From the Institut für Experimentalphysik, Freie
Universität Berlin, Arnimallee 14, 14195 Berlin, Germany,
§ Lehrstuhl Organische Chemie und Biochemie, Technische
Universität München, Lichtenbergstrasse 4, 85747 Garching,
Germany, ¶ Institut für Botanik,
Ludwig-Maximilians-Universität München, Menzinger Strasse
64, 80638 München, Germany, and
Institut für
Zellbiologie, Ludwig-Maximilians-Universität München,
Schillerstrasse 42, 80336 München, Germany
Received for publication, June 4, 2002, and in revised form, December 26, 2002
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ABSTRACT |
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The LOV2 domain of Avena sativa
phototropin and its C450A mutant were expressed as recombinant fusion
proteins and were examined by optical spectroscopy, electron
paramagnetic resonance, and electron-nuclear double resonance. Upon
irradiation (420-480 nm), the LOV2 C450A mutant protein gave an
optical absorption spectrum characteristic of a flavin radical even in
the absence of exogenous electron donors, thus demonstrating that the
flavin mononucleotide (FMN) cofactor in its photogenerated triplet
state is a potent oxidant for redox-active amino acid residues within
the LOV2 domain. The FMN radical in the LOV2 C450A mutant is
N(5)-protonated, suggesting that the local pH close to the FMN is
acidic enough so that the cysteine residue in the wild-type protein is
likely to be also protonated. An electron paramagnetic resonance
analysis of the photogenerated FMN radical gave information on the
geometrical and electronic structure and the environment of the FMN
cofactor. The experimentally determined hyperfine couplings of the FMN
radical point to a highly restricted delocalization of the unpaired
electron spin in the isoalloxazine moiety. In the light of these
results a possible radical-pair mechanism for the formation of the
FMN-C(4a)-cysteinyl adduct in LOV domains is discussed.
Numerous phenomena in the life cycle of plants such as circadian
timing, regulation of gene expression, and phototropism (the adaptive
process whereby plants bend toward a light source to maximize light
capture for photosynthesis) are responses to ambient light levels in
the UV-A and blue spectral regions comprising wavelengths of about
320-500 nm (1-3). Currently, two classes of blue light photoreceptors
have been identified in plants; they are the cryptochromes (4-6) and
the phototropins (7, 8), both of which are flavoproteins. Phototropins,
the blue light photoreceptors for phototropic bending (9, 10),
chloroplast relocation (11-13), and stomatal opening (14), have been
identified in several plant species including Arabidopsis
thaliana, Avena sativa (oat), Oryza sativa
(rice), and Zea mays (corn) (8). Phototropin of A. sativa is a protein comprising 923 amino acids, which is specified
by the phot1 gene (previously designated
nph-1) (1, 9). The protein contains two 12-kDa flavin
mononucleotide (FMN) binding domains. The FMN binding modules belong to
the PAS (PER/ARNT/SIM) domain superfamily (15, 16) occurring in many regulatory proteins and have been designated as
LOV1 domains; the acronym is
based on the involvement in the signaling of light,
oxygen, or voltage levels (10, 17). Very
recently, the presence of a LOV domain with similarity to the
photoactive LOV domains of the phototropin of higher plants has been
identified in the non-photosynthetic soil bacterium Bacillus
subtilis (18).
After illumination by blue light, recombinant LOV domains of
phototropin undergo a transient and fully reversible bleaching of their
optical absorption at 400-500 nm accompanied by an increase of
absorption at 390 nm (19, 20). Based on the similarity of the spectral
characteristics of the photoproduct 19 and that of a kinetically
competent intermediate in mercuric ion reductase (21), Vincent
Massey2 suggested that the LOV
photocycle comprises a light-induced addition of a thiol group
(cysteine 450 of phototropin in the case of the LOV2 domain from
A. sativa) to the C(4a) position of the flavin chromophore
followed by the spontaneous fragmentation of the adduct in the dark
(see Fig. 1). This hypothesis could later
be confirmed by 13C NMR spectroscopy (22).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
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Fig. 1.
The formation of a cysteinyl-flavin-C(4a)
covalent adduct in LOV2 after the absorption of blue light by
the FMN cofactor.
Recently, the crystal structure of the LOV2 domain of the phytochrome/phototropin chimeric photoreceptor phy3 from the fern Adiantum capillus-veneris was solved at a 2.7 Å resolution (23). The single LOV2 cysteine residue is located 4.2 Å from the flavin atom C(4a). Until now, however, the details of the mechanism of adduct formation have not been conclusively established. The ground state, an intermediate state resembling the FMN triplet (3FMN), and the adduct (19, 20) have been observed in phototropin by optical spectroscopy, whereas with 13C and 31P NMR (22) and x-ray crystallography (24) the ground state and the adduct could be characterized.
Swartz et al. (20) propose an ionic reaction pathway for
FMN-C(4a)-thiol adduct formation with the Cys-450 residue initially present as a thiolate, thus requiring a proton donor other than Cys-450
to protonate the light-generated FMN triplet (20). Recently, however,
it has been shown by Fourier transform infrared spectroscopy that the
cysteine residue in LOV2 is protonated in the ground state (25),
implying that an ionic mechanism, which relies on the presence of a
thiolate, is flawed. Thus other potential reaction pathways including a
concerted mechanism and a radical-pair mechanism have to be considered.
We have therefore investigated both the wild-type A. sativa
LOV2 domain and a C450A mutant by electron paramagnetic resonance (EPR)
spectroscopy. The formation of the FMN-thiol adduct is not possible in
the C450A mutant due to the absence of a thiol group. In this case, as
will be shown, 3FMN undergoes a photoreduction resembling
that observed in photolyases (26-28) and other flavoenzymes with the
possible participation of a redox-active amino acid residue, resulting
in the formation of a neutral flavin radical, FMNH·. The
implications of the observation of FMNH· are discussed in terms
of a possible radical-pair mechanism for adduct formation in LOV
domains. The characterization of FMNH· by EPR and
electron-nuclear double resonance (ENDOR) spectroscopy (29) allowed us
to probe the geometrical and electronic structure and the environment
of the flavin cofactor in the LOV2 domain using the paramagnetic state
as a natural spin label.
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EXPERIMENTAL PROCEDURES |
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Construction of an Expression Vector-- The open reading frame specifying hisactophilin of Dictyostelium discoideum was amplified using the oligonucleotides described in Table I as primers and plasmid pIMS5/c516 as template. The amplified DNA fragment was digested with EcoRI and HindIII and ligated into a pNCO vector (30) that had been treated with the same restriction enzymes. The resulting plasmid pNCO-HISACT-BNH was transformed into Escherichia coli strain XL1-Blue.
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Construction of Expression Plasmids--
The gene segment
specifying the A. sativa LOV2 domain was excised from the
plasmid pCAL-LOV2 (19) by restriction with BamHI and
HindIII. The fragment was ligated into the vector
pNCO-HISACT-BNH, which had been treated with the same enzymes. The
resulting plasmid designated pNCO-HISLOVWT specifies a fusion protein
comprising the LOV2 domain of phot1 of A. sativa
and hisactophilin from D. discoideum (Fig.
2). An expression plasmid specifying the
corresponding C450A mutant was obtained by the same approach starting
from the plasmid pCAL-LOV2C450A (19). The plasmids were
electro-transformed into E. coli strain M15[pREP4].
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Protein Expression and Purification--
Bacterial cells were
grown in LB medium supplemented with ampicillin (180 mg/liter) and
kanamycin (15 mg/liter) to an optical density of 0.6 at 600 nm.
Isopropyl-thio--D-galactopyranoside was added to a final
concentration of 1 mM. The cultures were incubated for
5 h, harvested by centrifugation, and stored at
20 °C. Frozen
cell mass (5 g) was thawed in 15 ml of buffer A (50 mM Tris
hydrochloride, pH 8.0, containing 100 mM NaCl and 2 mM CaCl2) supplemented with 20 mg of lysozyme.
The suspension was sonicated and centrifuged. The supernatant was
applied to a column of Chelating Sepharose Fast Flow (column volume, 15 ml; Amersham Biosciences), which had been equilibrated with
buffer B (50 mM sodium phosphate, pH 8.0, containing 300 mM NaCl) supplemented with 10 mM imidazole. The
column was washed with 150 ml of buffer B and was then developed with a
linear gradient of 10-500 mM imidazole in buffer B (total
volume, 100 ml). Yellow fluorescent fractions were combined and
concentrated by ultrafiltration (10-kDa membrane, Pall Gelman, Ann
Arbor, MI). The solution was applied to a column of Superdex S75-prep
grade (2.6 × 60 cm, Amersham Biosciences), which was developed
with 50 mM potassium phosphate, pH 7.5. Yellow fluorescent
fractions were combined and concentrated by ultrafiltration to a final
concentration of 1 mM as determined photometrically (
447 = 13,800 M
1cm
1 (19)).
Cleavage of the LOV2 Cys-450 Hisactophilin Fusion Protein by Thrombin-- 10 mg of LOV2 C450A hisactophilin fusion protein in buffer B supplemented with 2.5 mM CaCl2 were mixed with 50 units of thrombin (Sigma) and incubated overnight at room temperature. The protein solution was applied to a column of Chelating-Sepharose Fast Flow (column volume, 15 ml) that had been equilibrated with buffer B supplemented with 10 mM imidazole. Cleaved LOV2 C450A protein was collected in the flow-through, whereas hisactophilin protein and uncleaved fusion protein remained bound to the column. Yellow fluorescent fractions were combined and concentrated by ultrafiltration through microconcentrators (1-kDa membrane, Pall Gelman, Ann Arbor, MI). Protein homogeneity was monitored by SDS-PAGE electrophoresis.
UV/Visible Measurements-- Protein samples (concentration ~0.05 mM) were transferred into an optical cuvette (path length, 1 mm; Hellma, Müllheim, Germany) and supplemented with 0.5 mM EDTA depending on the sample and/or deoxygenated and then illuminated for up to 40 min with 420-480-nm light from a filtered Xe lamp (ILC, PS800SW-1) at room temperature. Decay of the radical was measured in a Shimadzu UV-1601PC (Shimadzu Scientific Instruments, Columbia, MD) spectrophotometer.
Buffer Exchange-- Samples were transferred into the desired buffer (usually 70 mM sodium/potassium phosphate, pH 7.0) in H2O or D2O by dilution and ultrafiltration through C10 microconcentrators at 4 °C. The cycle was repeated 5 times to give a final D2O enrichment of 93-97%.
EPR Sample Preparation-- The enzyme preparations were transferred into EPR quartz tubes (3-mm inner diameter for X-band (9-10 GHz) EPR; 0.6-mm inner diameter for W-band (95 GHz) EPR) under an argon inert gas atmosphere. They were then illuminated with light of 420-480 nm from a filtered Xe lamp and frozen rapidly in liquid nitrogen.
EPR Instrumentation-- Continuous-wave (cw) EPR spectra at X-band frequencies (9-10 GHz) were obtained using a laboratory-built spectrometer. It consists of a Bruker ER041MR microwave (mw) bridge (Bruker, Rheinstetten, Germany) and an AEG-20 electromagnet. Samples were placed in a Bruker ER4118X-MS-5W1 dielectric resonator, which was immersed in a laboratory-built helium gas flow cryostat controlled by a LakeShore 321 temperature controller.
W-band cw-EPR spectra were recorded with a laboratory-built high field EPR spectrometer operating at 94-96 GHz and equipped with a cylindrical TE011 cavity. The six-line EPR signal of a Mn(II)/MgO standard, placed near the sample in the cavity, was recorded simultaneously for g-factor calibration.
ENDOR Instrumentation--
X-band cw-ENDOR spectra were recorded
using a laboratory-built spectrometer consisting of an AEG-20
electromagnet and a Bruker ER041MR mw bridge. A radio frequency
synthesizer (Hewlett Packard 8647A) in conjunction with a high power
radio frequency amplifier (ENI A-300) was used to generate the cw radio
frequency field in the laboratory-built TM110 ENDOR
resonator (Q 1800, 1 turn/mm of NMR coil). The
temperature was adjusted using a nitrogen-gas flow controlled by a
Bruker ER4111VT temperature controller.
Calculations--
To assist in the assignment of experimentally
determined hyperfine couplings (hfcs) to individual nuclei in the
7,8- dimethyl isoalloxazine moiety of FMNH·,
density-functional theory calculations were performed with
lumiflavin (7,8,10-trimethyl isoalloxazine) using the program
package Gaussian 98 (31). Lumiflavin is considered as a valid model
because the ribityl side chain has no significant influence on the
electronic structure of FMNH·, which is bound in an extended
conformation in the LOV domain. The geometry of the FMNH· state
was optimized at the unrestricted B3LYP/EPR-II level of theory, and
single-point calculations of hfcs and electron densities were performed
at the same level.
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RESULTS |
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Optical Spectroscopy-- Expression vectors specifying fusion proteins comprising the A. sativa LOV2 domain (amino acid residues 412-516) or its C450A mutant and the actin binding hisactophilin protein from D. discoideum were constructed as described under "Experimental Procedures." E. coli strains harboring one of these plasmids formed copious amounts of the recombinant fusion protein (about 15% of soluble cell protein), which could be purified easily by affinity chromatography using a nickel-chelate column. The fusion proteins bind tightly to the chelating-Sepharose matrix as a consequence of the abundant histidine residues at the surface of hisactophilin. LOV2 C450A domain, from which the hisactophilin module was cleaved, was prepared with a yield of ~80% as described under "Experimental Procedures." LOV2 C450A domain fused with calmodulin-binding protein was expressed and purified according to published procedures (19).
The optical absorption spectra of the LOV2 C450A domains from the
samples under investigation before illumination and after irradiation
with blue light (420 < < 480 nm) are shown in Fig. 3. Both fusion proteins have virtually
the same ground-state absorbance properties (Fig. 3, B and
C, solid lines) as LOV2 protein
cleaved from hisactophilin (Fig. 3A, solid line)
but show enhanced stability and solubility. All three proteins have
absorption maxima at 363 and 447 nm, characteristic of an FMN
chromophore in the oxidized redox state. Shoulders at 425 and 474 nm
are vibrational contributions that are well resolved. This is
indicative of tight binding between the noncovalently bound FMN and the
highly ordered protein structure as well as of the nonpolar nature of
the flavin binding pocket.
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After continuous blue light irradiation all three LOV2 C450A proteins show the formation of a flavin radical characterized by absorption maxima at 570 and 605 nm (Fig. 3, dashed and dotted lines). Radical formation occurs in the absence (Fig. 3, A-C) and in the presence of an exogenous electron donor such as EDTA (Fig. 3D). Furthermore, the radical formation could be observed either in the presence (Fig. 3, dotted lines) or in the absence (Fig. 3, dashed lines) of oxygen. However, inspection of Fig. 3 clearly shows that the yield of flavin radical differs depending on the protein construct and on the photoreduction conditions. By comparison with the absorption spectra of other flavin radicals in a protein matrix (see, e.g. Ref. 27), we conclude that in the fusion protein comprising the LOV2 C450A domain and hisactophilin, supplemented with a 10-fold excess of EDTA, the FMN cofactor is fully converted from the oxidized to the one-electron reduced semiquinone form (Fig. 3D, dashed line). To compare the amount of flavin semiquinone radical formed in the various other samples, we compare their absorbances at 605 nm normalized to the LOV2 C450A hisactophilin fusion protein sample supplemented with EDTA (Fig. 3D, dotted line). The salient points of these data are that the presence of oxygen reduces the radical yield, whereas the presence of either EDTA or the protein fusion partner hisactophilin enhances the radical yield and reduces the oxygen dependence (see Table II).
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In all samples investigated, the flavin radical is completely
reoxidized in the dark to regenerate oxidized FMN on a time scale of
minutes depending on the protein construct and buffer conditions (see
Table II). We have monitored this process by observing the absorbance
changes at 605 nm as a function of time after blue light illumination
(see Fig. 4). Under anaerobic conditions
(squares) the flavin radical in the cleaved LOV2 C450A
domain (Fig. 4A) slowly decays with a 1/e time
constant of (78 ± 7) min, whereas under aerobic conditions
(circles) the radical decay is much faster (7 ± 1 min). In the presence of excess EDTA, the decay times in the absence
(triangles) or presence (diamonds) of oxygen are
virtually identical (see Table II).
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For the fusion protein comprising the LOV2 C450A domain and hisactophilin, similar trends are observed (Fig. 4B). Under anaerobic conditions (squares) the flavin radical decays with a 1/e time constant of (43 ± 5) min, which is somewhat shorter than that observed for the cleaved LOV2 C450A domain. Under aerobic conditions (circles) the radical decays with a 1/e time constant of only (8 ± 2) min, which is very close to the value observed for the cleaved LOV2 C450A. In the presence of excess EDTA, the decay times are extended depending on the absence (31 ± 4 min, triangles) or presence (28 ± 4 min, diamonds) of oxygen.
EPR Spectroscopy--
To characterize in more detail the
light-induced FMN radical in LOV2 C450A domains, EPR experiments were
performed with a sample that was frozen in liquid nitrogen immediately
after blue light irradiation. The X-band cw-EPR signal (Fig.
5) reveals a radical signature centered
at g = 2.0032 ± 0.0001, which is characteristic for a neutral flavin radical (32, 33). Neutral flavin radicals protonated at N(5) can be distinguished from anion flavin radicals by
means of their characteristic peak-to-peak EPR line widths of 2.0
and
1.5 mT, respectively, which is due to the presence or absence of
the large hfc of the proton at N(5) (29, 34). The observed peak-to-peak
line width of 2.00 ± 0.01 mT is typical of a neutral flavin
radical, FMNH·. The overall line width and line shape of the
signal is attributed to the mostly unresolved contributions of hfcs of
the unpaired electron spin with 1H and 14N
nuclei of the isoalloxazine moiety of FMN as well as of the protein
environment.
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To try to detect the EPR signal of a possible second radical species
that ought to be created upon electron transfer to FMN in the LOV2
C450A mutant protein, EPR experiments at higher microwave frequencies
(95 GHz) and correspondingly larger magnetic fields were also performed
(Fig. 6), because overlapping signal
contributions from organic radicals are better separated in high
magnetic fields due to their differences in g-factor.
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Even at 95 GHz, only the spectrum arising from one radical species was observed. Scanning over a wider magnetic field range did not reveal any additional signals except for the hyperfine lines of the Mn(II)/MgO standard used for g-factor calibration. The signal shown in Fig. 6A resembles the frozen-solution spectrum of a neutral flavin radical and is similar to the flavin adenine dinucleotide semiquinone radical observed in DNA photolyase (32). From spectral simulation of the rhombic symmetry of the signal, the principal values of the g-matrix, gX = 2.0042 ± 0.0001, gY = 2.0035 ± 0.0001, and gZ = 2.0020 ± 0.0001 (X, Y, and Z are the principal axes of the g-tensor) may be extracted.
Some hyperfine structure emerges in both W-band and X-band EPR spectra
(Figs. 5A and 6A). Although not fully resolved, a
spacing of 1.05 mT (29 ± 3 MHz) between adjacent shoulders in the
signals could be determined. After the procedure outlined previously
(32), this hyperfine splitting has been assigned to the hfc tensor
component Az of H (5) (x, y,
and z are the principal axes of the H (5) hyperfine
tensor based on the following findings. (a) The
splitting disappears when H (5) is replaced with a deuteron in an
exchange of the protonated for deuterated buffer (Figs. 5B
and 6B); (b) in the high field EPR experiment,
the center of the hyperfine structure shifts toward resonance field
values where molecules with their gz axis aligned
parallel to the external magnetic field are in resonance (Fig.
6A), which is characteristic for an -proton; and
(c) the exchangeable
-proton at N (3) is expected to
contribute only marginally to the overall EPR line width due to its
small hfc in the 1-2-MHz range (35).
ENDOR Spectroscopy--
To characterize in greater detail the
electronic structure of the FMN radical, we have also performed ENDOR
experiments. For doublet-state radicals, two ENDOR lines are expected
per proton hfc tensor component, A. When A < 2 H , the resonance frequencies are
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(Eq. 1) |
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(Eq. 2) |
In Fig. 7, A and B,
the X-band cw-ENDOR spectra of the FMN radical cofactor in protonated
and deuterated buffer are shown, recorded at a magnetic field of 333.86 mT, corresponding to giso = 2.0032. The frozen
samples give rise to powder-type spectra that are symmetrically
centered at the proton Larmor frequency, H.
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A group of overlapping signals with hfcs A 2.0 MHz
forms the so-called "matrix" ENDOR line and is the sum of
contributions from the weakly coupled protons at C(7
) and C(9) of
the isoalloxazine moiety as well as protons of solvent water and amino
acid residues near the cofactor-binding site. Their individual signal
contributions are not readily assigned. One conclusion, however, may be
drawn from a comparison of the matrix lines of the protein in
protonated and deuterated buffer (Fig. 7, A and
B); the remarkably small differences in intensity and shape
indicate that the cofactor-binding site comprises mostly
non-exchangeable protons.
A pair of lines with an hfc of 2.3 ± 0.1 MHz is located next to the matrix region (Fig. 7A). Based on the disappearance of these signals upon buffer deuteration (Fig. 7B) they are assigned to the proton at N(3), see Table III.
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When comparing the protonated and deuterated samples in the region
where the resonances of H(6) and the methyl protons H(8) are
expected, two changes become obvious. First, there is a feature at 10.2 MHz that does not appear at 18.2 MHz in the protonated sample (Fig.
7A). At first glance this might be due to a large nitrogen
hfc tensor. Upon deuteration, however, the spectrum is restored to
symmetry, demonstrating that this feature is due to a proton hfc tensor
component (Fig. 7B). It must be a very large hfc tensor
component that has been reflected through zero frequency, because it
only appears on one side of
H. From Equation 2, a hfc of
48.8 ± 1 MHz may be estimated. This can only be the Ay component of H (5). There should be a partner
transition at 38.6 MHz, but it has not been detected. Second, the
features at 10.7 and 17.7 MHz decrease in intensity on going from
protonated to deuterated buffer. This indicates that another hfc
component, again from an exchangeable proton, has resonances at these
frequencies. This hfc component of 7.0 ± 0.5 MHz is, therefore,
assigned to the smallest component, Ax, of H (5).
-Protons in a CH segment of a planar
system are predicted to
resonate at Ax
0.5 × Aiso, Az
Aiso, and Ay
1.5 × Aiso (36). For the H(5) of FMNH· in LOV2
C450A, the anisotropy is even larger, almost approaching 70% of
Aiso.
The remaining pronounced hfcs are expected to come from the methyl
protons at C(8). Yet the pattern that is observed in the spectrum of
the sample in deuterated buffer is still not typical of the axial
symmetry expected from a freely rotating methyl group. Rather it is
also an overlap of two roughly axial symmetric proton hfc tensors. This
is clear from the inflections observed at 10.7 and 17.7 MHz. The only
other proton that can have such a large hfc and with the correct axial
symmetry is one of the
-protons at C(1').
Two different hfcs are expected for the two -protons at position
C(1'). The distance of these protons to N(10) (which carries a high
spin density) is rather large, and the anisotropic dipolar coupling is,
therefore, relatively small. They may have a large Fermi contact hfc
interaction, Aiso, however, which varies with the twist angle
between the N(10)s 2pz orbital and the
plane defined by the N(10)-C(1') bond and the respective C(1')-H bond. A cos2 dependence of the splitting is expected if the
coupling is caused by hyperconjugation (37, 38). From the x-ray
structure of the LOV2 domain (23),
is 4.9°. Taking this angle
into account in a model structure of the isoalloxazine ring and the
ribityl side chain and calculating hfcs using density-functional theory gives values of 7.4 and 2.6 MHz for Aiso of the
more strongly and the more weakly coupled proton at C(1'), respectively.
Deconvolution of A and
A
of the methyl groups at C(7
) and C(8
)
and the
-protons at C(1') is accomplished utilizing orientation-selection effects that appear when ENDOR spectra are recorded at off-center magnetic field positions (Fig. 7C) in
the EPR spectrum, as indicated in Fig. 5. The outer parts of the
FMNH· EPR signal predominantly arise from molecules with
14N nuclear quantum number values of either +1 or
1 that have their molecular plane perpendicular to the magnetic
field. Hence, by desaturation of the spectral wings of the EPR signal,
single-crystal like ENDOR spectra can be expected where only those
components of the proton hfc tensors are detected that have their axes
parallel to the respective component of the dominating 14N
hfc tensors, N(5) and N(10). In the case of methyl protons this is the
A
component (perpendicular to the cylindrical
axis of the rotating methyl group). Similar selection principles apply for the
-protons at C(1'). From the two ENDOR lines with hfcs of 8.7 and 6.8 MHz, only the latter may be detected at off-center field
settings. Therefore, these two signals may be assigned as the
A
= 8.7 ± 0.3 MHz and
A
= 6.8 ± 0.1 MHz components of the hfc
tensor of H(8
) and the more strongly coupled proton at C(1'), giving
Aiso = 7.4 ± 0.2 MHz (see Table III).
The off-center field ENDOR experiment has enabled the identification of
the hfc due to the methyl group at C(7) even though the assignment
is more difficult due to the small size of this coupling. Nevertheless,
a value of 1.6 ± 0.1 MHz for Aiso (see Table III) could be obtained for this nucleus in accordance with the
following evidence. First, both the A
and
A
features can still be observed after
H2O/D2O buffer exchange (see Fig. 7,
A and B), demonstrating that the corresponding
proton is neither exchangeable nor a true matrix proton. In the
orientation-selection experiment (see Fig. 7C) only the
A
component should be observed. Indeed, in
the correct region around 2 MHz, there is a major change upon
orientation selection. This feature may, therefore, be assigned to
A
.
Finally, the hfc 5.9 ± 0.1 MHz due to H(6) could also be detected
and assigned from the orientation-selection ENDOR experiment. This is
an -proton and would be expected to be of rhombic shape. Typically,
only the central crossing point in the powder ENDOR spectrum of this
hfc tensor is visible. It is usually assigned to
Aiso, which is a good assumption as long as the
tensorial pattern is symmetric around Aiso.
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DISCUSSION |
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Blue light irradiation converts the wild-type LOV2 domain into an
FMN-C(4a)-cysteinyl adduct via the FMN triplet state. Illumination of
the LOV2 C450A mutant with blue light has also been shown to produce a
flavin triplet (20), 3FMN; however, the subsequent
generation of a flavin radical has not yet been reported. Here we have
shown that in the LOV2 C450A mutant, electron transfer to
3FMN takes place, with the participation of an as yet to be
identified amino acid residue (X' in Fig.
8), resulting in the formation of a
neutral FMN radical, FMNH·.
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The observation of FMNH· is independent of protein construct and buffer conditions such as the presence of an exogenous reductant. This is in contrast to the well known photoreduction of flavoproteins (described by Massey and Palmer (39)) in which supplemented electron donors are required for flavin semiquinone generation. Therefore, we conclude that in the LOV2 C450A domains the flavin radical is formed by intraprotein electron transfer from a redox-active amino acid. Tryptophan, histidine, or tyrosine are likely candidates for electron transfer to flavin triplets (due to their lower redox potential compared with that of 3FMN) as has been shown previously by laser flash photolysis (40, 41).
The amount of FMNH· produced is different for the various LOV2
samples, and these variations deserve some comment. In principle, the
radical yield reflects the competition between radical formation and
its decay, the latter occurring via either back electron transfer or
reoxidation of FMNH· by molecular oxygen to form FMN and
O
Reoxidation of FMNH· by molecular oxygen is expected to be similar for all three LOV2 samples, as is indeed observed experimentally (Fig. 4). Under aerobic conditions, this is clearly the preferred process for regeneration of oxidized FMN and, therefore, dominates flavin radical decay kinetics. Under anaerobic conditions, radical decay occurs via back electron transfer to regenerate the thermodynamically more stable oxidized FMN. Again, the hisactophilin-fused LOV2 domain provides more efficient electron transfer pathways compared with the cleaved LOV2 domain, and therefore, radical decay is enhanced in the recombinant fusion protein.
The situation is different in the presence of EDTA. EDTA is a well known reducing agent for photoexcited flavin triplet states (39). In solution studies, triplet flavin abstracts a hydrogen atom from EDTA, forming a neutral flavin radical, whereas the ETDA then decarboxylates and fragments to produce stable products (44, 45). However, in the photoreduction of LOV2 C450A, EDTA has only minor influence on the measured radical yields (Fig. 3, C and D), which indicates that FMNH· is almost exclusively formed by intraprotein electron transfer rather than by reaction of 3FMN with EDTA. On the other hand, in the presence of EDTA, the reoxidation of FMNH· becomes independent of oxygen content (Fig. 4, diamonds and triangles). Qualitatively, this finding may be understood in terms of the EDTA sequestering trace amounts of transition metal ions (that are unavoidably present as impurities from the protein preparation) that, if not chelated, may act as catalysts in the activation of molecular oxygen (46, 47). Thus, EDTA plays its well known antioxidant role by preventing reoxidation of the flavin.
Once FMNH· is generated, its decay is extremely slow (Fig. 4). This is even more evident when taking into account the small size of the LOV2 domain (23). Electron transfer time constants in the sub-microsecond range are observed in other flavoproteins; for example, in photolyases in which electrons are transferred over distances exceeding the diameter of LOV domains (28). This demonstrates that, in contrast to many other flavoproteins, low activation energy electron transfer pathways do not exist in the LOV2 domain or that endergonic steps in the electron transfer pathway have to be considered. Nevertheless, both redox forms of the FMN chromophore, oxidized and one-electron reduced, are well stabilized by the protein matrix. Hence, overall the excited-state flavin is optimized to undergo electron transfer but the electron transfer pathways leading to the surface of the LOV2 domain are inefficient. This may be necessary so that adduct formation with Cys-450 in the wild-type, which takes place on a microsecond time scale (20, 48), could dominate alternative electron transfer processes. From this study we cannot directly draw conclusions about the time scale or quantum yield of the forward electron transfer observed in the LOV2 C450A domain. A slowdown of electron transfer by roughly an order of magnitude is expected for every 2-Å increase of distance between the redox partners, if other parameters such as protein packing density, free energy, and reorganization energy remain comparable (49). Thus, in comparison to a possible electron transfer from Cys-450 (which is 4.2 Å away from C(4a) of FMN) to 3FMN in the wild-type a roughly 104-fold decreased rate of electron transfer from one of the other (more distant) redox-active amino acids in the LOV2 domain to 3FMN would be expected. This decrease, however, is predicted to make the electron transfer rate so slow that 3FMN would be mostly converted back to the ground state before electron transfer could happen. Thus, the quantum yield of electron transfer should be rather low. This is most likely the reason why in previous laser-flash photolysis studies no flavin radical formation was observed, whereas by using continuous irradiation in this study, a flavin radical was generated at high yield. Taken together, these observations have several major implications for adduct formation in LOV domains, which will now be considered.
Swartz et al. (20) propose an ionic reaction pathway with
the Cys-450 residue initially present as a thiolate. In this mechanism, 3FMN is protonated at N(5) by a nearby and as yet
unidentified proton-donating group in the protein to give the
FMNH+ cation. Upon protonation of N(5) (which occurs in the
ground state FMN only at pH 0), the electron density
distribution of the isoalloxazine ring is altered due to the
non-bonding pair of electrons at N(5) becoming a bonding pair with the
additional proton. The FMNH+ carbocation, which formally
has a positive charge at C(4a), is the electrophile that can then form
a bond with the nucleophilic thiolate, thus generating the adduct. The
hypothetical ionic mechanism also requires that Cys-450 is present as a
thiolate and simultaneously that 3FMN is protonated before
adduct formation occurs. Yet in solution it has been demonstrated that
flavin triplet protonation occurs only at pH < 4.4 (50-52). At
higher pH, 3FMN undergoes electron transfer followed by
protonation when the pH is <8.3 to form a neutral flavin radical (50)
rather than the anion radical, FMN·-. These facts
seem to be somewhat contradictory, and although it is not unreasonable
that the local pH at the FMN-binding site could be rather low given the
proximity of FMN and C450, it seems unlikely that the latter could at
the same time be deprotonated (pKa of cysteine in
solution, 8.37). Furthermore, it has recently been shown by Fourier
transform infrared spectroscopy that the cysteine residue is protonated
in the ground state (25).
An alternative that is consistent with our experimental observations is a radical pair mechanism (see Fig. 8B; for review, see Ref. 53). 3FMN is an extremely efficient oxidizing agent in the presence of electron donors such as EDTA or redox-active amino acids in solution (40, 41) and in various proteins, including the LOV2 domain, as demonstrated in this study. A flavin semiquinone is formed from 3FMN in a one-electron photoreduction that may be followed by proton transfer depending on the pH to give the anionic or neutral flavin radical (39).
In the wild-type LOV2 domain 3FMN could abstract an
electron from Cys-450 and a spin-correlated ionic radical pair
consisting of an anionic flavin radical, FMN·, and
a sulfur-centered radical, RS·+H, would be formed.
This could undergo subsequent proton transfer to give the neutral
flavin radical, FMNH·, and a sulfur-centered radical,
RS·. Alternatively, 3FMN could directly abstract a
hydrogen atom from cysteine, giving the same product, a neutral radical
pair. A radical pair created from a triplet state precursor has
initially the same spin state (i.e.
3[FMNH· ··· RS·]), due to the
conservation of angular momentum, and thus cannot form a covalent bond.
At first glance this might be thought to rule out adduct formation via
a radical-pair mechanism or that radical pairs should have been
detected in the wild type, which has not been the case to date. A
covalent bond and, hence, the FMN-cysteinyl adduct may only form if the
spin state of the radical pair evolves to obtain a singlet character by
singlet-triplet mixing: 3[FMN· ···
RS·]
1[FMN· ··· RS·].
Because of strong spin-orbit coupling in sulfur-centered radicals (spin-orbit coupling constant, 382 cm
1), however, a
mechanism for extremely rapid singlet-triplet interconversion exists.
At room temperature, in solution, these radicals have been shown to
have relaxation times in the nanosecond regime (54). Hence, if spin
mixing occurs on this time scale or even faster, the covalent adduct
will be formed on the same time scale, which implies that the lifetime
of the intermediate radical pair will be too short for detection by
EPR. That they were not detected by optical methods simply implies that
their short-lived absorption may be swamped by the background of
relatively long-lived 3FMN and adduct.
The spin chemistry is also important for the hypothetical ionic mechanism discussed by Swartz et al. (20) and for a possible concerted mechanism. Given that adduct formation proceeds via 3FMN, then FMNH+ and the adduct must also be created in the triplet state. That the adduct would be created in a triplet state is rather unlikely, but if it were, its spin state would have been evident from its effect on the NMR line widths, yet no line broadening was observed (22). The only possibility is that FMNH+ converts to the singlet ground state before the adduct is formed. This is certainly possible under the influence of the sulfur atom, but the singlet ground state would almost certainly deprotonate before adduct could form (recall that the ground-state flavin is a much weaker base than the excited state (55)).
In a hypothetical radical-pair mechanism these requirements are relaxed. Electron transfer may occur with either a protonated or deprotonated cysteine residue (40, 41). Flavin protonation at N(5) from either the cysteine or another donor group can occur either during the radical-pair lifetime or after the adduct has been formed. An important point is that protonation of 3FMN is not necessarily the rate-determining step in a radical-pair mechanism, whereas it should be in an ionic mechanism.
The minimal differences in the matrix region of the ENDOR spectra in
protonated and deuterated buffer are also of interest. Although the
x-ray structure does not show the presence of any water molecules close
to the FMN isoalloxazine ring, this does not necessarily prove that the
cofactor is in a water-free environment. Often in flavoenzymes the
matrix region collapses upon buffer deuteration. This is especially
true where a substrate approaches the flavin cofactor very closely to
facilitate, for example, hydrogen ion transfer (56). In DNA
photolyases, the matrix region is slightly altered, and it is known
from its x-ray structure (57) that the cofactor is mostly buried in the
protein and isolated from solvent water. In the C450A mutant of
phototropin, the changes in the ENDOR spectrum are minimal. No
exogenous substrate needs to approach the FMN cofactor, because the
cysteine is an integral part of the protein. The flavin triplet state
is a very reactive species that must be isolated so that it may only
perform its biological function; that is, adduct formation. In LOV2
domains, 3FMN is protected from reacting with
(a) molecular oxygen with which it may generate either
singlet oxygen or a FMN-C(4a)-peroxide adduct (hence, the slow
reoxidation of FMNH· to form FMN and O
The proton hfcs also allow some information on the electronic structure
of the cofactor to be drawn. First, the Aiso hfc
from the methyl H(8) protons determined by ENDOR, which is usually considered a guide to the overall spin-density distribution (due to its
ease of determination) is one of the smallest yet determined (32, 58).
This further supports the rather isolated situation of the FMN cofactor
in the LOV2 domain and points to a rather nonpolar cofactor-binding
site (59). Second, Aiso for H (5) determined by
EPR is substantially larger than that observed in DNA photolyase (32).
Taken together, these hfcs indicate that the unpaired electron is,
rather, localized toward the pyrimidine ring of the isoalloxazine
moiety. A localization of the unpaired electron would be expected to
aid the efficient formation of a bond between the flavin radical and
the cysteinyl radical in a radical pair mechanism.
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CONCLUSIONS |
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Our work unambiguously shows that the flavin cofactor of the LOV2 domain has electron transfer properties. Hence, in the wild type there must be a competition between adduct formation and the electron transfer reaction with another as yet unidentified amino acid residue. Nevertheless, adduct formation is the overwhelmingly dominant process in wild-type LOV2; no radicals are observed in the wild type that may be irradiated (reversibly) for long periods without suffering degradation (22). Previous optical studies have determined that adduct formation occurs on the microsecond time scale (20, 48). Hence, the competing electron transfer reaction observed in the mutant must be much less efficient and, hence, slower to ensure that only FMN-cysteinyl adduct formation occurs. This is in contrast to other flavoenzymes such as the structurally unrelated photolyases, where the photoreduction from the fully oxidized flavin proceeds on a nanosecond time scale (28). It would be surprising, therefore, if the crucial step in adduct formation were, as previously postulated (20), a proton transfer rather than an electron transfer, given that it should be significantly faster than the alternative electron transfer process observed in the C450A mutant.
The observation of a neutral flavin radical rather than an anion flavin radical gives information on the local pH of the flavin moiety. In aqueous solution, a neutral flavin radical is only formed at pH values < 8.3. Consequently, we conclude, in agreement with a recent Fourier-transform infrared study, that it is likely that the cysteine is at least partially protonated in the wild-type protein.
Although the evidence presented here is consistent with a radical-pair
mechanism for adduct formation, it is still not proven. This question
has to be addressed by time-resolved EPR and chemically induced dynamic
nuclear polarization experiments that are presently being performed
in our laboratory.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Michael Salomon and Elke Knieb for generously providing the plasmids pCAL-LOV2 and pCAL-LOV2C450A. We thank Professor Robert Bittl (Free University Berlin) and Professor Maria-Elisabeth Michel-Beyerle (Technical University Munich) for helpful discussions. Continued support by Professor Möbius is gratefully acknowledged.
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FOOTNOTES |
---|
* This work was supported by VolkswagenStiftung Grant I/77100 (to S. W.) and by the Deutsche Forschungsgemeinschaft, the Fonds der Chemischen Industrie, and the Hans-Fischer-Gesellschaft.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AF544403.
This paper is dedicated to the memory of Vincent Massey.
** To whom correspondence may be addressed. Tel.: 49-30-838-56139; Fax: 49-30-838-56046; E-mail: Stefan.Weber@physik.fu-berlin.de.
To whom correspondence may be addressed. Tel.: 49-89-289-13336;
Fax: 49-89-289-13363; E-mail: Gerald.Richter@ch.tum.de.
Published, JBC Papers in Press, January 13, 2003, DOI 10.1074/jbc.M205509200
2 A contribution to discussion at the 13th International Congress on Flavinsand Flavoproteins, August 29-September 4, 1999, Konstanz, Germany.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: LOV, light, oxygen, and voltage levels; EPR, electron magnetic resonance; ENDOR, electron-nuclear double resonance; cw, continuous-wave; mw, microwave; hfc, hyperfine coupling; mT, millitesla.
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