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Role of heme propionates of myoglobin in vibrational energy relaxation
Chemical Physics Letters 430 (2006) 404–408 www.elsevier.com/locate/cplett
Role of heme propionates of myoglobin in vibrational energy relaxation Mai Koyama a, Saburo Neya b, Yasuhisa Mizutani a
a,c,*
Department of Chemistry, Graduate School of Science and Technology, Kobe University, 1-1 Rokkodai, Nada, Kobe 657-8501, Japan b Graduate School of Pharmaceutical Sciences, Chiba University, Inage-Yayoi, Chiba 263-8522, Japan c Molecular Photoscience Research Center, Kobe University, 1-1 Rokkodai, Nada, Kobe 657-8501, Japan Received 18 May 2006; in final form 20 July 2006 Available online 12 September 2006
Abstract The role of heme propionates of myoglobin in vibrational energy relaxation was studied by time-resolved resonance Raman spectroscopy. Time-resolved anti-Stokes spectra were measured to monitor the vibrational energy relaxation of the heme. The decay rates of the band intensities were compared between wild-type myoglobin and etioheme-substituted myoglobin where the heme lacks hydrogenbonding side chains. The decay rates of the anti-Stokes intensities of the latter were less than those of the former, providing strong support for a theoretical proposal that the propionates and their coupling to solvent bath play an important role in the dissipation of excess energy of the excited heme in solvated wild-type myoglobin. Ó 2006 Elsevier B.V. All rights reserved.
1. Introduction Myoglobin (Mb) is a small, well-characterized heme protein, which serves as an intracellular oxygen storage site. It contains 153 amino acids and an iron–protoporphyrin IX complex (protoheme, Fig. 1a). Histidine 93 is bound to the iron on the proximal side of the heme. In the ferrous state Mb can reversibly bind oxygen, carbon monoxide, or nitric oxide on the distal side of the heme [1]. Upon photoexcitation, the bound ligand to the heme is dissociated almost instantaneously [2–7]. This property has allowed us to study Mb dynamics by using a variety of timeresolved spectroscopic techniques [8–12]. Since the instantaneous photodissociation deposits a large amount of excess vibrational energy to the heme, an account of the energy flow serves as a model system for studying the energy relaxation path in complex molecules [13–18]. Both ultrafast spectroscopy and molecular dynamics simulation [19–21] have been used to study the vibrational energy relaxation processes after photoexcitation. Geminate * Corresponding author. Present address: Department of Chemistry, Graduate School of Science, Osaka University, 1-16 Machikaneyama, Toyonaka 560-0043, Japan. Fax: +81 6 6850 5785. E-mail address: [email protected] (Y. Mizutani).
0009-2614/$ - see front matter Ó 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2006.09.018
recombination of O2, which is a physiological substrate, is fast and occurs in the subnanosecond regime, while that of CO is slow and occurs in the submicrosecond regime. By using CO as the ligand instead of O2, it is possible to avoid complicated side reactions and associated population dynamics [22]. The heme is embedded within the protein and is sustained in a cavity by approximately 90 van der Waals contacts with the protein [23]. Although the heme and the protein constitute a single molecule, the heme prosthetic group is relatively isolated from the protein and approximates to a solute molecule dissolved in a ‘protein solvent’. Although, much of the heme remains buried in a hydrophobic pocket within the protein matrix, however, the heme has significant contacts with the solvent. The two propionate side chains of the heme are highly solvated and extend away from the protein. Lian et al. first suggested the possibility that the vibrational energy could be transferred directly from the heme to the solvent through the two heme propionate side chains [24]. Subsequently, several groups studied the heme ‘cooling’ in Mb using molecular dynamics simulation [19–21]. Considerably, Sagnella and Straub [20] suggested that the dominant channel for vibrational energy relaxation of the heme involved two steps: (1) the redistribution of the excess energy through a
M. Koyama et al. / Chemical Physics Letters 430 (2006) 404–408
N
N
N
N
N
Fe
Fe N
N
CO2 -
N
CO2 -
a
b
Fig. 1. Molecular structures of protoheme (a) and etioheme (b).
rapid intraheme vibrational relaxation process and (2) the transfer of excess energy to the surrounding solvent directly through the two propionate side chains. Several theoretical works have suggested the participation of the propionates in the heme cooling [19–21] and an experimental work showed that removal of all the heme side chains (including the propionates) slows down the heme cooling [25]. However, no experimental evidence has been obtained to show that energy is transferred to the surrounding solvent directly through the two propionate side chains. Time-resolved anti-Stokes Raman spectroscopy is selective for vibrationally excited modes, and it is therefore, considered as a powerful method for measuring vibrational energy relaxation [26–28]. In the present work, we studied the vibrational energy relaxation of reconstituted Mb (rMb) in which heme was replaced by etioheme. Etioheme has side chains on the b-positions, which is similar to protoheme; however, it lacks hydrogen-bonding side chains (Fig. 1b). Our results strongly support the conjecture that the two propionate side chains and their coupling to the solvent bath play a significant role in the dissipation of excess energy of the excited heme in solvated wild-type Mb (wtMb). 2. Experimental The details of the time-resolved resonance Raman apparatus are described elsewhere [6]. A picosecond modelocked Ti:sapphire oscillator (Spectra-Physics, Tsunami 3950), pumped by a diode-pumped solid state laser (Spectra-Physics, Millennia Vs), produced approximately 1.5 ps pulses with a repetition of 82 MHz. The seed pulse was amplified by a regenerative amplifier (Positive Light, Spitfire) operated at 1 kHz by pumping it with the 527 nm output of an intracavity frequency-doubled Nd:YLF laser (Spectra-Physics, Evolution X). In the pump arm, a pump pulse of 540 nm was generated with a home-built optical parametric generator (OPG) and amplifier (OPA). In the probe arm, a probe pulse of 442 nm was generated as the first Stokes stimulated Raman scattering from compressed
405
methane gas (50 kg/cm2) excited by the second harmonic of the 784 nm output. The pump and probe beams were made collinear and coaxial using a dichroic mirror. The sample solution was contained in a 10 mm B NMR tube and spun with a spinning cell device. The pump and probe pulses were attenuated by a Cr-coated quartz ND filter. At the sample point, the energies of the probe and pump pulses were 0.1 and 15 lJ, respectively. Spherical and cylindrical lenses were used to focus the pump and probe beams on the sample in line-focusing condition. Raman scattering was dispersed by a single spectrometer (Spex, 500M), equipped with a blazed-holographic grating (2400 grooves/mm), and detected by a liquid-nitrogen-cooled charge-coupled device (CCD) detector (Roper Scientific, Spec-10:400B/LN). The Raman shifts were calibrated with cyclohexane or carbon tetrachloride. The peak positions of Raman bands were accurate within ±2 cm 1. Data acquisition of picosecond time-resolved resonance Raman spectra was as described previously [6]. A cross-correlation trace of the pump and probe pulses was measured with a 1 mm bbarium borate (BBO) crystal, which indicated a width of 2.3 ps. The 0.0 ps of delay time (uncertainty < 0.2 ps) was calibrated using sum frequency mixing in the same crystal. The time-resolved anti-Stokes Raman spectra were accumulated over 24 and 20 min for wtMb and rMb, respectively. The probe-without-photolysis spectrum was subtracted from the probe-with-photolysis spectrum to yield the photoproduct spectrum. The subtraction parameter was determined by subtracting the probe-without-photolysis spectrum from the probe-with-photolysis spectrum until negative features were seen at the site of prominent bands of the CO-bound form. The subtraction parameter was then reduced until these negative peaks were eliminated, thereby accounting for the depletion of the CObound form caused by the pump pulse. Horse skeletal Mb (Sigma, M-0630) was used without further purification. The rMb with ferric etioheme was prepared according to the reported procedure [29] with some modifications. The crude mixture of apoMb and ferric etioheme chloride was dialyzed against several changes of 10 mM Tris buffer (pH 6.5) at 4 °C for 12–16 h and was loaded onto a carboxy-methylated-cellulose column equilibrated with the same buffer. The purified Mb was eluted from the column with a linear gradient of Tris buffer (10–100 mM) at pH 7.0 and 4 °C. Fractions with an absorption ratio A394 nm/A280 nm of 6.0 or greater were collected. The CO-bound forms of Mb were prepared by exposing dithionite-reduced samples to CO in tightly sealed Raman cells. 3. Results and discussion Fig. 2 shows the anti-Stokes time-resolved resonance Raman difference spectra of photodissociated wtMb (a) and rMb (b) for various values of delay times of the probe pulse with respect to the pump pulse. The delay times of the probe pulse from the pump pulse are indicated on the left
406
M. Koyama et al. / Chemical Physics Letters 430 (2006) 404–408
a
ν7
ν5
ν4
-5 ps -2 ps -1 ps 0 ps
ν7
ν5
669
1130
ν4
-5 ps -2 ps -1 ps 0 ps
669 1 ps 2 ps
b
1355
1356
1125 1 ps
3 ps 4 ps
2 ps
6 ps 8 ps
3 ps
10 ps
6 ps
15 ps 20 ps 30 ps 50 ps
8 ps 10 ps 15 ps 20 ps 30 ps 50 ps
4 ps
600 800 1000 1200 1400 1600 Raman shift / cm
-1
600 800 1000 1200 1400 1600 Raman shift / cm
-1
Fig. 2. Picosecond time-resolved anti-Stokes resonance Raman spectra of photodissociated forms of wtMb (a) and of rMb (b) in the 700–1700 cm region.
side of each spectrum. In these spectra, the contribution of unreacted species has been subtracted. Anti-Stokes intensities are highest at a 1 ps delay, when the m4, m5, and m7 bands were observed at 1355, 1130, and 669 cm 1 for wtMb and at 1355, 1125, and 671 cm 1 for rMb, respectively. The intensity of these bands decreased as the delay time increased and reached their equilibrium intensities at a 20 ps delay. It is well known that recombination of CO to the heme in Mb takes place in the time regime of microseconds to milliseconds [30]. In fact, we have previously reported that there was no significant intensity change in the Stokes m4 and m7 bands in the 3–50 ps time range [5,6,17]. Therefore, the observed intensity decay in the anti-Stokes m4 and m7 bands can be ascribed to vibrational energy relaxation. At 50 ps, recognizable anti-Stokes intensities were observed for the m4 and m7 modes. These intensities can be attributed to contribution from the population in thermal equilibrium at room temperature for following reasons. First, the observed relative Stokes and anti-Stokes intensities of the m4 band are estimated to be 1000, which is close to the value of Boltzmann factor for this mode at room temperature. Second, the anti-Stokes m4 band due to MbCO was observed in the probe-without-photolysis spectrum (data not shown). The observed anti-Stokes intensity should originate from thermal population of MbCO [6] because the quantum yield of the CO photodissociation is nearly unity [11] and the rebinding of the dissociated CO does not take place in the picosecond time regime [30]. An anti-Stokes band due to the thermal population at room temperature was observed also for a C@C
1
stretching band at 1520 cm 1 of canthaxanthin [27,31]. Therefore, it is reasonable to attribute the persistent antiStokes intensities of the CO-dissociated form to contribution from the population in thermal equilibrium at room temperature. Fig. 3 shows the temporal changes of anti-Stokes m4 and m7 band intensities for wtMb (a) and rMb (b). The anti-Stokes intensities were found to develop within the instrument response time. This is consistent with the photodissociation of CO from the heme taking place within 50 fs [4]. The instrument response was deconvoluted from the decay of the anti-Stokes intensity using a Gaussian fit to the cross-correlation signal. This analysis obtained the decay constants of 1.2 ± 0.4 ps for the m4 band [32] and 1.8 ± 0.4 ps for the m7 band of wtMb and of 1.8 ± 0.3 ps for the m4 band and 7.0 ± 1.9 ps for the m7 band of rMb. Simpson et al. suggested that the m7 mode of the heme couples quite well to its environment [16]. This mode readily losses its excess vibrational energy and rapidly reflects the environmental temperature. Therefore, we discuss the energy dissipation mechanism based on the difference of decay rates of anti-Stokes m7 intensities between wtMb and rMb. The intensity decay rate of the m7 band of rMb was less than those of wtMb. Because frequencies and intensity patterns of in-plane Raman bands of wtMb and rMb are very similar to each other, the vibrational characters of the m7 mode is considered similar in the etioheme and the protoheme. If the excess energy of the etioheme in Mb dissipated as efficiently as that of protoheme in Mb, the intensity decays of anti-Stokes m7 band would be
M. Koyama et al. / Chemical Physics Letters 430 (2006) 404–408
a
0
10
20
30
40
50
b
0
10
20 30 Delay time / ps
40
50
Fig. 3. Temporal changes in Raman intensity of the anti-Stokes m4 (circles) and m7 (triangles) bands of photodissociated forms of wtMb (a) and rMb (b). The solid lines are fits using an exponential function of the form A[exp( t/sdecay) + B] (an instantaneous rise and an exponential decay) convoluted with an instrument response function. The lines shown in (a) were obtained with the parameters of sdecay = 1.2 ± 0.4 and 1.8 ± 0.4 ps, B = 0.07 ± 0.03 and 0.21 ± 0.03 for the m4 and m7 bands, respectively. The lines shown in (b) were obtained with the parameters of sdecay = 1.7 ± 0.2 and 7.0 ± 1.9 ps, B = 0.01 ± 0.01 and 0.32 ± 0.03 for the m4 and m7 bands, respectively.
virtually the same in both wtMb and rMb, which is not the case. The difference in the decay rate indicates that the vibrational cooling of the etioheme is slower than that of the protoheme in Mb. This strongly suggests that the heme propionates accelerate the vibrational cooling in Mb. These results provide experimental evidence suggesting that the two propionate side chains play an important role in releasing excess energy from the heme of wild-type Mb. Vibrational energy relaxation of heme proteins has been studied using ultrafast spectroscopic techniques. The picosecond temporal changes of resonance Raman spectra [6,17,18,33] and band III [34] were measured in order to monitor the energy dissipation of the heme. The time constant of 1.8 ps for the population decay of the m7 mode of wtMb gives rise to time constant of 2 ps for the vibrational temperature decay in the present study. This is somewhat smaller than the value reported by Lim et al. [34]. The difference in the time constant may be due to the difference in the preparation of the vibrationally excited heme in Mb. We prepared the vibrationally excited heme by the photodissociation of CO for MbCO. Lim et al. prepared the vibrationally excited heme by the photoexcitation of deoxyMb. The excess energy deposited in the heme is much larger for the latter than for the former. Structural changes
407
in the heme and protein moiety accompany to the CO photodissociation. On the other hand, no structural change accompanies to the photoexcitation of deoxyMb. The fast energy dissipation to the water bath has been noted in the femtosecond TRIR study, which monitored the heating of water caused by the photoexcitation of deoxyMb [24]. The observed kinetics was fitted with a model having two time constants. The fast component was best fitted by a Gaussian rise function with a time constant of 7.5 ± 1.5 ps, and the slow component was described by a time constant of ca. 20 ps with 40% of the total amplitude. A comparison of the heme cooling and water heating studies suggests that there are two channels of energy dissipation from the protein to the water bath. One is a classical diffusion process, and it is responsible for the slow component, while the other is through the collective motions of the protein or through the heme side chains to the surrounding solvent, and is responsible for the fast component. Based on the results of molecular dynamics simulations of solvated deoxyMb, Sagnella and Straub have indicated that the strong electrostatic interaction of the propionate side chains and the solvating water is a most important ‘doorway’ for dissipation of excess energy in the heme [20]. Nagaoka and his coworkers carried out molecular dynamics simulations for the photodissociation of MbCO in vacuo and demonstrated that the vibrationally excited heme brings about excitation of the vibrational modes of the propionate groups [19]. A molecular dynamics simulation study by Bu and Straub [21] predicted that the relaxation time of heme cooling in Mb with the heme lacking propionate side chains decreased by 50% compared to native Mb. Champion and coworkers reported that the heme vibrational relaxation reduces in porphine-substituted Mb, where all the protoheme side chains are replaced by hydrogens [25]. This result suggested that the side chain structure affects the heme vibrational relaxation. A transient grating study by Miyata and Terazima suggested that most (83%) of the energy of the heme in deoxyMb is first transferred to the protein part and subsequently dissipates into the solvent and the remaining energy is directly transferred to the solvent [35]. The present results show that the heme vibrational relaxation reduces in rMb as compared to wtMb: the amputation of the heme’s propionates results in an increase in the time constant of the m7 band by factor of 3.9 ± 1.6. In the etioheme-substituted Mb, we assume that the strong electrostatic interaction between side chain and solvating water is lost, since the methyl and ethyl groups replace the propionates. This suggests that the vibrational relaxation will reduce, as confirmed by the present study. These results are the first direct observation that the heme vibrational relaxation is significantly affected by amputation of the propionates and strongly support the theoretical proposal that a possible doorway for energy release from the vibrationally excited heme involves the interaction of its propionate groups with neighboring solvent molecules.
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Acknowledgements This work was supported by a Grant-in-Aid for Specially Promoted Research (Grant No. 14001004) from the Ministry of Education, Culture, Sports, Science and Technology of Japan and a Grant-in-Aid for Scientific Research (B) (Grant No. 17350009) from the Japan Society for the Promotion of Science. References [1] E. Antonini, M. Brunori, Hemoglobin and Myoglobin in their Reactions with Ligands, North-Holland Publishing Co., Amsterdam, London, 1971. [2] L. Zhu, J.T. Sage, P.M. Champion, Science 266 (1994) 629. [3] P.A. Anfinrud, C. Han, R.M. Hochstrasser, Proc. Natl. Acad. Sci. USA 86 (1989) 8387. [4] J.W. Petrich, C. Poyart, J.L. Martin, Biochemistry 27 (1988) 4049. [5] Y. Mizutani, T. Kitagawa, J. Phys. Chem. B 105 (2001) 10992. [6] Y. Mizutani, T. Kitagawa, Chem. Rec. 1 (2001) 258. [7] S. Franzen, B. Bohn, C. Poyart, J.L. Martin, Biochemistry 34 (1995) 1224. [8] A. Ansari et al., Biochemistry 25 (1986) 3139. [9] J.P. Ogilvie, M. Plazanet, G. Dadusc, R.J.D. Miller, J. Phys. Chem. B 106 (2002) 10460. [10] P.A. Cornelius, R.M. Hochstrasser, A.W. Steele, J. Mol. Biol. 163 (1983) 119. [11] E.R. Henry, J.H. Sommer, J. Hofrichter, W.A. Eaton, J. Mol. Biol. 166 (1983) 443. [12] W.D. Tian, J.T. Sage, V.V. Srajer, P.M. Champion, Phys. Rev. Lett. 68 (1992) 408. [13] R.J.D. Miller, Annu. Rev. Phys. Chem. 42 (1991) 581. [14] P. Li, J.T. Sage, P.M. Champion, J. Chem. Phys. 97 (1992) 3214. [15] P. Li, P.M. Champion, Biophys. J. 66 (1994) 430. [16] M.C. Simpson, E.S. Peterson, C.F. Shannon, D.D. Eads, J.M. Friedman, C.M. Cheatum, M.R. Ondrias, J. Am. Chem. Soc. 119 (1997) 5110. [17] Y. Mizutani, T. Kitagawa, Science 278 (1997) 443.
[18] R. Lingle Jr., X. Xu, H. Zhu, S.-C. Yu, J.B. Hopkins, J. Phys. Chem. 95 (1991) 9320. [19] I. Okazaki, Y. Hara, M. Nagaoka, Chem. Phys. Lett. 337 (2001) 151. [20] D.E. Sagnella, J.E. Straub, J. Phys. Chem. B 105 (2001) 7057. [21] L. Bu, J.E. Straub, J. Phys. Chem. B 107 (2003) 10634. [22] Q.H. Gibson, J.S. Olson, R.E. McKinnie, R.J. Rohlfs, J. Biol. Chem. 261 (1986) 10228. [23] E.R. Henry, W.A. Eaton, R.M. Hochstrasser, Proc. Natl. Acad. Sci. USA 83 (1986) 8982. [24] T. Lian, B. Locke, Y. Kholodenko, R.M. Hochstrasser, J. Phys. Chem. 98 (1994) 11648. [25] X. Ye et al., J. Phys. Chem. A 107 (2003) 8156. [26] K.T. Schomacker, O. Bangcharoenpaurpong, P.M. Champion, J. Chem. Phys. 80 (1984) 4701. [27] H. Okamoto, T. Nakabayashi, M. Tasumi, J. Phys. Chem. A 101 (1997) 3488. [28] A.P. Shreve, R.A. Mathies, J. Phys. Chem. 99 (1995) 7285. [29] S. Neya, N. Funasaki, K. Imai, Biochim. Biophys. Acta 996 (1989) 226. [30] Q.H. Gibson, J.S. Olson, R.E. McKinnie, R.J. Rohlfs, J. Biol. Chem. 261 (1986) 10228. [31] T. Nakabayashi, H. Okamoto, M. Tasumi, J. Phys. Chem. A 101 (1997) 3494. [32] The time constant for the m4 mode previously reported by our group is 1.9 ± 0.6 ps [18], which is slightly larger than that reported in the present study. It is likely that apparent discrepancy in the time constant resulted from the incomplete curve fitting. In the previous report [18], the fit to the experimental points to obtain the time constant was not the best. There is a small deviation between the experimental points and the fitted curve in Fig. 4C of Ref. [18]. The fit is good enough in Fig. 3 of the present study. The present value of the time constant of the anti-Stokes m4 decay is more accurate. The time constant of 1.3 ps is consistent with the value reported in our recent paper [6]. [33] J.W. Petrich, J.L. Martin, D. Houde, C. Poyart, A. Orszag, Biochemistry 26 (1987) 7914. [34] M. Lim, T.A. Jackson, P.A. Anfinrud, J. Phys. Chem. 100 (1996) 12043. [35] R. Miyata, M. Terazima, Bull. Chem. Soc. Jpn. 76 (2003) 1707.
Role of heme propionates of myoglobin in vibrational energy relaxation Mai Koyama a, Saburo Neya b, Yasuhisa Mizutani a
a,c,*
Department of Chemistry, Graduate School of Science and Technology, Kobe University, 1-1 Rokkodai, Nada, Kobe 657-8501, Japan b Graduate School of Pharmaceutical Sciences, Chiba University, Inage-Yayoi, Chiba 263-8522, Japan c Molecular Photoscience Research Center, Kobe University, 1-1 Rokkodai, Nada, Kobe 657-8501, Japan Received 18 May 2006; in final form 20 July 2006 Available online 12 September 2006
Abstract The role of heme propionates of myoglobin in vibrational energy relaxation was studied by time-resolved resonance Raman spectroscopy. Time-resolved anti-Stokes spectra were measured to monitor the vibrational energy relaxation of the heme. The decay rates of the band intensities were compared between wild-type myoglobin and etioheme-substituted myoglobin where the heme lacks hydrogenbonding side chains. The decay rates of the anti-Stokes intensities of the latter were less than those of the former, providing strong support for a theoretical proposal that the propionates and their coupling to solvent bath play an important role in the dissipation of excess energy of the excited heme in solvated wild-type myoglobin. Ó 2006 Elsevier B.V. All rights reserved.
1. Introduction Myoglobin (Mb) is a small, well-characterized heme protein, which serves as an intracellular oxygen storage site. It contains 153 amino acids and an iron–protoporphyrin IX complex (protoheme, Fig. 1a). Histidine 93 is bound to the iron on the proximal side of the heme. In the ferrous state Mb can reversibly bind oxygen, carbon monoxide, or nitric oxide on the distal side of the heme [1]. Upon photoexcitation, the bound ligand to the heme is dissociated almost instantaneously [2–7]. This property has allowed us to study Mb dynamics by using a variety of timeresolved spectroscopic techniques [8–12]. Since the instantaneous photodissociation deposits a large amount of excess vibrational energy to the heme, an account of the energy flow serves as a model system for studying the energy relaxation path in complex molecules [13–18]. Both ultrafast spectroscopy and molecular dynamics simulation [19–21] have been used to study the vibrational energy relaxation processes after photoexcitation. Geminate * Corresponding author. Present address: Department of Chemistry, Graduate School of Science, Osaka University, 1-16 Machikaneyama, Toyonaka 560-0043, Japan. Fax: +81 6 6850 5785. E-mail address: [email protected] (Y. Mizutani).
0009-2614/$ - see front matter Ó 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2006.09.018
recombination of O2, which is a physiological substrate, is fast and occurs in the subnanosecond regime, while that of CO is slow and occurs in the submicrosecond regime. By using CO as the ligand instead of O2, it is possible to avoid complicated side reactions and associated population dynamics [22]. The heme is embedded within the protein and is sustained in a cavity by approximately 90 van der Waals contacts with the protein [23]. Although the heme and the protein constitute a single molecule, the heme prosthetic group is relatively isolated from the protein and approximates to a solute molecule dissolved in a ‘protein solvent’. Although, much of the heme remains buried in a hydrophobic pocket within the protein matrix, however, the heme has significant contacts with the solvent. The two propionate side chains of the heme are highly solvated and extend away from the protein. Lian et al. first suggested the possibility that the vibrational energy could be transferred directly from the heme to the solvent through the two heme propionate side chains [24]. Subsequently, several groups studied the heme ‘cooling’ in Mb using molecular dynamics simulation [19–21]. Considerably, Sagnella and Straub [20] suggested that the dominant channel for vibrational energy relaxation of the heme involved two steps: (1) the redistribution of the excess energy through a
M. Koyama et al. / Chemical Physics Letters 430 (2006) 404–408
N
N
N
N
N
Fe
Fe N
N
CO2 -
N
CO2 -
a
b
Fig. 1. Molecular structures of protoheme (a) and etioheme (b).
rapid intraheme vibrational relaxation process and (2) the transfer of excess energy to the surrounding solvent directly through the two propionate side chains. Several theoretical works have suggested the participation of the propionates in the heme cooling [19–21] and an experimental work showed that removal of all the heme side chains (including the propionates) slows down the heme cooling [25]. However, no experimental evidence has been obtained to show that energy is transferred to the surrounding solvent directly through the two propionate side chains. Time-resolved anti-Stokes Raman spectroscopy is selective for vibrationally excited modes, and it is therefore, considered as a powerful method for measuring vibrational energy relaxation [26–28]. In the present work, we studied the vibrational energy relaxation of reconstituted Mb (rMb) in which heme was replaced by etioheme. Etioheme has side chains on the b-positions, which is similar to protoheme; however, it lacks hydrogen-bonding side chains (Fig. 1b). Our results strongly support the conjecture that the two propionate side chains and their coupling to the solvent bath play a significant role in the dissipation of excess energy of the excited heme in solvated wild-type Mb (wtMb). 2. Experimental The details of the time-resolved resonance Raman apparatus are described elsewhere [6]. A picosecond modelocked Ti:sapphire oscillator (Spectra-Physics, Tsunami 3950), pumped by a diode-pumped solid state laser (Spectra-Physics, Millennia Vs), produced approximately 1.5 ps pulses with a repetition of 82 MHz. The seed pulse was amplified by a regenerative amplifier (Positive Light, Spitfire) operated at 1 kHz by pumping it with the 527 nm output of an intracavity frequency-doubled Nd:YLF laser (Spectra-Physics, Evolution X). In the pump arm, a pump pulse of 540 nm was generated with a home-built optical parametric generator (OPG) and amplifier (OPA). In the probe arm, a probe pulse of 442 nm was generated as the first Stokes stimulated Raman scattering from compressed
405
methane gas (50 kg/cm2) excited by the second harmonic of the 784 nm output. The pump and probe beams were made collinear and coaxial using a dichroic mirror. The sample solution was contained in a 10 mm B NMR tube and spun with a spinning cell device. The pump and probe pulses were attenuated by a Cr-coated quartz ND filter. At the sample point, the energies of the probe and pump pulses were 0.1 and 15 lJ, respectively. Spherical and cylindrical lenses were used to focus the pump and probe beams on the sample in line-focusing condition. Raman scattering was dispersed by a single spectrometer (Spex, 500M), equipped with a blazed-holographic grating (2400 grooves/mm), and detected by a liquid-nitrogen-cooled charge-coupled device (CCD) detector (Roper Scientific, Spec-10:400B/LN). The Raman shifts were calibrated with cyclohexane or carbon tetrachloride. The peak positions of Raman bands were accurate within ±2 cm 1. Data acquisition of picosecond time-resolved resonance Raman spectra was as described previously [6]. A cross-correlation trace of the pump and probe pulses was measured with a 1 mm bbarium borate (BBO) crystal, which indicated a width of 2.3 ps. The 0.0 ps of delay time (uncertainty < 0.2 ps) was calibrated using sum frequency mixing in the same crystal. The time-resolved anti-Stokes Raman spectra were accumulated over 24 and 20 min for wtMb and rMb, respectively. The probe-without-photolysis spectrum was subtracted from the probe-with-photolysis spectrum to yield the photoproduct spectrum. The subtraction parameter was determined by subtracting the probe-without-photolysis spectrum from the probe-with-photolysis spectrum until negative features were seen at the site of prominent bands of the CO-bound form. The subtraction parameter was then reduced until these negative peaks were eliminated, thereby accounting for the depletion of the CObound form caused by the pump pulse. Horse skeletal Mb (Sigma, M-0630) was used without further purification. The rMb with ferric etioheme was prepared according to the reported procedure [29] with some modifications. The crude mixture of apoMb and ferric etioheme chloride was dialyzed against several changes of 10 mM Tris buffer (pH 6.5) at 4 °C for 12–16 h and was loaded onto a carboxy-methylated-cellulose column equilibrated with the same buffer. The purified Mb was eluted from the column with a linear gradient of Tris buffer (10–100 mM) at pH 7.0 and 4 °C. Fractions with an absorption ratio A394 nm/A280 nm of 6.0 or greater were collected. The CO-bound forms of Mb were prepared by exposing dithionite-reduced samples to CO in tightly sealed Raman cells. 3. Results and discussion Fig. 2 shows the anti-Stokes time-resolved resonance Raman difference spectra of photodissociated wtMb (a) and rMb (b) for various values of delay times of the probe pulse with respect to the pump pulse. The delay times of the probe pulse from the pump pulse are indicated on the left
406
M. Koyama et al. / Chemical Physics Letters 430 (2006) 404–408
a
ν7
ν5
ν4
-5 ps -2 ps -1 ps 0 ps
ν7
ν5
669
1130
ν4
-5 ps -2 ps -1 ps 0 ps
669 1 ps 2 ps
b
1355
1356
1125 1 ps
3 ps 4 ps
2 ps
6 ps 8 ps
3 ps
10 ps
6 ps
15 ps 20 ps 30 ps 50 ps
8 ps 10 ps 15 ps 20 ps 30 ps 50 ps
4 ps
600 800 1000 1200 1400 1600 Raman shift / cm
-1
600 800 1000 1200 1400 1600 Raman shift / cm
-1
Fig. 2. Picosecond time-resolved anti-Stokes resonance Raman spectra of photodissociated forms of wtMb (a) and of rMb (b) in the 700–1700 cm region.
side of each spectrum. In these spectra, the contribution of unreacted species has been subtracted. Anti-Stokes intensities are highest at a 1 ps delay, when the m4, m5, and m7 bands were observed at 1355, 1130, and 669 cm 1 for wtMb and at 1355, 1125, and 671 cm 1 for rMb, respectively. The intensity of these bands decreased as the delay time increased and reached their equilibrium intensities at a 20 ps delay. It is well known that recombination of CO to the heme in Mb takes place in the time regime of microseconds to milliseconds [30]. In fact, we have previously reported that there was no significant intensity change in the Stokes m4 and m7 bands in the 3–50 ps time range [5,6,17]. Therefore, the observed intensity decay in the anti-Stokes m4 and m7 bands can be ascribed to vibrational energy relaxation. At 50 ps, recognizable anti-Stokes intensities were observed for the m4 and m7 modes. These intensities can be attributed to contribution from the population in thermal equilibrium at room temperature for following reasons. First, the observed relative Stokes and anti-Stokes intensities of the m4 band are estimated to be 1000, which is close to the value of Boltzmann factor for this mode at room temperature. Second, the anti-Stokes m4 band due to MbCO was observed in the probe-without-photolysis spectrum (data not shown). The observed anti-Stokes intensity should originate from thermal population of MbCO [6] because the quantum yield of the CO photodissociation is nearly unity [11] and the rebinding of the dissociated CO does not take place in the picosecond time regime [30]. An anti-Stokes band due to the thermal population at room temperature was observed also for a C@C
1
stretching band at 1520 cm 1 of canthaxanthin [27,31]. Therefore, it is reasonable to attribute the persistent antiStokes intensities of the CO-dissociated form to contribution from the population in thermal equilibrium at room temperature. Fig. 3 shows the temporal changes of anti-Stokes m4 and m7 band intensities for wtMb (a) and rMb (b). The anti-Stokes intensities were found to develop within the instrument response time. This is consistent with the photodissociation of CO from the heme taking place within 50 fs [4]. The instrument response was deconvoluted from the decay of the anti-Stokes intensity using a Gaussian fit to the cross-correlation signal. This analysis obtained the decay constants of 1.2 ± 0.4 ps for the m4 band [32] and 1.8 ± 0.4 ps for the m7 band of wtMb and of 1.8 ± 0.3 ps for the m4 band and 7.0 ± 1.9 ps for the m7 band of rMb. Simpson et al. suggested that the m7 mode of the heme couples quite well to its environment [16]. This mode readily losses its excess vibrational energy and rapidly reflects the environmental temperature. Therefore, we discuss the energy dissipation mechanism based on the difference of decay rates of anti-Stokes m7 intensities between wtMb and rMb. The intensity decay rate of the m7 band of rMb was less than those of wtMb. Because frequencies and intensity patterns of in-plane Raman bands of wtMb and rMb are very similar to each other, the vibrational characters of the m7 mode is considered similar in the etioheme and the protoheme. If the excess energy of the etioheme in Mb dissipated as efficiently as that of protoheme in Mb, the intensity decays of anti-Stokes m7 band would be
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a
0
10
20
30
40
50
b
0
10
20 30 Delay time / ps
40
50
Fig. 3. Temporal changes in Raman intensity of the anti-Stokes m4 (circles) and m7 (triangles) bands of photodissociated forms of wtMb (a) and rMb (b). The solid lines are fits using an exponential function of the form A[exp( t/sdecay) + B] (an instantaneous rise and an exponential decay) convoluted with an instrument response function. The lines shown in (a) were obtained with the parameters of sdecay = 1.2 ± 0.4 and 1.8 ± 0.4 ps, B = 0.07 ± 0.03 and 0.21 ± 0.03 for the m4 and m7 bands, respectively. The lines shown in (b) were obtained with the parameters of sdecay = 1.7 ± 0.2 and 7.0 ± 1.9 ps, B = 0.01 ± 0.01 and 0.32 ± 0.03 for the m4 and m7 bands, respectively.
virtually the same in both wtMb and rMb, which is not the case. The difference in the decay rate indicates that the vibrational cooling of the etioheme is slower than that of the protoheme in Mb. This strongly suggests that the heme propionates accelerate the vibrational cooling in Mb. These results provide experimental evidence suggesting that the two propionate side chains play an important role in releasing excess energy from the heme of wild-type Mb. Vibrational energy relaxation of heme proteins has been studied using ultrafast spectroscopic techniques. The picosecond temporal changes of resonance Raman spectra [6,17,18,33] and band III [34] were measured in order to monitor the energy dissipation of the heme. The time constant of 1.8 ps for the population decay of the m7 mode of wtMb gives rise to time constant of 2 ps for the vibrational temperature decay in the present study. This is somewhat smaller than the value reported by Lim et al. [34]. The difference in the time constant may be due to the difference in the preparation of the vibrationally excited heme in Mb. We prepared the vibrationally excited heme by the photodissociation of CO for MbCO. Lim et al. prepared the vibrationally excited heme by the photoexcitation of deoxyMb. The excess energy deposited in the heme is much larger for the latter than for the former. Structural changes
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in the heme and protein moiety accompany to the CO photodissociation. On the other hand, no structural change accompanies to the photoexcitation of deoxyMb. The fast energy dissipation to the water bath has been noted in the femtosecond TRIR study, which monitored the heating of water caused by the photoexcitation of deoxyMb [24]. The observed kinetics was fitted with a model having two time constants. The fast component was best fitted by a Gaussian rise function with a time constant of 7.5 ± 1.5 ps, and the slow component was described by a time constant of ca. 20 ps with 40% of the total amplitude. A comparison of the heme cooling and water heating studies suggests that there are two channels of energy dissipation from the protein to the water bath. One is a classical diffusion process, and it is responsible for the slow component, while the other is through the collective motions of the protein or through the heme side chains to the surrounding solvent, and is responsible for the fast component. Based on the results of molecular dynamics simulations of solvated deoxyMb, Sagnella and Straub have indicated that the strong electrostatic interaction of the propionate side chains and the solvating water is a most important ‘doorway’ for dissipation of excess energy in the heme [20]. Nagaoka and his coworkers carried out molecular dynamics simulations for the photodissociation of MbCO in vacuo and demonstrated that the vibrationally excited heme brings about excitation of the vibrational modes of the propionate groups [19]. A molecular dynamics simulation study by Bu and Straub [21] predicted that the relaxation time of heme cooling in Mb with the heme lacking propionate side chains decreased by 50% compared to native Mb. Champion and coworkers reported that the heme vibrational relaxation reduces in porphine-substituted Mb, where all the protoheme side chains are replaced by hydrogens [25]. This result suggested that the side chain structure affects the heme vibrational relaxation. A transient grating study by Miyata and Terazima suggested that most (83%) of the energy of the heme in deoxyMb is first transferred to the protein part and subsequently dissipates into the solvent and the remaining energy is directly transferred to the solvent [35]. The present results show that the heme vibrational relaxation reduces in rMb as compared to wtMb: the amputation of the heme’s propionates results in an increase in the time constant of the m7 band by factor of 3.9 ± 1.6. In the etioheme-substituted Mb, we assume that the strong electrostatic interaction between side chain and solvating water is lost, since the methyl and ethyl groups replace the propionates. This suggests that the vibrational relaxation will reduce, as confirmed by the present study. These results are the first direct observation that the heme vibrational relaxation is significantly affected by amputation of the propionates and strongly support the theoretical proposal that a possible doorway for energy release from the vibrationally excited heme involves the interaction of its propionate groups with neighboring solvent molecules.
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Acknowledgements This work was supported by a Grant-in-Aid for Specially Promoted Research (Grant No. 14001004) from the Ministry of Education, Culture, Sports, Science and Technology of Japan and a Grant-in-Aid for Scientific Research (B) (Grant No. 17350009) from the Japan Society for the Promotion of Science. References [1] E. Antonini, M. Brunori, Hemoglobin and Myoglobin in their Reactions with Ligands, North-Holland Publishing Co., Amsterdam, London, 1971. [2] L. Zhu, J.T. Sage, P.M. Champion, Science 266 (1994) 629. [3] P.A. Anfinrud, C. Han, R.M. Hochstrasser, Proc. Natl. Acad. Sci. USA 86 (1989) 8387. [4] J.W. Petrich, C. Poyart, J.L. Martin, Biochemistry 27 (1988) 4049. [5] Y. Mizutani, T. Kitagawa, J. Phys. Chem. B 105 (2001) 10992. [6] Y. Mizutani, T. Kitagawa, Chem. Rec. 1 (2001) 258. [7] S. Franzen, B. Bohn, C. Poyart, J.L. Martin, Biochemistry 34 (1995) 1224. [8] A. Ansari et al., Biochemistry 25 (1986) 3139. [9] J.P. Ogilvie, M. Plazanet, G. Dadusc, R.J.D. Miller, J. Phys. Chem. B 106 (2002) 10460. [10] P.A. Cornelius, R.M. Hochstrasser, A.W. Steele, J. Mol. Biol. 163 (1983) 119. [11] E.R. Henry, J.H. Sommer, J. Hofrichter, W.A. Eaton, J. Mol. Biol. 166 (1983) 443. [12] W.D. Tian, J.T. Sage, V.V. Srajer, P.M. Champion, Phys. Rev. Lett. 68 (1992) 408. [13] R.J.D. Miller, Annu. Rev. Phys. Chem. 42 (1991) 581. [14] P. Li, J.T. Sage, P.M. Champion, J. Chem. Phys. 97 (1992) 3214. [15] P. Li, P.M. Champion, Biophys. J. 66 (1994) 430. [16] M.C. Simpson, E.S. Peterson, C.F. Shannon, D.D. Eads, J.M. Friedman, C.M. Cheatum, M.R. Ondrias, J. Am. Chem. Soc. 119 (1997) 5110. [17] Y. Mizutani, T. Kitagawa, Science 278 (1997) 443.
[18] R. Lingle Jr., X. Xu, H. Zhu, S.-C. Yu, J.B. Hopkins, J. Phys. Chem. 95 (1991) 9320. [19] I. Okazaki, Y. Hara, M. Nagaoka, Chem. Phys. Lett. 337 (2001) 151. [20] D.E. Sagnella, J.E. Straub, J. Phys. Chem. B 105 (2001) 7057. [21] L. Bu, J.E. Straub, J. Phys. Chem. B 107 (2003) 10634. [22] Q.H. Gibson, J.S. Olson, R.E. McKinnie, R.J. Rohlfs, J. Biol. Chem. 261 (1986) 10228. [23] E.R. Henry, W.A. Eaton, R.M. Hochstrasser, Proc. Natl. Acad. Sci. USA 83 (1986) 8982. [24] T. Lian, B. Locke, Y. Kholodenko, R.M. Hochstrasser, J. Phys. Chem. 98 (1994) 11648. [25] X. Ye et al., J. Phys. Chem. A 107 (2003) 8156. [26] K.T. Schomacker, O. Bangcharoenpaurpong, P.M. Champion, J. Chem. Phys. 80 (1984) 4701. [27] H. Okamoto, T. Nakabayashi, M. Tasumi, J. Phys. Chem. A 101 (1997) 3488. [28] A.P. Shreve, R.A. Mathies, J. Phys. Chem. 99 (1995) 7285. [29] S. Neya, N. Funasaki, K. Imai, Biochim. Biophys. Acta 996 (1989) 226. [30] Q.H. Gibson, J.S. Olson, R.E. McKinnie, R.J. Rohlfs, J. Biol. Chem. 261 (1986) 10228. [31] T. Nakabayashi, H. Okamoto, M. Tasumi, J. Phys. Chem. A 101 (1997) 3494. [32] The time constant for the m4 mode previously reported by our group is 1.9 ± 0.6 ps [18], which is slightly larger than that reported in the present study. It is likely that apparent discrepancy in the time constant resulted from the incomplete curve fitting. In the previous report [18], the fit to the experimental points to obtain the time constant was not the best. There is a small deviation between the experimental points and the fitted curve in Fig. 4C of Ref. [18]. The fit is good enough in Fig. 3 of the present study. The present value of the time constant of the anti-Stokes m4 decay is more accurate. The time constant of 1.3 ps is consistent with the value reported in our recent paper [6]. [33] J.W. Petrich, J.L. Martin, D. Houde, C. Poyart, A. Orszag, Biochemistry 26 (1987) 7914. [34] M. Lim, T.A. Jackson, P.A. Anfinrud, J. Phys. Chem. 100 (1996) 12043. [35] R. Miyata, M. Terazima, Bull. Chem. Soc. Jpn. 76 (2003) 1707.