Specific Ca 2+ -binding motif in the LH1 complex from photosynthetic bacterium Thermochromatium tepidum as revealed by optical spectroscopy and structural modeling Fei Ma 1,3 , Yukihiro Kimura 2 , Long-Jiang Yu 2 , Peng Wang 1 , Xi-Cheng Ai 1 , Zheng-Yu Wang 2 and Jian-Ping Zhang 1 1 Department of Chemistry, Renmin University of China, Beijing, China 2 Faculty of Science, Ibaraki University, Mito, Japan 3 Beijing National Laboratory for Molecular Science, State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Institute of Chemistry, Chinese Academy of Sciences, China Light-harvesting (LH) complexes are transmembrane proteins that are involved in the primary steps of bac- terial photosynthesis: capturing the sun light and trans- ferring the energy, in the form of electronic excitation, to the reaction center (RC). Most purple bacteria con- tain two basic types of LH complexes, i.e. the periph- eral antenna LH2 and the core antenna LH1 [1–3]. X-ray crystallographic structures of LH2 are available for Rhodopseudomans (Rps.) acidophila strain 10050 [4] and Rhodospirillum (Rs.) molischianum [5] with resolu- tions of 2.0–2.5 A ˚ . Although the highest available reso- lution for LH1 [6], 4.8 A ˚ , is not sufficient to display the structural details, it clearly shows that bacterio- chlorophyll (BChl) dimers are sandwiched between a- and b-helices of 15 or 16 subunits arranged in a ring-like manner around the RC. In addition, the Keywords 3D structural modeling; light-harvesting– reaction center core complex (LH1–RC); photosynthetic purple bacterium; Raman spectroscopy; Thermochromatium (Tch.) tepidum Correspondence Z Y. Wang, Faculty of Science, Ibaraki University, Mito 310 8512, Japan Fax: +81 29 2288352 Tel: +81 29 2288352 E-mail: wang@mx.ibaraki.ac.jp J P. Zhang, Department of Chemistry, Renmin University of China, Beijing 1000872, China Fax: +86 10 62516444 Tel: +86 10 62516604 E-mail: jpzhang@chem.ruc.edu.cn (Received 25 November 2008, revised 14 January 2009, accepted 14 January 2009) doi:10.1111/j.1742-4658.2009.06905.x Native and Ca 2+ -depleted light-harvesting–reaction center core complexes (LH1–RC) from the photosynthetic bacterium Thermochromatium (Tch.) tepidum exhibit maximal LH1–Q y absorption at 915 and 889 nm, respec- tively. To understand the structural origins of the spectral variation, we performed spectroscopic and structure modeling investigations. For the 889 nm form of LH1–RC, bacteriochlorophyll a (BChl a) in the native form was found by means of near-infrared Fourier-transform Raman spec- troscopy, a higher degree of macrocycle distortion and a stronger hydrogen bond with the b-Trp )8 residue. SWISS-MODEL structure modeling sug- gests the presence of a specific coordination motif of Ca 2+ at the C-termi- nus of the a-subunit of LH1, while MODELLER reveals the tilt of a- and b-polypeptides with reference to the structural template, as well as a change in the concentric orientation of BChl a molecules, both of which may be connected to the long-wavelength LH1–Q y absorption of the 915 nm form. The carotenoid spirilloxanthin shows a twisted all-trans configuration in both forms of LH1 as evidenced by the resonance Raman spectroscopic results. With regard to the thermal stability, the 915 nm form was shown by the use of temperature-dependent fluorescence spectroscopy to be approximately 20 K more stable than the 889 nm form, which may be ascribed to the specific Ca 2+ -binding motif of LH1. Abbreviations BChl a, bacteriochlorophyll a; Car, carotenoid; fwhm, full width at half maximum; LH1, light-harvesting complex 1; Q y , the absorptive optical transition to the lowest excited state of BChl a; RC, reaction center. FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1739 structures of a- and b-polypeptides in solution were determined for Rs. rubrum by means of 2D-NMR spectroscopy [7]. The purple photosynthetic bacterium Thermochro- matium (Tch.) tepidum was first identified in Mammoth Hot Springs in the Yellowstone National Park [8]. It is a moderate thermophile with an optimal temperature range of 48–50 °C and an upper limit of 55 °C, and its pigment–protein complexes show considerably higher thermal stability than those from its mesophilic counterparts such as Allochromatium (Ach.) vinosum, Rhodobacter (Rb.) sphaeroides and Blastochloris (Bl.) viridis, which grow at temperatures below approxi- mately 30 °C [9]. The light-harvesting–reaction center core complex (LH1–RC) from Tch. tepidum is peculiar with respect to its long-wavelength Q y absorption of BChl a at 915 nm, which shifts to approximately 885 nm when eluted in presence of NaCl, KCl, KBr, NaCl or MgCl 2 (150 mm). Interestingly, the 885 nm LH1–RC complex can be fully converted back to the 915 nm form by adding CaCl 2 [10,11]. Recently, polypeptides of LH1 from Tch. tepidum have been purified and the amino acid sequences deter- mined [12]. In addition, the dimeric feature and the highly symmetric ring assembly of BChls in LH1, as well as the interaction between BChl a and carotenoid molecules, have been confirmed [13]. It has been shown that spirilloxanthin is the major carotenoid (approxi- mately 92.3%), and that the 889 nm form of LH1–RC is thermally less stable than the 915 nm form [11,13]. Furthermore, Ca 2+ has been proven to coordinate in a ratio of 1 : 1 to an a-, b-subunit when the 889 to 915 nm transformation is induced [11]. Our recent study on the excitation dynamics of the two forms has shown similar LH1-to-RC excitation trapping kinetics, as well as similar efficiency of the transfer of excitation energy from carotenoid to BChl despite some differences in the BChl-to-carotenoid molecular orientation [14]. Ca 2+ plays vital roles in biological activities, e.g. as messengers of signal transduction in the cell, and for structural stabilization of proteins, etc. [15]. Ca 2+ in protein usually coordinates seven oxygen atoms from amino acid residues and water molecules, which accordingly form a pentagonal bi-pyramid cavity. However, coordination with 6, 8 or even up to 12 atoms is also possible. A helix-loop-helix structural domain constituting the Ca 2+ binding motif is found in a large number of Ca 2+ -binding proteins, and is also known as the EF hand [16]. Proteins containing the EF hand are divided into two classes according to their functions: signaling and buffering ⁄ transport proteins. The former undergoing Ca 2+ -dependent conformational changes, constitute the largest family, including well-known members such as the Ca 2+ -AT- Pase from skeletal muscle sarcoplasmic reticulum whose transmembrane helices tilt approximately 30° when transformed from the Ca 2+ -bound form (E1Ca 2+ ) to the Ca 2+ -free form [E2(TG)] [17,18]. The interchangeable 915 and 889 nm forms of LH1– RC from Tch. tepidum provide us with a unique opportunity to investigate the structure–function rela- tionship of these proteins. In the present study, we used near-infrared Fourier-transform Raman spectros- copy (FT-Raman) to assess the structural differences in BChl a molecules between the two forms of LH1– RC. Compared to the 889 nm form, the 915 nm form shows a stronger hydrogen bond (H-bond) interaction between the C 10a acetyl carbonyl and the tryptophan (Trp) residue from the b-polypeptide, b-Trp )8 , and more severe distortion of the BChl a macrocycle. Fur- thermore, the twist of all-trans spirilloxanthin was found to be similar between the two LH1–RC forms by use of resonance Raman spectroscopy. The results of 3D structural modeling reveal a specific Ca 2+ -coor- dination cavity that may induce configurational changes in the polypeptides, and, as a result, in BChl a molecules. The results are discussed in terms of the long-wavelength Q y absorption of native LH1–RC. Furthermore, the systematic shift of fluorescence spec- tra against temperature shows that the thermal stabil- ity of the intact LH1–RC is approximately 20 K higher than that of Ca 2+ -depleted LH1–RC. Results and Discussion Steady-state absorption and fluorescence spectroscopy The 915 nm form of LH1–RC exhibits much higher thermal stability than the 889 nm form. As shown in Fig. 1A, the absorption spectra of 915 nm LH1–RC vary slightly from 273 to 323 K, i.e. the LH1–Q y absorbance decreases approximately 3% with little change in band width. In contrast, for the 889 nm form under similar experimental conditions, dramatic decreases in the LH1–Q y and carotenoid absorption are seen (Fig. 1B), together with emergence of a new absorption maximum at 770 nm that is ascribed to monomeric BChl a. When the temperature exceeds 303 K, a large spectral change is seen, most likely due to disassembly of the LH1 complex. Upon increasing temperature, the fluorescence peak wavelengths of both the 915 and 889 nm forms shift to blue, and the emission bands get broader (Fig. 2). For the 915 nm form, the peak wavelength shifts from 945.4 to 939.5 nm, and the bandwidth increases from Ca 2+ -binding motif in an LH1 complex F. Ma et al. 1740 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 494 to 553 cm )1 [full width at half maximum (fwhm)] on raising the temperature from 273 to 323 K. The increase in spectral shift or bandwidth in response to the increase in temperature may indicate the involve- ment of more thermally populated excitonic states in the Q y -state manifold of BChl a. Using an energy dif- ference of 120 cm )1 between the lowest and the second lowest excitonic states [19], the population increase in the second lowest excitonic state in response to a temperature increase of 50 K was estimated to be 6.4%. Given the amount of spectral shift (66 cm )1 ) and band broadening (57 cm )1 ), it is reasonable to ascribe the fluorescence spectral changes to a new ther- mal equilibrium in the Q y state. As shown in the inset to Fig. 2A, the spectra shift slowly to blue against a temperature increase below 293 K, and the shift is faster and shows linear temperature dependence above 293 K. In addition, the decrease in fluorescence inten- sity may be due to the increased rate of internal con- version. On the other hand, when the temperature increases from 273 to 303 K, the fluorescence maxi- mum of the 889 nm form shifts from 918.8 to 914.1 nm, while the bandwidth increases from 534 to 569 cm )1 (fwhm). When the temperature exceeds 303 K, the fluorescence intensity decreases consider- ably due to dissociation of the LH1–RC assembly. The tendency of spectral shift appears to be signifi- cantly different between the two LH1–RC forms, i.e. nonlinear and linear temperature dependence are observed for the 915 and 889 nm forms, respectively, which may reflect their structural differences. The 915 nm LH1–RC form exhibits slower (273–293 K) and faster (293–323 K) phases of band shift (Fig. 2A); however, the 889 nm complex shows monophasic behavior (273–303 K; Fig. 2B) with a slope compara- ble to the faster phase of the 915 nm form. Comparing 300 400 500 600 700 800 900 1000 300 400 500 600 700 800 900 1000 0.00 0.05 0.10 0.15 0.20 323 K 293 K 273 K LH1-Q y Car Wavelength (nm) Wavelength (nm) 0.00 0.05 0.10 0.15 323 K 313 K 303 K 293 K 283 K 770 nm 273 K Car LH1-Q y Absorbance Absorbance A B Fig. 1. Steady-state UV-visible spectra of the 915 nm (A) and 889 nm (B) LH1–RC preparations from Tch. tepidum at the indi- cated temperatures. Arrows in (B) indicate the direction of absor- bance change upon temperature increase from 273 to 323 K. 900 950 1000 0 1000 2000 3000 10 580 10 600 10 620 10 640 Wavelength (nm) Wavelength (nm) 900 950 0 1000 2000 3000 270 280 290 300 270 280 290 300 310 320 10 880 10 900 10 920 10 940 Fluorescence intensity / a.u. Fluorescence intensity / a.u. A B T/K T/K ν m ·cm –1 ν m ·cm –1 Fig. 2. Fluorescence emission spectra recorded at various temper- atures for the 915 nm (A) and 889 nm (B) LH1–RC preparations from Tch. tepidum. Arrows show the direction of temperature change from 273 to 323 K in (A) and from 273 to 303 K in (B). Insets show the change of emission maxima (in wave number) against temperature. The excitation wavelength was 590 nm. F. Ma et al. Ca 2+ -binding motif in an LH1 complex FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1741 the 915 and 889 nm forms, a difference of 20 K in the starting temperature of the faster phases was found (293 versus 273 K), indicating that the pigment–pro- tein assembly of the 915 nm complex is more stable, most likely because of the binding of Ca 2+ . This result is in agreement with a recent differential scanning calo- rimetry study on the same core complexes [20], in which the dissociation temperature of the 915 nm form was found to be 15 K higher than that of the 889 nm form, and the enthalpy change for the former was found to be approximately 28% larger than that for the latter. The Stokes shifts between absorption and fluores- cence maxima are 28.9–24.5 nm for the 915 nm form and 29.8–25.1 nm for the 889 nm form over the tem- perature ranges 273–323 and 273–303 K, respectively, and are considerably larger than those of mesophilic purple bacteria such as Rs. rubrum (approximately 15 nm). Therefore, for Tch. tepidum, the spectral over- lap integral between LH1 emission and RC absorption (maximum at 865 nm) must be much smaller. How- ever, the rates of LH1-to–RC excitation energy trans- fer are rather similar from the thermophilic to the mesophilic species [14], implying that the rate is not strictly proportional to the spectral overlap integral. Resonance Raman spectroscopy Figure 3A shows the resonance Raman spectrum of a 915 nm form with spirilloxanthin as the major caro- tenoid component (approximately 92.3%). The key Raman lines at 1504 cm )1 (m 1 , C=C stretching) and 1143 cm )1 (m 2 , C–C stretching) can be assigned to all-trans spirilloxanthin in LH1. The Raman bands from 15-cis spirilloxanthin in the RC normally seen at 1528, 1239 and 1160 cm )1 [21] do not show up because the majority of spirilloxanthin molecules associate with LH1 and only a minor amount in the 15-cis configura- tion binds preferentially to the RC [22]. The Raman band at approximately 965.3 cm )1 is characteristic of the out-of-plane movement of C–H (m 4 ), which becomes symmetry-allowed only when the polyene backbone experiences nonplanar distortion [23]. As the m 4 mode is localized to and originates from the twists at C 11 =C 12 and C 7 =C 8 and their conjugates, C 11¢ C 12¢ and C 7¢ C 8¢ , it is concluded that all-trans spirilloxan- thin bound to LH1 takes on a twisted configuration, similarly to the case for LH1 of Rs. rubrum [21,23]. The Raman spectra do not change appreciably between the 915 and 889 nm LH1–RC forms, indicat- ing that the configuration of spirilloxanthin does not vary despite a large difference in the Q y absorption wavelength of BChl a (26 nm). A similar conclusion was reached in a recent investigation of the same com- plexes by means of circular dichromism spectroscopy [11]. Near-infrared FT-Raman spectroscopy Figure 4 shows the FT-Raman spectra for the 915 and 889 nm LH1–RC forms from Tch. tepidum, and Table 1 lists the assignments based on recent work by Frolov et al. [24]. The key Raman lines labeled with carotenoid correspond to the m 1 (1504 cm )1 ), m 2 (1147 cm )1 ) and m 3 (1023 cm )1 ) modes of spirilloxan- thin (see above), while those labeled R1–R4 originate ν 4 ν 3 ν 2 Intensity / a.u. ν 1 Raman shift·cm –1 800 1000 1200 1400 1600 967.4 997 1145 1187 1278 1352 1387 1447 1504 997 965.3 1143 1185 1276 1352 1392 1444 1504 A B Fig. 3. Room-temperature resonance Raman spectra for the 915 nm (A) and 889 nm (B) LH1–RC preparations from Tch. tepi- dum. The excitation wavelength was 514 nm. ∗ ∗ 1675 1671 1065 1641 1641 ∗ ∗ ∗ ∗ ∗ 1171 (R4) 1444 (R3) 1534 (R2) 1609 (R1) 1170 (R4) 1436 (R3) 1540 (R2) Intensity / a.u. 1610 (R1) 1000 1200 1400 1600 Raman shift·cm –1 1024 (ν 3 ) 1147 (ν 2 ) 1147( ν 2 ) 1023 (ν 3 ) 1504 (ν 1 ) 1504 (ν 1 ) A B Fig. 4. Room-temperature FT-Raman spectra for the 915 nm (A) and 889 nm (B) LH1–RC core complexes from Tch. tepidum. The excitation wavelength was 1064 nm. Ca 2+ -binding motif in an LH1 complex F. Ma et al. 1742 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS from BChl a, and are sensitive to the core size of bac- teriochlorin and the molecular conformation of BChl a. These modes are known to be conserved in various LHs [25,26]. It is worthy of noting that the band at 1065 cm )1 in the Raman spectrum of 915 nm LH1–RC is not seen in the 889 nm form (Fig. 4A,B), probably due to variation in the resonance conditions of Raman excitation [24]. For both forms of LH1–RC, the presence of meth- ane bridge stretching at approximately 1610 cm )1 (R1) confirms the penta-coordination of BChl a molecules [27] that is often seen when the a- and b-polypeptides of LH1 have higher flexibility [28]. Raman lines R5 or R6 overlapped with the intense carotenoid band (m 3 ) and therefore cannot be resolved. For both the 915 and 889 nm forms, the R1 and R4 Raman lines appear at similar frequencies (Table 1); however, the R2 and the R3 frequencies vary considerably, i.e. 1540 versus 1531 cm )1 and 1436 versus 1444 cm )1 , respectively. The R1–R4 lines of the LH1 complexes from Rb. sph- aeroides 2.4.1 and Rhodospirillum (Rsp.) rubrum G 9 [26] are conserved in the 889 nm LH1–RC form from Tch. tepidum. Therefore, the macrocycle configurations of BChl a are most likely similar among these com- plexes. However, the R2 and R3 lines and those with asterisks in the Raman spectrum of the 915 nm form are distinctly different from those of the 889 nm form, both in frequency and intensity, suggesting significant differences in the BChl a conformations between the two LH1–RC forms of Tch. tepidum. According to recent theoretical studies on the peridinin–chlorophyll– protein complex and the light-harvesting complex II Table 1. Raman shifts obtained from the near-infrared FT-Raman spectra of BChl a in the 915 and 889 nm LH1–RC forms from Tch. tepi- dum (see Fig. 4) and the corresponding assignments. ‘Carotenoid’ indicates that the Raman lines of BChl a overlap with those originating from carotenoid. Intensities are indicated after the Raman shifts. Raman shift (cm )1 ) Key Raman lines b Assignments c 915 nm LH1 889 nm LH1 LH1 ⁄ LH2 a 1671 s 1676 m 1640–1680 mC 9 =O, mC 10a =O 1641 m 1641 m 1630–1660 mC 2a =O 1610 w 1609 vw 1608–1609 R1 as mC a C m (a, b, c, d) 1585 w 1594 vw 1570–1590 as mC a C m (c, d) 1567 w 1567 w as mC a C m (a, b) 1540 sh 1534 sh 1530–1537 R2 mC b C b ,smC a C m (c), mCN(III) 1456 vw — — s mC a C m (a), mCN(II) 1436 vw 1444 w 1444–1445 R3 CH 3 bend, s mC a C m (d), mCN(IV) — 1408 sh 1406–1409 CH 3 bend, C 6 C 16 1394 sh 1391 w 1408–1415 mCN(I), dC m H(a, d), CH 3 bend 1372 m 1370 w 1385–1396 dC m H(d), CH 3 bend 1354 sh 1350 vw 1371–1376 mCN(III), dC m H(b), CH 3 bend, d defs 1331 vw 1333 sh 1346–1348 mCN(III), dC m H(b), CH 3 bend, CH 2 bend, CH bend1291 m 1291 m 1284–1288 1279 m 1281 m 1273–1277 CH 3 bend, CH bend 1258 w 1253 m 1252–1257 mCN(IV), mC 7 C 17 , d defs 1236 m 1206 sh 1235–1237 dC m H(d), mC a C b (II), CH 2 bend, CH bend1194 sh 1209–1212 1170 sh 1171 sh 1173–1175 R4 dC m H(b) carotenoid vs carotenoid vs 1142 ⁄ 1137 ⁄ carotenoid R5 mCN(III), mC 5 C 5a , CH bend 1117 sh 1116 m 1116–1119 mCN(I) 1093 vw 1089 vw 1090–1095 1065 s — 1065–1066 d (IV), CH 3 bend, CH 2 bend carotenoid s carotenoid s 1024 ⁄ 1029 ⁄ carotenoid R6 CH 3 bend, mCC (saturated) 1000 m 998 vw 1000–1003 CH 3 bend, mC 2a =O 969 w 959 m 967–969 mC 10b O, mC 10 C 10a , 948 w 949–952 d defs — 927 w 925–927 s dNCC m (d) a These Raman frequencies are from reference [25] and are given for comparison. b R1–R4 are key Raman lines that are sensitive to the core size of BChl a [26]. c Assignments based on reference [24]. See Scheme 1 for the numbering system of BChl a . F. Ma et al. Ca 2+ -binding motif in an LH1 complex FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1743 (LHCII) chlorophylls [29], distortion of the Chl a mac- rocycle is the key structural factor governing the Q y absorptive transition energy. As seen in Fig. 4 (and with reference to the number- ing system of BChl a in Scheme 1), the three Raman lines above 1600 cm )1 for the 915 nm ⁄ 889 nm forms may be ascribed to the stretching modes of the methane bridge (1610 ⁄ 1609 cm )1 ), the C 2 acetyl (1641 ⁄ 1641 cm )1 ) and the C 9 keto–C 10a acetyl carbo- nyls (1671 ⁄ 1676 cm )1 ). It is known that, for free BChl a in nonpolar solvent, lines for the two carbonyl stretching modes appear at 1663 and 1685 cm )1 , but downshift as much as approximately 40 cm )1 for BChl a bound to protein via an H-bond, and, impor- tantly, a downshift of the C 2 acetyl stretching correlates linearly with the red shift of the Q y absorption [25]. As the frequency of the particular mode at 1641 cm )1 is identical between the 915 and 889 nm forms, the H-bond interaction with the C 2 acetyl carbonyl cannot be responsible for the shift of Q y absorption from 915 to 889 nm. Compared with the 889 nm form, the C 9 keto ⁄ C 10a acetyl carbonyl stretching of the 915 nm form shows a downshift of 5 cm )1 , indicating a stron- ger H-bond between the Trp )8 residue of the b-subunit and the C 10a acetyl carbonyl (see below). 3D modeling of Ca 2+ -binding motifs The amino acid sequences of LH1 from Tch. tepidum show the highest homology to the LH1 peptides from Rs. rubrum, i.e. the 50.0% and 53.3% (E-value, 2 · 10 )11 ⁄ 0) identity for a- and b-polypeptides, respec- tively. For comparison, the corresponding identities to LH2 of Rs. molischianum are 35.0% and 40.5% (E-value, 2 · 10 )4 ⁄ 6 · 10 )9 ), respectively, and those to the LH2 peptides of Rps. acidophila are 28.6% and 34.2% (E-value, 3 · 10 )3 ⁄ 2 · 10 )5 ). However, as the available structures of the a- and b-polypeptides of H CH 2 H = R Scheme 1. BChl a chemical structure and numbering. Right, numbering of carbon atoms according to the Fischer system. Left, genetic labeling of meso and pyrrolic carbon atoms. Ca 2+ -binding motif in an LH1 complex F. Ma et al. 1744 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS LH1 from Rs. rubrum were determined independently in solution [7], they are not suitable to serve as tem- plates for the LH1 of Tch. tepidum with sufficient structural details of BChl a molecules and the loop domain at the C-terminus. Figure 5A shows the BChl a binding sites and the loop motif of the LH1 polypeptides of Tch. tepidum based on SWISS-MODEL modeling using the LH2 template from Rs. molischianum; those obtained using the LH2 template from Rps. acidophila are presented in Fig. 5B. With regard to the possible H-bond to BChl a, the NH 2 of b-Trp )8 falls into close proximity to BChl a, i.e. 3.63 A ˚ to O 10a when the LH2 crystallo- graphic structure of Rs. molischianum is used as a tem- plate (Fig. 5A, upper right) and 3.62 and 5.86 A ˚ to O 10b and O 10a , respectively, when the LH2 of Rps. aci- dophila was used (Fig. 5B, upper right). In the LH2s of Rs. molischianum and Rps. acidophila, the amino acid corresponding to b-Trp )8 in LH1 of Tch. tepidum is phenylalanine (Phe), which cannot form an H-bond with the acetyl carbonyl of BChl a. Previous 3D struc- tural modeling of LH1 of Roseospirillum parvum 930I proved that the H-bonds between the thiol groups of cysteine (a-Cys +3 , b-Cys )4 ) and BChl a are responsible for the long-wavelength LH1–Q y absorption (909 nm) [30]. Similarly, the H-bonds found in the LH1 of Tch. tepidum may be responsible for the extremely red absorption of BChl a (915 nm), although other factors such as BChl–BChl excitonic interactions are certainly also in operation. The possible Ca 2+ coordinations optimized by means of SWISS-MODEL modeling based on the LH2 templates of Rs. molischianum and Rps. acidophila, respectively, are shown in the lower right parts of Fig. 5A,B. In both cases, the Ca 2+ -binding cavities are localized in the C-termini, which comprise O of Leu )4 , Ser )5 , Thr )6 , OD1 of Asp )7 and OD1 and OD2 of Asp )13 in Fig. 5A, O of Val )3 , Leu )4 , Ser )5 , Thr )6 , Asp )7 and OG of Ser )5 in Fig. 5B (O, OD1 ⁄ 2 and OG are oxygen atoms of the backbone carbonyl, side-chain acetyl or hydroxyl carbonyl, and side-chain hydroxyl, respectively). The Ca 2+ chelation motifs agree well with the EF-hand characteristics, i.e. they tend to localize to the helix-loop-helix motifs with a coordina- tion number of 6 or 7. Figure 6 shows the results of MODELLER model- ing based on an averaged template (the LH1 from Rs. rubrum and the LH2s from Rs. molischianum and Rps. acidophila). The H-bond between b-Trp )8 and BChl a and the presence of a Ca 2+ coordination cavity (consisting of O of Val )3 , Leu )4 , Ser )5 , Thr )6 , Asp )7 and OD2 of Ser )13 ) within a helix-loop-helix motif are predicted, which is similar to the results of A B α α α α β α β β β Fig. 5. Three-dimensional models of a polypeptide subunit of LH1 of Tch. tepidum obtained by SWISS-MODEL modeling using the crystallographic structures of LH2 from Rs. molischianum (A) and Rps. acidophila (B) as templates. In each panel, the structures within circles were magnified to show more detail and these are shown at upper and lower right. Color codes: deep blue, secondary structure of polypeptide subunit; orange, amino acid; green, BChl a; red, His; pink, Trp )8 ; yellow, hydrogen atoms in NH 2 of Trp )8 ; light grey, oxygen atoms of BChl a; purple, oxygen atoms most probably coordinating to Ca 2+ . F. Ma et al. Ca 2+ -binding motif in an LH1 complex FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1745 SWISS-MODEL modeling. To our surprise, a consid- erable tilt of the optimized polypeptides with respect to the template is predicted, especially for the b-poly- peptide. Furthermore, the orientation of the histidine (His) coordinating to BChl a changes significantly in a- and b-polypeptides as seen in the top right of Fig. 6. This implies a large difference in the orientation of BChl a molecules between the LH1 of Tch. tepidum and the template, and, as a result, a large difference in the BChl–BChl excitonic interactions. In addition, the results show that the locations of a- and b-His residues are rather different between the LH1 of Tch. tepidum and the template. It is therefore expected that coordi- nation of His residues to BChl a induces considerable structural heterogeneity in the BChl a molecules bound to a- and b-polypeptides, and this is supported by a recent transient spectroscopic study of LH–RC forms from Tch. tepidum [14]. Although the modeling results for Ca 2+ -induced conformational changes in the LH1 of Tch. tepidum are preliminary and qualitative, they reveal basic struc- tural differences between the 915 and 889 nm forms of LH1–RC from Tch. tepidum, e.g. the strength of the H-bond between the b-Trp )8 residue and the C 10a ace- tyl carbonyl of BChl a, the excitonic interaction among the BChl a molecules in LH1, and deformation of the BChl a macrocycle induced by Ca 2+ binding, all of which may account for the low absorptive transition energy of BChl a molecules in native LH1. We pro- pose that the presence of a specific Ca 2+ -binding motif in the a-, b-subunit of LH1 is responsible for the long- wavelength LH1–Q y absorption of Tch. tepidum,as well as for the high thermal stability of this particular pigment–protein assembly. Conclusion Based on the spectroscopic and 3D structural modeling results for the 915 and 889 nm forms of LH1–RC from Tch. tepidum, this paper proposes a specific Ca 2+ -coordination cavity localized to the C-terminus of the a-subunit of LH1, which agrees with the EF-hand characteristics. This Ca 2+ -binding motif may be responsible for the reversible conformation change in the a- and b -polypeptides between the forms, which in turn lead to changes in the arrangement of BChl a molecules, in the strength of the H-bond between b-Trp )8 and the O 10 of BChl a, and in distortion of BChl a macrocycle. All of these structural variations are helpful to understand the long-wavelength Q y absorption of the native LH1–RC complex from Tch. tepidum. Furthermore, thermal equilibrium among the α α α α β β β Fig. 6. Three-dimensional model of the LH1 polypeptides of Tch. tepidum obtained by MODELLER modeling based on the aver- aged template (LH1 from Rs. rubrum and the LH2s from Rs. molischianum and Rps. acidophila). The model at the top right shows details of the BChl a binding motifs of the optimized LH1 and the template, while that at the bottom right shows details of the possible Ca 2+ -binding cavity; both models are re-oriented for clarity with respect to the model on the left. Color code: dark green, secondary structure of LH1 polypeptides; blue, secondary structure of the template; orange, amino acids of LH1 polypeptides; red, His coordinating to BChl a in LH1 of Tch. tepidum; purple, His coordi- nating to BChl a in the template; yellow, hydrogen atoms coordinating to BChl a; light blue: oxygen atoms coordinating to Ca 2+ . Ca 2+ -binding motif in an LH1 complex F. Ma et al. 1746 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS excitonic states of BChls and other structural changes in the 915 nm form of LH1, as reflected by the temper- ature-dependent fluorescence spectra, reveal higher dissociation enthalpy of this complex with respect to the 889 nm form, which may account for the higher thermal stability of the native LH1–RC complex from Tch. tepidum. Experimental procedures Preparation of LH1–RC complexes Chromatophore was isolated by sonication of the Tch. tepi- dum cells suspended in 20 mm Tris ⁄ HCl buffer (pH 8.5) followed by differential centrifugation at 4 °C for 15 min (5000 g). Chromatophores thus obtained were extracted with 0.35% w ⁄ v lauryldimethylamine N-oxide at 25 °C for 60 min, followed by centrifugation at 4 °C for 90 min (150 000 g). The LH1–RC core complex with an LH1–Q y absorption maximum at 915 nm (Fig. 1A) was prepared as described previously [24]. The final concentration of LH1– RC was determined to be approximately 10 lm by using a molar extinction coefficient for BChl a of e 915 nm = 4.3 · 10 3 mm )1 cm )1 [11]. As the preparation was eluted using a linear gradient of CaCl 2 from 10 to 25 mm, the ionic force was estimated be approximately 75 mm. This LH1–RC preparation is considered to be intact as the LH1–Q y absorption maximum, 915 nm, is similar to that of the chromatophore. The 889 nm preparation, i.e. the LH1–RC complex with an LH1–Q y absorption maximum at approxi- mately 889 nm (Fig. 1B), was prepared by adding 200 mm EDTA to the intact 915 nm form. For spectroscopic measurements, these preparations were suspended in buffer containing 20 mm Tris ⁄ HCl and 0.8% w ⁄ v n-octyl-b-d- glucopyranoside (pH 7.5). Steady-state UV-visible and near-infrared fluorescence spectroscopy UV-visible absorption spectroscopy with a spectral resolu- tion of 0.5 nm was performed using a U-3310 spectropho- tometer (Hitachi, Japan). Near-infrared fluorescence spectra (spectral resolution of 0.25 nm) were recorded using a liquid nitrogen-cooled linear photodiode array (OMA V: 10242.2 Princeton Instruments, Trenton, NJ, USA) attached to an imaging polychromater (SpectraPro 2300i; Acton Research, Acton, MA, USA), for which excitation pulses at 590 nm (approximately 2 mJÆpulse )1 , approximately 7 ns, 10 Hz) were supplied by an optical parametric oscillator (Quanta- Ray MOPO-SL; Spectra Physics, Mountain View, CA, USA) driven by an Nd 3+ :YAG laser (Quanta-Ray PRO-230; Spec- tra Physics). Sample temperatures were controlled exactly in the range 273–323 K using a water-flow type thermostat (RTE-110; Neslab Instruments Inc., Newington, NH, USA). Resonance Raman and near-infrared FT-Raman spectroscopy Room-temperature resonance Raman spectra (spectral reso- lution of 1.4 cm )1 ) were recorded with a liquid nitrogen- cooled CCD detector (SPEC-10-400B ⁄ LN; Roper Scientific Research, Trenton, NJ, USA) attached to a 0.5 m poly- chromator (grating density 1200 grooves per mm, Spectro- pro 550i; Acton Research Corporation). A continuous-wave Ar + laser (2060-10S; Spectra Physics) provided the Raman excitation power of 1.8 mW at 514 nm. Raman scattering light was collected in a backscattering geometry, and was focused onto the entrance slit of the polychromator after passing through a Raman notch filter (HSNF-514.0-1.5; Kaiser Optical Systems, Ann Arbor, MI, USA). The Raman spectra were obtained using an exposure time of 15 s and a spectral resolution of 1.4 cm )1 . The absorbance of the LH1–RC samples was 5 cm )1 at 514 nm. Raman spectra, with pre-resonance to the Q y transition of BChl a, were recorded on a FT-Raman spectrometer (DIGILAB FTS-3500; Bio-Rad, Krefeld, Germany) with a resolution of 0.5 cm )1 ; the excitation source was a continu- ous-wave Nd 3+ :YAG laser operated at 1064 nm. The spec- tra were obtained by averaging of 200 scans. The optical densities of the two forms of LH1–RC were adjusted to 120 cm )1 at the maximal Q y absorption. 3D structural modeling of LH1 The 3D modeling was performed using two methods: SWISS- MODEL, accessed using the Deep View Swiss-PDB Viewer version 4.0 [31–33], and MODELLER [34,35]. SWISS- MODEL superimposes a template with the target sequence, and is fully automated by a homology-modeling server (http://www.expasy.ch/spdbv/). The a-, b-polypeptides of LH2 from Rps. acidophila (PDB record 1nkz) and Rs. molis- chianum (PDB record 1lgh) as well as those of LH1 from Rs. rubrum (PDB records 1xrd and 1wrg) were used as the templates. MODELLER is used for homology or compara- tive 3D modeling of protein structures. It implements com- parative modeling by satisfaction of spatial restraints, and can perform additional tasks such as de novo modeling of loops, etc. We used an average of the three templates above (multiple-model) to increase the accuracy. Five models were thus obtained, and the one with the lowest discrete optimized protein energy (DOPE potential) was chosen. Sequence identity between target LH1 polypeptides from Tch. tepidum and each template was calculated using MODELLER. Acknowledgements This work has been supported by the Natural Science Foundation of China (grant nos NSFC 20703067 and 20673144, and NSFC-JSPS. 20711140133) and by the F. Ma et al. Ca 2+ -binding motif in an LH1 complex FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1747 National Basic Research Program of China (grant no. 2009CB220008). References 1 Scheuring S, Francia F, Busselez J, Melandri BA, Rigaud JL & Le ´ vy D (2004) Structural role of PufX in the dimerization of the photosynthetic core com- plex of Rhodobacter sphaeroides. J Biol Chem 279, 3620–3626. 2 Siebert CA, Qian P, Fotiadis D, Engel A, Hunter CN & Bullough PA (2004) Molecular architecture of photo- synthetic membranes in Rhodobacter sphaeroides: the role of PufX. EMBO J 23, 690–700. 3 Cogdell RJ, Southall J, Gardiner AT, Law CJ, Gall A, Roszak AW & Isaacs NW (2006) How purple photo- synthetic bacteria harvest solar energy. C R Chim 9, 201–206. 4 McDermott G, Prince SM, Freer AA, Hawthornthwa- ite-Lawless AM, Papiz MZ, Cogdell RJ & Isaacs NW (1995) Crystal structure of an integral membrane light- harvesting complex from photosynthetic bacteria. Nature 374, 517–521. 5 Koepke J, Hu XC, Muenke C, Schulten K & Michel H (1996) The crystal structure of the light-harvesting com- plex II (B800–850) from Rhodospirillum molischianum. Structure 4, 581–597. 6 Roszak AW, Howard TD, Southall J, Gardiner AT, Law CJ, Isaacs NW & Cogdell RJ (2003) Crystal struc- ture of the RC–LH1 core complex from Rhodopseudo- monas palustris. Science 302, 1969–1972. 7 Wang ZY, Gokan K, Kobayashi M & Nozawa T (2005) Solution structures of the core light-harvesting a and b polypeptides from Rhodospirillum rubrum: impli- cations for the pigment–protein and protein–protein interactions. J Mol Biol 347, 465–477. 8 Madigan MT (1984) A novel photosynthetic purple bac- terium isolated from a Yellowstone hot spring. Science 225, 313–315. 9 Watson AJ, Hughes AV, Fyfe PK, Wakeham MC, Hol- den-Dye K, Heathcote P & Jones MR (2005) On the role of basic residues in adapting the reaction center– LH1 complex for growth at elevated temperatures in purple bacteria. Photosynth Res 86, 81–100. 10 Fathir I, Ashikaga M, Tanaka K, Katano T, Nirasawa T, Kobayashi M, Wang ZY & Nozawa T (1998) Bio- chemical and spectral characterization of the core light harvesting complex 1 (LH1) from the thermophilic pur- ple sulfur bacterium Chromatium tepidum. Photosynth Res 58, 193–202. 11 Kimura Y, Hirano Y, Yu LJ, Suzuki H, Kobayashi M & Wang ZY (2008) Calcium ions are involved in the unusual red-shift of the light-harvesting 1 Q y transition of the core complex in thermophilic purple sulfur bacterium Thermochromatium tepidum. J Biol Chem 283, 13867–13873. 12 Wang ZY, Shimonaga M, Suzuki H, Kobayashi M & Nozawa T (2003) Purification and characterization of the polypeptides of core light-harvesting complexes from purple sulfur bacteria. Photosynth Res 78, 133– 141. 13 Suzuki H, Hirano Y, Kimura Y, Takaichi S, Kobayashi M, Miki K & Wang ZY (2007) Purification, character- ization and crystallization of the core complex from thermophilic purple sulfur bacterium Thermochroma- tium tepidum. Biochim Biophys Acta 1767, 1057–1063. 14 Ma F, Kimura Y, Zhao XH, Wu YS, Wang P, Fu LM, Wang ZY & Zhang JP (2008) Excitation dynamics of two spectral forms of the core complexes from photosynthetic bacterium Thermochromatium tepidum. Biophys J 95, 3349–3357. 15 Krebs J & Micnalak M (2007) Calcium: A Matter of Life or Death. Elsevier Press, Amsterdam. 16 Kretsinger RH (1972) Gene triplication deduced from the tertiary structure of a muscle calcium binding pro- tein. Nat New Biol 240, 85–88. 17 Toyoshima C & Nomura H (2002) Structural changes in the calcium pump accompanying the dissociation of calcium. Nature 418, 605–611. 18 Toyoshima C, Nomura H & Sugita Y (2003) Structural basis of ion pumping by Ca 2+ -ATPase of sarcoplasmic reticulum. FEBS Lett 555, 106–110. 19 Wu HM, Ra ¨ tsep M, Jankowiak R, Cogdell RJ & Small GJ (1998) Hole-burning and absorption studies of the LH1 antenna complex of purple bacteria: effect of pressure and temperature. J Phys Chem B 102, 4023–4034. 20 Kimura Y, Yu LJ, Hirano Y, Suzuki H & Wang ZY (2009) Calcium ions are required for the enhanced ther- mal stability of the light-harvesting-reaction center core complex from thermophilic purple sulfur bacterium Thermochromatium tepidum. J Biol Chem 284, 93–99. 21 Kuki M, Naruse M, Kakuno T & Koyama Y (1995) Resonance Raman evidence for 15-cis to all-trans photoisomerization of spirilloxanthin bound to a reduced form of the reaction center of Rhodospirillum rubrum S1. Photochem Photobiol 62, 502–508. 22 Qian P, Saiki K, Mizoguchi T, Hara K, Sashima T, Fujii R & Koyama Y (2001) Time-dependent changes in the carotenoid composition and preferential binding of spirilloxanthin to the reaction center and anhydro- rhodovibrin to the LH1 antenna complex in Rhodobium marinum. Photochem Photobiol 74, 444–452. 23 Papagiannakis E, Das SK, Gall A, van Stokkum IHM, Robert B, van Grondelle R, Frank HA & Kennis JTM (2003) Light harvesting by carotenoids incorporated into the B850 light-harvesting complex from Rhodo- bacter sphaeroides R-26.1: excited-state relaxation, Ca 2+ -binding motif in an LH1 complex F. Ma et al. 1748 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS [...]... binding-site and electronic properties in light-harvesting proteins of purple bacteria J Phys Chem B 101, 7227–7231 Fiedor L (2006) Hexacoordination of bacteriochlorophyll in photosynthetic antenna LH1 Biochemistry 45, 1910–1918 Zucchelli G, Brogioli D, Casazza AP, Garlaschi FM & Jennings RC (2007) Chlorophyll ring deformation 30 31 32 33 34 35 modulates Qy electronic energy in chlorophyll–protein complexes.. .Ca2+-binding motif in an LH1 complex F Ma et al 24 25 26 27 28 29 ultrafast triplet formation, and energy transfer to bacteriochlorophyll J Phys Chem B 107, 5642–5649 Frolov D, Gall A, Lutz M & Robert B (2002) Structural asymmetry of bacterial reaction centers: a Qy resonant Raman study of the monomer bacteriochlorophylls J Phys Chem A 106, 3605–3613... chlorophyll–protein complexes and generates spectral forms Biophys J 93, 2240–2254 Tuschak C, Beatty JT & Overmann J (2004) Photosynthesis genes and LH1 proteins of Roseospirillum parvum 930I, a purple non-sulfur bacterium with unusual spectral properties Photosynth Res 81, 181– 199 Peitsch MC (1995) Protein modeling by E-mail Nat Biotechnol 13, 658–660 Guex N & Peitsch MC (1997) SWISS-MODEL and the Swiss-Pdb viewer:... Simo¨ nin I, Scheer H & Robert B (1997) Resonance Raman spectroscopy of metal-substituted bacteriochlorophylls: characterization of Raman bands sensitive to bacteriochlorin conformation J Raman Spectrosc 28, 599–604 Lapouge K, Naveke A, Gall A, Ivancich A, Seguin J, ¨ Scheer H, Sturgis JN, Mattioli TA & Robert B (1999) Conformation of bacteriochlorophyll molecules in photosynthetic proteins from purple... Swiss-Pdb viewer: an environment for comparative protein modeling Electrophoresis 18, 2714–2723 Schwede T, Kopp J, Guex N & Peitsch MC (2003) SWISS-MODEL: an automated protein homology -modeling server Nucleic Acids Res 31, 3381–3385 ´ Martı´ -Renom MA, Stuart AC, Fiser A, Sanchez R, Melo F & Sali A (2000) Comparative protein structure modeling of genes and genomes Annu Rev Biophys Biomol Struct 29, 291–325... Comparative protein structure modeling of genes and genomes Annu Rev Biophys Biomol Struct 29, 291–325 Sali A & Blundell TL (1993) Comparative protein modeling by satisfaction of spatial restraints J Mol Biol 234, 779–815 FEBS Journal 276 (2009) 1739–1749 ª 2009 The Authors Journal compilation ª 2009 FEBS 1749 . Specific Ca 2+ -binding motif in the LH1 complex from photosynthetic bacterium Thermochromatium tepidum as revealed by optical spectroscopy and structural modeling Fei Ma 1,3 ,. of LH1 from Tch. tepidum have been purified and the amino acid sequences deter- mined [12]. In addition, the dimeric feature and the highly symmetric ring assembly of BChls in LH1, as well as the. is also known as the EF hand [16]. Proteins containing the EF hand are divided into two classes according to their functions: signaling and buffering ⁄ transport proteins. The former undergoing Ca 2+ -dependent conformational