Fourier transform resonance Raman spectroscopy of bacterial reaction center proteins: The primary electron donor

ELSEVIER

Journal of MOLECULAR STRUCTURE Journal of Molecular Structure 347 (1995) 459-466

Fourier transform resonance Raman spectroscopy of bacterial reaction center proteins: T h e p r i m a r y electron donor Tony A. MATTIOLI D6partement de Biologie Cellulaire et Mo16culaire, SBPM, CEA and URA CNRS 1290 C.E. de Saclay 91191 Gif-sur-Yvette cedex FRANCE 1. INTRODUCTION Photosynthesis involves the conversion of light energy into chemical potential energy. The primary events in photosynthesis occur in specialized membrane proteins known as reaction centers, RCs. RCs contain several cofactors which are, in general, chlorophyll and carotenoid molecules. Man.y of these cofactors can be selectively probed using resonance Raman spectroscopy using various excitation wavelengths. The physicochemical (e.g. spectroscopic, redox) properties of these cofactors are largely dependent on the protein environment and specific pigment-protein interactions. This type of information can readily be obtained by resonance Raman spectroscopy both in resting states and intermediate functional states of the proteins [1-3]. The structural elucidation of two purple photosynthetic bacteria, Rhodopseudomonas viridis [4] and Rhodobacter sphaeroides [5-8] has revealed the spatial arrangement of the cofactors that mediate the initial photoinduced transmembrane charge separation. The RC from Rb. sphaeroides 2.4.1 consists of three polypeptides named L, M, and H. The L and M polypeptides noncovalently bind several cofactors: a pair of excitonically coupled bacteriochlorophyll a (BChl a) molecules constituting the primary electron donor (P), two monomeric BChl a molecules, two .bacteriopheophytin a (BPhea) molecules, two ubiquinone molecules, a non-heme iron atom, and a carotenoid molecule. These bacteriochlorin and quinone cofactors are arranged in pairs along a C2 symmetry axis resulting in two branches called L and M. Despite these two possible branches, normal electron transfer occurs asymmetrically along the L branch of cofactors. Recently, we have shown that near infrared Fourier transform Raman spectroscopy, which usually utilizes 1064 nm as excitation wavelength, provides excellent Raman spectra of chlorophyll molecules in preresonance with their Qv absorption band [9-12]. Application of this technique to study purple bacterial reacti6n centers demonstrated that near IR excited FT Raman spectroscopy is an ideal tool to obtain the selective vibrational contributions of P in its resting and oxidized states [10,11]. These selective observations of the P and P +" vibrational modes permit the direct determination of the protein interactions with P as well as the degree of + charge localization on one of the two BChl a molecules constituting P upon its oxidation. Furthermore, application of this technique to mutant RCs where site-specific alterations have been genetically introduced in the P binding pocket permits the ready determination of changes in H-bonding interactions [13,14]. In the following, we present some recent work on the primary electron donor in native and mutant reaction centers of purple photosythetic bacteria. The direct vibrational information obtained on these systems using FT Raman spectroscopy allow us to identify specific pigment-protein interactions (e.g. hydrogen bonding) which change physicochemical properties of P. Results to date indicate that the chemical nature of the H459-466

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460 bond donor to conjugated carbonyl groups of P plays a significant r61e in the tuning redox properties of P. 2. METHODS Room temperature FT Raman spectra of RC proteins are recorded using a Bruker IFS 66 interferometer coupled to a Bruker FRA 106 Raman module equipped with a diodepumped Nd:YAG continuous wave laser. Typical laser powers used are 150-200 mW. Raman l~hotons are collected using a 180 ° backscattefing geometry and spectral resolution is 4 cm -1. Reaction center samples were usually held in a sapphire cell [15]. Low temperature experiments (10 K) are performed using a helium-circulating cryostat (SMC, France). Samples are deposited as droplets on a glass slide and frozen. No window separates the sample from the laser beam nor the helium flow. For the experiments involving variable near IR excitation, a cw Ti 4 + :laser (Spectra Physics 3900 S) pumped by an Ar + laser (Coherent Innova 100) was used. In these experiments, the FT Raman module was fitted with custom-made holographic notch filters (Kaiser Optical Systems, Inc.) to reject the Rayleigh line [16]. RC samples were poised in their P and p T . states by the addition of ascorbate and ferricyanide, respectively. This chemical treatment ensured that 100 % of the RCs were in the desired redox state during measurement. Typical concentrations used were ca. 0.5 mM of protein. 3. RESULTS AND DISCUSSION 3.1. Purple Bacterial Reaction Centers. Resonance Raman spectroscopy of reaction centers excited in the visible and near UV has recently been reviewed [17]. Excitation of purple bacterial reaction centers using 1064 nm offers two major advantages: i) the vibrational modes of P are preresonantly enhanced via its nea~ IR Qv absorption band at ca. 865 nm, and ii) the vibrational modes of the cation radical p T . are resonantly enhanced via its 1250 nm absorption band. 3.1.1. Preresonance FT Raman Spectra of P. Excitation of BChl a-containing RCs using 1064 nm results in a strong preresonant enhancement of the primary donor whose lower exciton Qy~ component absorbs at ca. 865 nm. The preresonance enhancement is greater for P than for the other two monomeric BChl a molecules which absorb at ca. 800 nm and greater still for the two BPhe a molecules found in the RC [11]. For RCs from Rb. sphaeroides R26 in which P is reduced, it has been estimated that at least 65 % of the Raman intensity of the BChl a modes arises from P [I0]. This means that the vibrational spectrum of reduced P may be directly observed without resorting to difference techniques. Furthermore, since 1064 nm excitation is essentially transparent for reduced RCs, the experiments may be performed at physiological temperatures for prolonged periods of time without sample degradation. The FT Raman spectrum, in the high frequency region, of reduced reaction centers is shown in Fig. 1. The features of this spectrum have been described in detail elsewhere [10,11,13,14]. A schematic diagram presenting the pigment-protein interaction structural model for the primary donor in Rb. sphaeroides as deduced from its FT Raman spectrum is also shown in Fig. 1. The vibrational frequency assignments of the acetyl carbonyl and keto carbonyl groups of of the two BChl a molecules (named PL and PM) constituting P were deduced from the X-ray crystal structure and FT Raman spectra; these assignments have been confirmed using site-directed mutagenesis [14]. In the native reaction center, only one conjugated carbonyl of P is observed to be engaged in a hydrogen bond, consistent with the X-ray crystallographic structure [6,8] which places a hydrogen bond on the C 2 acetyl carbonyl of PL donated by His L168. This H-bond, as judged by the vibrational frequency of the acetyl carbonyl observed in the FT Raman spectra, is quite strong

461 (estimated H-bond enthalpy of 0.207 eV [14]). The donor of this strong H-bond, namely histidine L168, is strictly conserved in the RCs of purple bacteria whose primal, amino acid sequences are known, and may be important in some structural or functional role.

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.

~

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Raman shift (cm-l) Figure 1. (Top) Schematic diagram presenting the pigment-protein interaction structural model for the primary electron donor in Rb. sphaeroides. Indicated are the vibrational frequency assignments of the acetyl and keto carbonyl groups as deduced from the crystal structure and FT Raman spectroscopy [14]. (Below) FT Raman spectrum of reaction centers from Rb. sphaeroides in the presence of ascorbate, excited using 1064 nm radiation [10]. The dominant contributions to this spectrum are from the primary electron donor in its ground, neutral state. See structure above for the assignments of the Pvaman bands.

462 3.1.2. FT Resonance Raman Spectrum of P + ". To date, excitation with 1064 nm radiation has been the only manner toj~bserve the resonance Raman spectrum of the primary ~lonor in its cation radical state, p T . [10,11]. Relative t 9 the observed maximum of the P-'-" absorption band at 1250 nm, 1064 nm is ca. 1400 cm -~ higher in energy and thus falls within the vibronic region of this absorption band; this constitutes a genuine resonance condition. Stqucturally informa[ive bands arising from P+" include those at 1600, 1641, and 1717 cm -~. The 1717 cm -~ band arises from the slretching mode of the C 9 keto carbonyl of PL [10,11] which has been upshifted by 26 cm -~ upon ,the one-electron oxidation of P. The magnitude of this oxidation-induced upshift of 26 cm -1 of the P ketp carbonyl band as compared to that of monomeric BChl a in nonprotic solvents, 32 cm -~ [18,19] indicates that the resulting + charge on the dimer is not equally shared but resides primarily on one of the two BChl a molecules izonstituting P, namely PL [10,11]. Assuming the 32 cm -~ upshift observed in vitro represents 100 % localization on a BChl a molecule and a linear dependence of the magnitude of the C 9 keto carbonyl upshift upon P oxidation and the degree of + charge localization, it was estimated that 80 % of the + charge in p T . resides on the PL cofactor [10] in Rb. sphaeroides carotenoidless strain R26. This value corresponds gratifyingly well with that of 68 % obtained using ENDOR/TRIPLE techniques [20] for another strain of Rb. sphaeroides. More recently, we have measured the FT resonance Raman spectra of RCs from the same strain which was used in the ENDOR studies and have obtained ca. 72 % + charge localization on PL [14], which is in quantitative agreement with the ENDOR measurements; this point illustrates the co_[nplementarity of the two techniques in obtaining electronic structural information of the p T . species. 3.2 Variable Wavelength Excitation At present, commercial FT Raman spectrometers provide only 1064 nm from a Nd:YAG laser as possible excitation wavelength. This restriction is usually dictated by the need of a very efficient but "non-tunable" notch filter to rej~cj_ the Rayleigh line from the Raman spectrum. Recently, we have reported the use of a Ti "*T'sapphire laser and custommade holographic notch filters to permit the excitation and recording of FT spectra from ca. 800 nm down to 1090 nm [16,21]. This apparatus is currently being used at Saclay as a sensitive near IR fluorescence and Raman FT spectrometer. In Figure 2 we show the FT Raman spectrum of an aspirin tablet, excited with 1064 nm radiation usj_ng the standard diqd$-pumped Nd:YAG laser and excited at 9 6 6 n m using an Ar T laser-pumped Ti'*T:saphhire laser. The 966 nm-excited spectrum was recorded using a grazing incidence geometry. This spectrum illustrates that FT Raman spectrum of highly scattering samples can be obtained using our apparatus. 3.3. Antenna-less M u t a n t Systems with Wildtype RCs. (Collaboration with M. Jones, University of Sheffield). We have recently used the above-described apparatus to obtain FT Raman spectra of reaction centers from Rb. sphaeroides in situ, in their native membrane. This is not easily achieved in wild-type systems because of the many bacteriochlorophyll pigments in the antenna proteins which are present in the photosynthetic membrane. However, using a genetically modified system [22] where the antenna proteins have been deleted this problem can be circumvented. In such antenna-less membranes, the only chromophores absorbing in the near IR are those from the reaction centers. Thus, exciting the Raman spectrum of these antenna-less membranes with 1064 nm radiation yields the contributions of the RC as well as some contributions of the membrane; the vibrational features of the primary donor are clearly visible [23]. By changing the excitation from 1064 nm to 940 nm there results a marked increase in the preresonance enhancement of the vibrational spectrum of P (absorbing at 860 nm). Under these conditions of excitation, the contributions of the membrane compared to P are greatly reduced. A comparison of low temperature (15 K) FT Raman spectra of RCs, both isolated and in their native membranes is shown in Figure 3. The bottom spectrum in Fig. 3 clearly

463

Aspirin tablet t,=~ i

b

,-p.

t

2

15'oo Raman

shift

[era-l]

Figure 2. Room temperature FT Raman spectra of an ja~pirin tablet excited at 1064 nm (using Nd:YAG laser) and at 966 nm (using Ti"-r :sapphire laser). Top: excited at 1064 nm, 100 mW laser power, 250 interferograms, 4 cm -1 resolution, 180 ° backscattering geometry. Bottom: excited at 966 nm, 100 mW laser power, 1000 interferograms, 4 cm -1 resolution, grazing incidence geometry. 15K

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-o

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16'00 Raman

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[cm-1]

Figure 3. Low temperature (15 K) FT Raman spectra of reaction centers from

Rb. sphaeroides. Top: Purified RCs excited at 1064 nm. Bottom: RCs in situ, in antenna-less membranes, ~xcited at 940 nm. 200 mW laser power, 2000 interferograms, 4 cm-Xresolution.

464 illustrates that the spectral features which dominate the spectrum are those of the reaction center protein and specifically those of the primary donor. The two spectra in Fig. 3 are ostensibly the same demonstrating that, for this case, the isolation and purification of the reaction center protein has no observable effects on the vibrational spectum of P, indicating no gross perturbations, environmental changes, and no alteration of the pigment-protein interactions. Thus, this combined technique of using molecular biology to produce antennaless mutants, and the use of an excitation-variable near IR FT Raman spectrometer permits the study of the structure of the primary donor of the reaction center in its native membrane environment. This is a powerful tool in determining the integrity of some purified RCs which may not be stable towards detergent solubilization. In general, the FT Raman spectrum of P in reaction centers from Rb. sphaeroides show minor shifts of some bands at 15 K as compared to room temperature [24]. In particular, the most noticeable difference is the loss in resolution, at 10 K (Figure 3) of the 1607 cm -1 band of the CaC m stretching mode (see Fig. 1) and the 1620 cn~-I band of the PL C2 acetyl carbonyl mo@. As well, there is an upshift of the 1691 cm -~ band at room temperature up to 1696 c m - l a t 10 K. 3.3 FT Raman Studies of Mutant Reaction Centers. Because FT Raman spectroscopy of purple bacterial reaction centers excited with 1064 nm provides a direct, high-quality vibrational spectrum of the primary electron, the technique is a powerful tool in determining structural consequences of genetically replaced amino acid residues near P. In this way, the r61e of i) specific amino acid residues interacting with P in native RCs or ii) new interactions produced by the genetic replacement of amino acids near P, can be assessed in their modulation of the physicochemical properties of P and chlorophylls in general. The first use of FT Raman spectroscopy to study mutant RCs was reported by Wachtveitl et al. [25]. In this study, a set of mutants were constructed in order to make the protein environment of the primary donor of Rb. sphaeroides more similar to that of the primary donor of Rps. viridis. One of these mutations, namely the replacement of phenylalanine M197 (see Fig. 1) with a tyrosine residue resulted in the formation of a new hydrogen bond on the C 2 acetyl carbonyl of PM, as is the case in Rps. viridis [4]. This new H-bond, surprisingly, only modestly modified the spectral and redox properties of the primary donor [25]. For example, this tyrosin~-donated H-bond resulted in a 30 mV increase in the redox midpoint potential of the p / p T couple. A similar mutant at the M197 position was constructed by Williams and A11en [14] but instead of tyrosine, these workers genetically introduced a histidine residue. FT Raman spectroscopy confirmed that a similar H-bond was formed in this mutant, that of slightly greater strength as compared to the tyrosine-donated H-bond. This former mutant, however, increased the redox potential of P by 125 mV. The results of these studies suggest that the chemical nature of the H-bond donor to conjugated carbonyls of P plays an important role in modifying its redox properties. 3.4 Hydrogen-Bonding and Physicochemical Properties of P. Williams and Allen have also genetically replaced amino acids in the vicinity of P with histidine residues at positions L131 and M160, i.e. near the C 9 keto carbonyls of P (see Fig. 1). These mutants were designed to introduce H-bonds to the keto carbonyls of P, as was the M197 mutant, described above, to introduce a H-bond on the acetyl carbonyl of PM; They also constructed a mutant by replacing histidine L168 (which is engaged in a strong H-bond with the C 2 acetyl carbonyl of PL in wildtype) by a phenylalanine residue incapable of H-bonding. FT Raman spectroscopy of these mutant reaction centers confirmed the rupture and formation of H-bonds to P as designed [14]. In addition, this study permitted us to unambiguously identify each of the C 2 and C 9 carbonyl vibrators of P in Rb. sphaeroides, consistent with our previous attributions [10] and X-ray crystal structure [6,8]. One great advantage of near IR FT Raman spectroscopy is that no difference techniques are required and the vibrational spectrum of P is directly obtained. This means

465 that the spectral bands directly give the frequency of the corresponding vibrational mode; this is not the case for differential spectra. Thus, with the FT Raman technique that we use, we can accurately measure the vibrational frequency shifts of the conjugated carbonyl bands of P when their H-bonding interactions are modified. This allows us to easily compare strengths of H-bonds formed or broken in the mutant RCs as well as estimate thermodynamic parameters such as H-bond enthalpies [14]. It is interesting to compare the mutant Rb. sphaeroides RC where phenylalanine M197 was replaced by a tyrosine (FY(M197) mutant) residue to that where the same residue is replaced by a histidine residue (FH(M197) mutant. Both of these mutations resulted in the formation of a new H-bond to the C 2 acetyl carbonyl of PM but in one case the donor is a tyrosine residue and in the other it is a histidine residue. As measured from the FT ,Raman spectra of these mutant RCs, the C 2 acetyl carbonyl vibrator downshifted by 23 c m - ' f o r the histidine-donatfA H-bond, whereas for the tyrosine-donated H-bond, the observed downshift is 17 cm-', indicating that the histidine-donated H-bond is slightly stronger [14]. Using empirical Badger-type relations, estimated enthalpies of 0.145 and 0.114 eV were calculated for the histidine- and tyrosine-donated H-bonds, respectively. In principle, the formatior~ of a H-bond should preferentially stabilize the ground state of P and thus raise the P/P-'- redox potential. Thus a stronger H-bond should raise the redox potential of P more than a weaker one. This general trend is observed for the two mutan_~ but while the estimated bond enthalpies differ by a factor of 1.3, the increase in p / p T redox potential for these two mutants compared to wildtype differ by a factor of six. Thus, if the strength of the H-bond is the sole factor underlying this effect, then this effect is not strictly linear and the nature of the H-bond donor and its environment may also need to be considered.

ACKNOWLEDGEMENTS I wish to thank my colleagues at Saclay, Marc Lutz, Bruno Robert, Diane Spiedel, Delphine Albouy, and James N. Sturgis for helpful and stimulating discussions. I would also like to thank Alain Boussac and Crcile Roselli at Saclay for their invaluable r61es in the variable excitation work. I am also grateful to my collaborators who have made the work presented here possible: James P. Allen and JoAnn C. Williams of Arizona State University, Michael R. Jones of the University of Sheffield, and Josef Wachtveitl at the Ludwig Maximilian-Universit~it, Munich. REFERENCES 1. M. Lutz, in: Advances in Infrared and Raman Spectroscopy, Vol. 11 (R.J.H. Clark and R.E. Hester, eds.), pp 211-300, Wiley-Heyden, London, 1984. 2. M. Lutz and B. Robert, in: Biological Applications of Raman Spectroscopy, (T.G. Spiro, ed.), pp 347-411, John Wiley & Sons, New York, 1988. 3. M. Lutz and W. M~tele, in: Chlorophylls, (H. Scheer, ed.), pp 855-902, CRC Press, Boca Raton, 1991. 4. H. Michel, O. Epp, and J. Deisenhofer, EMBO J., 5 (1986), 2445. 5. J.P. Allen, G. Feher, T.O. Yeates, H. Komiya, and D.C. Rees, Proc. Natl. Acad. Sci. U.S.A., 84 (1987) 5730. 6. O. E1-Kabbani, C.-H. Chang, D. Tiede, J. Norris, and M. Schiffer, Biochemistry, 30 (1991) 5361. 7. U. Ermler, G. Fritzsch, S. Buchannan, and H. Michel, in: Research in Photosynthesis (N. Murata, ed.), pp 341-347, Kluwer Academic, The Netherlands, 1992. 8. A.J. Chirino, E.J. Lous, M. Huber, J.P. Allen, C. Schenck, M.L. Paddock, G. Feher, and D.C. Rees, Biochemistry, 33 (1994), 4584. 9. T.A. Mattioli, A. Hoffmann, M. Lutz, and B. Schrader, Comptes Rendus Acad. Sci. Paris (s6rie III), 310 (1990), 441.

466 10. T.A. Mattioli, A. Hoffmann, B. Robert, B. Schrader, and M. Lutz, Biochemistry, 30 (1991), 4648. 11. T.A. Mattioli, A. Hoffmann, D.G. Sockalingum, S. Schrader, B. Robert, and M. Lutz, Spectrochim. Acta, 49A (1993), 785. 12. U. Feiler, T.A. Mattioli, I. Katheder, H. Scheer, M. Lutz, and B. Robert, J. Raman Spectrosc., 25 (1994), 365. 13. J. Wachtveitl, J.W. Farchaus, R. Das, M. Lutz, B. Robert, and T.A. Mattioli, Biochemistry, 32 (1993), 12875. 14. T.A. Mattioli, J.C. Williams, J.P. Allen, and B. Robert, Biochemistry, 33 (1994), 1636. 15. B. Schrader, A. Hoffmann, A. Simon, R. Podschadlowski, and M. Tischer, J. Mol. Struct., 217 (1990), 207. 16. C. Roselli, A. Boussac, and T.A. Mattioli, Proc. Natl. Acad. Sci. U.S.A., in press; C.L. Schoen, S.H. Sharma, C.E. Helsley, and H. Owen, Appl. Spectrosc., 47 (1993), 305. 17. B. Robert, Biochim. Biophys. Acta, 1017 (1990), 99. 18. W.G. M~intele, A.M. Wollenweber, E. Nabedryk, and J. Breton, Proc. Natl. Acad. Sci. U.S.A., 85 (1988), 8468. 19. R.L. Heald and T.M. Cotton, J. Phys. Chem., 94 (1990), 3968. 20. J. Rautter, Ch. Gef~ner, F. Lendzian, W. Lubitz, J.C. Williams, H.A. Murchison, S. Wang, N.W. Woodbury, and J.P. Allen, in: The Photosynthetic Bacterial Reaction Center II (J. Breton and A. Vermeglio, eds.), pp 99-108, Plenum, New York, 1992. 21. T.A. Mattioli, C. Roselli, and A. Boussac, Biochim. Biophys. Acta, 1101 (1992), 121. 22. M.R. Jones, R.W. Visschers, R. van Grondelle, and C.N. Hunter, Biochemistry, 31 (1992), 4458. 23. L.M.P. Beekman, R. Visschers, R. Monhouser, F. van Mourik, M. Heer-Dawson, T.A. Mattioli, P. McGlynn, C.N. Hunter, B. Robert, R. van Grondelle, and M.R. Jones, Biochim. Biophys. Acta, submitted for publication. 24. T.A. Mattioli, D. Sockalingum, M Lutz, and B. Robert, in: Research in Photosynthesis (N. Murata, ed.), pp 405-408, Kluwer Academic, The Netherlands, 1992. 25. J. Wachtveitl, J.W. Farchaus, R. Das, M. Lutz, B. Robert, and T.A. Mattioli, Biochemistry, 32 (1993), 12875.