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Originally published In Press as doi:10.1074/jbc.C400137200 on April 19, 2004

J. Biol. Chem., Vol. 279, Issue 23, 23855-23858, June 4, 2004
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Resonance Raman Evidence for Two Conformations Involved in the L Intermediate of Photoactive Yellow Protein*

Masashi Unno{ddagger}§, Masato Kumauchi¶, Norio Hamada||, Fumio Tokunaga¶, and Seigo Yamauchi{ddagger}§

From the {ddagger}Institute of Multidisciplinary Research for Advanced Materials, Tohoku University, Sendai 980-8577, Japan, the Department of Earth and Space Science, Graduate School of Science, Osaka University, Toyonaka, Osaka 560-0043, Japan, and ||JST, CREST, Osaka University, Suita, Osaka 565-0871, Japan

Received for publication, April 2, 2004 , and in revised form, April 14, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 REFERENCES
 
The blue light receptor photoactive yellow protein (PYP) displays a photocycle that involves several intermediate states. Here we report resonance Raman spectroscopic investigations of the short-lived red-shifted intermediate denoted PYPL. We have found that the Raman bands of the carbonyl C=O stretching mode {nu}11 as well as the C=C stretching mode {nu}13 for the chromophore can be resolved into two peaks, and the ratio of the two components varies as a function of pH with pKa ~6. The isotope effects on the resonance Raman spectra have confirmed a deprotonated cis-chromophore for the two components. The results indicate the presence of two conformations in the active site of PYPL. The normal coordinate calculations based on the density functional theory provide a structural model for the two conformations, where the low pH form is possibly an active structure for the protonation reaction generating a following intermediate in the photocycle.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 REFERENCES
 
The photoactive yellow protein (PYP)1 from the purple bacterium Halorhodospira halophila is a small water-soluble photoreceptor (1), and it has been an attractive model for studying protein structures and dynamics. Recently PYP gained further attention as the structural prototype for the PAS (Per-ARNT-Sim) and LOV (light, oxygen, or voltage) domains of a large class of receptor proteins (2). In a dark state (PYPdark), the phenolate anion of the trans-4-hydroxycinnamyl chromophore (3, 4) is stabilized by hydrogen bonds with Tyr42 and Glu46 (5, 6). Lowering the pH induces a formation of a bleached state denoted PYPM,dark, which contains protonated trans-chromophore (6). Photoexcitation of PYPdark triggers a photocycle that involves two major intermediate states denoted PYPL (also called I1 or pR) and PYPM (also called I2 or pB) (7, 8). Although it is well established that the PYPL -> PYPM process involves protonation of the phenolic O1 (9, 10), it is still not clear what structural factors promote the protonation reaction. To address this issue, we have performed resonance Raman (RR) investigations of PYPL and report a novel finding for two conformations in the active site of this intermediate state. The two conformations are in a pH-dependent equilibrium with pKa ~6, and a normal coordinate analysis based on the density functional theory (DFT) indicates that the low pH conformation lacks a hydrogen bond between Glu46 and the phenolic O1 (Fig. 1). We propose that the newly found low pH form is an active conformation for the protonation reaction during the PYPL -> PYPM process, whereas the high pH form is a direct photointermediate from PYPdark.



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  		FIG. 1.
Proposed active site structure of PYPL at neutral pH (Ref. 10).

 

   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS AND DISCUSSION
 REFERENCES
 
Sample Preparations—Expression of wild-type (WT) and Glu46 -> Gln (E46Q) mutant apoproteins from Escherichia coli, chromophore reconstitution, and protein purification were performed as described previously (11). PYP in buffered D2O (90% D2O/10% H2O) was prepared by proper dilution of a concentrated protein into D2O, and then the sample was incubated overnight at room temperature before the measurements. The 13C-labeled 4-hydroxycinnamic acids were synthesized as described previously (12). 13C-Labeled PYPs were prepared by reconstitution of apoprotein with 4-hydroxycinnamic anhydride whose carbonyl carbon atom (C9) or ring carbon atoms (C1–C6) were labeled with 13C. These labeled samples are denoted as 13C=O and 13C6-ring isotopomers, respectively.

Resonance Raman Spectroscopy—RR spectra were obtained as described previously (6, 10, 12, 13, 14). A liquid nitrogen-cooled CCD detector (Instrument S. A., Inc.) recorded the Raman spectra after a Triax190 spectrometer (Instrument S. A., Inc.) removed the excitation light, and a Spex 500M spectrometer dispersed the scattered light. Samples were excited with the 413.1 nm line available from a krypton ion laser (BeamLok 2065, Spectra-Physics Lasers, Inc.). The measurements were made on samples contained in a quartz spinning cell (10-mm diameter, 800 rpm), and a transit time of the sample in the beam was ~200 µs. We performed the measurement of PYPdark with a low laser power (0.24 mW), while the higher laser power (2.9 mW) was used for PYPL. The lifetimes of the photocycle intermediates preceding the PYPL are shorter than that of the PYPL more than 50-fold (15), so that their populations are negligible under the present experimental conditions. RR spectra of PYPL were obtained by subtracting a fraction of the normalized low power spectrum from the normalized high power one such that the sharp {nu}11 mode (1631 cm–1) of PYPdark disappeared without introducing spurious derivative band shapes or negative peaks. Note that a higher laser power (8.9 mW) was used to measure the high power spectrum in our previous work (10). We have found, however, that the spectrum of PYPL exhibits laser power dependence, e.g. the frequency of {nu}25 (see below) was affected by a high laser power, possibly due to a formation of a photoproduct of PYPL. Thus we used lower laser powers in the present study.

DFT Calculations—The optimized geometry and the harmonic vibrational frequency were calculated via Gaussian98 program (16). The hybrid functional B3LYP and the 6-31G** basis set were used for the DFT calculations. The calculated frequencies were scaled using a factor 0.9613. For some cases, we used the Onsager reaction field method (17) to take into account the dielectric constant in protein.


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
REFERENCES
 
Effects of pH on the Resonance Raman Spectra of PYPL—Fig. 2 depicts the transient RR spectra of WT PYPL with a 200-µs time resolution at pH 8.1–4.5 (traces a–e). The RR spectrum at neutral pH (trace b) is in agreement with that reported previously (10, 14, 18), and the assignments of important Raman bands are designated in the figure (10, 12).2 Fig. 2 also demonstrates significant changes in the spectrum as a function of pH. Above pH 7.0 (traces a and b), a carbonyl C=O stretching vibration {nu}11 and a coupled C=C stretching mode {nu}13 are characterized by a peak at 1672 cm–1 with a shoulder of ~1655 cm–1 and a peak at 1553 cm–1 with a shoulder of ~1575 cm–1, respectively. Lowering pH increases the intensity of the shoulders, and both {nu}11 and {nu}13 bands become unresolved doublets at pH 4.5. Similarly, a coupled C=C stretching mode {nu}14 at 1530 cm–1 and a C=C stretching/C–H rocking mode {nu}17 at 1441 cm–1 are upshifted by ~10 cm–1 upon lowering pH (Table I). We note that the RR spectrum of PYPdark exhibits little changes in the examined pH region (see supplemental Fig. 1S). In Fig. 2, we also find significant effects of Glu46 -> Gln (E46Q) mutation on the spectrum of PYPL (trace f). Similar to the case of lowering pH, E46Q mutation causes an increase in intensity around 1655 ({nu}11) and 1576 cm–1 ({nu}13) as well as an upshift of {nu}14 and {nu}17.



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  		FIG. 2.
Resonance Raman spectra of WT and E46Q mutant of PYPL. The sample concentration was 80 µM. The following list gives a buffer composition and pH: 10 mM Tris-HCl, pH 8.1 (trace a); 10 mM MOPS, pH 7.0 (trace b); 20 mM MES, pH 6.0 (trace c); 20 mM acetate, pH 5.0 (trace d); 20 mM acetate, pH 4.5 (trace e); and 10 mM MOPS, pH 6.5 (trace f).

 


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  		TABLE I
Observed and calculated vibrational frequency (cm–1) of PYPL and its models

 
Fig. 3A shows the {nu}11 region of the RR spectra of PYPL at pH 7.4, whose chromophore is unlabeled or labeled with 13Catthe carbonyl carbon atom, as well as the results of peak-fitting. The 12C/13C isotope substitution downshifts the whole doublet by ~35 cm–1, demonstrating that both components originate from the {nu}11 mode. This observation implies that there are two conformations where the {nu}11 frequency is different. Here we denote the high and low pH conformations as PYP hL and PYP lL, respectively. In Fig. 3B, the integrated intensities of the {nu}11 band for PYP lL are plotted relative to that for PYP hL by changing pH and fitted to the Henderson-Hasselbalch equation (19) with pKa = 6.0 ± 0.4. We also plot analogous relative intensities for {nu}13 in the figure. The latter plot is fitted with pKa = 6.4 ± 0.2. The reasonable agreement of the two pKa values indicates that the observed spectral changes are due to a pH-dependent equilibrium between two species, PYPLh and PYPL1. To examine the chromophore structure for these conformations, we have performed further investigations as described below.



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  		FIG. 3.
A, resonance Raman spectra of WT PYPL at pH 7.4 in the {nu}11 region. The spectra for natural abundance (trace a) and 13C=O samples (trace b) are shown. The peaks around the {nu}11 region are fitted by using two Voigt functions with a fixed width (FWHM = 15 cm–1) on a polynomial base line. B, pH dependence of the relative peak area of {nu}11 ({circ}) (A1655/A1672) and {nu}13 () (A1576/A1553).

 
Protonation State of Chromophore—The deuteration effects on the {nu}33 band around 720 cm–1 and the {nu}32 + 2{gamma}12 doublet around 850 cm–1 act as markers for the protonation state of the chromophore (6, 12). These Raman bands are examined in Fig. 4A, which displays the 600–950 cm–1 region of the RR spectra of WT PYPL at pH 7.4 and those of the E46Q mutant at pH 6.5. These samples contain mainly PYPLh and PYPL1, respectively. For comparison, the figure also illustrates the spectra of WT PYPM (12). The {nu}33 mode is observed at ~720 cm–1 for the protonated chromophore in PYPM,dark and PYPM (traces f–h) (6, 12). Because this mode includes O1-C1 and C4-C7 stretching coordinates such as Y13 of tyrosine (20), it exhibits ~3 and ~10 cm–1 downshifts upon deuterium substitution of the hydroxyl group (trace f -> g) and 13C6-ring labeling (trace f -> h), respectively. The {nu}32 + 2{gamma}12 doublet is observed as a broad Raman band at ~840 cm–1 for PYPM,dark and PYPM (6, 12). This doublet arises from the Fermi resonance between the ring-breathing vibration {nu}32 and the overtone of an out-of-plane ring-bending vibration {gamma}12. Because of the removal of the Fermi resonance, the band shape of the doublet significantly changes upon deuteration of the phenolic OH group (trace f -> g) or 13C6-ring substitution (trace f -> h).



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  		FIG. 4.
A, resonance Raman spectra of PYPL and PYPM in the {nu}33 and {nu}32 region. The experimental conditions are the same as those described in the legend to Fig. 2. The spectra for PYPM are taken from Ref. 12. B, resonance Raman spectra of PYPL at pH 7.4 and 5.5 in the {nu}29 region.

 
In the case of WT PYPL at pH 7.4, we assign a band at 737 cm–1 and a broad feature around 840 cm–1 to {nu}33 and the {nu}32 + 2{gamma}12 doublet, respectively, on the basis of their –11 and –22 cm–1 13C6-ring shifts (trace a -> c). In contrast to PYPM,dark and PYPM, the deuterium substitution of exchangeable protons little affect these Raman bands for WT PYPL at pH 7.4 (trace a -> b). Analogous experimental data for the E46Q mutant (trace d -> e) do not exhibit any significant H/D isotope substitution effects in the 600–950 cm–1 region nor do WT PYP at pH 5.5 (not shown). This demonstrates that the chromophore is deprotonated for the PYP hL and PYP lL species. We can therefore exclude a possibility that the low pH component observed here is the other intermediate such as PYPM or its preceding intermediate (21), whose chromophore is protonated (10, 12, 13, 18). In addition, the extinction coefficient of these blue-shifted intermediates at the 413.1 nm excitation is essentially zero (8, 22), so their contribution to the RR spectra will be negligible.

We comment on the other Raman bands that are sensitive to the protonation state of the chromophore. Previous Raman studies on PYP and a chromophore model (4, 6, 10) have shown that the protonation state of the chromophore affects the frequency of the {nu}25 and {nu}14/{nu}13 modes. However, these modes are also sensitive to different factors such as trans/cis isomerization of the chromophore (10) or the dielectric constant in the vicinity of the chromophore (6). Thus the deuteration effects on {nu}33 and the {nu}32 + 2{gamma}12 doublet are more reliable to determine the protonation state of the phenolic OH group. Another possible marker for the chromophore protonation is the combined C1-O1 stretching and HC7 = C8H bending modes {nu}22 and {nu}23 near 1250–1300 cm–1 (12). Although these modes were recently used to characterize the chromophore structure in PYPL (18), a strong overlap with the nearby Raman bands complicates the interpretation of the data. We will discuss the details of the assignments in this region elsewhere.2

Isomerization State of Chromophore—Fig. 4B shows the 900–1150 cm–1 region of the RR spectra of WT PYPL. The spectra of natural abundance (trace i) and 13C=O-labeled (trace j) PYPs at pH 7.4 and their difference spectrum (trace k) confirm a previous result (10) that the C8-C9 stretching mode {nu}29 appears as a Raman band near 1000 cm–1. The figure also demonstrates the data for WT PYP at pH 5.5 (traces l–n). The {nu}29 frequency is sensitive to trans/cis isomerization around the C7=C8 bond (10, 12). The trans configuration exhibits the {nu}29 band at ~1050 cm–1, whereas {nu}29 for the cis form is 990–1000 cm–1. As can be seen in the figure, lowering pH does not affect the {nu}29 frequency, indicating a cis configuration chromophore both for PYP hL and PYP lL.

Structural Models for PYP hL and PYP lL—As discussed above, the results presented in Fig. 4 indicate that both PYP hL and PYP lL contain the deprotonated cis-chromophore. This implies that the two conformations differ in chromophore-protein interactions. To discuss the observed differences, we have performed normal coordinate calculations based on DFT. For these calculations, deprotonated cis-4-hydroxycinnamyl methyl thiolester was employed as a chromophore model. In addition, methanol and acetic acid were included to mimic the hydrogen bonds of the phenolic O1 with Tyr42 and Glu46, respectively. These components were arranged on the basis of the crystal structure of a cryotrapped intermediate preceding PYPL (23) and subsequently optimized to yield the structure illustrated in supplemental Fig. 2S and Table 1S (model 1). This model has been shown to be a good model for the active site of PYP hL (10), and the calculated frequencies are summarized in Table I.

We used the following optimized structures as models for PYP lL (supplemental Fig. 2S and Table 1S). Models 2 and 3 contain only one hydrogen bond with Tyr42 and Glu46, respectively. The two hydrogen bonds are removed in model 4, whereas model 5 considers an additional hydrogen bond at the carbonyl O2 with water. Model 6 is similar to model 1, but the rest of the protein is treated by continuum electrostatics with {epsilon} = 20. As can be seen in Table I, model 2 explains the main features for PYP lL, i.e. the removal of a hydrogen bond with Glu46 shifts {nu}11 and {nu}13 by –9 and +12 cm–1, respectively. Furthermore, the observed upshifts of {nu}14 and {nu}17 by ~10 cm–1 are consistent with the calculations. In contrast, the other models failed to reproduce the difference between PYP hL and PYP lL, suggesting that model 2 is a plausible active site structure of PYP lL. The observation that the RR spectrum of E46Q PYP is similar to that of WT PYP lL (Fig. 2) supports an importance of the hydrogen bond between the phenolic O1 and Glu46. Note that model 4, where the hydrogen bond with Glu46 is also missing, explains some features of PYP lL, but the predicted effect of removing the two hydrogen bonds is significantly larger compared with the observed difference between PYP hL and PYP lL.

Here we should discuss a titratable group that regulates the equilibrium between PYP hL and PYP lL with pKa ~6. One of the possible candidates is Glu46 because of an importance of its hydrogen bond with the phenolic O1 of the chromophore. This explanation implies that the carboxyl group of Glu46 is protonated in PYP lL, whereas it is deprotonated in PYP hL. This idea, however, is unlikely, because previous Fourier transform infrared studies (21, 24, 25) showed that Glu46 is protonated at pH 7, where PYP hL is a dominant fraction. Another possible residue is His3 or His108, because the pKa of the imidazole group in aqueous solution is about 6 (19). However, it seems that these residues are situated in positions far a bit from the active site. For example, the {alpha} carbon of His3 and His108 is located more than 10 Å from the carboxyl group of Glu46 (3). Further experiments of PYP mutants are needed to identify a residue that controls the equilibrium between the two conformations.

The PYPL -> PYPM process involves the protonation of the phenolic O1 of the chromophore (9, 10, 12, 13, 18). In this study, we suggest that the absence or weakening of the hydrogen bond between the phenolic O1 and Glu46 in PYP lL. Because previous studies using site-directed mutagenesis showed that the removal of a hydrogen bond at the phenolic O1 favors a protonation of the chromophore (5, 6), the formation of PYP lL potentially promotes the protonation process. In fact, both lowering pH and E46Q mutation increase the fraction of PYP lL and accelerate the PYPL -> PYPM transition by a factor of 3–6-fold (26). In addition, although the actual PYPL -> PYPM transition exhibits a complex pH dependence (22), this process was characterized by a pKa around 6 (26). These results suggest that the photoexcitation of PYPdark initially produce PYP hL (10), and the low pH form PYP lL could be an active structure for the PYPL -> PYPM process. To examine this idea, further studies using time-resolved RR studies of WT and some PYP mutants are currently in progress.


   FOOTNOTES
 
* This work was supported by grants from the Association for the Progress of New Chemistry (to M. U.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1S and 2S and Table 1S. Back

§ To whom correspondence may be addressed. Tel.: 81-22-217-5618; Fax: 81-22-217-5616; E-mail: unno{at}tagen.tohoku.ac.jp.

1 The abbreviations used are: PYP, photoactive yellow protein; DFT, density functional theory; PYPdark, dark-state PYP; PYPL, red-shifted L intermediate of PYP; PYPM, blue-shifted M intermediate of PYP; PYPM,dark, acid-induced blue-shifted state of PYP; RR, resonance Raman; WT, wild-type; mW, milliwatt; MOPS, 4-morpholinepropanesulfonic acid; MES, 4-morpholineethanesulfonic acid. Back

2 M. Unno, M. Kumauchi, N. Hamada, F. Tokunaga, and S. Yamauchi, manuscript in preparation. Back


   ACKNOWLEDGMENTS
 
We are grateful to R. Nakamura (Osaka University) for helpful discussion and K. Yoshihara (Suntory Institute for Bioorganic Research) for assistance in preparing the 13C-labeled compounds.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 REFERENCES
 

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