From the Department of Chemistry and Biochemistry,
University of California, Santa Cruz, California 95064 and the
§ Department of Plant Biology, Carnegie Institution of
Washington, Stanford, California 94305
Received for publication, September 6, 2002, and in revised form, October 29, 2002
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ABSTRACT |
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The phototropins are a family of
membrane-associated flavoproteins that function as the primary blue
light receptors regulating phototropism, chloroplast movements,
stomatal opening, and leaf expansion in plants. Phot1, a member of this
family, contains two FMN-binding domains, LOV1 and LOV2, within the
N-terminal region and a C-terminal serine-threonine protein kinase
domain. Light irradiation of oat phot1 LOV2 produces a cysteinyl adduct (Cys-39) at the flavin C(4a) position, which decays thermally back to the dark state. We measured pH and isotope effects on the
photocycle. Between pH 3.7 and 9.5, adduct formation showed minimal pH
dependence, and adduct decay showed only slight pH dependence,
indicating that the pK values of mechanistically relevant groups are outside this range. LOV2 showed a nearly 5-fold slowing of
adduct formation in D2O relative to H2O,
indicating that the rate-limiting step involves proton transfer(s).
Light-induced changes in the far UV CD spectrum of LOV2 revealed
putative protein structural perturbations. The light minus dark CD
difference spectrum resembles an inverted The phototropins are a family of blue light receptors that are
responsible for phototropism (1, 2) and are involved in light-induced
chloroplast movements (3) as well as blue light-stimulated stomatal
opening (4) and leaf expansion in higher plants (5). Phototropin-like
proteins have recently been identified in the green alga
Chlamydomonas reinhardtii (6, 7) and in bacteria (8). The
physiological roles of such proteins in these latter systems are not
yet elucidated.
The phototropin phot1 (9, 10), which becomes autophosphorylated in
response to blue light, is a membrane-associated protein that contains
two 12-kDa, FMN-binding LOV (light, oxygen,
voltage) (11, 12) domains (LOV1 and LOV2) in its N-terminal
region and a typical serine-threonine kinase domain in its C-terminal region (11). LOV domains belong to the PAS domain superfamily, which
are found in a variety of sensor proteins in organisms ranging from
archaea to eukaryotes (13).
Upon light excitation, the isolated LOV2 domain of phot1 undergoes a
cyclic photoreaction (14). The photocycle of phot1 LOV2 has been
elucidated in part (15). Blue light irradiation excites the FMN
chromophore to a triplet state that absorbs maximally around 660 nm
(designated LOV2L660) (15), resulting in
electronic redistribution that increases the basicity of N-5 of
the FMN chromophore (16). Protonation at N-5 and attack of the
Cys-39 1 sulfur at the C(4a)
position produce a flavin-cysteinyl covalent adduct (17-20) absorbing
maximally at around 390 nm (designated LOV2S390). Formation of the adduct occurs on
the microsecond time scale (15). The adduct thermally decays back to
the ground state on a time scale of many seconds and is proposed to be
the protein signaling state (15).
Despite the above progress made in understanding the phot1 LOV2
photochemistry, specific mechanistic and conformational steps of the
photocycle remain to be elucidated. Adduct decay for the LOV2 domain of
oat phot1 has been shown to be 3 times slower in D2O than
in H2O (15), indicating that proton transfer reactions, probably involving at least N-5, are rate-limiting components of the
back reaction. To date, no comparable information has been established
regarding the mechanism of adduct formation. Furthermore, negligible
protein motions were detected during the photocycle in crystals of the
LOV2 domain of fern phy3 (19), whereas major shifts in the amide proton
region were reported in a one-dimensional NMR study of the LOV2
photocycle of oat phot1 (18).
To elucidate the photocycle mechanism and structural changes further,
we measured the D2O and pH dependences of oat phot1 LOV2
photocycle kinetics and monitored light-induced changes of the circular
dichroism spectrum in the far UV region, where spectral bands are
typically attributed to protein secondary structure. Significant
D2O effects were observed but only minor pH effects. In
addition, changes in the far UV CD spectrum of LOV2 were observed and
may represent a reversible change in Sample Preparation--
Protein samples (oat phot1 LOV2 and
LOV2C39A)1 were prepared as
previously described (14). The LOV domains were expressed in
Escherichia coli and purified by calmodulin
affinity chromatography (Stratagene). Purified samples were washed into
clean buffer (5 mM Tris, 5 mM NaCl, pH 8) using
centrifugal concentrators (Continental Lab Products, San Diego, CA) and
were centrifuged and/or filtered through a 0.2-µm filter prior to
use. Protein concentrations were determined by absorption spectroscopy
using a Hewlett Packard HP8452 diode array spectrophotometer with
Light-induced Absorption Changes at Short Times--
Difference
spectra in the 30-ns to 690-ms time window were collected on an
instrument described previously (21). A dye laser pumped by the third
harmonic of an Nd:YAG laser provided a 10-ns, 80-µJ/mm2
light pulse at 477 nm. A fresh sample was provided for each laser flash, allowing the averaging of absorbance data of several samples. The laser pulse traversed the sample perpendicular to the path of white
light from a flashlamp that was used to probe the absorbance change in
the sample. The optical path lengths for the probe light and laser were
2 and 0.5 mm, respectively. The white light used to probe absorbance
was linearly polarized at the magic angle (54.7°) relative to the
laser polarization axis to prevent rotational diffusion artifacts (22).
The temperature for all measurements was about 20 °C.
Light-induced Changes at Long Times--
As described previously
(15), difference spectra in the 1-100-s time range were collected on
an HP8452A diode array spectrophotometer at room temperature. The
optical path length was 1 cm, and the excitation pulse was provided by
a white light camera strobe flash.
Global Analysis--
Data were analyzed using programs written
in a Matlab environment (The Mathworks, Natick, MA) as described
previously (15). Briefly, data matrices were constructed and subjected
to singular value decomposition followed by global exponential fitting
(23, 24). Kinetic changes at all measured wavelengths are decomposed into a sum of exponential components. The exponents contain the apparent rate constants for the observed kinetic changes, and the
amplitudes at different wavelengths represent the spectral changes
associated with the exponential process and are called the b-spectra
(23, 25).
D2O Exchange--
Protein samples were divided into
two equivalent aliquots and were lyophilized overnight in the dark as
described previously (15). One aliquot was then reconstituted in
H2O and the other in D2O.
pH Titrations--
Protein samples (5 mM Tris, 10 mM NaCl) began at pH 8. The pH was changed in a stepwise
manner using 0.5-1 M HCl or NaOH and was monitored with a
Corning Digital 110 meter and a Beckman Futura (model 511063;
Fullerton, CA) semimicro AgCl combination electrode.
CD Spectroscopy--
Protein CD spectra were recorded using an
Aviv 60DS CD spectrometer. A lid for the CD instrument was constructed
containing shutters for blocking the CD lamp and detector, and a hole
was made for a light guide that fit directly into the top of the
cuvettes. Thus, samples placed into the CD sample chamber were in
complete darkness while the CD lamp and detector were blocked and the
external lamp was off. The adduct form was induced by 20 s of
white light irradiation from an external 100-watt tungsten halogen lamp
via the light guide. We observed no spectral or kinetic differences resulting from the use of white or blue light. The CD measuring lamp
itself was not observed to be actinic. Full spectra were recorded at
2 °C in order to prolong the adduct form (14). Spectra were smoothed
using the smoothing program in SigmaPlot (SPSS Science). Using the same
irradiation method described above, we monitored relaxation kinetics by
CD at wavelengths of 450, 290, 222, and 208 nm at 20 °C. Control
experiments were conducted at 500 and 250 nm, wavelengths at which
little difference is observed between dark and light spectra (data not
shown). Points were recorded every 1 s for 700 s at 20 °C.
In the visible/near UV region (260-500 nm) 15 µM protein
was used in a 1-cm path length rectangular cuvette; in the far UV
region (190-250 nm), 7 µM protein was used in a 0.1-cm
path length rectangular cuvette. Relaxation kinetics by CD were also
measured at 10 and 4 °C for calculation of activation energy.
FMN-cysteinyl Adduct Formation from the FMN Triplet State Is Nearly
5 Times Slower in D2O than in H2O--
We
previously demonstrated by nanosecond laser spectroscopy (see
"Materials and Methods" and Ref. 15) that light irradiation of LOV2
produces the FMN triplet state, characterized by a broad absorption
band at about 660 nm (LOV2L660) and,
subsequently, the flavin-cysteinyl adduct with characteristic
absorption at 390 nm (LOV2S390) (15). Using the
same technique, we measured changes in the absorption difference
spectrum of LOV2 in H2O and in D2O between 30 ns and 690 ms after an excitation flash. The flavin triplet state
decayed in H2O with an apparent time
constant2 of 3 µs (data not
shown), slightly slower than our previously reported value (15). A
small difference in temperature could account for the difference in
rate. Fig. 1A shows the
absorption difference spectra of LOV2 in D2O. Global
exponential analysis of the measured difference spectra indicates a
single first order process with an apparent time constant of 14 µs,
almost 5 times slower than in H2O. The residuals (the
difference between spectral data and calculated data from the
exponential fitting) show no significant spectral features. The
inset of Fig. 1A shows the relative first-order
rates in H2O and D2O, approximated by the decay
of the 660-nm band alone.
Fig. 1B shows the corresponding b-spectra that
resulted from the global exponential analysis. The single exponential
fit indicates that two unique spectra (represented by
b1 and b0) exist over the
time course of the measurements, where each spectrum represents at
least one species. The b1-spectrum represents
the difference between the spectra of the first (present at 30 ns) and
final (persisting at 690 ms) intermediates and corresponds to the
apparent decay time constant of 14 µs. The
b0-spectrum is the spectrum of the final
intermediate referenced against the ground form. We showed previously
that the b1-spectrum resembles the difference spectrum of the FMN triplet state and that the
b0-spectrum is spectrally consistent with a
flavin-cysteinyl adduct (15). The presence of an isosbestic point (at
about 420 nm) is also consistent with a two-state system, as observed
previously (15). Therefore, the b1- and
b0-spectra in Fig. 1B, which are
nearly identical to those produced in H2O (data not shown)
(15), represent the same transitions of the triplet state to the
FMN-cysteinyl adduct and the ground form, respectively, as reported
earlier (15).
Absolute spectra for the triplet state
(LOV2L660) and adduct form
(LOV2S390) were calculated previously (15) by
adding to each intermediate spectrum (calculated from the
b-spectra) enough ground state absorption spectrum to remove
the bleach at 450 nm fully. Twice as much ground state absorption
spectrum was required to remove the bleach in the first intermediate
spectrum (calculated from b1) as in the second
intermediate spectrum (calculated from b0).
Hence, twice as much bleach at 450 nm exists in the triplet state as in
the adduct form, and therefore the triplet state apparently decays into
equal amounts of ground state and adduct. The apparent time constant of
3 µs for triplet decay in H2O therefore represents two
equivalent time constants2 of 6 µs (for return to ground
state and for adduct formation), as observed previously (15). In
D2O, the triplet state was also observed to decay equally
to the adduct form and to the ground state, resulting in time constants
of 28 µs for both adduct formation and triplet relaxation to the
ground state.
We showed previously that light irradiation of the LOV2C39A mutant,
which lacks the sulfur group required to form the adduct, produces the
triplet state, which then relaxes back to the ground state (15). In
contrast to the wild type, only a negligible D2O effect was
observed for the truncated photocycle of the LOV2C39A mutant. The
collected difference spectra are shown in Fig.
2A. The time constant observed
for decay of the triplet state to the ground state is 72 µs in
H2O and 80 µs in D2O; the relative rates are
illustrated in the inset of Fig. 2A. As can be
seen from the difference spectra, the decay of the 660-nm band is
concomitant with the full recovery of the bleaching at 450 nm,
indicating that no spectrally observable intermediates are formed
during triplet decay. As shown in Fig. 2B, global analysis
results in a b1-spectrum that is consistent with
the one from wild type LOV2 and a b0-spectrum of
zero.
Kinetics of FMN-cysteinyl Adduct Formation in H2O Are
pH-insensitive between pH 6.3 and 9.5--
To probe the dependence of
light-induced adduct formation on proton transfers further, we
monitored triplet decay at the same time values used above from pH 6.3 to 9.5. We stayed within this range to minimize
pH-dependent aggregation of LOV2 that occurs in solution at
protein concentrations sufficient to show significant changes by
absorption spectroscopy. The difference spectra and b-spectra obtained were the same as previously observed
(data not shown). As shown in Fig.
3A, only a slight effect on
the rate of adduct formation was observed from pH 6.3 to pH 9.5. The
time constant for triplet decay is near 2.5 µs at all pH values
measured. The relative pH insensitivity of adduct formation indicates
that titratable groups in the chromophore and mechanistically relevant protein residues are not titrated within this pH range. This finding is
consistent with our previous results that showed no change in
chromophore fluorescence intensity between pH 5.5 and 10 (15).
Kinetics of FMN-cysteinyl Adduct Decay Show Minor Change between pH
3.7 and 9.8--
We showed previously by absorption difference
spectroscopy that the FMN-cysteinyl adduct decays back to the ground
form with a half-life2 of about 45 s (15). Here, we
measured the rate of decay of the adduct form between pH 3.7 and 9.8. Slow chromophore release occurred below pH 5, as previously reported
(15). However, the back-reaction kinetics were sufficiently slow to
allow for longer signal integration times than were possible when
measuring adduct formation. Further, samples restored to neutral pH
showed recovered spectral and kinetic properties except for the
expected loss of bleaching intensity due to some irreversible
chromophore release. We were therefore able to acquire difference
spectra down to pH 3.7.
The difference spectra were unaffected by the presence of released FMN,
because it does not undergo a photocycle on the slow time scale of the
protein. As shown in Fig. 3B, the rate of adduct decay was
constant from high pH to about pH 7, below which a reproducible decrease in rate of about only 40% was apparent.
Circular Dichroism Spectrum of Phot1 LOV2 in the 190-500-nm Range:
Dark and Light-irradiated States--
The CD spectrum of the LOV2
ground form in the visible/near UV range was previously reported (14).
We extended the measurement to the far UV range, which contains protein
secondary structural information. The solid lines
in Fig. 4 represent the CD spectra obtained for the ground state of LOV2 from 500 to 260 nm (Fig. 4A) and from 250 to 190 nm (Fig. 4B). As
previously shown (14), there are negative CD bands in the 350-500-nm
range that coincide in wavelength with all of the absorption bands of
the chromophore. The visible wavelength bands represent only the FMN
chromophore, because there are no protein groups absorbing at these
wavelengths. In addition to these optically active chromophore bands,
there are CD-active positive and negative bands in the protein aromatic residue region (
In the far UV range (Fig. 4B), LOV2 shows a CD spectrum
typical of a protein that contains significant fractions of
The primary absorption of the exciton split
The CD spectrum for LOV2 in the far UV range (Fig. 4B) is
therefore expected to reflect primarily CD from the peptide bond and
thus reflects protein secondary structure. The spectrum is consistent
with the
The dashed lines in Fig. 4 correspond to
light-irradiated CD spectra (i.e. the CD spectra of the
FMN-cysteinyl adduct) in the visible and UV ranges. The visible/near UV
spectrum shows spectral changes consistent with the disappearance of a
450-nm species and the appearance of a 390-nm species. These changes are consistent with those reported previously (14). In the near UV
(aromatic) region, a large, positive band appears at about 290 nm, also
consistent with the previous report. Aromatic protein residues
typically contribute to CD in this region. The LOV2 fusion protein
contains several aromatic residues, which absorb maximally at about 280 nm but with low extinction coefficients. Upon formation of the adduct
the C(4a) carbon becomes a chiral center, which is expected to give
rise to new or increased optical activity. Because FMN absorption at
270 nm is substantially stronger than individual protein (aromatic)
absorption at 280 nm, the large positive CD band around 290 nm may be
dominated by the chiral chromophore contribution.
Upon adduct formation, the far UV CD spectrum remains qualitatively
similar to the dark state, but the ellipticity extremes (discussed above) lose 10-15% intensity. Shown in the
inset of Fig. 4B is the light minus dark
difference spectrum, which resembles an inverted Protein and Chromophore CD Bands of Light-irradiated LOV2 Relax
Synchronously in the Dark--
We also used CD spectroscopy to
determine whether protein structural perturbations could be
distinguished kinetically from chromophore activity during the
photocycle. As discussed above, changes in the visible and near UV
regions of the CD spectrum reflect primarily chromophore activity,
whereas changes in the far UV region probably reflect primarily protein
secondary structure. From these spectra, we selected two wavelengths
showing considerable changes upon light-irradiation from each region,
namely 450, 390, 222, and 208 nm. We then monitored dark relaxation of
the irradiated state of LOV2 by CD spectroscopy to see whether the
expected protein bands (208 and 222 nm) show different rates from the
chromophore bands (390 and 450 nm). As shown in Fig.
6, relaxation monitored at each of the
four key wavelengths showed the same rate, with a half-life of about
45 s, consistent with absorption measurements. Relaxation was also
monitored at these wavelengths in D2O, resulting in a
slowing of the rate by a factor of about 3 (Fig.
7), consistent with our previous
absorption measurement (15). No changes in CD were observed in control
experiments in which dark relaxation was monitored at 250 and 500 nm,
where no light-induced absorption changes occur in LOV2 (data not
shown). Finally, the decay profile at 222 nm was also measured at 10 and 4 °C and was found to fit a single exponential with a rate of
decay about 3 and 9 times slower than at room temperature (data not
shown), respectively.
D/H Exchange Demonstrates That the Rate-limiting Steps
in Both FMN-cysteinyl Adduct Formation and Decay Involve Proton
Transfer(s)--
Hydrogen transfer reactions require the
breakage of a bond between hydrogen and the donor atom and the
formation of a new bond between hydrogen and the acceptor atom.
Deuterium transfer reactions are generally slower due to the larger
mass of deuterium relative to hydrogen. Such kinetic slowing, referred
to as a primary deuterium isotope effect, leads to a relatively large
effect on the observed reaction rate, especially in the case of
rate-limiting proton transfer(s) (30-32).
Secondary isotope effects, which include hydrogen-bonding and
structural solvent effects, also contribute to observed solvent kinetic
isotope effects (30). Such effects are often quite small in magnitude
compared with primary effects. However, the effects are multiplicative,
and therefore a large number of secondary effects can cumulatively
contribute considerably to the observed kinetic isotope effect.
The results presented show that FMN-cysteinyl adduct formation in phot1
LOV2 shows a large (5-fold) solvent D/H exchange kinetic effect,
suggesting that formation or breakage of bonds involving hydrogen atoms
are rate-limiting steps in this reaction. The crystal structure of phy3
LOV2 (19) indicates that multiple hydrogen bond perturbations occur in
response to light, possibly constituting secondary isotope effects that
are reflected in the observed 5-fold decrease in rate. No significant
D2O-induced structural changes were observed by absorption
and CD spectroscopies in the visible and UV ranges (data not shown),
indicating that no secondary effects arise from this source.
The data presented here show that the rate of triplet decay in LOV2C39A
shows a minimal D2O effect. The LOV2C39A mutant lacks the
sulfur group necessary to form the adduct and therefore decays directly
from the triplet state to the ground form (15). The observed
D2O independence therefore indicates that triplet decay to
the ground state does not involve rate-limiting proton transfers in the
absence of the sulfur group.
The data presented also show that the triplet state in wild type LOV2
decays equally to the ground form and to the adduct form in
D2O, although the respective rates are slowed about 5-fold relative to H2O. The preservation of the 50/50 split is
consistent with our previous observation that the relative quantum
yields of adduct formation are the same in H2O and in
D2O (15). These data indicate that the processes of triplet
decay to the ground form and adduct formation share a common
rate-limiting step involving proton transfer(s). It is possible that a
single molecular event, such as thiolate attack at C(4a) triggered by
N-5 protonation, results in either adduct formation or intersystem
crossing to the ground form. Such a mechanism is consistent with the
observed 50/50 split between triplet decay to the ground form and
adduct formation in both H2O and D2O. In the
absence of this putative molecular event, such as when the sulfur group
is not present, adduct formation would be precluded and triplet decay
to the ground form would proceed by other (slower) processes. The
observation that triplet decay in the LOV2C39A mutant is slow (time
constant of 80 µs) is also consistent with a mechanism involving such
a single molecular event.
The rate of decay of the adduct form measured by CD spectroscopy in the
chromophore and protein spectral regions was found to be 3 times slower
in D2O (data not shown) than in H2O. Further, this 3-fold rate effect is identical to that observed by light-induced absorbance (15) changes of the chromophore. These findings indicate that protein and chromophore relaxation processes are governed by
common, rate-limiting proton transfer reaction(s). In addition, from
the temperature dependence of the rate constants in H2O, we
calculated that the activation energy barrier for the proton transfer
rate-limiting step of adduct decay is about 55 kJ/mol, assuming
Arrhenius behavior. This activation energy is within the energy range
of a few hydrogen bonds.
The Minimal pH Sensitivity for FMN-cysteinyl Adduct Formation and
Decay Suggests That Important Groups in the Rate-limiting Steps Have pK
Values outside the Measured Range--
Within the pH range studied, we
observed neither significant changes in the absorption spectrum of the
flavin chromophore in LOV2 nor substantial changes in the kinetics of
adduct formation or adduct decay. Titration of residues that are
involved in mechanistically relevant proton transfer(s) would be
expected to show a relatively large rate effect, which was not observed
for adduct formation or decay in the pH range studied. A broader pH
range could not be tested due to slow release of the chromophore and/or
protein precipitation that occurs at high and low pH.
The apparent pH independence of adduct formation between pH 5.5 and pH
9.5 was not unexpected, because most titratable protein groups
including glutamate, aspartate, lysine, tyrosine, and arginine, have
normal pK values outside this range. We have shown evidence that Cys-39 of LOV2 has an atypical pK of less than 4 (15). In addition
to cysteine, the only other relevant group that shows apparent
pK values in proteins in the range 5.5-9.5 is histidine (pKa ~6). The absence of significant pH effects on
adduct formation indicates that either these groups do not interact
with the chromophore and their ionization changes are inconsequential, or they also exhibit in LOV2 unusual pKa values
outside our experimental range. An additional possibility is that these groups are isolated from the bulk solution protons. The latter is
unlikely because of the overwhelming evidence that the chromophore pocket is reachable by bulk protons and bulk deuterons; (a)
we observed the titration of the N-3 proton of FMN in LOV2 at a
pK almost identical to the solution value (15),
(b) D/H exchange at N-3 and N-5 of the FMN chromophore was
observed by Fourier transform infrared (20), and (c) we have
observed that a molecule as large as iodoacetamide (but not
N-phenylmaleimide (14)) can eventually react with Cys-39 in
LOV2, releasing the FMN chromophore, whereas in the LOV2C39A mutant the
FMN chromophore is stable to such treatment (15).
The small pH effect on the adduct decay kinetics shows an apparent
pKa of about 6.8. This value is in the range of typical pK values for histidine (as discussed above) and for
the first ionization of phosphate on the FMN chromophore and is at the
extreme of the pK range for the deprotonation of typical
carboxyls. Protonation of the FMN phosphate could cause some structural
change in the chromophore binding pocket or in the chromophore binding affinity, resulting in this slight kinetic perturbation. Previous modeling of oat phot1 LOV2 (14) and comparison with the crystal structure of phy3 LOV2 (29) indicates that the FMN phosphate is located
near the molecular surface and thus is likely to be accessible to bulk
medium. At this time, we cannot identify a histidine or carboxyl group
neighboring the FMN, suggesting that the 40% slowing of adduct decay
at low pH is likely to be due to protonation of the FMN phosphate.
Does the weak pH sensitivity of decay of the adduct form indicate that
the mechanism is not base-catalyzed? The rates of general acid/base
catalysis reactions typically show pH dependences of 1 order of
magnitude or more. A pH dependence of this magnitude was reported for
adduct decay of C. reinhardtii phot LOV1 (33) and was
interpreted to indicate that the adduct decay reaction is
base-catalyzed. However, a structure for it has not been solved, and
the identity of a potential catalytic residue has not been proposed. In
contrast, the crystal structures of dark- (29) and light-irradiated
(19) phy3 LOV2 indicate an absence of basic residues near the
chromophore. The back reaction of phy3 LOV2 was not proposed to be
base-catalyzed (29), presumably for this reason. Given the high degree
of sequence homology between the phy3 LOV2 and oat phot1 LOV2 (Fig.
5B), it might be expected that the latter also has no basic
residues that are clear candidates for catalysis of the back reaction.
However, as shown, the LOV2 fusion protein in this study contains over
50 native phot1 residues that are not present in the crystallized
domain, allowing for the possibility of currently unidentified
contributions to the photocycle. In fact, adduct decay in the phy3 LOV2
domain used for crystallography showed a time constant for adduct decay
in solution (19) that is about 6 times slower than that observed for
our oat LOV2 fusion protein. This strongly supports the idea that
residues outside the 107-residue LOV2 domain (11) are important in the
photocycle. Hence, it is possible that the phot1 LOV2 back reaction is
base-catalyzed with participation from protein residue(s) outside the
LOV2 domain itself and that the relevant pK value lies
outside the pH range studied.
The Circular Dichroism Spectral Changes Suggest Reversible Protein
Structural Changes That Share a Common Rate-limiting Step with
Chromophore--
We have shown that the far UV CD bands change with
the same kinetic profile as that of the visible bands, suggesting a
common rate-limiting step for protein and chromophore structural
changes. The same phenomenon has been observed for photoactive yellow
protein (34), which is a PAS domain protein that contains a
p-coumaric acid chromophore and shows a folded structure
similar to that of phy3 LOV2 (29). It was observed by CD spectroscopy
that photoactive yellow protein loses a substantial amount of its
tertiary fold and a small amount of secondary structure upon light
irradiation, and these structural changes relax back with the same time
constant as observed for the p-coumaric acid chromophore
(34).
We considered the possibility that the light-induced far UV CD changes
observed for LOV2 are exclusively contributed to by the chromophore.
Although free FMN shows extremely minor CD activity in far UV range, as
discussed above, it is conceivable that FMN in the adduct form acquires
absorption and strong CD activity in this wavelength region. In this
case, the difference spectrum shown in the inset of Fig.
4B would represent the change in CD of the chromophore.
However, as discussed above, it is unlikely that FMN could acquire the
extremely large magnitude (at least a 100-fold increase) that would be
needed to compete with the CD activity of the ~200 amino acid
residues. Further, the double minimum present in both the far UV CD
spectrum and the far UV CD difference spectrum is strongly indicative
of helical structure and of mixed helix/sheet structure. A CD profile
from FMN in the adduct form possessing coincidentally the identical
electronic transitions with similar relative proportions is unlikely.
Finally, light-induced protein structural perturbations in oat phot1
LOV2 are indicated by NMR (18) and by infrared (20).
We also considered the possibility that the far UV CD difference
spectrum includes contributions from N- and C-terminal residues outside
the LOV2 domain. As discussed above, the LOV2 fusion protein used in
this work contains an expression vector segment and calmodulin-binding peptide at the N terminus of the LOV2 domain plus an additional 59 native phot1 residues (Fig. 5A). Protein secondary
structural changes revealed by CD spectroscopy in the far UV range
could include structural perturbations of these regions as well as of the LOV2 domain itself. If this were the case, remarkable signal transduction capability (from chromophore to extraneous protein segments) would be indicated. Indeed, fusion proteins containing both
LOV1 and LOV2 domains from Arabidopsis (35), with LOV1 mutated to C39A, did show kinetic differences from the LOV2 fusion protein used here, suggesting that extension of the N-terminal region
of LOV2 may impact light-induced protein behavior. We therefore interpret the observed light-induced far UV CD changes to reflect changes in protein secondary structure. These protein structural changes may be part of the signal transduction mechanism for oat phot1 LOV2.
-helix spectrum,
suggesting that
-helicity is reversibly lost upon light irradiation.
Decay kinetics for CD spectral changes in the far UV region occur at
the same rate as those in the visible region, indicating synchronous
relaxation of protein and chromophore structures.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-helicity during the photocycle. We also show that the light-induced changes in the far UV
CD spectrum relax on the same time scale as those in the visible region.
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
447(LOV2) = 13,800 M
1
cm
1 (14).
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
A, difference spectra of phot1 LOV2 in
D2O at 0.03, 0.13, 0.33, 1, 3, 10, 30, and 100 µs and 690 ms. The 30-ns spectrum has the highest absorption intensity at 660 nm
and maximum bleach at 450 nm; the arrows indicate the
direction of amplitude change over time. The inset
illustrates the nearly 5-fold slowing of triplet decay in
D2O ( ) relative to H2O (
) calculated by
the decay of the respective 660-nm absorption bands. B,
results of global exponential fitting of difference spectra from
A. The b-spectrum b1 has
an apparent time constant of 14 µs, and b0 is
the difference spectrum of the product formed.
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Fig. 2.
A, difference spectra of the LOV2C39A
mutant in D2O scanned at 0.03, 0.13, 0.33, 1, 30, and 100 µs and 1 ms; the arrows indicate the direction of
amplitude change over time. The inset shows that the triplet
decay rate is nearly identical in D2O ( ) and in
H2O (
), calculated by the decay of the respective 660-nm
absorption bands. B, results of global exponential fitting
of difference spectra from A. The b-spectrum
b1 has an apparent time constant of 80 µs, and
b0 is the difference spectrum of the product
formed (none).
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Fig. 3.
Minimal pH sensitivity of the rate of adduct
formation (A) and adduct decay
(B). The line in B
represents a Henderson-Hasselbach fit to the data and indicates a
pK of 6.8.
max ~280 nm), where flavin
chromophores have substantially stronger absorption (
max
~270 nm) than individual aromatic protein residues.
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Fig. 4.
CD spectra of LOV2 in the visible/near UV
(A) and far UV (B) ranges.
Solid lines represent ground state spectra
(LOV2D450), and dashed
lines represent light-irradiated spectra
(LOV2S390). The inset of
B shows the far UV CD light minus dark difference spectrum,
which resembles an inverted -helix.
-helical and
-sheet structure (26). The peptide bond has strong absorption bands at about 215 nm and below 200 nm. CD bands in this region arise
from electronic transitions of the peptide bond in particular structural conformations. The observed double minimum at 222 and 208 nm
and maximum around 195 nm are signature features of helical secondary
structure. These bands represent the n
* transition (~222 nm) and exciton split
* transition (~190 and 208 nm) of peptide bonds and have high extinction coefficients in the
-helical conformation. For the
-sheet conformation, the
transitions occur at 215 nm and at 175 and 198 nm, respectively, with
lower extinction coefficients than observed for the
-helix conformation.
* transition
occurs around 190 nm, such that its intensity is typically stronger than that at 208 nm. This ratio is not observed in the absolute far UV
spectrum of LOV2, suggesting that there are contributions to the
spectrum other than
-helix and
-sheet. The LOV2 fusion protein
used in this work, shown in Fig.
5A, contains a 46-residue segment from the cloning expression vector and calmodulin-binding peptide, plus 10 native oat phot1 residues, that are N-terminal to the
107-residue LOV2 domain (11) and 50 additional native oat phot1
residues C-terminal to the LOV2 domain. The peptide bonds of these
additional residues contribute to the far UV CD spectrum. The FMN
chromophore is also expected to contribute to the spectrum, although
only weakly. Free FMN in aqueous and organic solvents shows a positive
CD band (
max ~222 nm) that is similar in magnitude to
a single peptide bond (this work, data not shown) (27, 28). For
comparison, Fig. 4B shows the molar ellipticity (CD
magnitude) normalized to the concentration of protein (left axis) and peptide bonds (right axis).
The LOV2 domain binds a single FMN chromophore such that the protein
concentration is the same as the FMN concentration, whereas the peptide
bond concentration is about 200 times larger because there is about one
peptide bond per amino acid residue. The right
axis in Fig. 4B, which represents the per residue
CD intensity, is therefore about 200 times smaller in magnitude than
that of the left axis. In order for FMN to make a
significant contribution to the LOV2 far UV spectrum, the protein-bound FMN would have to acquire a CD magnitude large enough to compete with
about 200 amino acid residues; in other words, the FMN would have to
increase its CD magnitude by a few orders of magnitude.
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Fig. 5.
A, sequence of the oat phot1 LOV2 fusion
protein. There are 46 nonnative residues from the pCAL-n-EK expression
vector and the calmodulin-binding peptide at the N terminus and 7 residues from the expression vector at the C terminus. The defined LOV2
domain spans residues 413-520. The conserved cysteine residue is
shaded. B, sequence alignment of oat phot1 LOV2,
fern phy3 LOV2, and C. reinhardtii phot LOV1. Conserved
residues are shaded. The segment of phy3 LOV2 studied by
crystallography (19, 29) is indicated by arrowheads.
+
tertiary structure classification (26), which is
consistent with PAS domain structures in general and specifically with
the predicted structure of oat phot1 LOV2 (14) and the crystal
structure of phy3 LOV2 from the fern Adiantum capillus-veneris (29).
-helix spectrum,
suggesting that a small amount of helicity is lost upon adduct
formation. As discussed above, the
-helix conformation has the
highest molar absorptivity at its characteristic wavelengths, meaning
that other types of secondary structure that exist in only moderate
amounts contribute much less to the observed CD spectra. Therefore, the
10-15% loss of intensity of the far UV CD bands upon light
irradiation of LOV2 would correspond to a maximum loss of 10-15%
helicity. We fit the far UV CD difference spectrum to a linear
combination of CD reference spectra (26) of various types of secondary
structure. The fits were not unique (did not favor a particular
combination of formed secondary structure); however, the loss of
10-15% helicity was necessary in all fits. The crystal structure (29)
indicates that the LOV2 domain contains ~25%
-helical structure,
so the overall change observed by CD is 10-15% of this
(i.e. about 3% of the LOV2 domain secondary structure).
However, the fusion protein used in this work has almost twice the
molar mass of the crystallized LOV2 domain, and the secondary structure
of the additional protein segments is unknown. Therefore, we cannot
quantitate the fraction of lost helicity for the total fusion protein.
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Fig. 6.
Dark relaxation of the LOV2 adduct monitored
by circular dichroism at visible, near UV, and far UV wavelengths.
Symbols represent raw data, and lines are
first-order exponential fits that gave relaxation half-lives of 46 s at 450 nm, 47 s at 290 and 222 nm, and 51 s at 208 nm.
These values are consistent with the half-life measured from absorption
difference spectroscopy (45 s), which monitors only chromophore
activity.
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Fig. 7.
The D2O effect on dark relaxation
of the LOV2S390 (adduct) measured by circular
dichroism. The line in H2O represents
relaxation monitored at 450 nm. All four wavelengths monitored showed
the same 3-fold slowing in D2O, further suggesting
that protein and chromophore relaxation share a common rate-limiting
step. The single exponential fits calculated by SigmaPlot (see
"Materials and Methods") showed correlation coefficients of at
least 0.98 with S.E. values in the relaxation half times of 14 ± 4%.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. John Christie for critical reading of the manuscript and for assistance with Fig. 5. We also thank Dr. David Kliger for generously allowing use of the spectroscopy facilities.
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FOOTNOTES |
---|
* This work was supported by faculty research funds granted by the University of California, Santa Cruz (to R. A. B.), research funds provided by Covalent Partners LLC (to R. A. B. and S. B. C.), and National Science Foundation Grants IBN9940546 and MSB-0091384 (to W. R. B.) and DMB-0090817 (to R. A. B.).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.
¶ To whom correspondence should be addressed: Dept. of Chemistry and Biochemistry, University of California, Santa Cruz, 1156 High St., Santa Cruz, CA 95064. Tel.: 831-459-4294; Fax: 831-459-2935; E-mail: bogo@chemistry.ucsc.edu.
Published, JBC Papers in Press, October 30, 2002, DOI 10.1074/jbc.M209119200
1 Position 39 in the isolated LOV2 domain of oat phot1 (14) corresponds to position 450 in the full-length sequence.
2
In this work, the time constant (denoted ) is
used interchangeably with lifetime, which refers to 63%
(e.g. (1/e)) reaction completion and is equal to
the reciprocal of the apparent rate constant,
kapp. Accordingly, for branched reactions,
kapp = 1/
app = 1/
1 + 1/
2 + ... + 1/
n and
app = (t1/2) (ln 2).
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