dot), Proc. Natl. Acad. Sci. U. S. A. 98 (2001)
6623–6628.
[54] S. Un, A. Boussac, M. Sugiura, Characterization of the tyrosine-Z radical and its
environment in the spin-coupled S(2)Tyr(Z)(center dot) state of Photosystem II
from Thermosynechococcus elongatus, Biochemistry 46 (2007) 3138–3150.
1121
[55] C.W. Hoganson, D.F. Ghanotakis, G.T. Babcock, C.F. Yocum, Mn-2+ reduces Y+
Z in
manganese-depleted Photosystem-II preparations, Photosyn. Res. 22 (1989) 285–293.
[56] N.E. LeBrun, A.M. Keech, M.R. Mauk, A.G. Mauk, S.C. Andrews, A.J. Thomson, G.R.
Moore, Charge compensated binding of divalent metals to bacterioferritin: H+
release associated with cobalt(II) and zinc(II) binding at dinuclear metal sites,
FEBS Lett. 397 (1996) 159–163.
[57] F. Bou-Abdallah, P. Arosio, S. Levi, C. Janus-Chandler, N.D. Chasteen, Defining metal
ion inhibitor interactions with recombinant human H- and L-chain ferritins and
site-directed variants: an isothermal titration calorimetry study, J. Biol. Inorg.
Chem. 8 (2003) 489–497.
[58] J. Dasgupta, G.M. Ananyev, G.C. Dismukes, Photoassembly of the water-oxidizing
complex in Photosystem II, Coord. Chem. Rev. 252 (2008) 347–360.
[59] S. Baaghil, A.J. Thomson, G.R. Moore, N.E. Le Brun, Studies of copper(II)-binding to
bacterioferritin and its effect on iron(II) oxidation, J. Chem. Soc. Dalton (2002)
811–818.
[60] E. Freire, O.L. Mayorga, M. Straume, Isothermal titration calorimetry, Anal. Chem.
62 (1990) A950–A959.
[61] H.L. Liu, H.N. Zhou, W.M. Xing, H.F. Zhao, S.X. Li, J.F. Huang, R.C. Bi, 2.6 Angstrom
resolution crystal structure of the bacterioferritin from Azotobacter vinelandii,
FEBS Lett. 573 (2004) 93–98.
[62] S. Baaghil, A. Lewin, G.R. Moore, N.E. Le Brun, Core formation in Escherichia coli
bacterioferritin requires a functional, Biochemistry 42 (2003) 14047–14056.
[63] X.S. Tang, M. Zheng, D.A. Chisholm, G.C. Dismukes, B.A. Diner, Investigation of
the differences in the local protein environments surrounding tyrosine radicals
Y–Z(center dot) and Y–D(center dot) in Photosystem II using wild-type and the
D2-Tyr160Phe mutant of Synechocystis 6803, Biochemistry 35 (1996)
1475–1484.
[64] Y. Miloslavina, M. Szczepaniak, M.G. Muller, J. Sander, M. Nowaczyk, M. Rogner,
A.R. Holzwarth, Charge separation kinetics in intact Photosystem II core
particles is trap-limited. A picosecond fluorescence study, Biochemistry 45
(2006) 2436–2442.
[65] M.A. Carrondo, Ferritins, iron uptake and storage from the bacterioferritin
viewpoint, EMBO J. 22 (2003) 1959–1968.
[66] S. Groni, P. Dorlet, G. Blain, S. Bourcier, R. Guillot, E. Anxolabehere-Mallart,
Reactivity of an aminopyridine [LMnII](2+) complex with H2O2. Detection of
intermediates at low temperature, Inorg. Chem. 47 (2008) 3166–3172.
[67] B. Zhang, J.N. Harb, R.C. Davis, J.W. Kim, S.H. Chu, S. Choi, T. Miller, G.D. Watt, Kinetic
and thermodynamic characterization of the cobalt and manganese oxyhydroxide
cores formed in horse spleen ferritin, Inorg. Chem. 44 (2005) 3738–3745.
[68] A. Abragam, B.I. Bleaney, Electron Paramagnetic Resonance of Transition Ions,
Oxford University Press, Oxford, 1970.
[69] S.A. Al'tshuler, B.M. Kozyrev, Electron Paramagnetic Resonance in Compounds of
Transition Elements, 2nd ed. Halsted-Wiley, New York, NY, USA, 1974.
[70] H.J. Gerritsen, E.S. Sabisky, Paramagnetic resonance of trivalent manganese in
rutile (TiO2), Phys. Rev. 132 (1963) 1507–1512.
[71] J.S. Griffith, Theory of Transition: Metal Ions, Cambridge University Press, 1971.
[72] A.M. Tyryshkin, R.K. Watt, S.V. Baranov, J. Dasgupta, M.P. Hendrich, G.C. Dismukes,
Spectroscopic evidence for Ca2+ involvement in the assembly of the Mn4Ca
cluster in the photosynthetic water-oxidizing complex, Biochemistry 45 (2006)
12876–12889.
[73] J.W. Whittaker, M.M. Whittaker, Active-site spectral studies on manganese
superoxide-dismutase, J. Am. Chem. Soc. 113 (1991) 5528–5540.
SUPPORTING INFORMATION
The general Hamiltonian used for the MnII dimer takes the form:
H  J  S1  S2  S1  D  S2   H  g i  Si  Si  D ZFi  Si  Si  A i  I 
Where:
Eq.1
J
exchange tensor (assumed to be isotropic)
D
the dipolar tensor (fixed by the geometry of the system i.e. the Mn-Mn
inter-distance)
DZFi
zero-field tensors
Ai
Hyperfine tensors
The orientation of the two zero-field tenors, dipolar tensor, etc., were assumed to be colinear.
The hyperfine component was approximated by an isotropic line-broadening. The resultant
Hamiltonian (36x36 matrix) was solved numerically using Scilab-4.4.1, an open source
vector-based linear algebra package available from www.scilab.org. This package can be
implemented on most computer platforms. Typical powder pattern averages used 10-100 
angles (defined below). A least squares minimization routine was employed to find the
optimal solutions for the above parameters.
Powder Pattern Averaging
The paramagnetic centers of a frozen ESR sample take all possible orientations (relative to
the field axis), often referred to as a powder pattern. The simulation code allows for this
effect by using combination of Euler rotations ( and ). This system only requires two
Euler rotations to define all unique orientations as rotation about the field axis does not alter
the eigenvalues of the system.
A generalized powder pattern code was adopted (shown below). The typical number of 
angles used (n) ranged from 20-100. Dummy variables x and y were used to uniformly
sample the hemisphere. For a given x () angle 1 to p(x+1) y () angles were sampled (with
the entries of p(x) as defined below). Note: for x =0 (,  was also set to zero. 
For x = 0 to n-1
For y = 1 to p(x+1)




360
  90
 
p( x  1)  
  sin  (n  1 2)   x 
90

 

 (n  1 2) 


 90 

  x 
n 1 
2



 360 

  y  1  
2  p( x  1) 
 and  angles define the rotation matrix as below:
cos()  cos() sin()  cos()  cos()
R  
sin()
cos()
0 

cos()
sin()  sin() cos() 
The unitary transformation T *  R  T  R ' was applied to the tensors:
D*  R  D  R '
g Fe*  R  g Fe  R '
J*  R  J  R '
Transition Intensities
The intensity of transition between the ith and jth state is given the equation below:
I  i  H1  g Fe  S Fe  g Q  H1  SQ j
2
Where the oscillating H1 field lies in the x-y plane (perpendicular to H). All of its possible
orientations must be included to correctly estimate transition intensities (requires all three
Euler angles). This is achieved by averaging over the third Euler angle (). H1 can be
expressed as:
cos 
H1   sin  
 0 
hence
H 2 2
cos     sin     j
I i
2
2
where




  g yx  ig yy  S   g yx  ig yy  S   2g yzS z
  g xx  ig xy  S   g xx  ig xy  S   2g xz S z
Integrating with respect to  yields:
2
2
0 (  cos(  )    sin(  )) d
 02
2
2
 (1  cos(2  )) 
 (1  cos(2  ))   sin( 2  )d
2
2
2
2


(   1 sin( 2  )) 
(   1 sin( 2  )) 
cos(2  )
2
2
2
2
2
2
2
0
2
 (   )
The transition intensity can therefore be expressed as:


H 2 2  2   2
I i
j
2
2
Simulation parameters for the MnII dimers
Mn2II,II dimer
Mn2II,II dimer
(structured no ZnCe6)
(unstructured with ZnCe6)
J
1.3 K
~0.001
r
3.95 Å
5.30 Å
D1
0.15 K
0.15 K1
E1/D1
0.33
0.33
D2
0.02 K
0.02 K4
E2/D2
0.12
0.14
width
280 G
280 G
D and E values are assumed to be the same for the simulation of the unstructured Mn2II,II
dimer as for the structured Mn2II,II dimer signal.
2
Can take any value 0-0.33 (i.e. all allowed values for E/D). The parameter is non-unique.
1
Figure S1: UV-Visible spectrum of ZnCe6 free in solution (red) and bound to BFR1 (black).
All samples were in 25mM tricine, 100mM KCl pH7.7.
Figure S2: a.) Perpendicular X-band EPR spectra of the BFR1-ZM complex with an
underlying multiline signal apparent. b.) Cubic spline subtraction of the baseline to more
clearly visualize the multiline signal (magnified 2 times). Protein concentration 400μM,
ZnCe6 200μM, Mn 800μM. Microwave frequency 9.44 GHz, microwave power 2 mW,
modulation amplitude 2 mT, temperature 5 K.