Function of a PscD Subunit in a Homodimeric Reaction Center Complex of the Photosynthetic Green Sulfur Bacterium Chlorobium tepidum Studied by Insertional Gene Inactivation
2004; Elsevier BV; Volume: 279; Issue: 49 Linguagem: Inglês
10.1074/jbc.m410432200
ISSN1083-351X
AutoresYusuke Tsukatani, Ryo Miyamoto, Shigeru Itoh, Hirozo Oh‐oka,
Tópico(s)Spectroscopy and Quantum Chemical Studies
ResumoThe PscD subunit in the homodimeric "type I" photosynthetic reaction center (RC) complex of the green sulfur bacterium Chlorobium tepidum was disrupted by insertional mutagenesis of its relevant pscD gene. This is the first report on the use of the direct mutagenic approach into the RC-related genes in green sulfur bacteria. The RC complex of C. tepidum is supposed to form a homodimer of two identical PscA subunits together with three other subunits: PscB (FA/FB-containing protein), PscC (cytochrome cz), and PscD. PscD shows a relatively low but significant similarity in its amino acid sequence to PsaD in the photosystem I of plants and cyanobacteria. We studied the biochemical and spectroscopic properties of a mutant lacking PscD in order to elucidate its unknown function. 1) The RC complex isolated from the mutant cells showed no band corresponding to PscD on SDS-PAGE analysis. 2) The growth rate of the PscD-less mutant was slower than that of the wild-type cells at low light intensities. 3) Time-resolved fluorescence spectra at 77 K revealed prolonged decay times of the fluorescence from bacteriochlorophyll c on the antenna chlorosome and from bacteriochlorophyll a on the Fenna-Matthews-Olson antenna protein in the mutant cells. The loss of PscD led to a much slower energy transfer from the antenna pigments to the special pair bacteriochlorophyll a (P840). 4) The mutant strain exhibited slightly less activity of ferredoxin-mediated NADP+ photoreduction compared with that in the wild-type strain. The extent of suppression, however, was less significant than that reported in the PsaD-less mutants of cyanobacterial photosystem I. The evolutionary relationship between PscD and PsaD was also discussed based on a structural homology modeling of the former. The PscD subunit in the homodimeric "type I" photosynthetic reaction center (RC) complex of the green sulfur bacterium Chlorobium tepidum was disrupted by insertional mutagenesis of its relevant pscD gene. This is the first report on the use of the direct mutagenic approach into the RC-related genes in green sulfur bacteria. The RC complex of C. tepidum is supposed to form a homodimer of two identical PscA subunits together with three other subunits: PscB (FA/FB-containing protein), PscC (cytochrome cz), and PscD. PscD shows a relatively low but significant similarity in its amino acid sequence to PsaD in the photosystem I of plants and cyanobacteria. We studied the biochemical and spectroscopic properties of a mutant lacking PscD in order to elucidate its unknown function. 1) The RC complex isolated from the mutant cells showed no band corresponding to PscD on SDS-PAGE analysis. 2) The growth rate of the PscD-less mutant was slower than that of the wild-type cells at low light intensities. 3) Time-resolved fluorescence spectra at 77 K revealed prolonged decay times of the fluorescence from bacteriochlorophyll c on the antenna chlorosome and from bacteriochlorophyll a on the Fenna-Matthews-Olson antenna protein in the mutant cells. The loss of PscD led to a much slower energy transfer from the antenna pigments to the special pair bacteriochlorophyll a (P840). 4) The mutant strain exhibited slightly less activity of ferredoxin-mediated NADP+ photoreduction compared with that in the wild-type strain. The extent of suppression, however, was less significant than that reported in the PsaD-less mutants of cyanobacterial photosystem I. The evolutionary relationship between PscD and PsaD was also discussed based on a structural homology modeling of the former. Photosynthetic organisms convert light energy into electrochemical free energy by carrying out a series of light-driven electron transfer reactions. This process, which is fundamental for life, is mediated by reaction center (RC) 1The abbreviations used are: RC, reaction center; BChl, bacteriochlorophyll; Chl, chlorophyll; Fd, ferredoxin; FMO, Fenna-Matthews-Olson; FNR, ferredoxin:NADP+ oxidoreductase; MOPS, 4-morpholinepropanesulfonic acid. complexes. The RCs are primarily grouped into two types based on their terminal electron acceptors, type I (FeS-type) RCs and type II (quinone-type) RCs. Purple photosynthetic bacteria contain only type II RCs, which do not evolve oxygen, whereas oxygenic cyanobacteria and plants utilize both type I (photosystem I) and type II (photosystem II) RCs, which are connected in-line through the b6f complex. Green sulfur bacteria and heliobacteria have unique type I RCs, so-called "homodimeric" RCs, which are made of two identical core polypeptides. The "heterodimeric" type I and II RCs, which are found in all photosynthetic organisms other than green sulfur bacteria and heliobacteria, consist of a set of two partially different core polypeptides that produce a slightly asymmetric arrangement of cofactors. A three-dimensional structure of the heterodimeric RC was first obtained from the type II RC of the purple bacterium Blastochloris viridis in 1985 (1Deisenhofer J. Epp O. Miki K. Huber R. Michel H. Nature. 1985; 318: 618-626Crossref PubMed Scopus (2610) Google Scholar). Recently, those of photosystem I and II RCs from the thermophilic cyanobacterium Synechococcus elongatus were determined at high resolutions (2Jordan P. Fromme P. Witt H. Klukas O. Saenger W. Krauss N. Nature. 2001; 411: 909-917Crossref PubMed Scopus (2105) Google Scholar, 3Zouni A. Witt H. Kern J. Fromme P. Krauss N. Saenger W. Orth P. Nature. 2001; 409: 739-743Crossref PubMed Scopus (1789) Google Scholar). These structures have clarified that the folding motifs of membrane-spanning α-helices within core proteins as well as the spatial configurations of the electron transfer components are essentially identical in type I and II RCs, suggesting that they have a common origin. Various methods of gene manipulation have been applied to pursue research on heterodimeric RCs on the molecular level, and the results have led us to a comprehensive understanding of the electron transfer mechanisms and molecular architectures as well. The electron transfer reactions within these heterodimeric RCs occur via one of the two pathways that contain almost symmetrically arranged cofactors. On the other hand, homodimeric type I RCs have never been successful targets for crystal structural analysis and mutagenic approach due to the obligate anaerobic and photoautotrophic nature of green sulfur bacteria and heliobacteria. In fact, the RC complexes isolated from them are very fragile when exposed to oxygen, and many RC-related genes are essential. However, a study on the molecular level of the homodimeric RCs, which have a simple architecture and still retain the features of an ancestral RC, will offer valuable information to understand the mechanism and evolution of photosynthetic apparatuses. The RC complex of the green sulfur bacterium Chlorobium tepidum consists of four subunits, PscA, PscB, PscC, and PscD with an antenna size of about 30 bacteriochlorophyll (BChl) molecules, whereas the heterodimeric photosystem I complex is made of 12 polypeptides with a much larger antenna size of nearly 100 chlorophyll (Chl) molecules (4Hauska G. Schoedl T. Rémigy H. Tsiotis G. Biochim. Biophys. Acta. 2001; 1507: 260-277Crossref PubMed Scopus (144) Google Scholar, 5Heathcote P. Jones M.R. Fyfe P.K. Phil. Trans. R. Soc. Lond. B. 2003; 358: 231-243Crossref PubMed Scopus (40) Google Scholar). The functions of PscA, PscB, and PscC have been studied intensively using biochemical and spectroscopic methods. A pair of PscA forms a core protein, which is supposed to make up a homodimeric (PscA)2 structure, unlike a PsaA/PsaB heterodimeric one in photosystem I (6Büttner M. Xie D. Nelson H. Pinther W. Hauska G. Nelson N. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 8135-8139Crossref PubMed Scopus (159) Google Scholar). This core protein contains a special pair, P840, a primary electron acceptor, Chl a-670 (A0), and an iron-sulfur center, FX, although the existence of the acceptor quinone (A1) remains controversial (4Hauska G. Schoedl T. Rémigy H. Tsiotis G. Biochim. Biophys. Acta. 2001; 1507: 260-277Crossref PubMed Scopus (144) Google Scholar). PscB ligates two iron-sulfur clusters that are comparable with the FA and FB centers in PsaC of photosystem I but these proteins show almost no significant similarity to each other in their amino acid sequences (6Büttner M. Xie D. Nelson H. Pinther W. Hauska G. Nelson N. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 8135-8139Crossref PubMed Scopus (159) Google Scholar, 7Vassiliev I.R. Antonkine M.L. Golbeck J.H. Biochim. Biophys. Acta. 2001; 1507: 139-160Crossref PubMed Scopus (103) Google Scholar). PscC is the monoheme-type cytochrome cz, which serves as a direct electron donor to the special pair of BChl a, P840 (8Okkels J.S. Kjær B. Handdon O. Svendsen I. Møller B.L. Schller H.V. J. Biol. Chem. 1992; 267: 21139-21145Abstract Full Text PDF PubMed Google Scholar, 9Oh-oka H. Kamei S. Matsubara H. Iwaki M. Itoh S. FEBS Lett. 1995; 365: 30-34Crossref PubMed Scopus (60) Google Scholar, 10Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1998; 37: 12293-12300Crossref PubMed Scopus (33) Google Scholar), and has no counterpart in photosystem I. This cytochrome cz mediates the electron transfer between the cytochrome bc and RC complexes without participation of other soluble cytochromes (10Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1998; 37: 12293-12300Crossref PubMed Scopus (33) Google Scholar). However, the function of PscD has been unknown because of the absence of any cofactors within it. Three-dimensional imaging analysis with the electron microscopic technique (STEM) suggested that PscD was in contact with the Fenna-Matthews-Olson (FMO) protein on the cytoplasmic side (11Rémigy H. Stahlberg H. Fotiadis D. Muller S.A. Wolpensinger B. Engel A. Hauska G. Tsiotis G. J. Mol. Biol. 1999; 290: 851-858Crossref PubMed Scopus (50) Google Scholar). Although PscD has been assumed to function in the same way as PsaD in photosystem I, as judged from their sequence similarities (12Hager-Braun C. Xie D. Jarosch U. Herold E. Büttner M. Zimmermann R. Deutzmann R. Hauska G. Nelson N. Biochemistry. 1995; 34: 9617-9624Crossref PubMed Scopus (51) Google Scholar), there is no definite evidence for this speculation. The PsaD of photosystem I has been demonstrated to stabilize FA/FB-containing PsaC and facilitate the electron transfer reaction to ferredoxin (Fd). The Chlorobium RC complex also donates electrons to Fd (13Seo D. Tomioka A. Kusumoto N. Kamo M. Enami I. Sakurai H. Biochim. Biophys. Acta. 2001; 1503: 377-384Crossref PubMed Scopus (24) Google Scholar), and NADPH is finally produced by Fd:NADP+ oxidoreductase (FNR) (13Seo D. Tomioka A. Kusumoto N. Kamo M. Enami I. Sakurai H. Biochim. Biophys. Acta. 2001; 1503: 377-384Crossref PubMed Scopus (24) Google Scholar, 14Kjær B. Scheller H.V. Photosynth. Res. 1996; 47: 33-39Crossref PubMed Google Scholar). Furthermore, the Chlorobium RC complex is associated with antenna protein, the FMO protein. The FMO protein is water-soluble and contains seven molecules of BChl a. X-ray crystallographic analyses conducted by two research groups have shown that this protein forms a trimeric structure (15Matthews B.W. Fenna R.E. Bolognesi M.C. Schmid M.F. J. Mol. Biol. 1979; 131: 259-285Crossref PubMed Scopus (217) Google Scholar, 16Li Y.-F. Zhou W. Blankenship R.E. Allen J.P. J. Mol. Biol. 1997; 271: 456-471Crossref PubMed Scopus (179) Google Scholar). It is estimated that one or two FMO trimers have been included in the RC preparations so far reported, probably depending on the chemical properties of the detergents used in their isolation procedures (4Hauska G. Schoedl T. Rémigy H. Tsiotis G. Biochim. Biophys. Acta. 2001; 1507: 260-277Crossref PubMed Scopus (144) Google Scholar, 17Sakurai H. Kusumoto N. Inoue K. Photochem. Photobiol. 1996; 64: 5-13Crossref Scopus (37) Google Scholar, 18Rémigy H. Hauska G. Müller S.A. Tsiotis G. Photosynth. Res. 2002; 71: 91-98Crossref PubMed Scopus (24) Google Scholar). In the photosynthesis of green sulfur bacteria, light energy is captured by a supramolecular antenna complex, so-called chlorosome, which contains self-aggregates formed by large amounts of BChl c, d, or e depending on the species (e.g. the chlorosomes of C. tepidum contain BChl c aggregates) (19Blankenship R.E. Olson J.M. Miller M. Blankenship R.E. Madigan M.T. Bauer C.E. Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands1995: 399-435Google Scholar, 20Olson J.M. Photochem. Photobiol. 1998; 67: 61-75Crossref Scopus (326) Google Scholar). The BChl aggregates in chlorosomes make rod structures that are surrounded by a lipid monolayer envelop attached to the cytoplasmic surface of the inner membrane. The light energy absorbed by the BChl c aggregates in chlorosomes is transferred to the baseplate, which contains a small amount of BChl a, and then to the FMO protein, which seems to be located between chlorosomes and the RC complex and finally to the RC complex. In this study, we constructed a C. tepidum mutant lacking PscD with the recently established natural transformation system in this organism (21Frigaard N.-U. Bryant D.A. Appl. Environ. Microbiol. 2001; 67: 2538-2544Crossref PubMed Scopus (78) Google Scholar). PscD was shown to be involved in the efficient energy transfer from chlorosomes to P840, probably by allocating FMO proteins properly, and to enhance the NADP+ photoreduction via Fd. PscD was suggested to have an important role in the survival of green sulfur bacteria, even in the light-limited environment in which they are found in nature. Growth Conditions—The strain of C. tepidum used in this study was WT2321 (22Wahlund T.M. Madigan M.T. J. Bacteriol. 1995; 177: 2583-2588Crossref PubMed Google Scholar). For liquid cultivation and plating incubation, CL and CP media were prepared, respectively, as previously described (21Frigaard N.-U. Bryant D.A. Appl. Environ. Microbiol. 2001; 67: 2538-2544Crossref PubMed Scopus (78) Google Scholar). The growth temperature was set at 40 °C during the present work. The anaerobic growth on CP plates was carried out in an anaerobic jar (model G, code HP31; OXOID, Ltd.) equipped with a palladium catalyst andaH2/CO2-generating gas pack (BBL Gas Pack; BD Biosciences) and supplemented with 0.1 g of thioacetoamide to generate H2S by adding 1 ml of 0.5 m HCl. Plasmid Construction and Transformation of C. tepidum—A 1.2-kb fragment containing the pscD gene of C. tepidum was amplified by PCR, using a pscdF primer (5′-CTGAATTCAGTGGTACGAGAAGGCCATCC) and a pscdR primer (5′-CGGAATTCCGCTTGGCTGCAATTGCATCG), and cloned into the EcoRI site of pUC18, yielding plasmid pCD1. A 2.0-kb fragment containing the aadA gene, which was produced by SmaI digestion of pHP45Ω (23Prentki P. Krisch H.M. Gene (Amst.). 1984; 29: 303-313Crossref PubMed Scopus (1353) Google Scholar), was inserted into the NaeI site of pCD1, yielding plasmid pCDA (Fig. 1). Plasmid pCDA was then prepared in large amounts by using a MIDI-prep kit (Invitrogen). About 1 μg of AhdI-digested pCDA was used for the transformation. A natural transformation method was performed as described previously by Frigaard and Bryant (21Frigaard N.-U. Bryant D.A. Appl. Environ. Microbiol. 2001; 67: 2538-2544Crossref PubMed Scopus (78) Google Scholar). Transformants grown on selective (Smr/Spr) CP plates were restreaked three times onto selective CP plates. Each single colony on the third CP plate was inoculated into a fresh liquid CL medium containing selective antibiotics. The cultures to start growth were cultivated two or three times and used for further investigations. Preparation of Genomic DNA and Hybridization—Cell lysates were prepared essentially as previously reported (24Tsukatani Y. Matsuura K. Masuda S. Shimada K. Hiraishi A. Nagashima K.V.P. Photosynth. Res. 2004; 79: 83-91Crossref PubMed Scopus (35) Google Scholar). An equal volume of phenol was added to the cell lysate and centrifuged for 3 min at 15,000 × g, and the supernatant was then mixed with an equal volume of chloroform. After recentrifugation, nucleic acids in the aqueous phase were precipitated with 99.5% (v/v) ethanol supplemented with 300 mm sodium acetate, washed with 70% (v/v) ethanol, and suspended in TE buffer (10 mm Tris-HCl, 1 mm EDTA, pH 8.0). RNase was added to a final concentration of 50 μgml–1, and the mixture was incubated for 30 min at 37 °C. The suspension was mixed with 3 times the volume of the polyethylene glycol solution (13% (w/v) polyethylene glycol, 1.6 m NaCl) and incubated for 30 min on ice. Genomic DNA was precipitated, washed, and suspended in TE buffer. One microgram of genomic DNA was digested by HincII and used for hybridization analysis. Southern hybridization was carried out according to a manual on molecular cloning (25Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar) or instructions supplied by the manufacturer. PCR analysis was also used as a convenient means to confirm the insertional inactivation of the targeted gene. DNA fragments containing the pscD gene were amplified by the same primers as those used for cloning. The aadA marker in the mutant genome was detected by using the primers aadA946 and aadA947 (26Frigaard N.-U. Voigt G.D. Bryant D.A. J. Bacteriol. 2003; 184: 3368-3376Crossref Scopus (64) Google Scholar). Preparation of the RC Complex and SDS-PAGE—The RC complex was isolated from the wild-type and mutant cells as described previously by Oh-oka et al. (27Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1997; 36: 9267-9272Crossref PubMed Scopus (45) Google Scholar). All procedures for the preparation were carried out under anaerobic conditions in a chamber (Coy Laboratory Products, Ann Arbor, MI). SDS-PAGE was performed according to Laemmli (28Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (212360) Google Scholar). After electrophoresis, the separated protein bands were stained with Coomassie Brilliant Blue. A protein marker kit for the molecular mass estimation was purchased from New England BioLabs Inc. Spectral Analysis—The steady-state fluorescence emission spectra and absorption spectra were measured with a spectrofluorophotometer RF-1500 (Shimadzu) and spectrophotometer UV-3101PC (Shimadzu), respectively. The cells were suspended in 40 mm MOPS-NaOH buffer (pH 7.0) in an anaerobic chamber and placed in an air-tight anaerobic cuvette. The concentrations of the samples were adjusted to give an absorbance of 0.5 at 750 nm when measuring the fluorescence spectra. The time-resolved fluorescence spectra and kinetics were measured by a streak scope (Hamamatsu Photonics, Hamamatsu) with an excitation flash from a laser diode of 50-ps full width at half-maximum intensity operated at 1 MHz (Hamamatasu Photonics, Hamamatsu). The fluorescence excited by a laser beam was focused onto the entrance slit of the 30-cm monochrometer and dispersed depending on the wavelength inside the monochrometer along the horizontal axis. An image of the spectrum of the fluorescence at the exit slit of the monochrometer then excited the image intensifier at the entrance of the streak camera. A number of electrons produced by each photon in the image intensifier were biased by a time-dependent electric field along the vertical axis with respect to the delay of arrival and made an image on the phosphor plate. The images of each photon, whose location was biased due to wavelength along the x axis and the arrival time along the y axis, were captured by a CCD camera. The location of the trace of each photon on each image of CCD was counted and accumulated in the memory of a computer as shown in Fig. 6A. The ESR spectra were recorded using a Bruker ESP-300 EPR spectrometer equipped with a standard resonator (TE102). A gas flow temperature control system (CF935; Oxford Instruments) was used. Samples for measurements were chlorosome-free membranes prepared by disruption of cells and subsequent centrifugations before solubilization with detergents (10Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1998; 37: 12293-12300Crossref PubMed Scopus (33) Google Scholar). NADP+Photoreduction—The rates of ferredoxin-mediated NADP+ photoreduction were measured using membrane preparations (A810 = 0.7, corresponding to ∼3 μg of BChl a/ml) in a 1-ml reaction mixture containing 50 mm Tris-HCl (pH 7.8), 10 mm MgCl2, 8 mm sodium ascorbate, 100 μm 2,6-dichloroindophenol, 50 μm 2,3,5,6-tetramethyl-p-phenylenediamine, 0.05% n-dodecyl-β-maltoside, 5 mm glucose, 2 units of glucose oxidase, 20 units of catalase, 0.5 mm NADP+, 0.2 μm FNR, and various concentrations of ferredoxin, basically according to the compositions reported by two research groups (13Seo D. Tomioka A. Kusumoto N. Kamo M. Enami I. Sakurai H. Biochim. Biophys. Acta. 2001; 1503: 377-384Crossref PubMed Scopus (24) Google Scholar, 14Kjær B. Scheller H.V. Photosynth. Res. 1996; 47: 33-39Crossref PubMed Google Scholar). Membranes free from chlorosomes were prepared as previously described (10Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1998; 37: 12293-12300Crossref PubMed Scopus (33) Google Scholar). The C. tepidum ferredoxin was a generous gift of Dr. D. Seo and Prof. H. Sakurai (Waseda University) (13Seo D. Tomioka A. Kusumoto N. Kamo M. Enami I. Sakurai H. Biochim. Biophys. Acta. 2001; 1503: 377-384Crossref PubMed Scopus (24) Google Scholar). This ferredoxin was named FdC in their previous study (13Seo D. Tomioka A. Kusumoto N. Kamo M. Enami I. Sakurai H. Biochim. Biophys. Acta. 2001; 1503: 377-384Crossref PubMed Scopus (24) Google Scholar). The ferredoxin and FNR from Spirulina sp. were purified by Cui et al. (29Cui J.Y. Wakabayashi S. Wada K. Fukuyama K. Matsubara H. J. Biochem. (Tokyo). 1989; 105: 390-394Crossref PubMed Scopus (3) Google Scholar). Reduction of NADP+ was measured by monitoring absorption changes at 340 nm with a UV-3101PC spectrophotometer (Shimadzu) and a combination of appropriate filters. The samples were illuminated by actinic light with wavelengths longer than 490 nm. The photomultiplier was protected by a 340-nm interference filter with a bandwidth of 10 nm. Construction of the PscD-less Mutant—The pscD gene was inactivated by insertion of the aadA streptomycin/spectinomycin resistance cassette (Fig. 1) as described in Ref. 21Frigaard N.-U. Bryant D.A. Appl. Environ. Microbiol. 2001; 67: 2538-2544Crossref PubMed Scopus (78) Google Scholar. Southern hybridization analysis verified the insertion of a 2.0-kb aadA marker into the genome of the mutant (Fig. 2). A pscD probe hybridized with a 2.2-kb fragment in the wild type and with a 4.2-kb fragment in the mutant. In addition, an aadA probe hybridized with a 4.2-kb fragment in the mutant but not with the wild-type genomic DNA. PCR analysis using pscdF and pscdR primers (Fig. 1) also exhibited a 2.0 kb longer fragment in the mutant than in the wild type (data not shown). These results indicated that the aadA cassette was introduced correctly into the targeted pscD gene. The RC complexes were isolated from the mutant and wild-type strains according to the method described previously (27Oh-oka H. Iwaki M. Itoh S. Biochemistry. 1997; 36: 9267-9272Crossref PubMed Scopus (45) Google Scholar). The SDS-PAGE analysis (Fig. 3) clearly indicated the absence of PscD in the mutant strain. It is characteristic that no other subunits appeared to be affected prominently in their contents due to the deletion of PscD, suggesting that PscD would be loosely associated with and/or functionally independent of other subunits within the Chlorobium RC complex. Growth of the ΔpscD Strain—Growth rates were measured under high and low light conditions (Fig. 4). At a high light intensity of 164 μmol photons/m2/h, both the wild-type and mutant cells showed the same rates of photosynthetic growth. On the other hand, at a low light intensity of 2.5 μmol photons/m2/h, the growth rate of the ΔpscD mutant strain was almost the same as that of the wild-type strain until their culture turbidity, measured at 660 nm (OD660), reached 0.5. However, the former growth rate declined to about one-third beyond OD660 = 0.5, whereas the latter continued to grow at the same rate. A possible explanation of this rate transition during growth is that the light intensity penetrating into the interior portion of the culture would decrease far below 2.5 μmol photons/m2/h after the culture finally became turbid. Therefore, these results suggested that the ΔpscD mutant strain was incapable of collecting extremely low light energy at full efficiency, probably due to some damage in its antenna system. Another possibility might be that a subtle difference in some metabolic activity might be amplified during their growth, resulting in the delayed growth of the ΔpscD mutant strain compared with the normal one of the wild-type strain. Absorption and Fluorescence Spectra at Room Temperature—Absorption spectra of the wild-type and mutant cells are shown in Fig. 5A. They showed similar absorption spectra with high amounts of chlorosome BChl c pigments, although the absorption peak of chlorosome in the mutant cells was blue-shifted only by 3–4 nm. The contents of BChl c and BChl a did not show a significant change in the wild-type and mutant cells. Steady-state fluorescence emission spectra were measured in the wild-type and mutant cells at room temperature (Fig. 5B). Both types of cells exhibited emission peaks at 768 and 807 nm, which were derived from BChl c and BChl a, respectively. The spectrum with the high 807-nm BChl a peak in Fig. 5B was somewhat different from that reported in the isolated chlorosomes, which gave higher BChl c fluorescence compared with that of BChl a in the baseplates (19Blankenship R.E. Olson J.M. Miller M. Blankenship R.E. Madigan M.T. Bauer C.E. Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands1995: 399-435Google Scholar). The major portion of the 807-nm BChl a peak was thus ascribable to the fluorescence derived from BChl a in FMO protein instead of the baseplate BChl a in chlorosomes. On the other hand, the 768-nm peak was ascribable to that derived from BChl c aggregates in chlorosomes. In the ΔpscD cells, the emission peaks of both BChl c and a were larger than those in the wild type when measured at the same turbidity of OD750 = 0.5. The ratio of the 768- to 807-nm fluorescence intensity was also larger in the ΔpscD strain. These results implied that the light energy absorbed by chlorosomes and FMO proteins was transferred to P840 less efficiently in the mutant. Time-resolved Analysis of Fluorescence Spectra at 77 K—The time-resolved fluorescence spectra were measured in the picosecond time scale at 77 K in the presence of excess dithionite to determine the efficiency of the energy transfer from chlorosomes to the RC. Fig. 6A shows a two-dimensional (wavelength-time) image of the fluorescence decay in the wild-type cells of C. tepidum. The emission peaks detected at 785 and 825–829 nm (points a–d in Fig. 6A) at 77 K were ascribable to those of BChl c in chlorosomes and BChl a in the baseplate and FMO protein, respectively. The fluorescence peak in FMO protein is known to shift to the longer-wavelength side upon cooling to 77 K; therefore, the 829-nm peak was ascribable to the emission from FMO protein (19Blankenship R.E. Olson J.M. Miller M. Blankenship R.E. Madigan M.T. Bauer C.E. Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands1995: 399-435Google Scholar). After the selective excitation of BChl c in chlorosomes with a 645-nm laser flash of 50-ps width, the BChl c emission band at 785 nm rose first (point a in Fig. 6A), and then the BChl a one at 825 nm increased, indicating the energy transfer to the baseplate, as shown by point b (the fastest energy migration inside BChl c aggregates was not resolved with the present experimental time resolution). Fig. 6A further shows that the BChl a peak at 825 nm then shifted to 829 nm within 100–300 ps (point c), showing the energy transfer to the longer wavelength BChl a molecules in FMO protein. The fluorescence from FMO protein decayed slowly, presumably because of its equilibration with the slow energy dissipation in the RC (point d). These measurements were performed in the presence of excess dithionite, whose condition suppressed the fluorescence quenching by quinones in chlorosomes (30Wang J. Brune D.C. Blankenship R.E. Biochim. Biophys. Acta. 1990; 1015: 457-463Crossref PubMed Scopus (65) Google Scholar) and allowed only a primary electron transfer process, such as the reversible reduction of a primary acceptor A0 by P840 in the RC. The time-resolved spectra in Fig. 6B, calculated from slices along the horizontal wavelength axis of the two-dimensional image in Fig. 6A, clearly indicate the energy transfer processes mentioned above. The decay kinetics of the fluorescence at 785 and 830 nm were also calculated from slices along the vertical axis of the two-dimensional image in Fig. 6A (see Fig. 6C). The kinetics exhibited the fast decay of the BChl c fluorescence and the slow rise and decay of the BChl a fluorescence. All of these results indicate the fast energy transfer from BChl c aggregates to the RC via BChl a molecules in chlorosome baseplate and FMO protein, as has been reported elsewhere (19Blankenship R.E. Olson J.M. Miller M. Blankenship R.E. Madigan M.T. Bauer C.E. Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands1995: 399-435Google Scholar). On the other hand, in the ΔpscD cells, the energy transfer process occurred in a similar sequence, but both the fluorescence from BChl c and BChl a gave slower decay kinetics than those in the wild-type cells (Fig. 6D). After the fast rise of the fluorescence at 785 nm (point a), the BChl c fl
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