The Pennsylvania State University

The Graduate School

Eberly College of Science

ORGANIZATION AND FUNCTION OF CHLOROSOME PROTEINS

IN THE GREEN SULFUR BACTERIUM CHLOROBIUM TEPIDUM

A Thesis in

Biochemistry, Microbiology and Molecular Biology

by

Hui Li

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy May 2006

ii

The thesis of Hui Li has been reviewed and approved* by the following:

Donald A. Bryant Ernest C. Pollard Professor of Biotechnology Professor of Biochemistry and Molecular Biology Thesis Advisor Chair of Committee

John H. Golbeck Professor of Biochemistry and Biophysics Professor of Chemistry

Teh-hui Kao Professor of Biochemistry and Molecular Biology

Juliette T. J. Lecomte Associate Professor of Chemistry

Robert A. Schlegel Professor of Biochemistry and Molecular Biology Head of the Department of Biochemistry and Molecular Biology

* Signatures are on file in the Graduate School.

iii ABSTRACT

Chlorosomes are the light-harvesting antennae of the green sulfur bacteria, which usually live in extremely light-limited environments. Chlorosomes are sac-like structures with highly aggregated bacteriochlorophyll (BChl) c aggregates surrounded by a galactolipid/protein monolayer envelope. In the chlorosomes of Chlorobium tepidum, ten kinds of proteins (CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ, and CsmX) are located on the monolayer envelope membrane. Cross-linking experiments were performed to detect the relative locations and interactions among the chlorosome envelope proteins. The cross-linking reagent EDC (1-ethyl-3-(3- dimethylaminopropyl) carbodiimide) was used to cross-link the proteins of wild-type chlorosomes and chlorosomes from mutants lacking a single chlorosome protein, and the products were separated by SDS-PAGE and detected with antibodies against specific chlorosome proteins. CsmA forms dimers, trimers, and other multimers up to octamers, and is found in the baseplate region where it also interacts with the Fenna-Matthews-Olson (FMO) protein. The pre-CsmA, CsmB and CsmF proteins, which interact with CsmA to form heterodimers, might be located on the edge of the CsmA baseplate. CsmC forms homomultimers and it might be located on the opposite side from the baseplate facing the cytoplasm. Iron-sulfur chlorosome proteins CsmI and CsmJ form heterodimers, and both interact with CsmB. A model of the protein organization of the chlorosome membrane is proposed based on the cross-linking information. Three chlorosome proteins, CsmI, CsmJ, and CsmX, have strong sequence similarity in the amino-terminal domains to [2Fe-2S] ferredoxins of the adrenodoxin/putidaredoxin subfamily. The roles of the three iron-sulfur proteins were tested in single, double and triple knock-out mutants in all combinations. The mutant strains lacking the iron-sulfur chlorosome proteins grew at similar rates as the wild type under standard conditions. They were much more sensitive to oxygen than the wild-type cells as demonstrated by cell viability test after oxygen exposure. Fluorescence quenching and restoration experiments in chlorosomes and cells of the mutants suggest that CsmI and CsmJ are the most likely candidates for transferring electrons to and from the quencher within the chlorosome (most probably chlorobiumquinone) when the

iv environmental oxygen concentration is changed. CsmX, given its low concentration (~5%) compared to CsmI and CsmJ, makes little contribution to the quenching and restoration of fluorescence. The other chlorosome proteins can be divided into three groups according to their amino acid sequence similarity: CsmA/CsmE, CsmB/CsmF and CsmC/CsmD. CsmH contains two structural domains related in sequence to CsmB/CsmF and CsmC/CsmD. The functions of the chlorosome proteins were tested in double and triple mutants lacking members of the CsmB/CsmF or CsmC/CsmD motif family. The mutants exhibited apparent growth defects under limiting light intensities and contained significantly reduced amounts of cellular BChl c and/or carotenoids as indicated by absorption spectroscopy and HPLC analyses. Chlorosomes of the mutants also contained reduced amounts of BChl c and/or carotenoids, and exhibited significant differences in chlorosome size, shape, and absorption properties. These phenotypic effects strongly suggest that chlorosome proteins play roles in pigment incorporation into the chlorosomes, and that inactivation of chlorosome proteins probably inhibits pigment biosynthesis as a feedback effect. Recombinant CsmH, which contains two structural motifs specific to chlorosome proteins, was overexpressed in E. coli and purified by affinity chromatography for X-ray crystallographic analysis. Recombinant CsmA was also overexpressed in E. coli and purified from inclusion bodies. Binding analysis were performed with recombinant CsmA and Roseobacter-extracted BChl a. Evidence for in vitro binding was obtained on the basis of shifts of the BChl a absorption maximum observed by absorption spectroscopy.

v TABLE OF CONTENTS

LIST OF FIGURES …………………………………………………………….. ix LIST OF TABLES ……………………………………………………………… xii LIST OF ABBREVIATIONS ………………………………………………...... xiii ACKNOWLEDGEMENTS ……………………………………………………. xv

Chapter 1 Green Sulfur Bacteria and the Photosynthetic Apparatus ………. 1 1.1 Green sulfur bacteria …………………………………………………………. 2 1.1.1 Natural habitats and morphology …………………………………... 2 1.1.2 Metabolic characterization ………………………………………….3 1.1.3 Phylogeny properties ………………………………………………..4 1.1.4 Chlorobium tepidum … ……………………………………………..5 1.2 The photosynthetic apparatus of green sulfur bacteria ………………………..6 1.2.1 Pigment contents and cellular localization ………………………….6 1.2.2 The chlorosome …………………………………………………….. 8 1.2.2.1 The bacteriochlorophyll c aggregates ……………………. 8 1.2.2.2 The chlorosome proteins …………………………………. 12 1.2.3 Fenna-Matthews-Olson protein ……………………………………. 13 1.2.4 The reaction center …………………………………………………. 15 1.2.5 Energy transfer pathway and kinetics …………………………….... 17 1.2.6 Electron transfer and NAD(P)+ reduction ………………………….. 19

Chapter 2 Molecular Contacts for Chlorosome Envelope Proteins Revealed by Cross-linking Studies with Chlorosomes from Chlorobium tepidum …….. 33 2.1 Abstract ………………………………………………………………………. 34 2.2 Introduction …………………………………………………………………... 35 2.3 Materials and methods ……………………………………………………….. 38 2.3.1 Chlorobium tepidum strain and growth conditions ……………….... 38 2.3.2 Isolation of light harvesting antenna ……………………………….. 39 2.3.3 Cross-linking of chlorosome proteins …………………………….... 40

vi 2.3.4 SDS-PAGE and immunoblotting analysis …………………………. 41 2.4 Results ………………………………………………………………………... 43 2.4.1 Cross-linking of chlorosome proteins ……………………………… 43 2.4.2 Organization of CsmA ……………………………………………... 44 2.4.3 Organization of CsmC and CsmD …………………………………. 47 2.4.4 Interaction between CsmI/CsmJ and CsmB ……………………….. 48 2.4.5 Interactions for other chlorosome proteins ……………………….... 49 2.5 Discussion ……………………………………………………………………. 50

Chapter 3 [2Fe-2S] Proteins in the Chlorosome: Construction and Characterization of Mutants Lacking CsmI, CsmJ and CsmX in the Chlorosome Envelope of Chlorobium tepidum ………………………………... 70 3.1 Abstract ………………………………………………………………………. 71 3.2 Introduction …………………………………………………………………... 72 3.3 Materials and methods ……………………………………………………….. 75 3.3.1 Molecular manipulation in Escherichia coli…………………. ……... 75 3.3.2 Mutant construction and confirmation in Chlorobium tepidum …… 77 3.3.3 Chlorosome isolation and SDS-PAGE …………………………...... 79 3.3.4 Pigment and quinone analysis ……………………………………… 79 3.3.5 Fluorescence spectroscopy and cell viability test ………………….. 80 3.3.6 Reverse transcription PCR …………………………………………. 81 3.4 Results ………………………………………………………………………... 82 3.4.1 Construction and verification of mutants lacking CsmI, CsmJ and CsmX …………………………………………………………………….. 82 3.4.2 Pigment and quinone contents ……………………………………... 83 3.4.3 Fluorescence quenching and restoration …………………………… 84 3.4.4 Growth rates and viability after aerobic exposure …………………. 87 3.4.5 mRNA and protein level of CsmI and CsmJ in the csmX mutant ….. 89 3.5 Discussion ……………………………………………………………………. 90

vii Chapter 4 Chlorosome Proteins and Chlorosome Assembly: Construction and Characterization of Mutants Lacking CsmB/F or CsmC/D Motifs in the Chlorosome Envelope of Chlorobium tepidum ………………………………... 108 4.1 Abstract ………………………………………………………………………. 109 4.2 Introduction …………………………………………………………………... 111 4.3 Materials and methods ……………………………………………………….. 115 4.3.1 Plasmids construction in Escherichia coli …………………………. 115 4.3.2 Growth condition and mutant construction and confirmation in Chlorobium tepidum ………………………………………………………116 4.3.3 Chlorosome isolation ………………………………………………. 117 4.3.4 Analysis of protein contents ………………………………………... 117 4.3.5 Spectroscopy and pigment content determination ………………..... 118 4.3.6 Electron microscopy ……………………………………………….. 119 4.4 Results ………………………………………………………………………... 121 4.4.1 Construction of Chlorobium tepidum mutants ……………………... 121 4.4.2 Growth defects of the mutated cells ………………………………... 122 4.4.3 Cellular absorption profile and pigment contents ………………….. 123 4.4.4 Chlorosome isolation and protein component analysis ……………. 124 4.4.5 Absorption profile and pigment contents of the chlorosomes ……... 127 4.4.6 Electron microscopy of chlorosomes …...... 128 4.4.7 Chlorosome protein composition of mutants lacking the BChl c and carotenoids biosynthesis enzymes ……………………………………….. 129 4.5 Discussion ……………………………………………………………………. 131

Chapter 5 Purification and Characterization of Recombinant CsmH, and Pigment Binding Analysis of Recombinant CsmA …………………………… 153 5.1 Abstract ………………………………………………………………………. 154 5.2 Introduction …………………………………………………………………... 155 5.3 Materials and methods ……………………………………………………….. 157 5.3.1 Escherichia coli strains and growth conditions ……………………. 157 5.3.2 Construction of the expression vectors …………………………….. 157

viii 5.3.3 Overproduciton and purification of the recombinant proteins ……... 158 5.3.4 SDS-PAGE, protein staining and immunoblotting analysis ……….. 159 5.3.5 Characterization of recombinant CsmH …………………………..... 160 5.3.6 Pigment binding analysis of recombinant CsmA …………………... 161 5.4 Results ………………………………………………………………………... 162 5.4.1 Overproduction and purification of the recombinant proteins ……... 162 5.4.2 Characterization of recombinant CsmH …………………..……...... 163 5.4.3 Pigment binding analysis of recombinant CsmA ……………..……. 165 5.5 Discussion ……………………………………………………………………. 167

Chapter 6 Summary of Organization and Function of Chlorosome Proteins in Chlorobium tepidum ………………………………………………………… 179

Bibliography …………………………………………………………………….. 186

ix LIST OF FIGURES

Fig. 1-1. Sulfide and thiosulfate oxidation pathways of Chlorobium. ……………21

Fig. 1-2. The reductive tricarboxylic acid cycle of Chlorobium. ………………... 22

Fig. 1-3. Phylogenetic tree of green sulfur bacteria based on 16S rDNA sequences. ……………………………………………………………………………………... 23

Fig. 1-4. Circular representation of the C. tepidum genome. ……………………. 24

Fig. 1-5. Simplified chlorosome model showing interactions between the reaction center, the FMO protein and the chlorosome. …………………………... 25

Fig. 1-6. Structure of various bacteriochlorophylls (BChls). ……………………. 26

Fig. 1-7. Model of the chlorosome interior structure with bilayer tubes as the building block. …………………………………………………………………… 27

Fig. 1-8. Model of the chlorosome interior structure with BChl c lamellae as the building block. …………………………………………………………………… 28

Fig. 1-9. Structure of the Fenna-Matthews-Olson protein from Chlorobium tepidum. ………………………………………………………………………….. 29

Fig. 1-10. Structural organization of the Photosystem I reaction center from Synechococcus elongatus, and proposed organization of the reaction center in Chlorobium limicola. …………………………………………………………….. 30

Fig. 1-11. Redox potentials and rates of electron transfer in the P840-reaction center. ……………………………………………………………………………..32

Fig. 2-1. Wild type chlorosomes cross-linked with EDC for various time ranges and at various temperatures. ……………………………………………………... 59

Fig. 2-2. Wild type chlorosomes cross-linked at room temperature for 5 min with glutaraldehyde. …………………………………………………………………… 60

Fig. 2-3. Wild type chlorosomes cross-linked for various time ranges at room temperature. ……………………………………………………………………… 61

Fig. 2-4. SDS-treated chlorosomes cross-linked for various time ranges at room temperature. ……………………………………………………………………… 62

x Fig. 2-5. Carotenosomes cross-linked for various time ranges at room temperature. ……………………………………………………………………………………... 63

Fig. 2-6. Cross-linked chlorosomes detected by antibodies against CsmA, CsmF and FMO protein. ………………………………………………………………… 64

Fig. 2-7. Cross-linked chlorosomes detected by antibodies against CsmC. ……... 65

Fig. 2-8. Cross-linked chlorosomes detected by antibodies against CsmD. ……...66

Fig. 2-9. Cross-linked chlorosomes detected by antibodies against CsmI and CsmJ. ……………………………………………………………………………... 67

Fig. 2-10. Cross-linked chlorosomes detected by antibodies against CsmH. …….68

Fig. 2-11. Model of protein organization on the chlorosome envelope. …………. 69

Fig. 3-1. Restriction maps showing the structure of the gene inactivation constructs. ………………………………………………………………………... 100

Fig. 3-2. Protein composition analysis of chlorosomes isolated from the wild-type strain and the single, double and triple mutants lacking CsmI, CsmJ, and CsmX. .. ……………………………………………………………………………………... 101

Fig. 3-3. The quenching of fluorescence emission in isolated chlorosomes. ……. 102

Fig. 3-4. The restoration of fluorescence emission in isolated chlorosomes. ……. 103

Fig. 3-5. The restoration of fluorescence emission in whole cells of wild type and the csmI csmJ csmX mutant. ……………………………………………………... 105

Fig. 3-6. Model of electron transfer pathways in cellular fluorescence restoration. ……………………………………………………………………………………... 106

Fig. 3-7. RT-PCP and immunoblotting to detect mRNA and protein levels of CsmI/CsmJ in the csmX mutant. …………………………………………………. 107

Fig. 4-1. Restriction maps showing the construction and structure of the gene inactivation plasmids. ……………………………………………………………. 141

Fig. 4-2. Absorption spectra of whole cells of the wild type and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. ……………………… 142

Fig. 4-3. Chlorosome isolation from the csmB csmF and csmC csmD csmH mutants. …………………………………………………………………………... 143

xi Fig. 4-4. Protein composition analysis of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. .. ……………………………………………………………………………………... 145

Fig. 4-5. Absorption spectra and fluorescence spectra of chlorosomes from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. …………………………………………………...... 146

Fig. 4-6. Transmission electron micrographs of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. ………………………………………………………………. 148

Fig. 4-7. Protein composition analysis of chlorosomes from mutants lacking the BChl c and carotenoid biosynthesis enzymes. …………………………………… 150

Fig. 4-8. Correlation between the growth rates under limited light intensities and the cellular BChl c contents. ……………………………………………………... 152

Fig. 5-1. The deduced amino acid sequences of encoded proteins from the expression vectors. ……………………………………………………………….. 171

Fig. 5-2. Overproduction and purification of recombinant Chlorobium tepidum CsmH and CsmA in E. coli. ……………………………………………………… 173

Fig. 5-3. Light scattering of the N-terminal His-tagged CsmH. …………………. 175

Fig. 5-4. Gel filtration chromatography of N-terminal His-tagged CsmH and protein markers on Sephacryl-200 column. ……………………………………… 176

Fig. 5-5. Purified N-terminal His-tagged CsmH cross-linked with 5 mM EDC and detected by immunoblotting with the anti-CsmH antibodies. …………………… 177

Fig. 5-6. Binding analysis of extracted BChl a and the recombinant CsmA. ……. 178

xii LIST OF TABLES

Table 2-1. Interactions and locations of chlorosome proteins. ………………….. 57

Table 2-2. Distribution of chlorosome proteins in the genomes of all sequenced green sulfur bacteria other than Chlorobium tepidum. …………………………... 58

Table 3-1. Pigment and quinone contents of chlorosomes isolated from the wild-type strain and the mutants lacking CsmI, CsmJ and CsmX. ………………. 97

Table 3-2. Chlorosome fluorescence restoration with 12.5 mM and 25 mM dithionite shown by increasing rates and increasing rate percentage compared to the wild type. ……………………………………………………………………... 98

Table 3-3. Growth rates of the wild-type strain and the mutants lacking CsmI, CsmJ and CsmX under different light intensities. ……………………………….. 99

Table 4-1. Growth rates of the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif under different light intensities. … 136

Table 4-2. Characterization of cells of the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. ……………………… 137

Table 4-3. Characterization of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. …………. 138

Table 4-4. Genes encoding enzymes involved in BChl c and carotenoid biosynthesis and pigment compositions in the mutant strains with the corresponding genes inactivated. ………………………………………………… 139

Table 4-5. Lengths, widths and length-to-width ratios of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif, observed by transmission electron microscopy. ……………. 140

xiii LIST OF ABBREVIATIONS

ATP adenosine triphosphate BChl bacteriochlorophyll bp basepairs Chl chlorophyll CL Chlorobium Liquid (Medium) CP Chlorobium Plate (Medium) Cyt cytochrome Da Dalton dd double-distilled dATP deoxyadenosine triphosphate dCTP deoxycytosine triphosphate dGTP deoxyguanosine triphosphate dTTP deoxythymidine triphosphate DNA deoxyribonucleic acid DTT dithiothreitol EDC 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride EDTA ethylenediamine tetraacetic acid Fd ferredoxin Fe-S iron-sulfur FMO protein Fenna-Matthews-Olson protein h hour IgG Immunoglobulin G IPTG isopropyl-β-D-thiogalactopyranoside kbp kilobasepairs LB Luria-Bertani MES 2-[N-morpholino]ethane sulfonic acid min minutes MOPS 3-[N-morpholino]propanesulfonic acid

xiv NADH nicotinamide adenine dinucleotide NADP nicotinamide dinucleotide phosphate nt nucleotide OD optical density ORF open reading frame PAGE polyacrylamide gel electrophoresis PCR polymerase chain reaction pI isoelectric point PMSF phenylmethylsulfonyl fluoride PSI Photosystem I PSII Photosystem II psi pounds per square inch RNA ribonucleic acid s seconds SDS sodium dodecyl sulfate TBS Tris-buffered saline Tricine N-tris(hydroxymethyl)methylglycine Tris tris(hydroxymethyl)amino-methane

Tm melting (annealing) temperature WT wild type × g gravitational constant

xv ACKNOWLEDGEMENTS

I would like to thank my advisor, Dr. Bryant, for all of his assistance, support and encouragement throughout my graduate study and research. I would like to thank Dr. Niels-Ulrik Frigaard in Bryant’s lab and Dr. Thomas W. Johnson in Dr. Golbeck’s lab for their contributions to the characterization of chlorosome protein mutants. My acknowledgement also goes to my committee members, Dr. John Golbeck, Dr. James Ferry, Dr. Teh-hui Kao and Dr. Juliette Lecomte for their guidance and support. I would also like to thank Dr. Murakami for his assistance of the CsmH crystal screen, and Dr. Doba Jackson in Dr. Song Tan’s lab and Dr. Hemant Yennawar in the crystallography facility for their assistance of the light scattering experiment. I also thank the past and present members of the Bryant’s lab in no particular order for their technical help, discussion and friendship: Dr. Niels-Ulrik Frigaard, Dr. Elena V. Vassilieva, Dr. Gaozhong Shen, Dr. Tao Li, Dr. Yumiko Sakuragi, Ramakrishnan Balasubramanian, Joel E. Graham, Aline Gomez Maqueo Chew, Julia Maresca, Zhao Jin and Yingxian Wu. Finally, I would like to thank my family in China, and my husband Hao Wang, for their support throughout my studies.

1

Chapter 1

Green Sulfur Bacteria and the Photosynthetic Apparatus

2 1.1 Green sulfur bacteria

Green sulfur bacteria (Chlorobiaceae) are a group of obligate photoautotrophs that utilize light energy to synthesize organic compounds by photosynthesis. Distinguished from the other photosynthetic microorganisms (cyanobacteria, green nonsulfur bacteria, heliobacteria, purple bacteria), green sulfur bacteria survive under strictly anaerobic and sulfide-rich environments with limited light intensities, and exhibit specialized physical and biochemical properties.

1.1.1 Natural habitats and morphology

Since the first observation at the end of nineteen century, the presence of green sulfur bacteria has been repeatedly reported in aquatic environments all over the world (Pfennig and Trüper, 1992). Without exception, these environments possess three distinguishing characteristics: absence of oxygen, presence of light and presence of reduced sulfur compounds. Habitats of green sulfur bacteria are detected on the top millimeters of aquatic sediments or sandy sediments close to the shore, surrounded by various types of proteobacteria. They are also found at the top of the anoxic zone in freshwater or marine environments, where the light penetration is severely hampered due to the depth and the light intensity is limited to a few μmol photons m-2 s-1 (van Gemerden and Mas, 1995). A layer of sulfate-reducing proteobacteria is usually found underlying and serves as the source of reduced sulfur compounds (Dyer, 2003). Cells of green sulfur bacteria exhibit a variety of shapes --- spherical, ovoid, rod and spiral. All species lack flagella, and all species of Pelodictyon and Chloroherpeton have gas vesicles (Imhoff, 1995). Elemental sulfur produced by metabolism in these organisms may remain attached to the outside of the cells, appearing as tiny white-yellow globules (Dyer, 2003). Species of Prosthecochloris are characterized by the long stemlike appendages, named prosthecae, which modify the cells to a star-like morphology (Guyoneaud, 2001). Some freshwater green sulfur bacteria form spindle-shaped consortia with other bacteria, and these consortia have been given the genus names “Chlorochromatium” and “Pelochromatium”. In the consortia, green sulfur bacteria are

3 typically arranged in rows around a colorless bacterium in the center, which belongs to β- proteobacteria and carries a single polar flagellum (Overmann and Schubert, 2002). Signal exchange occurs between the two bacteria and the intact consortia exhibit photophilous response with the epibionts sensing the light and the central bacterium conferring motility (Overmann and Schubert, 2002).

1.1.2 Metabolic characterization

As anoxygenic photoautotrophs, green sulfur bacteria do not use light energy to oxidize water as an electron donor. Instead, they use inorganic sulfur compounds to provide electrons at a moderately low redox potential. All species except C. ferrooxidans in the phylum Chlorobi are able to oxidize hydrogen sulfide to elemental sulfur and sulfate, and some species have the additional ability to utilize thiosulfate (Brune, 1995). None of Chlorobiaceae is known to use sulfite as an electron donor probably because of the impermeability of the cell membrane (Brune, 1995). Two intermediates, elemental sulfur and sulfite, are produced in the process of sulfide oxidation. Elemental sulfur is occasionally exported and attached to the outside of the cells as sulfur globules (Madigan, 1988). The other intermediate, sulfite, is not observed in most species since its formation and consumption in the cytoplasm is too fast to escape to the external environment (Brune, 1995). The pathways of sulfide and thiosulfate oxidation are shown in Fig 1-1.

All species of green sulfur bacteria grow photolithotrophically with CO2 as the sole carbon source. Using reduced sulfur compounds or H2 as electron donors, two

molecules of CO2 are assimilated via reactions of the reductive tricarboxylic acid cycle into one molecule of acetyl-CoA (Evans et al., 1966; Fuchs et al., 1980 a, b; Ivanovsky et al., 1980). Two essential enzymes, α-ketoglutarate-ferredoxin oxidoreductase and pyruvate-ferredoxin oxidoreductase, make the key steps of the reversal TCA cycle possible. Acetyl-CoA produced by the cycle can be converted to carbohydrates and further to polysaccharide as carbonaceous reserve materials (Sirevåg, 1975). Other products and intermediates of the cycle (pyruvate, oxaloacetate, succinyl CoA and α- ketoglutarate) can be used in synthesis of fatty acids as well as amino acids. The reverse TCA cycle pathway is shown in Fig. 1-2.

4 Nitrogen fixation has been observed in all species of Chlorobium and multiple species of Pelodictyon, Prosthecochloris and Chloroherpeton (Bergstein et al., 1981; Heda and Madigan, 1988; Rodionov et al., 1986; Gibson et al., 1984). Chlorobium sp. cells assimilate ammonia mainly via the glutamine synthetase-glutamate synthase pathway (Wahlund and Madigan, 1993). Interestingly, the nitrogenase activity in cultures of Chlorobium species can be switched off by addition of ammonia and switched on again when ammonia is consumed (Wahlund and Madigan, 1993).

1.1.3 Phylogeny properties

The 16S rDNA sequences of green sulfur bacteria share a high degree of similarity, indicating that a close phylogenetic relatedness exists among all the strains. As shown in Fig. 1-3, five phylogenetic clusters can be distinguished in the green sulfur bacteria phylum: a truly marine branch with average G+C values between 52.2 and 53.5 mol%, a second cluster of saltwater strains with higher mol% G+C values, a third cluster comprising freshwater strains, a forth cluster including thermophilic strain Chlorobium tepidum ATCC 49652T and some strains of Chlorobium vibrioforme and Chlorobium limicola, and a fifth group containing Chloroherpeton thalassium (Figueras et al., 1997; Alexander et al., 2002). The phylogenetic tree was constructed based on 16S rDNA sequences and later confirmed by amino acid sequences of the Fenna-Matthews-Olson (FMO) protein, a characteristic protein from the photosynthesis apparatus in green sulfur bacteria (Figueras et al., 1997; Alexander et al., 2002). Green sulfur bacteria are sometimes mentioned together with green nonsulfur bacteria because both of them contain chlorosomes as light-harvesting antenna. However, the two families of green bacteria appear to be only very distantly related based on 16S rDNA analysis as well as physiological properties. The group of green nonsulfur bacteria branched early on the family tree soon after that of the ancient hyperthermophiles, and is distant from green sulfur bacteria (Gibson et al., 1985; Woese, 1987). Although green nonsulfur bacteria can produce chlorosomes and perform photosynthesis under anaerobic conditions, they act as heterotrophs when oxygen and organic molecules are available (Pierson and Castenholz, 1992). Green nonsulfur bacteria require organic molecules or

5

H2 instead of sulfur compounds as electron donors, utilize the unique 3-

hydroxypropionate pathway to fix CO2, and apparently cannot perform nitrogen fixation (Pierson and Castenholz, 1992; Holo, 1989; Heda and Madigan, 1988). All these properties clearly support the isolated phylogenetic position of green sulfur bacteria. To explain the presence of chlorosomes in two such widely separated groups, lateral transformation of the genes coding for chlorosome components has been suggested (Blankenship, 1992).

1.1.4 Chlorobium tepidum

Chlorobium tepidum was originally isolated from high-sulfide hot springs in New Zealand (Wahlund et al., 1991). When cultured in the laboratory, C. tepidum grows optimally at 47-48°C and pH 6.8-7.0 with a doubling time of about two hours. High cell yields were only obtained when thiosulfate was supplied as the major electron source, while some sulfide was also required (Wahlund et al., 1991). The whole genome sequence of C. tepidum has been determined, which represents the first genome sequence from the phylum Chlorobi (Fig. 1-4) (Eisen et al., 2002). The complete genome size is 2.15 Mb, with a G+C content of 56.5% and more than 2,000 predicted protein coding sequences (Eisen et al., 2002). The genes encoding specific enzymes of reverse TCA cycle (pyruvate synthase and α-ketoglutarate synthase) were identified (Atomi, 2002; Yoon et al., 2001). A homolog of the large subunit of RubisCO (ribulose-1,5-bis- phosphate carboxylase) is encoded although the RubisCO activity is not present in C. tepidum (Hanson and Tabita, 2001). Only one set of nitrogen fixation genes is present in the genome; they are closely related to those in Archaea and might have originated from lateral gene transfer (Eisen et al., 2002). The sulfur metabolism genes are localized in four gene clusters, and many of them have been duplicated (Eisen et al., 2002). The large number of sulfur oxidation genes supports the fact that C. tepidum is capable of using various sulfur compounds as electron sources, although the detailed metabolic pathway remains unclear.

6 1.2 The photosynthetic apparatus of green sulfur bacteria

The photosynthetic apparatus of phototrophic organisms includes three parts: the light- harvesting antenna, the reaction center and the electron transfer pathway. The light- harvesting antennae are arrays of pigment-protein complexes localized in or on the cytoplasmic surface of the cytoplasmic membrane. The reaction centers are situated in the cytoplasmic membrane and catalyze the photochemical events of photosynthesis. Light energy is gathered by light-harvesting antenna and delivered by Förster resonance transfer to the reaction centers via specialized microenvironments. The excitation energy causes charge separation on a pair of chlorophyll molecules (called the special pair) in the reaction center; and the excited electron is transferred via a few cofactors finally to soluble ferredoxins which can be used for production of ATP and reducing forces. In green sulfur bacteria, light energy is harvested by unique antennae known as chlorosomes, and transferred via the Fenna-Matthews-Olson protein layer to the Type I reaction centers (Fig. 1-5). Pigment molecules, especially bacteriochlorophylls, play critical role in the process of photosynthesis.

1.2.1 Pigment contents and cellular localization

The most widely studied green sulfur bacteria are a deep emerald-green color, with bacteriochlorophyll c or d and the carotenoid chlorobactene and OH-chlorobactene as light-harvesting pigments (Imhoff, 1995). Other brown-colored species contain bacteriochlorophyll e and the carotenoids isorenieratene and β-isorenieratene, showing a broad absorption between 480 and 550 nm (Gloe et al., 1975; Imhoff, 1995). The roles of the pigment molecules in the photosynthesis systems include several aspects: efficiently absorbing light energy, degrading excess energy to heat, stabilizing the structure of photosynthetic apparatus, transferring excitation energy to reaction centers, and generating electron transfer by charge separation. Bacteriochlorophyll (BChl) is the name given to the photosynthetic pigments belonging to the cyclic tetrapyrrole group in photosynthetic bacteria. BChls a, b and g are true bacteriochlorins (7,8,17,18-tetrahydro-porphyrins) with two rings, B and D, reduced

7 (Fig. 1-6 Panel A). BChl a has an acetyl group at C3, and is mostly esterified with phytol at the C17 propionyl group (Scheer, 1991). BChl b differs from BChl a by the presence of an ethylidene group at C8, and BChl g has a vinyl group at C3 in addition to a C8 ethylidene group (Scheer et al., 1974; Michalski et al., 1987). The Qy-absorption band of these BChls is 750-800 nm in organic solvents and even more red-shifted in situ. The Soret-band absorption occurs at wavelengths less than 400 nm. The primary donors in the reaction centers of photosynthetic bacteria are exclusively BChls a, b or g (Scheer, 2003). BChl a, the most widely distributed BChl, is also present in the core and peripheral light- harvesting antennas. BChl c, d, and e are chemically chlorins (17,18-dihydroporphyrin) just like chlorophylls, in spite of the name BChl (Fig. 1-6 Panel B). These BChls are the main constituents in the light-harvesting antenna of green bacteria, with absorption bands at the edges of the visible spectrum (400-500 nm and 650-680 nm) in organic solvents (Imhoff, 1995). They characteristically lack the 132-carboxymethyl group and have an α- hydroxyethyl group at C3 (Scheer, 2003). BChls c and e are also unique in the presence of a methyl group at C20 (Scheer, 2003). Each of the three BChls exhibits a remarkable structural diversity with various modifications at C8 (isobutyl, n-propyl or ethyl) and C12 (ethyl or methyl), different esterified alcohols at C173 (farnesyl or stearol) and variable chirality at C31 (R- or S-configuration) (Caple et al., 1978; Smith and Simpson, 1986; Tamiaki et al., 1994). The variations are so great in number that they account for more than 50% of the currently known chlorophyll structures (Scheer, 2003). Various kinds of carotenoids have been identified in green sulfur bacteria, including carotenes and their oxidized xanthophyll derivatives. The basic hydrocarbon structure of carotenoids is a chain of 32 carbon atoms, bearing eight methyl side-chains, which is formed by two C20-units (originally geranyl-geraniol) joined tail-to-tail. Common modifications involve cyclization at one or both ends, desaturation of the carbon-carbon bonds, rearrangements of the double bonds, and even shortening and

extension of the C40-skeleton (Scheer, 2003). Most carotenoids of green sulfur bacteria are located in chlorosomes, accompanied by BChl, while carotenoid glucoside esters are mainly located in the membranes and reaction centers (Takaichi, 1999). In addition to their contribution to light harvesting (absorption band 450-550 nm), the most critical role

8 of carotenoids is to protect photosynthesis against direct and indirect damage by light, such as quenching BChl triplet states and scavenging toxic radicals.

1.2.2 The chlorosome

Photosynthetic organisms have evolved multiple light-harvesting systems to collect light energy and transmit that energy to photosynthetic reaction centers, such as phycobilisomes of cyanobacteria and red algae, peridinin-chlorophyll proteins of dinoflagellates, and caroteno-chlorophyll a/b binding proteins (CAB/LHC) of higher plants (Blankenship, 2002). The green sulfur bacteria also have their solution to the light- harvesting problem: these organisms harbor unique organelles known as chlorosomes to collect light energy at extraordinarily low light intensities. Chlorosomes are flattened, ellipsoid-shaped structures containing highly aggregated BChl c rods, small amounts of BChl a, carotenoids and quinones enclosed by protein-stabilized monolayer galactolipid membranes (Blankenship and Matsuura, 2003). The dimensions of chlorosomes are not strictly defined; they are usually 100 to 200 nm long, 70 to 90 nm wide and 30 to 40 nm thick, with variation depending on the light intensity and other growth conditions (Staehelin et al., 1980; Oelze and Golecki, 1995; Montaño et al., 2003a). Unlike the other light-harvesting antennae, in which chromophores are held and oriented by proteins, BChl c molecules in chlorosomes are organized by intermolecular hydrophobic interactions, and the protein to chlorophyll mass ratio is remarkably low (Vassilieva et al., 2000). Green nonsulfur bacteria can also develop chlorosomes and perform photosynthesis under anaerobic conditions, but their chlorosomes are much smaller and contain fewer kinds of proteins than those of green sulfur bacteria (Staehelin et al., 1978; Vassilieva et al., 2000).

1.2.2.1 The bacteriochlorophyll c aggregates

The membrane envelope of chlorosomes is a glycolipid monolayer about 10 Å thick that consists mainly of monogalactosyl diglyceride (Staehelin et al., 1978). The polar head groups of the galactolipids point to the chlorosome outer surface while the fatty acid tails

9 sticking toward the interior (Holo et al., 1985). Thus the interior of the chlorosomes is believed to be very hydrophobic. The main pigments in chlorosomes are BChl c, d or e depending on the species in which the chlorosomes occur. Earlier estimates suggested that roughly 10,000 molecules of BChl c, d or e are enclosed in each chlorosome (Olson, 1980; Olson, 1998). Recent measurements in chlorosomes of Chlorobium tepidum using fluorescence correlation spectroscopy indicated that approximately 215,000 BChl c molecules are contained per chlorosome (Montaño et al., 2003a; Frigaard and Bryant, 2004). The molecules of BChl c are not protein-bound but self-aggregate to create higher order structures. BChl c oligomers can be formed by π- π stacking interactions between the rings, coordination of Mg atoms by the C31-OH of the stacked neighbor and a hydrogen bond between the C31- OH and C131=O of the neighbor in the same chain (Rossum et al., 2001). One important

clue of BChl c self-aggregation in the chlorosomes is that the Qy absorption spectrum of chlorosome BChl c is red-shifted about 70 nm compared with the spectrum of monomeric BChl c in organic solvents. The red-shift of the absorption spectrum can also be produced in vitro by BChl c aggregates in non-polar solvents (Blankehship et al., 1995; Tamiaki, 1996; Olson, 1998) or pigment-lipid aggregates in aqueous phase with BChl c and galactolipids (Hirota et al., 1992; Miller et al., 1993). Moreover, SDS-extraction of chlorosome proteins has shown no effects on the spectral properties of the BChl c, indicating that the BChl c aggregates are self-organized instead of being bound to chlorosome proteins (Griebenow et al., 1989, 1991; Holzwarth et al., 1990; Bryant et al., 2002). On the other hand, the aggregated BChl c can be converted reversibly to the monomeric form by treating intact chlorosomes with 1-hexanol (Brune et al., 1987b; Matsuura and Olson, 1990; Zhu et al., 1995). The interior structure of chlorosomes continues to be a controversial topic. Because of their large size and unusual organization, crystallization of chlorosomes or aggregated BChl c has not been possible, and high-resolution structural information has not yet been obtained. Early freeze-fracture electron microscopy studies showed that chlorosomes are filled with 10 to 30 rod-shaped elements with a diameter of 10 nm (Staehelin et al., 1980). The rod elements are closely packed and oriented along the longest axis of the chlorosome (Staehelin et al., 1980). Chlorosome models based on rod

10 elements have been proposed over the years, either by in vitro aggregation tests of bacteriochlorophylls or by computer modeling studies (Matsuura et al., 1993; Nozawa et al., 1994; Holzwarth and Schaffner, 1994; Prokhorenko et al., 2000). The Holzwarth models are generally considered to be the best ones by most researchers in this field. In the earliest model proposed by Holzwarth and Schffaner, the rod element is formed by a monolayer tubular sheet of aggregated BChl c with the farnesyl tails pointing outward (Holzwarth and Schaffner, 1994). However, this arrangement leaves a large amount of space inside the tube and is not in agreement with the electron microscopy observations that the central cavity is only about 3 nm in diameter (Cruden and Stanier, 1970). The model was later refined to a bilayer cylinder with the farnesyl chains of the outer layer extending outward and farnesyl chains of the inner layer filling the central cavity of the rod (Fig. 1-7) (Rossum et al., 2001). Solid-state nuclear magnetic resonance studies showed that two structurally different arrangements of aggregated BChl c exist in the chlorosome, named anti and syn stacks, with a molar ratio of about 6:4 (Rossum et al., 2001). Existence of the two stack configurations are also supported by solution nuclear magnetic resonance studies (Mizoguchi et al., 1998), linear dichroism and circular dichroism spectra (Matsuura et al., 1993; Somsen et al., 1996), and spectral changes during treatment in acidic buffer (Steensgaard et al., 1997). The syn and anti stacks are mirror images to each other with opposite sliding directions. The bilayer tube model with an outer layer of anti-aggregated BChl c and an inner layer of syn-aggregated BChl c has thus been proposed in keeping with the solid-state NMR results and the microscopy observations. Although the rod-element models have been widely accepted over the years, the high-resolution images by cryoelectron microscopy published recently provide evidence that suggest another model. Chlorosomes from Chlorobium tepidum were bedded in vitreous ice and imaged directly without further treatment or staining (Pšenčík et al., 2004). A striation pattern with a repeat dimension of about 20 Å was observed in nearly all chlorosomes, and extended parallel to the long axis of the chlorosome. X-ray scattering from chlorosomes exhibited a similar feature with ~ 20 Å spacing. The scattering curve contained a prominent peak corresponding to a spacing of 20.9 Å and a broad minor peak with a spacing of 4.5 Å, which was likely to reflect short-range order

11 such as stacking of chlorine rings. Notably, no peak was observed reflecting the rod-like elements, which should be ~100 Å in diameter. Pšenčík and his collaborators argued that the observed spacing of 20 Å is compatible with a lamellar model rather than a model with rod elements (Fig. 1-8) (Pšenčík et al., 2004). Firstly, rod elements built up by BChl c molecules would be unlikely to have a diameter of 20 Å, which requires very close packing and unrealistically high BChl densities. Secondly, the observed spacing perfectly matches the spacing of semicrystalline lateral arrays with antiparallel BChl dimers as the building unit. Pšenčík et al. (2004) suggest that the dimer chains are assembled together by hydrophobic interactions between the esterifying alcohol tails extending out of the plane, and the aggregates are highly likely to be bent or disordered by undulation. It could be possible that the 20 Å striation is difficult to detect in freeze-fracture electron microscopy and the rod-like elements observed by Staehelin et al. (1980) were generated from the undulating planes within the lamellae. Thus, the detailed interior structure of chlorosomes remains to be determined and obtaining high-resolution structural information might be the only way leading to a final conclusion about the BChl c aggregation in chlorosomes. In addition to the BChl c (d or e), chlorosomes from green sulfur bacteria contain small amounts of BChl a (1% w/w of BChl c), carotenoids (7% w/w of BChl c) and quinones (7% w/w of BChl c) (Blankenship and Matsuura, 2003). BChl a is mostly located in the baseplate region together with CsmA protein (Sakuragi et al., 1999; Bryant et al., 2002; Montaño et al., 2003b). The Qy absorption band of BChl a is red shifted from 770 nm of the monomeric form to 795 nm due to pigment-protein interactions. The location and organization of carotenoids and quinones have been unclear. Detergent treatment of chlorosomes demonstrated that quinones are inaccessible to SDS and that they are probably located in the interior of the chlorosomes (Frigaard et al., 1998). Some studies suggest that the carotenoids are closely associated with the BChl aggregates, whereas others indicate that they are assembled with the baseplate complex (Arellano et al., 2000). Recent studies with BChl c and carotenoids in aqueous buffer showed that the

BChl c Qy transition is further red-shifted in presence of carotenoids, indicating that carotenoids are incorporated into the BChl c aggregates and induce further aggregation (Klinger et al., 2004). The precise composition values of the pigments depend on the

12 species and are easily affected by variation in growth conditions and environments (Oelze and Golecki, 1995).

1.2.2.2 The chlorosome proteins

Although proteins are not involved in BChl c aggregation, they are still considered to be essential components in the biosynthesis and organization of the chlorosome. At least ten different polypeptides have been identified in highly purified chlorosomes from Chlorobium tepidum: CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ, and CsmX (Chung and Bryant, 1996 a, b; Vassilieva et al., 2002b). Subcellular localization studies proved that all ten chlorosome proteins copurified proportionally with BChl c, and that none of them were substantially detected in other subcellular fractions (Vassilieva et al., 2002b). The proteins are exposed at the surface of isolated chlorosomes to various extents, as indicated by protease susceptibility mapping and chlorosome agglutination experiments using antibodies (Chung and Bryant, 1996b; Vassilieva et al., 2002b). However, the functions of these proteins remain unclear except in the case of the CsmA protein, which has recently been shown to bind BChl a (Sakuragi et al., 1999; Bryant et al., 2002; Montaño et al., 2003b). CsmA is the most abundant chlorosome protein in Chlorobium tepidum and has a relatively small molecular weight of 6.2 kDa (Chung et al., 1994). The CsmA homolog in green nonsulfur bacterium Chloroflexus aurantiacus is about 5.7 kDa (Wechsler et al., 1985). Gold labeling electron microscopy demonstrated that CsmA in Chloroflexus aurantiacus is located in the lipid envelope of the chlorosome (Wullink et al., 1991). Recent results clearly indicate that the CsmA homologs bind to BChl a in the baseplate, which is located along the base of the chlorosome and serves as an intermediate in energy transfer to the reaction center. Experiments using proteolytic digestion and detergent treatment provided solid evidence that BChl a is associated with the CsmA protein in Chloroflexus aurantiacus (Sakuragi et al., 1999). Isolated baseplates from the chlorosomes of Chloroflexus aurantiacus was shown to contain BChl a, β-carotene and CsmA with a ratio of 1.0 CsmA/1.6 BChl a/4.2 β-carotene (Montaño et al., 2003b). Selective protein extraction by hexanol and SDS also demonstrated that CsmA can be

13 released along with BChl a from the chlorosomes of Chlorobium tepidum, and quantitative estimates showed that the complex probably has a one-to-one ratio of BChl a and CsmA (Bryant et al., 2002). The regions with sequences of G-H-W and I-N-R/Q-N- A-Y are highly conserved in the CsmA homologs (Wagner-Huber et al., 1988, 1990). The conserved histidine residue in the CsmA protein probably provides a suitable ligand for binding BChl a. In Chlorobium tepidum, the genes encoding CsmA and CsmC, the genes encoding CsmD and CsmE, the genes encoding CsmX and CsmJ are on the same transcription unit, while the genes of other chlorosome proteins are scattered in the genome. Sequence alignments for chlorosome proteins of Chlorobium tepidum indicate that CsmE is 49% identical to CsmA, and that the C-terminal motifs of CsmI, CsmJ and CsmX are distantly related to CsmA and CsmE. The N-terminal domains of CsmI, CsmJ, and CsmX have strong sequence similarity to adrenodoxin-type [2Fe-2S] ferredoxins, and both CsmI and CsmJ overexpressed in Escherichia coli were shown by EPR spectroscopy to contain iron-sulfur clusters (Vassilieva et al., 2001). The remaining five chlorosome proteins can be divided into two structural families: CsmB/F and CsmC/D. The C-terminus of CsmH is similar to CsmC/D, while its N-terminus is related to CsmB/F (Vassilieva et al., 2000). The sequence relationships between chlorosome proteins might be generated by gene duplication and divergence that occurred among a small number of protein types (Vassileva et al., 2002). Although CsmA is the only polypeptide with obviously conserved sequences between green sulfur bacteria and green nonsulfurs, the Csm proteins are highly conserved in Chlorobium species such as C. vibrioforme and C. phaeobacteroides (Vassilieva et al., 2000). None of the chlorosome proteins shares significant sequence similarity with other photosynthesis-related proteins. Little information has been obtained about the function and detailed location of chlorosome proteins other than CsmA.

1.2.3 Fenna-Matthews-Olson protein

The Fenna-Matthews-Olson (FMO) protein, also known as the bacteriochlorophyll a- binding protein, is found only in green sulfur bacteria (Daurat-Larroque et al., 1986;

14 Dracheva et al., 1992). It was first isolated and characterized by Olson et al. in 1966, and crystallized for structural analysis as the first chlorophyll-containing protein (Olson, 1966; Fenna and Matthews, 1975; Matthews et al., 1979; Matthews and Fenna, 1980). The structure of the FMO protein is currently available from two species, Prosthecochloris aestuarii 2K and Chlorobium tepidum (Tronrud et al., 1986, 1993; Li et al., 1997). The quaternary structure of the FMO protein is trimeric, consisting of three identical subunits related by a 3-fold axis of crystallographic symmetry (Fenna et al., 1974; Li et al., 1997). Each monomer is a 36 kDa polypeptide with 17 β-strands extending and curving into a “taco shell” structure (Fig. 1-9). The β-strands have an average length of 12 residues, and the front ones are approximately perpendicular to the ones at the back. Six α-helices, each containing about ten residues, are located in the open side of the taco shell, facing the center of the trimer. Salt bridges between arginine and aspartic acid, in addition to polar-polar interactions, contribute to holding the three subunits together (Li et al., 1997). Seven BChl a molecules are enclosed in the β-sheets of the monomer protein and are arranged into two clusters: BChl 1 and 2 are exposed to the surface while BChls 3, 4, 5, 6 and 7 are buried in the core. Each of the BChl a molecules is five-coordinate via a fifth ligand from a histidine residue (BChls 1, 3, 4, 6 and 7), backbond oxygen (BChl 5) or bound water molecule (BChl 2) (Li et al., 1997). There are no carotenoids associated with this complex. The FMO protein is localized between the chlorosome and the membrane-bound reaction center complex, providing a microenvironment for energy transfer. Linear dichroism spectroscopy showed that the trimeric FMO complex lies flat on the membrane, with the threefold symmetry axis perpendicular to the membrane surface (Melkozernov et al., 1998). Although Li et al. suggested that the bottom side of the trimeric complex is relatively more hydrophobic than the top view and might associate with the membrane, it is more likely that both the membrane-associated and chlorosome-associated surfaces are hydrophobic and that the hydrophobicity of the top and bottom sides doesn’t differ greatly (Li et al., 1997; Blankenship and Matsuura, 2003). Thus, the orientation of the FMO complex remains tentative. A portion of the FMO trimers is tightly associated with the reaction center and can be co-purified with the reaction center by gradient ultracentrifugation (Hager-Braun et al., 1995). Images obtained by scanning transmission

15 electron microscopy suggested that at least one FMO trimer is associated per reaction center complex and a second FMO trimer is likely to be accommodated to yield a symmetric complex (Rémigy et al., 1999). Low-temperature absorption spectroscopy indicated that two FMO trimers are contained in the FMO-reaction center complex of both Prosthecochloris aestuarii 2K and Chlorobium tepidum (Permentier et al., 2000). The FMO protein has been found to be required for the stabilization of the photoactive reaction centers (Hager-Braun et al., 1995). The essential role of the FMO protein is also confirmed by the fact that attempts to delete this protein have not been successful (Hu and Blankenship, 1999).

1.2.4 The reaction center

Photosynthetic reaction centers are pigment-protein complexes embedded in the cytoplasmic membrane. Light energy is used by the reaction centers to drive electron transport reactions leading to the production of proton gradient for ATP synthesis and/or reducing power. The reaction centers are classified as type I and type II based on their terminal electron acceptors (Blankenship, 1992). Green sulfur bacteria and heliobacteria contain type I reaction centers, which utilize an iron-sulfur cluster as the terminal electron acceptor. Purple photosynthetic bacteria have type II reactions centers, in which the terminal electron acceptor is a quinone. Both type I and type II reaction centers exist in oxygenic cyanobacteria and plants, named separately as Photosystem I and Photosystem II. The three dimensional structures of type II reaction centers was first obtained in the mid-1980s from the purple bacteria Rhodopseudomonas viridis and Rhodobacter sphaeroides (Deisenhofer et al., 1985; Allen et al., 1987). Recently, the crystal structures of Photosystem I and Photosystem II from the cyanobacterium Synechococcus elongatus were also determined at high resolutions (Jordan et al., 2001; Zouni et al., 2001). However, the structure of the type I reaction centers in green sulfur bacteria has not been successfully analyzed by crystallography due to their sensitivity to oxygen. Low- resolution images by STEM demonstrated that the reaction center particles from green sulfur bacteria have similar dimensions to the core of Photosystem I (Rémigy et al., 1999).

16 Using sequence alignment and structural coordinates, the structure of the type I reaction center from green sulfur bacteria has been modeled and described (Fig. 1-10) (Heathcote et al., 2003). The core of the reaction centers in green sulfur bacteria is built by two completely identical polypeptides termed PscA, which is a significant difference from the “heterodimeric” reaction centers made up of two related, but different polypeptides as is the case for Photosystem I and all type II complexes (Büttner et al., 1992b; Fromme et al., 2001). This is because the genomes of green sulfur bacteria only contain one gene for the core polypeptide, instead of two genes seen in cyanobacteria and higher plants. The PscA from C. tepidum is an 82 kDa protein, which includes 731 amino acids that bind the

primary electron donor P840, the primary electron acceptor A0 and the [4Fe-4S] cluster

FX (Hauska et al., 2001). The transmembrane helices for holding the redox compoments are conserved between PscA of green sulfur bacteria and the corresponding PsaA/B of Photosystem I (Büttner et al., 1992a; Liebl et al., 1993; Hauska et al., 2001). PscB, the 23

kDa protein binding the two terminal [4Fe-4S] electron acceptors FA and FB, is located at the stromal surface of the homodimeric PscA complex (Kjær et al., 1994; Hauska et al. 2001, Heathcote et al. 2003). The pscB gene is found to be co-transcribed with pscA forming the transcription unit pscAB in C. limicola f. sp. thiosulfatophilum and C. tepidum (Bütter et al., 1992; Hager-Braun et al., 1999). The primary structure of PscB is highly conserved in Chlorobium species, and it retains a similarity to PsaC of Photosystem I in its C-terminal region, which contains eight cysteines harboring the two [4Fe-4S] clusters (Hauska et al., 2001). PscC and PscD are also involved in forming the reaction centers. PscC is a monoheme cytochrome c molecule with three transmembrane helices at its N-terminus (Okkels et al., 1992). Electron transfer kinetics studies showed that two molecules of PscC are symmetrically arranged around the reaction center core (Kusumoto et al., 1999). PscD is a 15 kDa protein resides in the vicinity of PscB, which might facilitate the stabilization of PscB and/or the interaction with ferredoxin (Hager- Braun et al., 1995; Seo et al., 2001). Compared to Photosystem I, which accommodates around 90 light-harvesting chlorophylls, the chlorophyll density in the reaction center of green sulfur bacteria is much lower. The reaction center core of green sulfur bacteria contains only 4 Chl a and

17 16 BChl a, with each PscA protein binding two and eight (Griesbeck et al., 1998; Permentier et al., 2000). Two of the 16 BChl a molecules are 132-epimers (BChl a’), and are considered to form the special pair of the primary donor P840 (Kobayashi et al., 2000; Hauska et al., 2001). The four chlorophyll a molecules are esterified at C17 to 2,6- phytadienol instead of the normal phytol group (van de Meent et al., 1992; Feiler et al., 1994; Hauska et al., 2001). Two chlorophyll a molecules are known to be the primary electron acceptor A0, and the other two are proposed to be located at the positions of the accessory chlorophylls (A) in Photosystem I (Heathcote et al., 2002). However, no conclusive evidence has been obtained for the participation of a quinone electron

acceptor A1 between A0 and the iron-sulfur cluster FX (Heathcote et al., 2002). The carotenoid content in the reaction center of green sulfur bacteria is also substantially lower than that in the aero-tolerant Photosystem I. Only two carotenoids are contained in the reaction center core, for a molar ratio of one per 10 Chls/BChls (Hauska et al., 2001).

1.2.5 Energy transfer pathway and kinetics

Up to 200,000 BChl c, d or e molecules are contained in each chlorosome of green sulfur bacteria, with more than 5000 BChl molecules harvesting light for a single reaction center (Montaño et al., 2003a). The energy transfer pathway has been revealed by absorption and fluorescence spectra studies as well as biochemical resolution and separation. The main absorption peaks of BChl molecules are between 720 to 750 nm, and the excitation energy is transferred via the so-called baseplate containing BChl a-795 to the FMO-BChl a complex, with an absorption maximum at 808 nm. FMO further transfers the excitation energy to the reaction center, with a pair of BChl a serving as the primary electron donor P840. Thus, the excitation energy harvested by BChl c is transferred to pigments with progressively lower energy levels: BChl a on the baseplate, BChl a combined with FMO protein, and finally to the P840 BChl a dimer within the reaction center (Blankenship and Matsuura, 2003). Early time-resolved fluorescence measurements in Chlorobium limicola indicated that about 90% of the BChl c molecules transferred energy to BChl a in 20-50 ps (Borisov et al., 1977; Fetisova and Borisov 1980 a, b). Later fluorescence measurements

18 in isolated chlorosomes reported fluorescence lifetimes of less than 30 ps for BChl c emission (Brune et al., 1987a). However, energy transfer from FMO protein to the reaction center can not be observed in isolated FMO-reaction center complexes, and the kinetics is still poorly understood (Oh-oka et al., 1998b; Neerken et al., 1998). The energy transfer from carotenoids to BChl has hardly been studied. The carotenoids in the chlorosomes are considered to transfer light energy to BChl c and the carotenoids in the baseplate are considered to transfer light energy directly to BChl a (Blankenship and Matsuura, 2003). Interestingly, the energy transfer process in the chlorosomes of green sulfur bacteria appears to be regulated by redox potential in either isolated chlorosomes or whole cells. The energy transfer efficiency can be reduced from nearly 100% to less than 10% by adding oxidizing reagents, with a 10- to 50-fold drop in BChl c fluorescence and a decreased lifetime of the excited-state of BChl c (Wang et al., 1990; Blankenship et al., 1993; Blankenship et al., 1995). The quenching is a reversible process and energy transfer efficiency can be restored to 100% by adding reducing reagents such as sodium dithionite (Wang et al., 1990). Flash-induced electron transfer reactions showed that electron transfer in reaction centers was triggered by selective excitation of BChl c but not BChl a (Frigaard and Matsuura, 1999). Thus, the quenching mechanism in chlorosomes effectively regulates energy transfer in the reaction center. The quenching mechanism can also be observed in isolated FMO proteins. The fluorescence lifetime of the FMO protein dropped from approximately 2 ns in strongly reducing conditions to approximately 60 ps in neutral or oxidizing conditions (Zhou et al., 1994). Conformational or structural changes of the FMO protein can also be induced by changes in the redox potential (Zhou et al., 1994). A 60 ps component is proposed to be related to energy or electron transfer between excited BChl a and a redox active group or quencher (Zhou et al., 1994). The quenching of BChl c in chlorosomes and FMO proteins might be totally different mechanisms that operate on two separate levels. For quenching in chlorosomes, it is proposed that chlorosomes contain a component that acts as a highly quenching center of excited BChl c under oxidizing conditions (Wang et al., 1990). A unique isoprenoid quinone called chlorobiumquinone (1’-oxomenaquinone-7) with a redox

19 midpotential of 36 mV is suggested to be such a quencher molecule (Powls and Redfearn, 1969; Frigaard et al. 1997). Chlorobiumquinone is mainly located in the chlorosome and could be responsible for, or at least involved in, the redox-dependent quenching in chlorosomes. It is notable that Chloroflexus aurantiacus does not contain chlorobiumquinone and does not exhibit the redox-dependent quenching mechanism (Blankenship and Matsuura, 2003). The quenching/restoration might be a protection mechanism against reactive oxygen species which can be formed by oxidation of cellular reductants when oxygen is in the environment (Frigaard and Matsuura, 1999). Quenchers (most probably chlorobiumquinone) within the chlorosomes are oxidized and activated under oxidizing conditions, and they efficiently quench the energy transfer to decrease the formation of cytotoxic oxygen species under transient exposure to air. Chlorobium tepidum can withstand very long exposure to oxygen in darkness but is extremely sensitive to oxygen when cells are subjected to even weak illumination (Bryant et al., unpublished data).

1.2.6 Electron transfer and NAD(P)+ reduction

Time-resolved optical spectroscopy provided details in primary charge separation and electron transfer steps in the reaction centers of green sulfur bacteria. The primary charge separation of P840 was observed after a time of 10-30 ps (Nuijs et al., 1985; Shuvalov et

al., 1986). The primary electron acceptor A0 is located approximately halfway across the + - membrane. The decay time of the primary radical pair P840 A0 to the triplet state of

P840 was measured to be 20-35 ns (Swarthoff et al., 1981). Between A0 and P840 is a second chlorophyll pair (A) occupying the position of the accessory chlorophylls in Photosystem I. The A chlorophyll probably acts as an electron transfer intermediate

between P840 and A0 (Heathcote et al., 2003). The forward electron transfer from A0 occurs within about 600 ps as determined by laser-flash spectroscopy (Nuijs et al., 1985; Shuvalov et al., 1986). Redox potentials and halftimes of electron transfer in the P840 reaction center are summarized in Fig. 1-11. The participation of a quinone acceptor A1 has not been confirmed, and A1 is not shown between A0 and FX but is placed into a side path (Heathcote et al., 2003). The electron transfer chain terminates with three bound

20

[4Fe-4S] iron-sulfur clusters FX, FA and FB (Vassiliev et al., 2001). Two copies of the PscC (cytochrome c) subunit copurify with the P840 reaction center and act as the electron donors to P840+. The half-life of electron donation is 7 μs in whole cells, and 100 μs in isolated reaction centers (Prince and Olson, 1976; Oh-Oka et al., 1995; Kusumoto et al., 1999). The iron-sulfur clusters in the reaction center of green sulfur bacteria have very low mid-point potentials and can directly reduce ferredoxins at high rates (Feiler and Hauska, 1995; Sakurai et al., 1996; Rémigy et al., 1999). Soluble 2[4Fe-4S] ferredoxins have been purified from Chlorobium tepidum, and have been shown to act as electron acceptors from purified reaction center preparations (Seo et al., 2001). The reduced ferredoxins diffuse freely in the cytoplasm and are utilized in the reverse TCA cycle for carbon reduction (Buchanan and Arnon, 1990; Sirevåg, 1995). Green sulfur bacteria can reduce NAD(P)+ directly via the photo-reduced ferredoxins without additional energy requirement (Buchanan and Evans, 1969; Kusai and Yamanaka, 1973; Shioi et al., 1976). The ferredoxin:NAD(P)+ oxidoreductase (FNR) capable of producing NAD(P)H directly from photo-reduced ferredoxins was recently identified in Chlorobium tepidum (Seo and Sakurai, 2002).

21

Fig. 1-1. Sulfide and thiosulfate oxidation pathways of Chlorobium. (Slightly modified from Brune, 1989).

22

Fig. 1-2. The reductive tricarboxylic acid cycle of Chlorobium as proposed by Evans et al. (1966) (Adapted from Sirevåg, 1995)

23

Fig. 1-3. Phylogenetic tree of green sulfur bacteria based on 16S rDNA sequences. Tree and branch lengths were calculated by distance methods using DNADIST and FITCH from the PHYLIP program package. (Adapted from Alexander et al., 2002)

24

Fig. 1-4. Circular representation of the C. tepidum genome. Circles 1 and 2: predicted protein-coding regions on the plus and minus strand. Circles 3, (red), 4 (blue), and 5 (green): proteins with top matches to proteins from Archaea, photosynthetic species, and C. tepidum, respectively. Other circles: 6, GC skew; 7, percent G+C; 8, 2 value for trinucleotide composition in a 2,000-bp window; 9, tRNA genes; and 10, rRNA genes (blue) and sRNA genes (red). (Adapted from Eisen et al., 2002)

25

Fig. 1-5. Simplified chlorosome model showing interactions between the reaction center, the FMO protein and the chlorosome. BChl c aggregation is shown in the common favored rod model (left side, Nozawa et al., 1994) and the recently proposed alternative lamellar model (right side, Pšenčík et al., 2004). Light and excitation energy transfer is shown by thick arrows. (Adapted from Frigaard et al., 2005b).

26

Fig. 1-6. Structures of various bacteriochlorophylls (BChls). Panel A. Bacteriochlorophylls of the bacteriochlorin type, with single bonds between C7/C8 and C17/C18: bacteriochlorophylls a, b and g. Panel B. Bacteriochlorophylls of the chlorin type, with a single bond between C17 and C18: bacteriochlorophylls c, d and e. Abbreviations: et, ethyl; me, methyl; ib, isobutyl; np, neopentyl; pr, propyl. (Slightly modified from Scheer, 2003).

27

Fig. 1-7. Model of the chlorosome interior structure with bilayer tubes as the building block. A single rod consists of double layers of neighboring syn- (inner ring) and anti-stacks (outer ring) of BChls. The dotted lines indicate hydrogen bonds between 131-C=O and 31-OH of adjacent stacks. (Slightly modified from Rossum et al., 2001)

28

Fig. 1-8. Model of the chlorosome interior structure with BChl c lamellae as the building block. Panel A. The undulating lamellae extend along the long axis of the chlorosome (z), and through the height of the chlorosome (y). The arrangement of antiparallel dimers of BChl c (Panels B and C) is compatible with the observed lattice dimensions. (Adapted from Pšenčík et al., 2004)

29

Fig. 1-9. Structure of the Fenna-Matthews-Olson protein from Chlorobium tepidum as determined by Li et al. (1997). The β-strands are blue, the α- helices are green, and the bacteriochlorophyll structures, with phytyl chains removed for clarity, are red with a central yellow Mg. (Adapted from Li et al., 1997)

30 Fig. 1-10. Structural organization of the Photosystem I reaction center from Synechococcus elongatus, and proposed organization of the reaction center in Chlorobium limicola. Panel A. The electron transfer cofactors of the S. elongatus Photosystem I complex, including the primary electron donor (P700), the primary electron acceptors (A and A0), the secondary electron acceptor (A1) and three iron-sulfur clusters (FX, FA and FB). Panel B. The protein component of the S. elongatus Photosystem I complex. The PsaA and PsaB subunits are shown in maroon and green, the PsaC subunit is shown in cyan. Panel C. Proposed organization of the reaction center from C. limicola. The homodimer of PscA is shown in green and maroon, the PscB subunit is shown in cyan, the PscD subunit is shown in orange.

31

32

Fig. 1-11. Redox potentials and halftimes of electron transfer in the P840- reaction center, measured in isolated reaction centers at room temperature. Fd: ferredoxin. (Adapted from Hauska et al., 2001)

33

Chapter 2

Molecular Contacts for Chlorosome Envelope Proteins Revealed by Chemical Cross-linking Studies with Chlorosomes from Wild-type and Mutant Strains of Chlorobium tepidum

34 2.1 Abstract

Chlorosomes are unique light-harvesting antenna in the green sulfur bacteria with BChl c aggregates enclosed by a monolayer of protein/galactolipid membrane. In the chlorosomes of Chlorobium tepidum, ten proteins (CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ, and CsmX) are embedded in the monolayer envelope. Wild- type chlorosomes and chlorosomes from mutants lacking a single protein were cross- linked with zero-length cross-linker 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) and analyzed by SDS-PAGE; the cross-linked products were detected by immunoblotting with polyclonal antibodies raised against chlorosome proteins. CsmA was found to form dimers, trimers, and other multimers up to dodecamers in wild-type chlorosomes, SDS-treated chlorosomes and carotenosomes (the vestigial chlorosomes in bchK knock-out mutant). Given the biochemical evidence that CsmA binds bacteriochlorophyll (BChl) a (Bryant et al., 2002), CsmA is most likely to be localized in the baseplate region that interacts with the Fenna-Matthews-Olson (FMO) layer between the chlorosome baseplate and the cytoplasmic membrane. Consistent with this hypothesis, CsmA and FmoA in isolated chlorosomes could be cross-linked. The precursor form of CsmA, denoted pre-CsmA, was cross-linked to CsmA, CsmB, and CsmF. This suggests that pre-CsmA is probably located on the edge of the CsmA baseplate and that the interactions between pre-CsmA, CsmB, and CsmF might only occur at the edges of the baseplate array of CsmA. CsmC is another chlorosome protein that forms homo- multimers. Chlorosome proteins CsmI and CsmJ, which have N-terminal domains similar to adrenodoxin-type ferredoxins and ligate [2Fe-2S] clusters, form heterodimers, and each protein additionally interacts with CsmB. Based on the information obtained from cross-linking experiments, a model for protein organization and interactions within the chlorosome envelope membrane is proposed.

35 2.2 Introduction

Photosynthetic organisms have evolved many different light-harvesting systems, whose functions are to absorb light energy and transmit that energy efficiently to photosynthetic reaction centers. Examples include phycobilisomes of cyanobacteria and red algae, peridinin-chlorophyll proteins of dinoflagellates, and caroteno-chlorophyll a/b binding proteins (CAB/LHC) of higher plants (Blankenship, 2002). The green sulfur bacteria (Chlorobi) and green gliding bacteria (Chloroflexi) have their own unique solution to the light-harvesting problem. These organisms harbor unique organelles known as chlorosomes to collect light energy at extraordinarily low light intensities (a few μmol photons m-2 s-1). Chlorosomes are flattened ellipsoid-shaped structures containing highly aggregated BChl c/d/e, a small amount of BChl a, carotenoids, and quinones, and they are enclosed by a protein-stabilized, galactolipid-containing monolayer membrane (Blankenship and Matsuura, 2003). The dimensions of chlorosomes are not strictly defined, but they are usually 100-200 nm long, 30 to 70 nm wide, and 30 to 40 nm thick (Oelze and Golecki, 1995; Staehelin et al., 1978, 1980; Frigaard et al., 2005). Unlike other light-harvesting antenna complexes, in which chromophores are rigidly held and oriented by proteins, the BChl c/d/e molecules in chlorosomes are organized by hydrogen bonding, magnesium chelation, and intermolecular hydrophobic interactions, and the protein-to-chlorophyll mass ratio in chlorosomes is remarkably low. Highly purified chlorosomes of the green sulfur bacteria Chlorobium tepidum contain at least ten different polypeptides: CsmA (6.2 kDa), CsmB (7.5 kDa), CsmC (14.3 kDa), CsmD (11.1 kDa), CsmE (6.7 kDa), CsmF (7.7 kDa), CsmH (21.6 kDa), CsmI (25.9 kDa), CsmJ (21.8 kDa), and CsmX (24.0 kDa). The N-terminal domains of CsmI, CsmJ, and CsmX have strong sequence similarity to adrenodoxin-type [2Fe-2S] ferredoxins (Vassilieva et al., 2001). The remaining chlorosome proteins can be divided into three structural families: CsmA/E, CsmB/F, and CsmC/D. The C-termini of CsmI, CsmJ and CsmX are distantly related to CsmA/E; the C-terminus of CsmH is similar to CsmC/D, while its N-terminus is related to CsmB/F (Vassilieva et al., 2000; Vassilieva et al., 2002b). Both CsmA and CsmE are produced as pre-proteins with carboxyl-terminal extensions of ~20 residues, which are proteolytically removed during chlorosome

36 biogenesis (Chung et al., 1994; Chung et al., 1995). The genes encoding all ten proteins were cloned, and polyclonal antisera were raised against the purified recombinant proteins (Chung et al., 1994; Chung et al., 1995; Chung and Bryant, 1996a, b; Vassilieva et al., 2002b). Subcellular localization experiments showed that all ten chlorosome proteins copurified proportionally with BChl c, and that none of them were substantially associated with any other subcellular fractions (Vassilieva et al., 2002b). Protease susceptibility mapping suggests that the chlorosome proteins are exposed at the surface of isolated chlorosomes to various extents (Chung and Bryant, 1996 a, b). Isolated chlorosomes were incubated in phosphate buffer with proteases such as trypsin, pronase and proteinase K. CsmA was resistant to any of the three proteases under control conditions, but became quite protease-sensitive after chlorosomes were treated with 0.1% Triton X-100 to disrupt the membrane envelope integrity. (Detergent treatments are known to extract most proteins of the chlorosome envelope (Bryant et al., 2002)). CsmB and CsmE were not readily digested with trypsin, but these proteins were sensitive to protease K and pronase. CsmC and CsmD were quite readily accessible to all three proteases. Chlorosome agglutination experiments using polyclonal antisera to the chlorosome proteins confirmed that all proteins are exposed on the chlorosome envelope (Chung and Bryant, 1996 a, b; Vassilieva et al., 2002b). Antisera to CsmA, CsmB, CsmC, CsmD, CsmE, CsmH, CsmI, CsmJ and CsmX are all capable of agglutinating and precipitating isolated chlorosomes, indicating that some portion of these chlorosome proteins are exposed to solvent in isolated chlorosomes. The only exception is that anti- CsmF antiserum failed to cause precipitation of chlorosomes; however, CsmF is obviously related in sequence to an abundant chlorosome protein CsmB, and CsmF was proved to copurify with chlorosomes by subcellular localization analysis. Detergent treatments of isolated chlorosomes disrupts the integrity of the chlorosome envelope, increases the protease susceptibility of chlorosome proteins, and even releases chlorosome proteins from the envelope (Vassilieva and Bryant, 1998; Bryant et al., 2002). CsmC, CsmD and CsmH were very easily extracted from the chlorosome envelope by incubation with 0.01% (w/v) SDS. Increasing the SDS concentration to 0.05% (w/v) resulted in the extraction of all of the chlorosome proteins except CsmA and a small proportion of CsmB, and incubation with 0.1% (w/v) SDS or

37 higher released virtually all of the chlorosome proteins except CsmA. In spite of the removal of the chlorosome envelope proteins, SDS-treated chlorosomes retained their characteristic size and shape (Bryant et al., 2002). In addition to the SDS-treated chlorosomes, the vestigial chlorosomes from bchK mutant also lack a full complement of chlorosome proteins. The bchK mutant lacks BChl c synthase and thus totally lacks BChl c (Frigaard et al., 2002). However, the bchK mutant produces vestigial chlorosomes, called carotenosomes, which contain a nearly unchanged amount of CsmA; a small amount of CsmD; trace amounts of CsmB, CsmE, CsmF, CsmI; and no other proteins (Frigaard et al., 2002; Frigaard et al., 2005). Carotenosomes, along with SDS-treated chlorosomes, provide interesting possibilities to gain insights into the organization of chlorosome proteins, especially CsmA. The main goal of the studies reported in this chapter is to detect the location and organization of chlorosome proteins by chemical cross-linking experiments. Two cross- linking reagents were used: glutaraldehyde and EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide). Glutaraldehyde results in amine-to-amine cross-linking reactions, whereas EDC results in amine-to-carboxylic acid (zero-length) cross-linking reactions. Cross- linked wild-type chlorosomes and chlorosomes from mutants lacking a single chlorosome protein (Frigaard et al., 2004a) were analyzed by SDS-polyacrylamide gel electrophoresis, and immunoblotting with antibodies against chlorosome proteins was employed to detect protein multimerization or interactions between various protein species. Based upon the information obtained from these cross-linking experiments, a model is proposed that shows the protein locations, organization and interactions on the chlorosome envelope membrane.

38 2.3 Materials and methods

2.3.1 Chlorobium tepidum strain and growth conditions

The wild-type strain of Chlorobium tepidum used in these studies was WT2321 (Wahlund and Madigan, 1995). The mutant strains lacking single chlorosome proteins were constructed as described by Frigaard et al., 2004a. The medium for C. tepidum was the same as described by Frigaard and Bryant (Frigaard and Bryant, 2001). The liquid

medium (CL) per liter contained 20 mL of salts A (per liter: 0.64 g Na2EDTA.2H2O, 10 g

MgSO4.7H2O, 2.5 g CaCl2.2H2O, 20 g NaCl); 20 mL of salts B (per liter: 25 g

NH4CH3COO, 20 g NH4Cl, 115 g Na2S2O3.5H2O); 20 mL of CL buffer (per liter: 25 g

KH2PO4, 105 g MOPS); and 1 mL trace elements (per liter: 5.2 g Na2EDTA.2H2O, 0.19 g

CoCl2.6H2O, 0.1 g MnCl2.4H2O, 1.5 g FeCl2.4H2O, 6 mg H3BO3, 17 mg CuCl2.2H2O,

188 mg Na2MoO4.2H2O, 25 mg NiCl2.6H2O, 70 mg ZnCl2, 2 mg Na2WO4.2H2O, 2 mg -1 -1 NaHSeO3); 50 μL of 10 mg resazurin mL ; and 20 μL of 1 mg vitamin B12 mL . A fresh solution of 0.6 g Na2S.9H2O and 2.0 g NaHCO3 was dissolved in 50 mL of water, and filter-sterilized, and added to 1 L of autoclaved CL medium to obtain the proper redox potential and pH. Small volume growth in medium CL was performed in a 42 ºC anaerobic chamber (Coy Laboratory Products, Grass Lake, MI). Large-volume cultures were grown in completely filled and tightly sealed 2 L bottles in a 45-48 ºC water-bath. Plating medium (CP) contained the same components as the liquid medium except for the addition of 0.36 g L-cysteine L-1 and 1.5% (w/v) agar or 1.0% (w/v) phytagel. The pH of

the CP medium was adjusted to 7.6 with NaOH, and no fresh Na2S solution was added after autoclaving. Once inoculated, the plates were incubated in an anaerobic jar (BBL

GasPak 100 system, Becton Dickinson, Sparks, MD) containing one disposable H2-CO2 generating envelope (BBL GasPak, Becton Dickinson) and a small test tube containing

0.1 g thioacetamide and 1 mL of 1 M HCl to generate H2S.

39 2.3.2 Isolation of light harvesting antenna complexes

Isolation of chlorosomes. Chlorosomes were isolated by sucrose-gradient ultracentrifugation as described by Vassilieva et al., 2002b. The C. tepidum cells were harvested from a 2-L culture by centrifugation and were disrupted by three passages through a chilled French press (124 MPa, 4 ºC) after lysozyme incubation (3 mg mL-1, room temperature for more than 20 min) in isolation buffer (2 M NaSCN, 10 mM Tris- HCl, 5 mM EDTA, 0.5 mM PMSF, 1 mM DTT, pH 7.5). Chlorosomes and cell debris were pelleted by ultracentrifugation at 220,000 × g for 2 h at 4 ºC, and the resuspended pellet was purified on 7 to 47% (w/v) continuous sucrose gradients by ultracentrifugation again at 220,000 × g for 18 h at 4 ºC. The chlorosome fraction was then collected, diluted 4-fold with phosphate-buffered saline (10 mM K-phosphate, 150 mM NaCl, 0.5 mM PMSF, 1 mM DTT, pH 7.2) and centrifuged at 240,000 × g for 1.5 h. The resulting pellet was resuspended in the same buffer and centrifuged again, and the firmly pelleted chlorosomes were resuspended in a small volume of phosphate-buffered saline with 1 mM PMSF and 2 mM DTT. The purified chlorosomes were aliquoted and stored at -80 ºC for future use.

Isolation of carotenosomes. Carotenosomes were isolated by a modified method of chlorosome isolation. After three passages through French press, the extract from the bchK mutant cells was clarified by centrifugation at 13,000 × g for 20 min at 4 ˚C. The supernatant was brought to 20% (w/v) sucrose, transferred to screw-capped ultracentrifugation tubes, and overlaid with one-half volume of isolation buffer containing 5% (w/v) sucrose. After centrifugation at 270,000 × g for 2 h at 4 ˚C, the carotenosomes floated as an orange band on top of the 5% sucrose solution. The carotenosome fraction was removed using a Pasteur pipette, aliquoted and stored at -80 ˚C until required.

40 2.3.3 Cross-linking of chlorosome proteins

Cross-linking of chlorosome proteins. Chlorosome proteins were cross-linked with glutaraldehyde or EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride). For cross-linking with glutaraldehyde, chlorosome samples were diluted with chlorosome buffer about three times to protein concentration of 1 mg mL-1. Glutaraldehyde was then added from 2% (w/v) stock to desired final concentrations (0.003% -1.5% (w/v)). The cross-linking mixtures were shaken at room temperature for 1 h and removed to 4˚C before SDS polyacrylamide gel electrophoresis. The cross-linking reagent employed more frequently was EDC, a compound that belongs to the carbodiimide family and that can cross-link an amine group to a carboxylic acid group at zero distance. Chlorosomes were diluted two-fold with conjugation buffer (0.1 M 2-[N-morpholino]ethane sulfonic acid (MES), pH 5.5) and incubated at room temperature for 10 min. One-ninth volume of 50 mM EDC was added to reach the final concentration of 5 mM, and the chlorosomes were incubated with gentle shaking at room temperature for variable times (5 min to 3 h). The cross-linking reaction was stopped by addition of 1/10 volume of 1 M ammonium acetate. Carotenosomes were washed twice with chlorosome buffer, concentrated ten-fold with centrifugal microconcentrator (12,000 × g for 30 minutes at 4˚C), and cross-linking was then performed as described above for chlorosomes.

SDS treatment of chlorosomes. Isolated chlorosomes were diluted 12-fold in Tris-wash buffer (25 mM Tris-HCl, pH 8.5, 150 mM NaCl), and sodium dodecyl sulfate (SDS) was added to a final concentration of 0.4% (w/v). The chlorosomes were incubated in SDS solution for 45 min, and the extracted proteins were separated from the retained chlorosomes using Microcon YM-100 centrifugal filtration devices (Millipore, Billerica, MA) by centrifugation at 13,000 rpm at room temperature. The retained chlorosomes were washed twice with 0.2 ml Tris-wash buffer to ensure the complete separation. Finally the retained chlorosomes were washed off from the Microcon filter membrane with a small volume of phosphate buffered saline.

41 2.3.4 SDS-PAGE and immunoblotting analysis

SDS polyacrylamide gel electrophoresis. The protein composition of various protein samples was analyzed by Tris-Tricine-buffered SDS polyacrylamide gel electrophoresis (Schägger and van Jagow, 1987). The stacking gel was 4% T and 2.6% C, and the resolving gel was 16% or 8% T and 3.3% C. Protein samples were precipitated from chlorosomes with cold acetone (sometimes optional); solubilized in 1X sample buffer (0.1 M Tris-HCl, pH 6.8, 24% (v/v) glycerol, 1% (w/v) SDS, 2% (v/v) β-mercaptoethanol, and 0.02% (w/v) Coomassie Blue G-250); and heated at 50 ˚C for 5 min or boiled for 1 min; and loaded onto gels for analysis. Protein samples containing ~15 μg BChl c were loaded for silver staining and samples containing ~60-200 μg BChl c were loaded for immunoblotting experiments. The gels were typically electrophoresed for 16-20 h under constant amperage using Tris-Tricine electrode buffer containing 0.1 M Tris, 0.1 M Tricine and 0.1% SDS (w/v).

Silver staining. Silver staining was performed as described by Blum et al. (1987). The gel was shaked in fixing solution (50% methanol (v/v), 12% acetate acid (v/v), 0.5 mL 37% (w/v) formaldehyde L-1 for more than 1 h and washed with 50% (v/v) ethanol three times for 20 min each. After 3 min of pretreatment in a solution containing -1 Na2S2O3·5H2O (0.2 g L ), the gel was incubated in impregnation solution (2 g AgNO3 L-1, 0.75 mL 37% (w/v) formaldehyde L-1, rinsed with water, and developed in a solution -1 -1 -1 of 60 g Na2CO3 L , 0.5 mL 37% (w/v) formaldehyde L and 4 mg Na2S2O3·5H2O L . The gel was developed until protein bands appeared with desired intensities (~5-10 min), was washed with water, and then scanned with an Epson Expression 1680 scanner (Epson America Inc., Long Beach, CA).

Immunoblotting analysis. For immunoblotting, proteins on the gels were transferred onto 0.45 μm nitrocellulose membranes (Schleicher & Schuell, Keene, NH) or 0.45 μm “Immunobilon-P” PVDF (polyvinylidene fluoride) membranes (Millipore, Billerica, MA) using a semi-dry transfer cell (Bio-Rad, Richmond, CA). Membranes were blocked with 5% (w/v) nonfat milk in Tris-buffered saline and incubated with rabbit-raised antibodies

42 against recombinant chlorosome proteins. The dilutions of antibodies against chlorosome proteins were as follows: anti-CsmA 1:2000; anti-CsmB 1:180; anti-CsmC 1:1000; anti- CsmD 1:800; anti-CsmE 1:180; anti-CsmF 1:1000; anti-CsmH 1:3000; anti-CsmI 1:5000; anti-CsmJ 1:2500; anti-CsmX 1:1000. Membranes were later incubated with goat anti- (rabbit-IgG) antibodies conjugated with horseradish peroxidase at a dilution of 1:7000 and immunoreactions were detected by enhanced chemiluminescence according to the instructions of the manufacturer (Amersham Pharmacia Biotech, Piscataway, NJ).

43 2.4 Results

2.4.1 Cross-linking of chlorosome proteins

Chlorosome proteins were cross-linked with 5 mM EDC for different times and at different temperatures, analyzed by SDS-PAGE, and detected by silver staining (Fig. 2-1). As seen from the figure, cross-linked products appeared within 5 min at 25 °C (Fig. 2-1, compare lanes 1 and 2) and were largely complete by 1 h (Fig. 2-1, lane 3). The cross- linking results obtained at different temperatures (Fig. 2-1, lanes 7 to 11) were similar. In all cases, CsmC, CsmD, pre-CsmA, and CsmF rapidly disappeared and these proteins were undetectable after cross-linking for 1 h at 25 °C. Cross-linked products with apparent masses of 12 kDa, 16 kDa, and 18 kDa accumulated rapidly, and many larger products could also be observed. Approximately 90% of the two most abundant chlorosome proteins, CsmA and CsmB, were cross-linked after 2 h at 25 ºC. Trace amounts of CsmI, CsmJ, CsmX and CsmH are still observable after cross-linking of 3 h at 25 ºC or 1 h at 50 ºC. Although performed for differing times and at different temperatures, the results shown in the left and the right halves of Fig. 2-1 can be overlaid and show nearly identical patterns of disappearance and appearance of protein bands. New protein bands at 12 kDa, 16 kDa, 18 kDa, 22 kDa, 30 kDa appeared under all conditions, and less well- resolved species with larger apparent masses also were observed. In many cases, cross- linked products increased in abundance and subsequently decreased, suggesting that larger cross-linking products were being formed. Interestingly, some cross-linking products (e.g., the product with apparent mass of 16 kDa) did not appear as well-resolved bands, and some products showed a trend of increasing apparent mass as time increased or as the temperature was increased. This could be due to intrachain cross-linking reactions or to modification to differing extents of sidechain groups with EDC. The cross- linking intermediate O-acylisourea can rearrange to N-acylurea, and this could lead to an altered electrophoretic mobility of the modified protein (Wong, 1991). The multiple bands appearing at about 7 kDa in Fig. 2-1, lane 6, are likely to be modified forms of CsmA and/or CsmB instead of other unmodified proteins.

44 Chlorosome proteins were also cross-linked by various concentration of glutaraldehyde (0.02-0.8% v/v) at room temperature for 5 min and detected by silver staining (Fig. 2-2). At the lowest concentration tested, 0.02% (v/v) glutaraldehyde, the bands of CsmC, CsmD and CsmF decreased only slightly after 5 min. The disappearance of these proteins was greater for chlorosomes treated with 0.04% (v/v) glutaraldehyde, and new protein species (e.g., at 12 kDa) could be observed. The 12-kDa cross-linking product was increased in abundance as the glutaraldehyde concentration increased but was somewhat lower in abundance at the highest glutaraldehyde concentration tested (0.8 % v/v). The bands of CsmI, CsmJ, CsmX and CsmH were significantly lower in intensity after treatment with 0.2% to 0.8% (v/v) glutaraldehyde. Moreover, the CsmA/CsmB band decreased in intensity when higher concentrations of glutaraldehyde were used. At 0.8% (v/v) glutaraldehyde, most of the CsmA/CsmB had disappeared completely and new species with apparent molecular masses of 12, 18, and 24 kDa had appeared. Although the results shown in Figs. 2-1 and 2-2 were obtained with different chemical reagents, the overall pattern of protein disappearance and the appearance of cross-linking products were similar. The cross-linking pattern thus does not appear to be dependent upon the chemical nature of the cross-linking reagent, but rather appears to be dependent upon the structure, location and organization of the chlorosome proteins. Because the electrophoretic resolution of the cross-linked products was substantially greater for proteins cross-linked with EDC, immunoblotting analyses of cross-linked proteins was only performed using this reagent.

2.4.2 Organization of CsmA

The 6.15-kDa protein CsmA is the most abundant protein of the chlorosome envelope (Vassilieva et al., 2000). To simplify the cross-linking experiments, chlorosomes were treated with 0.4% (w/v) SDS to extract all proteins except CsmA (Bryant et al., 2002). The remaining chlorosomes containing only CsmA were cross-linked with 5 mM EDC at room temperature for various times, and the cross-linked proteins were detected by silver staining and immunoblotting with anti-CsmA antibodies. As shown in Fig. 2-3, a regular pattern of new protein species, with masses of 12 kDa, 18 kDa, 24 kDa, and higher

45 molecular masses was detected by silver staining (Panel A), and immunoblotting analysis confirmed that all of these new species are due to cross-linking of CsmA (Panel B). These new protein species most likely represent dimers, trimers, tetramers and higher multimers of CsmA. The amounts of monomeric CsmA decreased and CsmA multimers increased steadily as the cross-linking time increased. CsmA dimers were highly abundant after only 5 min of cross-linking; the amount of dimeric CsmA continued to increase up to 0.5 h, but its abundance decreased as larger multimers were formed at later times. Fig. 2-4 clearly shows that CsmA multimers up to octamers were formed, and close inspection of the gels suggested that species up to dodecamers were probably formed at lower yield. Chlorosomes that had not been treated with SDS were also cross-linked in the same manner as described above, and cross-linking products were analyzed by SDS- PAGE and detected by silver staining and immunoblotting (Fig. 2-4). All of the multimeric CsmA species observed in the SDS-extracted chlorosomes (Fig. 2-3) can also be detected in the untreated chlorosomes (Fig. 2-4). This result establishes that the formation of these multimeric species is not the result of an artifact caused by the SDS treatment employed to remove the nine other chlorosome proteins, but rather the CsmA multimers reflect the true quaternary structure of CsmA in the chlorosome envelope. However, the immunoblotting results in Fig. 2-4, Panel B, have an apparently greater complexity due to the presence of cross-linked species containing the precursor form of CsmA (pre-CsmA). A bchK mutant of C. tepidum lacks BChl c synthase and makes vestigial chlorosomes that contain carotenoids but no BChl c (Frigaard et al., 2002; Frigaard et al., 2005b). Isolated carotenosomes contain a nearly unchanged amount of CsmA; a greatly reduced amount of CsmD; trace amounts of CsmB, CsmE, CsmF, CsmI; and no other proteins (Fig. 2-5). Carotenosomes were cross-linked with 5 mM EDC at room temperature for different times, and proteins were separated by SDS-PAGE and detected by silver staining and immunoblotting with anti-CsmA antibodies. CsmA dimers, trimers and multimers were detected by silver staining (Fig. 2-5, panel A) and confirmed with anti-CsmA antibodies (Fig. 2-5, Panel B). Since the antibodies were raised to the precursor form of CsmA, these antibodies have a much stronger cross-reaction with pre-

46 CsmA than CsmA (compare Fig 2-5, Panels A and B, lane 1). Interestingly, as revealed by immunoblotting, CsmA dimer and pre-CsmA dimer (12 kDa and 14 kDa, respectively) were present even in the control chlorosome sample that had not been cross-linked (Fig. 2-5, Lane 1). These naturally formed dimers even survived heating at 50 ˚C for 5 min in SDS sample buffer. The formation of such natural dimers is not understood, but it has been observed on several occasions, especially for samples that have been stored at –80 ˚C for a long time. To investigate the interactions of CsmA and pre-CsmA with other proteins, immunoblotting experiments with antibodies to CsmA were used to detect cross-linked species in chlorosomes isolated from mutants lacking single chlorosome proteins (Fig. 2- 6, Panel A, left). Different cross-linking patterns appeared in the lanes of mutant chlorosomes: the 15.7-kDa and faint 13.6-kDa products were not detected after cross- linking chlorosomes from the csmB mutant, and the 16-kDa and 13.9-kDa bands were not detected after cross-linking chlorosome proteins from the csmF mutant. These same bands were also detected by anti-CsmF antibodies, which cross-react with both CsmB and CsmF, in chlorosomes of the wild type or csmB and csmF single mutants (Fig. 2-6, Panel A, right). The missing bands provide direct evidence that both pre-CsmA and CsmA interact with CsmB and CsmF. The 14.5-kDa product found in all lanes analyzed with anti-CsmA is probably a dimer of pre-CsmA and CsmA (Fig. 2-6, Panel A, left). More complex hetero-multimers with molecular masses of ≥20 kDa formed from pre- CsmA, CsmA, CsmB and CsmF were also detected by immunoblotting. Chlorosome preparations contain variable amounts of contaminating FmoA (the FMO protein in C. tepidum) (Fig. 2-6, Panel B, right, lane ctrl). When proteins of wild- type chlorosomes were cross-linked, anti-FmoA antibodies detected the formation a 50- kDa product (Fig. 2-6, Panel B, right, lane wt). This same cross-linked species could be detected after cross-linking chlorosome proteins from mutants specifically lacking CsmB, CsmC, CsmD, CsmE, and CsmF (Fig. 2-6, Panel A, right). Although the results are not as clear because of the formation of CsmA multimers, a 50-kDa product was also detected with antibodies to CsmA (Fig. 2-6, Panel B, left, lane wt). These results show that CsmA can be cross-linked to FmoA. They are consistent with the expectation that CsmA in the

47 chlorosome baseplate must interact with FmoA, which in turn is bound to the reaction centers that reside in the cytoplasmic membrane.

2.4.3 Organization of CsmC and CsmD

Fig. 2-7 shows the cross-linking products of wild type and mutant chlorosomes detected by anti-CsmC antibodies. As shown in the left part of Fig. 2-7, the products with masses of approximately 28, 43, and 57 kDa were detected in the wild type samples as well as mutants lacking CsmB, CsmC, CsmD and CsmE. These results suggested that these products represent dimers, trimers and tetramers of CsmC that do not contain other proteins. Thus, like CsmA, CsmC is second chlorosome protein that forms homo- multimers. When the cross-linking time was increased to 30 min (Fig. 2-7, right), the yield of CsmC homotetramer increased, but this species was less abundant at longer cross-linking times of 1 to 3 h. The ~22-kDa and ~36-kDa products disappeared in the chlorosomes lacking CsmB but were observed after cross-linking chlorosome proteins from mutants lacking CsmD, CsmE, and CsmF. These results show that CsmB can interacts with CsmC monomers and dimers (Fig. 2-7, left). When this immunoblot was over-exposed, a faint ~25 kDa band with an electrophoretic mobility slightly greater than the CsmC homodimer was detected. This product was not detected from chlorosomes lacking CsmD. Although this was a faint band suggesting a low-abundance product, this result provides evidence that CsmC and CsmD may be weakly interacting. The cross-linking products from wild-type chlorosomes as well as those from mutants lacking CsmB, CsmC, CsmE, and CsmF were also blotted with antibodies against CsmD (Fig. 2-8). Cross-linking products of ~16 and ~21.5 kDa were found for all of the mutants as well as for carotenosomes. These results suggest that CsmD forms homodimers and that this protein also weakly interacts with CsmA in both chlorosomes and carotenosomes. However, this product was not detected with antibodies to CsmA (see Fig. 2-6). Additionally, in all samples except for carotenosomes, a product of ~33 kDa was observed that might represent CsmD homotrimers. A cross-linked product of ~18-kDa was found for all samples except for those lacking CsmB, namely the chlorosomes of the csmB mutant and the carotenosomes that only contain trace amounts

48 of CsmB. These results suggest that CsmD may interact with CsmB to form heterodimers. One interesting property distinguishes CsmD from other chlorosome proteins. After cross-linking, the samples are usually dissolved in 1.4 mL acetone to extract the lipids and pigments (mainly BChl c) and to pellet the proteins. This step helps to give better resolution during SDS-PAGE analyses and usually does not affect the amount of protein detected. However, in the absence of acetone extraction, nearly no CsmD (less than 10%) could be detected by immunoblotting from chlorosomes, even if the samples were boiled for 5 min before loading. Surprisingly, the acetone extraction was not necessary to detect CsmD in carotenosomes. These results suggest that large amounts of BChl c may interfere with the electrophoretic properties of CsmD during SDS-PAGE.

2.4.4 Interactions between CsmI/CsmJ and CsmB

Fig. 2-9 shows cross-linking products obtained with chlorosomes from the wild type and various mutant strains when probed with anti-CsmI and anti-CsmJ antibodies. A ~48-kDa product was detected with both anti-CsmI and anti-CsmJ antibodies in chlorosomes of wild type; these results suggest that this could be heterodimer of CsmI (25.9 kDa) and CsmJ (21.8 kDa). In chlorosomes of a mutant lacking CsmJ (Fig. 2-9, lane csmJ csmX ) the 48-kDa product was not detected but a CsmI homodimer (~52 kDa) was observed. Similarly, in chlorosomes of a mutant lacking CsmI (Fig. 2-9, lane csmI csmX) a broad band of ~42-kDa, presumed to represent a CsmJ homodimer, was detected. The broad nature of this product may reflect the sensitivity of CsmJ to proteolytic degradation and is consistent with the broad nature of CsmJ that was not cross-linked. Since all three cross- linking products are detected in chlorosomes of the wild type and mutant strains lacking CsmB, CsmD, CsmE, and CsmF, it is probable that CsmI and CsmJ form heterotetramers

that are a dimer of dimers: (CsmI2)(CsmJ2). When chlorosomes from csmB mutant were cross-linked, products of ~33 and ~40 kDa detected by anti-CsmI and of ~29 kDa detected by anti-CsmJ were not detected. Since these products were observed for chlorosomes from the wild type and other mutant strains, these results indicate that CsmB interacts with both CsmI and CsmJ.

49 2.4.5 Interactions for other chlorosome proteins

Cross-linking products of the remaining chlorosome proteins (CsmH, CsmE, and CsmX) were also detected. CsmH could be cross-linked with both CsmA and CsmB (Fig. 2-10). Although the level of CsmH is significantly reduced in chlorosomes of a csmC mutant (Frigaard et al., 2004a), no direct interaction between CsmH and CsmC was detected by cross-linking. A possible explanation for this unexpected result is that CsmC could play a role in the incorporation of CsmH into chlorosome envelope but not remain associated with it. CsmE cross-linking products were not well characterized because of the small amount of CsmE in chlorosomes and the low specificity and titer of the anti-CsmE antibodies. Nevertheless, a 12-kDa dimer between CsmB and CsmE was detected (data not shown). No major cross-linking products, as detected using other antibodies, were missing from chlorosomes lacking CsmE, so interactions between CsmE and other proteins seem unlikely. Finally, homodimers of CsmX and 35-kDa heterodimers of CsmX and CsmB were detected when wild-type chlorosomes were cross-linked (data not shown).

50 2.5 Discussion

Glutaraldehyde, one of the most extensively used cross-linking reagents, contains two reactive aldehyde groups that can react with primary and secondary amino groups. However, glutaraldehyde easily forms polymers in solution, and the distance between the two cross-linked groups cannot be estimated. The other cross-linking reagent used in these studies, EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide), is a zero length cross-linker and does not polymerize. EDC belongs to the carbodiimide family, which promotes the direct joining of carboxyl and primary amino groups to form amide bonds without the introduction of any extrinsic atoms. Cross-linking reactions between two polypeptides only occur when the free carboxyl and amino groups are on separate polypeptides and interacting through a salt bridge (zero distance). As a carboxyl group activating reagent, EDC first forms a highly reactive O-acylisourea intermediate with the carboxyl group, and a subsequent nucleophilic attack by an amino group results in the formation of a new amide bond and the elimination of the activating moiety (Wong, 1991). Competing reactions from the intermediate O-acylisourea include rearrangement to form a stable N-acylurea derivative, or hydrolysis to regenerate the original protein and a urea derivative (Wong, 1991). Unless specified otherwise, all of the cross-linking experiments described in this chapter were performed with EDC. The chlorosome envelope is described as a monolayer membrane of monogalactosyl diglyceride with the galactosyl head groups exposed on the outer surface (Staehelin et al., 1978; Holo et al., 1985; Vassilieva et al. 2002). The envelope galactolipid monolayer was reported to be 10 Å in thickness, much less than that of typical phospholipid bilayer membranes (Staehelin et al., 1978; Staehelin et al., 1980). Chlorosome proteins, which are embedded in the envelope membrane, are somewhat hydrophobic in nature but are clearly predicted to lack the trans-membrane α-helical structure that is typical of all proteins destined for the cytoplasmic membrane (Chung and Bryant, 1996 a, b). Thus, the chlorosome envelope is a highly unusual and asymmetric membrane system. Although it cannot be excluded that the galactolipids and proteins are mobile in the envelope, the majority of the cross-linking results do not appear to be the results of

51 membrane fluidity and random protein collisions. First of all, CsmA is responsible for binding BChl a and transferring light energy to reaction centers via FmoA, as been shown by proteolytic digestion and detergent treatments in either Chlorobium or Chloroflexus sp. (Sakuragi et al., 1999; Bryant et al., 2002, Montaño et al., 2003b). As the most abundant chlorosome protein, CsmA covers roughly 20-35% of the chlorosome surface, and as shown here, CsmA forms large paracrystalline arrays in which CsmA principally interacts with itself and not with other proteins. CsmA is not likely to diffuse freely around the envelope, since this would likely interfere with energy transfer to the reaction centers and might result in formation of BChl a triplets that could lead to cell damage (Blankenship and Matsuura, 2003). Secondly and most importantly, little evidence for the random formation of cross-linking products was observed in our results. In addition to CsmA, several other chlorosome proteins, including CsmC, CsmD, CsmI, CsmJ, and CsmX, also apparently form multimers, which are unlikely to have resulted from chance collisions. Membrane fluidity increases as the temperature increases (Los and Murata, 2004). If collisional cross-linking was occurring, there should be more types of random products formed at high temperature or after extended periods of cross-linking. However, similar of cross-linking patterns were observed at high or low temperatures and after both long and short cross-linking times (see Fig 2-1). Moreover, similar patterns of cross-linking were observed in mutants lacking individual chlorosome proteins. The immunoblotting results showed clearly that only specific proteins were cross-linked and that if cross-linking of proteins was occurring as a result of random collisions that these events are relatively rare in the chlorosome envelope. CsmA, the most abundant chlorosome protein, comprises roughly one-third to one-half of the total protein of chlorosomes in Chlorobium tepidum (Bryant et al., 2002). Although proteins other than CsmA were extremely easily extracted from the chlorosome envelope by ionic and nonionic detergents (SDS, Lubrol PX and Triton X-100), CsmA was only released by detergents in combination with a low concentration of 1-hexanol (Bryant et al., 2002). Moreover, CsmA was only protease sensitive when 0.1% Triton X- 100 was added to disrupt the integrity of the chlorosome envelope (Chung and Bryant, 1996b). Thus, CsmA is either compactly organized or deeply embedded in the chlorosome envelope. This conclusion is further supported by the observation of matrix-

52 assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS) of protease-digested chlorosome proteins, which provides direct evidence that the N- terminal portion of CsmA (residues 2-38, mainly hydrophobic) is not accessible to proteases and is most likely buried in the chlorosome envelope membrane (Milks et al., 2005). Cross-linking experiments with wild type chlorosomes showed that CsmA readily forms multimers, including dimers, trimers, tetramers and other multimeric species to perhaps dodecamers and beyond. CsmA must be organized into rather homogeneous, perhaps paracrystalline arrays, since homomultimers are the predominant species observed after cross-linking. The same cross-linking pattern was detected for untreated chlorosomes, SDS-treated chlorosomes and carotenosomes, indicating that CsmA maintains its quaternary structure even in the absence of all other chlorosome proteins. Given the evidence that isolated baseplate from chlorosomes of Chloroflexus aurantiacus is comprised of CsmA, BChl a and carotenoids (Sakuragi et al., 1999; Montaño et al., 2003b), and that the contents of BChl a and CsmA in chlorosomes of C. tepidum are equimolar (Bryant et al., 2002; Frigaard et al., 2004), CsmA in Chlorobium species is likely to be located in the chlorosome baseplate region, facing the FmoA layer and making direct contact with it. Consistent with this idea, CsmA could be cross-linked to the FmoA present in chlorosome preparations. Nine mutants of Chlorobium tepidum, each lacking a single chlorosome protein, were previously constructed and characterized (Frigaard et al., 2004a). Using chlorosomes isolated from the mutants and multiple antibodies against recombinant chlorosome proteins, the protein composition of the cross-linked products were characterized to detect molecular contacts of the proteins of the chlorosome envelope. A summary of the results is shown in Table 2-1. The most abundant chlorosome protein, CsmA, was shown to make contacts mostly with itself, with its precursor form, pre- CsmA, and with FmoA. In addition to its contacts with CsmA, Pre-CsmA could be cross- linked to CsmB and CsmF. Pre-CsmA is the precursor of CsmA and contains 20 more hydrophilic amino acids at the carboxyl-terminus of the polypeptide. The presence of pre- CsmA in chlorosomes indicates that the cleavage of this carboxyl-terminal extension happens after insertion of pre-CsmA into the chlorosome envelope. Since CsmA is mostly cross-linked into homomultimers, and since most of the cross-linking products

53 between CsmA and Pre-CsmA are low-molecular-mass products, it seems likely that cross-linking between CsmA and pre-CsmA probably occurs on the edges of the CsmA baseplate where pre-CsmA is initially inserted. CsmB is the second most abundant chlorosome protein for Chlorobium tepidum (Chung and Bryant, 1996a). CsmF has a very high degree of sequence similarity (63%) to CsmB (Vassilieva et al., 2000), but is present at much lower copy number per chlorosome (Frigaard et al., 2004a). Analyses of CsmB and CsmF indicate that these proteins are slightly hydrophobic throughout nearly their entire length (Højrup et al., 1991; Chung and Bryant, 1996a). After 5 min of cross-linking, the majority of CsmF was cross-linked, and the 13.9-kDa CsmA-CsmF product appeared to be more abundant than the 16-kDa pre-CsmA-CsmF product (Fig. 2-6, csmB lanes). This result suggests that CsmF is more readily to be cross-linked to CsmA than to pre-CsmA. However, the cross- linking between CsmB and pre-CsmA greatly exceeded the cross-linking between CsmB and CsmA (Fig. 2-6, csmF lanes), suggesting that CsmB principally interacts with pre- CsmA. Assuming that pre-CsmA is located along the edges of CsmA baseplate, then CsmF is most likely to be located between pre-CsmA and the edge of CsmA, while CsmB is more likely to be located outside of the CsmF and pre-CsmA molecules at the edges of the CsmA baseplate. Since CsmB can be cross-linked to nearly all other proteins, the cross-linking studies suggest that CsmB is probably distributed over much of the chlorosome envelope surface except for the baseplate region, from which it would be excluded by the paracrystalline nature of CsmA. Unlike CsmA, however, CsmB does not appear to form homo-multimers. CsmC and CsmD are the most readily released chlorosome proteins during detergent treatments with SDS, Lubrol PX and Triton X-100 (Bryant et al., 2002). These proteins are also quite accessible to proteases such as trypsin, proteinase K and pronase (Chung and Bryant, 1996b), and antibodies to these proteins readily agglutinate chlorosomes (Vassilieva et al., 2002b). These results indicate that significant portions of the CsmC and CsmD proteins are exposed at the surface of the chlorosome envelope. In the cross-linking experiments, CsmC and CsmD are the two protein species that were most readily cross-linked. CsmC was rapidly cross-linked into homo-dimers and homotrimers, and homo-tetramers were also detected after longer cross-linking times.

54 Chlorosomes from a csmC mutant were ~25% smaller than wild-type chlorosomes (Frigaard et al., 2004a). This observation suggests that multimeric CsmC plays a role in extending the chlorosome structure in the longitudinal direction. CsmC must be localized along the sides or the cytoplasmic surface of chlorosomes opposite to the baseplate. CsmD is hydrophobic throughout its entire length except near its amino- and carboxy-termini (Chung and Bryant, 1996b). Trypsin digestion suggested that both termini are exposed near the chlorosome surface (Chung and Bryant, 1996b). CsmD dimers were detected in both chlorosomes and carotenosomes. The latter contain nearly an unchanged amount of CsmA, a reduced amount of CsmD, trace amounts of CsmB, CsmE, CsmF, CsmI and no other proteins (Frigaard et al., 2002). Given that CsmD is the second-most abundant protein in the flattened carotenosomes, CsmD must be located in those regions of the chlorosome envelope that face the cytoplasm. CsmH is another protein that is very easily extracted from the chlorosome envelope by detergents (Vassilieva and Bryant, 1998). CsmH antiserum agglutinates chlorosomes with high efficiency, indicating that at least the most immunogenic epitopes of CsmH are highly exposed at the chlorosome envelope surface (Vassilieva et al., 2002b). CsmH did not form multimers, but this protein could be cross-linked to both CsmA and CsmB. These results suggest that CsmH is localized along the periphery of the baseplate. Mutants lacking CsmH also form somewhat smaller chlorosomes, and it is thus possible that, together with CsmC, CsmH plays some role in the longitudinal extension of the chlorosomes during their biogenesis. CsmI, CsmJ and CsmX are the three [2Fe-2S] proteins found in the chlorosome envelope (Vassilieva et al., 2001). Protein sequencing of chlorosome proteins initially only detected CsmI and CsmJ; the csmX gene was detected by genome sequencing and sequence alignment with csmI and csmJ, and immunological analyses indicate that CsmX is present at much lower levels than the other two proteins (Vassilieva et al., 2001). Antibody agglutination studies suggested that the surface-exposed regions of these three proteins might be relatively small compared to their overall sizes (Vassilieva et al., 2002b). The cross-linking studies performed here show that CsmI and CsmJ can form homodimers, and that heterodimers of these two proteins are readily formed during EDC cross-linking. These results are most easily interpreted as showing that these proteins

55

form hetero-tetramers (CsmI2)(CsmJ2) in wild-type chlorosomes but homodimers when one protein is missing in the csmI and csmJ mutants. CsmI, CsmJ and CsmX each interact with one or two molecules of CsmB. Table 2-1 and Fig. 2-11 summarize the conclusions drawn from the cross-linking experiments and show an overall model for chlorosome protein organization. Distribution of chlorosome proteins in the genomes of eight sequenced green sulfur bacteria other than C. tepidum is shown in Table 2-2. These sequenced green sulfur bacteria include Pelodictyon luteolum DSMZ 273(T), Chlorobium limicola DSMZ 245(T), Chlorobium phaeobacteroides DSMZ 266(T), Chlorobium vibrioforme f. thiosulfatophilum DSMZ 265(T), Chlorochromatium aggregatum, Pelodictyon phaeoclathratiforme BU-1, Prosthecochloris aestuarii SK413, and Prosthecochloris sp. BS1. Sequence identities are compared between the homolog proteins in C. tepidum and the other species of green sulfur bacteria. CsmA and CsmC are the two most highly conserved proteins present in the genomes of all the eight species, and CsmC, CsmE, CsmF, CsmH, CsmI, CsmJ and CsmX are conserved with moderately high percent-identity values in most of the organisms. The homolog of CsmD is only detected in Prosthecochloris aestuarii SK413. With most of the chlorosome proteins conserved, the protein organization in chlorosomes of C. tepidum proposed in this chapter is likely to be universal for the protein organization in the chlorosomes of the eight species of green sulfur bacteria listed in Table 2-2. In the nine mutants lacking single chlorosome proteins except CsmA, no significant changes in bacteriochlorophyll c content, mutant growth rates, or gross structural organization was observed (Frigaard et al., 2004a). These results indicate that chlorosomes are extremely robust structures that can tolerate considerable changes in their protein composition. Gene duplication and divergence is clearly the reason for the sequence relationships observed among chlorosome proteins. Although compensatory increases in the abundance of chlorosome proteins were not observed in the single mutants (Frigaard et al., 2004a), some structural and functional substitution might nevertheless occur when a single chlorosome protein is eliminated by mutation. Double and triple mutants, in which all chlorosome proteins belonging to a specific motif class

56 have been eliminated, have been constructed to provide further information about functions of the chlorosome proteins. These studies will be described in Chapter 4.

57 Table 2-1. Interactions and locations of chlorosome proteins

Protein Mass Cross-linking properties Comments/Localization (kDa) CsmA 6.2 Dimers, trimers, multimers up to Baseplate dodecamers; cross-linked to pre- CsmA, CsmB, CsmF, CsmD, FmoA Pre-CsmA 8.3 Cross-linked to CsmA, CsmB, CsmF Edges of the baseplate

CsmB 7.5 Cross-linked to CsmA, CsmC, Everywhere except the baseplate CsmD, CsmE, CsmH, CsmI, CsmJ, CsmX CsmC 14.3 Dimers, trimers and tetramers; cross- Cytoplasmic surface opposite the linked to CsmB baseplate or along the sides CsmD 11.1 Dimers and trimers; cross-linked to Cytoplasmic surface opposite the CsmA and CsmB baseplate or along the sides CsmE 7.5 Cross-linked to CsmB No evidence; BChl a-binding? CsmF 7.7 Cross-linked to CsmA and Pre-CsmA Edges of the baseplate CsmH 21.8 Cross-linked to CsmB and CsmA Cytoplasmic surface opposite the baseplate or along the sides CsmI 25.9 Cross-linked to CsmI, CsmJ and Forms homo-dimers and CsmI-CsmJ CsmB hetero-tetramers; location uncertain CsmJ 23.9 Cross-linked to CsmJ, CsmI and Forms homo-dimers and CsmI-CsmJ CsmB hetero-tetramers; location uncertain CsmX 24.0 Cross-linked to CsmX and CsmB Forms homo-dimers; location uncertain

58 Table 2-2. Distribution of chlorosome proteins in the genomes of all sequenced green sulfur bacteria other than Chlorobium tepidum. The values shown here are percent identity between the homolog proteins of C. tepidum and the listed species.

59

Fig. 2-1. Wild-type chlorosomes cross-linked with EDC for various times and at various temperatures. Proteins corresponding to ~15 μg BChl c were loaded for each lane, separated by 16% SDS-PAGE and detected by silver staining. Lane 1: uncross-linked chlorosomes as control. Lane 2-5: chlorosomes cross-linked at 25 °C for 5 min, 0.5 h, 1 h, 2 h and 3 h. Lane 7-11: chlorosomes cross-linked at 0 °C, 25 °C, 37 °C, 50 °C and 70 °C for 1 h.

60

Fig. 2-2. Wild-type chlorosomes cross-linked at room temperature for 5 min with various concentrations of glutaraldehyde. Protein samples corresponding to ~15 μg BChl c were loaded for each lane, separated by 8% SDS-PAGE and detected by silver staining. Lane 1: uncross-linked chlorosomes as control. Lane 2-8: chlorosomes cross-linked with glutaraldehyde at concentrations of 0.02%, 0.04%, 0.08%, 0.1%, 0.2%, 0.4% and 0.8% (v/v), respectively.

61

Fig. 2-3. SDS-treated chlorosomes cross-linked for various times at room temperature. Protein samples corresponding to ~15 μg BChl c were loaded for silver staining (Panel A) and samples corresponding to ~200 μg BChl c were loaded for immunoblotting detected with anti-CsmA antibodies (Panel B). Lane 1: uncross-linked SDS-treated chlorosomes as control. Lanes 2-5: SDS-treated chlorosomes cross-linked for 5 min, 0.5 h, 1 h, 2 h and 3 h.

62

Fig. 2-4. Wild-type chlorosomes cross-linked for various time ranges at room temperature. Proteins corresponding to ~15 μg BChl c were loaded for silver staining (Panel A), and samples corresponding to ~200 μg BChl c were loaded for immunoblotting detected with anti-CsmA antibodies (Panel B). Lane 1: uncross-linked chlorosomes as control. Lanes 2-5: chlorosomes cross-linked for 5 min, 0.5 h, 1 h, 2 h and 3 h.

63

Fig. 2-5. Carotenosomes, isolated from a bchK mutant of C. tepidum, were cross-linked for various times at room temperature. Protein samples (~3 μg) were loaded for silver staining (Panel A), and protein samples ~40 μg) were loaded for immunoblotting detected with anti-CsmA antibodies (Panel B). Lane 1: uncross-linked carotenosomes as control. Lanes 2-5: carotenosomes cross-linked for 5 min, 0.5 h, 1 h, 2 h and 3 h.

64

Fig. 2-6. Chlorosome cross-linked products detected by antibodies against CsmA (Panels A and B, left), CsmF (Panel A, right) and FmoA protein (Panel B, right). Chlorosomes from wild type and the indicated mutants were cross-linked at room temperature for 5 min before polyacrylamide electrophoresis. Protein samples corresponding to ~100 μg BChl c were loaded for each lane. Proteins and cross-linking products are indicated at the sides of the figure (see text). CsmA and pre-CsmA interact with CsmB and CsmF, and CsmA interacts with FmoA.

65

Fig. 2-7. Cross-linked products for chlorosomes detected with antibodies against CsmC. Chlorosomes from wild type and the indicated mutants were cross-linked at room temperature for 5 min (left) or various times (right) before polyacrylamide electrophoresis. Protein samples corresponding to ~200 μg BChl c were loaded for each lane. Proteins and products are identified at the right of the figure. CsmC forms homo- multimers and interacts with CsmB.

66

Fig. 2-8. Cross-linked products from chlorosomes detected by antibodies against CsmD. Chlorosomes from wild type and the indicated mutants were cross-linked at room temperature for 5 min before polyacrylamide electrophoresis. Protein samples corresponding to ~100 μg BChl c were loaded for the wt, csmB, csmC, csmE and csmF lanes, and and a ~25-μg protein sample was loaded for the carotenosome lane. Proteins and cross- linked products are identified to the right of the figure. CsmD forms homo-dimers, homo-trimers, and interacts with CsmA and CsmB.

67

Fig. 2-9. Cross-linked products from chlorosomes detected by antibodies against CsmI (Panel A) and CsmJ (Panel B). Chlorosomes from wild type and the indicated mutants were cross-linked at room temperature for 1 h before polyacrylamide electrophoresis. Protein samples corresponding to ~60 μg of BChl c were loaded for each lane. Proteins and cross-linked products are indicated at the right edges of the figures. CsmI and CsmJ form homo-dimers and hetero-dimers. CsmB interacts with both CsmI and CsmJ.

68

Fig. 2-10. Cross-linked products from chlorosomes detected by antibodies against CsmH. Chlorosomes from wild type and the indicated mutants were cross-linked at room temperature for 1 h before polyacrylamide electrophoresis. Protein samples corresponding to ~60 µg BChl c were loaded for each lane. Proteins and cross-linked products are indicated at the right of the figure. CsmH interacts with both CsmA and CsmB.

69

Fig. 2-11. Model of protein organization of the chlorosome envelope (assumes a rod-like organization of the BChl c).

70

Chapter 3

[2Fe-2S] Proteins in the Chlorosome: Construction and Characterization of Mutants Lacking CsmI, CsmJ and CsmX in the Chlorosome Envelope of Chlorobium tepidum

71 3.1 Abstract

Chlorosomes of the green sulfur bacteria are formed from highly aggregated bacteriochlorophyll c rods surrounded by galactolipid/protein monolayer envelopes. Analyses of isolated chlorosomes from Chlorobium tepidum has shown the presence of ten proteins, and three chlorosome proteins, CsmI, CsmJ, and CsmX have strong sequence similarity in their amino-terminal domains to [2Fe-2S] ferredoxins of the adrenodoxin/putidaredoxin subfamily. Insertional inactivation of the three iron-sulfur proteins has been made in all combinations. Mutant strains lacking the iron-sulfur proteins presented similar or slightly slower growth rates compared with the wild type. Chlorosomes were isolated from the mutant strains, and comparison of the wild-type and mutant chlorosomes in fluorescence quenching/restoration strongly suggests that the iron-sulfur proteins, especially CsmI and CsmJ, play important roles in the redox regulation of energy transfer within the chlorosome. Under oxygen stress conditions, CsmI/J might transfer electrons from the quencher within the chlorosome (most probably chlorobiumquinone) to molecular oxygen to active the quencher and reduce the energy transfer from BChl c rods to reaction centers. Under oxygen release conditions, CsmI/J might transfer electrons in the other direction: from reductant outside of chlorosome to the chlorosome quencher to inactivate the quencher and restore the energy transfer. Similar quenching/restoration mechanisms also exist in whole cells. The quinone contents of chlorosomes have been measured by HPLC analysis using BChl c amounts as the standard. Although the total quinone contents were similar within chlorosomes of the wild type and the mutants, chlorosomes of the csmI csmJ double and csmI csmJ csmX triple mutants contain significantly less chlorobiumquinones and more menaquinone-7. The CsmX protein, given the low concentration (~5%) compared to CsmI and CsmJ, makes little contribution to fluorescence quenching/restoration. Although redox titration in the csmX mutant showed that the protein ratio of CsmI:CsmJ changed from 1:1 (wild type) to 2:1 (Johnson et al., data unpublished), RT-PCR and immunoblotting analysis detected no obvious variation in csmI and csmJ transcription and expression. The changed protein ratio observed by redox titration could be due to partial degradation of CsmJ in that particular batch of chlorosomes.

72 3.2 Introduction

Green sulfur bacteria are obligate photolithoautotrophs that normally grow under light- limited and strictly anaerobic conditions. They contain unique light-harvesting antenna complexes, known as chlorosomes, which effectively harvest light in extremely low-light environments. The sac-like chlorosomes are filled with bacteriochlorophylls (BChl) c, d or e in a highly aggregated state (Oelze and Golecki, 1995; Blankenship and Matsuura, 2003). In model green sulfur bacterium Chlorobium tepidum, the monolayer chlorosome envelope contains ten proteins, which are named CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ and CsmX (Chung and Bryant, 1996 a, b; Vassilieva et al., 2002b). Among these chlorosome proteins, three (CsmI, CsmJ and CsmX) share sequence similarities to adrenodoxin-type ferredoxins at their amino termini (Vassilieva et al., 2000). The cysteine motifs responsible for binding the [2Fe-2S] clusters are completely conserved in CsmI, CsmJ and CsmX. The carboxyl termini of these proteins has been suggested to be related to the precursor form of CsmA and CsmE (Vassilieva et al., 2000). In addition to the obvious sequence similarity to ferredoxins, several lines of evidence indicate that CsmI and CsmJ (probably CsmX) contain [2Fe-2S] clusters. CsmI and CsmJ from C. tepidum were overproduced in E. coli as inclusion bodies (Vassilieva et al., 2001). Under reducing conditions, inclusion bodies for both proteins exhibited characteristic, nearly axial EPR spectra characteristic of [2Fe-2S] ferredoxins. EPR characterization of reconstituted, recombinant CsmJ showed a spectrum with principal g- values of 2.018, 1.936 and 1.930, and the temperature dependent properties of this spectrum strongly imply that it arise from a [2Fe-2S] cluster. An EPR spectrum of purified CsmI was not obtained because of the very limited solubility of the CsmI protein. Moreover, chemically reduced chlorosomes from C. tepidum have an EPR spectrum with principal g-values of 2.018, 1.935, and 1.933, which are virtually identical to those of recombinant CsmI and CsmJ proteins in inclusion bodies or in solution (Vassilieva et al., 2001). The EPR properties of the chlorosomes are consistent with the observation that iron-sulfur clusters with a g-value of 1.94 existed in the membrane fragments (with chlorosomes attached) in Chlorobium limicola f. thiosulfatophilum (Knaff and Malkin,

73 1976). All of these data strongly imply that CsmI and CsmJ (probably CsmX) are [2Fe- 2S] proteins on the chlorosome envelope. The redox potential of the CsmJ from inclusion bodies was determined to be -194 mV, and isolated chlorosomes were shown to contain Fe/S clusters with redox potentials of -201 mV and +92 mV (Vassilieva et al., 2001). However, the intracellular midpoint potential in strictly anaerobic organisms like Chlorobium sp. might be as low as -400 mV to allow the reverse TCA cycle reactions to occur. Thus, chlorosome proteins with relatively high redox potentials are not likely to be involved in electron transfer to electron transfer proteins that participate in the reverse TCA cycle reactions. Alternative roles for CsmI, CsmJ and presumably CsmX might be to participate in the redox regulation of energy transfer in the chlorosomes. In green sulfur bacteria, energy transfer is strongly regulated by redox potential in both isolated chlorosomes and whole cells. The energy transfer efficiency can be reduced from nearly 100% to less than 10% by exposing cells or chlorosomes to oxygen or other oxidizing reagents, which cause a 10- fold to 50-fold drop in BChl c fluorescence and a decreased lifetime of the excited state of BChl c (Wang et al., 1990; Blankenship et al., 1993; Blankenship et al., 1995). Fluorescence quenching is a reversible process, and energy transfer efficiency can be restored to 100% by adding reducing reagents such as sodium dithionite (Wang et al., 1990). This quenching (and its restoration) might provide a temporary and highly protective mechanism against the formation of reactive oxygen species, which can be formed by oxidation of the strong, cellular reductants when oxygen is present in the environment (Frigaard and Matsuura, 1999). Quenchers, most probably chlorobiumquinone, within the chlorosomes are oxidized and activated under such conditions, and the high concentrations of oxidized quinones could efficiently quench the energy transfer to decrease the formation of cytotoxic oxygen species under transient exposure to oxic conditions (Wang et al., 1990; Frigaard et al., 1997). CsmI, CsmJ, and CsmX were proposed to play a role in activating and inactivating the quencher by transferring electrons under oxidative stress or stress-release conditions (Vassilieva et al., 2001). In this study, the functions of iron-sulfur proteins, CsmI, CsmJ and CsmX, were characterized by insertional inactivation of their genes. Single mutants as well as all

74 combinations of double and triple mutants were constructed. Chlorosomes were isolated from each mutant, and comparisons of fluorescence quenching and energy transfer restoration were made for the wild-type and mutated chlorosomes. The results indicate that iron-sulfur proteins CsmI and CsmJ play important roles in the redox regulation of energy transfer by transferring electrons to and from the quencher (presumably chlorobiumquinone) within the chlorosomes. Because of the small amounts of CsmX in the chlorosome membrane, CsmX appears to play a negligible role in the redox regulation of energy transfer under the conditions tested.

75 3.3 Materials and methods

3.3.1 Molecular manipulation in Escherichia coli

Escherichia coli strains and growth conditions. E. coli DH5α [genotype F- ф80dlacZ - + - ΔM15 Δ(lacZYA-argF)U169 deoR recA1 endA1 hsdR17 (rK , mK ) supE44 λ thi-1 gyrA96 relA1 from Bethesda Research Laboratories, Gaithersburg, MD] and ElectroMAX DH10B [genotype F- mcrA Δ(mrr-hsdRMS-mcrBC) ф80dlacZ ΔM15 ΔlacX74 deoR recA1 endA1 araD139 Δ(ara, leu)7697 galU galK λ- rpsL nupG from Gibco BRL Products, Gaithersburg, MD] were used for recombinant DNA manipulations. The cells were grown at 37 °C in liquid cultures or on 1.5% (w/v) agar plates of Luria- Bertani medium containing 1% (w/v) bacto-tryptone, 0.5% (w/v) bacto-yeast extract, and 1% (w/v) NaCl, pH 7.0 (adjusted with NaOH). Cells containing plasmids were selected by addition of appropriate antibiotics at final concentrations of 100 μg ampicillin mL-1, 30 μg kanamycin mL-1, 10 μg chloramphenicol mL-1 or 10 μg gentamicin mL-1.

Plasmid isolation from Escherichia coli. Mini-preparations of plasmids from E. coli were performed using a modification of alkaline lysis protocol described by Birnboin and Doly (1979). E. coli cells harboring plasmids of interest (~5 mL) were pelleted and resuspended in 100 μL of GTE solution (50 mM glucose, 25 mM Tris-HCl, 10 mM EDTA, pH 8.0). Freshly made NaOH-SDS solution (200 μL) was added (0.2 M NaOH, 1% w/v SDS), and as soon as the suspension cleared, 150 μL of potassium acetate solution (3M K-acetate, 5% (v/v) formic acid, pH 5.5) was added. Cell debris was pelleted by centrifugation, and plasmid DNA in the supernatant was precipitated by adding 2.5 volumes of 95% ethanol or by adding 0.6 volume of isopropanol. Contaminating protein was extracted with equal volume of phenol:chloroform:isoamyl alcohol (25:24:1) if required. The DNA pellet was dried and resuspended in TE buffer

(pH 8.0) or sterile ddH2O. The alkaline SDS extraction method was also employed for large-scale preparations of plasmid DNA from 100-500 mL cultures. DNA concentrations were determined spectrophotometrically by measuring absorption at 260 -1 nm and using the relationship: dsDNA (μg mL ) = A260·50

76

Preparation of electro-competent cells. Cultures of E. coli DH10B (~500 mL) were grown in SOB medium (2% (w/v) bacto-tryptone, 0.5% (w/v) bacto-yeast extract, 10 mM NaCl, 2.5 mM KCl) at 37 °C with vigorous shaking until the OD550 nm was 0.8. The cells were chilled on ice, harvested by centrifugation at 4 °C and washed twice with an

equal volume of ice cold WB (10% (v/v) glycerol in ddH2O). The cells were finally resuspended in a small volume of WB, aliquoted and frozen in liquid nitrogen before storage at -80°C until required.

Transformation of Escherichia coli. Electro-competent cells were thawed on ice and mixed with 1-2 μL of plasmids or ligation products. The mixture of competent cells and DNA was transferred into an E-5011-1 0.1 cm Cuvette (ISCBioExpress, Kaysville, UT), placed in a micro-electroporation chamber and pulsed once with charging voltage of 1.50 kV. The pulsed cells were immediately diluted with 1 mL SOC (SOB plus 10 mM MgCl2,

10 mM MgSO4, 20 mM glucose) and incubated with shaking at 37 °C for 1 h prior to spreading on selective plates.

Restriction enzyme digestions and ligation. Restriction enzymes and T4 DNA ligase were purchased from New England Biolabs (Beverly, MA) or Promega Corporation (Madison, WI). Digestions and ligation were performed according to the directions of the enzyme supplier.

Electrophoresis and isolation of DNA from agarose gels. DNA fragments were separated on 0.8-1.5% (w/v) agarose gels by electrophoresis at constant voltage (100V) with Tris- acetate buffer (40 mM Tris, 20 mM acetic acid, 1 mM EDTA, pH 8.0). Gels were stained in an ethidium bromide solution and imaged on a UV light box using a Gel Print 2000i (Biophotonics, Ann Arbor, MI) imaging system. For isolation of DNA fragments, bands corresponding to the desired DNA fragments were excised from the gel and purified using Perfectprep Gel Cleanup Kit (Eppendorf AG, Hamburg, Germany) according to the instructions of the manufacturer.

77 Construction of plasmids pCI and pCJ. Three antibiotic markers were used in construction of csmI, csmJ and csmX mutants: gentamicin resistance cartridge, aacCI (GmR), from plasmid pMS255 (Becker et al., 1995); streptomycin and spectinomycin resistance cartridge, aadA (SmR, SpR), from plasmid pHP45Ω (Prentki and Krisch, 1984); and chloramphenicol and erythromycin resistance cartridge, cat-ermC (CmR, EmR), from plasmid pRL409 (Elhai and Wolk, 1988). For the last marker cat-ermC, the chloramphenicol-resistance marker is effective for selection in E. coli while the erythromycin-resistance marker is effective for selection in C. tepidum. The plasmids pET3d::csmI::Ω and pET3d::csmX::Ω were constructed by insertion of the aadA marker from pHP45Ω into the XcmI site of pET3d::csmI and the MfeI site of pET32a::csmX site within the coding sequences of the csmI or csmX genes (Frigaard et al., 2004a). The plasmids pCI was constructed by deletion of aadA marker from pET3d::csmI::Ω by BamHI digestion, and insertion of cat-ermC marker at the same BamHI site. The plasmid pCJ was constructed by insertion of aacC marker into the ApoI site of pET3d::csmJ. The EcoRI site (sequence GAATTC) on pET3d::csmJ was deleted before digestion with ApoI (recognition sequence (G/A)AATT(C/T)), which cuts within the csmJ coding sequence.

3.3.2 Mutant construction and confirmation in Chlorobium tepidum

Chlorobium tepidum strain and growth conditions. The WT2321 strain of Chlorobium tepidum, which is a plating strain derived from C. tepidum strain ATCC 49652, was used for all experiments (Wahlund and Madigan, 1995) (Wahlund et al., 1991). The medium for C. tepidum was the same as described in Chapter 2.3.1. Manipulations and small volume growth were performed in an anaerobic chamber (Coy Laboratory Products, Grass Lake, MI) at 42 ºC. Large-volume cultures were grown in completely filled, tightly sealed 2-L bottles in a water-bath at 42-45 ºC. For freeze storage, Chlorobium cultures were grown to the mid- or late- exponential growth phase. Autoclaved glycerol was added to the cell cultures to reach a final glycerol concentration of 10%, and the mixed solution was aliquoted into sterile screw-capped tubes for storage at -80°C. For revival, the frozen glycerol stock was melted on ice and 50 μL culture was inoculated into 1 mL of liquid medium. The stock

78 was transferred back to the -80 °C freezer immediately after inoculation. For growth rates studies, the cells were grown in 20 mL CL medium in screw-capped glass tubes on a radial rotator. The culture turbidity was continuously measured at 600 nm at different times and used to calculate the doubling time or growth rate.

Natural transformation of Chlorobium tepidum. The restriction enzyme AhdI was used to linearize the plasmids pET3d::csmI::Ω, pET32a::csmX::Ω, pCI and pCJ. Chlorobium tepidum was grown to late-exponential phase, and cells (1 mL) were harvested by microcentrifugation. The cells were resuspended in 100 μL of CL medium containing 1- 10 μg of linearized DNA, and spotted on a non-selective CP plates for 10-18 h incubation. The cell patch was then scraped off, streaked on selective CP plates and incubated for 5-6 days to allow transformant colonies to appear. After two rounds of streaking on selective CP plates, single colonies were inoculated into medium, and the dense cultures were used for genomic DNA isolation or freeze storage. The csmI, csmJ and csmX single mutants were first made by transformation of pET3d::csmI::Ω (EmR), pCJ (GmR) and pET32a::csmX::Ω (SmR, SpR) into wild-type C. tepidum cells. pCJ plasmid was subsequently used to transform the csmI and csmX mutants to construct the csmI csmJ and csmJ csmX double mutants. pCI was also used to transform the csmX mutant to obtain the csmI csmX double mutant. A triple mutant csmI csmJ csmX was constructed by transformation the csmI csmX double mutant with pCJ.

Total DNA isolation from Chlorobium tepidum. Small amounts of genomic DNA were prepared from 1 mL of dense Chlorobium tepidum cultures. The pelleted cells were resuspended in 50 μL TE, and lysed by addition of 0.5 mL GES reagent (5 M guanidinium isothiocyanate, 0.1 M EDTA, 0.5% (w/v) Na-sarkosyl, stored at room temperature in dim light). Cold 7.5 M ammonium acetate was added at a volume of 0.25 mL before an extraction with chloroform:isoamyl alcohol (24:1) was performed. The clarified aqueous phase was removed to a fresh microcentrifuge tube, and DNA was precipitated by addition of 0.6 volume of cold isopropanol. The pellets were washed twice with 70% (v/v) ethanol and dried under vacuum before being dissolved in ddH2O or TE buffer.

79

Polymerase chain reaction. Polymerase chain reaction (PCR) with primers of csmI, csmJ and csmX (sequences described in Vassilieva et al., 2001 and 2002b) was performed using an Eppendorf Gradient Mastercycler (Eppendorf Scientific Inc., Westbury, NY) to analyze the segregation of the transformants. Chlorobium tepidum genomic DNA (50- 500 ng) or plasmids (1-10 ng) served as templates, and dNTPs (0.2 mM each), forward and reverse primers (0.5 μM each), Taq polymerase (GeneChoice Inc., Frederick, MD) (2-5U) were present in the final reaction mixture. Reactions were carried out with a denaturation temperature of 95 °C, an extension reaction temperature of 72 °C, and various annealing temperatures between ± 5 °C of the calculated Tm of the primers. The program cycle was generally repeated 30-35 times.

3.3.3 Chlorosome isolation and SDS-PAGE

Chlorosomes were isolated on 7-47% (w/v) continuous sucrose gradients by ultracentrifugation at 220,000 × g for 18 h as described in Chapter 2.3.2. The chlorosome fraction was then pelleted twice using phosphate-buffered saline by ultracentrifugation at 240,000 × g for 1.5 h, and resuspended in a small volume of phosphate buffer with 1 mM PMSF and 2 mM DTT. The purified chlorosomes were aliquoted and stored at -80 ºC for future use. Chlorosomes proteins were analyzed on Tris-Tricine buffered SDS polyacrylamide gels and detected by silver staining or immunoblotting as described in Chapter 2.3.4. The antibody dilutions used were as follows: anti-CsmI, 1:5000; anti- CsmJ, 1:2500; anti-CsmX, 1:1000; and goat anti-(rabbit-IgG), 1:7000.

3.3.4 Pigment and quinone analysis

Pigments and quinones from chlorosomes were extracted by mixing aliquots of chlorosomes with acetone:methanol (7:2, v/v). The extracted mixture was centrifuged in a microcentrifuge, filtered through a 0.45 μm PTFE syringe filter (Whatman, Maidstone, UK) and mixed with 0.1 volume of 1 M ammonium acetate before immediate injection onto the HPLC. The HPLC system used was an Agilent 1100 series instrument (Agilent

80 Technologies, Palo Alto, CA) that included a solvent degasser, a binary pump and a diode-array spectrophotometer detector controlled with Agilent ChemStation software. The chromotography column was a 4.5 mm × 25 cm Discovery C18 column (Supelco, Bellefonte, PA). The elution solvents was methanol:acetonitrile:water 42:33:25 by volume (solvent A) and methanol:acetonitrile:ethyl acetate 50:20:30 by volume (solvent B). The elution program began with solvent B at 30% and included a linear increase of solvent B to 100% over 52 min. After 6 min elution with 100% B, the percentage of solvent B was decreased to the starting point during 2 min. The flow rate was 1.0 mL min-1 during the whole program. The pigment composition was determined by absorption spectroscopy using the following absorption coefficients: BChl c, 20 L g-1 cm-1 at 635 nm (Stanier and Smith, 1960); BChl a, 60 L g-1 cm-1 at 770 nm (Tsuji et al., 1995); carotenoids, 265 L g-1 cm-1 at 491 nm (Holo et al., 1985); chlorobiumquinones (1’- oxomenaquinone-7 and 1’-hydroxymenaquinone-7), 17 L g-1 cm-1 at 270 nm (Frydman and Rappaport, 1963); menaquinone-7, 26 L g-1 cm-1 at 270 nm (Dunphy and Brodie, 1971).

3.3.5 Fluorescence spectroscopy and cell viability test

Fluorescence quenching and restoration of chlorosomes. The fluorescence signals were measured with SLM-AMINCO 8100 Series 2 spectrofluorometer (SLM Instruments, Urbana, IL) at 15 ºC, with excitation light generated by ELXE-500 Xenon lamp at 460 nm (SLM Instruments, Urbana, IL). For the quenching experiments, chlorosomes were -1 diluted to 20 μg BChl c mL with anaerobic KH2PO4 buffer (10 mM KH2PO4, pH 7.0, saturated with nitrogen) containing 1 mM dithionite and held for 1 h for fluorescence recovery. The anaerobic chlorosomes were mixed with aerobic KH2PO4 buffer (saturated with air) using an SFA-20 rapid-kinetics stopped-flow accessory (Hi-Tech Limited, Salisbury, UK) within a silica observation cell. For fluorescence restoration experiments, -1 chlorosomes were diluted to 200 μg BChl c mL with aerobic KH2PO4 buffer and mixed with anaerobic buffer containing 12.5, 25 or 50 mM dithionite using the stopped-flow accessory. The fluorescence emission was recorded at the 772-nm emission maximum at time resolution of 0.5 s.

81

Fluorescence restoration of intact cells. C. tepidum cells in late-exponential growth

phase were pelleted by microcentrifugation and resuspended in aerobic KH2PO4 buffer and held in darkness for various times (from 1 to 72 h). The cells were pelleted and

resuspended in anaerobic KH2PO4 buffer containing or buffer supplemented with 2 mM 2- 2- S2O3 or S . Fluorescence restoration was performed under several conditions: darkness, low light (0.2 μE m-2 s-1) and room light (~50 μE m-2 s-1). Fluorescence emission measurements were performed for ~2 h as described above.

Cell viability test upon oxygen exposure. Cells in late exponential phase (OD600 nm = ~1.5) were diluted 100-fold in both anaerobic and aerobic KH2PO4 buffer. The anaerobic dilutions were wrapped in foil (to maintain dark conditions) and held in an anaerobic chamber for 3 and 6 days. The aerobic dilutions were placed in dark or room light conditions with the lids half-open for 3 and 6 days. The cells were then diluted 1×104- fold, and diluted cells (30 μL) were plated. Colonies were counted after 6 days incubation under standard growth conditions.

3.3.6 Reverse transcription PCR

Total RNA was extracted from 5-10 mL of a dense C. tepidum culture with GeneChoice RNA Spin Mini Kit (PGC Scientific Corporation, Frederick, MD) and further purified using an RNeasy Mini Kit (QIAGEN Inc., Valencia, CA). RNA (400 ng) was reverse- transcribed using M-MLV reverse transcriptase (Promega, Madison, WI) and random hexamer primers (Promega, Madison, WI) following the protocol of the manufacturer. cDNA samples were subjected to PCR amplification with csmI and csmJ primers (sequences described in Vassilieva et al., 2001) under the following conditions: denaturation at 94 ºC for 30 s, annealing at 57 ºC for 30 s, and elongation at 72 ºC for 1 min (27-33 cycles). A housekeeping gene, rnpB, encoding the ribonucleaseP B unit, was amplified simultaneously as a positive control. The PCR products were separated on a 1.5% (w/v) agarose gel and products were quantified by phosphor-imaging using ImageQuant software (Molecular Dynamics, Sunnyvale, CA).

82 3.4 Results

3.4.1 Construction and verification of mutants lacking CsmI, CsmJ and CsmX

The plasmids pET3d::csmI::Ω and pET32a::csmX::Ω were constructed as described in Frigaard et al. 2004a, and the other two plasmids pCI and pCJ were constructed as shown in Fig. 3-1. The plasmid sequences were confirmed by DNA sequencing. Wild-type C. tepidum and csmI, csmJ, and csmX mutant strains were transformed with linearized plasmids pCI and pCJ as described in the Materials and Methods. Transformants were selected and streaked two to three times to allow segregation of alleles to occur. PCR analyses with primers specific for csmI, csmJ and csmX suggested that the various antibiotic-resistant transformant strains were homozygous for the appropriate insertionally inactivated target genes (data not shown). Chlorosomes were prepared from each of the single, double and triple mutants, and their chlorosome proteins were analyzed by SDS-PAGE and detected by silver staining and immunoblotting (Fig 3-2). As shown in Fig. 3-2, Panel A, the levels of other chlorosome proteins (CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH) were unchanged in single mutants lacking CsmI (lane 2), CsmJ (lane 3), or CsmX (lane 4), in double mutants lacking CsmI and CsmX (lane5), CsmI and CsmJ (lane 6), or CsmJ and CsmX (lane 7), and in a triple mutant unable to synthesize CsmI, CsmJ, and CsmX (lane 8). Immunoblotting of chlorosome proteins with polyclonal antibodies to recombinant CsmI, CsmJ, and CsmX (Fig. 3-2, Panel B) confirmed the results from PCR analyses of the mutants. Chlorosomes isolated from the csmI mutant (Fig. 3-2, Panel B, lane 2) lacked CsmI but contained CsmJ and CsmX at levels similar to the wild-type chlorosomes (Fig. 3-2, Panel B, lane 1). Correspondingly, the chlorosomes from the csmJ mutant (Fig. 3-2, Panel B, lane 3) lacked CsmJ but contained CsmI and CsmX, and chlorosomes from the csmX mutant (Fig. 3-2, Panel B, lane 4) lacked CsmX but contained CsmI and CsmJ. Chlorosomes from the csmI csmX double mutant only contained CsmJ (Fig. 3-2, Panel B, lane 5); chlorosomes from the csmI csmJ double mutant only contained CsmX (Fig. 3-2, Panel B, lane 6); and chlorosomes from the csmJ csmX double mutant only contained CsmI (Fig. 3-2, Panel B, lane 7). As expected,

83 chlorosomes from the triple mutant lacked CsmI, CsmJ, and CsmX (Fig. 3-2, Panel B, lane 8).

3.4.2 Pigment and quinone contents

The pigment and quinone contents of isolated chlorosomes of the wild type and mutant strains, as measured by reverse-phase HPLC analyses, are shown in Table 3-1. When compared to the BChl c content, the contents of BChl a and carotenoids showed little variation compared to wild type, suggesting that the absence of iron-sulfur proteins in the chlorosome envelope does not cause a modification of the amounts of light-harvesting pigments inside the chlorosome. Chlorosomes of C. tepidum contain two major isoprenoid quinones, chlorobiumquinone and menaquinone-7, and small amounts of 1’- hydroxy-chlorobiumquinone are usually detected as well (Frigaard et al., 1997). The quinones in chlorosomes account for more than half of the total cellular quinones, and are present at an approximate ratio of 1:10 relative to BChl c (Frigaard et al., 1997). Although the total quinone contents of were similar for chlorosomes from the wild type and all mutant strains, the csmI csmJ and csmI csmJ csmX mutants contain significantly less chlorobiumquinone (~65% of the average of the others) and substantially more menaquinone-7 (~180% of the average of the others). This variation could possibly be due to subtle differences in the individual growth conditions and growth phase of the cultures used for chlorosome isolation, although standard growth conditions were employed for the production of all cell material. Another alternative interpretation of the results might be that the absence of the Fe/S proteins from the chlorosome envelope, especially the most abundant proteins CsmI and CsmJ, is directly responsible for the reduced levels of chlorobiumquinone in the chlorosomes of these two mutants. However, at this time it is not clear why the ratio of the two quinone species change so markedly in mutants lacking both CsmI and CsmJ.

84 3.4.3 Fluorescence quenching and restoration

Fluorescence quenching and restoration of chlorosomes. Fig. 3-3 shows the time-course of fluorescence quenching for isolated chlorosomes when anaerobic chlorosomes (200 μg BChl c mL-1) were mixed with air-saturated (aerobic) buffer. Fig. 3-3, Panel A shows the fluorescence decay during the first 500 s for chlorosomes of the wild type and of three single mutants, csmI, csmJ, and csmX. Fig. 3-3, Panels B and C show the fluorescence quenching for chlorosomes of the wild type as well as of the double and triple mutants csmI csmX, csmJ csmX, csmI csmJ and csmI csmJ csmX. As shown in the figures, fluorescence emission from chlorosomes rapidly decreased for the wild type and all of the mutants. These data show that the Fe/S proteins are not required for the formation of

the quencher. However, the fold-decrease in the fluorescence emission (i.e., Fmax:Fmin, the ratio of the maximum fluorescence emission and the minimum fluorescence emission) is

different for the wild type and the mutants. At the end of 500 s, the Fmax:Fmin ratio of wild

type chlorosomes is greater than 40, while the Fmax:Fmin ratios of the single and double mutants (except csmI csmJ) are around 20. The mutants of csmI csmJ and csmI csmJ

csmX, having Fmax:Fmin ratios of about 10 and the fluorescence decay kinetics from these two mutants are notably slower compared with wild type (Panel C). The decreasing rates are also different among these samples. For wild-type chlorosomes, it took ~130 s for the

fluorescence to decay to an Fmax:Fmin value of 10. However, it took 180 to 250 s for the single and double mutants (except the csmI csmJ mutant) to be quenched to this same

extent, and it took ~360 s to reach an Fmax:Fmin value of 10 for the csmI csmJ and csmI csmJ csmX mutants. Since the mutants lacking the CsmI and CsmJ proteins have the most distinguishable quenching phenotype, it seems likely that CsmI and CsmJ play some role in the fluorescence quenching process and that these proteins are at least partly responsible for the rapid and nearly complete complete fluorescence decay observed for wild-type chlorosomes. However, it is also notable that the amounts of chlorobiumquinone in the csmI csmJ and csmI csmJ csmX mutants were significantly decreased relative to the other mutants (~65% of the average). Chlorobiumquinone is generally considered to be the quencher within the chlorosome (Frigaard et al., 1997). Redox regulation of fluorescence and energy transfer was observed in chlorosome-like

85 BChl c aggregates upon the addition of exogenous chlorobiumquinone (Frigaard et al., 1997). Moreover, the chlorobiumquinone contents in various C. tepidum mutants lacking a single chlorosome protein are inversely related to the fluorescence emission intensity under oxidizing conditions (Frigaard et al., 2004a). Thus, the chlorobiumquinone contents might be another important feature affecting the rate and magnitude of the fluorescence decay. Fig. 3-4 shows the time-course of fluorescence emission recovery of isolated chlorosomes when the chlorosomes under oxic conditions (air-saturated buffer) were mixed with nitrogen-saturated anoxic buffer in the presence of different concentrations of dithionite. At low dithionite concentrations (12-50 mM), the rate of fluorescence recovery is roughly linear during the first 200 s. Panels A-C of Fig. 3-4 also show that the fluorescence recovery of chlorosomes occurs faster at higher dithionite concentrations. For example, the fluorescence emission of wild type chlorosomes increased at 28 unit s-1 at 12.5 mM dithionite, 39 unit s-1 at 25 mM dithionite, and 42 unit s-1 at 50 mM dithionite. The fluorescence emission of the csmI csmJ chlorosomes increased at rates of 3 unit s-1 at 12.5 mM dithionite, 6 unit s-1 at 25 mM dithionite, and 8 unit s-1 at 50 mM dithionite. The rates of fluorescence emission recovery for chlorosomes from the csmI csmJ csmX mutant had similarly increased rates: 3 unit s-1, 5 unit s-1, and 7 unit s-1 at12.5, 25 and 50 mM dithionite, respectively. Panels D-F and Panels G-I separately compare the fluorescence recovery of wild-type chlorosomes with that for all of the mutants under 12.5 and 25 mM dithionite. The greatest difference between the results for the two dithionite concentrations is that rates of increase at 12.5 mM dithionite are about 50% slower than those at 25 mM dithionite. If chlorosomes were incubated for a longer time (e.g., 500 s instead of 250 s), the fluorescence restoration under 12.5 mM dithionite was nearly identical to that under 25 mM. Because the fluorescence recovery rates are essentially linear at 12.5 to 25 mM dithionite, the rates have been tabulated and compared as percentages of the wild-type rate in Table 3-2. The data in Fig. 3-4 and Table 3-2 show that the csmI csmJ and csmI csmJ csmX mutants have the greatest defects: their fluorescence recovery rates are only about 10% of that for wild-type chlorosomes. The recovery rates of the csmI csmX and csmJ csmX double mutants are about 35% of that for wild-type chlorosomes, while the recovery rates of chlorosomes from the csmI

86 and csmJ single mutants are higher (~45% and ~70%). The absence of CsmX had the smallest effects on the rate of fluorescence recovery, which suggests that this protein plays only a minor role in the fluorescence recovery process. However, the presence of either CsmI alone or CsmJ alone is sufficient to allow the fluorescence recovery process to occur at about 30% of the rate observed for wild-type chlorosomes. The differences in the rates of fluorescence recovery observed for chlorosomes from the single, double and triple mutants greatly exceeded the differences in their chlorobiumquinone contents. Thus, CsmI and CsmJ play important roles in reactivating energy transfer (inactivating fluorescence quenching) in chlorosomes.

Fluorescence restoration of cells. Numerous studies of the behavior of fluorescence quenching in whole cells of the wild-type and mutant strains were made under a variety of conditions. Chlorosome fluorescence emission was very rapidly quenched when wild- type cells or any of the mutant cells were diluted into oxic buffers. The quenching process was so rapid that differences among the various strains could hardly be discerned. On the other hand, differences in the rates of fluorescence emission recovery (quencher inactivation) of whole cells of the wild type and mutant strains were more easily observed. Cellular fluorescence emission decreased from a maximum value of ~130 units

(at a cell concentration equal to OD600 nm = 0.100) to its minimum value of ~6-8 after the cells were resuspended in oxic buffer for only 15 min. However, fluorescence emission recovered fully with similar rates under dark conditions when no external reductants were added after 1 h of air-exposure for cells of the wild type or the csmI csmJ csmX mutant. This observation suggested that there might a cellular reductant pool (either a storage polysaccharide or some reduced sulfur compound(s)) that could provide the electrons to reduce the quencher within the chlorosome once oxygen stress is removed. Thus, the oxygen-exposure time was increased to 72 h in an attempt to decrease the cellular reductant pool and reduce this background restoration. Fig. 3-5 shows the time course of fluorescence emission recovery with or without light and reductant for wild type and the csmI csmJ csmX mutant after 72 h of oxygen exposure. As shown in Fig. 3-5, Panel A, without light and added external reductant, no fluorescence emission recovery was observed for the wild type or the csmI csmJ csmX

87 mutant. Fig. 3-5, Panel B shows the fluorescence emission recovery in the dark when 2 mM thiosulfate was added as an electron donor. Surprisingly, the triple mutant recovered 35% of the maximal fluorescence value during the period of observation and recovered more rapidly and to a greater extent than the wild type. When 2 mM sulfide was provided as the electron donor, similar results were observed. The triple mutant recovered more rapidly and to a greater extent than the wild type (data not shown). Fig. 3-5, Panel C shows the fluorescence emission recovery under a very low light intensity (0.2 μmol photons m-2 s-1) with no added external reductant. During the first 0.5 h, the fluorescence recovery of the wild type and csmI csmJ csmX mutant remained similarly slow and minimal, but subsequently, the wild-type recovered over the next 1.5 h to 82% of the maximal value, while the mutant continued to recover more slowly and only recovered ~30% of the maximal emission value. Fluorescence recovery experiments were also performed without added reductants under room light (~50 μmol photons m-2 s-1). The recovery rates for both the wild type and the csmI csmJ csmX mutant were faster than under the very low light intensity, and the difference between the two types of cells was less significant. The wild type recovered about 30% faster than the mutant (data not shown). Finally, Fig. 3-5, Panel D shows the restoration of the wild type and the csmI csmJ csmX mutant with both 2 mM thiosulfate as electron donor and very low light intensity (0.2 μmol photons m-2 s-1). The most rapid recovery of the fluorescence emission occurred under these conditions, but the results for the wild type and the csmI csmJ csmX mutant were nearly identical. In both cases, the cells recovered about 82-84% of the maximal fluorescence emission value, and the lag times and rates of recovery were essentially identical.

3.4.4 Growth rates and viability after aerobic exposure

Growth of the mutants lacking CsmI, CsmJ and CsmX. Growth rates of the wild type and all of the mutant strains were tested under three light intensities: 8 μmol photons m-2 s-1, 30 μmol photons m-2 s-1, and 150 μmol photons m-2 s-1 (Table 3-3). Except for the csmI csmJ mutant, the growth rates of the mutants were only slightly slower (5-15%) than that of the wild type. The first csmI csmJ mutant strain that was characterized did not grow at

88 high light intensity and grew more slowly (50% of the rate of wild type) under very low light intensity. Further studies showed that this strain of csmI csmJ is both light-sensitive and temperature-sensitive (data not shown). In contrast to the wild type, which grows optimally at 47-48 ºC, the optimal growth temperature of this csmI csmJ strain was only 39-40 ºC. High light intensities (>60 μmol photons m-2 s-1 at 45 ºC) significantly decreased the growth rate of this csmI csmJ strain even more. However, two other independently constructed csmI csmJ strains had a less severe phenotype than this initial strain. Their growth rates were 50-60% of the wild-type rate at high light intensity and at the normal growth temperature (growth rates of one of these strains is shown in Table 3- 3). It is important to note that the csmI csmJ csmX mutant has a similar growth rate similar to that of the wild type. Thus, the phenotype of the first characterized csmI csmJ strain is more likely to be the consequence of some random, secondary mutation than due to the inactivation of CsmI and CsmJ.

Cell viability after aerobic exposure. Cells of the wild type and the csmI csmJ csmX mutant were exposed to air-levels of oxygen for 3 and 6 days under dark or light conditions, and cells were then plated on CP plates to determine cell viability as colony forming units. An aliquot of the same cells were kept under dark anaerobic conditions for 3 days or 6 days as the control sample for this experiment. After 3 days of air-exposure in darkness, 96% of the wild type cells and 92% of the csmI csmJ csmX mutant cells were still alive and produced colonies. The percentage of viable cells decreased further to 88% for wild type and 86% for the triple mutant after 6 days of air-exposure. After 3 days aerobic treatment under room light, the 31% of the wild type cells were still viable, about twice the percentage for the csmI csmJ csmX mutant (16%). No colonies were detected for either strain after 6 days of air-exposure under light. These data demonstrate that the csmI csmJ csmX mutant is more sensitive to oxygen than the wild type when light is provided. The cells treated by air and room light for 3 days were also applied to 2- fluorescence restoration experiments with 2 mM S2O3 and 0.2 μmol photons light provided. The fluorescence of both wild type cells and the triple mutant cells could only recover to ~20% of the maximum after 8000 s, which indicates that the light-harvesting and energy transfer system were seriously damaged by oxygen in combination with light.

89

3.4.5 mRNA and protein level of CsmI and CsmJ in the csmX mutant

Redox titrations of chlorosomes from the csmX mutant suggested that the CsmI:CsmJ protein ratio had increased to 2:1 instead of the 1:1 ratio typically seen in wild-type chlorosomes (Johnson et al., unpublished data). This suggested that CsmX might play some regulatory or stabilizing role in determining the amounts of CsmI and CsmJ present in chlorosomes. To investigate the effects of the csmX mutation on transcription, RT- PCR with primers specific for csmI and csmJ was performed (Fig. 3-7 Panel A). The product levels for the csmI and csmJ amplicons were slightly greater after 27, 30, and 33 cycles in the csmX mutant than in the wild type. The csmJ amplicon was about 1.2-fold more abundant for the mutant, and the csmI amplicon was about 1.3-fold more abundant for the mutant than for the wild type. Immunoblotting analyses of chlorosomes isolated from the wild type and the csmX mutant showed that CsmI and CsmJ contents were similar in the two samples of chlorosomes. These results indicate that CsmI and CsmJ should be present in similar amounts in the chlorosomes of the csmX mutant. Since CsmJ appears to be more sensitive to proteolytic degradation than CsmI, it is possible that the altered protein ratio implied from the redox titration results may have resulted from partial degradation of CsmJ in the chlorosomes used for the redox titration experiments.

90 3.5 Discussion

The moderately thermophilic green sulfur bacterium C. tepidum was originally isolated from slightly acidic, high-sulfide hot springs in New Zealand (Wahlund et al., 1991). As obligately anaerobic photoautotrophs that commonly live in extremely light-limited environments, the Chlorobia have developed distinctive mechanisms of photosynthesis and metabolism (Eisen et al., 2002). These bacteria harvest light energy with unique photosynthetic antenna complexes, chlorosomes, which contain mainly BChl c and carotenoids and which transfer the absorbed light energy to Type I, homodimeric reaction centers to drive ATP synthesis and reducing power production (Hauska et al., 2001). Carbon fixation in the Chlorobi occurs by the reductive tricarboxylic acid (TCA) cycle instead of Calvin cycle, which occurs in other purple photosynthetic bacteria, cyanobacteria and higher plants (Evans et al., 1966; Sirevåg, 1974; Buchanan and Arnon,

1990). Acetyl-CoA is synthesized by reduction of CO2 using electrons derived from hydrogen, reduced sulfur compounds, or rarely ferrous iron. Numerous [4Fe-4S] ferredoxins with very low redox potentials are involved in these unique photosynthetic and metabolic pathways. The PscB subunit in the reaction centers of green sulfur bacteria harbors two [4Fe-4S] clusters analogous to FA and FB in PS I in oxygenic phototrophs, and these serve as the terminal electron acceptors (Vassiliev et al., 2001). One of these two clusters was found to have a midpoint potential of -550 mV in Chlorobium limicola (Knaff and Malkin, 1976; Jennings and Evans, 1977).

Midpoint potentials of FA and FB were later reported to be -502 mV and -450 mV in isolated reaction centers from Chlorobium vibrioforme (Scott et al., 1997). Evidence for

a third [4Fe-4S] cluster analogous to FX was identified by peptide sequence similarity and EPR spectroscopic studies (Büttner et al., 1992a; Vassiliev et al., 2001). Strongly reducing electrons generated by reaction centers using light energy are delivered to the enzymes of the reductive TCA cycle for carbon fixation by soluble ferredoxins. Two novel 2[4Fe-4S] ferredoxins with redox potentials of -514 and -584 mV were recently isolated from Chlorobium tepidum and shown to be electron donors to crucial enzymes in carbon fixation, α-ketoglutarate synthase and pyruvate synthase (Yoon et al., 1999; Yoon et al., 2001). They appear more electronegative than any previously studied ferredoxins

91 in which the two [4Fe-4S] clusters display a redox potential value (Smith and Feinberg, 1990; Camba and Armstrong, 2000; Yoon et al., 2001). The highly electronegative cytoplasm of green sulfur bacteria determines that these bacteria can only survive in strictly anoxic conditions. Green sulfur bacteria in nature typically occur in anoxic aquatic layers or sediments rich in reduced sulfur compounds (Wahlund et al., 1991). However, this does not mean that the cells are never challenged by chance encounters with oxygen. For example, vertical mixing of oxic and anoxic water layers could cause green sulfur bacteria to encounter oxic conditions, and scenarios could likewise cause cells to come in contact with oxygen (Frigaard and Matsuura, 1999). The C. tepidum genome encodes several oxygen protection enzymes, including superoxide dismutase, rubredoxin oxygen oxidoreductase and cytochrome bd quinol oxidase (Eisen et al., 2002). Even if only trace amounts of oxygen were transiently present, the highly reduced electron transfer proteins in the C. tepidum cytoplasm would rapidly react with molecular oxygen to generate superoxide and hydrogen peroxide as products, which are stronger oxidants than oxygen and which are extremely damaging to metabolic enzymes and ferredoxins (Imlay, 2002). One of the principal protection mechanisms against oxygen stress in C. tepidum is that energy transfer in the chlorosomes can be effectively regulated by redox potential (Blankenship et al., 1993). The energy transfer efficiency in the chlorosomes is reduced from nearly 100% to less than 10% under oxidizing conditions, which can easily be observed as a large drop in BChl c fluorescence emission and by a decreased lifetime for the excited state of BChl c. Thus, the photosynthetic apparatus is effectively protected by suppressing the activity of reaction centers, by decreasing the level of reduction of ferredoxins, and by minimizing the formation of reactive oxygen species. Energy transfer in isolated chlorosomes can be reversibly restored with 100% efficiency by the addition of reducing reagents such as sodium dithionite (Wang et al., 1990). Chlorosome proteins CsmI, CsmJ and/or CsmX share obvious sequence similarity to adrenodoxin-type [2Fe-2S] ferredoxins, a family of small soluble iron-sulfur proteins that transfer electrons from ferredoxin reductases to cytochromes P450 in mammalian mitochondria and some bacteria (Müller et al., 1998; Grinberg et al.., 2000; Sevrioukova et al., 2003). Mammalian adrenodoxin, bacterial putidaredoxin and bacterial terpredoxin

92 are all classified as the adrenodoxin-type ferredoxins (Chang et al., 1988; Peterson et al., 1990; Mo et al., 1999). Structure studies of adrenodoxin-type ferredoxins suggest that the N-terminal and the C-terminal half of the polypeptide sequences form two “lobes” and have shown that the iron-sulfur cluster is located between these lobes near one end of the molecule (Kostic et al., 2002). Each iron is coordinated tetrahedrally by two acid-labile

sulfurs and two sulfurs from cysteine residues within the cluster binding motif C-X5-C-

X2-C-X35-37-C (Kostic et al., 2002). The N-termini of chlorosome proteins CsmI, CsmJ and CsmX contain an identical cluster-binding motif except that there are only 34 residues between the third and the fourth cluster-liganding cysteine residues (Vassilieva et al., 2001). The EPR properties of these three [2Fe-2S] proteins were recently studied in wild type chlorosomes and in chlorosomes containing a single [2Fe-2S] protein isolated from appropriate mutant strains (Johnson et al., unpublished data). Purified chlorosomes containing only CsmI or CsmJ displayed axial spectra with distinctive g-values for the midfield derivative peaks (1.947 for CsmI and 1.940 for CsmJ). The amount of CsmX in the chlorosomes is much lower (~5%) than CsmI and CsmJ (Vassilieva et al., 2002b), and CsmX could not be directly observed by EPR spectroscopy. The contribution of CsmX to the EPR spectral properties is negligible in wild-type chlorosomes, and combining the spectra of CsmI and CsmJ in a 1:1 ratio accurately simulates the EPR spectrum of chlorosomes. The redox potential for CsmI was determined to be -346 mV in wild-type chlorosomes and -204 mV in chlorosomes from the csmJ csmX mutant, which only contain CsmI. The redox potential of CsmJ was determined to be + 90 mV in wild- type chlorosomes and -3 mV in chlorosomes from the csmI csmX mutant, which only contain CsmJ. These redox potential shifts are likely to result from the different multimerization patterns for these [2Fe-2S] proteins. The cross-linking studies presented in Chapter 2 indicate that CsmI forms CsmI2CsmJ2 heterotetramers in wild-type

chlorosomes and CsmI2 homodimers in the csmJ csmX mutant. The redox potentials of CsmJ in chlorosomes are much more oxidizing than the redox potential of CsmJ in inclusion bodies (-194 mV) (Vassilieva et al., 2001). Moreover, these values are certainly out of the typical redox potential range of adrenodoxin-type ferredoxins, which usually vary from -235 to -273 mV (Grinberg et al., 2000). The iron-sulfur clusters in the Fe/S

93 proteins of chlorosomes are stable to oxygen. These proteins are not inactivated by oxygen even in the presence of the strong chaotrope, 2M sodium thiocyanate. The EPR properties of chlorosomes prepared aerobically are identical to the properties of chlorosomes isolated and kept under anoxic conditions (Vassilieva et al., 2001). These observations indicate that the Fe/S proteins of the chlorosome envelope are well suited to play an important role under fully oxic conditions, since they will retain their correct conformation and function even under oxygen-stress conditions. Analyses of the restoration of fluorescence emission with isolated chlorosomes after addition of dithionite showed clearly that CsmI and CsmJ play essential roles in inactivating the quencher and in the recovery of energy transfer and fluorescence emission. When either CsmI or CsmJ is missing from the chlorosome envelope, the recovery rate for fluorescence emission decreases to 30-70% that of wild type. When both CsmI and CsmJ are missing in the chlorosomes, the recovery rate was only ~10% that of wild type. In this restoration process, the dithionite acted as the electron donor, and electrons were presumably transferred to the quencher within the chlorosomes by CsmI and CsmJ. Fluorescence recovery was also observed with whole cells whose fluorescence had been quenched to minimal levels by suspending cells in aerobic buffer for minutes. Interestingly, the fluorescence could recover in both the wild type as well as the mutant lacking all three [2Fe-2S] chlorosome proteins, even when no light or external reductant was provided. In such cases, it is likely that a polysaccharide storage material within the cells provides the electrons required for reduction of the quencher within the chlorosomes. The most common polysaccharide storage in C. tepidum is glycogen (Sirevåg, 1995), a polymer in which numerous residues are joined together by α-(1, 4)- glucosidic linkages into long chains and α-(1, 6) linkages into branches (Candy, 1980). It was observed that glycogen serves as a source of reducing power when the cells were incubated without a hydrogen donor, either in the dark or in the light (Sirevåg and Ormerod, 1977). To completely consume the intracellular reductant pool, the C. tepidum cells were exposed to air for 3 days. The cells were still alive after a 3-day air exposure, but the endogenous restoration of fluorescence emission was no longer observed. The fluorescence emission still recovered when cells were illuminated or when external reductants such as thiosulfate or sulfide were added. Notably, the wild type recovered

94 much more rapidly than the mutant lacking all [2Fe-2S] chlorosome proteins when only light was provided. Thus, electron transfer through the [2Fe-2S] proteins of the chlorosome envelope is probably a light-dependent (reaction center-dependent) process. Elemental sulfur and polysulfide, the intermediates of thiosulfate and sulfide oxidation, which are attached to the exterior of cells, might serve as electron donors under such conditions. Electrons might be transferred to the quencher through the reaction center and then to CsmI/CsmJ/(CsmX). When only external reductants are provided, the restoration of fluorescence emission in mutant cells lacking all [2Fe-2S] proteins in the chlorosome envelope was even faster than for the wild-type cells. Since no light was provided, the reaction centers did not participate in the transfer of electrons to the quencher under these conditions. These results suggest that at least two electron transfer pathways to the quencher exist: one for external reductants that is independent of reaction centers, and one that is light-dependent and requires the participation of the reaction centers. These two possibilities are illustrated in Fig. 3-6. The results for investigations of fluorescence emission recovery described above showed that the defect of the csmI csmJ csmX mutant in the restoration process is not observed, or may not be very significant under some conditions (Fig. 3-5, Panels B and D). However, as shown in Fig. 3-5 Panel C, the superiority of wild type is easily demonstrated under some conditions. These results differ from those obtained for fluorescence emission recovery of oxygen-treated chlorosomes. In those studies, the mutants showed the greatest phenotype under all experimental conditions tested. Therefore, the electron transfer pathways are likely to be different in the case of fluorescence emission restoration in cells and in fluorescence emission restoration of chlorosomes. In the experiments with isolated chlorosomes, the electrons are probably transferred from the reductant (dithionite) directly to CsmI/CsmJ/(CsmX) proteins, and then to the quencher, and this might represent the only pathway for reduction of the quencher other than bulk chemical reduction of the quencher at high dithionite concentrations. In the cellular fluorescence emission recovery experiments, the electrons might be transferred through the CsmI/CsmJ/(CsmX) to the quencher, or they might be transferred to the quencher by other pathways, which also exist and compete with the one that includes CsmI/CsmJ/(CsmX). The greatest difference between the wild type and the

95 csmI csmJ csmX mutant was obtained under very low light intensity (0.2 μmol photons m-2 s-1) indicates that the electron transfer through CsmI/CsmJ/(CsmX) might be a light- dependent process, i.e., a reaction-center dependent process. Fig. 3-6 presents a model that includes two different electron transfer pathways in cellular fluorescence restoration. If only light but no external reductant is provided, electrons presumably are transferred from internal or external residual storage materials through the reaction centers via CsmI/CsmJ/(CsmX) to the quencher. Elemental sulfur/polysulfide attached to the outside of the cells and residual storage carbohydrates might serve as the electron donors in this case (Fig. 3-5, Panel C). From the results in Fig. 3-5, light greatly stimulates the energy transfer (fluorescence emission) recovery, but the csmI csmJ csmX mutant requires the addition of an external reductant (Fig. 3-5, Panel D) while the wild type does not (Fig. 3- 5, Panel C). One possibility is that in cells, quinones can shuttle between the cytoplasmic membrane and the chlorosomes. Reduced menaquinone-7 might be capable of reducing the residual chlorobiumquinone or might act as a quencher, albeit a less efficient one. The suggestion that the alternative electron transfer pathway in Fig. 3-5 involves quinone shuttling is supported by the observation that chlorosomes of the csmI csmJ and csmI csmJ csmX mutants contain ~two-fold more menaquinone than other chlorosomes, although the total quinone contents of the chlorosomes from these two mutants are similar to those of the wild type and other mutants (Table 3-1). The existence of energy-transfer (fluorescence) quenchers in chlorosomes was proposed in early 1990’s and proved by redox titration of fluorescence emission (Wang et al., 1990; Blankenship et al., 1993). These quenchers are most likely to be quinones, which have been shown to quench chlorophyll fluorescence by accepting an electron from excited-state chlorophyll and by then performing extremely rapid charge recombination to restore the chlorophyll ground state (Natarajan and Blankenship, 1983). Chlorosomes of green sulfur bacteria contain large amounts of a unique isoprenoid quinone, chlorobiumquinone (1’-oxomenaquinone-7), which has a rather oxidizing redox potential of +36 mV (Powls and Redfearn, 1969; Frigaard et al., 1997). In C. tepidum cells, chlorobiumquinone is mostly localized in the chlorosomes, where it constitutes about 70% of the total quinones of the chlorosome (the other 30% is mainly menaquinone-7) (Frigaard et al., 1997). The ratio of chlorobiumquinone to BChl c is

96 approximately 1:15 (Frigaard et al., 1997). The quinone content of chlorosomes is not affected by SDS treatment, which indicates that the quinones are probably located in the interior of the chlorosome rather than in the chlorosome envelope (Frigaard et al., 1998; Bryant et al., 2002). When quinones were added to artificial, chlorosome-like BChl c aggregates in an aqueous solution, the fluorescence and energy transfer properties of the artificial aggregates were highly dependent upon the redox potential of the solution, and chlorobiumquinone was the most effective quinone in quenching energy transfer and fluorescence (Frigaard et al., 1997). When chlorobiumquinone and menaquinone-7 were extracted from the isolated chlorosomes with hexane, the redox dependence of the energy transfer and fluorescence emission was no longer detected (Frigaard et al., 1998). Chlorosomes from the green filamentous bacterium Chloroflexus aurantiacus only contain menaquinone-10 and do not exhibit redox-dependent energy transfer regulation (Wang et al., 1990). Given all of the above evidence, it has been concluded that chlorobiumquinone plays a significant role in quenching the fluorescence emission and in inhibiting the energy transfer in chlorosomes in C. tepidum under oxic conditions (Frigaard et al., 1997; Frigaard et al., 1998; Tokita et al., 2000). By studying the quenching of BChl c fluorescence emission using many exogenously added quinones, Tokita et al. (2000) found evidence to suggest that the 1’-oxo group chlorobiumquinone might be essential to its behavior as a highly effective fluorescence quencher (Tokita et al., 2000).

97 Table 3-1. Pigment and quinone contents of chlorosomes isolated from the wild-type strain and the mutants lacking CsmI, CsmJ and CsmX. The amounts of pigments and quinones were shown in mg per g of BChl c. All values are the averages and standard deviations of at least two measurements for two separate chlorosome preparations from different cell cultures.

BChl a Carot- Chlorobium Mena- Total enoids -quinones quinone-7 Quinones wild type 11±1 57±7 54±4 10±2 64±6 csmI 12±1 70±3 46±7 17±5 63±12 csmJ 15±4 64±2 52±9 14±2 66±11 csmX 12±3 67±2 51±11 12±0 63±11 csmI csmX 11±1 56±6 60±11 13±5 73±16 csmJ csmX 13±0 57±2 66±10 14±4 82±14 csmI csmJ 14±2 52±5 36±8 25±5 61±13 csmI csmJ csmX 14±1 53±3 38±8 28±7 66±15

98 Table 3-2. Chlorosome fluorescence emission recovery with 12.5 mM and 25 mM dithionite. The results are reported as the rate of recovery and as the percentage of the wild-type recovery rate.

Rate (unit s-1) Percentage of Rate (unit s-1) Percentage of of fluorescence wild-type rate of fluorescence wild-type rate recovery at 12.5 recovery at 25 mM dithionite mM dithionite wild type 28±2 --- 39±3 --- csmI 12±2 43% 20±3 51% csmJ 20±2 71% 27±2 69% csmX 27±3 96% 37±3 95% csmI csmX 9±2 32% 13±2 33% csmJ csmX 10±1 36% 15±2 38% csmI csmJ 3±1 11% 6±2 15% csmI csmJ csmX 3±1 11% 5±2 13%

99 Table 3-3. Growth rates of the wild type and mutants lacking CsmI, CsmJ and CsmX under three light intensities.

Growth Light Intensity (μmol photons m-2 s-1) Strain 8 30 150 wild type 0.047±0.004 0.203±0.007 0.277±0.008 csmI 0.039±0.005 0.203±0.005 0.263±0.010 csmJ 0.039±0.002 0.170±0.005 0.236±0.005 csmX 0.044±0.004 0.193±0.006 0.275±0.004 csmI csmX 0.041±0.003 0.174±0.012 0.248±0.007 csmJ csmX 0.043±0.003 0.168±0.007 0.229±0.006 csmI csmJ 0.040±0.004 0.168±0.005 0.163±0.007 csmI csmJ csmX 0.042±0.005 0.174±0.004 0.254±0.009

100

Fig. 3-1. Restriction maps showing the structures of the gene inactivation constructs.

101

Fig. 3-2. Protein composition analysis of chlorosomes isolated from the wild-type and the single, double and triple mutants lacking CsmI, CsmJ, and CsmX. Panel A. SDS-PAGE with proteins detected by silver staining. Proteins corresponding to 15 µg of BChl c were loaded on each lane. The electrophoretic positions of the various chlorosome (Csm) proteins are indicated at the left of the figure by single letters for their gene loci. Panel B. Immunoblots probed with anti-CsmI, anti-CsmJ and anti-CsmX polyclonal antibodies. Lane1, wild-type chlorosomes; Lanes 2-8, chlorosomes from the csmI, csmJ, csmX, csmI csmX, csmI csmJ, csmJ csmX, and csmI csmJ csmX mutants, respectively. Proteins corresponding to 200 µg of BChl c were loaded on each lane.

102

Fig. 3-3. Fluorescence emission quenching in isolated chlorosomes. Each sample contained chlorosomes corresponding to a BChl c concentration of 20 μg mL-1. Fluorescence emission was recorded at 772 nm with a time resolution of 0.5 s. Panel A. Chlorosomes from the wild type (black line), the csmI mutant (red line), the csmJ mutant (green line), and the csmX mutant (blue line). Panel B. Chlorosomes from the wild type (black line), the csmI csmX mutant (red line), and the csmJ csmX mutant (green line). Panel C. Chlorosomes from the wild type (black line), the csmI csmJ mutant (red line), and the csmI csmJ csmX mutant (green line).

103 Fig. 3-4. The fluorescence emission recovery in isolated chlorosomes. Chlorosomes corresponding to a BChl c concentration of 200 μg mL-1 were used in each experiment. Fluorescence emission was recorded at 772 nm with a time resolution of 0.5 s. Panel A-C. Fluorescence emission recovery of chlorosomes from the wild type (Panel A), the csmI csmJ mutant (Panel B) and the csmI csmJ csmX mutant (Panel C) at 12.5 mM dithionite (black lines), 25 mM dithionite (red lines) and 50 mM dithionite (green lines). Panels D-F. Fluorescence emission recovery for various chlorosome samples in the presence of 25 mM dithionite. Panel D. Chlorosomes from wild type (black line) and the csmI (red line), csmJ (green line), and csmX mutants (blue line). Panel E. Chlorosomes from wild type (black line) and the csmI csmX (red line) and csmJ csmX mutants (green line). Panel F. Chlorosomes from the wild type (black line) and the csmI csmJ (red line) and csmI csmJ csmX mutant (green line). Panels G-I. Fluorescence emission recovery for various chlorosome samples in the presence of 12.5 mM dithionite. Panel G. Chlorosomes from the wild type (black line) and csmI (red line), csmJ (green line), and csmX mutants (blue line). Panel H. Chlorosomes from the wild type (black line) and the csmI csmX (red line) and the csmJ csmX mutants (green line). Panel I. Chlorosomes from the wild type (black line) and the csmI csmJ (red line) and the csmI csmJ csmX mutants (green line).

104

105

Fig. 3-5. Fluorescence emission recovery in whole cells of wild type (black line) and the csmI csmJ csmX mutant (red line). Fluorescence emission was recorded at 772 nm at various times with a resolution of 1 s.

Cells corresponding to an OD600 nm = 0.100 were used for all measurements. Panel A. Recovery in the dark with no external reductant added. Panel B. Recovery in the dark in the presence of 2 mM thiosulfate 2- (S2O3 ). Panel C. Recovery in the presence of very weak light (0.2 μmol photons m-2 s-1) with no external reductant added. Panel D: Recovery in the presence of very weak light (0.2 μmol photons m-2 s-1) light in the 2- presence of 2 mM thiosulfate (S2O3 ).

106

Fig. 3-6. Model for electron transfer during recovery fluorescence emission in cells. Panel A. Electrons are transferred via reaction centers and CsmI/CsmJ/(CsmX) to the quencher only when light is provided. Panel B. Electrons are transferred from external reductants, thiosulfate and sulfide, to the quencher via some unknown cellular component(s) in darkness. Panel C. Electrons are transferred to the quencher by both pathways if both light and external reductants are provided.

107

Fig. 3-7. RT-PCR and immunoblotting analyses to detect mRNA levels for csmI and csmJ, as well as CsmI and CsmJ chlorosome protein levels, in the csmX mutant. Panel A. RT-PCR with primers for csmI and csmJ was performed for the cycle numbers specified, and amplicons were separated on a 1.5% (w/v) agarose gel. Panels B and C. Immunoblots to detect the CsmJ (Panel B) and CsmI (Panel C) protein levels in chlorosomes from the wild type and the csmX mutant at the indicated dilution ratios. Antibodies to CsmC served as an internal control served as an internal loading control. Chlorosomes corresponding to a BChl c concentration of 50 μg mL-1 were loaded for the 1:1 sample.

108

Chapter 4

Chlorosome Proteins and Chlorosome Assembly: Construction and Characterization of Mutants Lacking CsmB/CsmF or CsmC/CsmD Motifs on the Chlorosome Envelope of Chlorobium tepidum

109 4.1 Abstract

The ten chlorosome proteins in the green sulfur bacterium Chlorobium tepidum are made up of only four structural motifs: the CsmA/CsmE motif, the CsmB/CsmF motif, the CsmC/CsmD motif, and the iron-sulfur motif. Nine mutants lacking a single chlorosome protein except for CsmA were previously created. Chlorosomes of these mutants were generally similar to the wild-type chlorosomes with respect to size, shape and pigment content, suggesting that chlorosome proteins of the same motif family might functionally substitute for one another. Thus, we proceeded to inactivate each member of the CsmB/CsmF motif and CsmC/CsmD motif by making double and triple mutants. The mutant strains contained significantly reduced amounts of cellular BChl c and/or carotenoids compared to the wild-type strain, and showed apparent growth defects at limiting light intensities. During chlorosome isolation, an additional layer was isolated right below the chlorosome fraction by gradient ultracentrifugation for the mutant strains. These second fraction samples presented distinguishable biochemical and biophysical properties from the normal chlorosomes, and probably provided evidence relating to chlorosome biosynthesis and pigment incorporation. The absorption maximum of the

BChl c Qy band was blue-shifted in the chlorosomes lacking CsmC and CsmD, but red- shifted in the chlorosomes lacking CsmB and CsmF, presumably as a result of altered BChl c aggregation. Electron microscopy of isolated chlorosomes revealed that chlorosomes from the mutant strains deviated from the wild-type chlorosomes in sizes and shapes. In combination, these results confirm that proteins of the same motif family functionally substitute for one another in single mutants, and strongly suggest that chlorosome proteins affect pigment biosynthesis and chlorosome assembly in the green sulfur bacteria. At the same time, multiple C. tepidum mutant strains with modified pigment compositions were constructed in our lab by inactivating enzymes in the BChl c and carotenoids biosynthesis pathways. Chlorosomes were isolated from these mutants, and protein compositions of chlorosomes were analyzed by SDS polyacrylamide gel electrophoresis. This study provides clues of how the altered pigment composition affects

110 the chlorosome protein contents, and supports the conclusion that chlorosome proteins play roles in pigment incorporation and chlorosome assembly in other ways.

111 4.2 Introduction

Green sulfur bacteria, obligate photolithotrophs, utilize chlorosomes as their light harvesting antennae for the photosynthetic reactions. Some green-nonsulfur bacteria, known as filamentous anoxygenic phototrophs, also produce chlorosomes when they grow phototrophically. Although both contain chlorosomes, the two groups of green bacteria appear to be widely divergent based on physiological and biochemical properties, and the presence of chlorosomes might be due to lateral gene transfer (Blankenship, 1992). Chlorosomes from the green sulfur bacterium Chlorobium tepidum contain at least ten chlorosome proteins named CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ and CsmX. Four families of the chlorosome proteins are apparent upon close inspection of the amino acid sequences: CsmA/CsmE, CsmB/CsmF, and CsmC/CsmD (Vassilieva et al., 2000). CsmA and CsmE are 49% identical, and both proteins are synthesized as precursors and are processed by removing the 20 amino acids at their carboxy-termini to generate the mature polypeptides. CsmB and CsmF are 29% identical and 63% similar in sequence. Moreover, the amino-terminal domain of CsmH is clearly related in sequence to these two proteins. The CsmC and CsmD proteins are 26% identical and 45% similar in sequence, and share substantial sequence similarity to the carboxy-terminus of CsmH. The other three chlorosome proteins (CsmI, CsmJ and CsmX) share sequence similarities to the precursor forms of CsmA and CsmE in the carboxyl- terminal regions, while their amino-terminal regions are related in sequence to adrenodoxin-type [2Fe-2S] ferredoxins (Vassilieva et al., 2001). Thus, the ten chlorosome proteins are clearly made up of only four structure motifs. Sequence relationships among chlorosome proteins indicate that gene duplication and divergence must have occurred among a small number of gene types in C. tepidum (Vassilieva et al., 2002b). On the other hand, chlorosomes in green nonsulfur bacterium C. aurantiacus were initially reported to contain only four polypeptides with masses of 5.7 kDa to 18 kDa (Feick and Fuller, 1984). The smallest polypeptide (5.7 kDa) is obviously a homolog of CsmA in green sulfur bacteria, and this protein is also synthesized as a precursor with a carboxyl-terminal extension of 27 amino acids (Theroux et al., 1990). Two other

112 polypeptides, CsmN and CsmM with apparent molecular masses of 11 and 18 kDa, are distantly related in sequence to the CsmC and CsmD proteins of C. tepidum (Niedermeier et al., 1994; Vassilieva et al., 2000). In addition, a 17.2 kDa protein, denoted CsmP, has been detected in chlorosome fractions from C. aurantiacus (Frigaard et al., 2001). The homolog of csmP has not been found in the genome C. tepidum, but related proteins exist in the cyanobacteria Synechocystis sp. PCC 6803 and Synchococus sp. PCC 7002 and occur as the bacterio-opsin linked product in Halobacterium sp. (Frigaard et al., 2001). The chlorosome proteins are exposed at the surface of isolated chlorosomes at various extents, as indicated by protease susceptibility mapping, immunoelectron microscopy and chlorosome agglutination experiments using antibodies against chlorosome proteins (Wullink et al., 1991; Chung and Bryant, 1996b; Vassilieva et al., 2002b). Although the essential protein CsmA is probably assembled at an early stage of chlorosome assembly, it is not clear at what stage the other chlorosome proteins are assembled onto the chlorosome envelope. Likewise, it is not clear whether the other chlorosome proteins are essential for pigment incorporation. Earlier experiments in acetylene-treated Chlorobium vibrioforme cultures indicated that the synthesis of chlorosome proteins and the assembly of the chlorosome envelope are largely independent of pigment synthesis (Vassilieva et al., 2002a). As a strong inhibitor to BChl c biosynthesis, acetylene decreased the cellular level of BChl c in the treated cells to as low as 10% of the level in the control cells. However, the synthesis and accumulation of chlorosome proteins were essentially unaffected in the acetylene-treated cultures. Moreover, in C. aurantiacus, the BChl c and chlorosome protein contents increased at different rates (76-fold versus 10-fold) when the cultures were transferred from chemotrophic (aerobic) to phototrophic (anaerobic) conditions and developed chlorosomes (Foidl et al., 1998). These results suggested that chlorosome envelopes and their components proteins are constitutively synthesized and that the synthesis of chlorosome proteins is not dependent on the BChl c contents. Recent studies in a mutant totally lacking BChl c also revealed valuable information on the chlorosome biosynthesis and protein functions (Frigaard et al., 2002). Constructed by inactivation of the bchK gene, which encodes BChl c synthase, the mutant produces vestigial chlorosomes, denoted “carotenosomes”, which only contain carotenoids, isoprenoid quinones and

113 protein-associated BChl a (Frigaard et al., 2002). Except for CsmA and CsmD, the protein contents of these carotenosomes were significantly reduced to trace amounts, while CsmC, CsmH, CsmJ and CsmX were totally absent as indicated by immunoblotting analysis (Frigaard et al., 2005b). These results argue that the incorporation of several chlorosome proteins is dependent on the presence of BChl c and that most chlorosome proteins are not present at the beginning of chlorosome assembly before BChl c incorporation. The most abundant and smallest chlorosome protein, CsmA, has been identified as the BChl a-binding protein in both green sulfur and green nonsulfur bacteria. Detergent treatments can release CsmA along with BChl a from the chlorosomes in C. tepidum, indicating that BChl a is associated with the CsmA protein (Bryant et al., 2002). Evidence obtained in C. aurantiacus also demonstrates that CsmA binds BChl a, and the CsmA-BChl c complex in the baseplate serves as an intermediate in energy transfer to the reaction center (Sakuragi et al., 1999; Montaño et al., 2003b). However, little information has been generated about the function of the chlorosome proteins apart from the CsmA protein. SDS-treated chlorosomes retain their typical shape and size but lack all proteins except CsmA; this observation suggests the possibility that none of the nine proteins are absolutely required for maintaining the shape and integrity of chlorosomes, at least not after the BChl c aggregates and baseplate have been formed (Bryant et al. 2002). In recent studies, nine mutants lacking a single chlorosome protein except CsmA were created in C. tepidum (Frigaard et al., 2004a). Only four of the csm mutants exhibited convincing phenotypes. Compared to the wild-type strain, the csmB mutant cells and chlorosomes contained about 25% less carotenoids, the csmH chlorosomes were about 20% shorter in the longest dimension, and the csmJ strain grew ~10% more slowly at saturating and inhibitory light intensities. Chlorosomes from the csmC mutant were 25% smaller than those from the wild type, and the BChl c absorbance maximum was blue shifted from 750 nm to 743 nm (Frigaard et al., 2004a). Other than these differences, no distinguishing characteristics were found in the csmD, csmE, csmF, csmJ and csmX mutant cells and chlorosomes. A csmD csmE double mutant was also obtained by inactivating the adjacent csmD and csmE genes together. The csmD csmE double mutant produced normal chlorosomes and did not exhibit any obvious growth or pigmentation

114 phenotype. Based upon the above results, none of these nine chlorosome proteins is essential for the biogenesis, light harvesting, or the structural organization of the chlorosomes, and chlorosomes can obviously tolerate considerable changes in protein composition. Since classes of chlorosome proteins are related in sequence, there is the possibility that proteins of the same motif family might functionally substitute for one another when a given protein is missing. In this study, each member of the CsmB/CsmF family and CsmC/CsmD family of genes were inactivated by making double and triple mutants. Six mutant strains were constructed by natural transformation and antibiotic resistance selection: csmC csmD, csmC csmD csmE, csmC csmD csmH, csmC csmD csmE csmH, csmB csmF and csmB csmF csmH. The mutant cells contained significantly reduced BChl c and/or carotenoids levels, and the isolated chlorosomes of the mutants were greatly modified in pigment composition, absorption properties and morphology. These results strongly suggest that the chlorosome proteins play a role in pigment incorporation into the chlorosomes, and that defects in chlorosome assembly affect the regulation of pigment biosynthesis. In addition, the protein compositions of chlorosomes of mutants lacking the BChl c or carotenoid biosynthesis enzymes were analyzed; the data obtained show that the pigment contents affect the chlorosome protein contents in other ways.

115 4.3 Materials and methods

4.3.1 Plasmids construction in Escherichia coli

Escherichia coli ElectroMAX DH10B cells [genotype F- mcrA Δ(mrr-hsdRMS-mcrBC) ф 80dlacZ ΔM15 ΔlacX74 deoR recA1 endA1 araD139 Δ(ara, leu)7697 galU galK λ- rpsL nupG from Gibco BRL Products, Gaithersburg, MD] were used for recombinant DNA manipulations. Recombinant DNA experimental methods with E. coli cells and DNA fragments were performed as described in Chapter 3.3.1. The plasmids pCT720 (containing the csmC gene fragment) and pCBS11 (containing the csmB gene fragment interrupted by the aadA resistance marker) were constructed by previous lab members (Chung, 1995; Frigaard et al., 2004a). The plasmid pUCB::aacC was constructed from pCBS11 by deletion of the aadA resistance marker and insertion of the aacC resistance marker (GmR) at the same PstI restriction sites. The plasmid pUCC::aacC was constructed from pCT720 by insertion of the aacC resistance marker (GmR) at the NsiI restriction site within the coding sequence of the csmC gene. For cloning of the csmH gene, the gene was modified by PCR with the primers 5’ CAA ATC AAA CCC ATA TGG CTA CCG AA (NdeI) and 5’ CGA ATG CCT TGG ATC CGT TCT ATT CA (BamHI), introducing an NdeI site and a BamHI site at the ends of the gene (mutated bases are in bold, and introduced restriction sites are underlined). The plasmid pUCH13 was constructed by cloning the modified csmH gene into pUC19 vector at the NdeI and BamHI restriction sites, and the plasmid pUCH::cat-ermC was constructed by inserting the cat-ermC resistance marker (EmR) into the pUCH13 NheI restriction site within the coding sequence of the csmH gene. For cloning of the csmA gene, the gene was modified by PCR with the primers 5’ AGG CAG TTA AGA GCT CTG CTG GCT C (SacI) and 5’ GCT CTG TCT GTC TAG AGA TAT AGA GAG (XbaI) introducing a SacI site and a XbaI site at the ends of the gene (mutated bases are in bold, and introduced restriction sites are underlined). The plasmid pUCA8 was constructed by cloning the modified csmA gene into pUC19 vector at the SacI and XbaI restriction sites, and the plasmid pUCA::cat-ermC was constructed by inserting the cat-ermC resistance marker (EmR) into the pUCA8 NcoI restriction site within the coding sequence of the csmA gene. The

116 construction and structure of pUCB::aacc, pUCC::aacC, pUCH::cat-ermC and pUCA::cat-ermC were shown in Fig. 4-1.

4.3.2 Growth condition and mutant construction and confirmation in Chlorobium tepidum

Mutant construction was accomplished in Chlorobium tepidum strain WT2321 (Wahlund and Madigan, 1995), which is a plating strain derived from C. tepidum strain ATCC 49652 (Wahlund et al., 1991). C. tepidum cells were grown in CL medium or on CP plates as described in Chapter 2.3.1. Manipulations and small volume growth were performed at 42 ºC in an anaerobic chamber (Coy Laboratory Products, Grass Lake, MI). Large-volume cultures were grown in sealed 2-L bottles in a water-bath at 42-45 ºC. For growth rates studies, the cells were grown in CL medium in 20-mL screw-capped glass tubes on a radial rotator. The culture turbidity was measured at 600 nm at different times for growth rate calculations. For natural transformation of C. tepidum, cells of the wild-type strain, the csmF mutant, the csmD mutant and the csmD csmE mutant were grown to late-exponential phase and harvested by microcentrifugation. The plasmids pUCB::aacC, pUCC::aacC, pUCH::cat-ermC and pUCA::cat-ermC were linearized by restriction enzyme AhdI. Pelleted cells from 1-mL culture were resuspended in 100 μL of CL medium, mixed with 10 μg linearized DNA, and spotted on a non-selective CP plate for 10-18 h incubation. The cell patch was then scraped off, streaked on selective CP plates and incubated for 5-6 days to allow single transformant colonies to appear. After two more rounds of selective streaking, the single colonies were inoculated into CL medium and grown into dense cultures. PCR with primers of the csm genes (csmB-F, 5’-CCG CTC CTG AAA TCT GTC AAA G; csmB-B, 5’-CGA CCG ATA CAC TGC CTT GG; csmC-F, 5’-AAA GAC GCC TCC TCC ACT C; csmC-B, 5’-GGA ACT TGA TTT TAC CGA CAG G; csmH-F, 5’-ATG GCT ACC GAA GAA ACA AAC; csmH-B, 5’-GCT GTG GGC TGA GGA GTC AT) was performed to analyze the segregation status using DNA from 1-2 μL of cells as templates. The linearized plasmid pUCA::cat-ermC (EmR) was transferred to wild type and the bchK mutant (SmR, SpR) in attempts to inactivate the csmA gene. The linearized plasmid pUCB::aacC (GmR) was transferred to the csmF mutant (SmR, SpR) to

117 construct the double mutant csmB csmF. The linearized plasmid pUCC::aacC (GmR) was transferred to the csmD and csmD csmE mutants (SmR, SpR) to construct double mutants lacking CsmC and CsmD. After confirmation of the csmB csmF, csmC csmD and csmC csmD csmE mutants, the linearized plasmid pUCH::cat-ermC (EmR) was transferred to these mutants (GmR, SmR, SpR) to construct triple mutants lacking CsmB, CsmF, and CsmH or CsmC, CsmD, and CsmH. Segregated mutant strains were stored as frozen stocks at -80 ºC as described in Chapter 3.3.2.

4.3.3 Chlorosome isolation

Chlorosomes were isolated by sucrose-gradient ultracentrifugation as described in Chapter 2.3.2. Pelleted cells from a 2-L culture were disrupted by three passages through a French press cell (124 MPa, 4 ºC) in isolation buffer containing 2 M NaSCN. Chlorosomes-enriched fractions were pelleted by ultracentrifugation at 220,000 × g at 4 ºC for 2 h, and further purified on 7 to 47% (w/v) continuous sucrose gradients by ultracentrifugation at 220,000 × g at 4 ºC for 18 h. The chlorosome fraction was then collected, diluted 4-fold with phosphate-buffered saline and centrifuged at 240,000 × g for 1.5 h. The resulting pellet was resuspended in the same buffer and centrifuged again, and the firmly pelleted chlorosomes were resuspended in a small volume of phosphate- buffered saline containing 1 mM PMSF and 2 mM DTT. The purified chlorosomes were aliquoted and stored at -80 ºC for future use.

4.3.4 Analysis of protein contents

SDS-PAGE and immunoblotting. Protein composition was analyzed on 16% T and 3.3% C gels by Tris-Tricine buffered, SDS polyacrylamide gel electrophoresis (Schägger and van Jagow, 1987). Chlorosome samples containing 10 μg BChl c were loaded for silver staining, and samples containing 100 μg BChl c were loaded for immunodetection. Silver staining and immunoblotting were performed as described in Chapter 2.3.4. The antibody dilutions were as follows: anti-CsmA, 1:2,000; anti-CsmB, 1:180; anti-CsmC, 1:1,000; anti-CsmD, 1:800; anti-CsmH, 1:3,000; anti-CsmK, 1:10,000; anti-CsmI, 1:5,000; anti-

118 CsmJ, 1:3,000; anti-CsmW, 1:5,000; anti-CsmX, 1:1,000; and goat anti-(rabbit-IgG), 1:7,000.

Lowry protein assay. The Lowry protein assay was used for determination of total protein amounts in C. tepidum cells and subcellular fractions. The assay was performed using a procedure described by Sigma (procedure P5656, Sigma, St. Louis, MO) with slight modifications. Proteins were precipitated from whole cells (0.5-1 mL of dense culture) or chlorosomes (10-20 μL) with methanol or acetone and pelleted by centrifugation at 13,000 × g for 15 min. The protein pellets were solubilized in 50 μL of 0.5 M NaOH by boiling and vortexing before addition of 0.5 mL Lowry reagent (Sigma, St. Louis, MO). The samples were then diluted to 1 mL with water and incubated at room temperature for 20 min. Folin and Ciocalteu’s phenol reagent (0.25 mL, Sigma, St. Louis, MO) was added with immediate mixing, and the samples were incubated for 30 min at room temperature before taking absorbance at 750 nm. Bovine serum albumin (0-400 μg, Pierce, Rockford, IL) was used as the standard in this method.

4.3.5 Spectroscopy and pigment content determination

Determination of BChl c concentrations. BChl c was extracted by diluting chlorosomes 1:100 in acetone:methanol (7:2 v/v) or resuspending pelleted cells in equal volume of acetone:methanol (7:2 v/v). Proteins were separated by centrifugation at 4°C for 10 min. The pigments were then diluted 1:100 (chlorosome pigments) or 1:10 (cell pigments) in organic solvents, and the concentration of BChl c was determined by absorption spectroscopy using the following specific absorption coefficients: 86 (g/L)-1 cm-1 at 669 nm for methanol (Stanier and Smith, 1960), 92.6 (g/L)-1 cm-1 at 662.5 nm for acetone (Stanier and Smith, 1960) or 91 (g/L)-1 cm-1 at 666 nm for acetone:methanol (7:2 v/v) (Oelze, 1985).

High performance liquid chromatography. Pigments and quinones were extracted from small amounts of chlorosomes or pelleted cells with 100-200 μL acetone:methanol (7:2 v/v). The extracted pigment mixture was microcentrifuged, filtered and mixed with 0.1

119 volume of 1 M ammonium acetate before immediate injection onto the HPLC. The HPLC system used was an Agilent 1100 series system (Agilent Technologies, Palo Alto, CA) with a 4.5 mm × 25 cm Discovery C18 column (Supelco, Bellefonte, PA) under the control of Agilent ChemStation software. The HPLC procedure was as described in Chapter 3.3.4. Pigment contents were determined by absorption spectroscopy using the following absorption coefficients: BChl c, 20 L g-1 cm-1 at 635 nm (Stanier and Smith, 1960); BChl a, 60 L g-1 cm-1 at 770 nm (Tsuji et al., 1995); carotenoids, 265 L g-1 cm-1 at 491 nm (Holo et al., 1985); chlorobiumquinone, 17 L g-1 cm-1 at 270 nm (Frydman and Rappaport, 1963); menaquinone-7, 26 L g-1 cm-1 at 270 nm (Dunphy and Brodie, 1971).

Absorption spectroscopy. Absorption spectra of chlorosomes and whole cells were measured from 350 to 900 nm with Visionlite Scan software (Thermo Electronic, Madison, WI) on a GENESYS 10 spectrophotometer (Thermo Spectronic, Rochester, NY). Chlorosomes were diluted at the ratio of 1:10,000 and cells were diluted at the ratio

of 1:10 in the phosphate buffered saline (10 mM KH2PO4, 50 mM NaCl, pH 7.0) before the absorption spectra were taken.

Fluorescence spectroscopy. Fluorescence emission spectra of chlorosomes or whole cells were measured under reducing conditions with a SLM-AMINCO 8100 Series 2 spectrofluoremeter (SLM Instruments, Urbana, IL). Excitation light was generated at 460 nm by an ELXE-500 Xenon lamp (SLM Instruments, Urbana, IL). Chlorosomes or cells

were diluted in KH2PO4 buffer (10 mM KH2PO4, pH 7.0) to ODmax 0.2 (at about 750 nm), incubated with 100 mM sodium dithionite in lid-closed cuvettes for at least one hour, and the fluorescence emission spectra were recorded at 15ºC from 700 to 850 nm.

4.3.6 Electron microscopy

Chlorosomes were diluted with ddH2O to ODmax 2-3 (at about 750 nm), and bound to Cu- coated formvar grids (400 mesh) by incubation at room temperature for 3 min. The chlorosomes were stained with uranyl acetate (2% (w/v) aqueous) for 30 s in the dark,

120 and images were photographed using a JEOL 1200 EXII transmission electron microscope (Peabody, MA).

121 4.4 Results

4.4.1 Construction of C. tepidum mutants

Transformation of the wild type and the bchK mutant with linearized plasmid pUCA::cat- ermC yielded no colonies on selective plates after incubation for 10 to 12 days under high light intensity (more than 150 μmol photons m-2 s-1). This result suggests that CsmA may be an essential component for light harvesting and photosynthesis and that inactivation of this protein might be lethal to the C. tepidum cells. Transformations of the csmD, csmDcsmE and csmF mutants with DNA fragments containing csmC::aacC or csmB::aacC yielded dark green colonies after 5-6 days incubation on selective plates. The colonies were restreaked two more times on selective plates, and isolated, individual colonies were inoculated into small volume of CL medium. Segregation of the transformants was confirmed by PCR on whole cells with primers of the corresponding csm genes as well as primers of the antibiotic resistance markers (data not shown). PCR with the various csm primers yielded the anticipated csm fragments of 0.5-0.8 kb with the wild-type strain, but products of 1.5-2.5 kb with the segregated transformants. PCR with primers of the antibiotic resistance markers yielded 0.5-0.8 kb products with the segregated transformants but no products with the wild-type strain. All colonies of the transformants csmC csmD and csmC csmD csmE were confirmed by PCR as homozygous for the insertionally inactivated gene alleles; however, the csmB csmF transformants were both segregated strains and merodiploid strains in the ratio of 5:2. Clonal isolates of the three segregated mutant strains were grown in CL medium to late exponential phase and used for the next round of transformation with linearized plasmid pUCH::cat-ermC. Dark green colonies were observed on selective plates after 5-6 days incubation, and the twice re-streaked individual colonies were confirmed by PCR to be segregated transformants with the insertionally inactivated csmH gene alleles. Thus, six mutants of C. tepidum were generated: csmC csmD, csmC csmD csmE, csmC csmD csmH, csmC csmD csmE csmH, csmB csmF and csmB csmF csmH. The mutants of csmC csmD csmH and csmC csmD csmE csmH lack all the chlorosome proteins of the CsmC/CsmD

122 family, and the mutant of csmB csmF csmH lacks all the chlorosome proteins of the CsmB/CsmF family (see Chapter 4.4.4).

4.4.2 Growth defects of the mutated cells

Growth rates are calculated by the equation

Nt μ = ln N0 t

where Nt is the optical density at time t, N0 is the reference optical density, t is the time elapsed between the two, and μ is the rate of growth. Table 4-1 shows the growth rates of the wild type and the mutants of C. tepidum lacking the CsmC/CsmD motif proteins or the CsmB/CsmF motif proteins at four different light intensities: 9 μmol photons m-2 s-1, 32 μmol photons m-2 s-1, 80 μmol photons m-2 s-1 and 210 μmol photons m-2 s-1. Surprisingly, none of the mutants showed a significant deviation in growth rates from the wild-type strain at saturating and inhibitory light intensities (>80 μmol photons m-2 s-1). At limiting light intensities, the csmC csmD csmE mutant exhibited a similar growth rate as the csmC csmD mutant, and the csmC csmD csmE csmH mutant exhibited a similar growth rate as csmC csmD csmH. These results indicate that the CsmE protein is not an essential chlorosome component, in agreement with the results for the csmE single mutant. The former two mutants grew about 15% slower than the wild-type strain at 9 μmol photons m-2 s-1, but showed no convincing growth defects at 32 μmol photons m-2 s- 1. The csmC csmD csmH and csmC csmD csmE csmH mutants grew about 25% slower than the wild-type strain at 9 μmol photons m-2 s-1, and about 15% slower at 32 μmol photons m-2 s-1. The csmB csmF and csmB csmF csmH mutants showed significant growth defects at the lowest light intensity (9 μmol photons m-2 s-1). The csmB csmF mutant grew about 75% as fast as the wild type, and the csmB csmF csmH mutant grew only about 55% as fast as the wild type. The growth rates of the two mutants increased to 95% and 92% of that of the wild type when the light intensity was increased to 32 μmol photons m-2 s-1.

123 4.4.3 Cellular absorption profile and pigment contents

Fig. 4-2 shows absorption spectra of cell suspensions of the wild type and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif determined at equal cell densities as determined by OD600 nm. The absorption spectrum of the wild-type strain

(shown as solid lines in all panels) exhibited two major peaks: the BChl c Qy absorption peak at 752 nm, and the associated BChl c Soret absorbance peak at 462 nm. The carotenoid absorbance was detectable as a weak shoulder at ~520 nm, and the FMO protein-bound BChl a peak was detectable as a weak shoulder at about 810 nm. As judged from the amplitudes of the BChl c Qy absorption band, the csmC csmD, csmC csmD csmE, csmC csmD csmH and csmC csmD csmE csmH mutants (shown as dashed lines in Panels A, B, C, D) apparently contained less BChl c or had BChl c aggregated

differently than the wild-type strain at the same OD600 nm value. Moreover, the BChl c Qy absorption maximum in the csmC csmD and csmC csmD csmE mutants was slightly blue- shifted from 752 nm in the wild type to 748 nm. The absorbance properties of the csmB csmF and csmB csmF csmH mutant cells were generally very similar to the wild-type cells except that the BChl c Qy absorption maximum was slightly red-shifted to 756 nm for the csmB csmF mutant and to 753 nm for the csmB csmF csmH mutant. Fluorescence emission spectra of all strains were recorded under reducing conditions, and all appeared to be very similar. The fluorescence emission maximum of BChl c was detected at about 771 nm (at 768 nm for the csmC csmD and csmC csmD csmE mutants), and the emission maximum of BChl a was detected at about 805 nm in all cases. The absorption maxima and fluorescence emission maxima of the wild-type and mutant strains are listed in Table 4-2. HPLC analyses were utilized to determine the cellular pigment contents in the wild type and the mutant strains. Measurements were performed at least twice with cells

from independent mid-exponential phase cultures (OD600 nm ~ 1.0), and the averages are shown in Table 4-2. Cellular protein amounts were determined by the Lowry protein assay and used as the standard in pigment content comparisons between the wild type and the mutant strains. The wild-type cells grown at ~30 μmol photons m-2 s-1 contained about 232 mg of BChl c, 2.5 mg of BChl a, 13.6 mg of carotenoids and 12 mg of

124 quinones per g of cellular protein. The contents of BChl a and quinones varied slightly in the cells of the mutant strains, which was probably caused by minor differences in the growth states or growth conditions. The carotenoid contents were slightly decreased to 11.8 and 12.0 mg per g of cellular protein in the mutants lacking the CsmB/CsmF motif (the csmB csmF and csmB csmF csmH mutants). However, the BChl c amounts were very significantly reduced to 144 and 123 mg of BChl c per g of cellular protein in the csmC csmD csmH and csmC csmD csmE csmH mutants, which were only 55-60% of the BChl c amount in the wild-type strain. At the same time, the BChl c contents in the csmC csmD, csmC csmD csmE, csmB csmF and csmB csmF csmH mutants were only reduced by 10- 15% compared to the wild type. Therefore, both absorption spectroscopy and HPLC pigment analysis indicated the mutant strains (especially csmC csmD csmH and csmC csmD csmE csmH) contained decreased amounts of cellular BChl c.

4.4.4 Chlorosome isolation and protein component analysis

C. tepidum cells from 2-L cultures were harvested for chlorosome isolation. After cell disruption and low-speed centrifugation to spin down the cell debris, the suspension containing chlorosomes was subjected to ultracentrifugation to pellet the chlorosome- enriched fraction and to separate the chlorosomes from the soluble components. At this step, an orange-colored fraction in the uppermost 2 mL banded at the top of the suspensions from all the mutants lacking the CsmC/CsmD or the CsmB/CsmF motif, whereas no such colored fraction was observed for the wild-type strain and any of the single mutant strains except the csmC mutant. Absorption spectra in methanol showed that this colored fraction contained mostly carotenoids (with absorption maxima at 460 nm and 490 nm) as well as small amounts of BChl c (with absorption maximum at 669 nm) and BChl a (with absorption maximum at 770 nm). The ratio of carotenoids to BChl c was about 20-fold greater than the ratio in typical chlorosomes. SDS polyacrylamide gel electrophoresis showed that CsmA and CsmB were also contained in this orange- colored fraction, along with a large number of other cellular proteins. Thus, the orange- colored fraction is likely to represent intermediate structures during chlorosome assembly before or near the beginning of BChl c incorporation. This fraction represented less than

125 1% of the total chlorosomes, as estimated up on the carotenoids and BChl a contents of this fraction. The chlorosome-containing pellet from the first ultracentrifugation step was resuspended and loaded onto 7-47% sucrose gradients for further purification. For the wild-type strain, the chlorosome fraction appeared as a highly concentrated, dark green band in the upper 1/3 to 1/4 of the sucrose gradient. A thin band of light green color was present at the 2/3 position of the gradient, which contained mainly membrane fragments. The other cell debris was precipitated at the bottom of the tube as a brownish pellet. The gradients for the csmC csmD and csmC csmD csmE mutants were similar in appearance to those of the wild-type strain, but the gradients for the csmC csmD csmH, csmC csmD csmE csmH, csmB csmF and csmB csmF csmH mutants differed from those of the wild type. For these mutants the chlorosome band was at the normal position, but an additional green layer was recovered right below the chlorosome fraction from the sucrose gradient. The second layer for the csmB csmF and csmB csmF csmH mutants was at the 1/2 to 3/4 position of the gradient and overlapped with the membrane fragment fraction. The second layer for the csmC csmD csmH and csmC csmD csmE csmH mutants was a well-defined band with relatively clear edges. The sucrose gradients of the wild-type strain, the csmB csmF mutant and the csmC csmD csmH mutant are shown in Fig 4-3, Panel A. In subsequent steps, the second fraction samples were further purified just as chlorosomes: they were collected from the sucrose gradients, pelleted by two more rounds of ultracentrifugation and resuspended in phosphate-buffered saline. The protein compositions of chlorosomes and second fraction samples from the wild-type strain, the csmB csmF mutant and the csmC csmD csmH mutant were analyzed by SDS polyacrylamide gel electrophoresis, and proteins were detected by silver staining as well as immunoblotting (Fig. 4-3, Panels B, C). Silver staining revealed many proteins between 14 kDa to 31 kDa in the second fraction samples from the csmB csmF and the csmC csmD csmH mutants. These proteins were probably contamination from the membrane fragments in the csmB csmF second fraction sample. The CsmA protein, the most abundant protein in wild-type chlorosomes, was contained in the second fraction sample of the csmB csmF mutant, but CsmA was hardly detectable in the second fraction sample of the csm CcsmD csmH mutant. Interestingly, the most abundant protein in the

126 second fraction sample of the csmB csmF mutant was a protein with an apparent mass slightly larger than CsmA, which had a characteristic greenish color when stained with silver. The most abundant protein in the second fraction sample of the csmC csmD csmH mutant is a protein migrating at the same position of CsmB/CsmE. Fig. 4-3 Panel C shows the immunoblotting results of the chlorosomes and the second fraction samples with anti-CsmA and anti-CsmF antibodies (anti-CsmF cross-reacts with both CsmF and CsmB). Immunoblotting with anti-CsmF antibodies confirmed that the abundant protein in the second fraction sample of the csmC csmD csmH mutant was CsmB. Thus, the amount of CsmB in this sample greatly exceeded the amount of CsmA. The greenish colored band in the second fraction sample of the csmB csmF mutant did not cross-react with either anti-CsmA or anti-CsmB antibodies. Further results showed that this polypeptide did not cross-react with any antibodies against chlorosome proteins. The polypeptide was later identified by matrix-assisted laser desorption ionization time-of- flight mass spectrometry (MALDI-TOF-MS) as a hypothetical protein encoded by the gene CT0426, which is related in sequence to the Flp/Fap pilin component in various bacteria species. Besides the second fraction samples of the csmB csmF and csmC csmD csmH mutants, the chlorosomes of each of the mutant strains were also analyzed by Tris- Tricine-buffered SDS polyacrylamide gel electrophoresis, and the proteins were detected by silver staining and immunoblotting (Fig. 4-4). Immunoblotting with antibodies against chlorosome proteins verified that the CsmB, CsmC, CsmD, CsmF and CsmH proteins were successfully inactivated in double and triple combinations in the mutant strains (Fig. 4-4 B). Silver staining indicated that the relative amounts of the remaining proteins (CsmA, CsmC, CsmD, CsmI and CsmJ) in the csmB csmF and csmB csmF csmH chlorosomes were about the same level as those in the wild type (Fig. 4-4 Panel A Lanes 6, 7). However, the amounts of CsmF and CsmB seemed to be slightly increased in chlorosomes from the csmC csmD, csmC csmD csmE, csmC csmD csmH and csmC csmD csmE csmH mutants, and there appeared to be a slight decrease in the amount of CsmA (Fig. 4-4, Panel A Lanes 2, 3, 4, 5).

127 4.4.5 Absorption profile and pigment contents of the chlorosomes

Panels A-H of Fig. 4-5 show the absorption spectra of wild-type chlorosomes and chlorosomes (or second fraction samples) from the mutant strains in solid and dashed lines, respectively. The absorption spectra were taken in aqueous buffer and normalized

for their contents of monomeric BChl c (OD669 nm in methanol = 1). The BChl c Qy absorption maximum in the wild-type chlorosomes was about 750 nm. Chlorosomes from the csmC csmD, csmC csmD csmE mutants had a BChl c Qy absorption maximum that was blue-shifted by about 12 nm (Fig. 4-5, Panels A and B). A blue-shift of 5 nm in the

BChl c Qy absorption maximum was also observed in chlorosomes from the csmC csmD csmH and csmC csmD csmE csmH mutants (Fig. 4-5, Panels C and D). In contrast to the csmC csmD csmH chlorosomes, the second fraction sample from the csmC csmD csmH

mutant had a BChl c Qy absorption maximum at 732 nm, which was blue-shifted by 18

nm from that of the wild type (Fig. 4-5, Panel E). In addition, the amplitude of the Qy absorption peak of this second fraction sample was much smaller than that of any other chlorosome sample, although all samples contained the same amounts of monomeric BChl c. These properties indicate that the BChl c organization was greatly modified in the second fraction sample of the csmC csmD csmH mutant. The BChl c Qy absorption maxima in the csmB csmF and csmB csmF csmH chlorosomes were red-shifted by 6 and 3 nm compared to the wild type (Fig. 4-5, Panels F and G). Consistent with the results of

the csmB csmF chlorosomes, a red-shift (about 4 nm) in the BChl c Qy absorption maximum was also observed in the second fraction sample from the csmB csmF mutant (Fig. 4-5, Panel H). A shoulder at 810 nm was detectable in this sample, which was probably caused by the protein-bound BChl a and FMO from contaminating membranes. Fluorescence emission spectra of the above samples were recorded under reducing conditions with 10 mM sodium dithionite (data not shown). The BChl c emission maximum was blue-shifted by 2-6 nm in the samples from the csmC csmD, csmC csmD csmE, csmC csmD csmH and csmC csmD csmE csmH mutants, and red-shifted by 2 nm in the samples from the csmB csmF and csmB csmF csmH mutants. The emission maximum of BChl a appeared as a shoulder at about 805 nm in all the samples, but was barely detected in the second fraction sample from the csmC csmD csmH mutant (Fig. 4-

128 5, Panel I). The absorption maxima and fluorescence maxima of the samples from the wild-type and mutant strains are summarized in Table 4-3. HPLC analysis was used to determine the pigment contents in the chlorosomes and the second fraction samples from the wild type and mutant strains (Table 4-3). Using the BChl c content as the reference standard, the BChl a content was 10-12 mg per g of BChl c in the wild type and mutant chlorosomes, but the BChl a content significantly increased (2-fold and 5-fold) in the second fraction samples from the csmC csmD csmH and csmB csmF mutants. The increased BChl a contents in the second fraction samples was probably due to FMO- and reaction center-associated BChl a contained in the membrane contamination. The carotenoid content was 57 mg per g of BChl c in the wild- type chlorosomes, and this value was increased about 30% in the csmC csmD csmH and csmC csmD csmE csmH chlorosomes. Considering the decreased cellular BChl c contents in the two mutants, the relatively high amounts of carotenoids were likely to be the result of the decreased cellular contents of BChl c. In contrast, the csmB csmF and csmB csmF csmH chlorosomes contained significantly reduced amounts of carotenoids (~25% less than that of the wild type). The quinone contents (menaquinone-7 and chlorobiumquinone) were similar in the wild-type and mutant samples except for the 2- fold increase in menaquinone contents in the csmC csmD csmH chlorosomes, the csmC csmD csmE csmH chlorosomes and the csmC csmD csmE csmH second fraction sample.

4.4.6 Electron microscopy of chlorosomes

Mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif exhibited no cell morphology phenotype by light microscopy (data not shown). However, electron microscopy revealed obvious size differences among the isolated chlorosomes from the wild-type and the mutant strains (Table 4-5 and Fig. 4-6). The measured dimensions of chlorosomes from the wild-type strain (sample size, 40) were: length, 165 ± 60 nm; width, 60 ± 25 nm; and length to width ratio, 2.7 ± 0.6 (Fig. 4-6, Panel A). Chlorosomes from the csmC csmD, csmC csmD csmE, csmC csmD csmH and csmC csmD csmE csmH mutants deviated significantly from the wild type with reduced dimensions of 100 ± 30 nm in length and 45 ± 15 nm in width (sample size, 25-40) (Fig. 4-6, Panels B, C, D, E).

129 Noticeably, the length to width ratios of chlorosomes from the csmC csmD and csmC csmD csmE mutants differed significantly from that of the wild type (2.1 ± 0.6 and 2.0 ± 0.5), but this difference was not observed in chlorosomes from the csmC csmD csmH and csmC csmD csmE csmH mutants (length to width ratio, 2.4 ±0.7 and 2.6 ± 0.8). The second fraction sample of the csmC csmD csmH mutant mostly contained very small chlorosomes (Fig. 4-6, Panel F). These very small chlorosomes were only 55 ± 10 nm in length and 27 ± 7 nm in width, much smaller (85% smaller in size) than wild-type chlorosomes. The chlorosomes from the csmB csmF mutant were about 25% longer and 15% wider than those from the wild-type strain (length, 210 ± 80 nm; width, 70 ± 20 nm), with a length to width ratio of 3.1 ± 0.6 (Fig. 4-6, Panel G). The dimensions of the csmB csmF csmH mutant were similar to those of the wild-type chlorosomes (Fig. 4-6, Panel H). The second fraction sample of the csmB csmF mutant contained a few rod-shaped chlorosomes with similar sizes as the csmB csmF chlorosomes (shown by the arrows), but this fraction was mostly enriched by the irregular-shaped structures with diameters ranging from 50 to 150 nm (Panel I). The irregular-shaped structures are probably membrane fragments and vesicles.

4.4.7 Chlorosome protein composition of mutants lacking the BChl c and carotenoids biosynthesis enzymes

Based on comparative analyses with known enzymes or sequence motifs, the genes encoding enzymes involved in BChl c and carotenoid biosynthesis were predicted in the genome sequence of C. tepidum (Eisen et al., 2002; Frigaard et al., 2004b; Frigaard et al., 2005a). The genes encoding these putative enzymes were inactivated by transformation, and the enzymatic functions were identified by analyzing the pigment compositions in the resulting mutants (Frigaard et al., 2004b; A. Gomez Maqueo Chew and J.A. Maresca, unpublished data). Nine genes encoding enzymes involved in BChl c biosynthesis and six genes encoding enzymes involved in carotenoid biosynthesis were identified in this way. The identified genes as well as pigment compositions in the resulting mutants are listed in Table 4-4.

130 Chlorosomes of these mutant strains were purified on sucrose gradients after over-night ultracentrifugation. The only exception was the bchS* sample, which was concentrated from the brown-colored supernatant after the first ultracentrifugation step. Proteins of chlorosomes were separated by SDS polyacrylamide gel electrophoresis and detected by silver-staining and immunoblotting analysis (Fig. 4-7). As seen from the silver staining figure (Fig. 4-7, Panel A), the chlorosome samples from the bchT, bchH, bchS and bchS* mutants, lacking the Mg chelatase subunit T, H and S respectively, contained less CsmC, CsmI, CsmJ and CsmH than the wild-type chlorosomes. In fact, immunoblotting with antibodies against chlorosome proteins confirmed that the amounts of CsmC and CsmH were decreased in the bchT and bchH samples and severely decreased in the bchS and bchS* samples, which also contained greatly reduced amounts of BChl c (Fig. 4-7, Panel B). The CsmK (18-20 kDa) and CsmW (18 kDa) proteins are two contaminants in chlorosomes of C. tepidum (Vassilieva et al., 2002b). The two proteins were both present in the bchT, bchH, bchS and bchS* samples, and a significant amount of CsmK was present in bchS samples as indicated by immunoblotting (Fig. 4-7, Panel B). Silver staining of chlorosomes from the BChl c methyltransferases mutants (bchU, bchQ bchU, bchR bchU, bchQ bchR bchU) revealed many protein bands above 14 kDa, even after the chlorosomes were purified twice by sucrose gradient ultracentrifugation (Fig. 4-7, Panel A). Further immunoblotting analysis also detected large amounts of CsmW and CsmK in these samples (Fig. 4-7, Panel C). At the same time, the contents of CsmI, CsmJ, CsmH and CsmC were obviously reduced, especially in the chlorosomes from the triple mutant bchQ bchR bchU (Fig. 4-7, Panel C). The crtB and crtP chlorosomes were isolated from mutant strains lacking phytoene synthase or phytoene dehydrogenase and that these mutants synthesize no carotenoids or phytoene only, separately. The contents of CsmC were lower in the crtB and crtP samples (Fig. 4-7, Panel A). Protein components of chlorosomes from the other mutants (bchQ, bchR, bchQR, bchO, bchP, bchV, crtC, crtH, crtQ, crtU) were also detected by silver staining of the polyacrylamide gels (Fig. 4-7, Panel A). No significant differences were observed between these samples and the wild-type chlorosomes.

131 4.5 Discussion

In this work, six mutants of C. tepidum lacking the CsmC/CsmD motif proteins or the CsmB/CsmF motif proteins were generated by insertional gene inactivation. The mutant cells were characterized by growth rate tests, and pigment components were analyzed by high performance liquid chromatography. Growth defects of these mutants were only detected at low light intensities, suggesting that these mutants have defects in light harvesting. The cellular contents of carotenoids were decreased (15-20%) in the mutants lacking the CsmB/CsmF motif (the csmB csmF and csmB csmF csmH mutants), and the cellular contents of BChl c were greatly reduced (40-45%) in the csmC csmD csmH and csmC csmD csmE csmH mutants and slightly reduced in the other four mutant strains (csmC csmD, csmC csmD csmE, csmB csmF, csmB csmF csmH). Interestingly, a correlation was found between the growth rates under low light intensities and the cellular BChl c contents per g of cellular protein (Fig. 4-8). The correlation strongly suggests that the growth defects in the mutant strains were mainly caused by decrease of the amounts of BChl c, which is the dominant light harvesting pigment in C. tepidum cells. Carotenoids are also involved in light harvesting, energy transfer and triplet quenching in C. tepidum, and the growth defects of the csmB csmF and csmB csmF csmH mutants might be partially due to their decreased carotenoid contents. The csmB csmF csmH mutant had the greatest defects in light harvesting, and had a growth rate much lower than expected for its BChl c content. On the other hand, the two mutants completely lacking the CsmC/CsmD motif grew faster than would be predicted solely upon their BChl c contents. In a previous study, nine mutants of C. tepidum each lacking a single chlorosome protein were constructed and characterized (Frigaard et al., 2004a). Although the csmC mutant was reported to grow approximately 90% as fast as the wild type under a limiting light intensity, the csmB, csmD, csmF, and csmH mutant strains showed no growth-rate defects under any light intensities tested. The absence of phenotypic effects in most of the single mutants was probably due to the large number of chlorosome proteins present in C. tepidum, and proteins belonging to the same motif families may functionally substitute for one another. In this study, significant defects on growth rate tests were detected in the

132 mutants lacking multiple chlorosome proteins of the CsmB/CsmF family or the CsmC/CsmD family (the csmB csmF, csmB csmF csmH, csmC csmD csmH and csmC csmD csmE csmH mutants). The evidence supports the hypothesis that chlorosome proteins are functionally related by containing similar structure motifs and that these proteins may substitute for one another when a given protein is inactivated. In addition, it is clearly shown that inactivation of multiple chlorosome proteins affects the growth rates by partially inhibiting the biosynthesis of BChl c and/or carotenoids, which impairs the light-harvesting ability of the cells. These inhibitory effects may arise from defects in chlorosome biogenesis. Chlorosomes were isolated from 2 L batch of cultures by sucrose gradient ultracentrifugation. Each mutant strain was able to develop chlorosomes of similar density as the wild-type chlorosomes, indicating that neither the CsmC/CsmD motif nor the CsmB/CsmF motif was absolutely required for chlorosome assembly. However, multiple phenotypic effects were detected in the mutant chlorosomes relating to their pigment contents, absorption properties and morphological properties. Pigment content analysis by HPLC showed that the carotenoid contents for all carotenoid species were decreased by ~25% in the csmB csmF and csmB csmF csmH chlorosomes. These results were in agreement with the observation that the carotenoid content was reduced by 25% in chlorosomes of the csmB mutant (Frigaard et al., 2004a). Thus, the CsmB protein is a possible candidate for playing a role in carotenoid incorporation in the chlorosomes. In previous studies with the single csm mutants, the absorption maximum of the csmC chlorosomes was blue-shifted by about 7 nm, and the dimensions of the csmC and csmH chlorosomes were decreased by 20-25% compared to the wild type chlorosomes (Frigaard et al., 2004a). Similar absorbance shifts and size variations were detected in the chlorosomes from the mutants lacking multiple proteins of the CsmC/CsmD family. The absorption maximum of the csmC csmD and csmC csmD csmE chlorosomes was blue- shifted by 12 nm to 738 nm, and the absorption maximum of the csmC csmD csmH and csmC csmD csmE csmH chlorosomes were blue-shifted by 5 nm to 745 nm. The mutant chlorosomes were about 50% smaller than the wild-type chlorosomes, and the length-to- width ratios were significantly reduced for the csmC csmD and csmC csmD csmE chlorosomes. These data suggest that the BChl c aggregation is significantly altered in the

133 chlorosomes from the mutants lacking the CsmC/CsmD motif, and that the proteins CsmC, CsmD and CsmH may play roles in facilitating BChl c organization. Interestingly, mutation of csmH seems to affect the effects of mutations of csmC and csmD, by restoring the shape of chlorosomes to something more similar to wild type. Moreover, the BChl c amounts were relatively low in the csmC csmD csmH and csmC csmD csmE csmH chlorosomes compared to the carotenoid contents, BChl a contents and quinone contents. Given the fact that the cellular BChl c amounts were greatly decreased in these two mutants, it is likely that the mutants lacking the CsmC/CsmD motif were impaired in incorporating BChl c molecules into the chlorosomes, and the reduced rate of BChl c incorporation feedbacks to BChl c biosynthesis to inhibit further synthesis. Since there is little apparent gene regulation at the transcriptional level in C. tepidum, it is not likely that the chlorosome proteins regulate BChl c biosynthesis directly. Chlorosomes of the other two mutants, csmB csmF and csmB csmF csmH, had red-shifted absorption maxima at 756 and 753 nm, and the csmB csmF chlorosomes were ~40% larger than the wild-type ones and had an increased length-to-width ratio. Thus, CsmB and CsmF together, may also be involved in BChl c organization. Differently from CsmC and CsmD, the absence of which results in a spectral blue-shift and reduced chlorosome size, the absence of CsmB and CsmF causes a red-shift of the absorption maximum and an increase of the chlorosome size. The fluorescence emission spectra of all mutant chlorosomes were similar to that of the wild type, indicating that the energy transfer from BChl c to BChl a was not significantly affected. Little information is known about the biogenesis of chlorosomes. It is not clear what components are required at the beginning, or in which order the chlorosome components are assembled. In this study, various subcellular fractions of the mutant cultures were recovered during the chlorosome isolation procedure, and these provided some clues about chlorosome biogenesis. An orange-colored fraction banded at the top of the suspensions of all the six mutants when the chlorosome-enriched fractions were pelleted by the first ultracentrifugation. This orange fraction contained CsmA and CsmB, small amounts of BChl c and BChl a, and large amounts of carotenoids. This fraction likely represents a small portion of the total chlorosomes before or at the beginning of BChl c incorporation, and it suggests that assembly of the chlorosome probably starts

134 with an empty chlorosome envelope containing only a few chlorosome proteins and pigment molecules. Notably, CsmA seems to be essential for chlorosome development as well as cell survival. As the most abundant chlorosome protein in C. tepidum, CsmA is the only chlorosome protein that cannot be inactivated by natural transformation. Although it has 49% sequence identity to CsmA, CsmE seems to be functionally unimportant in the chlorosome biosynthesis. The two mutants lacking CsmE (csmC csmD csmE and csmC csmD csmE csmH) exhibited similar phenotypes as the csmC csmD and csmC csmD csmH mutants containing CsmE. This is probably due to the relatively small content of CsmE in the chlorosome envelope (~5% that of CsmA). The chlorosome- enriched fractions were further purified on sucrose gradients by over-night ultracentrifugation. An additional green layer was separated from the chlorosome fraction due to higher densities on the gradients of the csmC csmD csmH, csmC csmD csmE csmH, csmB csmF and csmB csmF csmH mutants. The amount of BChl c contained in the second fraction was about 5-7% of that contained in the chlorosomes. The second fraction of the csmC csmD csmH mutant contained small, rod-shaped chlorosomes that were only 15% in size compared to the wild-type chlorosomes. The presence of these very small chlorosomes supports the conclusion above that the csmC csmD csmH mutant is impaired in pigment incorporation into chlorosomes, and suggests the CsmC, CsmD and CsmH proteins are involved in longitudinal extension of chlorosomes at the early stage of chlorosome biogenesis. The second fraction of the csmB csmF mutant contained relatively normal sized chlorosomes with membrane fragments. The presence of membrane contaminants suggests that a tighter association exists between the chlorosomes and the membranes, and that this association is not easy to break under the isolation conditions currently applied with the chaotrope 2 M NaSCN. Cross-linking experiments also suggested that the CsmB and CsmF proteins are most likely located on the sides of the baseplate region, which interacts with the FMO proteins and faces the cytoplasmic membrane (Chapter 2.4.2). As a supplement to the studies in mutants lacking chlorosome proteins, protein components were analyzed for the chlorosomes isolated from multiple mutant strains lacking specific enzymes in the BChl c and carotenoid biosynthesis pathways. These chlorosomes contained modified pigment contents as listed in Table 4-4. Of all ten kinds

135 of chlorosome proteins, CsmC is most readily affected by the altered pigment contents. The CsmC content was decreased in the three mutants lacking a Mg chelatase subunit (bchT, bchH and bchS), in the four mutants lacking BChl c methyltransferases (bchU, bchQ bchU, bchR bchU and bchQ bchR bchU), and in the two mutants lacking phytoene synthase or phytoene dehydrogenase (crtB and crtP). The mutants lacking Mg chelatase subunits contained less BChl c, and the mutants lacking BChl c methyltransferases contained homologs of BChl d instead of BChl c. The contents of CsmC, CsmH and the iron-sulfur proteins (CsmI, CsmJ and CsmX) were all decreased in these mutant chlorosomes. These data demonstrate that decrease of BChl c contents may also result in reduced levels of some chlorosome proteins. Many contaminants were revealed by silver staining in the chlorosomes of the bchQ bchU, bchR bchU and bchQ bchR bchU mutants. This could due to incomplete separation of chlorosomes and the attached membranes, which could be caused by significant alteration of pigment contents and chlorosome biogenesis. The crtB and crtP mutants synthesized no carotenoids or only phytoene. Although previous studies suggested that CsmB is the most likely candidate for carotenoid incorporation into chlorosomes, the contents of CsmB in the mutant chlorosomes of crtB and crtP were not significantly different from the wild type. In summary, the absence of multiple chlorosome proteins of the same motif family caused large effects on pigment biosynthesis and chlorosome assembly. Pigment incorporation, BChl c biosynthesis and BChl c organization were significantly altered in the mutant chlorosomes lacking proteins of the CsmC/CsmD and/or the CsmB/CsmF family. These chlorosome proteins, by facilitating pigment incorporation and longitudinal extension, play important roles in the biogenesis of chlorosomes.

136 Table 4-1. Growth rates of the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif under different light intensities. The growth rates shown here are averages of at least two independent measurements.

Growth Light Intensity (μmol photons m-2 s-1) Strain 9 32 80 210 wild type 0.036±0.004 0.214±0.004 0.244±0.005 0.264±0.007 csmC csmD 0.032±0.003 0.200±0.004 0.246±0.007 0.270±0.005 csmC csmD csmE 0.029±0.004 0.206±0.005 0.245±0.011 0.271±0.008 csmC csmD csmH 0.028±0.002 0.184±0.005 0.232±0.008 0.241±0.005 csmC csmD csmE csmH 0.027±0.005 0.189±0.002 0.238±0.002 0.254±0.004 csmB csmF 0.028±0.003 0.202±0.007 0.252±0.005 0.280±0.004 csmB csmF csmH 0.020±0.004 0.198±0.004 0.236±0.002 0.254±0.008

137 ~ 1.0). The levels of ~ 1.0). The levels 600 nm nd standard deviations of at least two mC/CsmD motif or the CsmB/CsmF motif. mC/CsmD motif mM sodium dithionite. to the mid-exponential phase (OD to the mid-exponential -1 s -2 mol photons m reduced conditions with 10 reduced conditions with ype strain and the mutants lacking the Cs the lacking the mutants and ype strain r proteins. All values are the averages a preparations from different cell cultures. ght intensity of about 30 μ Characterization of cells of the wild-t cells of Characterization The cells were grown at li Table 4-2. pigments are shown in mg per g of cellula measurements for two separate chlorosome for two separate chlorosome measurements

*: Fluorescence emission spectra were taken under

138 ing the CsmC/CsmD motif or CsmC/CsmD ing the . The values are the results of one measurement of one measurement . The values are the results c mM sodium dithionite. strain and the mutants lack and the strain reduced conditions with 10 reduced conditions with gments are shown in mg per g of BChl Characterization of chlorosomes isolated from the wild-type isolated from the chlorosomes of Characterization *: Fluorescence emission spectra were taken under Table 4-3. the CsmB/CsmF motif. The levels of pi motif. The levels of the CsmB/CsmF for one chlorosome preparation.

139 and carotenoid biosynthesis pigm ent compositions in the mutant c nding genes inactivated. Genes encoding enzymes involved in BChl strains with the correspo strains Table 4-4.

140 Table 4-5. Lengths, widths and length-to-width ratios of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif, observed by transmission electron microscopy. The values are the averages and standard deviations of 25-40 individual chlorosomes of the same chlorosome preparation.

Strain Length (nm) Width (nm) Length-to-width ratio wild type 165±60 60±25 2.7±0.6 csmC csmD 93±30 46±15 2.1±0.6 csmC csmD csmE 112±30 56±25 2.0±0.5 csmC csmD csmH 104±40 44±15 2.4±0.7 csmC csmD csmE csmH 90±30 35±15 2.6±0.8 csmC csmD csmH layer2 55±10 27±7 2.0±0.4 csmB csmF 210±80 70±20 3.1±0.6 csmB csmF csmH 160±50 70±20 2.4±0.7 csmB csmF layer2 200±45 65±20 3.1±0.6

141

Fig. 4-1. Restriction maps showing the construction and structure of the gene inactivation plasmids.

142

Fig. 4-2. Absorption spectra of whole cells of the wild type (solid line) and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif

(dashed lines). Spectra were recorded for cell suspensions at OD600 nm =

0.1~0.2 and normalized to OD600 nm = 1. The cells were grown at light intensity of 30 μmol photons m-2 s-1 to late exponential phase. Panel A. Wild type and the csmC csmD mutant. Panel B. Wild type and the csmC csmD csmE mutant. Panel C. Wild type and the csmC csmD csmH mutant. Panel D. Wild type and the csmC csmD csmE csmH mutant. Panel E. Wild type and the csmB csmF mutant. Panel F. Wild type and the csmB csmF csmH mutant.

143 Fig. 4-3. Chlorosome isolation from the csmB csmF and csmC csmD csmH mutants. Panel A. Chlorosome fractions on the ultracentrifuged 7-47% sucrose gradient solution. Lane 1: wild-type strain; Lane 2: the csmB csmF mutant; Lane 3: the csmC csmD csmH mutant. Panel B. Silver-stained SDS-PAGE showing the protein composition of the chlorosomes and the second fraction samples. Lane 1: wild-type chlorosomes; Lane 2: the csmB csmF chlorosomes; Lane 2*: the csmB csmF second fraction sample; Lane 3: the csmC csmD csmH chlorosomes; Lane 3*: the csmC csmD csmH second fraction sample. Samples containing 2 μg BChl c were loaded for Lanes 2* and 3*, and samples containing 10 μg BChl c were loaded for Lanes 1, 2 and 3. Panel C. Immunoblot probed with anti-CsmF and anti- CsmA polyclonal antibodies (The anti-F polyclonal antibodies cross-react with both CsmF and CsmB). Lane 1: wild-type chlorosomes; Lane 2: the csmB csmF chlorosomes; Lane 2*: the csmB csmF second fraction sample; Lane 3: the csmC csmD csmH chlorosomes; Lane 3*: the csmC csmD csmH second fraction sample. Samples containing 20 μg BChl c were loaded for Lanes 2* and 3*, and samples containing 100 μg BChl c were loaded for Lanes 1, 2 and 3.

144

145

Fig. 4-4. Protein composition analysis of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. Panel A. SDS-PAGE with proteins detected by silver staining. Chlorosomes containg 10 μg BChl c were loaded for each lane. Panel B. Immunoblot probed with anti-CsmH, anti-CsmC, anti-CsmD and anti-CsmF polyclonal antibodies (The anti-CsmF polyclonal antibodies cross-react with both CsmF and CsmB). Chlorosomes containing 100 μg BChl c were loaded for each lane. Lanes 1 and 8: wild-type chlorosomes; Lanes 2-5: chlorosomes from the csmC csmD, csmC csmD csmE, csmC csmD csmH and csmC csmD csmE csmH mutants; Lanes 6-7: chlorosomes from the csmB csmF and csmB csmF csmH mutants.

146 Fig. 4-5. Absorption spectra (A-H) and fluorescence emission spectra (I) of chlorosomes from the wild type (solid line) and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif (dashed lines) containing the same amounts of monomeric BChl c (OD669 nm in methanol = 1). Panel A. Absorption spectra of the wild-type and the csmC csmD chlorosomes. Panel B. Absorption spectra of the wild-type and the csmC csmD csmE chlorosomes. Panel C. Absorption spectra of the wild-type and the csmC csmD csmH chlorosomes. Panel D. Absorption spectra of the wild-type and the csmC csmD csmE csmH chlorosomes. Panel E. Absorption spectra of the wild-type chlorosomes and the csmC csmD csmH second fraction sample. Panel F. Absorption spectra of the wild-type and the csmB csmF chlorosomes. Panel G. Absorption spectra of the wild-type and the csmB csmF csmH chlorosomes. Panel H. Absorption spectra of the wild-type chlorosomes and the csmB csmF second fraction sample. Panel I. Fluorescence spectra of the wild-type chlorosomes and the csmC csmD csmH second fraction sample.

147

148 Fig. 4-6. Transmission electron micrographs of chlorosomes isolated from the wild-type strain and the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. Panel A. Wild-type chlorosomes. Panel B. The csmC csmD chlorosomes. Panel C. The csmC csmD csmE chlorosomes. Panel D. The csmC csmD csmH chlorosomes. Panel E. The csmC csmD csmE csmH chlorosomes. Panel F. The csmC csmD csmH second fraction sample. Panel G. The csmB csmF chlorosomes. Panel H. The csmB csmF csmH chlorosomes. Panel I. The csmB csmF second fraction sample. The samples were stained with 2% (w/v) aqueous uranyl acetate. Bars represent 200 nm.

149

150

c and

were g BChl

μ

g BChl c bchR bchU μ , e wild-type lane, and lanes. bchQ bchU ated proteins in chlorosomes , atase subunits BchT, BchH and bchU Panel B. Immunoblot detection of were loaded for th were loaded for and bchQ bchR bchU g BChl c , bchR bchU were loaded for the for the d were loaded lacking various BChl and carotenoid biosynthesis enzymes. enzymes. biosynthesis carotenoid BChl and lacking various and bchS* lanes, and chlorosomes containing 100 and chlorosome and chlorosome-associ g BChl bchS μ from mutants lacking the Mg chel the lacking mutants from , bchQ bchU were loaded for the other lanes. c bchU Chlorosomes containg 75 μ g BChl μ mponents separated by SDS-PAGE. Chlorosomes containing 1 were loaded for the Immunoblot detection of were loaded for the d were loaded g BChl c μ methyltransferases. methyltransferases. c g BChl μ lanes, chlorosomes containg 10 chlorosome and chlorosome-associated proteins in chlorosomes in chlorosomes proteins and chlorosome-associated chlorosome BchS. Chlorosomes containing 10 were loaded for the other lanes. Panel C. Protein composition analysis of chlorosomes from mutants mutants from chlorosomes of analysis composition Fig. 4-7. Protein Panel A. Silver staining of chlorosome protein co bchQ bchR bchU BChl the lacking from mutants chlorosomes containing 75 loaded for the bchS and bchS* lanes, chlorosomes containing 10

151

152

Fig. 4-8. Correlation between the growth rates under limiting light intensities and the cellular BChl c contents (mg per g of cellular protein). Panel A. Correlation under light intensity of 9 μmol photons m-2 s-1. Panel B. Correlation under light intensity of 32 μmol photons m-2 s-1. The plotted points represent the various strains of the wild type (wt) or the mutants lacking the CsmC/CsmD motif or the CsmB/CsmF motif. The labels indicate the missing chlorosome proteins for each data point.

153

Chapter 5

Purification and Characterization of Recombinant CsmH, and Pigment Binding Analysis of Recombinant CsmA

154 5.1 Abstract

CsmH is one of the proteins located on chlorosome monolayer envelope in the green sulfur bacterium Chlorobium tepidum. CsmH contains two structural motifs specific to chlorosome proteins: the CsmC/CsmD motif and the CsmB/CsmF motif. When overproduced in Escherichia coli, CsmH is the only one recovered in water-soluble form out of the ten Chlorobium tepidum chlorosome proteins. In this study, the recombinant CsmH proteins, with either amino- or carboxyl-terminal poly-histidine tags, were overproduced in E. coli and purified by affinity chromatography. Biochemical and biophysical characterization indicated that the recombinant CsmH existed in a multimeric form in solution. CsmA is the most abundant chlorosome protein in C. tepidum, and it has been shown to bind BChl a at a one-to-one ratio by detergent extraction experiments (Bryant et al., 2002). The absorption maximum of BChl a is shifted from 770 nm to 798 nm due to binding with CsmA. In this study, binding trials were performed in vitro between the precursor form of CsmA purified from inclusion bodies and BChl a extracted from

Roseobacter sp. Evidence for in vitro binding were obtained based on a shift of the Qy absorption maximum of BChl a observed by absorption spectroscopy.

155 5.2 Introduction

Chlorosomes are unique light-harvesting antennae present in green sulfur bacteria and green nonsulfur bacteria (Blankenship et al., 1995). The core of the chlorosome, consisting mainly of rod-like or lamellar aggregates of BChl c molecules, is surrounded by a galactolipid monolayer envelope stabilized by proteins. At least ten different proteins (CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ and CsmX) were incorporated into the chlorosome envelope in C. tepidum, and the chlorosome proteins contain four specific structural motifs: the CsmA/CsmE motif, the CsmB/CsmF motif, the CsmC/CsmD motif and the adrenodoxin motif (Vassilieva et al., 2000). The CsmH protein is a 21.6 kDa polypeptide with a predicted isoelectric point of 4.72 (Vassilieva et al., 2002b). CsmH is made up of two structural motifs: the N-terminal domain of CsmH is related in sequence to CsmB and CsmF, while the C-terminal domain of CsmH shares similarity to CsmC and CsmD (Vassilieva et al., 2000). CsmH appears to be highly exposed on the chlorosome envelope as shown by the detergent extraction, protease digestion and immunoprecipitation analyses (Vassilieva et al., 2002b). Furthermore, CsmH is the only chlorosome protein recovered in water-soluble form after overproduction in Escherichia coli (Vassilieva et al., 2002b). All these features make CsmH the best candidate for structural studies of chlorosome proteins. The amino acid sequence of CsmA is highly conserved within green sulfur bacteria and green nonsulfur bacteria (Wechsler et al., 1985; Chung et al., 1994). Recent studies indicate that CsmA is a BChl a-binding protein in both Chloroflexus aurantiacus and C. tepidum (Sakuragi et al., 1999; Montaño et al., 2003; Bryant et al., 2002). Proteolytic digestion and detergent treatment in chlorosomes of C. aurantiacus provided solid evidence that BChl a is associated with the CsmA protein (Sakuragi et al., 1999). In addition, purified baseplates from the chlorosomes of C. aurantiacus clearly consist of BChl a and CsmA but no other chlorosome proteins (Montaño et al., 2003). Selective protein extraction by hexanol and SDS also demonstrated that CsmA can be released along with BChl a from the chlorosomes of C. tepidum, and quantitative estimates showed that the binding is probably at a one-to-one ratio (Bryant et al., 2002). Due to the

association with CsmA, the characteristic Qy absorbance of BChl a is shifted from 770

156 nm to 798 nm in both chlorosomes and purified baseplates. The single conserved histidine residue in the CsmA protein probably provides a suitable ligand for binding BChl a. In this chapter, construction, overproduction and purification of amino- and carboxyl-terminal poly-histidine tagged CsmH are described. Biochemical and biophysical characterizations indicated that the recombinant CsmH existed in a multimeric form in solution. Crystallization trials were made with the CsmH sample, but no crystals were obtained. The precursor form of CsmA was overproduced in E. coli as inclusion bodies, and the protein was purified by gel filtration under denaturing conditions. Pigment binding trials were performed between renatured CsmA and BChl a extracted from Roseobacter sp. The BChl a-CsmA binding was monitored by absorption spectroscopy of the characteristic BChl a Qy absorption. Evidence of in vitro binding were obtained, although the ideal binding conditions remain to be further determined.

157 5.3 Materials and methods

5.3.1 Escherichia coli strains and growth conditions

Recombinant DNA manipulations were performed in Escherichia coli DH5α [genotype - - + F ф 80dlacZ ΔM15 Δ(lacZYA-argF)U169 deoR recA1 endA1 hsdR17 (rK , mK ) supE44 λ- thi-1 gyrA96 relA1 from Bethesda Research Laboratories, Gaithersburg, MD] and Escherichia. coli DH10B [genotype F- mcrA Δ(mrr-hsdRMS-mcrBC) ф 80dlacZ ΔM15 ΔlacX74 deoR recA1 endA1 araD139 Δ(ara, leu)7697 galU galK λ- rpsL nupG from Gibco BRL Products, Gaithersburg, MD]. Protein overproduction was performed using - - - Escherichia coli strains BL21(DE3) [genotype F ompT hsdSB(rB mB ) gal dcm (DE3) - - - from Novagen, Madison, WI], BL21(DE3) pLysS [genotype F ompT hsdSB(rB mB ) gal dcm (DE3) pLysS (CmR) from Novagen, Madison, WI] and BLR(DE3) [genotype F- - - R ompT hsdSB(rB mB ) gal dcm Δ(srl-recA) 306::Tn10 (Tc ) (DE3) from Novagen, Madison, WI]. The cells were grown at 37°C in liquid cultures or on 1.5% (w/v) agar plates of Luria-Bertani medium containing 1% (w/v) bacto-tryptone, 0.5% (w/v) bacto-yeast extract, and 1% (w/v) NaCl, pH 7.0. Cells containing plasmids were selected by addition of appropriate antibiotics at final concentrations of 100 μg ampicillin mL-1 or 30 μg kanamycin mL-1.

5.3.2 Construction of the expression vectors

Manipulations with E. coli cells and DNA fragments were performed as described in Chapter 3.3.1. The expression vectors pET11a/CsmA and pET42b(+)/CsmH encoding the CsmA precursor and the carboxyl-terminal His-tagged CsmH were constructed by previous lab members (Chung, 1995; Milks, 2002). For construction of the vector expressing amino-terminal His-tagged CsmH, the csmH gene from the genomic DNA was modified by PCR with the primers 5’ CAA ATC AAA CCC A/TA TGG CTA CCG AAG AA (NdeI) and 5’ CGA ATG CCT TG/G ATC CGT TCT ATT CA (BamHI) to introduce an NdeI site and a BamHI site at the ends of the gene (mutated bases are highlighted, and introduced restriction sites are underlined). For construction of the

158 vectors expressing mature CsmA, the csmA gene from genomic DNA was modified by PCR with the primers 5’ CCA AAA GGA GGA CAT ATG AGT GGA GG (NdeI) and 5’ CGG ATC CTT AAG AAC CAC GAA GGC TGC CAC C (BamHI) to introduce an NdeI site and a BamHI site at the ends of the gene (mutated bases are highlighted, and introduced restriction sites are underlined). The modified PCR products of csmH and csmA were digested with the appropriate restriction enzymes and ligated to the corresponding restriction sites in plasmids pET28a(+) and pET11a separately (Studier et al., 1990). The constructed expression vectors were used to transform BL21(DE3), BL21(DE3)pLysS or BLR(DE3) cells.

5.3.3 Overproduciton and purification of the recombinant proteins

Overproduction of recombinant proteins was accomplished according to the methods

supplied by Novagen (Madison, WI). Cells were grown at 37°C with shaking until OD600

nm was approximately 0.5-1.0, and protein production was induced by addition of IPTG to a final concentration of 1 mM. Cells were incubated for additional 3 h and harvested by centrifugation at 10,000 x g for 10 min at 4°C. The His-tagged CsmH proteins were purified using Ni-NTA Agarose (Qiagen Inc, Valencia, CA) under native conditions according to manufacturer’s instructions. The cells

in lysis buffer (50 mM NaH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0) were disrupted by three passes through a chilled French pressure cell, and 1 mM PMSF was added after the first passage through the French press. The whole-cell extract was clarified by centrifugation at 13,000 × g for 30 min at 4°C , and the supernatant was mixed with Ni-NTA resin and gently agitated at 4°C for at least an hour to allow chelation to occur. The resin was then loaded onto a gravity-flow column and washed

twice with 8 volume of chilled Wash buffer (50 mM NaH2PO4, 300 mM NaCl, 40 mM imidazole, pH 8.0). The combined protein was finally eluted with 4 volume of elution

buffer containing 50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8.0. Dialysis

was performed in buffer containing 50 mM NaH2PO4 and 300 mM NaCl, pH 8.0 to remove imidazole from the purified protein.

159 The poly-histidine residues were removed from the amino-terminal His-tagged CsmH by protease digestion with thrombin (Novagen, Madison, WI). The reaction was carried out at room temperature for 16 h in cleavage buffer (20 mM Tris-HCl, 150 mM

NaCl, 2.5 mM CaCl2, pH 8.4), with 1 U Thrombin per 8 mg recombinant protein. The digested protein (denoted CsmH2) was further purified by Ni-NTA affinity chromatography and was recovered in the flow-through fraction. To remove the protease from CsmH2, gel filtration chromatography on Sephacryl-200 column was employed. Eluted fractions were analyzed by SDS-PAGE, and fractions containing CsmH2 were concentrated by ultrafiltration through a YM-50 membrane centrifugal concentrator (Amicon, Beverly, MA). The recombinant CsmA protein was purified from inclusion bodies by gel filtration chromatography under denaturing conditions. The cells were disrupted by three passes through a chilled French pressure cell in Tris buffer (20 mM Tris-HCl, 10 mM NaCl, pH 7.0), and the cell lysate was centrifuged at 10,000 × g for 30 min at 4°C to pellet the inclusion bodies. The pellets were solubilized in Tris buffer containing 5 M urea, clarified by centrifugation at 13,000 × g for 20 min, and loaded onto Sephacryl-200 gel filtration column for further purification. Eluates were collected in small fractions and analyzed by SDS-PAGE.

5.3.4 SDS-PAGE, protein staining and immunoblotting analysis

The protein composition was analyzed by Tris-Tricine-buffered SDS polyacrylamide gel electrophoresis (Schägger and van Jagow, 1987). The stacking gel was 4% T and 2.6% C, the resolving gel was 12% or 16% T and 3.3% C. Both the stacking and the resolving gels were buffered with 1.0 M Tris-HCl, pH 8.4 and 0.1% (w/v) SDS. Protein samples were mixed with the denaturation buffer (1x denaturation buffer: 0.1 M Tris-HCl, pH 6.8, 24% (v/v) glycerol, 1% (w/v) SDS, 2% (v/v) β-mercaptoethanol, and 0.02% (w/v) Coomassie Blue G-250) and boiled for 1 min before loaded onto gels. Gels were electrophoresed at constant voltage (100 V) in the running buffer containing 0.1 M Tris, 0.1 M Tricine and 0.1% SDS (w/v).

160 Proteins in polyacrylamide gels were stained with the Coomassie staining solution (30% (v/v) methanol, 14% (v/v) acetic acid, and 0.18% (w/v) Coomassie Brilliant Blue R-250) for 30 min. Destaining of gels was accomplished by gently shaking the gels in destaining solution (20 % (v/v) methanol, 7% (v/v) acetic acid) at room temperature with a paper tower to absorb the excess dye. Immunoblotting analyses were performed as described in Chapter 2.3.4 with antibody dilutions of 1:3,000 for anti-CsmH and 1:7,000 for goat anti-(rabbit-IgG). Protein concentrations were determined by the Coomassie blue assay using the BCA reagents supplied by Pierce Chemical Company (Rockford, IL).

5.3.5 Characterization of recombinant CsmH

The recombinant CsmH was diluted to 50 μM in ddH2O and analyzed by mass spectrometry in Dr. Daniel Jones’s laboratory. Dynamic light scattering studies were performed in Dr. Song Tan’s lab using the DynaPro MS/X (Protein solutions, Charlottesville, VA). The N-terminal His-tagged CsmH at about 2 mM was filtered through a 2.2 μm filter membrane before being applied to the instrument. Data collection was accomplished at 4°C utilizing the DYNAMICS V6 program. Gel filtration chromatography was performed in a Sephacryl-200 column with buffer containing 50

mM NaH2PO4 and 300 mM NaCl (pH 8.0). Protein markers (alcohol dehydrogenase, 150 kDa; bovine serum albumin, 66 kDa; carbonic anhydrase, 29 kDa from Sigma, St. Louis, MO) were loaded onto the gel filtration column together with the N-terminal His-tagged CsmH. For the cross-linking studies, the N-terminal His-tagged CsmH was diluted to 10 μg mL-1 in conjugation buffer (0.1 M MES (2-[N-morpholino]ethane sulfonic acid), pH 5.5), and incubated with 5 mM EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride) at room temperature for 5 min or 1 h. The cross-linked CsmH sample was analyzed on SDS-PAGE and multimers were detected by immunoblotting with anti- CsmH antibodies.

161 5.3.6 Pigment binding analysis of recombinant CsmA

Pigments were extracted from pelleted cells of Roseobacter (a species isolated by Maresca, J.A. in Woods Hole, MA) by adding a small volume of organic solvent to pelleted cells (acetone:methanol 7:2 v/v) and sonicating. The extracted pigments containing mainly BChl a were dried under vacuum and stored at -20°C for further use. The concentration of BChl a was determined by absorption spectroscopy using an absorption coefficient of 76 L g-1 cm-1 at 770 nm (Tsuji et al., 1995). Dialysis of CsmA was performed with 3 kDa cut-off membrane (Spectrum, Houston, TX) in buffer containing 20 mM Tris-HCl, 10 mM NaCl, pH 7.0. The renatured CsmA was mixed with extracted BChl a at a molar ratio of 1:1 and held in aerobic or anaerobic dark conditions at room temperature. Buffer mixed with the same amount of BChl a was used as a control. The binding between CsmA and BChl a was monitored by absorption spectroscopy from 350 nm to 900 nm.

162 5.4 Results

5.4.1 Overproduction and purification of the recombinant proteins

Construction of the expression vectors. The expression vectors pET11a/CsmA and pET42b(+)/CsmH encoding the CsmA precursor and the C-terminal His-tagged CsmH were constructed by previous lab members (Chung, 1995; Milks, 2002). To express N- terminal His-tagged CsmH, the pET28a(+) vector with His-tag coding sequence was employed. To express mature CsmA, a stop codon was introduced immediately after the serine residue at position 59 by the csmA primer (5’ CGG ATC CTT AAG AAC CAC GAA GGC TGC CAC C). The csmH and csmA genes were modified and amplified by PCR, digested with the appropriate restriction enzymes NdeI and BamHI, and inserted into pET28a(+) or pET11a vectors separately (Studier et al., 1990). The nucleotide sequences of constructed vectors pET28a(+)/CsmH and pET11a/CsmAm were verified by sequencing analysis using primers for the T7 promoter and terminator. The deduced amino acid sequences of C- and N-terminal His-tagged CsmH, precursor and mature forms of CsmA are shown in Fig. 5-1.

Overproduction and purification of CsmH and CsmA. E. coli strain BL21(DE3) was transformed with the plasmids pET28a(+)/CsmH, pET42b(+)/CsmH, pET11a/CsmA and pET11a/CsmAm. Cultures were grown with appropriate antibiotics until OD600 nm was between 0.4 and 1.0, and protein expression was induced by addition of 1 mM IPTG. Induction was continued for 3 h on a rotary shaker at 37ºC. The C-and N-terminal His-tagged CsmH were overproduced at a relatively high level as shown in Fig. 5-2 (Panels A and B, Lane 2). After cell disruption and centrifugation, the CsmH proteins were recovered in the supernatant fraction as water- soluble proteins (Fig. 5-2, Panels A and B, Lane 4), and purified by affinity chromatography as shown in Fig. 5-2 Panels A and B, Lane 5. A polypeptide band of ~14 kDa was also found in the eluted C-terminally His-tagged CsmH, and the band was confirmed by immunoblotting analysis to be a CsmH degradation product. Thrombin digestion was performed to remove the poly-histidine extension from the N-terminal His-

163 tagged CsmH protein. Thrombin cleaves immediately after the arginine residue in the peptide sequence LVPRGS. The resulting product, CsmH2, was separated from the poly- histidine peptide and the protease by affinity chromatography and gel filtration chromatography. With a smaller molecular weight, CsmH2 electrophoresed slightly faster than the His-tagged CsmH on SDS-polyacrylamide gel. (Fig. 5-2, Panel B, Lane 6). Overproduction and purification of the precursor form of CsmA is shown in Fig. 5-2 Panel C. As shown in Lane 2, the CsmA precursor with an apparent mass of 8.5 kDa was expressed in significant amounts in the E. coli cells. After cell disruption and centrifugation, the CsmA precursor was recovered in the pellet fraction as inclusion bodies (Fig. 5-2, Panel C, Lane 3). Interestingly, a protein species of about 7.0 kDa was detected in the pellet fraction along with the CsmA precursor. The 7.0 kDa protein was also observed in previous CsmA overproduction studies, and was verified by N-terminal sequencing to be CsmA with the first methionine removed (Chung, 1995). For purification of the CsmA precursor, the inclusion bodies were solubilized in denaturing buffer containing 5 M urea, clarified by centrifugation, and purified by gel filtration chromatography on Sephacryl S-200 column (Fig. 5-2, Panel C, Lane 5). Expression trials for the mature form of CsmA were performed in E. coli strains BL21(DE3), BL21(DE3)pLysS and BLR(DE3). However, the mature CsmA at 6.2 kDa was not detected on SDS polyacrylamide gel by silver staining in any of the above strains. Immunoblotting analysis was performed with the anti-CsmA antibodies. A protein band at 8 kDa was detected in the pelleted cells as well as the LB medium before and after IPTG induction. This protein was likely to be an E. coli protein species that cross-reacted nonspecifically with anti-CsmA antibodies. It is not clear why CsmA cannot be overproduced in its mature form. It is possible that the mature CsmA has some toxicity effects on E. coli, or the mature CsmA was degraded so rapidly in E. coli that the protein could not be detected.

5.4.2 Characterization of recombinant CsmH

Mass spectrometry was utilized to determine the molecular mass of the recombinant CsmH proteins. With molecular masses of 22806 and 23808 Dalton, the C- and N-

164 terminal His-tagged CsmH matched perfectly with the deduced sequences except that the first methionine was removed (Fig. 5-1). Dynamic light scattering (DLS) was performed to detect the hydrodynamic radius as well as the homogeneity of the N-terminal His- tagged CsmH sample (Fig. 5-3). In DLS, scattering intensity fluctuations are monitored across μs time scales and then correlated. The intensity fluctuations reflect the particle motion, and the measured property in the correlation analysis reflects the distribution of diffusion coefficients. The particle size is then calculated using the Stokes-Einstein equation,

kT

RH = 6πηD

where RH is the hydrodynamic radius, k is the Boltzmann constant, T is the temperature, η is the solvent viscosity and D is the diffusion coefficient. As shown in Fig. 5-3 B, DLS indicated that the sample was dominated by particles with a hydrodynamic radius of 24 ± 3 nm. The result strongly suggests that CsmH exists in a highly aggregated form in the solution. With a Pd (polydispersity) value of about 45.2%, the CsmH sample was an ensemble of multimeric states or non-specific aggregates. To verify the molecular mass of the aggregated CsmH, gel filtration chromatography was performed on Sephacryl-200 column with buffer containing 50 mM

NaH2PO4 and 300 mM NaCl (pH 8.0). Samples containing alcohol dehydrogenase (150 kDa), bovine serum albumin (66 kDa), carbonic anhydrase (29 kDa) and N-terminal His- tagged CsmH were applied in a narrow zone at the top of the column and eluted through the column by the mobile phase. Molecules in the samples were separated according to the sizes, with the largest molecules eluting first and the smallest last. The eluted fractions were collected and analyzed by SDS-polyacrylamide gel electrophoresis (Fig. 5- 4). A protein species of about 24 kDa was detected in the earliest eluted fractions (Fig. 5- 4, Panel A, Lanes 1-4), and the protein was confirmed to be N-terminal His-tagged CsmH by immunoblotting analysis with anti-CsmH antibodies (Fig. 5-4, Panel B). Thus, the CsmH aggregates eluted earlier than alcohol dehydrogenase and should have a molecular mass greater than 150 kDa.

165 Multimers of CsmH were clearly detected after cross-linking with the zero-length chemical cross-linker EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride) (Fig. 5-5). The N-terminal His-tagged CsmH was diluted to10 μg mL-1 and incubated with 5 mM EDC at room temperature for 5 min or 1 h, then analyzed by SDS-PAGE and immunoblotted with anti-CsmH antibodies. Dimers, trimers, tetramers, and probably pentamers of CsmH appeared after 5 min of cross-linking (Fig. 5-5, Lane 2), but disappeared after 1 h of cross-linking due to the formation of higher aggregation state products (Fig. 5-5, Lane 3). Interestingly, a dimer band of CsmH was also detected in the uncross-linked sample at low intensity (Fig. 5-5, Lane 1).

5.4.3 Pigment binding analysis of recombinant CsmA

Pigments were extracted from cells of Roseobacter sp. (an organism isolated by J.A. Maresca from Woods Hole, MA) by sonicating in organic solvents (acetone:methanol 7:2 v/v). The extracted pigments contained small amounts of carotenoids, phaeophytin and mainly BChl a. Absorption spectra of the extracted pigments are shown in Fig. 5-6, Panel

A. The Qy absorption of the extracted BChl a appeared as a strong peak at 770 nm in acetone:methanol mix (7:2 v/v), but was around 780 nm at a much lower amplitude in

ddH2O due to the limited solubility of BChl a in aqueous solutions. Recombinant CsmA was purified under denaturing conditions and renatured in buffer containing 20 mM Tris-HCl, 10 mM NaCl, pH 7.0 by dialysis. The renatured CsmA, at a concentration of 20 μM, was mixed with the same molar concentration of extracted BChl a and incubated under aerobic or anaerobic dark conditions for 10 days at room temperature. Tris buffer containing no CsmA was also mixed with the same amount of BChl a as a control. The binding between BChl a and CsmA was monitored by absorption spectroscopy from 350 nm to 900 nm (Fig. 5-6, Panels B and C). Green- colored pellets of insoluble BChl a formed at the bottom of the tubes after 10 days incubation, and about 40-80% of the BChl a remained in the solution as indicated by the

Qy absorption intensities. After aerobic incubation, the BChl a Qy absorption were observed at 778 nm and 774 nm separately for the binding and the control sample. The maximum difference between the two samples was probably due to the association

166

between BChl a and CsmA which resulted in a red-shift of the BChl a Qy absorption. A peak at 795 nm clearly appeared when the absorption spectrum of the control sample was

subtracted from that of the binding one. After anaerobic incubation, the Qy absorption maximum of the BChl a was shifted to 835 nm for both the binding and the control samples, which might be a consequence of BChl a dimerization. Interestingly, BChl a appeared to be more soluble in aqueous solutions when CsmA was present under either incubation condition.

167 5.5 Discussion

Chlorosomes exhibit multiple characteristic features that are not common in the light harvesting organelles of other photosynthetic organisms. The BChl c molecules in chlorosomes are self-organized into aggregates by pigment-pigment interactions instead of pigment-protein interactions, and they are enclosed by a monolayer of monogalactosyl diglyceride with the polar head groups pointing outside and the fatty acid tails sticking inside (Staehelin et al., 1978; Staehelin et al., 1980; Holo et al., 1985). The chlorosome proteins are incorporated into the monolayer envelope membrane. At least some portions of the chlorosome proteins are exposed to the chlorosome surface, as indicated by protease digestion, immuno-agglutination and gold labeling electron microscopy (Wullink et al., 1991; Chung and Bryant, 1996b; Vassilieva et al., 2002b). Detergent treatment of chlorosomes showed that CsmH was easily extracted from the chlorosome envelope, and immuno-agglutination studies indicated that chlorosomes could be agglutinated very efficiently by anti-CsmH antibodies (Vassilieva and Bryant, 1998; Vassilieva et al., 2002b). Both results suggest that CsmH is highly exposed at the chlorosome envelope surface. The amino- and carboxyl-termini of CsmH share sequence similarity to two structural motifs that have only been identified in chlorosome proteins: the CsmB/CsmF motif and the CsmC/CsmD motif (Vassilieva et al., 2000). Beyond that, little structural information is known about CsmH. Secondary structural predictions with the PredictProtein Internet Server (Rost et al., 2004) suggests that 6 helixes exist in CsmH, including the amino acid residues 30-67, 75-92, 100-124, 129-144, 162-166 and 181-187. The protein sequence shows low probability of membrane association throughout, except for a short region WTGMISNLNAMV from position 113 to position 124. The amino- and carboxyl-terminal sequences (about 25-30 amino acid residues) are predicted to be a loop/coil structure readily accessible to the solvent. No cysteine is contained in CsmH, and thus disulfide bonds are not involved in the tertiary or quaternary structure. Structural studies of chlorosome protein would likely lead to a greater understanding of the chlorosome and might also define a new class of proteins inserted into a lipid monolayer instead of a bilayer membrane. All the chlorosome proteins from C.

168 tepidum have been overproduced in E. coli λDE3 lysogen cells by IPTG induction. CsmH (as its natural form) was the only protein that could be easily overproduced as a water- soluble protein instead of inclusion bodies. Proteins sharing sequence similarity to domain of CsmH (CsmB, CsmF, CsmC and CsmD) were all recovered as inclusion bodies and exhibited low solubility after denaturation and renaturation attempts (Chung, 1995; Vassilieva et al., 2002b). In this study, CsmH was fused with an amino- or carboxyl-terminal poly-histidine tag, overproduced in E. coli and purified by affinity chromatography for structural analysis. However, biophysical and biochemical characterization revealed that CsmH formed aggregates in solutions. Dynamic light scattering showed a hydrodynamic radius of ~23.8 nm for the CsmH aggregates, and the deduced molecular mass was more than 5 mDa assuming the aggregates had a sphere shape. Gel filtration chromatography confirmed that the molecular mass of the CsmH aggregates was more than 150 kDa. Direct evidence of aggregation was obtained from cross-linking experiments with the zero-length cross-linker EDC. Under the experimental conditions using a concentration of 10 μg protein mL-1, the yields of intramolecular cross-linking should have greatly exceeded the yields of intermolecular cross-linking, and only polypeptides situated next to each other could be linked together (Wong, 1991). CsmH multimers including dimers, trimers, tetramers and pentamers were detected by SDS-PAGE and immunoblotting analysis, which also provides evidence of CsmH aggregation in solution. The reasons why CsmH forms aggregates in solution remains unclear. It is not likely that poly-histidine tag caused protein aggregation, since the CsmH2 protein with His-tag removed by protease digestion behaved as the His-tagged CsmH on gel filtration column, indicating they had similar molecular weights and particle sizes. Other causes for protein aggregation include hydrophobic patches on the surface of the protein, differently charged isoforms, as well as electrostatic interactions (Howard and Brown, 2002). In some cases, protein molecules can be crystallized even from amorphous aggregates if the crystal form is more stable than the nonspecific aggregated form (Dr. K. Murakami, personal communication). Crystal screening is being carried out with the CsmH2 samples. At the same time, trials are being performed with chemical additives such as detergents, chaotropes, electrostatic reagents, alcohols or salts to manipulate sample-solvent

169 interactions and prevent the aggregation. Disorder analysis by DisEMBL (http://dis.embl.de/) suggests that the amino- and carboxyl-terminal residues 1-31 and 189-212 have significantly high disorder probability. These two regions are rich in alanine and may have flexible structure that causes amorphous aggregation. Truncated CsmH without the amino- and carboxyl-terminal extensions may also be overproduced in E. coli and used for crystallization trial. The CsmA proteins, 6.2 kDa in C. tepidum and 5.7 kDa in Chloroflexus aurantiacus, have been well studied in both organisms. Although the amino acid sequence of CsmA in C. tepidum is only distantly related to the one in Chloroflexus aurantiacus (about 30% sequence identity), it is more than 90% conserved among the green sulfur bacteria including Chlorobium vibrioforme, Chlorobium limicola, Chlorobium phaeovibrioides, Pelodictyon luteolum, and Prosthecochloris aestuarii. (Wagner-Huber et al., 1988; Zuber and Brunisholz, 1991). Interestingly, CsmA proteins in both C. tepidum and Chloroflexus aurantiacus are synthesized as precursors of 79 amino acids and they are subsequently processed at their carboxyl-termini to mature proteins of 52 residues (Chloroflexus aurantiacus) or 58 to 59 residues (C. tepidum). The carboxyl-terminal processing is not unusual for bacterial proteins but usually occurs in the periplasm (Chung et al., 2002), and the similarity of this feature in the two groups of green bacteria is probably significant. The CsmA protein has recently been proven to be the BChl a-binding protein by proteolytic digestion and detergent treatment of chlorosomes (Sakuragi et al., 1999; Montaño et al., 2003; Bryant et al., 2002). The amino acid sequence Gly-His-Trp with its surrounding hydrophobic regions is conserved in the CsmA proteins in both Chlorobium and Chloroflexus species. The conserved histidine residue is a suitable ligand for binding BChl a, as seen in other BChl a-binding

photosynthetic proteins (Blankenship and Matsuura, 2003). The Qy absorption of BChl a is red-shifted from 770 nm to 795 nm in this complex, probably due to pigment-protein interactions. Located on the bottom side of the chlorosome envelope and facing the cytoplasmic membrane (in a structure denoted as the baseplate), the BChl a-CsmA complex mediates energy transfer from BChl c to the BChl a of the FMO protein and the reaction centers (Gerola and Olson, 1986).

170 In this study, pigment binding analysis was carried out in vitro with E. coli- overproduced CsmA and Roseobacter sp.-extracted BChl a. The genes encoding the CsmA precursor or mature CsmA of C. tepidum were cloned into pET vectors and overexpressed in E. coli cells by IPTG induction. The CsmA precursor was expressed as inclusion bodies at a significant level (50 mg L-1) in E. coli BL21(DE3) strain, and purified under denaturing conditions by gel filtration chromatography. However, expression of mature CsmA was not detected in E. coli strains BL21(DE3), BL21(DE3) pLysS (which inhibits the background expression of the target protein) or BLR(DE3) (which helps to stabilize the target plasmids). The reasons for different expression patterns of CsmA precursor and mature CsmA are not clear. The conformation of the CsmA protein might be greatly altered without the carboxyl-terminal sequence and might make the mature form of CsmA more toxic to the expression host or make it more readily degraded than the precursor form. The mixture of BChl a and CsmA precursor was incubated under aerobic or anaerobic conditions and monitored by absorption spectroscopy. The presence of CsmA enhanced the BChl a solubility by about 50% under

both conditions. After aerobic incubation, the BChl a Qy absorption in the binding sample was red-shifted by 4 nm from the control one. Subtraction of the absorption spectrum of

the control sample from that of the binding one revealed a BChl a Qy absorption peak at

795 nm, which matched perfectly with the BChl a Qy absorption in isolated baseplates. Under anaerobic conditions, BChl a dimerization happened rather than BChl a-CsmA association. Further binding trials will be made with low concentration of detergents, alcohols or additional pigments (such as carotenoids) to increase the BChl a solubility as well as stability to determine the most suitable binding conditions.

171 Fig. 5-1. The deduced amino acid sequences of encoded proteins from the expression vectors. His-tags are underlined. Asterisks indicate the C- terminal end of the deduced polypeptide sequences. C-CsmH, the C- terminal His-tagged CsmH encoded by pET42b/CsmH; N-CsmH, the N- terminal His-tagged CsmH encoded by pET28a/CsmH; CsmA, the precursor form of CsmA encoded by pET11a/CsmA; CsmAm, the mature form of CsmA encoded by pET11a/CsmAm.

172 10 20 30 40 C-CsmH MATEETNMPAAEAPKAAAGAPN N-CsmH MGSSHHHHHHSSGLVPRGSHMATEETNMPAAEAPKAAAGAPN

50 60 70 80 C-CsmH TSAGNGDMAHLIGNMGILIDSTIESVQGVISTVSSATGQIIEGVTTT N-CsmH TSAGNGDMAHLIGNMGILIDSTIESVQGVISTVSSATGQIIEGVTTT

90 100 110 120 130 C-CsmH INSEPVKEIINNVNSVSGQIIEGVTNTLKSEQIQNSFNELGKFWTGM N-CsmH INSEPVKEIINNVNSVSGQIIEGVTNTLKSEQIQNSFNELGKFWTGM

140 150 160 170 180 C-CsmH ISNLNAMVNSNQVKNLFDNVSAGINQLAGGIFPQGMPPMFMGAS N-CsmH ISNLNAMVNSNQVKNLFDNVSAGINQLAGGIFPQGMPPMFMGAS

190 200 210 220 C-CsmH SGEEKRKVVHQIPVVHTSESGAATLKAMTPQPTAAPAAPAAPAAP N-CsmH SGEEKRKVVHQIPVVHTSESGAATLKAMTPQPTAAPAAPAAPAAP

230 240 C-CsmH KNKPELEHHHHHHHH* N-CsmH KNKPEGE*

10 20 30 40 CsmA MSGGGVFTDILAAAGRIFEVMVEGHWETVGMLFDSLGKGTMRIN CsmAm MSGGGVFTDILAAAGRIFEVMVEGHWETVGMLFDSLGKGTMRIN

50 60 70 CsmA RNAYGSMGGGSLRGSSPEVSGYAVPTKEVESKFAK* CsmAm RNAYGSMGGGSLRGS*

173 Fig. 5-2. Overproduction and purification of recombinant C. tepidum CsmH and CsmA in E. coli. Panel A. C-terminal His-tagged CsmH. Panel B. N-terminal His-tagged CsmH. Panel C. the precursor form of CsmA. Lane 1: whole-cell extract of BL21(DE3) cells transformed with pET42b/CsmH (for Panel A), pET28a/CsmH (for Panel B) and pET11a/CsmA (for Panel C) before IPTG induction; Lane 2: whole-cell extract of BL21(DE3) cells transformed with pET42b/CsmH (for Panel A), pET28a/CsmH (for Panel B) and pET11a/CsmA (for Panel C) after IPTG induction for 3 h; Lane 3: pellets after cell disruption and centrifugation; Lane 4: supernants after cell disruption and centrifugation; Lane 5: purified proteins after Ni-NTA affinity chromatography (for Panels A and B) or gel filtration chromatography (for Panel C); Lane 6: CsmH2 obtained after Thrombin digestion, Ni-NTA affinity chromatography and centrifugal concentration to remove the poly-histidine peptide and protease.

174

175

Fig. 5-3. Light scattering of the N-terminal His-tagged CsmH. Panel A. The light-scattering autocorrelation function curve. Panel B. The hydrodynamic radius.

176

Fig. 5-4. Gel filtration chromatography of N-terminal His-tagged CsmH and protein markers on Sephacryl-200 column. The protein markers include alcohol dehydrogenase (AD, 150 kDa), bovine serum albumin (BSA, 66 kDa) and carbonic anhydrase (CA, 29 kDa). Panel A. Coomassie blue stained SDS polyacrylamide gel showing the protein compositions in the eluted fractions numbered 1 to 9. Panel B. Immunoblot probed with anti-CsmH polyclonal antibodies of the eluted fractions 1 to 9.

177

Fig. 5-5. Purified N-terminal His-tagged CsmH (10 μg/mL) cross-linked with 5 mM EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride) and detected by immunoblotting with the anti-CsmH antibodies. Lane 1: the uncross-linked CsmH; Lane 2: CsmH cross-linked for 5 min; Lane 3: CsmH cross-linked for 1 h.

178

Fig. 5-6. Binding analysis of extracted BChl a and the recombinant CsmA. Panel A. Absorption spectra of extracted BChl a in acetone:methanol (7:2 v/v) (solid line) and ddH2O (dotted line). Panel B. Binding analysis of BChl a and CsmA in aerobic conditions. Solid line, sample containing BChl a and CsmA at a 1:1 ratio; dotted line, control sample containing BChl a only. Panel C. Analysis of BChl a and CsmA binding under anaerobic conditions. Solid line, sample containing BChl a and CsmA at a 1:1 ratio; dotted line, control sample containing BChl a only.

179

Chapter 6

Summary of Organization and Function of Chlorosome Proteins in Chlorobium tepidum

180 Photosynthetic organisms utilize light-harvesting antennae to absorb light and transfer light energy to reaction centers. Green photosynthetic bacteria, which usually live in environments with extremely low light intensities, contain an antenna complex known as the chlorosome which is well adapted to harvest light energy under these conditions (Blankenship et al., 1995). Differing from all other antenna complexes, in which the chlorophyll and other pigments are specifically associated with proteins, the chlorosome contains self-organized BChl pigments, and its proteins appear to play only minor roles in the organization of the pigments. The absorption maximum is shifted far to the red due to direct and strong pigment-pigment interactions that lead to rapid, exciton- coupled energy transfer. The protein composition of chlorosomes was controversial in the 1980s and early 1990s. During this time, the chlorosomes of green non-sulfur bacteria and chlorosomes of green sulfur bacteria were considered as though they were identical, and chlorosomes were isolated by different procedures in the presence of detergents (Feick and Fuller, 1984; Griebenow and Holzwarth, 1989; Holzwarth et al., 1990; Stolz et al., 1990). Later studies demonstrated that chlorosomes of the green nonsulfur bacteria and chlorosomes of the green sulfur bacteria have distinctly different protein compositions and that the use of detergents causes severe loss of chlorosome proteins (Blankenship et al., 1995; Vassilieva et al., 2000; Bryant et al., 2002). Gerola and Olson developed an isolation procedure with a chaotropic agent (NaSCN) to release chlorosomes from the membrane and obtained a reproducible preparation of chlorosomes (Gerola and Olson, 1986). Using this method, Chung et al. first showed that the chlorosomes of Chlorobium tepidum contained at least ten distinct protein species, named CsmA, CsmB, CsmC, CsmD, CsmE, CsmF, CsmH, CsmI, CsmJ and CsmX (Chung et al., 1994). A rough estimate of protein copy numbers was made based upon the protein mass and the staining intensities of protein bands on SDS-PAGE (Frigaard et al., 2004). There are about 2,700 copies of CsmA, 1,000 copies of CsmB, 300 copies of CsmC and CsmD, 200 copies of CsmE and CsmF, 100 copies of CsmH, CsmI, CsmJ, and 20 copies of CsmX per chlorosome. Thus, an average chlorosome envelope contains about 5,000 protein molecules. CsmA (6.2 kDa) comprises about one-third to one-half of the total chlorosome proteins in chlorosomes of C. tepidum (Bryant et al., 2002). CsmA is synthesized as an

181 8.3 kDa precursor (pre-CsmA) and subsequently processed at the carboxyl-terminus to the mature form (Chung et al., 1994). The amino acid sequence of CsmA is moderately conserved in the green nonsulfur bacterium Chloroflexus aurantiacus but highly conserved in the green sulfur bacteria (Wagner-Huber et al., 1988; Zuber and Brunisholz, 1991; Table 2-2). CsmA is not easily extracted from the chlorosome envelope by low concentrations of ionic or nonionic detergents and is only accessible to protease digestion after addition of 0.1% Triton X-100 (Chung and Bryant, 1996b; Bryant et al., 2002). Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI- TOF-MS) suggests that the N-terminal portion of CsmA (residues 2-38, mainly hydrophobic) is not accessible to proteases and is most likely buried in the envelope membrane (Milks et al., 2005). Although originally proposed to be a BChl c-binding protein, CsmA has been identified as the BChl a-binding protein of chlorosomes by proteolytic digestion and detergent treatments in both green sulfur bacteria and green nonsulfur bacteria (Sakuragi et al., 1999; Bryant et al., 2002; Montaño et al., 2003). Cross-linking studies presented here demonstrate that CsmA forms many distinct homo- multimer species and that CsmA interacts with the FMO protein. These results suggest that CsmA is homogeneously organized as a paracrystalline array and that it is localized on the bottom side of the chlorosome envelope that faces the FMO protein layer and the cytoplasmic membrane (Chapter 2.4.2). Interactions between CsmA and pre-CsmA most likely occur on the edges of the CsmA baseplate, where pre-CsmA is initially inserted into the chlorosome envelope (Chapter 2.4.2). The BChl a-CsmA complex mediates energy transfer from BChl c aggregates in the chlorosome to the reaction center on the cytoplasmic membrane (Chapter 2.5). Inactivation of CsmA has not been successful in C. tepidum, and CsmA might be required for the viability of the cells in green sulfur bacteria (Chung, 1995; unpublished data). CsmE is similar to CsmA in the amino acid sequence and in carboxyl-terminal processing (Vassilieva et al., 2000). CsmE is present in a relatively low amount on the chlorosome envelope (~5% of the amount of CsmA) (Chung and Bryant, 1996b), and mutant strains lacking CsmE did not show any growth defect (Chapter 4.4.2). CsmE is certainly not an essential chlorosome component, and while it is not clear if CsmE binds

182 BChl a as CsmA does, this certainly seems to be likely given the high degree of sequence similarity between the two proteins. CsmB is the second most abundant protein in chlorosomes of C. tepidum (Chung and Bryant, 1996a). CsmF shares 63% sequence similarity to CsmB, but CsmF is present at a much lower copy number per chlorosome (Vassilieva et al., 2000; Frigaard et al., 2004a). Hydrophilicity analyses of CsmB and CsmF indicate that the proteins are slightly hydrophobic throughout nearly their entire length (Højrup et al., 1991; Chung and Bryant, 1996a). Although CsmB is not readily digested with trypsin, anti-CsmB antibodies are highly effective in agglutinating whole chlorosomes; this suggests that some portion of CsmB is exposed to solvent in isolated chlorosomes (Chung and Bryant, 1996a). In cross- linking experiments, both CsmB and CsmF can be cross-linked to the CsmA species (CsmA or pre-CsmA), suggesting that they are present on the edge of the CsmA baseplate (Chapter 2.4.2). In addition to the interaction with CsmA species, CsmB can be cross-linked to nearly all other chlorosome proteins (Chapter 2.5). Thus, it is most likely that CsmB is distributed over much of the chlorosome envelope surface except for the baseplate region, from which it would be excluded by the paracrystalline interactions of CsmA. Single mutants lacking CsmB or CsmF, double mutants lacking CsmB and CsmF and triple mutants lacking CsmB, CsmF, and CsmH were constructed (Chapter 4.4.1). The carotenoid contents were decreased by 25% in the chlorosomes of the csmB, csmB csmF and csmB csmF csmH mutants, and CsmB might play a role in carotenoid incorporation into the chlorosomes (Chapter 4.4.3; Chapter 4.4.5). Chlorosomes of the csmB csmF and csmB csmF csmH mutants had absorption maxima that were red-shifted by 3 to 6 nm, and the csmB csmF chlorosomes were ~40% larger than the wild-type ones and had an increased length-to-width ratio. Thus, CsmB and CsmF together may also play roles in BChl c organization (Chapter 4.4.5; Chapter 4.4.6). CsmC, CsmD and CsmH can be easily extracted from the chlorosome envelope by low concentration of detergents (Vassilieva and Bryant, 1998; Bryant et al., 2002). In addition, CsmC and CsmD are quite readily accessible to various proteases, and antisera to CsmC, CsmD and CsmH are all highly effective at agglutinating chlorosomes (Chung and Bryant, 1996b; Vassilieva et al., 2002). The above results suggest that significant portions of CsmC, CsmD and CsmH are exposed at the chlorosome envelope surface. In

183 cross-linking experiments, CsmC and CsmD are the two protein species that are most readily cross-linked (Chapter 2.4.3). CsmC and CsmD can be rapidly cross-linked into homo-dimers and homo-trimers, and homo-tetramers of CsmC are also detected after longer cross-linking times. These proteins are most likely to be localized along the sides or the cytoplasmic surface of chlorosomes opposite to the baseplate (Chapter 2.5). CsmH does not form multimers, but this protein can be cross-linked to both CsmA and CsmB (Chapter 2.4.5). These results suggest that CsmH is localized along the periphery of the baseplate (Chapter 2.5). Double and triple mutants lacking CsmC and CsmD or CsmC, CsmD, and CsmH were also constructed (Chapter 4.4.1). The cellular BChl c amounts were greatly decreased in the triple mutant strains csmC csmD csmH and csmC csmD csmE csmH and slightly reduced in the double mutant strains csmC csmD and csmC csmD csmE (Chapter 4.4.3). The absorption maximum was blue-shifted by 12 nm in the chlorosomes of the double mutants and by 5 nm in the chlorosomes of the triple mutants, and the mutant chlorosomes were ~50% smaller than the wild-type ones and had modified length-to-width ratios (Chapter 4.4.5; Chapter 4.4.6). These data suggest that the BChl c aggregation is significantly altered in the chlorosomes of the mutants lacking CsmC, CsmD and CsmH, and that CsmC, CsmD and CsmH play roles in facilitating BChl c incorporation into the chlorosomes and BChl c organization (Chapter 4.5). It is likely that the mutants lacking CsmC, CsmD and CsmH are somehow impaired in the incorporation of BChl c molecules into the chlorosomes, and the slow-down of BChl c incorporation probably feeds back to inhibit BChl c biosynthesis (Chapter 4.5). CsmI, CsmJ and CsmX are related in sequence to [2Fe-2S] ferredoxins at their amino termini, and CsmI and CsmJ overproduced in E. coli exhibited the characteristic EPR spectra of [2Fe-2S] ferredoxins. Antibody agglutination studies suggest that the surface-exposed regions of these three proteins might be relatively small compared to their overall sizes (Vassilieva et al., 2002b). Cross-linking studies showed that CsmI and CsmJ homo-dimers and hetero-dimers were readily formed during cross-linking with EDC. The most reasonable interpretation of this observation is that these proteins form hetero-tetramers (CsmI2)(CsmJ2) in wild-type chlorosomes but only homo-dimers when one protein is missing in the csmI and csmJ mutants. CsmI, CsmJ and CsmX each could be cross-linked to one or two molecules of CsmB. Single, double and triple mutants

184 lacking CsmI, CsmJ and CsmX were constructed in all combination (Chapter 3.4.1). The mutants were characterized by fluorescence emission spectroscopy, and CsmI and CsmJ were found to participate in the redox regulation of energy transfer in both isolated chlorosomes and whole cells (Chapter 3.4.3). Under oxygen stress conditions, CsmI and CsmJ might transfer electrons from the quencher within the chlorosome (most probably chlorobiumquinone) to molecular oxygen to activate the quencher and reduce the energy transfer from BChl c aggregates to reaction centers. Upon the return to anoxic conditions, CsmI and CsmJ probably transfer electrons in the other direction: from reductants outside of chlorosome to the chlorosome quencher to inactivate the quencher and restore the energy transfer. Similar quenching and restoration mechanisms also exist in whole cells. The redox regulation of energy transfer effectively protects the photosynthetic apparatus under aerobic conditions by suppressing the activity of reaction centers, decreasing the level of reduction of ferredoxins, and minimizing the formation of reactive oxygen species (Frigaard and Matsuura, 1999). CsmX is present at a much lower amount than CsmI and CsmJ and appears to play a negligible role in the redox regulation of energy transfer (Chapter 3.5). In future studies, biochemical and biophysical methods will be further applied to analyze the structure and organization of chlorosome proteins. Immunoelectron microscopy and atomic force microscopy could be performed with anti-Csm antibodies and isolated chlorosomes to determine the exact distribution of proteins on the chlorosome envelope. Chlorosome proteins would be cross-linked on isolated chlorosomes and digested with proteases, and Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS) could be used to determine the cross-linking relationship among the peptide fragments, and this could lead to a more detailed model of chlorosome protein organization. Truncated forms of CsmH, in which the alanine-rich extensions have been deleted from the amino and carboxyl termini, can be overproduced in E. coli. If the truncated forms of CsmH are soluble and do not aggregate, then structural studies by either nuclear magnetic resonance (NMR) or X-ray crystallography should be performed to determine the structure of this protein. These studies will hopefully lead to an improved understanding of the structure and

185 organization of chlorosome proteins and the mechanism of chlorosome biogenesis and pigment incorporation.

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VITA

HUI LI

EDUCATION Ph.D., Biochemistry, Microbiology, and Molecular Biology, May 2006 The Pennsylvania State University B.S., Biochemistry and Molecular Biology, Jul 2000 Peking University, China

HONORS AND AWARDS Graduate Student Travel Award, The Pennsylvania State University, Aug 2004 Excellent Graduate Award of Pilot Program in Science, Peking University, China, May 2000

PUBLICATIONS Li, H. and Bryant, D. A. 2006. Roles of chlorosome proteins in pigment incorporation and chlorosome assembly: evidence revealed by mutants lacking multiple chlorosome proteins of Chlorobium tepidum. Manuscript in preparation. Li, H., Frigaard, N.-U., and Bryant, D. A. 2006. Molecular contacts for chlorosome envelope proteins revealed by cross-linking studies with chlorosomes from Chlorobium tepidum. Biochemistry, to be submitted. Li, H., Frigaard, N.-U., and Bryant, D. A. 2006. [2Fe-2S] proteins in chlorosomes. I. Construction and characterization of mutants lacking CsmI, CsmJ, and CsmX in the chlorosome envelope of Chlorobium tepidum. Biochemistry, to be submitted. Johnson, T. W., Li, H., Frigaard, N.-U., Bryant, D. A. and Golbeck, J. H. 2006. [2Fe-2S] proteins in chlorosomes. II. Redox titration of the [2Fe-2S] clusters in the proteins CsmI, CsmJ, and CsmX in the chlorosome envelope of Chlorobium tepidum. Biochemistry, to be submitted. Frigaard, N.-U., Li, H., Martinsson, P., Kumar Das, S., Frank, H. A., Aartsma, T. J., and Bryant, D. A. 2005. Isolation and characterization of carotenosomes from a bacteriochlorophyll c- less mutant of Chlorobium tepidum. Photosynthesis Research, 86: 101-111. Li, H., Frigaard, N.-U., and Bryant, D. A. 2004. Locations and interactions of chlorosome proteins on the chlorosome envelope in Chlorobium tepidum: insights from cross-linking experiments. Photosynthesis: Fundamental Aspects to Global Perspectives, the Proceedings of the 13th International Congress on Photosynthesis, Section 3, p. 116-119, ACG Publishing, Lawrence, Canada. Frigaard, N.-U., Li, H. and Bryant, D. A. 2004. Nine mutants of Chlorobium tepidum each unable to synthesize a different chlorosome protein still assemble functional chlorosomes. Journal of Bacteriology, 186: 646-653. Frigaard, N.-U., Gomez Maqueo Chew, A., Li, H., A., Maresca, J. A. and Bryant, D. A. 2003. Chlorobium tepidum: insights into the physiology and biochemistry of green sulfur bacteria from the complete genome sequence. Photosynthesis Research, invited review, 78: 93-117. Bryant, D. A., Vassilieva, E. V., Frigaard, N.-U., and Li, H. 2002. Selective protein extraction from Chlorobium tepidum chlorosomes using detergents. Evidence that CsmA forms multimers and binds bacteriochlorophyll a. Biochemistry 41: 14403-14411. Frigaard, N.-U., Vassilieva, E. V., Li, H., Milks, K. J., Zhao, J. and Bryant, D. A. 2001. The remarkable chlorosome. PS2001 Proceedings, Proceedings of the 12th International Congress on Photosynthesis, Brisbane, Australia. Article S1-003, p. 6, CSIRO Publishing, Canberra, Australia.