The Pennsylvania State University
The Graduate School
CHARACTERIZATION OF PIGMENT BIOSYNTHESIS AND LIGHT-HARVESTING
COMPLEXES OF SELECTED ANOXYGENIC PHOTOTROPHIC BACTERIA
A Dissertation in
Biochemistry, Microbiology, and Molecular Biology and Astrobiology by
Jennifer L. Thweatt
2019 Jennifer L. Thweatt
Submitted in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
December 2019
ii
The dissertation of Jennifer L. Thweatt was reviewed and approved* by the following:
Donald A. Bryant Ernest C. Pollard Professor in Biotechnology and Professor of Biochemistry and Molecular Biology Dissertation Advisor Chair of Committee
Squire J. Booker Howard Hughes Medical Investigator Professor of Chemistry and Professor of Biochemistry and Molecular Biology Eberly Distinguished Chair in Science
John H. Golbeck Professor of Biochemistry and Biophysics Professor of Chemistry
Jennifer L. Macalady Associate Professor of Geosciences
Timothy I. Miyashiro Assistant Professor of Biochemistry and Molecular Biology
Wendy Hanna-Rose Professor of Biochemistry and Molecular Biology Department Head, Biochemistry and Molecular Biology
*Signatures are on file in the Graduate School
iii ABSTRACT
This dissertation describes work on pigment biosynthesis and the light-harvesting
apparatus of two classes of anoxygenic phototrophic bacteria, namely the green bacteria
and a newly isolated purple sulfur bacterium. Green bacteria are introduced in Chapter 1
and include chlorophototrophic members of the phyla Chlorobi, Chloroflexi, and
Acidobacteria. The green bacteria are defined by their use of chlorosomes for light harvesting. Chlorosomes contain thousands of unique chlorin molecules, known as bacteriochlorophyll (BChl) c, d, e, or f, which are arranged in supramolecular aggregates.
Additionally, all green bacteria can synthesize BChl a, the and green members of the
phyla Chlorobi and Acidobacteria can synthesize chlorophyll (Chl) a. The biosynthetic pathways leading to the chlorophylls (Chls) and bacteriochlorophylls (BChls) of green bacteria have been nearly fully elucidated over the past two decades. The current state of
knowledge of the biosynthetic pathways leading from protoporphyrinogen IX to the
production of the (B)Chls found in green bacteria, and the distribution of these biosynthetic enzymes across the green bacteria in relation to their physiology, is reviewed
Chapter 2. The biosynthetic pathway leading to BChlide e or f was known prior to this
work, with the exception of one reaction, and the enzyme required for this reaction is
characterized in Chapter 3. A gene encoding a putative radical S-adenosyl-L-methionine
(SAM) protein, bciD, was known to be required for BChl e biosynthesis in vivo; however, it was unknown whether BciD was sufficient to convert BChlide c into BChlide e. To determine the function of BciD a His-tagged version of the enzyme was produced in Escherichia coli, and the enzyme was characterized. Characterization of BciD iv indicated that it contains a [4Fe-4S] cluster. Biochemical assays showed that BciD catalyzed the conversion of SAM into 5ʹ-deoxyadenosine and BChlide c or d into
BChlide e or f, respectively. Additionally, a 71-(OH) BChlide c or d intermediate was also observed. These data led to the conclusion that BciD is a radical SAM enzyme that converts the methyl group of BChlide c or d into the formyl group of BChlide e or f likely via a mechanism involving consecutive hydroxylation reactions of the methyl group. The demonstration that BciD is sufficient to catalyze the conversion of BChlide c into
BChlide e completed the biosynthetic pathways for (B)Chl biosynthesis in green bacteria.
Additionally, in the process of writing a review of (B)Chl biosynthesis in green bacteria,
it became necessary to perform bioinformatic analyses to identify (B)Chl biosynthetic
genes in newly available genomes and to validate prior annotations. These data are
presented in Chapter 4, where special attention is paid to the enzymes involved in the following reactions: the anoxic coproporphyrinogen III oxidative decarboxylation,
protoporphyrinogen (Protogen) IX oxidation, Proto IX magnesium chelation; C13
propionate methylation of Mg-Proto IX, oxidative ring cyclization of Mg-Proto IX 13- monomethyl ester to form the isocyclic E-ring of the macrocyle, C3 vinyl hydration, C8
and C12 methylation, and esterification of (B)Chlides with alcohol pyrophosphates.
Finally, Chapter 5 focuses on the photosynthetic pigments and light harvesting
complexes of a new Thiohalocapsa sp. that was isolated from Mushroom Spring in
Yellowstone National Park. This organism is referred to as Thiohalocapsa MS
throughout Chapter 5 and is the first thermophilic member of the genus Thiohalocapsa
and only the third thermophilic species purple sulfur bacteria. The main light-harvesting pigments are identified as bacteriochlorophyll a, spirilloxanthin, and anhydro- v rhodovibrin. Additionally, two spectral variants of the peripheral light-harvesting
complex, LH2, were identified and co-purified from this organism, they are denoted
B800-B830 and B800-B855. The core light harvesting complex, LH1-RC, was also purified and shown to have an absorbance maximum around 900 nm. The near-IR absorbance of this LH1-RC experiences a blue shift of 5-8 nm in response to the loss of
Ca2+ ions, similar to other thermophilic purple sulfur bacteria. These results support the
hypothesis that, like other thermophilic PSB, the Thiohalocapsa MS LH1-RC complex
binds Ca2+ ions that enhance its thermostability. Together the studies presented in this
dissertation provide new insights into the light-harvesting pigments and physiology of specific anoxygenic phototrophs and provide a jumping off point for further work on these subjects.
vi TABLE OF CONTENTS List of Figures xiii
List of Tables xv
List of Abbreviations xvi
Acknowledgements xix
Chapter 1 Introduction to Green Bacteria ...... 1
1.1 Abstract ...... 2
1.2 Introduction ...... 3
1.3 Light Harvesting in Green Bacteria ...... 8
1.4 Chlorophylls and Bacteriochlorophylls of Green Bacteria ...... 11
1.5 Organization of This Dissertation ...... 15
References ...... 17
Chapter 2 Biosynthesis of Chlorophylls and Bacteriochlorophylls in Green Bacteria ..... 30
2.1 Abstract ...... 31
2.2 Introduction ...... 32
2.3 Early Steps ...... 32
2.4 Protoporphyrin IX to Chlorophyllide a...... 36
2.4.1 Magnesium Chelation ...... 36
2.4.2 C13 Propionate Methylation ...... 38
2.4.3 Isocyclic E-Ring Formation ...... 39 vii 2.4.5 Reduction of C17=C18 Double Bond ...... 41
2.4.6 Reduction of C8 Vinyl Group ...... 42
2.5 Chlorophyllide a to Bacteriochlorophyllide a ...... 45
2.5.1 Reduction of the C7=C8 Double Bond...... 46
2.5.2 Hydration of the C3 vinyl group...... 48
2.5.3 3-Hydroxyethyl Dehydrogenase ...... 49
2.6 Chlorophyllide a to Bacteriochlorophyllide c, d, e, and f ...... 50
2.6.1 Demethoxycarbonylation of the C132 methylcarboxyl group ...... 50
2.6.2 Methylation at C8 and C12 ...... 52
2.6.3 Hydration of the C31 Position ...... 53
2.6.4 Methylation of the C20 methine bridge ...... 56
2.6.5 Formation of the C7 Formyl Group of BChlide e ...... 58
2.7 The final steps in (B)Chl biosynthesis ...... 59
2.7.1 Esterification of (B)Chlide ...... 59
2.7.2 Reduction of the alcohol moiety ...... 63
2.8 Concluding Remarks ...... 65
References ...... 66
Chapter 3 Characterization of BciD from Cba. limnaeum ...... 82
3.1 Abstract ...... 83 viii 3.2 Introduction ...... 84
3.3 Experimental Procedures ...... 89
3.3.1 Strains Used in This Study ...... 89
3.3.2 Cloning and Inactivation of bciD in Cba. limnaeum ...... 90
3.3.3 BciD Purification ...... 92
3.3.4 Protein Analyses and Verification ...... 93
3.3.5 Reconstitution of Fe/S Cluster ...... 93
3.3.6 Spectroscopic Measurements ...... 94
3.3.7 Pigment Separation and Preparation of Substrate Compounds ...... 95
3.3.8 Mass Spectrometry ...... 96
3.3.9 Enzyme Activity Assay ...... 97
3.3.10 Phylogenetic Analyses ...... 98
3.4 Results ...... 99
3.4.1 Inactivation of bciD in Cba. limnaeum ...... 99
3.4.2 Purification and Characterization of BciD-His6 ...... 100
3.4.3 BciD-His6 Activity Assay with BChlide c ...... 102
3.4.4 BciD-His6 Activity Assay with BChlide d ...... 104
3.5 Discussion ...... 105
References ...... 112 ix Chapter 4 Bioinformatic Insights into (Bacterio)chlorophyll Biosynthesis in Green
Bacteria ...... 137
4.1 Abstract ...... 138
4.2 Introduction ...... 139
4.3 Experimental Procedures ...... 145
4.3.1 Genomes and Metagenomes Used in This Work ...... 145
4.3.2 Phylogenetic Tree Building ...... 146
4.3.3 Sequence Similarity Networks ...... 146
4.4 Results ...... 146
4.4.1 Identification of Genes Encoding (B)Chl Biosynthetic Enzymes in
Green Bacteria ...... 147
4.4.2 Decarboxylation of Coproporphyrinogen III ...... 148
4.4.3 Oxidation of Protoporphyrinogen IX ...... 149
4.4.4 Magnesium Chelation ...... 150
4.4.5 C13 Propionate Methylation ...... 151
4.4.6 Isocyclic E-Ring Formation ...... 153
4.4.7 Hydration of the C3 vinyl group...... 154
4.4.8 Methylation at C8 and C12 ...... 155
4.4.9 Esterification of (B)Chlide ...... 158
4.5 Discussion ...... 158 x 4.5.1 Early Steps ...... 159
4.5.2 Proto IX to Chlide a...... 162
4.5.3 Chlide a to BChlide a and BChlide c, d, e, and f ...... 165
4.5.4 Final Steps ...... 168
4.5.6 Concluding Remarks ...... 169
References ...... 171
Chapter 5 Light Harvesting Apparatus and Pigments of Thiohalocapsa MS ...... 200
5.1 Abstract ...... 201
5.2 Introduction ...... 202
5.2.1 Purple Sulfur Bacteria ...... 202
5.2.2 Newly Isolated Thiohalocapsa sp. from Mushroom Spring,
Yellowstone National Park...... 205
5.3 Experimental Procedures ...... 206
5.3.1 Strains used in this study ...... 206
5.3.2 Phylogeny of Purple Sulfur Bacteria ...... 207
5.3.3 Microscopy ...... 207
5.3.4 Spectroscopic Analysis ...... 208
5.3.5 Pigment Separation and Analysis ...... 208
5.3.6 Isolation and Purification of Light Harvesting Complexes ...... 209 xi 5.4 Results ...... 210
5.4.1 Identification, Growth, and Absorbance Spectra ...... 210
5.4.2 Pigment Analysis ...... 211
5.4.3 Purification and Characterization of LH Complexes ...... 212
5.4.4 Effect of Ca2+ on the LH1-RC Complex ...... 213
5.5 Discussion ...... 214
References ...... 219
Chapter 6 Concluding Remarks ...... 232
6.1 Overview and Discussion ...... 232
References ...... 236
Appendix A. Co-culture Growth Studies of Prosthecochloris sp. HL-130-GSB and
Geobacter sulfurreducens PCA ...... 239
A.1 Abstract ...... 239
A.2 Introduction ...... 240
A.3 Experimental Procedures ...... 241
A.3.1 Strains Used in This Work ...... 241
A.3.2 Growth Media ...... 241
A.3.3 Growth Studies ...... 242
A.4 Results ...... 242 xii A.5 Discussion ...... 243
References ...... 247
Appendix B.Genome Sequence of Thiohalocapsa MS ...... 254
B.1 Introduction ...... 254
B.2 Experimental Procedures...... 254
B.2.1 Strains Used in This Study ...... 254
B.2.2 Genomic DNA (gDNA) Preparation ...... 255
B.2.3 gDNA Sequenceing ...... 255
B.2.4 Bioinformatic Analyses ...... 255
B.3 Results and Discussion ...... 256
B.3.1 Overview of Genome ...... 256
B.3.2 Central Metabolism ...... 256
B.3.3 Photosynthesis Related Genes ...... 259
B.4 References ...... 261
xiii LIST OF FIGURES Figure 1.1 Phylogeny of Green Bacteria…………………………………………………………26
Figure 1.2 Localization of Chlorophylls and Bacteriochlorophyls in Green Bacteria………….28
Figure 1.3 Structure of Chlorophylls and Bacteriochlorophylls from Green Bacteria…………..29
Figure 2.1 Distribution of Biosynthetic Enzymes for (B)Chl Biosynthesis in Green Bacteria…78
Figure 2.2 Biosynthesis: 5-ALA to Chlide a…………………………………………………….79
Figure 2.3 Biosynthesis: Chlide a to BChlide a and BChlide c, d, e, or f……………………….80
Figure 2.4 Biosynthesis: Final Steps…………… ………………………………………………81
Figure 3.1 Scheme showing the biosynthetic pathway leading from Chlide a to BChl c……...119
Figure 3.2 Organization of the gene cluster encoding bciD and cruB in brown-colored GSB...120
Figure 3.3 Inactivation of bciD in Cba. limnaeum……………………………………………..121
Figure 3.4 Absorbance spectra of whole cells and pigment extracts…………………………..122
Figure 3.5 SDS-PAGE analysis of fractions from the purification of recombinant BciD-His6..123
Figure 3.6 UV-visible absorbance spectra of as-isolated and reconstituted recombinant BciD-
His6……………………………………………………………………………………..124
Figure 3.7 EPR spectra of as-isolated and reconstituted recombinant BciD-His6……………...125
Figure 3.8 Reversed-phase HPLC analysis of BciD reaction with BChlide c………………….126
Figure 3.9 Reversed-phase HPLC analysis of 5ʹ-deoxyadenosine produced by BciD during
reaction with BChlide c………………………………………………………………...127
Figure 3.10 Reversed-phase HPLC analysis of BciD reactions with BChlide d……………….128
Figure 3.11 Sequence alignment of BciD proteins……………………………………………..129
Figure 3.12 Proposed reaction scheme for BciD and BchK to convert BChlide c into BChl e..130
Figure 3.13 Reversed Phase HPLC Elution Profile of 40°C Experiment………………………131 xiv Figure 3.14 Phylogenetic tree showing the relationship among chlorophyll synthases………..132
Figure 4.1 Phylogeny of HemN Homologs in Green Bacteria…………………………………177
Figure 4.2 Multiple Sequence Alignment of Green Bacterial HemN Paralogs………………...178
Figure 4.3 Sequence Similarity Network of HemY Related Sequences in Green Bacteria……179
Figure 4.4 Phylogeny of HemY Related Protein in Green Bacteria……………………………180
Figure 4.5 Phylogeny of BchH Paralogs in Green Bacteria……………………………………181
Figure 4.6 Phylogeny of BchF Paralogs in Green Bacteria…………………………………….183
Figure 4.7 Sequence Similarity Network of BchQ, BchR and BchE Related Sequences in Green
Bacteria…………………………………………………………………………………184
Figure 4.8 Phylogeny of BchQ, BchR, and BchE in Green Bacteria…………………………..185
Figure 4.9 Phylogeny of ChlG, BchG and BchK in Green Bacteria…………………………...187
Figure 5.1 Structures of Light-Harvesting and Reaction Center Complexes of a PSB……...... 223
Figure 5.2 16S rRNA Phylogenetic Tree of the Order Chromatiales…………………………..224
Figure 5.3 Appearance of Thiohalocapsa MS………………………………………………….225
Figure 5.4 UV-Vis Absorbance Spectra………………………………………………………..226
Figure 5.5 Elution profile from reversed-phase HPLC monitored at 770 nm………………....227
Figure 5.6 Elution profiles from reversed-phase HPLC monitored at 491 nm………………...228
Figure 5.7 Chromatophores and LH complexes of Thiohalocapsa MS………………………..229
Figure 5.8 LH1-RC response to CaCl2 and chelators………………….……………………….230
Figure A.1 Co-culture Growth Experiment…………………………………………………….248
Figure A.2 Growth Experiment with Varied Additives………………………………………...249 xv LIST OF TABLES Table 3.1 Oligonucleotide primers used in this study…………………………………………..134
Table 3.2 Summary of properties and assignments of peaks in BciD reaction with BChlide c..135
Table 3.3 Summary of properties and assignments of peaks in BciD reaction with BChlide d..136
Table 4.1 Green Bacterial Genomes Analyzed in this Work…………………………………...189
Table 4.2 (B)Chl Genes in Green Bacteria-Early Steps……………………………………..…190
Table 4.3 (B)Chl Genes in Green Bacteria Leading from ProtoIX to Chlide a………………...192
Table 4.4 (B)Chl Genes in Green Bacteria Leading from Chlide a to BChlide c, d, e, or f……194
Table 4.5 (B)Chl Genes in Green Bacteria Final Steps………………………………………...196
Table 4.6 Metagenome-assembled genomes of ‘Candidatus Thermochlorobacter aerophilum’197
Table 4.7 Summary of O-methyltransferase Hits in the CDD from Ca. T. aerophilum………..198
Table 4.8 List C13 Propionate Methyltransferase Candidates in Ca. T. aerophilum OS………199
Table 5.1 Effect of Ca2+ on LH1-RC from Thermophillic PSB………………………………..231
Table A.1 Comparison of Basic Media Components of HL-Cl and mGB……………………..250
Table A.2 Comparison of Vitamin and Trace Element Components HL-CL and mGB……….252
Table B.1 Carbon Metabolism Genes…………………………………………………………..263
Table B.2 Calvin Benson Basham Cycle Genes………………………………………………..265
Table B.3 Nitrogen Fixation Genes…………………………………………………………….266
Table B.4 Sulfur Metabolism Genes……………………………………………………………268
Table B.5 Pigment Biosynthesis Genes………………………………………………………...270
TableB.6 Light Harvesting Apparatus Genes…………………………………………………..272 xvi LIST OF ABBREVIATIONS
3-HPP 3-hydroxypropionate 3V C3 vinyl 5-ALA 5-aminolevulinic acid 8V C8 vinyl 8VR 8 vinyl reductase A. Arabidopsis Ac acetyl Alc. Allochromatium ATP adenosine triphosphate BChl bacteriochlorophyll BChlide bacteriochlorophyllide BPhe bacteriopheophytin BPheide bacteriopheophorbide C. Chloroploca Ca. Candidatus Cab. Chloroacidobacterium CAO chlorophyllide a oxygenase Cba. Chlorobaculum Cfl. Chloroflexus Chl. Chlorobium Chl chlorophyll Chlide chlorophyllide Chp. Chloroherpeton Coprogen III coproporphyrinogen III COR chlorophyllide oxidoreductase xvii cryo-EM cryogenic electron microscopy DPOR dark operative protochlorophyllide reductase E. Escherichia Et ethyl EPR electron paramagnetic resonance F farnesyl FAD flavin adenine dinucleotide FAP filamentous anoxygenic phototrophs FMO Fenna-Matthews-Olsen protein G. Gemmatimonas GG geranylgeranyl GSB green sulfur bacteria HYSCORE hyperfine sublevel correlation IR infrared LH light harvesting complex MgCH magnesium chelatase Mch. Marichromatium Mg-PME Mg-Proto IX 13-monomethyl ester NAD nicotinamide adenine dinucleotide NADP nicotinamide adenine dinucleotide phosphate Osc. Oscillochloris P. Plasmodium P phytyl PChlide protochlorophyllide PD Δ2,6 phytadienyl Phe pheophytin xviii Pheide pheophorbide PP diphosphate Proto IX protoporphyrin IX Protogen IX protoporphyrinogen IX PSB purple sulfur bacteria Ptc. Prostecochloris Rba. Rhodobacter RC reaction center rRNA ribosomal ribonucleic acid Rsp. Rhodospirillum SAM S-adenosyl-L-methionine sp. species spp. species (plural) SSN sequence similarity network T. Thermochlorobacter TCA tricarboxylic acid Tch. Thermochromatium Thc. Thiohalocapsa UV ultra-violet V. Viridilinea Vis visible WT wild type xix ACKNOWLEDGEMENTS
I am incredibly indebted to a great many people who have helped me to succeed
throughout my graduate career. First, I am indebted to my advisor who provided the
funding and environment in which I was able to conduct this research, in addition to
invaluable ideas and feedback on my work. I am also grateful for my committee members
Drs. Squire Booker, John Golbeck, Tim Miyashiro, and Jennifer Macalady who supported me and put in extra effort during my tenure in graduate school. Next I would like to thank the administration of the BMMB graduate program that has always taken care of any needs I’ve had during my time here. I am also grateful to those with whom
I’ve collaborated on the work in this dissertation namely Drs. Daniel Canniffe, Marcus
Tank and Bryan Ferlez. I am also so grateful to have worked with a group of incredibly talented and motivated people over the years in the Bryant Lab and the Golbeck Lab and to have received invaluable advice from members of the Booker Lab on working with radical-SAM enzymes. Finally, I am incredibly grateful to the support I have received from so many friends and family over the years, some of whom are not here to see the completion of this dissertation but whose faith in me to succeed has kept me going over many long days and late nights in the lab. The work in this dissertation was supported by funding from the Photosynthetic Systems Program, Division of Chemical Sciences,
Geosciences, and Biosciences (CSGB), Office of Basic Energy Sciences of the United
States Department of Energy Grant DE-FG02-94ER20137 to Donald A. Bryant. 1
Chapter 1 Introduction to Green Bacteria
Publication: Jennifer L. Thweatt, Daniel P. Canniffe, and Donald A. Bryant (2019)
Biosynthesis of chlorophylls and bacteriochlorophylls in green bacteria. Advances in
Botanical Research. 90, 35-89.
Contributions: JLT wrote the text and made all figures. DAB and DPC provided
feedback, editing, and additions to the introduction from the original draft.
2 1.1 Abstract
Green bacteria include chlorophototrophic members of the phyla Chlorobi,
Chloroflexi, and Acidobacteria and are defined by their use of chlorosomes for light
harvesting. Despite their shared use of chlorosomes as light-harvesting antenna and their
chlorophototrophic metabolic mode, these organisms exhibit a surprising diversity in
their ecological and physiological characteristics. Chlorosomes contain thousands of
unique chlorin molecules, known as bacteriochlorophyll (BChl) c, d, e, or f, which are
arranged in supramolecular aggregates. Additionally, all green bacteria can synthesize
BChl a, the green members of the phyla Chlorobi and Acidobacteria can synthesize
chlorophyll (Chl) a, and Chloracidobacterium thermophilum uniquely synthesizes Zn-
BChl a′. In this chapter the general physiology, light-harvesting apparatus, and
(bacterio)chlorophylls of green bacteria will be introduced.
3 1.2 Introduction
Green bacteria are an eclectic group of chlorophototrophic (i.e., chlorophyll (Chl)
and/or bacteriochlorophyll (BChl)-dependent) bacteria, which are defined by their ability
to make large light-harvesting antenna complexes known as chlorosomes (Bryant and
Canniffe, 2018; Bryant et al., 2012; Bryant and Frigaard, 2006; Frigaard and Bryant,
2004, 2006; Oostergetel et al., 2010; Orf and Blankenship, 2013; Saer and Blankenship,
2017; Thiel et al., 2018). Chlorosomes contain self-assembled supramolecular aggregates
of BChl c, d, e, or f, which are chlorin molecules uniquely found in chlorosomes (Bryant and Canniffe, 2018; Gomez Maqueo Chew and Bryant, 2007). Green bacteria currently
include organisms from three different bacterial phyla: Chlorobi, Chloroflexi, and
Acidobacteria (Bryant et al., 2007, 2012; Thiel et al., 2018). (see Figure 1.1A). In addition to the green bacteria, chlorophototrophic bacteria are also found in the phyla
Proteobacteria (commonly called purple bacteria), Gemmatimonadetes, Cyanobacteria, and Firmicutes (Thiel et al., 2018). Recent studies also show sequence-based evidence for chlorophotrophy in the phyla Bacteroidetes and “Candidatus Eremiobacterota,” which if confirmed biochemically would make a total of nine bacterial phyla which contain chlorophototrophs (Tahon and Willems, 2017; Thiel et al., 2018; Ward et al.,
2019). When compared to the other major bacterial phyla, green bacteria are phylogenetically distant from each other and in some cases are more closely related to other chlorophototrophic bacteria than to each other (Figure 1.1A).
The term “green bacteria” was initially used in a non-specific fashion to differentiate green-colored chlorophototrophic bacteria from purple chlorophototrophic 4 bacteria. However, this common name eventually became synonymous with green sulfur
bacteria (GSB). GSB contain chlorosomes and are chlorophototrophic members of the
phylum Chlorobi, families Chlorobiaceae and Chloroherpetonaceae, whose first isolates
were green-colored (Imhoff, 2017; Imhoff and Thiel, 2010; Liu et al., 2012b; Nadson,
1906). When Chloroflexus (Cfl.) aurantiacus was initially isolated and shown to contain
chlorosomes, it and other chlorosome-containing members of the phylum Chloroflexi,
such as Oscillochloris (Osc.) trichoides, were included within the green bacteria (Bryant
et al., 2012; Gorlenko and Korotkov, 1979; Keppen et al., 1994; Pierson and Castenholz,
1974b, 1974a). Chlorosome-containing members of the Chloroflexi are now referred to as
green filamentous anoxygenic phototrophs, or green FAPs, which were formerly and
commonly known as green non-sulfur bacteria or green gliding bacteria (Bryant et al.,
2012; Bryant and Frigaard, 2006; Garrity et al., 2001). One of the most recent additions
to the green bacteria is Chloracidobacterium (Cab.) thermophilum (Bryant et al., 2007).
Cab. thermophilum produces chlorosomes and is currently the only known chlorophototrophic species of the phylum Acidobacteria (Garcia Costas et al., 2011,
2012a, 2012b, Tank et al., 2017, 2019, Tank and Bryant, 2015b, 2015a; Thiel et al.,
2018). Despite the common trait that they use chlorosomes for light harvesting, members of these three phyla are otherwise markedly different in their physiological properties and photosynthetic apparatus. I will briefly describe each group of green bacteria and their
photosynthetic apparatus before discussing the biosynthesis of their unique complements
of (B)Chls. 5 The phylum Chlorobi comprises two classes, the non-chlorophototrophic
Ignavibacteria and the chlorophototrophic Chlorobia (Iino et al., 2010). Ignavibacterium
album and Melioribacter roseus are the best described members of the Ignavibacteria
(Iino et al., 2010; Kadnikov et al., 2013; Liu et al., 2012a; Podosokorskaya et al., 2013).
The class Chlorobia contains all chlorophototrophic members of the phylum Chlorobi:
GSB as well as “Candidatus (Ca.) Thermochlorobacter aerophilum” (Liu et al., 2012b)
(Figure 1.1B). GSB are obligately anaerobic, anoxygenic Chl-dependent chlorophotolithoautotrophic organisms that include all members of the family
Chlorobiaceae as well as Chp. thalassium (and possibly other members of the family
Chloroherpetonaceae) (Liu et al., 2012a, 2012b). These organisms fix carbon dioxide via the reverse TCA cycle, fix dinitrogen with molybdenum-containing nitrogenase, and use reduced sulfur sources, molecular hydrogen, or ferrous iron as their electron sources
(Frigaard and Dahl, 2009; Heising et al., 1999; Wahlund and Madigan, 1993; Wahlund
and Tabita, 1997). Despite their name, GSB may be either green-colored or brown-
colored, depending on the type of BChl in their chlorosomes (see below). “Ca. T.
aerophilum” is unique within the Chlorobia. It is an aerobic, chlorophotoheterotophic
organism that contains chlorosomes, lacks the ability to fix CO2 and N2, and does not
oxidize sulfur compounds as electron sources (Liu et al., 2012a; Tank et al., 2017).
Because of these significant physiological and metabolic differences, a separate family,
Thermochlorobacteriaceae, has been proposed for this organism and its close relatives
(Liu et al., 2012b). All members of the Chlorobia have homodimeric, type-1 reaction centers (RCs), and employ the BChl a-binding Fenna-Matthews-Olson (FMO) protein as the conduit for energy transfer from the baseplate BChl a-binding protein, CsmA, to the 6 core antenna of the RC (Bryant et al., 2012; Bryant and Canniffe, 2018; Saer and
Blankenship, 2017).
The phylum Chloroflexi includes many non-chlorophototrophic members in
addition to FAPs (Thiel et al., 2018) (Figure 1.1B). Unlike the Chlorobia, FAPs lack
FMO and have type-2 RCs (Cardona, 2015; Thiel et al., 2018). Red FAPs include
members of the chlorosome-less genera Roseiflexus and Heliothrix, and the recently
discovered “Ca. Roseilinea gracile” (Tank et al., 2017; Thiel et al., 2018). The green
FAPs contain chlorosomes and include members of the genera Chloroflexus,
Oscillochloris, Chloronema, and “Ca. Chloroploca asiatica”, “Ca. Viridilinea
mediisalina” and “Ca. Chloranaerofilum” (Bryant et al., 2012; Grouzdev et al., 2018;
Tank et al., 2017; Thiel et al., 2018). Under oxic conditions Chloroflexus spp. grow chemoheterotrophically, but under anoxic conditions most are Chl-dependent photoheterotrophs and a few are photoautotrophs. Chloroflexus spp. do not fix N2; can
use organic carbon sources, H2, or H2S as electron donors; and can fix CO2 using the 3- hydroxypropionate pathway (3-HPP) (Fuchs, 2011; Klatt et al., 2007, 2013).
Oscillochloris spp. and the Candidatus strains listed above are part of a deeply branching
subclade of the suborder Chloroflexineae, but ‘Ca. Chloroploca’ and ‘Ca. Viridilinea’
likely differ from Oscillochloris at the family level (Grouzdev et al., 2018). Osc. trichoides is the best characterized member of this suborder. Oscillochloris spp. are unique among FAPs because they are strict anaerobes that can fix N2; moreover, they
grow chlorophotoautotrophically by fixing CO2 via the Calvin-Benson-Basham cycle with H2S (or H2) as the electron donor or chlorophotoheterotrophically using acetate or 7 pyruvate (Ivanovsky et al., 1999; Keppen et al., 1994). ‘Ca. C. asiatica’ and ‘Ca. V.
mediisalina’ have been studied via enrichment cultures and recent draft genomes. Like
Osc. trichoides both of these strains have been shown to grow as anaerobic
chlorophototrophs, however; like Chloroflexus spp. they have genes for the 3-HPP CO2
fixation pathway (Grouzdev et al., 2018). Green FAPs can actually be brown, green, orange or reddish in color depending on growth conditions, in particular the oxygen
concentration that controls their pigment composition (Thiel et al., 2018).
The phylum Acidobacteria comprises many physiologically diverse, non- phototrophic organisms but only a single chlorophototrophic species, Cab. thermophilum, has been described to date (Bryant et al., 2007; Tank et al., 2017, 2018; Tank and Bryant,
2015b, 2015a; Thiel et al., 2018) (Figure 1.1B). Cab. thermophilum is an anoxygenic, chlorophotoheterotrophic organism that grows optimally under microoxic conditions. It does not fix N2 and lacks the genes encoding key enzymes of all known pathways for
autotrophic CO2 fixation (Garcia Costas et al., 2012a; Thiel et al., 2018). Furthermore,
Cab. thermophilum cannot synthesize lysine, branched-chain amino acids, or vitamin B12
(cobalamin); however, it can degrade branched-chain amino acids, and it takes up and
assimilates all amino acids except aspartic acid and glutamic acid (Tank and Bryant,
2015b, 2015a; Tank et al. 2018). Like the Chlorobia, Cab. thermophilum has type-1
homodimeric RCs (Tsukatani et al., 2012). Homodimeric type-1 RCs are otherwise only found in strictly anaerobic members of the phyla Firmicutes (heliobacteria) and Chlorobi
(with the exception of “Ca. T. aerophilum” as noted above). An additional similarity to 8 the Chlorobia is the presence of the BChl a-binding FMO antenna protein (Tsukatani et
al., 2010; Wen et al., 2011).
1.3 Light Harvesting in Green Bacteria
As noted above, the defining property of green bacteria is the presence of
chlorosomes, light-harvesting organelles that are tightly appressed to the cytoplasmic
surface of the cell membrane (Cohen-Bazire et al., 1964; Staehelin et al., 1978, 1980).
Chlorosomes are supramolecular structures assembled from self-aggregated BChl c, d, e,
or f molecules within a protein-stabilized, phospholipid monolayer envelope (Frigaard
and Bryant, 2006; Tsukatani et al., 2016) (Figure 1.2). Chlorosomes can contain up to
250,000 BChl molecules and a single cell can contain up to ~250 chlorosomes (Frigaard
and Bryant, 2006; Montaño et al., 2003). Chlorosomes additionally contain large amounts
of carotenoids and quinones, and the envelope can contain up to ~10 different proteins
(Frigaard and Bryant, 2006); however, mutational studies in Chlorobaculum (Cba.)
tepidum have shown that only one chlorosome protein, CsmA, is essential (Bryant et al.,
2002, 2012; Frigaard et al., 2004; Garcia Costas et al., 2011; Li et al., 2013; Li and
Bryant, 2009). The extremely high pigment:protein ratio of chlorosomes makes them a
remarkably efficient light-harvesting antenna from a cost-benefit perspective, especially when compared to predominantly proteinaceous antenna complexes (e.g., the
phycobilisome; (Bryant and Canniffe, 2018; Frigaard and Bryant, 2006)). This efficiency, as well as the enormous number of BChls (up to 30% of the cell carbon), allows green bacteria to live in extremely low light intensities that exclude other chlorophototrophs, 9 which have antenna complexes with many fewer pigments per RC (Bryant and Canniffe,
2018; Frigaard and Bryant, 2006). After light absorption by BChl aggregates within the chlorosome, excitation energy is rapidly transferred to the BChl a molecules associated
with the baseplate (Bryant and Canniffe, 2018; Pšenčík et al., 2009, 2014; Saer and
Blankenship, 2017). The baseplate forms the interface between the chlorosome and
cytoplasmic membrane and is made up of an array of CsmA dimers (Bryant and
Canniffe, 2018; Li et al., 2006; Nielsen et al., 2016). Each CsmA subunit binds one BChl a molecule, and baseplate arrays of CsmA occur in the chlorosomes of all green bacteria
(Bryant et al., 2012; Garcia Costas et al., 2011; Li et al., 2006; Nielsen et al., 2016;
Pšenčík et al., 2009, 2014).
In Chlorobia and Cab. thermophilum, which have type-1 RCs, the baseplate
transfers excitation energy to the FMO protein, a BChl a-binding, peripheral antenna
complex that also binds to RCs (Huang et al., 2012; Tsukatani et al., 2010; Wen et al.,
2009, 2011). FMO functions as a homo-trimer, and each subunit binds 7 BChl a
molecules and 1 intersubunit BChl a molecule; thus, each FMO trimer binds 24 BChl a
molecules, and several trimers are probably associated with each RC (Busch et al., 2011;
Fenna and Matthews, 1975; Larson et al., 2011; Tronrud et al., 2009). The FMO protein
transfers excitation energy to the BChl a molecules associated with the core antenna of
the type-1 RC (Busch et al., 2011; Milder et al., 2010). The GSB RCs mostly contain
BChl a but a few (~4) Chl a molecules are also bound; the special pair, known as P840,
consists of two molecules of BChl a′, while Chl a is the primary electron acceptor. An
additional 14 BChl a and a total of 4 Chl a molecules, two forming A0 and two accessory 10 pigments, have been reported in the RC (Ohashi et al., 2010; Permentier et al., 2000).
This relatively low pigment content of the RC has recently been called into question. The
crystal structure of the heliobacterial RC had a much larger than expected number of
pigments (~60 BChl g and 81-OH-Chl a molecules), and this suggests that Chl numbers
in other homodimeric type-1 RCs should be reinvestigated (Gisriel et al., 2017). In Cab.
thermophilum the type-1 RCs contain BChl a, Zn-BChl a′, and Chl a with an estimated
ratio of 12.8:2:8, respectively (Tsukatani et al., 2012). The primary electron acceptor in
this RC was recently identified as Chl a, and thus, all type-1 RCs utilize a chlorin, either
Chl a or 81-OH-Chl a, as the electron acceptor (Zill et al., 2018). Very recent results from
HYSCORE analysis of P840+ indicate that Zn-BChl a′ is the special pair pigment in this
RC (Charles et al., 2019).
Unlike GSB and Cab. thermophilum, green FAPs lack the FMO protein and have
type-2 RCs. The chlorosome baseplate complex instead transfers excitation energy to a
B808-866 complex, which is similar to the LH1 complex of purple bacteria (see Chapter
5) (Novoderezhkin et al., 1998; Saer and Blankenship, 2017; Xin et al., 2005). The B808-
866 complex is a ring-shaped, integral membrane protein complex that binds ~45 BChl a
molecules (Collins et al., 2012; Majumder et al., 2016; Xin et al., 2018), and as in purple
bacteria, excitation energy is transferred from B808-866 to the type-2 RC. The heterodimeric RC comprises the PufL and PufM subunits that bind three BChl a and three bacteriopheophytin (BPhe) a molecules (Pierson and Thornber, 1983; Xin et al.,
2018). The special pair, known as P870, is made up of two BChl a molecules that act as the primary electron donor, and BPhe a is the primary acceptor (Blankenship, 1994; 11 Bruce et al., 1982; Woodbury et al., 1985). A “voyeur” BChl a molecule is present on the
active branch of the electron transport chain as in bacterial reaction centers, but the
inactive branch has two BPhe a molecules (Xin et al., 2018).
1.4 Chlorophylls and Bacteriochlorophylls of Green Bacteria
Like heliobacteria and cyanobacteria that also have type-1 RCs, Chlorobia and
Cab. thermophilum synthesize derivatives of Chl a. The C–C bond at the C17–C18 position of the D ring is a double bond in other porphyrins, but in chlorins, including Chl a, this is a single bond (Figure 1.3). In the RCs of both GSB and Cab. thermophilum, the
C173 carboxylate moiety of Chl a is esterified with Δ2,6-phytadienol, denoted here as Chl
aPD to differentiate it from Chl a esterified with phytol (Chl aP), which is the form of Chl
a commonly found in cyanobacteria, algae and plants. Because Chl aPD is only found in
small amounts in the RC, it is a relatively minor pigment in these organisms. For
example, it comprises only ~0.3% of the cellular (B)Chl content in the model GSB Cba.
tepdium but is somewhat more abundant in Cab. thermophilum (Frigaard and Bryant,
2004; Tsukatani et al., 2012).
BChl a derivatives occur in all green bacteria, most purple chlorophototrophic
bacteria and the recently discovered aerobic anoxygenic chlorophotoheterotroph,
Gemmatimonas phototrophica (Bryant et al., 2012; Garcia Costas et al., 2012b; Imhoff,
2017; Thiel et al., 2018; Zeng et al., 2014; Zeng and Koblížek, 2017). BChl a is a bacteriochlorin, meaning that, in addition to the reduced D ring present in chlorins, the 12 C7=C8 double bond of the B ring is also reduced to a single bond (Figure 1.3). The only
other difference between BChl a and Chl a occurs at the C3 position, where BChl a has an acetyl group but Chl a has a vinyl group. In green bacteria BChl a is esterified with
3 phytol at the C17 position and is denoted here as BChl aP. Additionally, BChl aP′ and
Zn-BChl aP′ are found in the RCs of GSB and Cab. thermophilum, respectively (Ohashi et al., 2010; Permentier et al., 2000; Tsukatani et al., 2012). BChl a′ differs from BChl a
in the stereochemistry at the C132 position, while Zn-BChl a derivatives differ by the
substitution of a Zn++ ion for the canonical Mg++ ion in the macrocycle (Figure 1.3).
BPhe a, which differs from BChl a by the absence of any metal ion in the macrocycle, is
found in type-2 RCs of purple bacteria, FAPs, and G. phototrophica.
Despite their names, BChls c, d, e and f are not bacteriochlorins. These molecules
are chlorins that are also commonly referred to as “Chlorobium Chls” and are only found
in green bacteria (Smith, 1994). They have unique structural features that allow them to
form the supramolecular structures found within chlorosomes (Balban et al., 2005).
Unlike all other (B)Chls, they lack the C132 methylcarboxyl group. Removal of this group decreases steric hindrance and enhances the ability of the molecules to self-
assemble. All “Chlorobium Chls” also have a hydroxyethyl group at the C3 position,
while most other (B)Chls have either a vinyl group or an acetyl group (Figure 1.3).
Within chlorosomes or in BChl aggregates in vitro, the hydroxyl group at C31 forms a
coordination bond with the central Mg of a neighboring BChl molecule that is crucial for
self-aggregation (Balaban et al., 2005; Ganapathy et al., 2009; Steensgaard et al., 2000).
The C31 carbon atom is a chiral center, and the hydroxyethyl group occurs with both R 13 and S stereochemistry at this position, which results in structural heterogeneity among the
BChl molecules in chlorosomes. Additional heterogeneity exists at the C82, C121, and
C173 positions of BChl c, d, e, and f. As a result of the structural heterogeneity of these
molecules, it is important to note that the terms BChl c, d, e, or f, describe a family of related homolog molecules and thus do not refer to molecules with a defined structure
(Smith, 1994; Gomez Maqueo Chew and Bryant, 2007). In Chlorobia, Cab. thermophilum, Osc. trichoides, (and probably ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’) but not Chloroflexus spp., the C82 and C121 positions are differentially methylated
(Bryant et al., 2012; Gaisin et al., 2019; Garcia Costas et al., 2012b; Grouzdev et al.,
2015, 2018; Gorlenko et al., 2014). The exact nature and extent of these modifications are
dependent on the organism and the growth conditions. The C8 side chains of these BChls
may be ethyl, n-propyl, isobutyl, or neopentyl groups, while the C12 side chain may be a
methyl or an ethyl moiety. The degree of methylation at C82 and C121 affects the
absorbance properties of the BChl aggregates in chlorosomes, with higher degrees of
methylation causing spectral broadening and correlating with greater amounts of BChl in
cells (Gomez Maqueo Chew et al., 2007). The C173 carboxyl group of BChls c, d, e, or f
can be esterified with a variety of alcohols in green bacteria. The esterifying alcohol for
these BChls in GSB is predominantly farnesol, but smaller amounts of other alcohols,
including geranylgeraniol, phytol and various straight-chain alcohols, often occur (Airs et
al., 2001; Glaeser et al., 2002; Gomez Maqueo Chew et al., 2008). In Cab. thermophilum
the C173 carboxyl group is mostly esterified with a mixture of straight-chain alcohols
between 15 and 18 carbons in length, predominantly octadecanol; smaller amounts of
farnesol and geranylgeraniol also occur (Garcia Costas et al., 2012b)). In green FAPs, the 14 C173 carboxyl moiety of BChl c is also mostly esterified with stearol and small amounts of geranylgeraniol. The chain length of the esterifying alcohols has been shown to affect the spacing of aggregated BChls in chlorosomes and may be modified in response to
changes in growth conditions (Pšenčík et al., 2010). The remarkable heterogeneity of
BChl homologs in green bacteria may be a mechanism to tune the absorption properties
of the BChls in chlorosomes, where no proteins are available to perform this function.
BChl c, d, e, and f are structurally similar, but the subtle differences among them
produce major differences in the absorption spectra and coloration of the organisms that
produce them (see Chapter 3). Green bacteria naturally synthesize either BChl c, d, or, e,
and mutational studies have produced organisms that synthesize BChl f, which has not
yet been shown to occur naturally (Harada et al., 2012; Orf et al., 2013; Vogl et al.,
2012). Organisms that synthesize BChl c or d are green-colored, while organisms
synthesizing BChl e or f are brown-colored (Thiel et al., 2018). BChl c is the most
common Chlorobium Chl and is found in most GSB, most if not all green FAPs, and in
Cab. thermophilum. BChl d is similar in structure to BChl c but lacks the methyl group
on the methine bridge at C20 (Figure 1.3). BChl d is found in GSB that occur closer to
the surface of stratified environments (Maresca et al., 2004; van Gemerden and Mas,
1995), and is found in some strains of Cba. parvum, “Ca. T. aerophilum,” and some
Chloronema spp. and has recently been reported in ‘Ca. V mediisalina which also makes
BChl c (Borrego et al., 1998; Dubinina and Gorlenko, 1975; Gaisin et al., 2019; Liu et
al., 2012b; Smith and Goff, 1985; Tank et al., 2017). BChl e, which differs from BChl c
by replacement of the C7 methyl group with a formyl group, is only made by brown- 15 colored GSB. Finally, BChl f, like BChl d, lacks a methyl group at C20, but has a formyl
group at C7 like BChl e (Tamiaki et al., 2011; Vogl et al., 2012)
1.5 Organization of This Dissertation
The remaining chapters of this dissertation focus on two main topics related to
anoxygenic phototrophic bacteria. The first focus is on the biosynthetic pathways of
(B)Chls in green bacteria. Chapter 2 reviews the state of knowledge of the biosynthetic
pathways of Chl a, BChl a, and BChls c, d, e, and f in green bacteria. Chapter 3
describes the biochemical characterization of BciD, the final enzyme in the pathways
leading to bacteriochlorophyllide (BChlide) e and f, in addition to confirming the pigment phenotype of a bciD mutant in Cba. limnaeum. The goal of this study was to test the
hypothesis that BciD is a radical-SAM enzyme that is sufficient to convert BChlide c or d
into BChlide e or f. This was tested via heterologous expression of His-tagged BciD from
Cba. limneaum in E.coli. BciD-His was then incubated with appropriate substrates and
the reaction products were monitored by reversed-phased HPLC and confirmed via LC-
MS. The data supported the initial hypothesis and additionally led to the hypothesis that
BciD converts the methyl group of BChlide c or d into the formyl group of BChlide e or f
via two subsequent hydroxylation reactions. Chapter 4 concludes the discussion of
(B)Chl biosynthesis in green bacteria, describing bioinformatic analyses across the green
bacteria which were used to examine (B)Chl biosynthetic genes encoded in currently
available green bacterial genomes. These analyses were performed while preparing a
review of (B)Chl biosynthesis in green bacteria in order to examine the biosynthetic
capabilities of newly available green bacterial genomes (Chapter 2). The second focus of 16 this dissertation is on a newly isolated thermophilic purple sulfur bacterium (PSB) from
Mushroom Spring in Yellowstone National Park. This organism is referred to as
Thiohalocapsa MS throughout Chapter 5. In Chapter 5 characterization of the pigments and light-harvesting apparatus of Thiohalocapsa MS is presented. Thiohalocapsa MS is of particular interest because it is the first thermophilic Thiohalocapsa spp. identified and only the third thermophilic PSB. As the first thermophilic Thiohalocapsa sp. it was predicted to share characteristics with other Thiohalocapsa and with thermophilic PSB.
The study presented here confirms that Thiohalocapsa MS is similar to other
Thiohalocapsa spp. in its morphology and BChl a content but that the carotenoid content and LH1-RC complex of Thiohalocapsa MS are more similar to other thermophilic PSB.
In particular the LH1-RC complex absorbs light beyond 890 nm and experiences a blue shift of 5-8 nm in response to the loss of Ca2+ ions. Thess data support the hypothesis
that, like other thermophilic PSB, the Thiohalocapsa MS LH1-RC complex binds Ca2+
ions which enhance its thermostability. In summary, the work described in this
dissertation contributes to our knowledge of the pigments and light-harvesting apparatus
of two classes of anoxygenic phototrophic bacteria.
17 References Airs, R. L., Borrego, C. M., Garcia-Gil, J., and Keely, B. J. (2001). Identification of the bacteriochlorophyll homologues of Chlorobium phaeobacteroides strain UdG6053 grown at low light intensity. Photosynth. Res. 70, 221–230. doi:10.1023/A:1015146304441. Balban, T.S., Tamiaki, H., Holzwarth, A.R. (2005) Chlorins programed for self- assembly. Top. Curr. Chem. 258, 1-38. doi: 10.1007/b137480 Blankenship, R. E. (1994). Protein-structure, electron-transfer and evolution of prokaryotic photosynthetic reaction centers. Antonie Van Leeuwenhoek Int. J. Gen. Mol. Microbiol. 65, 311–329. doi:10.1007/BF00872216. Borrego, C. M., Garcia-Gil, J., Cristina, X. P., Vila, X., and Abella, C. A. (1998). Occurrence of new bacteriochlorophyll d forms in natural populations of green photosynthetic sulfur bacteria. FEMS Microbiol. Ecol. 26, 257–267. doi:10.1016/S0168-6496(98)00038-5. Bruce, B. D., Fuller, R. C., and Blankenship, R. E. (1982). Primary photochemistry in the facultatively aerobic green photosynthetic bacterium Chloroflexus-aurantiacus. Proc. Natl. Acad. Sci. USA 79, 6532–6536. doi:10.1073/pnas.79.21.6532. Bryant, D. A., and Canniffe, D. P. (2018). How nature designs light-harvesting antenna systems: Design principles and functional realization in chlorophototrophic prokaryotes. J. Phys. B At. Mol. Opt. Phys. 51, 033001. doi:10.1088/1361- 6455/aa9c3c. Bryant, D. A., and Frigaard, N. U. (2006). Prokaryotic photosynthesis and phototrophy illuminated. Trends Microbiol. 14, 488–496. doi:10.1016/j.tim.2006.09.001. Bryant, D. A., Garcia Costas, A. M., Maresca, J. A., Gomez Maqueo Chew, A., Klatt, C. G., Bateson, M. M., et al. (2007). “Candidatus Chloracidobacterium thermophilum”: an aerobic phototrophic Acidobacterium. Science. 317, 523–526. doi:10.1126/science.1143236. 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. doi:10.1021/bi026599s. Bryant, D. A., Liu, Z., Li, T., Zhao, F., Garcia Costas, A. M., Klatt, C. G., et al. (2012). “Comparative and functional genomics of anoxygenic green bacteria from the taxa Chlorobi, Chloroflexi, and Acidobacteria,” in Advances in Photosynthesis and Respiration, Vol. 35, Functional Genomics and Evolution of Photosynthetic Systems, R. Burnap and W. Vermaas, eds. (Dordrecht: Springer Netherlands), pp. 47–102. doi:10.1007/978-94-007-1533-2_3. Busch, M. S. A., Mueh, F., Madjet, M. E.-A., and Renger, T. (2011). The eighth bacteriochlorophyll completes the excitation energy funnel in the FMO protein. J. Phys. Chem. Lett. 2, 93–98. doi:10.1021/jz101541b. Cardona, T. (2015). A fresh look at the evolution and diversification of photochemical reaction centers. Photosynth. Res. 126, 111–134. doi:10.1007/s11120-014-0065-x. Charles, P., Kalendra, V., He, Z., Khurshov, V., van der Est, A., Golbeck, J. H., et al. (2019). Elucidating the role of zinc-bacteriochlorophyll a’ in the primary 18 photochemistry of Chloroacidobacterium thermophilum reaction centers. Biophys. J. 116, 419a. Cohen-Bazire, G., Pfennig, N., and Kunisawa, R. (1964). The fine structure of green bacteria. J. Cell Biol. 22, 207–225. doi:10.1083/jcb.22.1.207. Cole, J. R., Wang, Q., Fish, J. A., Chai, B., McGarrell, D. M., Sun, Y., et al. (2014). Ribosomal Database Project: Data and tools for high throughput rRNA analysis. Nucl. Acids Res. 42, D633–D642. doi:10.1093/nar/gkt1244. Collins, A. M., Wen, J., and Blankenship, R. E. (2012). “Photosynthetic Light-Harvesting Complexes,” in Molecular Solar Fuels RSC Energy and Environment Series., TJ Wydrzynski, and W. Hillier, eds., (London, RSC Publishing) pp. 85–106. doi:10.1039/9781849733038-00085. Dubinina, G. A., and Gorlenko, V. M. (1975). New filamentous photosynthetic green bacteria containing gas vacuoles. Microbiology 44, 452–458. Fenna, R. E., and Matthews, B. W. (1975). Chlorophyll arrangement in a bacteriochlorophyll protein from Chlorobium-limicola. Nature 258, 573–577. doi:10.1038/258573a0. Frigaard, N. U., and Bryant, D. A. (2006). “Chlorosomes: Antenna organelles in photosynthetic green bacteria,” in Complex Intracellular Structures in Prokaryotes, ed. J. M. Shively (Berlin, Heidelberg: Springer), pp. 79–114. doi:10.1007/7171_021. Frigaard, N. U., and Bryant, D. A. (2004). Seeing green bacteria in a new light: Genomics-enabled studies of the photosynthetic apparatus in green sulfur bacteria and filamentous anoxygenic phototrophic bacteria. Arch. Microbiol. 182, 265–276. doi:10.1007/s00203-004-0718-9. Frigaard, N. U., Li, H., Milks, K. J., and Bryant, D. A. (2004). Nine mutants of Chlorobium tepidum each unable to synthesize a different chlorosome protein still assemble functional chlorosomes. J. Bacteriol. 186, 646–653. doi:10.1128/JB.186.3.646-653.2004. Frigaard, N., and Dahl, C. (2009). “Sulfur Metabolism in Phototrophic Sulfur Bacteria.” In Advances in Microbial Physiology Vol 54, ed. R. K. Poole, (Academic Press) pp. 103-200 doi:10.1016/S0065-2911(08)00002-7. Fuchs, G. (2011). Alternative pathways of carbon dioxide fixation: insights into the early evolution of life? Annu. Rev. Microbiol. 65, 631–658. doi:10.1146/annurev-micro- 090110-102801. Gaisin, V.A., Burganskaya, E.I., Grouzdev, D.S., Ashikhmin, A.A., Kostrikina, N.A., Bryantseva, I.A. et al., (2019). ‘Candidatus Viridilinea mediisalina’, a novel phototrophic Chloroflexi bacterium from a Siberian soda lake. FEMS Microbiol. Lett. 366, fnz043. doi:10.1093/femsle/fnz043 Garcia Costas, A. M., Liu, Z., Tomsho, L. P., Schuster, S. C., Ward, D. M., and Bryant, D. A. (2012a). Complete genome of Candidatus Chloracidobacterium thermophilum, a chlorophyll-based photoheterotroph belonging to the phylum Acidobacteria. Environ. Microbiol. 14, 177–190. doi:10.1111/j.1462-2920.2011.02592.x. Garcia Costas, A. M., Tsukatani, Y., Rijpstra, W. I. C., Schouten, S., Welander, P. V., Summons, R. E., et al. (2012b). Identification of the bacteriochlorophylls, carotenoids, quinones, lipids, and hopanoids of “Candidatus Chloracidobacterium thermophilum.” J. Bacteriol. 194, 1158–1168. doi:10.1128/JB.06421-11. 19 Garcia Costas, A. M., Tsukatani, Y., Romberger, S. P., Oostergetel, G. T., Boekema, E. J., Golbeck, J. H., et al. (2011). Ultrastructural analysis and identification of envelope proteins of “Candidatus chloracidobacterium thermophilum” chlorosomes. J. Bacteriol. 193, 6701–6711. doi:10.1128/JB.06124-11. Ganapathy, S., Oostergetel, G.T., Wawrzyniak, P.K., Reus, M., Gomez Maqueo Chew, A., Buda, F., et al., (2009). Alternating syn-anti bacteriochlorophylls form concentric helical nanotubes in chlorosomes. Proc. Natl. Acad. Sci. USA 106, 8525–8530. doi:10.1073/pnas.0903534106 Garrity, G. M., Holt, J. G., Castenholz, R. W., Pierson, B. K., Keppen, O. I., and Gorlenko, V. M. (2001). “Phylum BVI. Chloroflexi phy. nov.,” in Bergey’s Manual of Systematic Bacteriology, (Springer, New York, NY) pp. 427–446. doi:10.1007/978-0-387-21609-6_23. Gisriel, C., Sarrou, I., Ferlez, B., Golbeck, J. H., Redding, K. E., and Fromme, R. (2017). Structure of a symmetric photosynthetic reaction center-photosystem. Science 357, 1021–1025. doi:10.1126/science.aan5611. Glaeser, J., Bañeras, L., Rütters, H., and Overmann, J. (2002). Novel bacteriochlorophyll e structures and species-specific variability of pigment composition in green sulfur bacteria. Arch. Microbiol. 177, 475–485. doi:10.1007/s00203-002-0416-4. Gomez Maqueo Chew, A., and Bryant, D. A. (2007). Chlorophyll biosynthesis in bacteria: the origins of structural and functional diversity. Annu. Rev. Microbiol. 61, 113–129. doi:10.1146/annurev.micro.61.080706.093242. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2008). Identification of the bchP gene, encoding geranylgeranyl reductase in Chlorobaculum tepidum. J. Bacteriol. 190, 747–749. doi:10.1128/JB.01430-07. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2007). Bacteriochlorophyllide c C-82 and C-121 methyltransferases are essential for adaptation to low light in Chlorobaculum tepidum. J. Bacteriol. 189, 6176–6184. doi:10.1128/JB.00519-07. Gorlenko, V.M., Bryantseva, I.A., Kalashnikov, A.M., Gaisin, V.A., Sukhacheva, M.V., Grouzdev, D.S., et al., (2014) Candidatus “Chloroploca Asiatic gen. nov., sp. nov., a new mesophilic filamentous anoxygenic phototrophic bacterium. Microbiology 83, 838-848 Gorlenko, V. M., and Korotkov, S. A. (1979). Morphological and physiological features of the new filamentous gliding green bacteria Oscillochloris-trichoides nov. comb. Izv. Akad. Nauk Sssr Seriya Biol., 848. Grouzdev, D. S., Kuznetsov, B. B., Keppen, O. I., Krasil’nikova, E. N., Lebedeva, N. V., and Ivanovsky, R. N. (2015). Reconstruction of bacteriochlorophyll biosynthesis pathways in the filamentous anoxygenic phototrophic bacterium Oscillochloris trichoides DG-6 and evolution of anoxygenic phototrophs of the order Chloroflexales. Microbiology 161, 120–130. doi:10.1099/mic.0.082313-0. Grouzdev, D. S., Rysina, M. S., Bryantseva, I. A., Gorlenko, V. M., and Gaisin, V. A. (2018). Draft genome sequences of `Candidatus Chloroploca asiatica’ and `Candidatus Viridilinea mediisalina’, candidate representatives of the Chloroflexales order: phylogenetic and taxonomic implications. Stand. Genomic Sci. 13, 24. doi:10.1186/s40793-018-0329-8. 20 Harada, J., Mizoguchi, T., Tsukatani, Y., Noguchi, M., and Tamiaki, H. (2012). A seventh bacterial chlorophyll driving a large light-harvesting antenna. Sci. Rep. 2, 671. doi:10.1038/srep00671. Heising, S., Richter, L., Ludwig, W., and Schink, B. (1999). Chlorobium ferrooxidans sp. nov., a phototrophic green sulfur bacterium that oxidizes ferrous iron in coculture with a “Geospirillum” sp. strain. Arch. Microbiol. 172, 116–124. doi:10.1007/s002030050748. Huang, R. Y.-C., Wen, J., Blankenship, R. E., and Gross, M. L. (2012). Hydrogen- deuterium exchange mass spectrometry reveals the interaction of Fenna-Matthews- Olson protein and chlorosome CsmA protein. Biochemistry 51, 187–193. doi:10.1021/bi201620y. Iino, T., Mori, K., Uchino, Y., Nakagawa, T., Harayama, S., and Suzuki, K. I. (2010). Ignavibacterium album gen. nov., sp. nov., a moderately thermophilic anaerobic bacterium isolated from microbial mats at a terrestrial hot spring and proposal of Ignavibacteria classis nov., for a novel lineage at the periphery of green sulf. Int. J. Syst. Evol. Microbiol. 60, 1376–1382. doi:10.1099/ijs.0.012484-0. Imhoff, J. F. (2017). “Anoxygenic phototrophic bacteria from extreme environments,” in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, P.C. Hallenbeck, ed. (Berlin: Springer), pp. 427–480. doi:10.1007/978-3-319-46261- 5_13. Imhoff, J. F., and Thiel, V. (2010). Phylogeny and taxonomy of Chlorobiaceae. Photosynth. Res. 104, 123–136. doi:10.1007/s11120-009-9510-7. Ivanovsky, R. N., Fal, Y. I., Berg, I. A., Ugolkova, N. V., Krasilnikova, E. N., Keppen, O. I., et al. (1999). Evidence for the presence of the reductive pentose phosphate cycle in a filamentous anoxygenic photosynthetic bacterium, Oscillochloris trichoides strain DG-6. Microbiology 147, 1743–1748. doi:10.1099/13500872-145-7-1743. Kadnikov, V. V., Mardanov, A. V., Podosokorskaya, O. A., Gavrilov, S. N., Kublanov, I. V., Beletsky, A. V., et al. (2013). Genomic analysis of Melioribacter roseus, facultatively anaerobic organotrophic bacterium representing a novel deep lineage within Bacteriodetes/Chlorobi group. PLoS One 8, e53047. doi:10.1371/journal.pone.0053047. Keppen, O. I., Baulina, O. I., and Kondratieva, E. N. (1994). Oscillochloris trichoides neotype strain DG-6. Photosynth. Res. 41, 29–33. doi:10.1007/BF02184143. Klatt, C. G., Bryant, D. A., and Ward, D. M. (2007). Comparative genomics provides evidence for the 3-hydroxypropionate autotrophic pathway in filamentous anoxygenic phototrophic bacteria and in hot spring microbial mats. Environ. Microbiol. 9, 2067– 2078. doi:10.1111/j.1462-2920.2007.01323.x. Klatt, C. G., William, P., Herrgard, M. J., Jay, Z. J., Rusch, D. B., and Miller, S. R. (2013). Community structure and function of high-temperature chlorophototrophic microbial mats inhabiting diverse geothermal environments. Front. Microbiol. 4, 106. doi:10.3389/fmicb.2013.00106. Larson, C. R., Seng, C. O., Lauman, L., Matthies, H. J., Wen, J., Blankenship, R. E., et al. (2011). The three-dimensional structure of the FMO protein from Pelodictyon phaeum and the implications for energy transfer. Photosynth. Res. 107, 139–150. doi:10.1007/s11120-010-9604-2. 21 Letunic, I., and Bork, P. (2016). Interactive tree of life (iTOL) v3: an online tool for the display and annotation of phylogenetic and other trees. Nucl. Acids Res. 44, W242– W245. doi:10.1093/nar/gkw290. Li, H., and Bryant, D. A. (2009). Envelope proteins of the CsmB/CsmF and CsmC/CsmD motif families influence the size, shape, and composition of chlorosomes in Chlorobaculum tepidum. J. Bacteriol. 191, 7109–7120. doi:10.1128/JB.00707-09. Li, H., Frigaard, N. U., and Bryant, D. A. (2013). Proteins in chlorosomes: CsmI and CsmJ participate in light-dependent control of energy transfer in chlorosomes of Chlorobaculum tepidum. Biochemistry 52, 1321–1330. doi:10.1021/bi301454g. 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 45, 9095–9103. doi:10.1021/bi060776y. Liu, Z., Frigaard, N., Vogl, K., Iino, T., Ohkuma, M., Overmann, J., et al. (2012a). Complete genome of Ignavibacterium album , a metabolically versatile , flagellated , facultative anaerobe from the phylum Chlorobi. Front. Microbiol. 3, 185. doi:10.3389/fmicb.2012.00185. Liu, Z., Klatt, C. G., Ludwig, M., Rusch, D. B., Jensen, S. I., Kühl, M., et al. (2012b). “Candidatus Thermochlorobacter aerophilum”: An aerobic chlorophotoheterotrophic member of the phylum Chlorobi defined by metagenomics and metatranscriptomics. ISME J. 6, 1869–1882. doi:10.1038/ismej.2012.24. Majumder, E. L. W., Olsen, J. D., Qian, P., Collins, A. M., Hunter, C. N., and Blankenship, R. E. (2016). Supramolecular organization of photosynthetic complexes in membranes of Roseiflexus castenholzii. Photosynth. Res. 127, 117–130. doi:10.1007/s11120-015-0179-9. Maresca, J. A., Gomez Maqueo Chew, A., Ponsatí, M. R., Frigaard, N. U., Ormerod, J. G., and Bryant, D. A. (2004). The bchU gene of Chlorobium tepidum encodes the C- 20 methyltransferase in bacteriochlorophyll c biosynthesis. J. Bacteriol. 186, 2558– 2566. doi:10.1128/JB.186.9.2558-2566.2004. Milder, M. T. W., Brueggemann, B., van Grondelle, R., and Herek, J. L. (2010). Revisiting the optical properties of the FMO protein. Photosynth. Res. 104, 257–274. doi:10.1007/s11120-010-9540-1. Montaño, G. A, Bowen, B. P., LaBelle, J. T., Woodbury, N. W., Pizziconi, V. B., and Blankenship, R. E. (2003). Characterization of Chlorobium tepidum chlorosomes: a calculation of bacteriochlorophyll c per chlorosome and oligomer modeling. Biophys. J. 85, 2560–2565. doi:10.1016/S0006-3495(03)74678-5. Munoz, R., Yarza, P., Ludwig, W., Euzéby, J., Amann, R., Schleifer, K. H., et al. (2011). Release LTPs104 of the All-Species Living Tree. Syst. Appl. Microbiol. 34, 169–170. doi:10.1016/j.syapm.2011.03.001. Nadson, G. A. (1906). The morphology of inferior algae. III. Chlorobium limicola Nads., the green chlorophyll bearing microbe. Bull. du Jard. impérial Bot. St.-pétersbg. 6, 190. Nielsen, J. T., Kulminskaya, N. V, Bjerring, M., Linnanto, J. M., Ratsep, M., Pedersen, M. O., et al. (2016). In situ high-resolution structure of the baseplate antenna complex in Chlorobaculum tepidum. Nat. Commun. 7, 12454. doi:10.1038/ncomms12454. 22 Novoderezhkin, V. I., Taisova, A. S., Fetisova, Z., Blankenship, R. E., Savikhin, S., Buck, D. R., et al. (1998). Energy transfers in the B808-866 antenna from the green bacterium Chloroflexus aurantiacus. Biophys. J. 74, 2069–2075. doi:10.1016/S0006- 3495(98)77913-5. Ohashi, S., Iemura, T., Okada, N., Itoh, S., Furukawa, H., Okuda, M., et al. (2010). An overview on chlorophylls and quinones in the photosystem I-type reaction centers. Photosynth. Res. 104, 305–319. doi:10.1007/s11120-010-9530-3. Oostergetel, G. T., van Amerongen, H., and Boekema, E. J. (2010). The chlorosome: A prototype for efficient light harvesting in photosynthesis. Photosynth. Res. 104, 245– 255. doi:10.1007/s11120-010-9533-0. Orf, G. S., and Blankenship, R. E. (2013). Chlorosome antenna complexes from green photosynthetic bacteria. Photosynth. Res. 116, 315–331. doi:10.1007/s11120-013- 9869-3. Orf, G. S., Tank, M., Vogl, K., Niedzwiedzki, D. M., Bryant, D. A., and Blankenship, R. E. (2013). Spectroscopic insights into the decreased efficiency of chlorosomes containing bacteriochlorophyll f. Biochim. Biophys. Acta 1827, 493–501. doi:10.1016/j.bbabio.2013.01.006. Permentier, H. P., Schmidt, K. A., Kobayashi, M., Akiyama, M., Hager-Braun, C., Neerken, S., et al. (2000). Composition and optical properties of reaction centre core complexes from the green sulfur bacteria Prosthecochloris aestuarii and Chlorobium tepidum. Photosynth. Res. 64, 27–39. doi:10.1023/A:1026515027824. Pierson, B. K., and Thornber, J. P. (1983). Isolation and spectral characterization of photochemical reaction centers from the thermophilic green bacterium Chloroflexus aurantiacus strain J-10-fl. Proc. Natl. Acad. Sci. USA 80, 80–84. doi:10.1073/pnas.80.1.80. Pierson, B. K., and Castenholz, R. W. (1974a). Studies of pigments and growth in Chloroflexus aurantiacus, a phototrophic filamentous bacterium. Arch. Microbiol. 100, 283–305. doi:10.1007/BF00446324. Pierson, B. K., and Castenholz, R. W. (1974b). A phototrophic gliding filamentous bacterium of hot springs, Chloroflexus aurantiacus, gen. and sp. nov. Arch. Microbiol. 100, 5–24. doi:10.1007/BF00446302. Podosokorskaya, O. A., Kadnikov, V. V., Gavrilov, S. N., Mardanov, A. V., Merkel, A. Y., Karnachuk, O. V., et al. (2013). Characterization of Melioribacter roseus gen. nov., sp. nov., a novel facultatively anaerobic thermophilic cellulolytic bacterium from the class Ignavibacteria, and a proposal of a novel bacterial phylum Ignavibacteriae. Environ. Microbiol. 15, 1759–1771. doi:10.1111/1462-2920.12067. Price, M. N., Dehal, P. S., and Arkin, A. P. (2009). Fasttree: Computing large minimum evolution trees with profiles instead of a distance matrix. Mol. Biol. Evol. 26, 1641– 1650. doi:10.1093/molbev/msp077. Pruesse, E., Peplies, J., and Glöckner, F. O. (2012). SINA: Accurate high-throughput multiple sequence alignment of ribosomal RNA genes. Bioinformatics 28, 1823– 1829. doi:10.1093/bioinformatics/bts252. Pšenčík, J., Butcher, S. J., and Tuma, R. (2014). “Chlorosomes: Structure, Function and Assembly,” in Structural Basis of Biological Energy Generation Advances in 23 Photosynthesis and Respiration., eds. Govindjee and S. T. (Dordrecht, Netherlands: Springer), pp. 77–109. doi:10.1007/978-94-017-8742-0_5. Pšenčík, J., Collins, A. M., Liljeroos, L., Torkkeli, M., Laurinmäki, P., Ansink, H. M., et al. (2009). Structure of chlorosomes from the green filamentous bacterium Chloroflexus aurantiacus. J. Bacteriol. 191, 6701–6708. doi:10.1128/JB.00690-09. Pšenčík, J., Torkkeli, M., Zupčanová, A., Vácha, F., Serimaa, R. E., and Tuma, R. (2010). The lamellar spacing in self-assembling bacteriochlorophyll aggregates is proportional to the length of the esterifying alcohol. Photosynth. Res. 104, 211–219. doi:10.1007/s11120-010-9541-0. Saer, R. G., and Blankenship, R. E. (2017). Light harvesting in phototrophic bacteria: structure and function. Biochem. J. 474, 2107–2131. doi:10.1042/BCJ20160753. Smith, K. M., (1994). Nomenclature of the bacteriochlorophylls c, d, and e. Photosynth. Res. 41. 23-26. doi: 10.1007/BF02184141 Smith, K. M., and Goff, D. A. (1985). Bacteriochlorophylls-d from Chlorobium- vibrioforme - chromatographic separations and structural assignments of the methyl bacteriopheophorbides. J. Chem. Soc. Trans. 1, 1099–1113. doi:10.1039/p19850001099. Staehelin, L. A., Golecki, J. R., and Drews, G. (1980). Supramolecular organization of chlorosomes (Chlorobium vesicles) and of their membrane attachment sites in Chlorobium limicola. Biochim. Biophys. Acta 589, 30–45. doi:10.1016/0005- 2728(80)90130-9. Staehelin, L. A., Golecki, J. R., Fuller, R. C., and Drews, G. (1978). Visualization of the supramolecular architecture of chlorosomes (Chlorobium type vesicles) in freeze- fractured cells of Chloroflexus aurantiacus. Arch. Microbiol. 119, 269–277. doi:10.1007/BF00405406. Steensgaard, D.B., Wackerbarth, H., Hildebrandt, P., Holzwarth, A.R. (2000). Diastereoselective control of bacteriochlorophyll e aggregation. 31-S-BChl e is essential for the formation of chlorosome-like aggregates. J. Phys. Chem. B. 104, 10379–10386. Tamiaki H., Komada J., Kunieda M., Fukai K., Yoshitomi T., Harada J., Mizoguchi T. (2011). In vitro synthesis and characterization of bacteriochlorophyll-f and its absence in bacteriochlorophyll-e producing organisms. Photosynth. Res. 107, 133-8. doi: 10.1007/s11120-010-9603-3 Tank, M., and Bryant, D. A. (2015a). Nutrient requirements and growth physiology of the photoheterotrophic Acidobacterium, Chloracidobacterium thermophilum. Front. Microbiol. 6, 226. doi:10.3389/fmicb.2015.00226. Tank, M., and Bryant, D. A. (2015b). Chloracidobacterium thermophilum gen. nov., sp. nov.: An anoxygenic microaerophilic chlorophotoheterotrophic acidobacterium. Int. J. Syst. Evol. Microbiol. 65, 1426–1430. doi:10.1099/ijs.0.000113. Tank, M., Garcia Costas, A. M., and Bryant, D. A. (2018). “Chloracidobacterium,” in Bergey’s Manual of Systematics of Archaea and Bacteria, eds. W. B. Whitman, F. Rainey, P. Kämpfer, M. Trujillo, J. Chun, P. DeVos, et al. (American Cancer Society), pp. 1–9. doi:10.1002/9781118960608.gbm01534. Tank, M., Thiel, V., Ward, D. M., and Bryant, D. A. (2017). A panoply of phototrophs: An overview of the thermophilic chlorophototrophs of the microbial mats of alkaline 24 siliceous hot springs in Yellowstone National Park, WY, USA. In: Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, P. C. Hallenbeck, ed. (Berlin: Springer), pp. 87–137. doi:10.1007/978-3-319-46261-5_3. Thiel, V., Tank, M., and Bryant, D. A. (2018). Diversity of chlorophototrophic bacteria revealed in the omics era. Annu. Rev. Plant Biol. 69, 21–49. doi:10.1146/annurev- arplant-042817-040500. Tronrud, D. E., Wen, J., Gay, L., and Blankenship, R. E. (2009). The structural basis for the difference in absorbance spectra for the FMO antenna protein from various green sulfur bacteria. Photosynth. Res. 100, 79–87. doi:10.1007/s11120-009-9430-6. Tsukatani, Y., Mizoguchi, T., Thweatt, J., Tank, M., Bryant, D. A., and Tamiaki, H. (2016). Glycolipid analyses of light-harvesting chlorosomes from envelope protein mutants of Chlorobaculum tepidum. Photosynth. Res. 128, 235–241. doi:10.1007/s11120-016-0228-z. Tsukatani, Y., Romberger, S. P., Golbeck, J. H., and Bryant, D. A. (2012). Isolation and characterization of homodimeric type-I reaction center complex from Candidatus Chloracidobacterium thermophilum, an aerobic chlorophototroph. J. Biol. Chem. 287, 5720–5732. doi:10.1074/jbc.M111.323329. Tsukatani, Y., Wen, J., Blankenship, R. E., and Bryant, D. A. (2010). Characterization of the FMO protein from the aerobic chlorophototroph, Candidatus Chloracidobacterium thermophilum. Photosynth. Res. 104, 201–209. doi:10.1007/s11120-009-9517-0. van Gemerden, H., and Mas, J. (1995). “Ecology of phototrophic sulfur bacteria,” in Advances in Photosynthesis and Respiration, Vol. 2, Anoxygenic photosynthetic bacteria, R. E. Blankenship, M. T. Madigan, and C. E. Bauer, eds. (Dordrecht, Netherlands: Springer), pp. 49–85. Vogl, K., Tank, M., Orf, G. S., Blankenship, R. E., and Bryant, D. A. (2012). Bacteriochlorophyll f: Properties of chlorosomes containing the “forbidden chlorophyll.” Front. Microbiol. 3, 298. doi:10.3389/fmicb.2012.00298. Wahlund, T. M., and Madigan, M. T. (1993). Nitrogen fixation by the thermophilic green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 175, 474–478. doi:10.1128/jb.175.2.474-478.1993. Wahlund, T. M., and Tabita, F. R. (1997). The reductive tricarboxylic acid cycle of carbon dioxide assimilation: Initial studies and purification of ATP-citrate lyase from the green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 179, 4859–4867. doi:10.1128/jb.179.15.4859-4867.1997. Wen, J., Tsukatani, Y., Cui, W., Zhang, H., Gross, M. L., Bryant, D. A., et al. (2011). Structural model and spectroscopic characteristics of the FMO antenna protein from the aerobic chlorophototroph, Candidatus Chloracidobacterium thermophilum. Biochim. Biophys. Acta 1807, 157–164. doi:10.1016/j.bbabio.2010.09.008. Wen, J., Zhang, H., Gross, M. L., and Blankenship, R. E. (2009). Membrane orientation of the FMO antenna protein from Chlorobaculum tepidum as determined by mass spectrometry-based footprinting. Proc. Natl. Acad. Sci. USA 106, 6134–6139. doi:10.1073/pnas.0901691106. Woodbury, N. W., Becker, M., Middendorf, D., and Parson, W. W. (1985). Picosecond kinetics of the initial photochemical electron-transfer reaction in bacterial 25 photosynthetic reaction centers. Biochemistry 24, 7516–7521. doi:10.1021/bi00347a002. Xin, Y., Lin, S., Montaño, G. A., and Blankenship, R. E. (2005). Purification and characterization of the B808-866 light-harvesting complex from green filamentous bacterium Chloroflexus aurantiacus. Photosynth. Res. 86, 155–163. doi:10.1007/s11120-005-5103-2. Xin, Y., Shi, Y., Niu, T., Wang, Q., Niu, W., Huang, X., et al. (2018). Cryo-EM structure of the RC-LH core complex from an early branching photosynthetic prokaryote. Nat. Commun. 9, 1568. doi:10.1038/s41467-018-03881-x. Zeng, Y., Feng, F., Medova, H., Dean, J., and Kobli ek, M. (2014). Functional type 2 photosynthetic reaction centers found in the rare bacterial phylum Gemmatimonadetes. Proc. Natl. Acad. Sci. USA 111, 7795–7800. doi:10.1073/pnas.1400295111. Zeng, Y., and Koblížek, M. (2017). “Phototrophic Gemmatimonadetes: A new ‘purple’ branch on the bacterial tree of life,” in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, P. C. Hallenbeck, ed. (Berlin: Springer), pp. 163–192. doi:10.1007/978-3-319-46261-5_5. Zill, J. C., He, Z., Tank, M., Ferlez, B. H., Canniffe, D. P., Lahav, Y., et al. (2018). 15N photo-CIDNP MAS NMR analysis of reaction centers of Chloracidobacterium thermophilum. Photosynth. Res. 137, 295–305. doi:10.1007/s11120-018-0504-1.
26 A
Figure 1.1 Phylogeny of Green Bacteria A. Phylogenetic tree modified from SILVA living tree project 16S rRNA gene tree (release 128) to show the major phyla of Bacteria (Munoz et al., 2011). Boxed phyla contain at least one chlorophototrophic member. Phyla with type-1 RCs are boxed or highlighted in purple while phyla with type-2 RCs are boxed or highlighted in magenta; the 2 phyla which have not been biochemically characterized are highlighted in a lighter shade of magenta; cyanobacteria which contain PSI and PSII are highlighted in teal. Phyla containing at least one member of the green bacteria (indicating the use of chlorosomes for light harvesting) are additionally highlighted in green. B. Phylogenic tree of the class level branches which contain green bacteria. Green bacterial members within each class are highlighted in green. The 16S rRNA genes from cultured type strains available in the RDP database in addition to those from organisms with sequenced genomes (see Figure 2.1) were aligned with the SINA aligner, and the tree was computed with FastTree (Cole et al., 2014; Price et al., 2009; Pruesse et al., 2012). Phylogentic trees were modified using iTol (Letunic and Bork, 2016).
27 B
28
Figure 1.2 Localization of Chlorophylls and Bacteriochlorophyls in Green Bacteria Diagram of light-harvesting apparatus of green bacteria indicating localization of (B)Chls and path of excitation energy. The “*” indicates a molecule found only in Cab. thermophilum. The crystal structure of FMO (PDB:3ENI) from Cba. tepidum was used to represent the FMO proteins of Chlorobia and Cab. thermophilum. The structure of CsmA (PDB:2K37) from Cba. tepidum was used to represent the CsmA baseplate arrays of Chlorobia, Cab. thermophilum, and green FAPs. The cryo-EM structure of RC-LH core complex (PDB:5YQ7) of Roseiflexus castenholzii with the cytochrome subunit removed from view was used to represent the LH-RC of green FAPs.
29
Figure 1.3 Structure of Chlorophylls and Bacteriochlorophylls from Green Bacteria Structures of (B)Chls found in green bacteria. Numbering listed on Chl aPD molecule is used throughout review. Blue shading indicates structural differences from Chl aPD. Dashed boxes indicate differences in BChl d, e, and f from BChl c.
30 Chapter 2 Biosynthesis of Chlorophylls and Bacteriochlorophylls in Green Bacteria
Publication: Jennifer L. Thweatt, Daniel P. Canniffe, and Donald A. Bryant (2019)
Biosynthesis of chlorophylls and bacteriochlorophylls in green bacteria. Advances in
Botanical Research. 90, 35-89.
Contributions: JLT wrote the review of chlorophyll and bacteriochlorophyll biosynthesis in green bacteria, performed bioinformatic analyses related to this review (see Chapter
4) and made all figures. DPC drafted much of the section on reduction of the 8V group,
and discovery of the role of COR in the production of BChl b and BChl g. DAB and DPC
provided feedback, editing, and additions to the original draft.
31 2.1 Abstract
The biosynthetic pathways leading to the chlorophylls (Chls) and
bacteriochlorophylls (BChls) of green bacteria have been nearly fully elucidated over the
past two decades. This chapter reviews the biosynthetic pathways leading from
protoporphyrinogen IX to the production of Chls and BChls, specifically found in green
bacteria, in addition to discussing some earlier steps in these pathways. The physiological
diversity of green bacteria, which are distantly related, chlorosome-producing bacteria
(see Chapter 1), affects the particular combinations of biosynthetic enzymes that are used in different organisms. The distribution of these biosynthetic enzymes across the green bacteria in relation to their physiology is discussed. Additionally, the phylogenetic relationships and inferences about the evolution of (B)Chl biosynthesis are discussed in relation to some enzymes or enzyme classes. 32 2.2 Introduction
The biosynthetic pathways for Chl a and BChl a were initially studied in plants, algae, cyanobacteria, and purple bacteria. Homologs of the genes encoding enzymes for
BChl a synthesis in purple bacteria were subsequently found in the genomes of green bacteria (Bryant et al., 2012; Eisen et al., 2002; Garcia Costas et al., 2012a; Grouzdev et al., 2015; Tang et al., 2011). The unique enzymatic steps in the biosynthetic pathways of
BChl c, d, e, and f were characterized in the model GSB Cba. tepidum and Cba. limnaeum, and bioinformatic analyses and biochemical studies have confirmed that essentially the same pathways are present in other green bacteria as well. To date, finished or nearly complete permanent draft genomes are available for 17 Chlorobia, 7 green FAPs, and Cab. thermophilum (Figure 2.1). The availability of complete genomes for many green bacteria has made it possible to compare their biosynthetic capabilities
(Figure 2.1). In the following sections I will discuss the biosynthetic pathways of
(B)Chls in green bacteria (also see Chapter 4 for bioinformatic analyses conducted in the preparation of this review).
2.3 Early Steps
The biosynthesis of all hemes, Chls, BChls, cobalamin, and other tetrapyrroles proceeds via the same basic pathway from 5-aminolevulinate to uroporphyrinogen III.
The pathways for biosynthesis of Chls and BChls, and the porphyrin-dependent heme pathway, then proceed via the same enzymatic steps from uroporphyrinogen III to protoporphyrin IX (Proto IX) while the pathways for cobalamin and other tetrapyrroles, and the siroheme-dependent heme pathway diverge at uroporphyrinogen III, and the 33 coproporphyrin-dependent heme pathway diverges at coproporphyrinogen. (Dailey et al.,
2017). An in-depth discussion of these reactions can be found elsewhere (Fan et al.,
2019) and they will not be considered here except briefly to address distribution or other features important to green bacteria. A recent review of heme biosynthesis in prokaryotes has renamed several genes and gene products in this pathway (Dailey et al., 2017). Here I will use the old gene designations followed by the new ones in parentheses.
Two steps in the pathway from 5-aminolevulinic acid to Proto IX can be catalyzed by alternative enzymes (Figure 2.2). The first of these reactions is the oxidative decarboxylation of two of the four propionate groups of coproporphyrinogen III to produce the vinyl groups of protoporphyrinogen IX. In some aerobic organisms these reactions are performed by coproporphyrinogen decarboxylase, HemF (CgdC), which is an oxygen-dependent enzyme. In anaerobic organisms, an unrelated oxygen-independent, radical-SAM enzyme, coproporphyrinogen dehydrogenase, HemN (CgdH), forms protoporphyrinogen IX (Layer et al., 2005). Genes encoding both HemF (CgdC) and
HemN (CgdH) occur in the genomes of Chloroflexus spp., Cab. thermophilum, and Ca. T aerophilum that can grow aerobically or microaerophilically. In contrast, the strictly anaerobic GSB, Osc. trichoides, ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ only contain genes encoding HemN (CgdH) (Figure 2.1). Two copies of hemN (cgdH) are present in the genomes of all green bacteria examined for this review except Ca. T. aerophilum
(Figure 2.1), and the second paralog has been variably called hemN2 or hemZ (Eisen et al., 2002; Garcia Costas et al., 2012a; Tang et al., 2011). Recently, homologs of HemN
(CgdH) in other bacteria have been characterized as heme-binding proteins without 34 dehydrogenase activity and renamed HemW (Abicht et al., 2012; Haskamp et al., 2018).
Comparing the sequences of green bacterial HemN homologs to biochemically
characterized HemW, it is apparent that Ca. T. aerophilum lacks hemN and that all green
bacterial genomes including Ca. T. aerophilum have one copy of hemW (Figure 2.1)(see
Chapter 4).
The next reaction in the pathway, the 6-electron oxidative conversion of
protoporphyrinogen IX to Proto IX, is also performed by different isofunctional enzymes
(Figure 2.2). In some aerobic organisms an oxygen-dependent protoporphyrinogen IX
oxidase, HemY (PgoX), performs this oxidation (Dailey et al., 2017; Dailey and Dailey,
1996). HemY (PgoX) uses a single FAD molecule to perform three sequential, two-
electron oxidations using three molecules of O2 (the electron acceptor) and producing
Proto IX and three molecules of H2O2 (Dailey et al., 2017). Recently it has been shown that some homologs of HemY (PgoX) actually prefer coproporphrinogen III as a substrate. These enzymes have been designated coproporphyrinogen oxidases (CgoX), and they function in the coproporphyrin-dependent heme biosynthetic pathway that does not include Proto IX as an intermediate (Dailey et al., 2017). While these oxidases were originally described as oxygen-dependent enzymes, HemY from Plasmodium falciparum functions as a protoporphyrinogen oxidase under anoxic conditions in the presence of
FAD, NAD+, or NADP+ (Nagaraj et al., 2010). In some anaerobic organisms the oxygen-
independent enzyme protoporphryinogen dehydrogenase, HemG (PgdH1), is used. This
enzyme carries out the same six-electron oxidation as HemY but can use various electron acceptors including ubiquinone and menaquinone in vitro. It has been suggested that 35 HemG (PgdH1) activity may be coupled to the electron transport chain in vivo (Boynton
et al., 2009; Mobius et al., 2010; Sasarman et al., 1993). A third enzyme, known as HemJ
(PgdH2; note that PgdH1 and PgdH2 are not paralogs and do not belong to the same
protein family), has recently been identified that can perform this reaction in
cyanobacteria; however, no homologs of HemJ are encoded in any green bacterial
genomes (Boynton et al., 2011; Dailey et al., 2017; Kato et al., 2010; Skotnicová et al.,
2018).
Genes encoding HemY (PgoX) are found in the microaerophilic
chlorophototrophs, Chloroflexus spp. and in the aerobic Ca. T. aerophilum. Searches of the genomes of GSB for genes related to protoporphyrinogen IX oxidation have produced conflicting results. One study failed to identify any known gene(s) for protoporphyrinogen IX oxidation in GSB (Kato et al., 2010). However, more recent analyses show the presence of hemY (pgoX)-related genes in several GSB, while hemG
(pgdH1) was only found in Prosthecochloris (Ptc.) sp. BS-1 (Dailey et al., 2017;
Kobayashi et al., 2014). Surprisingly, the microaerophile, Cab. thermophilum, has a hemY homolog that is more closely related to those found in GSB, while the anaerobes,
Osc. trichoides, ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ have a hemY gene closely related to those of Chloroflexus spp. and Ca. T. aerophilum (Figure 2.1) (see Chapter
4). Considering that GSB are strict anaerobes, it is likely that their hemY-related genes encode an oxygen-independent enzyme for protoporphyrinogen oxidation, or that another as-yet unidentified enzyme performs this reaction. Further research on the putative HemY 36 enzymes of green bacteria is needed to understand how protoporphyrinogen IX oxidation occurs in these organisms.
2.4 Protoporphyrin IX to Chlorophyllide a
2.4.1 Magnesium Chelation
The biosynthesis of all (B)Chls in green bacteria proceeds via the same pathway from Proto IX to a branch point at chlorophyllide (Chlide) a. The first committed step of
(B)Chl biosynthesis is the insertion of the Mg++ ion into the macrocycle. This reaction requires an ATP-dependent enzyme, Proto IX magnesium (Mg) chelatase (MgCH).
MgCH uses ~15 ATP molecules, Mg++, and Proto IX to produce Mg-Proto IX (Reid and
Hunter, 2004) (Figure 2.2). This enzyme is made up of three subunits, denoted
ChlH/BchH, ChlD/BchD, and ChlI/BchI in all chlorophototrophic bacteria; conventionally, the Chl or Bch prefix is used depending on whether the gene/protein in question is associated with a Chl or BChl biosynthetic pathway, respectively (Willows et al., 1996). ChlH/BchH is the large subunit of MgCH which binds the Proto IX and Mg++ substrates (Sirijovski et al., 2008). ChlD/BchD and ChlI/BchI are smaller subunits with similar structures, which make homohexamers or homoheptamers, contain AAA+ domains, and associate with ChlH/BchH (Fodje et al., 2001; Lundqvist et al., 2010; Reid et al., 2003). Although both subunits have AAA+ domains, the ATPase activity resides in the ChlI/BchI subunit (Jensen et al., 1999).
Multiple copies of the chlH/bchH genes are present in most green bacteria
(Figure 2.1) (Bryant et al., 2012; Bryant and Liu, 2013; Eisen et al., 2002; Tang et al.,
2011). Because all green bacteria must make multiple types of (B)Chl molecules (see 37 above), it has been hypothesized that organisms may have different types ChlH/BchHDI
to channel Mg-Proto IX toward different end products and to allow differential feedback
regulation of the first committed step of (B)Chl biosynthesis (Gomez Maqueo Chew et
al., 2009). Mutational studies and in vitro gene expression of bchH paralogs from Cba.
tepidum seem to support this hypothesis. Three bchH genes, denoted bchH, bchS, and
bchT, are present in this GSB. Single mutants of all three paralogs were shown to have
MgCH activity producing all three (B)Chls found in GSB; however, a mutant lacking
BchS showed a significant decrease in BChl c content (Gomez Maqueo Chew et al.,
2009). Double mutants lacking bchH and bchT (functional BchS) or bchS and bchT
(functional BchH) also made all three (B)Chls; however, attempts to make a bchH bchS double mutant (functional BchT) failed. The bchS bchT double mutant made similarly low amounts of BChl c as the bchS single mutant (Gomez Maqueo Chew et al., 2009). In vitro studies with BchH, BchS, or BchT from Cba. tepidum produced in E. coli and reconstituted with BchD and BchI showed that BchHDI, BchSDI, and BchTDI all had
MgCH activity. The specific activity of BchSDI was two orders of magnitude higher than that of BchHDI and five orders of magnitude higher than that of BchTDI (Johnson and
Schmidt-Dannert, 2008). These data agree well with the in vivo studies and the substrate channeling hypothesis. Additionally, in Chloroflexus spp., which synthesize only two types of (B)Chl but experience both oxic and anoxic conditions, there are also three paralogs of bchH: two bchH-like paralogs and one bchS-like paralog. It has been suggested that the bchH-like paralogs might be differentially regulated on the basis of oxygen levels (Tang et al., 2011). 38 Interestingly, there is only one copy of the bchH gene in Cab. thermophilum,
which synthesizes Chl a, BChl a, Zn-BChl a′, and BChl c and probably experiences variations in oxygen levels (Bryant and Liu, 2013; Garcia Costas et al., 2012a, 2012b). At present the nature of the chelatase, if any, which inserts Zn++ into the macrocycle for the
biosynthesis of Zn-BChl a′ in Cab. thermophilum is unknown. In vitro studies have shown that Proto IX ferrochelatase of Rba. sphaeroides, which catalyzes Fe++ insertion
into Proto IX during heme biosynthesis, can insert various divalent metal cations into the
macrocycle, including Zn++ (Jones and Jones, 1970). However; it is unknown if this
enzyme performs this function in vivo in Cab. thermophilum. The mechanism of Zn++
insertion into the macrocycle is an open question in both purple bacteria and Cab.
thermophilum (Jaschke et al., 2011). It is possible that this reaction occurs spontaneously
in the latter organism, which only requires very small amounts of Zn-BChl a′ in RCs.
2.4.2 C13 Propionate Methylation
The insertion of Mg into Proto IX is followed by the conversion of Mg-Proto IX into Mg-Proto IX 13-monomethyl ester. This reaction involves methylation of the C13 propionate group of Mg-Proto IX and is catalyzed by Mg-Proto IX methyltransferase,
ChlM/BchM (Figure 2.2). ChlM/BchM is a SAM-dependent methyltransferase and is present in most chlorophototrophic bacteria (Bollivar et al., 1994a; Gibson and Hunter,
1994). Unlike the previous enzyme in the pathway, only one copy of chlM/bchM is present in the genomes of green bacteria and thus must be used to synthesize all (B)Chl molecules (Bryant and Liu, 2013). ChlHDI/BchHDI and ChlM/BchM have been shown to interact and possibly form a multi-enzyme complex (Sawicki and Willows, 2010). 39 Studies in cyanobacteria and purple bacteria established that ChlH/BchH affects the activity of ChlM/BchM (Hinchigeri et al., 1997; McLean and Hunter, 2009; Shepherd et al., 2005). In green bacteria, experiments comparing the effect of BchH, BchS, and BchT from Cba. tepidum on BchM showed that BchS and BchT increased BchM activity while
BchH decreased activity (Johnson and Schmidt-Dannert, 2008). These experiments further support the idea that the BchH subunits of green bacteria may be involved in substrate channeling and regulation of (B)Chl biosynthesis. The draft genome of Ca. T. aerophilum is notably lacking bchM and also lacks any known genes for isocyclic E-ring formation (see below, see Chapter 4; Figure 2.1). Possible explanations are that these genes are in a currently unsequenced region of the genome, that a different methyltransferase is used in lieu of BchM, or that the E-ring is formed via an uncharacterized reaction which does not require the formation of the methylester.
2.4.3 Isocyclic E-Ring Formation
Mg-Proto IX 13-monomethyl ester is converted to 8-vinyl-protochlorophyllide
(8V-PChlide) by one of several enzymes classified as oxidative isocylic ring cyclases, also known as Mg-Proto IX monomethylester oxidative cyclase. Like the oxidative reactions discussed previously, this reaction is performed by different enzymes in aerobic and anaerobic organisms, although many chlorophototrophs contain genes encoding more than one enzyme. Each enzyme performs a multi-step cyclization and oxidation which converts the C13 propionyl methyl ester group into the isocyclic E ring found in all
(B)Chls (Chen et al., 2017; Gough et al., 2000; Pinta et al., 2002) (Figure 2.2). In organisms inhabiting oxic environments the dominant cyclase enzyme is an oxygen- 40 dependent di-iron oxygenase known as AcsF (Pinta et al., 2002). The oxygen atom in the keto group of the E ring is derived from O2 (Walker et al., 1989). Plant and cyanobacterial AcsF enzymes require an additional subunit, Ycf54, for activity (Albus et al., 2012; Hollingshead et al., 2012), while AcsF paralogs found in purple chlorophototrophic members of the α-proteobacteria require BciE (Chen et al., 2017).
AcsF enzymes found in other chlorophototrophic purple bacteria do not require Ycf54 or
BciE. For example, AcsF from Rubrivivax gelatinosus encodes a functional cyclase requiring no additional subunit when produced in E. coli (Chen et al., 2018). Ycf54 and
BciE are not found together in the genome of any chlorophototroph studied to date (Chen et al., 2017). The acsF gene is not found in the strictly anaerobic GSB, Osc. trichoides, or
‘Ca. C. asiatica’ and but is present in ‘Ca. V. mediisalina’, Chloroflexus spp. and Cab. thermophilum (Bryant et al., 2012); however, ycf54 and bciE are absent from green bacteria containing acsF (Figure 2.1).
In anaerobic organisms the oxidative cyclase reaction is performed by an oxygen- independent and oxygen-sensitive enzyme, BchE. BchE is a radical-SAM enzyme, which contains an oxygen-labile [4Fe-4S] cluster and requires a cobalamin cofactor for catalysis
(Gough et al., 2000). As with HemN (CgdH) discussed above, this oxidation occurs under anoxic conditions via radical chemistry. The keto group on the isocyclic E ring derives its oxygen atom from H2O instead of O2 (Porra et al., 1996). BchE is a member of the BchE/P-methyltransferase family, which also includes C8 and C12 methyltransferases involved in the BChl c, d, e, and f biosynthetic pathway of some organisms (see below).
BchE is present in FAPs and Cab. thermophilum that also contain acsF (Figure 2.1). The 41 green FAP Cfl. aurantiacus primarily relies on BchE for the production of BChls; however, AcsF was detected in a chlorosome preparation under anoxic conditions, and unlike bchE, the expression of acsF does not change with O2 tension (Tang et al., 2009).
These observations led to the hypothesis that the Cfl. aurantiacus AcsF may have an alternative function; for example, this di-iron protein could play a role in electron transfer
or iron transport under anoxic conditions. The strictly anaerobic green bacteria, GSB,
Osc. trichoides, and ‘Ca. C. asiatica’ only contain the gene encoding the oxygen-
independent enzyme BchE and lack acsF (Liu and Bryant, 2012). As mentioned above,
the draft genome of ‘Ca. T. aerophilum’ did not contain bchM, bchE, or acsF (Figure
2.1). This raises the interesting prospect that this organism may form the isocyclic ring by a very different enzymatic mechanism than other organisms (Liu et al., 2012).
Furthermore, chlorophototrophic stramenopile algae also appear to lack acsF and bchE, providing additional evidence for the possible existence of a missing isocyclic ring cyclase enzyme, which may be shared with ‘Ca. T. aerophilum’ (Matsuo and Inagaki,
2018).
2.4.5 Reduction of C17=C18 Double Bond
The reduction of the C17=C18 double bond of the D ring results in the conversion of 8V-PChlide to 8V-Chlide a (or PChlide to Chlide a, see following section). In green bacteria the enzyme responsible for this reaction is the dark-operative (i.e., light- independent) PChlide reductase (DPOR) (Reinbothe et al., 2010) (Figure 2.1, 2.2).
DPOR is an ATP-dependent oxidoreductase made up of three subunits: BchN, BchB, and 42 BchL. The enzyme consists of a homodimer of BchL, which binds an intrasubunit [4Fe-
4S] cluster, and a heterotetramer of BchB and BchN which bind two [4Fe-4S] clusters; the electron donor for this reaction is reduced ferredoxin (Broecker et al., 2008a; Nomata et al., 2005). BchLBN is structurally similar to nitrogenase NifHDK (Broecker et al.,
2010; Muraki et al., 2010; Sarma et al., 2008). While BchLBN was initially described as a PChlide reductase, it was later demonstrated that 8V-PChlide can also be reduced by
DPOR (see below) (Broecker et al., 2008b).
2.4.6 Reduction of C8 Vinyl Group
Mutagenesis studies in purple bacteria described strains disrupted at the bchJ locus that were unable to grow phototrophically and that excreted a pigment that was later identified as 8V-PChlide (Bollivar et al., 1994b; Gomez Maqueo Chew and Bryant,
2007a, 2007b; Zsebo and Hearst, 1984). Although these mutants could still synthesize
BChl a carrying an ethyl group at C8, BchJ was nevertheless designated as an 8V- reductase (8VR) or at least a subunit of an 8VR. This result naturally led to the assumption that the substrate for 8VR was 8V-PChlide.
Mutants in the Arabidopsis thaliana AT5G18660 locus accumulate 8V-Chl a and
8V-Chl b (Nagata et al., 2005; Nakanishi et al., 2005). The recombinant protein encoded by this gene, which is unrelated to BchJ, is able to reduce the 8-vinyl group of 8V-Chlide a to an ethyl group to form Chlide a, confirming that this gene product is a functional
8VR. Subsequently, an ortholog of this gene from Cba. tepidum, bciA, was also shown to encode an 8VR. Heterologously produced BciA can reduce the 8-vinyl group of 8V- 43 PChlide to form PChlide a in the presence of NADPH (Gomez Maqueo Chew and
Bryant, 2007a).
The genomes of many cyanobacteria that synthesize Chl a do not contain orthologs of bciA. Mutants in open reading frame slr1923 of Synechocystis sp. PCC 6803
are unable to grow under high light and accumulate 8V-Chl a (Islam et al., 2008; Ito et
al., 2008). The ortholog of slr1923 from the GSB Chloroherpeton (Chp.) thalassium
(Ctha_1208) could complement the previously described ΔbciA mutant of Cba. tepidum
thereby demonstrating the activity of a second class of 8VR, BciB (Liu and Bryant,
2011a). In vitro characterization of this enzyme showed that BciB was able to convert
8V-PChlide to PChlide in the presence of either dithionite or ferredoxin, NADPH, and
ferredoxin:NADP+ oxidoreductase. BciB binds one FAD cofactor and two [4Fe-4S]
clusters, which are likely involved in the catalytic mechanism (Saunders et al., 2013).
All green bacteria have at least one gene encoding BciA or BciB, while some
have multiple paralogs for one or the other, or even genes for both enzymes (Figure 2.1)
(Liu and Bryant, 2011a). In organisms where the C82 position of BChl c, d, e, or f is
differentially methylated, the reduction of the C8 vinyl group is a prerequisite reaction
that must occur before the further modifications of the side chain by the BchQ
methyltransferase can occur. In mutants lacking 8VR activity, the C82 position cannot be
methylated, and this defect causes a decrease in the half bandwidth of the Qy absorbance
band compared to wild type. This defect also leads to lower total BChl c content in cells,
a blue-shift in the absorbance maximum of BChl c aggregates, and slower growth under
low irradiance conditions (Gomez Maqueo Chew and Bryant, 2007a; Liu and Bryant, 44 2011a). In low-light environments 8VR activity provides an important competitive
advantage for organisms that can methylate the C8 position, which may explain why
some organisms encode redundant 8VRs.
All tested higher plant enzymes demonstrate a preference for 8V-Chlide a as a substrate, as seen in the A. thaliana ortholog of BciA, and some are unable to reduce 8V-
PChlide entirely (Wang et al., 2013). Similarly, BciA from Rba. sphaeroides was shown to act on 8V-Chlide a, and could only reduce 8V-PChlide when the DPOR enzyme was absent (Canniffe et al., 2014). The same study also indicated that the BciB enzyme from
Synechocystis sp. PCC 6803 prefers 8V-Chlide a, but that enzyme is also able to reduce
8V-PChlide. Under pigment-accumulating conditions, only a small amount of reduced
PChlide could be detected in the WT (Canniffe et al., 2014). These studies indicate that the C8-vinyl reduction most likely occurs after the reduction of the C17=C18 bond, but that, as is common in pigment biosynthesis pathways, substrate flexibility is demonstrated by this enzyme, which could allow 8-vinyl reduction to occur under conditions that cause precursor pigments to accumulate.
Further analyses of bchJ mutants have been made. The deletion of this gene from the GSB Cba. tepidum and the purple bacterium Rba. sphaeroides did not perturb synthesis of (B)Chls, while disruption of bciA led to the accumulation of 8V-(B)Chl species (Canniffe et al., 2013; Gomez Maqueo Chew and Bryant, 2007a). There remains a lack of direct evidence for the activity of BchJ in this reaction. It has been suggested that BchJ may act as a substrate carrier/chaperone in the (B)Chl biosynthesis pathway in 45 anoxygenic chlorophototrophic bacteria, and thus play a role similar to that of Gun4 in plants and cyanobacteria (Sawicki and Willows, 2010).
A third enzyme with 8VR activity was hinted at in bciA mutants of Rba.
sphaeroides and Cba. tepidum, both of which lack bciB genes but still make normal BChl
a, (Canniffe et al., 2013; Mizoguchi et al., 2012). The finding that a bciA mutant of Cba.
tepidum can make 8-vinyl derivatives of Chl a and BChl c but reduced BChl a conflicted
with an earlier characterization of bciA mutants in Cba. tepidum that found 8-vinyl
derivatives of all three (B)Chls (Gomez Maqueo Chew and Bryant, 2007a; Mizoguchi et
al., 2012). Additionally, the red FAPs in the genus Roseiflexus have BChl a in their
photosynthetic apparatus, but lack orthologs of bciA or bciB in their genomes. These
observations strongly suggested that a third enzyme class capable of 8-vinyl group
reduction must exist (see discussion of Chlide reductase (COR) below).
2.5 Chlorophyllide a to Bacteriochlorophyllide a
As in purple bacteria, the conversion of Chlide a to BChlide a proceeds via the
action of three enzymes in all green bacteria, (Bollivar et al., 1994b; Harada et al., 2014;
Liu and Bryant, 2012; Suzuki et al., 1997). These enzymes catalyze the reduction of the
C7=C8 double bond (BchXYZ), hydration of the C3-vinyl group to form a hydroxylethyl group (BchF), and subsequent dehydrogenation of the C3-hydroxyethyl group to form an acetyl group (BchC) (Figure 2.3). Biosynthetic pathways commonly list the reactions in this order which was established based on substrate accumulation studies in various 46 studies of purple bacteria combined with limited in vitro characterization (Lange et al.,
2015). However, recent in vitro characterization of all three enzymes has shown that they exhibit expanded substrate reactivity; this suggests that multiple possible pathway orders may exist for the three terminal steps of BChlide a biosynthesis (Harada et al., 2014,
2015; Kiesel et al., 2015; Lange et al., 2015; Teramura et al., 2018).
2.5.1 Reduction of the C7=C8 Double Bond
The first committed step of BChlide a biosynthesis is the reduction of the C7=C8
double bond on ring B of the chlorin substrate producing a bacteriochlorin. The enzyme
responsible for this reaction is Chlide a oxidoreductase (COR), encoded by bchX, bchY,
and bchZ (Figure 2.1, 2.3). BchX, BchY, and BchZ are paralogs of the BchL, BchN, and
BchB subunits of DPOR and nitrogenase as described above. DPOR and COR are
structurally and functionally similar enzymes which catalyze the reduction of a double
bond on the macrocycle of a Chlide molecule in an ATP-dependent manner (Kiesel et al.,
2015; Nomata et al., 2006; Waetzlich et al., 2009). Importantly, duplication of the
DPOR-encoding genes leading to the evolution of COR, and thus the synthesis of
bacteriochlorins, permitted the use of near-infrared light for anoxygenic photosynthesis.
Initial in vitro characterization of COR showed that it could convert Chlide a into
3V-BChlide a (Nomata et al., 2006). Subsequently, substrate-specificity assays of
purified COR enzyme have shown that it is also active with other chlorins, including Zn-
containing analogs of 3-hydroxyethyl-Chlide a and 3-acetyl-Chlide a (Kiesel et al., 2015).
The specific activity of COR using the 3-hydroxy-ethyl-Chlide a analog as the substrate 47 was shown to be equal to that with Childe a as the substrate. In contrast, the specific
activity with the 3-acetyl-Chlide a analog was only 30% of that observed with Chlide a.
Based on these data, it seems equally likely that Chlide a or 3-hydroxyethyl-Chlide a may act as a substrate in vivo, while 3-acetyl-Chlide a may also be used but does not appear to be the preferred substrate for this enzyme.
Surprisingly, it was recently demonstrated that COR from the BChl a-producing purple chlorophototrophic bacterium, Rba. capsulatus, was also able to reduce the 8V group of Chlide a (Tsukatani et al., 2013b). This form of the enzyme (CORa) was able to catalyze the formation of 8-ethyl bacteriochlorin with both 8-vinyl and 8-ethyl Chlide
substrates. The COR enzymes from the BChl b- or BChl g-producing strains,
Blastochloris viridis or Heliobacterium modesticaldum, respectively (CORb) were not able to act on 8-ethyl Chlide a, but when 8V-Chlide a was provided as substrate, these enzymes formed a product carrying an ethylidene group at C8 (known as 3V-BChlide b or BChlide g) (Tsukatani et al., 2013b, 2013c). Replacement of the CORa-encoding genes of Rba. sphaeroides with the CORb orthologs from Blc. viridis led to the switch from the biosynthesis of BChl a to BChl b in this strain when the native 8VR, BciA, was absent(Canniffe and Hunter, 2014). These studies delineate a third class of 8VR enzyme, providing an explanation for the anomalous results mentioned previously. It is hypothesized that organisms using BChl a employ CORa to reduce the 8V group of any
Chlide molecules that have bypassed the conventional 8VR. This mechanism also accounts for absence of bciA or bciB in Roseiflexus spp. and clarifies why bciA mutants of Cba. tepidum and Rba. sphaeroides are able to synthesize BChl a (Canniffe et al., 48 2013; Mizoguchi et al., 2012). It is interesting to note that while COR enzymes may have
8V reductase activity, they require ATP for catalysis and are therefore more energetically
costly than the BciA and BciB enzymes. Considering the very large numbers of (B)Chls
produced by most green bacteria, this energetic penalty of the COR enzymes suggests
why they are probably not the preferred route of 8V reduction whenever BciA or BciB
are available. This penalty might also provide an explanation for why GSB have laterally
acquired an enzyme (BciA) that is quite similar to the enzymes found in higher plants
(Gomez Maqueo Chew and Bryant, 2007a). The 8VR activity of COR provides further
evidence that DPOR enzymes use 8V-PChlide instead of PChlide a as the substrate in
many circumstances. In fact, in the Roseiflexus spp. that lack bciA and bciB genes (see above), BChl a biosynthesis almost certainly proceeds from 8V-PChlide to 8V-Childe a catalyzed by DPOR, then from 8V-Chlide a to 3-vinyl-BChlide a, via a Chlide a intermediate, catalyzed by COR.
2.5.2 Hydration of the C3 vinyl group
BChl a has an acetyl group at the C3 position, while BChl c, d, e, and f have a hydroxyethyl group. The formation of each of these requires the action of a 3-vinyl
hydratase enzyme that can hydrate the vinyl group to form a hydroxyethyl moiety. In the
case of BChl a, this sidechain is further modified by a stereoselective, NAD+-dependent
dehydrogenase that results in the formation of the acetyl group.
The C3 hydratase in BChl a biosynthesis was first identified in purple bacteria, in
which the gene encoding this enzyme was denoted bchF (Burke et al., 1993). A single 49 homolog of bchF is found in green FAPs and Cab. thermophilum. For the most part,
green-colored GSB, such as Cba. tepidum, have two paralogs of bchF, denoted bchF and
bchV, and brown-colored GSB have a third paralog, denoted bchF3 (Figure 2.1) (Bryant and Liu, 2013). In green FAPs and Cab. thermophilum, BchF must perform C3 hydratase activity required for the synthesis of both BChl a and BChl c. Initial attempts to inactivate bchF in Cba. tepidum were unsuccessful, which led to the interpretation that bchF was required for BChl a biosynthesis making it an essential gene for GSB and that
BchV was specific to BChl c biosynthesis and did not function in BChl a biosynthesis
(Frigaard et al., 2003; Gomez Maqueo Chew et al., 2004; Liu and Bryant, 2012). Further discussion of the roles of BchF and BchV during the synthesis of BChlides c, d, e and f from Chlide a in green bacteria is found below.
2.5.3 3-Hydroxyethyl Dehydrogenase
Following formation of the C3 hydroxyethyl moiety, 3-hydroxyethyl BChlide a dehydrogenase, BchC, catalyzes the oxidation of the hydroxyethyl group at C3 to yield an acetyl group (Figure 2.1, 2.3). As with the bchF gene, bchC was first shown to encode the 3-hydroxyethyl dehydrogenase in purple bacteria (McGlynn and Hunter, 1993). In vitro experiments with heterologously expressed BchC from Cba. tepidum have examined the substrate specificity of this enzyme. The BchC dehydrogenase exhibited activity with a demetallated analog of 3-hydroxyethyl BChlide a and a Zn-containing analog of 3-hydroxyethyl Chlide a when NAD+, but not NADP+, was present as the
hydride acceptor (Lange et al., 2015). The specific activity with the Zn-containing analog of 3-hydroxyethyl Chlide a was about 20 times higher than that with the demetallated 50 analog of 3-hydroxyethyl BChlide a. The replacement of Mg by Zn is tolerated by many
enzymes in (B)Chl biosynthesis; however, the complete absence of a metal in the
macrocycle is often not tolerated (Harada et al., 2005; Kiesel et al., 2015; Teramura et al.,
2016a, 2016b, 2018, 2019). With this in mind it is difficult to say if BchC shows a
preference for 3-hydroxyethyl Chlide a or 3-hydroxyethyl BChlide a, because the analogs
used may not have allowed for a valid comparison. It is clear, however, that BchC has
significant activity with 3-hydroxyethyl Chlide a in addition to 3-hydroxyethyl BChlide
a. As a result of the expanded substrate reactivities of BchC, BchF, and COR, an
alternative reaction sequence has been proposed in which Chlide a is first acted upon by
BchF to form 3-hydroxyethyl Chlide a. This step is either followed by conversion of the
3-hydroxyethyl group to an acetyl group by BchC or by reduction of the C7=C8 double
bond by COR (BchXYZ) (Figure 2.3).
2.6 Chlorophyllide a to Bacteriochlorophyllide c, d, e, and f
2.6.1 Demethoxycarbonylation of the C132 methylcarboxyl group
The first committed step of the biosynthetic pathway leading from Chlide a to
BChls c, d, e, and f is the removal of the C132 methylcarboxyl group from Chlide a to
produce 3-vinyl-BChlide d (Figure 2.3). A similar reaction occurs in Chl degradation,
and two different enzymes are known to catalyze the removal of this group to produce
pheophorbide (Pheide) a in plants, algae, and some cyanobacteria. The enzyme
pheophorbidase catalyzes a methylesterase reaction, which is followed by spontaneous
decarboxylation, while an unknown enzyme in C. reinhardtii has been reported to have 51 demethyoxycarbonylase activity, which completes the removal of the C132
methylcarboxyl group in a single step (Hörtensteiner and Kräutler, 2011; Suzuki et al.,
2002). A candidate gene for this activity in green bacteria was identified by phylogenetic
profiling, and the corresponding deletion mutant in Cba. tepidum produced Chl a and
BChl a but no BChl c; the strain also accumulated (B)Pheides that retained the intact
C132 methylcarboxyl group (Liu and Bryant, 2011b). These data showed that the gene
was essential for removal of the C132 methylcarboxyl group, and it was designated bciC.
BciC is not homologous to known pheophorbidase enzymes or methylesterases, and so
the mechanism of the reaction remained unclear. Recently, the bciC gene from Cba. tepidum was heterologously expressed in E. coli, and the product catalyzed the removal of the C132 methylcarboxyl group of Chlide a and 8V-Chlide a, with a preference for
Chlide a. BciC could not catalyze this reaction when Pheide a, PChlide a, or 8V-PChlide a were tested as substrates (Teramura et al., 2016a). Further studies of BciC showed that the enzyme can use various Chlides modified at the C3 or C7 position, but at a reduced rate compared to Chlide a, and could also act on BChlide a and 3V-BChlide a at greatly reduced rates. However, no activity was observed with any porphyrins tested or any
Chlide with C132 S stereochemistry (Teramura et al., 2019). The substrate specificity data
supports the currently accepted branching point compound as Chlide a. BciC was not
inhibited by addition of methanol to the reaction, which suggests that it does not function
via the methylesterase mechanism seen in pheophorbidase enzymes but instead is a
demethoxycarbonylase. Homologs of the bciC gene are present in the genomes of all
green bacteria, and homologs are not found in any other organism (Figure 2.1). 52
2.6.2 Methylation at C8 and C12
Following removal of the methylcarboxyl group 3-vinyl-BChlide d may be
methylated at the C82 and C121 positions to form methylated-3-vinyl-BChlide d, which describes a mixture of methylated species (Figure 2.3). The methylations at these two positions are performed by independent methytransferases; each is a radical-SAM enzyme that derives its methyl-donating substrate from the cofactor S-adenosyl-L- methionine (Gomez Maqueo Chew et al., 2007; Huster and Smith, 1990; Kenner et al.,
1978). The C82-methyltransferase is encoded by bchQ while the C121-methytransferase is encoded by bchR; both are distant homologs of BchE, the oxygen-independent, oxidative ring cyclase discussed above (Gomez Maqueo Chew et al., 2007). The ethyl group at the C82 position may be methylated one to three times, sequentially producing n-
propyl, isobutyl, or neopentyl groups. The methyl side chain at C12 is only methylated
once to produce an ethyl moiety (Airs et al., 2001; Bobe et al., 1990; Borrego et al., 1999;
Glaeser et al., 2002; Gomez Maqueo Chew et al., 2007).
BchQ and BchR are found in Chlorobia, Cab. thermophilum, Osc. trichoides,
‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ but notably are absent in Chloroflexus spp.
(Figure 2.1) (Bryant et al., 2012). Brown-colored GSB have a second paralog of BchQ, denoted BchQ2. The bchQ2 gene is located just upstream from a conserved gene cluster
found in all brown-colored GSB. This cluster contains bchF3, a C31 hydratase, and bciD,
which is required for BChl e or f biosynthesis (see below and Chapter 3) (Thweatt et al.,
2017). While it is likely that bchQ2 is associated with BChl e biosynthesis, its exact 53 function remains unclear. Brown-colored GSB tend live in extremely low-light
environments which favor BChl species that are more highly methylated at the C82
position. It is possible bchQ2 is responsible for increased methylation activity in these
organisms. Because brown-colored GSB can live at lower light intensities than other
GSB, it is possible that increased methylation of BChl e by BchQ2 provides an evolutionary advantage by expanding the absorption cross-section of these organisms.
2.6.3 Hydration of the C31 Position
The BChls found in chlorosomes (BChl c, d, e or f) carry a hydroxyethyl group at
the C3 position, which is formed by the action of C31 hydratase on the C3 vinyl moiety of
a chlorin substrate. Green bacteria may have one, two, or three paralogs of the gene
encoding the C31 vinyl hydratase (Figure 2.1). In green FAPs and Cab. thermophilum, in
which only a single paralog occurs, BchF is responsible for the C31 hydration reaction in both BChl a and BChl c/d biosynthesis. However, in Chlorobia the situation is more complicated.
As mentioned above, an initial report led to the hypothesis that BchF was specific to BChl a biosynthesis and was thus an essential enzyme for members of the Chlorobia, and that BchV was specific to BChl c biosynthesis (Gomez Maqueo Chew et al., 2004).
However, individual mutants in the genes encoding these enzymes have now been successfully constructed in Cba. tepidum, and their roles have been characterized more thoroughly (Harada et al., 2015). The bchF mutant made significantly less BChl a, increased levels of 3-vinyl-BChl a, and less BChl c than the wild type. The BChl c 54 homologs in this mutant were primarily S-epimers, and when compared to the wild type, a higher proportion of these homologs were more highly methylated at the C82 position.
The bchV mutant contained less BChl c than the wild type, very little of which had the S-
configuration at C31. Ethyl and propyl groups at C8 were present, but unlike the wild
type, almost no homologs carried isobutyl sidechains. The cells contained more BChl a
and 3-vinyl-BChl c as a proportion of total (B)Chl. These results indicate that BchF is the
main C3 vinyl hydratase responsible for BChl a biosynthesis in Cba. tepidum, and it is
also important for the biosynthesis of BChl c homologs with ethyl or propyl groups at C8
in this organism. Additionally, these mutants show that the BchV hydratase must also
have some C3 hydratase activity in the BChl a biosynthetic pathway of GSB; that it
tolerates more highly methylated substrates; and that it contributes a greater ratio of S-
epimers while BchF contributes mostly R-epimers, which in turn provide the substrate for
the BchC dehydratase for biosynthesis of BChlide a.
Studies using lysates of E. coli expressing bchF and bchV from Cba. tepidum
showed that each enzyme has C3 hydratase activity when using a variety of substrates.
Hydroxyethyl groups could be formed with Chlide a, 8V-Chlide a, 3V-[8-ethyl,12-
methyl]-BChlide d, Zn-3V-[8-ethyl, 12-methyl]-BPheide d, Zn-3V-[8-ethyl/8-propyl/8- isobutyl,12-ethyl]-BPheide d, Zn-3V-[8-ethyl, 12-methyl]-BPheide c, and Zn-3V-[8- ethyl/8-propyl,12-ethyl]-BPheide c. Only BchF had activity with 3V-BChlide a, and neither enzyme could use Zn-3V-[8-isobutyl, 12-ethyl]-BPheide c (Harada et al., 2015;
Teramura et al., 2016b, 2018). The single BchF paralogs from Cfl. aurantiacus and Cab. thermophilum, and BchF, BchV and BchF3 from Cba. limnaeum were all able to use 55 Chlide a, Zn-3V-[8-ethyl, 12-methyl]-BPheide d and 3V-BChlide a as substrates
(Teramura et al., 2018).
On the basis of these results, it is likely that all three BchF homologs participate
in BChl a biosynthesis in Cba. limnaeum; however, their exact functions in vivo remain unclear (Teramura et al., 2018). The role of BchF3 in brown-colored GSB may be related to the fact that these organisms often live in very low-light environments in which higher degrees of methylation are favored. However, further studies are needed to determine if
BchF3 is important for the production of the C3 hydroxyethyl group in BChl e molecules carrying higher degrees of methylation.
It is interesting to note that brown-colored GSB, but not green-colored GSB, also possess an additional copy of bchQ, which could also be related to the prevalence of these neopentyl sidechains (see above). The apparent lack of in vitro hydratase activity
observed with BchV from Cba. tepidum when using 3V-BChlide a as the substrate suggests that either the activity was too low to be detected or that BchV contributes to
BChl a biosynthesis by hydration of chlorins instead of bacteriochlorins in vivo. The ability of C3 hydratase enzymes to use a variety of substrates, in combination with the expanded COR reactivity discussed above, suggests that the order of reactions within the
BChlide a pathway is currently ambiguous. However, these experiments were performed with crude lysates and did not yield specific activities. Based on in vitro hydratase activity with both 3V-BChlide c and 3V-BChlide d analogs, Teramura, Harada, &
Tamiaki (Teramura et al., 2016b, 2018) suggest a possible alternative reaction order in the biosynthetic pathway in which methylation of the C20 position comes before or 56 before and after C31 hydration in BChlide c homologs with C8 ethyl or C8 propyl groups.
However, this is in conflict with the crystal structure model of the C20 methyltransferase,
BchU, which modeled BChlide d in the active site and indicated hydrogen bonding
between the C31 hydroxyl group and the enzyme was likely to be important in substrate
binding and specificity (Wada et al., 2006)(see below). Further investigation of the
substrate specificities of C3 hydratases is needed to elucidate the preferred substrates for
each enzyme.
2.6.4 Methylation of the C20 methine bridge
In green bacteria that contain BChl c or BChl e, a C20 methyltransferase, BchU,
is required to methylate the C20 methine bridge of BChlide d to form BChlide c. As a
result BchU is encoded in the genomes of all green bacteria used in this review with the
exception of ‘Ca. T aerophilum’, an organism that produces BChl d and not BChl c or e
(Figure 2.1). It should also be noted that in Cba. parvum strains which produce BChl d there is a mutation in the gene encoding BchU. (Liu.et al., 2012). BchU is a SAM- dependent methyltraseferase, but it is not a radical-SAM methyltransferase like BchR and
BchQ, and it is not a homolog of the enzymes discussed above. Crystal structures of
BchU show that the enzyme is a homodimer and that the co-product, S- adenosylhomocysteine, binds near the C-terminal active site (Wada et al., 2006). The bchU gene was first identified in Cba. tepidum. When this gene was inactivated, the mutant produced BChl d instead of c (Maresca et al., 2004). Growth rate and competition studies showed that in high-light conditions Cba. tepidum containing BChl c or d grew at 57 the same rate, but in low-light conditions Cba. tepidum containing BChl c had a growth advantage; this agrees with other comparisons of BChl c or d-containing GSB. (Bobe et al., 1990; Broch-due and Ormerod, 1978; Maresca et al., 2004; Saga et al., 2003). BchU from Cba. tepidum produced by heterologous expression in E. coli could catalyze the conversion of Zn-BChlide d to Zn-BChlide c (Harada et al., 2005).
Mutants of bchU in the model brown-colored GSB, Cba. limnaeum, resulted in strains that produce BChl f, an unnatural BChl (Tamiaki et al., 2011; Tsukatani et al.,
2013a; Vogl et al., 2012). BChl f is able to form supramolecular structures and assemble functional chlorosomes (Niedzwiedzki et al., 2014). The bchU mutants of Cba. limnaeum containing BChl f exhibited a significantly slower growth rate under low-light conditions than wild-type strains containing BChl e (Vogl et al., 2012). A comparison of the energy transfer characteristics of chlorosomes containing BChl e or f by time-resolved fluorescence spectroscopy showed that energy transfer from the BChl f aggregates to the baseplate BChl a-binding CsmA complexes was much less efficient than from BChl e aggregates (Orf et al., 2013). Based on these data BChl f containing organisms would require anoxic, high-light conditions. Such environments are rare and may be more favorable for other phototrophs, which could explain why BChl f has not been found in nature (Orf et al., 2013). Furthermore, the triplet excitonic state of BChl f is energetically more favorable for the production of singlet oxygen than BChl c, d, or e (Hartzler et al.,
2014).
58 2.6.5 Formation of the C7 Formyl Group of BChlide e
Brown-colored GSB require an enzyme known as BChlide c C71 hydroxylase
which converts the C7 methyl group of BChlide c or d to the formyl group of BChlide e
or f, respectively, the enzyme is known as BciD (Thweatt et al., 2017) (see Chapter 3).
BciD is found exclusively in all brown-colored GSB (Figure 2.1). Similar to the
biosynthesis of Chl b, which also carries a formyl group at C7, this conversion is believed
to occur via two sequential hydroxylation reactions followed by spontaneous
dehydration. The formyl group of Chl b is formed via an oxygen-dependent enzyme
known as Chlide a oxygenase (CAO) (Tanaka et al., 1998). However, GSB are obligate
anaerobes and do not contain homologs of CAO. Brown-colored GSB use a radical-SAM enzyme, BciD, which contains a single [4Fe-4S] cluster and catalyzes the hydroxylation reactions via radical chemistry (Thweatt et al., 2017) (see Chapter 3). Characterization of BciD from Cba. limnaeum heterologously produced in E. coli is described in detail in
Chapter 3 and the main findings briefly summarized here. The bciD gene was first identified as a candidate gene related to BChl e and f biosynthesis via bioinformatic comparisons of the genomes of green-colored and brown-colored GSB which identified a gene cluster unique to brown-colored GSB. This cluster encodes cruB, a gene required for isorenieratene biosynthesis (Maresca et al., 2008); bciD, which encodes the C71
hydroxylase (Thweatt et al., 2017); bchF3, which encodes a C31 hydratase; and bchQ2,
which encodes a paralog of the C8 methyltransferase. Mutation of bciD in Cba. limnaeum resulted in cells producing BChl c instead of BChl e, while a double mutant of bchU and bciD resulted in cells producing BChl d (Harada et al., 2013; Mizoguchi et al.,
2015; Thweatt et al., 2017). 59
2.7 The final steps in (B)Chl biosynthesis
The final steps in the biosynthetic pathways of (B)Chls involve the formation of
the esterifying alcohol at the C173 position. After the formation of the (B)Chlide intermediate two types of enzymes are involve in forming the final product. A specific
Chl or BChl synthase esterifies an alcohol moiety to the C173 position and in the case of
phytyl or Δ2,6-phytadienol, geranylgeranyl reductase, ChlP/BchP, reduces the
geranylgeranyl precursor to the appropriate moiety (Figure 2.4). It remains uncertain whether reduction happens before or after esterification (or both), as in vitro assays have demonstrated that esterification can occur with diverse substrate alcohols at various stages of saturation (see below). However, the occurrence of BChl a carrying a geranylgeraniol moiety and BPhe a esterified with phytol in the purple bacterium
Rhodospirillum rubrum, and the subsequent identification of a mutation in its bchP gene
resulting in the production of BchP only able to reduce alcohols attached to demetallated
pigments, indicates that esterification occurs first in vivo in this organism (Addlesee and
Hunter, 2002). Therefore, the following sections are ordered in this manner, although
observations contradicting this somewhat will be mentioned.
2.7.1 Esterification of (B)Chlide
Green bacteria contain multiple (B)Chl synthase homologs known as Chl a
synthase (ChlG), BChl a synthase (BchG), and BChl c synthase (BchK). Green FAPs 60 have two types of BChl synthases, BchG and BchK, that esterify BChl a and BChl c or d,
respectively. Chlorobia and Cab. thermophilum additionally have a Chl a synthase,
ChlG, giving them three types of (B)Chl synthases. Some GSB have multiple copies of the bchK gene, which encodes BChl c, d, e, or f synthase (Figure 2.1) (Frigaard et al.,
2002; Thweatt et al., 2017). Phylogenetic analysis of these three types of synthases shows that the Chl/BChl synthase clade branches from other prenyl-transferases, and that BchG arose by a duplication from within an existing ChlG/BchK clade. This implies that the chlorin synthases BchK and ChlG existed prior to the branching of BchG in evolutionary history (see Chapter 4).
In Chl a-containing organisms, ChlG esterifies Chlide a at the C173 carboxylate
moiety to form Chl a by using an isoprenoid alcohol-diphosphate substrate. The chlG
genes of plants and cyanobacteria were first identified by homology to the bchG genes of
purple bacteria and were later used to identify chlG in green bacterial genomes (Bollivar
et al., 1994c; Eisen et al., 2002; Frigaard et al., 2003; Gaubier et al., 1995; Kaneko et al.,
1995). In vitro characterization of ChlG enzymes has shown that ChlG can esterify the
C173 position of Chlide a but not BChlide a, showing that ChlG is specific for the Chlide
a substrate (Oster et al., 1997). In experiments comparing different alcohol-diphosphate substrates, ChlG from cyanobacteria and plants, which produce Chl aP, can use both
geranylgeranyl-PP or phytyl-PP; some enzymes show a preference for geranylgeranyl-PP
and others show a preference for phytyl-PP (Oster et al., 1997; Rüdiger et al., 1980). No
in vitro study of ChlG from green bacteria has yet been reported; however, some hints at substrate specificity are available from bchP and bciC mutants in Cba. tepidum. The 61
bchP mutant was shown to produce Chl aGG, while the bciC mutant showed over-
production of Chl a derivatives resulting in mostly Chl aP in addition to the normal Chl aPD (Harada et al., 2008; Liu and Bryant, 2011b). These in vivo data indicate that ChlG in
Cba. tepidum can use geranylgeranyl-PP as a substrate. However, it is unclear if Chl aP
and Chl aPD are the result of reductions that occur after esterification with
geranylgeranyl-PP or esterification with reduced diphosphate counterparts.
BChl a synthase, BchG, was initially discovered and characterized in purple bacteria (Addlesee et al., 2000; Bollivar et al., 1994c; Oster et al., 1997). The activities of
homologs from green bacteria were first confirmed by expression of two BchG homologs
from Cfl. aurantiacus (Schoch et al., 1999). The first homolog (BchG) had synthase
activity with BChlide a derivatives but not Chlide a or BChlide c, d, or e derivatives; the
second, now known as BchK, exhibited activity with BChlide c, d, and e derivatives. The
bchG gene from Cba. tepidum was identified via complementation of BChl a synthase
activity in the purple bacterium Rba. capsulatus (Xiong et al., 2000). The bchG gene was
subsequently identified in the sequenced genomes of other green bacteria as well. A
recent study characterized the in vitro activity of heterologously produced BchG from
Cba. tepidum, which also showed specificity for BChlide a and that the enzyme is unable
to esterify Chlide a or BChlide c (Saga et al., 2015b). As observed with ChlG enzymes,
BchG enzymes are able to esterify BChlide a with a variety of diphosphate alcohols.
BchG from Cfl. aurantiacus could use the pyrophosphate derivatives of geranylgeraniol,
phytol, farnesol, and geraniol with a strong preference for geranylgeranyl-PP, while
BchG of Cba. tepidum was shown to use geranylgeranyl-PP or farnesyl-PP. Unlike the 62 studies with Cfl. aurantiacus, these experiments did not test phytyl-PP as a substrate or
show that BchG from Cba. tepidum preferentially used a particular esterifying alcohol.
The data confirm that BchG from Cba. tepidum is a BChl synthase specific for BChl a, but that it has broad specificity for the esterifying alcohol in vitro.
Surprisingly, BChl a derivatives in vivo almost exclusively have phytyl tails in
Cba. tepidum, except in the case of a bchP mutant that produced BChl aGG instead
(Gomez Maqueo Chew et al., 2008; Harada et al., 2008; Saga et al., 2015b). This
observation suggests that substrate channeling or sequestration may occur in vivo, or that
the pools of other alcohol-pyrophosphate derivatives must be too small to support rapid
esterification of BChlide a. However, the observation that BChl aP is still exclusively
produced by Cba. tepidum in the presence of exogenous alcohols, but BChl c contains
various exogenously added alcohols, suggests that the concentration of alcohol- pyrophosphates alone does not account for the selective nature of BChl aP production
(Larsen et al., 1995; Steensgaard et al., 1996).
BchK, BChl c (d/e/f) synthase, is the third type of (B)Chl synthase that occurs in
green bacteria. As noted above BChl c synthase activity was demonstrated with a BchG
paralog from Cfl. aurantiacus; however, the gene and gene product were not named at the
time (Schoch et al., 1999). Null mutants of the bchK gene in Cba. tepidum were unable to
synthesize BChl c but could still synthesize BChl a and Chl a and could grow
photoautotrophically (Frigaard et al., 2002). As a result, it was concluded that bchK
encodes BChl c synthase. It is important to note that in bchU mutants of Cba. tepidum,
BchK must function as a BChl d synthase. All green bacteria have at least one copy of 63 bchK in their genomes, while some have multiple copies. In this work, the BchK
homologs of Chlorobi have been designated as BchK1 and BchK2, while those of other
organisms are simply designated BchK (Figure 2.1). The phylogentic relationships
between the (B)Chl synthases of green bacteria and the implications of these relationships
will be discussed further in Chapter 3 and Chapter 4. Further studies are needed to
determine the substrate specificity of GSB BchK enzymes with relationship to the
BChlide substrate. Several in vivo studies feeding a wide variety of exogenous alcohols to
Cba. tepidum suggest that the substrate specificity of BchK for the esterifying alcohol is
extremely broad and is primarily driven by availability/concentration (Larsen et al., 1995;
Nishimori et al., 2011; Saga et al., 2014, 2015a, 2016; Steensgaard et al., 1996).
2.7.2 Reduction of the alcohol moiety
ChlP/BchP produces Δ2,6 phytadienol, or phytol moieties by sequential
reductions of geranylgeraniol for biosynthesis of Chl aPD and BChl aP, respectively. As
noted above, these reductions may occur either on the diphosphate derivative of geranylgeraniol or after the alcohol is esterified to the C173 position of (B)Chlide a
(Gomez Maqueo Chew et al., 2008; Harada et al., 2008; Liu and Bryant, 2012). Analysis
of the structures of intermediates in purple bacteria with a BchP variant that produces
phytyl, tetrahydrogeranylgeranyl, dihydrogeranylgeranyl, and geranylgeranyl BChl a, led
to the proposal of a pathway for subsequent reductions of the tail moiety. In this scheme
the double bonds of geranylgeranyl are sequentially reduced at C6–C7, C10–C11, and
C14–C15 (Harada et al., 2008; Mizoguchi et al., 2006). This pathway easily explains the 64 production of BChl aP; however, it does not account for the production of Chl aPD, which contains Δ2,6 phytadienol, whereas Δ2,14 phytadienol was identified. It is unclear if the
BchP enzymes of Chl a-containing green bacteria follow a different pathway for sequential reduction of the double bonds in geranylgeraniol, or if the Δ2,6 phytadienol originates from a phytyl intermediate that is re-oxidized at the C6–C7 bond (Harada et al., 2008).
Green FAPs and Cab. thermophilum have single copies of bchP, which encodes geranylgeranyl reductase. However, multiple copies of bchP are present in genomes in several species of GSB (Figure 2.1) (Canniffe et al., 2018). In Cba. tepidum, which has two bchP paralogs, null mutants showed that one gene, denoted bchP, was required for geranylgeranyl reductase activity in vivo, while the second gene, originally denoted bchO, had no phenotype affecting (B)Chl biosynthesis (Gomez Maqueo Chew et al.,
2008; Harada et al., 2008). More recent studies have shown that the bchO mutant of
Cba. tepidum cannot synthesize 1′,2′-dihydrochlorobactene (Canniffe et al., 2018).
Moreover, bchO from Cba. tepidum and two paralogs of bchO from Cba. limnaeum are able to catalyze reduction of 1,2 double bonds in carotenoid substrates produced in Rba. sphaeroides. Thus, these former bchO genes have been renamed cruI to reflect their demonstrated role in carotenoid biosynthesis. None of the other bchO paralogs tested in this way exhibited CruI activity or BchP activity when assayed in Rba. sphaeroides.
Thus, the roles of these other bchP paralogs remain unknown at this time (Canniffe et al.,
2018).
65 2.8 Concluding Remarks
Over the past two decades, the biosynthetic pathways of Chls and BChls of green bacteria have been nearly fully elucidated. This was largely due to the availability of complete genomes of many organisms, and similarities to the pathways for the synthesis
of these molecules in purple bacteria, cyanobacteria, and plants. Some remaining
questions in these biosynthetic pathways are also unresolved in other chlorophototrophs
while others relate largely to the unique properties of green bacteria. For example, the
insertion of Zn++ in the biosynthesis of Zn-BChl a′ in Cab. thermophilum and in
acidophilic purple bacteria has not been explained. Another major area of uncertainty
concerns the regulation of these pathways, particularly how green bacteria make and
channel the appropriate amounts of different (B)Chl precursors, all of which branch from
one biosynthetic pathway, to produce products required in vastly different amounts.
Mechanistic issues surrounding the different oxidative isocylic ring cyclases will
certainly be an interesting subject for future studies. Many of the answers likely lie in the
continued biochemical characterization of enzymes in vitro combined with carefully
planned genetic manipulations in vivo.
66 References Abicht, H. K., Martinez, J., Layer, G., Jahn, D., and Solioz, M. (2012). Lactococcus lactis HemW (HemN) is a haem-binding protein with a putative role in haem trafficking. Biochem. J. 442, 335–343. doi:10.1042/BJ20111618. Addlesee, H. A., Fiedor, L., and Hunter, C.N. (2000). Physical mapping of bchG, orf427, and orf177 in the photosynthesis gene cluster of Rhodobacter sphaeroides: Functional assignment of the bacteriochlorophyll synthetase gene. J. Bacteriol. 182, 3171–3182. doi:10.1128/JB.182.11.3175-3182.2000. Addlesee, H. A., and Hunter, C. N. (2002). Rhodospirillum rubrum possesses a variant of the bchP gene, encoding geranylgeranyl-bacteriopheophytin reductase. J. Bacteriol. 184, 1578–1586. doi:10.1128/JB.184.6.1578-1586.2002. Airs, R. L., Borrego, C. M., Garcia-Gil, J., and Keely, B. J. (2001). Identification of the bacteriochlorophyll homologues of Chlorobium phaeobacteroides strain UdG6053 grown at low light intensity. Photosynth. Res. 70, 221–230. doi:10.1023/A:1015146304441. Albus, C. A., Salinas, A., Czarnecki, O., Kahlau, S., Rothbart, M., Thiele, W., et al. (2012). LCAA, a novel factor required for magnesium protoporphyrin monomethylester cyclase accumulation and feedback control of aminolevulinic acid biosynthesis in tobacco. Plant Physiol. 160, 1923–1936. doi:10.1104/pp.112.206045. Bobe, F. W., Pfennig, N., Swanson, K. L., and Smith, K. M. (1990). Red shift of absorption maxima in Chlorobineae through enzymic methylation of their antenna bacteriochlorophylls. Biochemistry 29, 4340–4348. doi:10.1021/bi00470a012. Bollivar, D. W., Jiang, Z. Y., Bauer, C. E., and Beale, S. I. (1994a). Heterologous expression of the bchM gene product from Rhodobacter capsulatus and demonstration that it encodes S-adenosyl-L-methionine: Mg-protoporphyrin IX methyltransferase. J. Bacteriol. 176, 5290–5296. doi:10.1128/jb.176.17.5290- 5296.1994. Bollivar, D. W., Suzuki, J. Y., Beatty, J. T., Dobrowolski, J. M., and Bauer, C. E. (1994b). Directed mutational analysis of bacteriochlorophyll a biosynthesis in Rhodobacter capsulatus. J. Mol. Biol. 237, 622–640. doi:10.1006/jmbi.1994.1260. Bollivar, D. W., Wang, S., Allen, J. P., and Bauer, C. E. (1994c). Molecular genetic analysis of terminal steps in bacteriochlorophyll a biosynthesis: Characterization of a Rhodobacter capsulatus strain that synthesizes geranylgeraniol-esterified bacteriochlorophyll a. Biochemistry 33, 12763–12768. doi:10.1021/bi00209a006. Borrego, C. M., Gerola, P. D., Miller, M., and Cox, R. P. (1999). Light intensity effects on pigment composition and organization in the green sulfur bacterium Chlorobium tepidum. Photosynth. Res. 59, 159–166. doi:10.1023/A:1006161302838. Boynton, T. O., Daugherty, L. E., Dailey, T. A., and Dailey, H. A. (2009). Identification of Escherichia coli HemG as a novel, menadione-dependent flavodoxin with protoporphyrinogen oxidase activity. Biochemistry 48, 6705–6711. doi:10.1021/bi900850y. Boynton, T. O., Gerdes, S., Craven, S. H., Neidle, E. L., Phillips, J. D., and Dailey, H. A. (2011). Discovery of a gene involved in a third bacterial protoporphyrinogen oxidase 67 activity through comparative genomic analysis and functional complementation. Appl. Environ. Microbiol. 77, 4795–4801. doi:10.1128/AEM.00171-11. Broch-Due, M., and Ormerod, J. G. (1978). Isolation of a BChl c mutant from Chlorobium with BChl dby cultivation at low light intensity. FEMS Microbiol. Lett. 3, 305–308. doi:10.1111/j.1574-6968.1978.tb01953.x. Broecker, M. J., Virus, S., Ganskow, S., Heathcote, P., Heinz, D. W., Schubert, W.D., et al. (2008a). ATP-driven reduction by dark-operative protochlorophyllide oxidoreductase from Chlorobium tepidum mechanistically resembles nitrogenase catalysis. J. Biol. Chem. 283, 10559–10567. doi:10.1074/jbc.M708010200. Broecker, M. J., Waetzlich, D., Saggu, M., Lendzian, F., Moser, J., and Jahn, D. (2010). Biosynthesis of (Bacterio) chlorophylls ATP-dependent transient subunit interaction and electron transfer of dark operative protochlorophyllide oxidoreductase. J. Biol. Chem. 285, 8268–8277. doi:10.1074/jbc.M109.087874. Broecker, M. J., Waetzlich, D., Uliczka, F., Virus, S., Saggu, M., Lendzian, F., et al. (2008b). Substrate recognition of nitrogenase-like dark operative protochlorophyllide oxidoreductase from Prochlorococcus marinus. J. Biol. Chem. 283, 29873–29881. doi:10.1074/jbc.M805206200. Bryant, D. A., and Liu, Z. (2013). Green bacteria: insights into green bacterial evolution through genomic analyses. Elsevier doi:10.1016/B978-0-12-397923-0.00004-7. Bryant, D. A., Liu, Z., Li, T., Zhao, F., Garcia Costas, A. M., Klatt, C. G., et al. (2012). “Comparative and functional genomics of anoxygenic green bacteria from the taxa Chlorobi, Chloroflexi, and Acidobacteria,” in Advances in Photosynthesis and Respiration, Vol. 35 Functional Genomics and Evolution of Photosynthetic Systems, eds. R. Burnap and W. Vermaas (Dordrecht: Springer Netherlands), 47–102. doi:10.1007/978-94-007-1533-2_3. Burke, D. H., Alberti, M., and Hearst, J. E. (1993). bchFNBH bacteriochlorophyll synthesis genes of Rhodobacter capsulatus and identification of the third subunit of light-independent protochlorophyllide reductase in bacteria and plants. J. Bacteriol. 175, 2414–2422. doi:10.1128/jb.175.8.2414-2422.1993. Canniffe, D. P., Chidgey, J. W., and Hunter, C. N. (2014). Elucidation of the preferred routes of C8-vinyl reduction in chlorophyll and bacteriochlorophyll biosynthesis. Biochem. J. 462, 433–440. doi:10.1042/BJ20140163. Canniffe, D. P., Jackson, P. J., Hollingshead, S., Dickman, M. J., and Hunter, C. N. (2013). Identification of an 8-vinyl reductase involved in bacteriochlorophyll biosynthesis in Rhodobacter sphaeroides and evidence for the existence of a third distinct class of the enzyme. Biochem. J. 450, 397–405. doi:10.1042/BJ20121723. Canniffe, D. P., Thweatt, J. L., Chew, A. G. M., Hunter, C. N., and Bryant, D. A. (2018). A paralog of a bacteriochlorophyll biosynthesis enzyme catalyzes the formation of 1,2-dihydrocarotenoids in green sulfur bacteria. J. Biol. Chem. 293, 15233–15242. doi:10.1074/jbc.RA118.004672. Canniffe, D. P., and Hunter, C.N. (2014). Engineered biosynthesis of bacteriochlorophyll b in Rhodobacter sphaeroides. Biochim. Biophys. Acta 1837, 1611–1616. doi:10.1016/j.bbabio.2014.07.011. 68 Chen, G. E., Canniffe, D. P., and Hunter, C. N. (2017). Three classes of oxygen- dependent cyclase involved in chlorophyll and bacteriochlorophyll biosynthesis. Proc. Natl. Acad. Sci. 114, 6280–6285. doi:10.1073/pnas.1701687114. Chen, G. E., Canniffe, D. P., Barnett, S. F. H., Hollingshead, S., Brindley, A. A., Vasilev, C., et al. (2018). Complete enzyme set for chlorophyll biosynthesis in Escherichia coli. Sci. Adv. 4. doi:10.1126/sciadv.aaq1407. Dailey, H. A., Dailey, T. A., Gerdes, S., Jahn, D., Jahn, M., O’Brian, M. R., et al. (2017). Prokaryotic heme biosynthesis: multiple pathways to a common essential product. Microbiol. Mol. Biol. Rev. 81, e00048-16. doi:10.1128/MMBR.00048-16. Dailey, T. A., and Dailey, H. A. (1996). Human protoporphyrinogen oxidase: Expression, purification, and characterization of the cloned enzyme. Protein Sci. 5, 98–105. Eisen, J. A, Nelson, K. E., Paulsen, I. T., Heidelberg, J. F., Wu, M., Dodson, R. J., et al. (2002). The complete genome sequence of Chlorobium tepidum TLS, a photosynthetic, anaerobic, green-sulfur bacterium. Proc. Natl. Acad. Sci. USA 99, 9509–9514. doi:10.1073/pnas.132181499. Fan, T., Grimm, B., and Layer, G. (2019). Porphyrin and heme synthesis. In: Advances in Botanical Research. Vol. 91 Metabolism, Structure and Function of Plant Tetrapyrroles: Control Mechanisms of Chlorophyll Biosynthesis and Analysis of Chlorophyll-Binding Proteins, Bernhard Grimm, Ed. (Academic Press, London, UK), pp. 89-131. doi:10.1016/bs.abr.2019.06.001. Fodje, M. N., Hansson, A., Hansson, M., Olsen, J. G., Gough, S., Willows, R. D., et al. (2001). Interplay between an AAA module and an integrin I domain may regulate the function of magnesium chelatase. J. Mol. Biol. 311, 111–122. doi:10.1006/jmbi.2001.4834. Frigaard, N. U., Voigt, G. D., and Bryant, D. A. (2002). Chlorobium tepidum mutant lacking bacteriochlorophyll c made by inactivation of the bchK gene, encoding bacteriochlorophyll c synthase. J. Bacteriol. 184, 3368–3376. doi:10.1128/JB.184.12.3368. Frigaard, N. U., Gomez Maqueo Chew, A., Li, H., Maresca, J. A., and Bryant, D. A. (2003). Chlorobium tepidum: Insights into the structure, physiology, and metabolism of a green sulfur bacterium derived from the complete genome sequence. Photosynth. Res. 78, 93–117. doi:10.1023/B:PRES.0000004310.96189.b4. Garcia Costas, A. M., Liu, Z., Tomsho, L. P., Schuster, S. C., Ward, D. M., and Bryant, D. A. (2012a). Complete genome of Candidatus Chloracidobacterium thermophilum, a chlorophyll-based photoheterotroph belonging to the phylum Acidobacteria. Environ. Microbiol. 14, 177–190. doi:10.1111/j.1462-2920.2011.02592.x. Garcia Costas, A. M., Tsukatani, Y., Rijpstra, W. I. C., Schouten, S., Welander, P. V., Summons, R. E., et al. (2012b). Identification of the bacteriochlorophylls, carotenoids, quinones, lipids, and hopanoids of “Candidatus Chloracidobacterium thermophilum.” J. Bacteriol. 194, 1158–1168. doi:10.1128/JB.06421-11. Gaubier, P., Wu, H. J., Laudié, M., Delseny, M., and Grellet, F. (1995). A chlorophyll synthetase gene from Arabidopsis thaliana. Mol. Gen. Genet. 249, 58–64. doi:10.1007/BF00290236. Gibson, L. C. D., and Hunter, C. N. (1994). The bacteriochlorophyll biosynthesis gene, bchM, of Rhodobacter sphaeroides encodes S-adenosyl-L-methionine - Mg 69 protoporphyrin-IX methyltransferase. FEBS Lett. 352, 127–130. doi:10.1016/0014- 5793(94)00934-1. Glaeser, J., Bañeras, L., Rütters, H., and Overmann, J. (2002). Novel bacteriochlorophyll e structures and species-specific variability of pigment composition in green sulfur bacteria. Arch. Microbiol. 177, 475–485. doi:10.1007/s00203-002-0416-4. Gomez Maqueo Chew, A., and Bryant, D. A. (2007a). Characterization of a plant-like protochlorophyllide a divinyl reductase in green sulfur bacteria. J. Biol. Chem. 282, 2967–2975. doi:10.1074/jbc.M609730200. Gomez Maqueo Chew, A., and Bryant, D. A. (2007b). Chlorophyll biosynthesis in bacteria: the origins of structural and functional diversity. Annu. Rev. Microbiol. 61, 113–129. doi:10.1146/annurev.micro.61.080706.093242. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2009). Mutational analysis of three bchH paralogs in (bacterio-)chlorophyll biosynthesis in Chlorobaculum tepidum. Photosynth. Res. 101, 21–34. doi:10.1007/s11120-009-9460-0. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2008). Identification of the bchP gene, encoding geranylgeranyl reductase in Chlorobaculum tepidum. J. Bacteriol. 190, 747–749. doi:10.1128/JB.01430-07. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2007). Bacteriochlorophyllide c C-82 and C-121 methyltransferases are essential for adaptation to low light in Chlorobaculum tepidumJ. Bacteriol. 189, 6176–6184. doi:10.1128/JB.00519-07. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2004). “Identification of BchV, a C-31 hydratase specific for hypermethylated bacteriochlorophyll c in Chlorobium tepidum,” in Photosynthesis: Fundamental Aspects to Global Perspectives, eds. A. van der Est and D. Bruce (Lawrence, KS: Allen Press), 875– 877. Gough, S. P., Petersen, B. O., and Duus, J. Ø. (2000). Anaerobic chlorophyll isocyclic ring formation in Rhodobacter capsulatus requires a cobalamin cofactor. Proc Natl Acad Sci U S A 97, 6980–6913. doi:10.1073/pnas.97.12.6908. Grouzdev, D. S., Kuznetsov, B. B., Keppen, O. I., Krasil’nikova, E. N., Lebedeva, N. V., and Ivanovsky, R. N. (2015). Reconstruction of bacteriochlorophyll biosynthesis pathways in the filamentous anoxygenic phototrophic bacterium Oscillochloris trichoides DG-6 and evolution of anoxygenic phototrophs of the order Chloroflexales. Microbiol. (United Kingdom) 161, 120–130. doi:10.1099/mic.0.082313-0. Harada, J., Miyago, S., Mizoguchi, T., Azai, C., Inoue, K., Tamiaki, H., et al. (2008). Accumulation of chlorophyllous pigments esterified with the geranylgeranyl group and photosynthetic competence in the CT2256-deleted mutant of the green sulfur bacterium Chlorobium tepidum. Photochem. Photobiol. Sci. 7, 1179–1187. doi:10.1039/b802435a. Harada, J., Mizoguchi, T., Satoh, S., Tsukatani, Y., Yokono, M., Noguchi, M., et al. (2013). Specific gene bciD for C7-methyl oxidation in bacteriochlorophyll e biosynthesis of brown-colored green sulfur bacteria. PLoS One 8, e60026. doi:10.1371/journal.pone.0060026. 70 Harada, J., Mizoguchi, T., Tsukatani, Y., Yokono, M., Tanaka, A., and Tamiaki, H. (2014). Chlorophyllide a oxidoreductase works as one of the divinyl reductases specifically involved in bacteriochlorophyll a biosynthesis. J. Biol. Chem. 289, 12716–12726. doi:10.1074/jbc.M113.546739. Harada, J., Saga, Y., Yaeda, Y., Oh-Oka, H., and Tamiaki, H. (2005). In vitro activity of C-20 methyltransferase, BchU, involved in bacteriochlorophyll c biosynthetic pathway in green sulfur bacteria. FEBS Lett. 579, 1983–1987. doi:10.1016/j.febslet.2005.01.087. Harada, J., Teramura, M., Mizoguchi, T., Tsukatani, Y., Yamamoto, K., and Tamiaki, H. (2015). Stereochemical conversion of C3-vinyl group to 1-hydroxyethyl group in bacteriochlorophyll c by the hydratases BchF and BchV: Adaptation of green sulfur bacteria to limited-light environments. Mol. Microbiol. 98, 1184–1198. doi:10.1111/mmi.13208. Hartzler, D. A., Niedzwiedzki, D. M., Bryant, D. A., Blankenship, R. E., Pushkar, Y., and Savikhin, S. (2014). Triplet excited state energies and phosphorescence spectra of (Bacterio)chlorophylls. J. Phys. Chem. B 118, 7221–7232. doi:10.1021/jp500539w. Haskamp, V., Karrie, S., Mingers, T., Barthels, S., Alberge, F., Magalon, A., et al. (2018). The radical SAM protein HemW is a heme chaperone. J. Biol. Chem. 293, 2558–2572. doi:10.1074/jbc.RA117.000229. Hinchigeri, S. B., Hundle, B., and Richards, W. R. (1997). Demonstration that the bchH protein of Rhodobacter capsulatus activates S-adenosyl-L-methionine:magnesium protoporphyrin methyltransferase. FEBS Lett. 407, 337–342. doi:10.1016/S0014- 5793(97)00371-2. Hollingshead, S., Kopecná, J., Jackson, P. J., Canniffe, D. P., Davison, P. A., Dickman, M. J., et al. (2012). Conserved chloroplast open-reading frame ycf54 is required for activity of the magnesium protoporphyrin monomethylester oxidative cyclase in Synechocystis PCC 6803. J. Biol. Chem. 287, 27823–27833. doi:10.1074/jbc.M112.352526. Hörtensteiner, S., and Kräutler, B. (2011). Chlorophyll breakdown in higher plants. Biochim. Biophys. Acta - Bioenerg. 1807, 977–988. doi:10.1016/j.bbabio.2010.12.007. Huster, M. S., and Smith, K. M. (1990). Biosynthetic studies of substituent homologation in bacteriochlorophylls c and d. Biochemistry 29, 4348–4355. doi:10.1021/bi00470a013. Islam, M. R., Aikawa, S., Midorikawa, T., Kashino, Y., Satoh, K., and Koike, H. (2008). slr1923 of Synechocystis sp PCC6803 is essential for conversion of 3,8- divinyl(proto)chlorophyll(ide) to 3-monovinyl(proto)chlorophyll(ide). Plant Physiol. 148, 1068–1081. doi:10.1104/pp.108.123117. Ito, H., Yokono, M., Tanaka, R., and Tanaka, A. (2008). Identification of a novel vinyl reductase gene essential for the biosynthesis of monovinyl chlorophyll in Synechocystis sp. PCC6803. J. Biol. Chem. 283, 9002–9011. doi:10.1074/jbc.M708369200. Jaschke, P. R., Hardjasa, A., Digby, E. L., Hunter, C. N., and Beatty, J. T. (2011). A bchD (magnesium chelatase) mutant of Rhodobacter sphaeroides synthesizes zinc 71 bacteriochlorophyll through novel zinc-containing intermediates. J. Biol. Chem. 286, 20313–20322. doi:10.1074/jbc.M110.212605. Jensen, P. E., Gibson, L. C. D., and Hunter, C. N. (1999). ATPase activity associated with the magnesium-protoporphyrin IX chelatase enzyme of Synechocystis PCC6803: evidence for ATP hydrolysis during Mg2+ insertion. Biochem. J. 339, 127–134. doi:10.1042/0264-6021:3390127. Johnson, E. T., and Schmidt-Dannert, C. (2008). Characterization of three homologs of the large subunit of the magnesium chelatase from Chlorobaculum tepidum and interaction with the magnesium protoporphyrin IX methyltransferase. J. Biol. Chem. 283, 27776–27784. doi:10.1074/jbc.M804486200. Jones, M. S., and Jones, O. T. (1970). Ferrochelatase of Rhodopseudomonas spheroides. Biochem. J. 119, 453–462. doi:10.1042/bj1190453. Kaneko, T., Tanaka, A., Sato, S., Kotani, H., Sazuka, T., Miyajima, N., et al. (1995). Sequence analysis of the genome of the unicellular cyanobacterium Synechocystis sp. strain PCC6803. I. Sequence features in the 1 Mb region from map positions 64% to 92% of the genome. DNA Res. 2, 153–166. doi:10.1093/dnares/2.4.153. Kato, K., Tanaka, R., Sano, S., Tanaka, A., and Hosaka, H. (2010). Identification of a gene essential for protoporphyrinogen IX oxidase activity in the cyanobacterium Synechocystis sp. PCC6803. Proc. Natl. Acad. Sci. 107, 16649–16654. doi:10.1073/pnas.1000771107. Kenner, G. W., Rimmer, J., Smith, K. M., and Unsworth, J. F. (1978). Pyrroles and related compounds. 39. Structural and biosynthetic-studies of Chlorobium chlorophylls-660 (bacteriochlorophylls-c) - incorporations of methionine and porphobilinogen. J. Chem. Soc. Trans. 1, 845–852. doi:10.1039/p19780000845. Kiesel, S., Waetzlich, D., Lange, C., Reijerse, E., Broecker, M. J., Ruediger, W., et al. (2015). Iron-sulfur cluster-dependent catalysis of chlorophyllide a oxidoreductase from Roseobacter denitrificans. J. Biol. Chem. 290, 1141–1154. doi:10.1074/jbc.M114.617761. Kobayashi, K., Masuda, T., Tajima, N., Wada, H., and Sato, N. (2014). Molecular phylogeny and intricate evolutionary history of the three isofunctional enzymes involved in the oxidation of protoporphyrinogen IX. Genome Biol. Evol. 6, 2141– 2155. doi:10.1093/gbe/evu170. Lange, C., Kiesel, S., Peters, S., Virus, S., Scheer, H., Jahn, D., et al. (2015). Broadened substrate specificity of 3-hydroxyethyl bacteriochlorophyllide a dehydrogenase (BchC) indicates a new route for the biosynthesis of bacteriochlorophyll a. J. Biol. Chem. 290, 19697–19709. doi:10.1074/jbc.M115.660555. Larsen, K. L., Miller, M., and Cox, R. P. (1995). Incorporation of exogenous long-chain alcohols into bacteriochlorophyll c homologs by Chloroflexus aurantiacus. Arch. Microbiol. 163, 119–123. doi:10.1007/BF00381785. Layer, G., Grage, K., Teschner, T., Schünemann, V., Breckau, D., Masoumi, A., et al. (2005). Radical S-adenosylmethionine enzyme coproporphyrinogen III oxidase HemN: Functional features of the [4Fe-4S] cluster and the two bound S-adenosyl-L- methionines. J. Biol. Chem. 280, 29038–29046. doi:10.1074/jbc.M501275200. Liu, Z., Klatt, C. G., Ludwig, M., Rusch, D. B., Jensen, S. I., Kühl, M., et al. (2012). “Candidatus Thermochlorobacter aerophilum”: An aerobic chlorophotoheterotrophic 72 member of the phylum Chlorobi defined by metagenomics and metatranscriptomics. ISME J. 6, 1869–1882. doi:10.1038/ismej.2012.24. Liu, Z., and Bryant, D. A. (2012). “Bacteriochlorophyll biosynthesis and assembly in green chlorophototrophic bacteria: Theme and variations,” in Handbook of Porphyrin Science with Applications to Chemistry, Physics, Materials Science, Engineering, Biology and Medicine, Vol 20: Chlorophylls and Related Systems Handbook of Porphyrin Science., ed. R. Kadish, KM and Smith, KM and Guilard (Singapore: World Scientific Publishing Company Pte Ltd), 107–140. Liu, Z., and Bryant, D. A. (2011a). Multiple types of 8-vinyl reductases for (Bacterio)chlorophyll biosynthesis occur in many green sulfur bacteria. J. Bacteriol. 193, 4996–4998. doi:10.1128/JB.05520-11. Liu, Z., and Bryant, D. A. (2011b). Identification of a gene essential for the first committed step in the biosynthesis of bacteriochlorophyll c. J. Biol. Chem. 286, 22393–22402. doi:10.1074/jbc.M111.249433. Lundqvist, J., Elmlund, H., Wulff, R. P., Berglund, L., Elmlund, D., Emanuelsson, C., et al. (2010). ATP-induced conformational dynamics in the AAA+ motor unit of magnesium chelatase. Structure 18, 354–365. doi:10.1016/j.str.2010.01.001. Möbius, K., Arias-Cartin, R., Breckau, D., Haennig, A.L., Riedmann, K., Biedendieck, R., et al. (2010). Heme biosynthesis is coupled to electron transport chains for energy generation. Proc. Natl. Acad. Sci. U. S. A. 107, 10436–10441. doi:10.1073/pnas.1000956107. Maresca, J. A., Gomez Maqueo Chew, A., Ponsatí, M. R., Frigaard, N. U., Ormerod, J. G., and Bryant, D. A. (2004). The bchU gene of Chlorobium tepidum encodes the C- 20 methyltransferase in bacteriochlorophyll c biosynthesis. J. Bacteriol. 186, 2558– 2566. doi:10.1128/JB.186.9.2558-2566.2004. Maresca, J. A., Romberger, S. P., and Bryant, D. A. (2008). Isorenieratene biosynthesis in green sulfur bacteria requires the cooperative actions of two carotenoid cyclases. J. Bacteriol. 190, 6384–6391. doi:10.1128/JB.00758-08. Matsuo, E., and Inagaki, Y. (2018). Patterns in evolutionary origins of heme, chlorophyll a and isopentenyl diphosphate biosynthetic pathways suggest non photosynthetic periods prior to plastid replacements in dinoflagellates. PEER J 6, e5345. doi:10.7717/peerj.5345. McGlynn, P., and Hunter, C.N. (1993). Genetic analysis of the bchC and bchA genes of Rhodobacter sphaeroides. MGG Mol. Gen. Genet. 236, 227–234. doi:10.1007/BF00277117. McLean, S., and Hunter, C. N. (2009). An enzyme-coupled continuous spectrophotometric assay for magnesium protoporphyrin IX methyltransferases. Anal. Biochem. 394, 223–228. doi:10.1016/j.ab.2009.07.036. Mizoguchi, T., Harada, J., and Tamiaki, H. (2012). Characterization of chlorophyll pigments in the mutant lacking 8-vinyl reductase of green photosynthetic bacterium Chlorobaculum tepidum. Bioorganic Med. Chem. 20, 6803–6810. doi:10.1016/j.bmc.2012.09.054. Mizoguchi, T., Harada, J., and Tamiaki, H. (2006). Structural determination of dihydro- and tetrahydrogeranylgeranyl groups at the 17-propionate of bacteriochlorophylls-a. FEBS Lett. 580, 6644–6648. doi:10.1016/j.febslet.2006.11.020. 73 Mizoguchi, T., Harada, J., Yamamoto, K., and Tamiaki, H. (2015). Inactivation of bciD and bchU genes in the green sulfur bacterium Chlorobaculum limnaeum and alteration of photosynthetic pigments in the resultant mutants. J. Photochem. Photobiol. A Chem. 313, 52–59. doi:10.1016/j.jphotochem.2015.06.012. Muraki, N., Nomata, J., Ebata, K., Mizoguchi, T., Shiba, T., Tamiaki, H., et al. (2010). X-ray crystal structure of the light-independent protochlorophyllide reductase. Nature 465, 110–114. doi:10.1038/nature08950. Nagaraj, V. A., Arumugam, R., Prasad, D., Rangarajan, P. N., and Padmanaban, G. (2010). Protoporphyrinogen IX oxidase from Plasmodium falciparum is anaerobic and is localized to the mitochondrion. Mol. Biochem. Parasitol. 174, 44–52. doi:10.1016/j.molbiopara.2010.06.012. Nagata, N., Tanaka, R., Satoh, S., and Tanaka, A. (2005). Identification of a vinyl reductase gene for chlorophyll synthesis in Arabidopsis thaliana and implications for the evolution of Prochlorococcus species. Plant Cell Online 17, 233–240. doi:10.1105/tpc.104.027276. Nakanishi, H., Nozue, H., Suzuki, K., Kaneko, Y., Taguchi, G., and Hayashida, N. (2005). Characterization of the Arabidopsis thaliana mutant pcb2 which accumulates divinyl chlorophylls. Plant Cell Physiol. 46, 467–473. doi:10.1093/pcp/pci053. Niedzwiedzki, D. M., Orf, G. S., Tank, M., Vogl, K., Bryant, D. A., and Blankenship, R. E. (2014). Photophysical properties of the excited states of bacteriochlorophyll f in solvents and in chlorosomes. J. Phys. Chem. B 118, 2295–2305. doi:10.1021/jp409495m. Nishimori, R., Mizoguchi, T., Tamiaki, H., Kashimura, S., and Saga, Y. (2011). Biosynthesis of unnatural bacteriochlorophyll c derivatives esterified with alpha, omega-diols in the green sulfur photosynthetic bacterium Chlorobaculum tepidum. Biochemistry 50, 7756–7764. doi:10.1021/bi200994h. Nomata, J., Mizoguchi, T., Tamiaki, H., and Fujita, Y. (2006). A second nitrogenase-like enzyme for bacteriochlorophyll biosynthesis: Reconstitution of chlorophyllide a reductase with purified X-protein (BchX) and YZ-protein (BchY-BchZ) from Rhodobacter capsulatus. J. Biol. Chem. 281, 15021–15028. doi:10.1074/jbc.M601750200. Nomata, J., Swem, L. R., Bauer, C. E., and Fujita, Y. (2005). Overexpression and characterization of dark-operative protochlorophyllide reductase from Rhodobacter capsulatus. Biochim. Biophys. Acta 1708, 229–237. doi:10.1016/j.bbabio.2005.02.002. Orf, G. S., Tank, M., Vogl, K., Niedzwiedzki, D. M., Bryant, D. A., and Blankenship, R. E. (2013). Spectroscopic insights into the decreased efficiency of chlorosomes containing bacteriochlorophyll f. Biochim. Biophys. Acta - Bioenerg. 1827, 493–501. doi:10.1016/j.bbabio.2013.01.006. Oster, U., Bauer, C. E., and Rüdiger, W. (1997). Characterization of chlorophyll a and bacteriochlorophyll a synthases by heterologous expression in Escherichia coli. J. Biol. Chem. 272, 9671–9676. doi:10.1074/jbc.272.15.9671. Pinta, V., Picaud, M., Reiss-Husson, F., and Astier, C. (2002). Rubrivivax gelatinosus acsF (previously orf358) codes for a conserved, putative binuclear-iron-cluster- 74 containing protein involved in aerobic oxidative cyclization of Mg-protoporphyrin IX monomethylester. J. Bacteriol. 184, 746–753. doi:10.1128/JB.184.3.746-753.2002. Porra, R. J., Schafer, W., Gadon, N., Katheder, I., Drews, G., and Scheer, H. (1996). Origin of the two carbonyl oxygens of bacteriochlorophyll alpha - Demonstration of two different pathways for the formation of ring E in Rhodobacter sphaeroides and Roseobacter denitrificans, and a common hydratase mechanism for 3-acetyl group formation. Eur. J. Biochem. 239, 85–92. doi:10.1111/j.1432-1033.1996.0085u.x. Reid, J. D., and Hunter, C. N. (2004). Magnesium-dependent ATPase activity and cooperativity of magnesium chelatase from Synechocystis sp PCC6803. J. Biol. Chem. 279, 26893–26899. doi:10.1074/jbc.M400958200. Reid, J. D., Siebert, C. A., Bullough, P. A., and Hunter, C. N. (2003). The ATPase activity of the ChlI subunit of magnesium chelatase and formation of a heptameric AAA(+) ring. Biochemistry 42, 6912–6920. doi:10.1021/bi034082q. Reinbothe, C., Bakkouri, M. El, Buhr, F., Muraki, N., Nomata, J., Kurisu, G., et al. (2010). Chlorophyll biosynthesis: Spotlight on protochlorophyllide reduction. Trends Plant Sci. 15, 614–624. doi:10.1016/j.tplants.2010.07.002. Rüdiger, W., Benz, J., and Guthoff, C. (1980). Detection and partial characterization of activity of chlorophyll synthetase in etioplast membranes. Eur. J. Biochem. 109, 193– 200. doi:10.1111/j.1432-1033.1980.tb04784.x. Saga, Y., Oh-oka, H., Hayashi, T., and Tamiaki, H. (2003). Presence of exclusively bacteriochlorophyll-c containing substrain in the culture of green sulfur photosynthetic bacterium Chlorobium vibrioforme strain NCIB 8327 producing bacteriochlorophyll-d. Anal. Sci. 19, 1575–1579. doi:10.2116/analsci.19.1575. Saga, Y., Hayashi, K., Hirota, K., Harada, J., and Tamiaki, H. (2015a). Modification of the esterifying farnesyl chain in light-harvesting bacteriochlorophylls in green sulfur photosynthetic bacteria by supplementation of 9-decyn-1-ol, 9-decen-1-ol, and decan- 1-ol. J. Photochem. Photobiol. A Chem. 313, 44–51. doi:10.1016/j.jphotochem.2015.05.002. Saga, Y., Hayashi, K., Mizoguchi, T., and Tamiaki, H. (2014). Biosynthesis of bacteriochlorophyll c derivatives possessing chlorine and bromine atoms at the terminus of esterifying chains in the green sulfur bacterium Chlorobaculum tepidum. J. Biosci. Bioeng. 118, 82–87. doi:10.1016/j.jbiosc.2013.12.023. Saga, Y., Hirota, K., Harada, J., and Tamiaki, H. (2015b). In vitro enzymatic activities of bacteriochlorophyll a synthase derived from the green sulfur photosynthetic bacterium Chlorobaculum tepidum. Biochemistry 54, 4998–5005. doi:10.1021/acs.biochem.5b00311. Saga, Y., Yamashita, H., and Hirota, K. (2016). Introduction of perfluoroalkyl chain into the esterifying moiety of bacteriochlorophyll c in the green sulfur photosynthetic bacterium Chlorobaculum tepidum by pigment biosynthesis. Bioorganic Med. Chem. 24, 4165–4170. doi:10.1016/j.bmc.2016.07.004. Sarma, R., Barney, B. M., Hamilton, T. L., Jones, A., Seefeldt, L. C., and Peters, J. W. (2008). Crystal structure of the L protein of Rhodobacter sphaeroides light- independent protochlorophyllide reductase with MgADP bound: A homologue of the nitrogenase Fe protein. Biochemistry 47, 13004–13015. doi:10.1021/bi801058r. 75 Sasarman, A., Letowski, J., Czaika, G., Ramirez, V., Nead, M. A., Jacobs, J. M., et al. (1993). Nucleotide-sequence of the hemg gene involved in the protoporphyrinogen oxidase activity of Escherichia-coli K12. Can. J. Microbiol. 39, 1155–1161. doi:10.1139/m93-174. Saunders, A. H., Golbeck, J. H., and Bryant, D. A. (2013). Characterization of BciB: A ferredoxin-dependent 8-vinyl- protochlorophyllide reductase from the green sulfur bacterium Chloroherpeton thalassium. Biochemistry 52, 8442–8451. doi:10.1021/bi401172b. Sawicki, A., and Willows, R. D. (2010). BchJ and BchM interact in a 1:1 ratio with the magnesium chelatase BchH subunit of Rhodobacter capsulatus. FEBS J. 277, 4709– 4721. doi:10.1111/j.1742-4658.2010.07877.x. Schoch, S., Oster, U., Mayer, K., Feick, R., and Rudiger, W. (1999). Subtrate specificity of overexpressed bacteriochlorophyll synthase from Chloroflexus aurantiacus. In: Chloroplast: From Molecular Biology to Biotechnology, J. H. Argyroudi Akoyunoglou, H. Senger, Eds. (Netherlands: Springer), pp. 213–216. Shepherd, M., McLean, S., and Hunter, C. N. (2005). Kinetic basis for linking the first two enzymes of chlorophyll biosynthesis. FEBS J. 272, 4532–4539. doi:10.1111/j.1742-4658.2005.04873.x. Sirijovski, N., Lundqvist, J., Rosenbäck, M., Elmlund, H., Al-Karadaghi, S., Willows, R. D., et al. (2008). Substrate-binding model of the chlorophyll biosynthetic magnesium chelatase BchH subunit. J. Biol. Chem. 283, 11652–11660. doi:10.1074/jbc.M709172200. Skotnicová, P., Sobotka, R., Shepherd, M., Hájek, J., Hrouzek, P., and Tichý, M. (2018). The cyanobacterial protoporphyrinogen oxidase HemJ is a new b-type heme protein functionally coupled with coproporphyrinogen III oxidase. J. Biol. Chem. 293, 12394–12404. doi:10.1074/jbc.RA118.003441. Steensgaard, D. B., Cox, R. P., and Miller, M. (1996). Manipulation of the bacteriochlorophyll c homolog distribution in the green sulfur bacterium Chlorobium tepidum. Photosynth. Res. 48, 385–393. doi:10.1007/BF00029471. Suzuki, J. Y., Bollivar, D. W., and Bauer, C. E. (1997). Genetic analysis of chlorophyll biosynthesis. Annu. Rev. Genet. 31, 61–89. doi:10.1146/annurev.genet.31.1.61. Suzuki, Y., Doi, M., and Shioi, Y. (2002). Two enzymatic reaction pathways in the formation of pyropheophorbide a. Photosynth. Res. 74, 225–233. doi:10.1023/A:1020919929608. Tamiaki, H., Komada, J., Kunieda, M., Fukai, K., Yoshitomi, T., Harada, J., et al. (2011). In vitro synthesis and characterization of bacteriochlorophyll-f and its absence in bacteriochlorophyll-e producing organisms. Photosynth. Res., 133–138. doi:10.1007/s11120-010-9603-3. Tanaka, A., Ito, H., Tanaka, R., Tanaka, N. K., Yoshida, K., and Okada, K. (1998). Chlorophyll a oxygenase (CAO) is involved in chlorophyll b formation from chlorophyll a. Proc. Natl. Acad. Sci. 95, 12719–12723. doi:10.1073/pnas.95.21.12719. Tang, K. H., Barry, K., Chertkov, O., Dalin, E., Han, C. S., Hauser, L. J., et al. (2011). Complete genome sequence of the filamentous anoxygenic phototrophic bacterium Chloroflexus aurantiacus. BMC Genomics 12, 334. doi:10.1186/1471-2164-12-334. 76 Tang, K. H., Wen, J., Li, X., and Blankenship, R. E. (2009). Role of the AcsF protein in Chloroflexus aurantiacus. J. Bacteriol. 191, 3580–3587. doi:10.1128/JB.00110-09. Teramura, M., Harada, J., Mizoguchi, T., Yamamoto, K., and Tamiaki, H. (2016a). In vitro assays of BciC showing C132-demethoxycarbonylase activity requisite for biosynthesis of chlorosomal chlorophyll pigments. Plant Cell Physiol. 57, 1048– 1057. doi:10.1093/pcp/pcw045. Teramura, M., Harada, J., and Tamiaki, H. (2018). In vitro enzymatic assays of photosynthetic bacterial 3-vinyl hydratases for bacteriochlorophyll biosyntheses. Photosynth. Res. 135, 319–328. doi:10.1007/s11120-017-0415-6. Teramura, M., Harada, J., and Tamiaki, H. (2016b). In vitro stereospecific hydration activities of the 3-vinyl group of chlorophyll derivatives by BchF and BchV enzymes involved in bacteriochlorophyll c biosynthesis of green sulfur bacteria. Photosynth. Res. 130, 33–45. doi:10.1007/s11120-016-0220-7. Teramura, M., Harada, J., and Tamiaki, H. (2019). In vitro demethoxycarbonylation of various chlorophyll analogs by a BciC enzyme. Photosynth. Res. 139, 163–171. doi:10.1007/s11120-018-0573-1. Thweatt, J. L., Ferlez, B. H., Golbeck, J. H., and Bryant, D. A. (2017). BciD is a radical S-adenosyl-L-methionine (SAM) enzyme that completes bacteriochlorophyllide e biosynthesis by oxidizing a methyl group into a formyl group at C-7. J. Biol. Chem. 292, 1361–1373. doi:10.1074/jbc.M116.767665. Tsukatani, Y., Harada, J., Mizoguchi, T., and Tamiaki, H. (2013a). Bacteriochlorophyll homolog compositions in the bchU mutants of green sulfur bacteria. Photochem. Photobiol. Sci. 12, 2195–2201. doi:10.1039/c3pp50253h. Tsukatani, Y., Yamamoto, H., Harada, J., Yoshitomi, T., Nomata, J., Kasahara, M., et al. (2013b). An unexpectedly branched biosynthetic pathway for bacteriochlorophyll b capable of absorbing near-infrared light. Sci. Rep. 3, 1217. doi:10.1038/srep01217. Tsukatani, Y., Yamamoto, H., Mizoguchi, T., Fujita, Y., and Tamiaki, H. (2013c). Completion of biosynthetic pathways for bacteriochlorophyll g in Heliobacterium modesticaldum: The C8-ethylidene group formation. Biochim. Biophys. Acta - Bioenerg. 1827, 1200–1204. doi:10.1016/j.bbabio.2013.06.007. Wada, K., Yamaguchi, H., Harada, J., Niimi, K., Osumi, S., Saga, Y., et al. (2006). Crystal structures of BchU, a methyltransferase involved in bacteriochlorophyll c biosynthesis, and its complex with S-adenosylhomocysteine: implications for reaction mechanism. J. Mol. Biol. 360, 839–849. doi:10.1016/j.jmb.2006.05.057. Waetzlich, D., Broecker, M. J., Uliczka, F., Ribbe, M., Virus, S., Jahn, D., et al. (2009). Chimeric nitrogenase-like enzymes of (bacterio)chlorophyll biosynthesis. J. Biol. Chem. 284, 15530–15540. doi:10.1074/jbc.M901331200. Walker, C. J., Mansfield, K. E., Smith, K. M., and Castelfranco, P. A. (1989). Incorporation of atmospheric oxygen into the carbonyl functionality of the protochlorophyllide isocyclic ring. Biochem. J. 257, 599–602. doi:10.1042/bj2570599. Wang, P., Wan, C., Xu, Z., Wang, P., Wang, W., Sun, C., et al. (2013). One divinyl reductase reduces the 8-vinyl groups in various intermediates of chlorophyll biosynthesis in a given higher plant species, but the isozyme differs between species. Plant Physiol. 161, 521–534. doi:10.1104/pp.112.208421. 77 Willows, R. D., Gibson, L. C. D., Kanangara, C. G., Hunter, C. N., and von Wettstein, D. (1996). Three separate proteins constitute the magnesium chelatase of Rhodobacter sphaeroides. Eur. J. Biochem. 235, 438–443. doi:10.1111/j.1432-1033.1996.00438.x. Xiong, J., Fischer, W. M., Inoue, K., Nakahara, M., and Bauer, C. E. (2000). Molecular evidence for the early evolution of photosynthesis. Science . 289, 1724–1730. doi:10.1126/science.289.5485.1724. Zsebo, K. M., and Hearst, J. E. (1984). Genetic-physical mapping of a photosynthetic gene cluster from R. capsulata. Cell 37, 937. doi:10.1016/0092-8674(84)90428-8.
78
Figure 2.1- Distribution of Biosynthetic Enzymes for (B)Chl Biosynthesis in Green Bacteria. Color-coded array indicating the percentage of organisms in each category whose genomes encode the indicated biosynthetic enzyme. The term “HemY” refers to enzymes described as HemY-related in the text. In the ChlH/BchH column text indicates which paralogs are present in each group. In the green- colored Chlorobiaceae, Chl. chlorochromatii is missing ChlH/BchH but has BchS and BchT. In ‘Ca. T. aerophilum’ instead of a full length BchS the metagenom assembly appears to encode the N-terminus of the enzyme separately from the C-terminus, together the two genes encode a full length BchS. In the BchF column text indicates which paralogs are present. In the Brown Chlorobiaceae, Prosthecochloris sp. BS1 is missing the BchV paralog but contains BchF and BchF3. In the green-colored Chlorobiaceae, Ptc. bathyomarina and Chl. phaeovibriodes have a BchF3 paralog instead of BchV and Chl. limicola has 2 paralogs of BchV in addition to BchF. In the BchQ2 column it should be noted that BchQ and BchQ2 do not form clearly delineated clades, so for this figure a second copy of BchQ has been considered BchQ2. In the brown-colored Chlorobiaceae, Prosthecochloris sp. BS1 has only a single copy of BchQ, however the gene encoding BchQ is directly upstream of the gene cluster which encodes BciD, this is the same location where other brown-colored GSB encode BchQ2. This likely represents a loss of BchQ, as is true of BchV for this organism. Additionally, Chl. phaeobacteroides possesses a truncated version of BchQ which is encoded distally from the brown-colored GSB gene cluster. In green-colored Chlorobiaceae, Chl. limicola encodes a second copy of BchQ, as is true for BchV in this organism. Genomes analyzed for this figure can be found in Table 4.1. NP, not present.
79
Figure 2.2- Biosynthesis: 5-ALA to Chlide a. The biosynthetic pathway from 5-ALA to Chlide a is shown. Arrows indicate the reaction order, with arrow labels indicating the enzyme(s) responsible for catalysis. Grey dashed arrows indicate reactions which are less likely to occur in vivo based on experimental data (e.g., specific activity and substrate preference studies).
80
Figure 2.3- Biosynthesis: Chlide a to BChlide a and BChlide c, d, e, or f. The biosynthetic pathways from Chlide a to BChlide a and BChlide c, d, e, or f are shown. Arrows indicate the reaction order, with arrow labels indicating the enzyme(s) responsible for catalysis. Grey dashed arrows indicate reactions which are less likely to occur in vivo based on experimental data (e.g., specific activity). R8: ethyl, n-propyl, isobutyl, or neopentyl. R12: ethyl or methyl. 81
Figure 2.4- Biosynthesis: Final Steps. The final steps in the biosynthetic pathways of Chl aPD, Bchl aP, and Bchl cF are represented. Arrows indicate the reaction order, with arrow labels indicating the enzyme(s) responsible for catalysis. Dashed arrows represent possible reaction which have not be characterized. The “?” represents an unknown intermediate (in place of a molecule), or possible unknown dehydrogenase (in place of arrow label). GG: geranylgeranyl; P: phytyl; F: farnesyl; GGPP: geranylgeranyl diphosphate; DH: dihydro; TH: tetrahydro. 82 Chapter 3 Characterization of BciD from Cba. limnaeum
Publication: Jennifer L. Thweatt, Bryan H. Ferlez, John H. Golbeck and Donald A.
Bryant (2017). BciD is a radical S-adenosyl-L-methionine (SAM) enzyme that completes
bacteriochlorophyllide e biosynthesis by oxidizing a methyl group to a formyl group at
C-7. Journal of Biological Chemistry 292, 1361-1373.
Contributions: JLT designed, performed and interpreted the experiments, drafted the paper, and made all figures. BHF performed electron paramagnetic resonance (EPR) measurements and assisted with analysis of the EPR data. JHG assisted in analysis of
EPR data and editing of the manuscript. DAB conceived of the project, directed the research efforts, assisted in interpretation of data, drafted sections of the discussion and provided feedback and edits on the draft.
83 3.1 Abstract
Green bacteria are chlorophotorophs that synthesize bacteriochlorophyll (BChl) c,
d, or e, which assemble into supramolecular, nanotubular structures in large light-
harvesting structures called chlorosomes. The biosynthetic pathways of these
chlorophylls were known prior to this work, with the exception of one reaction (see
Chapter 2). Null mutants of bciD, which encodes a putative radical S-adenosyl-L-
methionine (SAM) protein, are unable to synthesize BChl e but accumulate BChl c;
however, it was unknown whether BciD is sufficient to convert BChl c (or its precursor,
bacteriochlorophyllide (BChlide) c) into BChl e (or BChlide e). To determine the
function of BciD, the bciD gene of Chlorobaculum limnaeum strain DSMZ 1677T was
expressed in Escherichia coli, and the enzyme was purified under anoxic conditions.
Electron paramagnetic resonance spectroscopy of BciD indicated that it contains a single
[4Fe-4S] cluster. In assays containing SAM, BChlide c or d, and sodium dithionite, BciD
catalyzed the conversion of SAM into 5-deoxyadenosine and BChlide c or d into BChlide
e or f, respectively. These analyses also identified intermediates that are proposed to be
71-OH-BChlide c and d. Thus, BciD is a radical SAM enzyme that converts the methyl
group of BChlide c or d into the formyl group of BChlide e or f. This probably occurs by
a mechanism involving consecutive hydroxylation reactions of the C-7 methyl group to
form a geminal diol intermediate, which spontaneously dehydrates to produce the final
products, BChlide e or BChlide f. The demonstration that BciD is sufficient to catalyze
the conversion of BChlide c into BChlide e completes the biosynthetic pathways for all
“Chlorobium chlorophylls.” 84
3.2 Introduction
Green bacteria are an eclectic ensemble of anoxygenic chlorophototrophic
bacteria that belong to three phyla: Chloroflexi, Chlorobi, and Acidobacteria (Bryant et
al., 2007, 2012; Bryant and Frigaard, 2006). Although physiologically and metabolically
diverse, green bacteria share the defining properties of synthesizing bacteriochlorophyll
(BChl) c, d, or e and assembling these BChls into very large light-harvesting complexes
known as chlorosomes (Frigaard and Bryant, 2006; Oostergetel et al., 2010; Orf and
Blankenship, 2013). Chlorosomes of green sulfur bacteria (GSB), which can contain up
to 250,000 BChl molecules (Adams et al., 2013; Bryant et al., 2002; Frigaard and Bryant,
2006; Montaño et al., 2003), are the largest known light-harvesting antenna complexes,
and correspondingly, these structures allow GSB to grow under extraordinarily low light
intensities. For example, a population of BChl e-producing Prosthecochloris sp. BS-1
(formerly Chlorobium phaeobacteroides BS-1) stably grows at the chemocline of the
Black Sea, which occurs at a depth of 100–110 m beneath the surface (Bryant et al.,
2012; Manske et al., 2005; Marschall et al., 2010). At this depth, the light intensity is nearly 106-fold lower than at the surface, 3 nmol of photons m–2 s–1, at the irradiance
levels where these bacteria occur (Maresca et al., 2004).
BChls c, d, and e (and BChl f, which does not occur naturally) (Harada et al.,
2012; Niedzwiedzki et al., 2014; Orf et al., 2013; Tsukatani et al., 2013; Vogl et al.,
2012) comprise the so-called “Chlorobium chlorophylls” (Chls). These molecules are not
actually BChls but are chlorins that are structurally and spectroscopically more similar to 85 Chl a than BChl a (Gomez Maqueo Chew and Bryant, 2007). Mutational analyses made
possible by the availability of genome sequences for numerous green bacteria led to the
establishment of the pathway for the synthesis of BChls c and d (Bryant and Liu, 2013;
Eisen et al., 2002; Frigaard et al., 2002; Gomez Maqueo Chew et al., 2004, 2007; Gomez
Maqueo Chew and Bryant, 2007; Liu and Bryant, 2011, 2012; Maresca et al., 2004).
These BChls are synthesized from a pathway that diverges from chlorophyllide (Chlide)
a (Figure 3.1, also see Chapter 2) (Gomez Maqueo Chew and Bryant, 2007). The first
committed step in the biosynthesis of BChl c, d, and e is the removal of the C-132
carboxymethyl group from Chlide a by BciC (Liu and Bryant, 2011; Teramura et al.,
2016a). BchQ and BchR methylate the resulting product at the C-82 and C-121 positions,
respectively, to produce a family of related homologs (Gomez Maqueo Chew and Bryant,
2007). These methylation homologs are then hydroxylated at C-31 by BchF or BchV to produce bacteriochlorophyllide (BChlide) d (Gomez Maqueo Chew et al., 2004; Harada et al., 2015; Teramura et al., 2016b), and in BChl c-producing strains, they are subsequently methylated by BchU at C-20 to produce BChlide c homologs (Harada et al.,
2005; Maresca et al., 2004; Wada et al., 2006). Finally, the BChlide c or BChlide d homologs are esterified with farnesol pyrophosphate by BchK to produce BChl c or BChl d homologs with differing numbers of methyl groups (Frigaared et al., 2002). The methylation reactions catalyzed by BchU, BchQ, and BchR help to tune the absorption properties of BChl c, d, and e in chlorosomes and also cause inhomogeneous broadening of the near-infrared absorption band of aggregated BChls in chlorosomes (Gomez
Maqueo Chew and Bryant, 2007; Maresca et al., 2004; Tsukatani et al., 2013). Naturally occurring or genetically engineered mutants of bchU in formerly BChl c-producing 86 strains exclusively synthesize BChl d (Bobe et al., 1990; Broch-Due and Ormerod, 1978;
Harada et al., 2004; Maresca et al., 2004; Saga and Tamiaki, 2004).
GSB that synthesize either BChl d or BChl c are green in color, whereas those
containing BChl e or the unnatural BChl f are brown. BChl e only differs from BChl c by
the presence of a formyl group instead of a methyl group at the C-7 position. Similarly,
BChl f is identical to BChl d except for the C-7 formyl group. The available genome sequences of brown-colored GSB contain homologs of all of the genes required for BChl c biosynthesis; thus, the BChl e biosynthetic pathway is predicted to proceed via the same reactions as that for BChl c until the oxidation of the methyl group at the C-7 position
(Bryant and Liu, 2013; Gomez Maqueo Chew and Bryant, 2007; Liu and Bryant, 2012).
A similar oxidation occurs at the C-7 position during the biosynthesis of Chl b from Chl a; Chl b differs from Chl a by the presence of a formyl group instead of a methyl group at the C-7 position (Scheer, 2006). In the case of Chl b, this reaction is catalyzed by an oxygen-dependent enzyme, chlorophyllide a oxygenase (Tanaka et al., 1998; Tomitani et al., 1999). However, because GSB are strictly anaerobic organisms and their genomes do not encode a homolog of this oxygenase, an as yet uncharacterized enzyme (or enzymes) capable of oxidizing the C-7 methyl group must be responsible for this reaction in the biosynthesis of BChl e.
Comparative genomics of brown-colored and green-colored GSB previously identified a conserved gene cluster that could potentially encode an enzyme involved in the conversion of BChlide c to BChlide e (Bryant et al., 2012; Bryant and Liu, 2013; Liu
and Bryant, 2012; Maresca, 2007). Similar versions of this gene cluster occur in all BChl 87 e-producing strains (Figure 3.2), and recent evidence suggests that this conserved gene cluster can be horizontally transferred among GSB by the action of bacteriophages
(Llorens-Marès et al., 2017). The smallest of these clusters contains about 11 genes extending approximately from a putative radical S-adenosyl-L-methionine (SAM) methyltransferase (bchQ2), similar to bchQ, on one border to a gene encoding a homolog of isoprenylcysteine carboxyl methyltransferase, an enzyme that methylates the C- terminal carboxyl group of prenylated proteins (Yang et al., 2011), near the other border.
Previous studies showed that one gene in this cluster, cruB, encodes a γ-carotene cyclase that is essential for the synthesis of β-carotene and isorenieratene, carotenoids that are uniquely and nearly universally produced by brown-colored GSB (Maresca et al., 2008).
This gene cluster also includes a third homolog of bchF and bchV, genes that encode C-3 vinyl hydratases (Gomez Maqueo Chew et al., 2004; Harada et al., 2015; Termaruma et al., 2016b).
The conserved gene cluster in brown-colored GSB notably contains an open reading frame, designated as bciD, which encodes a putative radical SAM enzyme.
Radical SAM enzymes are capable of performing a variety of radical-mediated reactions under anoxic conditions (Wang et al., 2014), and they often act as substitutes for oxygen- dependent enzymes in anaerobic organisms. For example, BchE is a radical SAM enzyme that catalyzes the 6-electron oxidation and cyclization of the isocylic E ring in chlorins in anaerobic chlorophototrophs (Bryant et al., 2012; Bryant and Liu, 2013; Gomez Maqueo
Chew and Bryant, 2007; Liu and Bryant, 2012). BchE catalyzes the same reaction as an oxygen-dependent monooxygenase, AcsF, or AcsF combined with Ycf54 or BciE, during 88 chlorophyll biosynthesis in aerobic and microaerophilic chlorophototrophs (Albus et al.,
2012; Bryant et al., 2012; Bryant and Liu, 2013; Chen et al., 2016, 2017, 2018;
Hollingshead et al., 2012; Liu and Bryant, 2012; Ouchane et al., 2004) (see Chapter 2).
When the bciD gene was inactivated by natural transformation in Chlorobaculum
limnaeum strain RK-j-1, the resulting strain could no longer synthesize BChl e but
instead produced BChl c (Harada et al., 2013). Although this result shows that BciD is
necessary for the synthesis of BChlide e, it does not show that BciD alone is sufficient to
catalyze this transformation. Repeated attempts to express the bciD gene, as well as some
of the surrounding genes from the conserved gene cluster shown in Figure 3.2, in
Chlorobaculum tepidum uniformly failed to produce a strain that could synthesize BChl
e. Because the phenotype of the bciD mutant reported by Harada et al. (2013) was not
verified by complementation, these failures led us to question whether BciD was directly
involved in the synthesis of BChl e.
This study verifies that inactivation of bciD in a second strain of Cba. limnaeum,
strain DSMZ 1677T, also led to the synthesis of BChl c in the resulting mutant. The bciD
gene was heterologously expressed in Escherichia coli, and the resultant protein was
purified and characterized. The purified enzyme contains a single [4Fe-4S] cluster, and when incubated with S-adenosyl-L-methionine (SAM) and sodium dithionite, it catalyzed the conversion of BChlide c into BChlide e and BChlide d into BChlide f, and it produced
5-deoxyadenosine as a by-product. These results demonstrate that BciD is a radical SAM enzyme and that it is sufficient to catalyze these two transformations. Thus, the biosynthetic pathway for all Chlorobium Chls is now complete. 89
3.3 Experimental Procedures
3.3.1 Strains Used in This Study
Cba. limnaeum strain DSMZ 1677T (Imhoff, 2003) was obtained from the culture
collection of Dr. Johannes Imhoff (University of Kiel, Germany). Cba. limnaeum is
capable of thiosulfate utilization (Vogl et al., 2012), and it was routinely grown on medium CL used for Cba. tepidum (Frigaard and Bryant, 2001; Wahlund and Madigan,
1995) at irradiances of 10–100 μmol of photons m–2 s–1 provided by either tungsten or
cool white fluorescent lamps. Cba. limnaeum cells were grown at room temperature in
CL medium in liquid and the same medium solidified with 1.5% (w/v) BactoTM Agar
(BD Biosciences) in a Coy anoxic chamber under an atmosphere of N2/CO2/H2
(80:10:10, v/v/v). Plates were further enclosed in Gas-Pak jars containing thioacetamide
as described previously (Frigaard and Bryant , 2001). The complete genome of this
organism has been sequenced and is available in GenBankTM as accession number
CP017305 (Tank et al., 2017a).
Routine cloning to produce constructions for gene inactivation were performed
using chemically competent α-select Escherichia coli cells (Bioline, Taunton, MA). The
E. coli strain used for conjugation was S17-1 (Simon et al., 1983), and E. coli strain
BL21 (DE3) was used for expression of the bciD gene. E. coli strains were grown in
Luria-Bertani (LB) medium containing antibiotics as required. For overproduction of 90 BciD, the medium was supplemented with ferric ammonium citrate (100 μM) and L-
cysteine (100 μM). Antibiotic concentrations for E. coli were as follows: spectinomycin,
100 μg ml–1; ampicillin, 100 μg ml–1; streptomycin, 50 μg ml–1; and kanamycin, 50 μg
ml–1. Antibiotic concentrations for Cba. limnaeum grown on solid medium were as follows: spectinomycin, 200 μg ml–1; streptomycin, 100 μg ml–1; kanamycin, 30 μg ml–1.
When grown in liquid medium, antibiotic concentrations for Cba. limnaeum were as follows: spectinomycin, 37.5 μg ml–1; streptomycin, 18.75 μg ml–1.
3.3.2 Cloning and Inactivation of bciD in Cba. limnaeum
Inactivation of the bciD gene of Cba. limnaeum strain DSMZ 1677T was
performed by conjugative gene transfer essentially as described (Vogl et al., 2012). A
gene-internal region of the bciD gene was amplified by PCR using oligonucleotide
primers CLbciDconSphIF and CLbciDconSalIR (Table 3.1) and cloned into the SphI and
SalI sites of the conjugation vector pCLCON (Vogl et al., 2012) to create vector
pCLCON::bciD (Figure 3.3A) The resulting plasmid was transformed into E. coli strain
S17-1. Conjugation between the S17-1 strain and wild-type Cba. limnaeum was performed as described previously (Vogl et al., 2012), and transconjugants were selected on solid medium containing spectinomycin, streptomycin, and kanamycin.
Transconjugant colonies were screened by PCR using oligonucleotide primers
CLbciDtestF and aadAtestR (Table 3.1) that flank one border of the gene insertion site; amplicons were analyzed by agarose gel electrophoresis (Figure 3.3 B), and positive 91 PCR amplicons were confirmed by DNA sequencing at the Genomics Core Facility
(Pennsylvania State University, University Park, PA).
Cloning and Heterologous Expression of bciD in E. coli—The bciD gene from
Cba. limnaeum was amplified via PCR using oligonucleotide primers CLbciDNdeIF and
CLbciDEcoRIR (Table 3.1). The resulting PCR amplicon and plasmid pET26b were digested with NdeI and EcoRI, and the products were ligated to insert the bciD gene upstream of the sequence for a C-terminal His6 tag. Site-directed mutagenesis was used to
change the bciD stop codon to a leucine codon using the QuikChange II mutagenesis kit
(Agilent Technologies, Santa Clara, CA) and oligonucleotide primers CLbciDstopLeuF
and CLbciDstopLeuR (Table 3.1). The complete coding sequence and the downstream
flanking region encoding the His6 tag in the resulting plasmid, pET26b(bciD-His6), were
confirmed by DNA sequencing at the Genomics Core Facility.
The expression and purification of BciD-His6 was modified from protocols used
for other radical SAM enzymes (Lanz et al., 2012). The expression plasmid
pET26b(bciD-His6) was transformed into E. coli strain BL21 (DE3) together with
plasmid pDB1282, which contains part of the Azotobacter vinelandii isc operon for [Fe-
S] cluster assembly under the control of an arabinose repressor and operator (plasmid
provided by the laboratory of Dr. Squire Booker) ( Delli-Bovi et al., 2010; Lanz et al.,
2012; Zheng et al., 1998). A starter culture of the expression strain was grown overnight
at 37 °C in LB medium supplemented with ampicillin and kanamycin. A 2.5-liter culture
in minimal medium containing 2% (v/v) glycerol, ampicillin, and kanamycin was
inoculated with a 25-ml overnight culture. The culture was grown at 37 °C with gentle 92
rotation to an OD600 of 0.2– 0.3, and expression of the isc operon of pDB1282 was
induced with 0.2% L-arabinose after supplementing the medium with ferric ammonium
citrate (100 μM) and cysteine (100 μM). After the addition of arabinose, cells were grown at 37 °C to an OD600 of 0.7– 0.8 and then cooled to 18 °C. Finally, BciD-His6
production was induced by adding isopropyl-β-D-1-thiogalactopyranoside (50 μM). After further incubation at 18 °C for 16–20 h, cells were harvested by centrifugation and either immediately transferred to an anoxic chamber or frozen and placed at 80 °C for short term storage before lysis.
3.3.3 BciD Purification
Buffers used for purification of BciD were made in an anoxic chamber by
dissolving the solid materials in anoxic water. The cell pellet was thawed on ice in the
anoxic chamber, and cells were resuspended in lysis buffer containing 50 mM HEPES,
pH 7.5, 300 mM NaCl, 20 mM imidazole, and 10 mM 2-mercaptoethanol. Lysozyme was
added to 1 mg ml–1, and the suspension was incubated with rocking for 30 min at room temperature. Cells were cooled on ice and then lysed by sonication. Lysed cells were transferred to ultracentrifuge tubes and sealed inside the anoxic chamber before pelleting the cell debris by ultracentrifugation. The tubes were returned to the anoxic chamber, and the soluble fraction was loaded onto a nickel affinity column pre-equilibrated with lysis
buffer. The column was washed three times with three volumes of wash buffer. Wash
buffer contained 50 mM HEPES, pH 7.5, 300 mM NaCl, 40 mM imidazole, 10 mM 2-
mercaptoethanol, and 20% glycerol. BciD was eluted with a minimal volume of elution 93 buffer, which was the same as the wash buffer except for the imidazole concentration
(250 mM). The dark brown-colored fractions were pooled and concentrated using an
Amicon stirred ultrafiltration cell with a YM-10 filter (10,000 molecular weight cut-off;
Millipore, Bedford, MA). The protein was exchanged into storage buffer containing 50 mM HEPES, pH 7.5, 100 mM NaCl, 10 mM dithiothreitol, and 20% glycerol using PD-
10 gel filtration columns (GE Healthcare) and stored in aliquots in liquid nitrogen.
3.3.4 Protein Analyses and Verification
The purity of the BciD-His6 protein was assessed by electrophoresis on 12% (w/v) polyacrylamide gels in the presence of SDS; proteins were stained with Coomassie Blue
R-250 (Shen and Bryant, 1995). The identity of purified BciD-His6 was verified by in-gel trypsin digestion (Ho et al., 2016) accompanied by mass spectrometry at the Proteomics and Mass Spectrometry Core Facility (The Pennsylvania State University, University
Park, PA). Protein concentrations were determined by the Bradford assay using a BSA standard (Bradford, 1976). Quantitative amino acid analysis performed by the University of California-Davis Proteomics Core Facility established that a correction factor of 0.87 was necessary for protein concentrations determined by the Bradford assay.
3.3.5 Reconstitution of Fe/S Cluster
The reconstitution procedure (Lanz et al., 2012; Lovenberg et al., 1963; Zhao et al., 1990) was carried out on ice with gentle stirring in an anoxic chamber. A final 94 concentration of dithiothreitol (2 mg ml–1) was added to reconstitution buffer (100 mM
HEPES, pH 7.5, 300 mM NaCl, 10% (v/v) glycerol), and the solution was incubated for
10 min before adding 300 μM FeCl3. Purified BciD-His6 was added to a final
concentration of 30 μM, and the solution was incubated for 1 h. A Na2S solution (100
mM) was slowly added to the protein solution over a 3-h period to produce a final
concentration of 300 μM, and the resulting reaction mixture was incubated on ice for 12 h. The protein solution was centrifuged at 10,000 × g for 10 min, and the supernatant was
concentrated and exchanged into storage buffer. To remove adventitiously bound iron,
BciD-His6 was chromatographed on a Superdex 75 10/300 GL size exclusion column
(GE Healthcare) maintained in an anoxic chamber.
3.3.6 Spectroscopic Measurements
UV-visible absorbance spectra of protein and pigment samples were measured on a GENESYS 10 spectrophotometer (ThermoFisher Scientific Corp., Waltham, MA).
Spectra of protein samples were measured in quartz cuvettes that were sealed inside an
anoxic chamber.
For EPR measurements, samples of as-isolated and reconstituted BciD-His6 were
transferred to EPR tubes in storage buffer. To characterize the presence of Fe/S cluster(s),
each protein preparation was measured in both the untreated and reduced states. To
reduce the Fe/S cluster(s) fully, dithionite in storage buffer was added to a final
concentration of 5 mM to the appropriate protein samples, and the samples were then
frozen in liquid nitrogen. EPR spectra were collected using a cylindrical TE 011 mode 95 resonator and a Bruker E500 spectrometer (Bruker Biospin Corp., Billerica, MA) at X- band (9.39 GHz). The temperature was maintained using an ESR 900 liquid helium cryostat and an ITC-4 temperature controller (Oxford Instruments, Concord, MA). EPR spectra were recorded at 15 K at 0.2-milliwatt microwave power with modulation amplitude of 20 G (2 millitesla) at a modulation frequency of 100 kHz. Sixteen scans were collected and averaged for each sample; the difference spectrum was calculated by subtracting the spectrum for the untreated protein from that for the dithionite reduced protein.
3.3.7 Pigment Separation and Preparation of Substrate Compounds
Pigments were separated and analyzed by reversed phase HPLC using a 25 cm ×
4.6-mm Discovery 5-μm C-18 column (Supelco, Bellefonte, PA) and an Agilent series
1100 HPLC system equipped with a diode array detector (Agilent Technologies, Palo
Alto, CA) (Frigaard et al., 1997). The data were analyzed using Agilent ChemStation software (revision B.02.01-SR1 6100 series). Pigment samples or reaction products were filtered through a 0.2-μm syringe filter, and 10 mM ammonium acetate was added before injection onto the column. Previously described separation methods (Borrego et al., 1994,
1999; Mallorqui et al., 2005) were modified slightly to separate the BChlide and bacteriopheophorbide (BPheide) components of interest. The column was pre- equilibrated with 12% solvent B (methanol, ethyl acetate, acetonitrile, 50:30:20 (v/v/v)) and 88% solvent A (methanol, 1 M ammonium acetate, 70:30 (v/v)) at a flow rate of 0.75 ml min–1 at injection and maintained for 5 min. After 5 min, solvent B was increased 96 linearly to 43%, and the flow rate was increased linearly to 1 ml min–1 at 44 min. Solvent
B was linearly increased to 100% from 44 to 55 min and then held constant for 10 min.
The column was then returned to 12% solvent B and washed extensively before the next
injection.
BChlide c was prepared from a 2-liter culture of the bciD mutant of Cba. limnaeum (see “Results”), and BChlide d was prepared from a 2-liter culture of the bchU
mutant of Cba. tepidum (Maresca et al., 2004). Cells were harvested by centrifugation,
and pigments were extracted with 7:2 acetone/methanol (40 ml) for 1 h at room
temperature in the dark with stirring. Cell debris was removed by centrifugation, and the
solubilized pigments were purified using a previously described, reversed-phase HPLC
method (Vogl et al., 2012). The BChl fractions were collected, pooled, dried under liquid nitrogen, washed with acetone, and finally dried again. To hydrolyze the esterifying alcohol groups of the BChls, the dried pigments were suspended in 4:1 solution of acetone/water containing 0.1 M NaOH, and the solution was incubated in the dark for 3 h
(Saga et al., 2015). One volume of hexane was added to extract unreacted BChls, and the acetone layer was dried under liquid nitrogen and washed three times with acetone. Dried samples were stored at 20 °C until required.
3.3.8 Mass Spectrometry
Mass spectrometric analyses were performed at the Mass Spectrometry Core
Facility (Huck Institutes for the Life Sciences, The Pennsylvania State University) on a
Waters Q-TOF Premier quadrupole/time-of-flight mass spectrometer (Waters Corp. 97 (Micromass Ltd.), Manchester, UK). MassLynxTM software version 4.1 was used to operate the mass spectrometer. Samples were introduced into the mass spectrometer using a Waters 2695 HPLC. The separation was performed using the same HPLC column and solvent gradient described above for analysis of the reaction products. The samples
were resuspended in acetonitrile (100 μl) and vortexed until dissolved. The injection
volume was 25 μl. The nitrogen drying gas temperature was set to 300 °C at a flow of 7
liters min–1, and the capillary voltage was 2.8 kV. The mass spectrometer was set to scan
from 400 to 700 m/z in positive ion mode, using electrospray ionization. Data acquisition
was performed in the middle of the run, dependent upon the anticipated elution time of
the analytes.
3.3.9 Enzyme Activity Assay
Enzyme assays were performed inside an anoxic chamber in the dark at room
temperature. The reaction buffer consisted of 100 mM MOPS, pH 8.0, 100 mM NaCl.
Tryptophan (200 μM final concentration) was added as an internal standard. In addition, a
final concentration of 700 μM SAM and 1 mM dithionite were added as cofactor and
reductant, respectively. Reactions contained a final concentration of 0.2 mg ml–1 purified
BciD; an equal volume of enzyme storage buffer was added to control reactions. BChlide
c (final concentration, 0.05 mg ml–1) or BChlide d (final concentration, 0.18 mg ml–1) in
acetone was added to the enzyme solution to start the reaction. At appropriate times,
aliquots were removed from each reaction for analysis. An equal volume of cold acetone
was added to stop the reaction, and precipitated protein was pelleted by centrifugation. 98 Samples were stored on ice or at 20 °C until the pigments were analyzed by reversed
phase HPLC.
To analyze SAM-related products, separate aliquots were taken at each time
point, and an equal volume of 100 mM H2SO4 was added to stop the reaction. SAM
products were separated by a reversed-phase HPLC procedure modified from Lanz et al.
(2012), using a 150 × 4.6-mm Kinetex 5-μm C-18 column (Phenomenex Inc., Torrance,
CA) and a Shimadzu UFLC system (Shimadzu Scientific Instruments, Columbia, MD).
Reaction products were filtered through a syringe filter (0.2 μm) before being injected
onto the column. The column was pre-equilibrated with 2% solvent B (HPLC grade
acetonitrile) and 98% solvent A (5% methanol, 40 mM ammonium acetate, pH 6.2) at a
flow rate of 0.5 ml min–1 at injection and maintained for 6.5 min. Solvent B was
increased linearly to 12% from 6.5 to 25 min, then to 24% from 25 to 33 min, and finally
to 50% from 33 to 40 min. Solvent B was maintained at 50% for 10 min and then
returned to the starting conditions. Products were monitored by UV-visible detection at
260 nm and were compared with authentic standards for SAM and 5ʹ-deoxyadenosine.
3.3.10 Phylogenetic Analyses
Phylogenetic relationships were inferred by the maximum likelihood method based on the JTT matrix-based model (Jones et al., 1992). The tree with the highest log likelihood value (10,582.4753) is shown. Based upon 100 bootstrap resamplings of the data, the percentage of trees in which the associated taxa clustered together is indicated next to the branches. Initial tree(s) for the heuristic search were obtained automatically by 99 applying neighbor-joining and BioNJ algorithms to a matrix of pairwise distances that
were estimated using a JTT model and then by selecting the topology with the highest log
likelihood value. The tree is drawn to scale, with branch lengths scaled to the number of
substitutions per site. The analysis employed 52 amino acid sequences. All positions
containing gaps due to insertions/deletions were eliminated from the alignment. The final
alignment data set contained 244 positions. Evolutionary analyses were conducted in
MEGA7 (Kumar et al., 2016).
3.4 Results
3.4.1 Inactivation of bciD in Cba. limnaeum
To confirm the phenotype of the bciD mutant described by Harada et al. (2013), the bciD gene in Cba. limnaeum 1677T was insertionally inactivated. The mutant was
made by conjugation from E. coli strain S17-1 carrying the plasmid pCLCON::bciD with
Cba. limnaeum 1677T. This plasmid contained a single homologous region completely
internal to the bciD gene. After a single homologous recombination event, this should
disrupt the bciD gene as shown in Figure 3.3A. Transconjugants were screened by PCR
with primers bciDtestF and aadAtestR flanking one border of the insertion site (Table
3.1). Positive transconjugants produced a PCR amplicon of 1.1 kb as expected (Figure
3.3B), whereas wild-type cells did not produce an amplicon. The resulting 1.1-kb PCR
product was verified by DNA sequencing. 100 Cultures of the bciD mutant of Cba. limnaeum were green in color, whereas wild-
type cultures are brown. To confirm that this color change was due to a change in the
BChl content of the bciD mutant strain, the UV-visible absorption spectra of whole cells and of the methanol-extracted pigments of wild-type Cba. limnaeum were compared to the bciD mutant of Cba. limnaeum, and wild-type Cba. tepidum, a closely related BChl c- containing GSB (Figure 3.4) (Imhoff, 2003). Whole-cell spectra (Figure 3.4A) of the bciD mutant very closely matched the spectrum of wild-type Cba. tepidum. The only notable difference was observed around 515 nm, where Cba. tepidum appears to have greater absorbance due to its greater carotenoid content. Similar to Cba. tepidum, the Qy
absorption peak of the green-colored, bciD mutant strain shifted to longer wavelength, whereas the Soret absorption peak of the mutant shifted to shorter wavelength when
compared to WT Cba. limnaeum. The bciD mutant also lacked the secondary absorption
peak at around 525 nm that is typical of brown-colored, BChl e-containing strains. The
UV-visible absorption spectra of the extracted pigments of the bciD mutant again
matched that of Cba. tepidum except for the carotenoid region (450–500 nm), where Cba.
tepidum again exhibited greater absorbance (Figure 3.4B). These observations agree with
those reported by Harada et al. (2013) and confirm that inactivation of bciD in the Cba.
limnaeum strain DSMZ 1677T results in loss of the ability to synthesize BChl e.
3.4.2 Purification and Characterization of BciD-His6
To clarify the role of BciD in the synthesis of BChl e, BciD from Cba. limnaeum
T strain DSMZ 1677 was overproduced as a C-terminally His6-tagged protein in E. coli. 101 The isc operon from plasmid pDB1282 was co-expressed with bciD to promote [Fe-S]
cluster assembly (Lanz et al., 2012). Protein purification was monitored by SDS-PAGE
(Figure 3.5). After induction, cells contained an abundant protein with an apparent mass
of about 44 kDa (Figure 3.5, lane 1), which is similar to the expected mass of BciD-His6
(47.3 kDa). Cells were lysed under anoxic conditions, and the lysate was subjected to
ultracentrifugation to separate the soluble fraction from unbroken cells, membranes, and
other debris. Although some of the protein formed inclusion bodies (Figure 3.5, lane 2),
a sufficient amount of the protein of interest was observed in the soluble fraction (Figure
3.5, lane 3) to justify continuing with the purification. The protein was purified by nickel
affinity chromatography under strictly anoxic conditions. After concentration and buffer
exchange, a relatively pure protein of roughly the expected size was obtained (Figure
3.5, lane 8). This protein was subjected to in-gel trypsin digestion, and the resulting peptides were analyzed by mass spectrometry, and the analysis confirmed that the protein was BciD-His6.
As expected for a radical SAM protein, which by precedent is predicted to contain
at least one [4Fe-4S] cluster, the purified BciD-His6 protein was brown in color. Figure
3.6 shows the UV visible absorption spectrum of the as-isolated protein, which had broad, weak absorbance characteristic of an S → Fe charge transfer band with a maximum at about 400 nm. After reconstitution with iron and sulfide as described under
“Experimental Procedures,” this absorbance band increased substantially (roughly 2- fold), indicating that the Fe/S cluster binding site in the as-isolated protein was not completely occupied. To confirm the presence of the predicted [4Fe-4S] cluster, we 102 measured the EPR spectra of as-isolated and reconstituted BciD. Figure 3.7 shows the difference spectra of protein samples reduced with 5 mM dithionite minus the spectra of the untreated samples (EPR measurements taken by Dr. Bryan Ferlez). In the untreated
(oxidized) samples, no EPR resonances were observed; however, after reduction, as seen in the difference spectra, both the as-isolated and the reconstituted protein showed spectral features with axial symmetry and g-values of 2.038 and 1.945. These spectra correspond to those typical of proteins with a single, reduced [4Fe-4S] cluster as predicted for this putative radical SAM enzyme.
3.4.3 BciD-His6 Activity Assay with BChlide c
Activity assays for BciD-His6 were performed under strictly anoxic conditions at
room temperature. Purified BciD-His6 was incubated with a mixture of methylation
homologs of BChlide c or BChlide d, SAM as a cofactor, and sodium dithionite as
reductant. Control reactions without added BciD-His6 were always prepared in parallel.
The pigments present in the mock control reaction were compared with those present
after incubation with the enzyme by reversed-phase HPLC, and the masses of the
products and reactants were determined by mass spectrometry in parallel reactions.
Figure 3.8A shows the HPLC elution profile of the pigments present in the samples containing BChlide c as the substrate. The dotted line contains a series of peaks that make
up the substrate; the peaks at 22.3, 26.1, 29.0, 32.3, and 35.0 min account for the various
methylation homologs of BChlide c, whereas the peaks at 36.9, 39.6, 40.3, and 43.0 min
represent the methylation homologs of BPheide c, a breakdown product of BChlide c 103 from which the central magnesium atom has been lost. As shown in Figure 3.8B, the
UV-visible spectra of BChl c in methanol and the in-line absorption spectrum of the
major BChlide c substrate peak eluting at 26 min are essentially identical. The solid line
in the HPLC elution profiles in Figure 3.8A shows a series of peaks that are derived from
reaction products and unreacted substrate molecules. The peaks at 12.1, 15.3, 16.6, 18.7,
20.0, and 21.6 min uniquely occur in the elution profile of the products (Figure 3.8A and
Table 3.2). The earlier elution times of the product peaks are consistent with the
increased hydrophilicity expected for BChlide e due to replacement of the methyl group
by the formyl group at the C-7 position. In Figure 3.8C, the UV-visible spectra of the
major 20-min product peak and the 26-min substrate peak are compared. Compared with
the substrate BChlide c, the Qy absorption peak of the product is blue-shifted, and its
Soret peak is red-shifted, as expected for BChlide e. In Figure 3.8D, the spectra of the
20-min product and BChl e in methanol are compared, which confirms that the reaction
product is BChlide e. Mass spectrometry showed that the masses of the BChlide c
substrate peaks increase in 14-Da increments as they become more hydrophobic because of the additional methyl groups of the homologs (Table 3.2). Correspondingly, the product peaks are shifted toward earlier elution times due to greater hydrophilicity, and these compounds had masses that were 14 Da larger than the corresponding substrate
peak, which diminished in intensity during the reaction (Table 3.2). This is consistent
with the results expected if the methyl group of BChlide c was transformed into a formyl
group to yield BChlide e. 104 In a separate analysis by HPLC, SAM was consumed, and 5ʹ-deoxyadenosine was
formed in the complete reaction containing BciD-His6, but these changes were not
observed in a control reaction lacking the enzyme (Figure 3.9). All of these observations
are consistent with the expected functioning of a radical SAM enzyme that converts
BChlide c into BChlide e.
The minor product peak at 15.2 min is particularly noteworthy. This product is
obviously more hydrophilic than the major BChlide e product peak at 20 min. The in-line
absorption spectrum of this product, with maxima at 441 and 666 nm (Figure 3.8E and
Table 3.2), more closely resembled the spectrum of the BChlide c substrate than the
BChlide e product but was nevertheless distinct. The mass of this compound was 2 Da
larger than that of the BChlide e product at 20 min. These properties are consistent with
those expected for 71-hydroxy-BChlide c. These observations strongly imply that BciD is a radical SAM enzyme that acts by catalyzing two consecutive hydroxylation reactions of the C-7 methyl group.
3.4.4 BciD-His6 Activity Assay with BChlide d
A similar activity analysis is shown in Figure 3.10 for the complete reaction and
control with BChlide d as the substrate. Similar to the results shown for BChlide c in
Figure 3.8, the substrate contained methylation homologs of BChlide d and small
amounts of BPheide d (Figure 3.10A), and as expected, the UV-visible spectrum of the major substrate peak at 21.6 min matched that of BChl d in methanol (Figure 3.10B).
The major product peak at 18.9 min differs from BChlide d (Figure 3.10C) but matches 105 the spectrum of BChl f, as expected for the conversion of BChlide d to BChlide f (Figure
3.10D). The main product peak appears at 18.9 min and appears to come from the second most abundant substrate peak at 25.4 min. The shift to an earlier elution time, from 25.4
min for the substrate peak to 18.9 min for the product, as well as the increase in mass by
14 Da, are the expected results if the C-7 methyl group of BChlide d has been transformed into a formyl group to produce BChlide f (Table 3.3). These data establish that BciD is not only necessary for the conversion of BChlide c and d into BChlide e and f, respectively, but it is sufficient to catalyze these transformations. As observed with
BChlide c as substrate, a product peak at 14 min was probably due to 71-hydroxy-
BChlide d. This product was more hydrophilic than the BChlide f products and had an
absorption spectrum with maxima at 433 and 653 nm (Table 3.3 and Figure 3.10E) that
was more similar to BChlide d than to BChlide f. These data strongly imply that BciD-
His6 sequentially hydroxylates the C-7 methyl group twice during the synthesis of
BChlide f.
3.5 Discussion
The goal of this study was to clarify the role of BciD in the biosynthetic pathway of BChl e. Previous studies (Harada et al., 2013) had shown that inactivation of the bciD
gene in Cba. limnaeum prevented the biosynthesis of BChl e, resulting in production of
BChl c in the mutant. However, repeated attempts to express bciD in Cba. tepidum failed
to produce strains capable of BChl e synthesis or related compounds (e.g. 71-hydroxy-
BChl c). Therefore, I first verified the phenotype reported for a bciD mutant of Cba. 106 limnaeum strain RK-j-1 (Harada et al., 2013). The results presented in Figure 3.4 agree
well with those from the previous study and show that inactivation of bciD of Cba.
limnaeum DSMZ 1677T also produced a BChl c-producing mutant strain.
The EPR spectra of recombinant BciD-His6 had features that are consistent with
the presence of a single [4Fe-4S] cluster in the enzyme. In radical SAM enzymes, the
SAM-activating [4Fe-4S] cluster has a binding motif consisting of three cysteines, and
the canonical motif is CXXXCXXC. Figure 3.11 shows a multiple sequence alignment of BciD from four brown-colored, BChl e-producing GSB. These proteins contain five conserved cysteine residues at positions 78, 124, 133, 136, and 261; we propose that the cysteines at positions 124, 133, and 136 are ligands to the observed [4Fe-4S] cluster. The proposed roles of these three cysteine residues in ligating the [4Fe-4S] cluster in BciD could be tested by site-specific mutagenesis in future studies. In the presence of SAM and dithionite, BciD-His6 converted BChlide c and BChlide d to BChlide e and BChlide f,
respectively. Additionally, the formation of more hydrophilic products was observed.
Based upon the elution properties, absorption spectra, and masses of these compounds, we propose that they are methylation homologs of 71-OH BChlide c and 71-OH BChlide d, respectively (Tables 3.2 and 3.3). Finally, BciD consumed SAM as a cofactor and produced 5ʹ-deoxyadenosine as a by-product of the reaction. Taken together, these observations lead to the conclusions that BciD is a radical SAM enzyme with a non- canonical cysteine motif of three cysteine residues that ligates a SAM-activating [4Fe-4S] cluster and that catalyzes the conversion of the C-7 methyl group of BChlide c or d to the 107 formyl group of BChlide e or BChlide f via consecutive hydroxylation reactions (Figure
3.12).
Given that BciD appears to be sufficient to convert BChlide c into BChlide e in
vitro, it is still a mystery why expression of bciD in the BChl c-containing GSB Cba.
tepidum has repeatedly failed to produce BChls containing either hydroxymethyl or
formyl side chains at C-7. One possible explanation is that Cba. limnaeum and Cba.
tepidum have significantly different optimal growth temperatures, and BciD might not be
active at higher temperature. Cba. tepidum is routinely grown at 40–48 °C in the
laboratory, whereas Cba. limnaeum is typically grown at 25–30 °C. However, BciD-His6
activity was still observed in vitro at 40 °C (Figure 3.13), which seemingly excludes this explanation. Alternatively, although it seems unlikely, one could postulate that there is no appropriate electron donor for this reaction in Cba. tepidum. Obviously, sodium dithionite is not the biological electron donor for this reaction in vivo in Cba. limnaeum.
GSB, including Cba. tepidum, contain small, highly abundant soluble ferredoxins that
typically serve this function (Bryant et al., 2012; Seo et al., 2001; Yoon et al., 2001). A
counterargument to this point is that there is no gene encoding a ferredoxin or other
potential electron donor protein encoded in the gene cluster that is specifically found in
brown-colored, BChl e-producing GSB strains. Although it again seems unlikely because
Cba. tepidum produces many Fe/S proteins, it is possible that the [4Fe-4S] cluster of
BciD is unable to assemble, or that BciD is unable to fold properly, in the context of the
Cba. tepidum cytoplasm. 108 An alternative explanation is that there is a biosynthetic problem downstream
from BciD and BChlide e that prevents the accumulation of BChl e in Cba. tepidum.
Specifically, the enzymes responsible for adding the esterifying alcohols to Chls and
BChls show considerable specificity for the (B)Chlide substrate but usually have broader
specificity for the esterifying alcohol (Frigaard et al., 2002; Oster et al., 1997; Saga et al.,
2015; Suzuki et al., 1997). In Cba. tepidum BchK is responsible for the addition of the
farnesyl tail to BChl c, and ChlG and BchG add phytadienol and phytol to Chl a and
BChl a, respectively (Frigaard et al., 2002). A comparative analysis of the genomes of
GSB shows that all have at least three esterifying enzymes: BchK, ChlG, and BchG.
Interestingly, Cba. limnaeum has two paralogs of the bchK gene; one paralog encodes
BchK1, which is very similar to the enzyme found in Cba. tepidum, but paralog BchK2 is
much more divergent (Figure 3.14). Moreover, all brown-colored, BChl e-producing
GSB species contain a BchK enzyme that is phylogenetically distant from the BchK of
Cba. tepidum. These observations suggest that a possible problem in producing BChl e in
Cba. tepidum might result from an incompatibility between the esterifying enzyme
(BchK) in Cba. tepidum and the BChlide e substrate. If this is correct, then it might be
possible to circumvent this problem by expressing the bchK2 gene of Cba. limnaeum
together with bciD in Cba. tepidum. The construction of a brown colored, BChl e- producing strain of Cba. tepidum would be very useful for physiological and biophysical
studies on the specific role(s) of the BChl e in adaptation of GSB to growth at very low light intensities. 109 When assembled into chlorosomes, BChls c, d, and e have distinctive absorption features that are associated with specific light niches in nature (Frigaard and Bryant,
2006; Orf and Blankenship, 2013). In aquatic systems, organisms with chlorosomes containing BChl d typically occur at shallower depths than organisms containing BChl c
(Maresca et al., 2004; Montesinos et al., 1983; van Gemerden and Mas, 1995; Vila and
Abella, 2001). A similar pattern is observed in benthic mat systems in chlorophototrophic mats associated with alkaline siliceous hot springs in Yellowstone National Park (Liu et al., 2012; Tank et al., 2017b). When BChl e assembles into supramolecular aggregates in
chlorosomes, an emergent absorption band appears at about 525 nm (see Figure 3.4) that
allows these organisms to very efficiently harvest blue light (Orf and Blankenship, 2013).
Such organisms are also typically able to grow at greater depths and at remarkably low
light intensities (Manske et al., 2005; Maresca et al., 2004; Marschall et al., 2010;
Montesinos et al., 1983; van Gemerden and Mas, 1995; Vila and Abella, 2001; Vogl et al., 2012). Thus, the ability to synthesize BChl e is associated with the ability to inhabit a light niche that is inaccessible to other chlorophototrophs. Should the bciD gene be
horizontally transferred to a recipient capable of BChl c synthesis, the ability to synthesize BChl e would allow that organism to modify its light-harvesting chlorosomes
and invade niches with lower light intensities and fewer competitors.
Evidence exists for horizontal gene transfer for the entire gene cluster that
includes bciD and cruB (Llorens-Marès et al., 2017). In Lake Cisó in Spain, the resident
population of GSB recently shifted from green to brown, and a metagenomic analysis
showed that the genome of the brown-colored organism was extremely similar to the type 110 strain of Chlorobium luteolum DSM 273T, a green-colored, BChl c-producing strain
(Bryant et al., 2012). Some evidence pointed to a DNA phage as a possible vector for this transfer (Llorens-Marès et al., 2017). Interestingly, Chlorobium luteolum DSM 273T has
two bchK genes, one of which is similar to the bchK gene of Cba. tepidum and one of
which (bchK2) belongs to the clade of enzymes that are found in brown-colored, BChl e-
producing GSB strains (Figure 3.14). The presence of a bchK2 gene, the product of
which can presumably esterify BChlide e with farnesol, might “predispose” Chlorobium
luteolum to transformation from a BChl c-producing into a BChl e-producing strain by
the acquisition of bciD, although the horizontally transferred, conserved gene cluster does
not include a gene for a chlorophyll synthase. These ideas could be tested in future
studies by inactivating the bchK1 and bchK2 genes in Cba. limnaeum as well as by
introducing the bciD gene into Cba. tepidum in combination with each of the two bchK
genes encoding the two paralogous esterification enzymes of Cba. limnaeum.
Questions remain about the detailed mechanism of hydroxylation, the source of
oxygen for this reaction (presumably water), and the structure of the BciD enzyme. BciD
is the first radical SAM enzyme of its subclass to be characterized. A blastp comparison
of BciD against the UCSF structure-function linkage database showed that BciD is most
closely related to uncharacterized subgroup 20 and uncharacterized subgroup 11 of the
radical SAM superfamily (Akiva et al., 2014). A blastp search of the NCBI protein database with the BciD query returned 928 sequences with an e-value <0.001; however,
none of these proteins have been characterized in vitro. The blastp search also showed
that BciD has similarity to the following conserved domains: Elp3, 111 rSAM_MSMEG_0568, Radical_SAM, and BioB. These conserved domains correspond to the superfamily denoted Elongator protein 3/MiaB/radical SAM, which contains several radical SAM enzyme families: the protein family radical SAM MSMEG_0568, which has no characterized function, the radical SAM superfamily, and a domain found in biotin synthase and related enzymes. The absence of obviously close relatives makes the BciD enzyme interesting for future study, not just from the perspective of BChl synthesis, but also from the perspective of radical SAM biochemistry more broadly. 112 References Adams, P. G., Cadby, A. J., Robinson, B., Tsukatani, Y., Tank, M., Wen, J., et al. (2013). Comparison of the physical characteristics of chlorosomes from three different phyla of green phototrophic bacteria. Biochim. Biophys. Acta - Bioenerg. 1827, 1235–1244. doi:10.1016/j.bbabio.2013.07.004. Akiva, E., Brown, S., Almonacid, D. E., Barber, A. E., Custer, A. F., Hicks, M. A., et al. (2014). The structure-function linkage database. Nucleic Acids Res. 42, D521–D530. doi:10.1093/nar/gkt1130. Albus, C. A., Salinas, A., Czarnecki, O., Kahlau, S., Rothbart, M., Thiele, W., et al. (2012). LCAA, a novel factor required for magnesium protoporphyrin monomethylester cyclase accumulation and feedback control of aminolevulinic acid biosynthesis in tobacco. Plant Physiol. 160, 1923–1936. doi:10.1104/pp.112.206045. Bobe, F. W., Pfennig, N., Swanson, K. L., and Smith, K. M. (1990). Red shift of absorption maxima in Chlorobineae through enzymic methylation of their antenna bacteriochlorophylls. Biochemistry 29, 4340–4348. doi:10.1021/bi00470a012. Borrego, C. M., and Garcia-Gil, L. J. (1994). Separation of bacteriochlorophyll homologues from green photosynthetic sulfur bacteria by reversed-phase HPLC. Photosynth. Res. 41, 157–164. doi:10.1007/BF02184156. Borrego, C. M., Gerola, P. D., Miller, M., and Cox, R. P. (1999). Light intensity effects on pigment composition and organization in the green sulfur bacterium Chlorobium tepidum. Photosynth. Res. 59, 159–166. doi:10.1023/A:1006161302838. Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. doi:10.1016/0003-2697(76)90527-3. Broch-due, M., and Ormerod, J. G. (1978). Isolation of a Bchl c mutant from Chlorobium with Bchl d by cultivation at low light intensity. FEMS Microbiol. Lett. 3, 305–308. doi:10.1111/j.1574-6968.1978.tb01953.x. Bryant, D. A., and Frigaard, N. U. (2006). Prokaryotic photosynthesis and phototrophy illuminated. Trends Microbiol. doi:10.1016/j.tim.2006.09.001. Bryant, D. A., Garcia Costas, A. M., Maresca, J. A., Gomez Maqueo Chew, A., Klatt, C. G., Bateson, M. M., et al. (2007). “Candidatus Chloracidobacterium thermophilum”: an aerobic phototrophic Acidobacterium. Science . 317, 523–526. doi:10.1126/science.1143236. Bryant, D. A., and Liu, Z. (2013). Green Bacteria. Insights into Green Bacterial Evolution through Genomic Analyses. Elsevier doi:10.1016/B978-0-12-397923- 0.00004-7. 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. doi:10.1021/bi026599s. Bryant, D. A., Liu, Z., Li, T., Zhao, F., Garcia Costas, A. M., Klatt, C. G., et al. (2012). “Comparative and Functional Genomics of Anoxygenic Green Bacteria from the Taxa Chlorobi, Chloroflexi, and Acidobacteria,” in Advances in Photosynthesis and Respiration, Vol. 35 Functional Genomics and Evolution of Photosynthetic Systems, 113 eds. R. Burnap and W. Vermaas (Dordrecht: Springer Netherlands), 47–102. doi:10.1007/978-94-007-1533-2_3. Chen, G. E., Canniffe, D. P., Martin, E. C., and Neil Hunter, C. (2016). Absence of the cbb3 terminal oxidase reveals an active oxygen-dependent cyclase involved in bacteriochlorophyll biosynthesis in Rhodobacter sphaeroides. J. Bacteriol. 198, 2056–2063. doi:10.1128/JB.00121-16. Chen, G. E., Canniffe, D. P., and Hunter, C. N. (2017). Three classes of oxygen- dependent cyclase involved in chlorophyll and bacteriochlorophyll biosynthesis. Proc. Natl. Acad. Sci. 114, 6280–6285. doi:10.1073/pnas.1701687114. Chen, G. E., Canniffe, D. P., Barnett, S. F. H., Hollingshead, S., Brindley, A. A., Vasilev, C., et al. (2018). Complete enzyme set for chlorophyll biosynthesis in Escherichia coli. Sci. Adv. 4, eaaq1407. doi:10.1126/sciadv.aaq1407. Delli-Bovi, T. A., Spalding, M. D., and Prigge, S. T. (2010). Overexpression of biotin synthase and biotin ligase is required for efficient generation of sulfur-35 labeled biotin in E. coli. BMC Biotechnol. 10, 73. doi:10.1186/1472-6750-10-73. Eisen, J. a, Nelson, K. E., Paulsen, I. T., Heidelberg, J. F., Wu, M., Dodson, R. J., et al. (2002). The complete genome sequence of Chlorobium tepidum TLS, a photosynthetic, anaerobic, green-sulfur bacterium. Proc. Natl. Acad. Sci. U. S. A. 99, 9509–9514. doi:10.1073/pnas.132181499. Frigaard, N. U., and Bryant, D. A. (2001). Chromosomal gene inactivation in the green sulfur bacterium Chlorobium tepidum by natural transformation. Appl Env. Microbiol 67, 2538–2544. doi:10.1128/AEM.67.6.2538. Frigaard, N. U., Voigt, G. D., and Bryant, D. A. (2002). Chlorobium tepidum mutant lacking bacteriochlorophyll c made by inactivation of the bchK gene, encoding bacteriochlorophyll c synthase. J. Bacteriol. 184, 3368–3376. doi:10.1128/JB.184.12.3368. Frigaard, N. U., and Bryant, D. A. (2006). “Chlorosomes: Antenna Organelles in Photosynthetic Green Bacteria,” in Complex Intracellular Structures in Prokaryotes, ed. J. M. Shively (Berlin, Heidelberg: Springer Berlin Heidelberg), 79–114. doi:10.1007/7171_021. Frigaard, N. U., Takaichi, S., Hirota, M., Shimada, K., and Matsuura, K. (1997). Quinones in chlorosomes of green sulfur bacteria and their role in the redox- dependent fluorescence studied in chlorosome-like bacteriochlorophyll c aggregates. Arch. Microbiol. 167, 343–349. doi:10.1007/s002030050453. Gomez Maqueo Chew, A., and Bryant, D. A. (2007). Chlorophyll biosynthesis in bacteria: the origins of structural and functional diversity. Annu. Rev. Microbiol. 61, 113–129. doi:10.1146/annurev.micro.61.080706.093242. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2004). “Identification of BchV, a C-31 hydratase specific for hypermethylated bacteriochlorophyll c in Chlorobium tepidum,” in Photosynthesis: Fundamental Aspects to Global Perspectives, eds. A. van der Est and D. Bruce (Lawrence, KS: Allen Press), 875– 877. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2007). Bacteriochlorophyllide c C-82 and C-121 methyltransferases are essential for 114 adaptation to low light in Chlorobaculum tepidum. J. Bacteriol. 189, 6176–6184. doi:10.1128/JB.00519-07. Harada, J., Mizoguchi, T., Satoh, S., Tsukatani, Y., Yokono, M., Noguchi, M., et al. (2013). Specific gene bciD for C7-methyl oxidation in bacteriochlorophyll e biosynthesis of brown-colored green sulfur bacteria. PLoS One 8, e60026. doi:10.1371/journal.pone.0060026. Harada, J., Mizoguchi, T., Tsukatani, Y., Noguchi, M., and Tamiaki, H. (2012). A seventh bacterial chlorophyll driving a large light-harvesting antenna. Sci. Rep. 2, 671. doi:10.1038/srep00671. Harada, J., Oh-oka, H., Saga, Y., and Tamiaki, H. (2004). “Natural reverse mutations of the inactivated bchU gene in Chlorobium vibrioforme NCIB 8327,” in Photosynthesis: Fundamental Aspects to Global Perspectives, eds. A. van der Est and D. Bruce (Lawrence, KS: Allen Press), 119–120. Harada, J., Saga, Y., Yaeda, Y., Oh-Oka, H., and Tamiaki, H. (2005). In vitro activity of C-20 methyltransferase, BchU, involved in bacteriochlorophyll c biosynthetic pathway in green sulfur bacteria. FEBS Lett. 579, 1983–1987. doi:10.1016/j.febslet.2005.01.087. Harada, J., Teramura, M., Mizoguchi, T., Tsukatani, Y., Yamamoto, K., and Tamiaki, H. (2015). Stereochemical conversion of C3-vinyl group to 1-hydroxyethyl group in bacteriochlorophyll c by the hydratases BchF and BchV: Adaptation of green sulfur bacteria to limited-light environments. Mol. Microbiol. 98, 1184–1198. doi:10.1111/mmi.13208. Ho, M. Y., Gan, F., Shen, G., and Bryant, D. A. (2017). Far-red light photoacclimation (FaRLiP) in Synechococcus sp. PCC 7335. II. Characterization of phycobiliproteins produced during acclimation to far-red light. Photosynth. Res. 131, 187–202. doi:10.1007/s11120-016-0303-5. Hollingshead, S., Kopecná, J., Jackson, P. J., Canniffe, D. P., Davison, P. A., Dickman, M. J., et al. (2012). Conserved chloroplast open-reading frame ycf54 is required for activity of the magnesium protoporphyrin monomethylester oxidative cyclase in Synechocystis PCC 6803. J. Biol. Chem. 287, 27823–27833. doi:10.1074/jbc.M112.352526. Imhoff, J. F. (2003). Phylogenetic taxonomy of the family Chlorobiaceae on the basis of 16S rRNA and fmo (Fenna-Matthews-Olson protein) gene sequences. Int. J. Syst. Evol. Microbiol. 53, 941–951. doi:10.1099/ijs.0.02403-0. Jones, D. T., Taylor, W. R., and Thornton, J. M. (1992). The rapid generation of mutation data matrices from protein sequences. Bioinformatics 8, 275–282. doi:10.1093/bioinformatics/8.3.275. Kumar, S., Stecher, G., and Tamura, K. (2016). MEGA7: Molecular evolutionary genetics analysis version 7.0. molecular biology and evolution. Mol. Biol. Evol. 33, 1870–1874. Lanz, N. D., Grove, T. L., Gogonea, C. B., Lee, K. H., Krebs, C., and Booker, S. J. (2012). RlmN and AtsB as models for the overproduction and characterization of radical SAM proteins. Methods Enzymol. 516, 125–152. doi:10.1016/B978-0-12- 394291-3.00030-7. 115 Liu, Z., and Bryant, D. A. (2012). “Bacteriochlorophyll Biosynthesis and Assembly in Green Chlorophototrophic Bacteria: Theme and Variations,” in Handbook of Porphyrin Science with Applications to Chemistry, Physics, Materials Science, Engineering, Biology and Medicine, Vol 20: Chlorophylls and Related Systems Handbook of Porphyrin Science., ed. R. Kadish, KM and Smith, KM and Guilard (Singapore: World Scientific Publishing Company, Pt Ltd), 107–140. doi 10.1142/8040-vol20 Liu, Z., and Bryant, D. A. (2011). Identification of a gene essential for the first committed step in the biosynthesis of bacteriochlorophyll c. J. Biol. Chem. 286, 22393–22402. doi:10.1074/jbc.M111.249433. Liu, Z., Klatt, C. G., Ludwig, M., Rusch, D. B., Jensen, S. I., Kühl, M., et al. (2012). “Candidatus Thermochlorobacter aerophilum”: An aerobic chlorophotoheterotrophic member of the phylum Chlorobi defined by metagenomics and metatranscriptomics. ISME J. 6, 1869–1882. doi:10.1038/ismej.2012.24. Llorens-Marès, T., Liu, Z., Allen, L. Z., Rusch, D. B., Craig, M. T., Dupont, C. L., et al. (2017). Speciation and ecological success in dimly lit waters: Horizontal gene transfer in a green sulfur bacteria bloom unveiled by metagenomic assembly. ISME J. 11, 201–211. doi:10.1038/ismej.2016.93. Lovenberg, W., Buchanan, B. B., and Rabinowitz, J. C. (1963). Studies on the chemical nature of Clostridial ferredoxin. J. Biol. Chem. 238, 3899–3913. Mallorquí, N., Arellano, J. B., Borrego, C. M., and Garcia-Gil, L. J. (2005). Signature pigments of green sulfur bacteria in lower Pleistocene deposits from the Banyoles lacustrine area (Spain). J. Paleolimnol. 34, 271–280. doi:10.1007/s10933-005-3731- 3. Manske, A. K., Glaeser, J., Kuypers, M. M. M., and Overmann, J. (2005). Physiology and phylogeny of green sulfur bacteria forming a monospecific phototrophic assemblage at a depth of 100 meters in the Black Sea. Appl. Environ. Microbiol. 71, 8049–8060. doi:10.1128/AEM.71.12.8049-8060.2005. Maresca, J. A. (2007). The genetic basis for pigment variation among green sulfur bacteria. Maresca, J. A., Gomez Maqueo Chew, A., Ponsatí, M. R., Frigaard, N. U., Ormerod, J. G., and Bryant, D. A. (2004). The bchU gene of Chlorobium tepidum encodes the C- 20 methyltransferase in bacteriochlorophyll c biosynthesis. J. Bacteriol. 186, 2558– 2566. doi:10.1128/JB.186.9.2558-2566.2004. Maresca, J. A., Romberger, S. P., and Bryant, D. A. (2008). Isorenieratene biosynthesis in green sulfur bacteria requires the cooperative actions of two carotenoid cyclases. J. Bacteriol. 190, 6384–6391. doi:10.1128/JB.00758-08. Marschall, E., Jogler, M., Henßge, U., and Overmann, J. (2010). Large-scale distribution and activity patterns of an extremely low-light-adapted population of green sulfur bacteria in the Black Sea. Environ. Microbiol. 12, 1348–1362. doi:10.1111/j.1462- 2920.2010.02178.x. Montaño, G. a, Bowen, B. P., LaBelle, J. T., Woodbury, N. W., Pizziconi, V. B., and Blankenship, R. E. (2003). Characterization of Chlorobium tepidum chlorosomes: a calculation of bacteriochlorophyll c per chlorosome and oligomer modeling. Biophys. J. 85, 2560–2565. doi:10.1016/S0006-3495(03)74678-5. 116 Montesinos, E., Guerrero, R., Abella, C., and Esteve, I. (1983). Ecology and physiology of the competition for light between Chlorobium limicola and Chlorobium phaeobacteroides in natural habitats. Appl. Environ. Microbiol. 46, 1007–1016. Niedzwiedzki, D. M., Orf, G. S., Tank, M., Vogl, K., Bryant, D. A., and Blankenship, R. E. (2014). Photophysical properties of the excited states of bacteriochlorophyll f in solvents and in chlorosomes. J. Phys. Chem. B 118, 2295–2305. doi:10.1021/jp409495m. Oostergetel, G. T., van Amerongen, H., and Boekema, E. J. (2010). The chlorosome: A prototype for efficient light harvesting in photosynthesis. Photosynth. Res. 104, 245– 255. doi:10.1007/s11120-010-9533-0. Orf, G. S., and Blankenship, R. E. (2013). Chlorosome antenna complexes from green photosynthetic bacteria. Photosynth. Res. 116, 315–331. doi:10.1007/s11120-013- 9869-3. Orf, G. S., Tank, M., Vogl, K., Niedzwiedzki, D. M., Bryant, D. A., and Blankenship, R. E. (2013). Spectroscopic insights into the decreased efficiency of chlorosomes containing bacteriochlorophyll f. Biochim. Biophys. Acta - Bioenerg. 1827, 493–501. doi:10.1016/j.bbabio.2013.01.006. Oster, U., Bauer, C. E., and Rüdiger, W. (1997). Characterization of chlorophyll a and bacteriochlorophyll a synthases by heterologous expression in Escherichia coli. J. Biol. Chem. 272, 9671–9676. doi:10.1074/jbc.272.15.9671. Ouchane, S., Steunou, A. S., Picaud, M., and Astier, C. (2004). Aerobic and anaerobic Mg-protoporphyrin monomethyl ester cyclases in purple bacteria: A strategy adopted to bypass the repressive oxygen control system. J. Biol. Chem. 279, 6385–6394. doi:10.1074/jbc.M309851200. Saga, Y., Hirota, K., Harada, J., and Tamiaki, H. (2015). In vitro enzymatic activities of bacteriochlorophyll a synthase derived from the green sulfur photosynthetic bacterium Chlorobaculum tepidum. Biochemistry 54, 4998–5005. doi:10.1021/acs.biochem.5b00311. Saga, Y., and Tamiaki, H. (2004). Comparison between chlorosomes containing bacteriochlorophyll-c and chlorosomes containing bacteriochlorophyll-d isolated from two substrains of green sulfur photosynthetic bacterium Chlorobium vibrioforme NCIB 8327. J. Photochem. Photobiol. B Biol. 75, 89–97. doi:10.1016/j.jphotobiol.2004.05.012. Scheer, H. (2006). “An overview of chlorophylls and bacteriochlorophylls : Biochemistry, biophysics, functions and applications,” in Advances in Photosynthesis and Respiration, Vol. 25, eds. B. Grimm, R. J. Porra, W. Rudiger, and H. Scheer (Dordrecht, Netherlands: Springer), 1–26. doi: 10.1007/1-4020-4516-6_1 Seo, D., Tomioka, A., Kusumoto, N., Kamo, M., Enami, I., and Sakurai, H. (2001). Purification of ferredoxins and their reaction with purified reaction center complex from the green sulfur bacterium Chlorobium tepidum. Biochim. Biophys. Acta - Bioenerg. 1503, 377–384. doi:10.1016/S0005-2728(00)00245-0. Shen, G., and Bryant, D. A. (1995). Characterization of a Synechococcus sp. strain PCC 7002 mutant lacking Photosystem I: Protein assembly and energy distribution in the absence of the Photosystem I reaction center core complex. Photosynth. Res. 44, 41– 53. doi:10.1007/BF00018295. 117 Simon, R., Priefer, U., and Pühler, A. (1983). A broad host range mobilization system for in vivo genetic engineering: Transposon mutagenesis in Gram negative bacteria. Nat. Biotechnol. 1, 784–791. doi:10.1038/nbt1183-784. Suzuki, J. Y., Bollivar, D. W., and Bauer, C. E. (1997). Genetic analysis of chlorophyll biosynthesis. Annu. Rev. Genet. 31, 61–89. doi:10.1146/annurev.genet.31.1.61. Tanaka, A., Ito, H., Tanaka, R., Tanaka, N. K., Yoshida, K., and Okada, K. (1998). Chlorophyll a oxygenase (CAO) is involved in chlorophyll b formation from chlorophyll a. Proc. Natl. Acad. Sci. 95, 12719–12723. doi:10.1073/pnas.95.21.12719. Tank, M., Liu, Z., Frigaard, N. U., Tomsho, L. P., Schuster, S. C., and Bryant, D. A. (2017a). Complete genome sequence of the photoautotrophic and bacteriochlorophyll e-synthesizing green sulfur bacterium Chlorobaculum limnaeum DSM 1677T. Genome Announc. 5, e00529-17. doi:10.1128/genomeA.00529-17. Tank, M., Thiel, V., Ward, D. M., and Bryant, D. A. (2017b). “A panoply of phototrophs: An Overview of the thermophilic chlorophototrophs of the microbial mats of Alkaline siliceous hot springs in Yellowstone National Park, WY, USA,” in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, ed. P. C. Hallenbeck (Berlin: Springer), 87–137. doi:10.1007/978-3-319-46261-5_3. Teramura, M., Harada, J., Mizoguchi, T., Yamamoto, K., and Tamiaki, H. (2016a). In vitro assays of BciC showing C132-demethoxycarbonylase activity requisite for biosynthesis of chlorosomal chlorophyll pigments. Plant Cell Physiol. 57, 1048– 1057. doi:10.1093/pcp/pcw045. Teramura, M., Harada, J., and Tamiaki, H. (2016b). In vitro stereospecific hydration activities of the 3-vinyl group of chlorophyll derivatives by BchF and BchV enzymes involved in bacteriochlorophyll c biosynthesis of green sulfur bacteria. Photosynth. Res. 130, 33–45. doi:10.1007/s11120-016-0220-7. Tomitani, A., Okada, K., Miyashita, H., Matthijs, H. C. P., Ohno, T., and Tanaka, A. (1999). Chlorophyll b and phycobilins in the common ancestor of cyanobacteria and chloroplasts. Nature 400, 159–162. doi:10.1038/22101. Tsukatani, Y., Harada, J., Mizoguchi, T., and Tamiaki, H. (2013). Bacteriochlorophyll homolog compositions in the bchU mutants of green sulfur bacteria. Photochem. Photobiol. Sci. 12, 2195–2201. doi:10.1039/c3pp50253h. van Gemerden, H., and Mas, J. (1995). “Ecology of phototrophic sulfur bacteria,” in Advances in Photosynthesis and Respiration Vol. 2 Anoxygenic photosynthetic bacteria, eds. R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Dordrecht, Netherlands: Springer), 49–85. Vila, X., and Abella, C. A. (2001). Light-harvesting adaptations of planktonic phototrophic micro-organisms to different light quality conditions. Hydrobiologia 452, 15–30. doi:10.1023/A:1011909330390. Vogl, K., Tank, M., Orf, G. S., Blankenship, R. E., and Bryant, D. A. (2012). Bacteriochlorophyll f: Properties of chlorosomes containing the “forbidden chlorophyll.” Front. Microbiol. 3, 1–12. doi:10.3389/fmicb.2012.00298. Wada, K., Yamaguchi, H., Harada, J., Niimi, K., Osumi, S., Saga, Y., et al. (2006). Crystal structures of BchU, a methyltransferase involved in bacteriochlorophyll c 118 biosynthesis, and its complex with S-adenosylhomocysteine: implications for reaction mechanism. J. Mol. Biol. 360, 839–849. doi:10.1016/j.jmb.2006.05.057. Wahlund, T. M., and Madigan, M. T. (1995). Genetic transfer by conjugation in the thermophilic green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 177, 2583– 2588. Wang, J., Woldring, R. P., Román-Meléndez, G. D., McClain, A. M., Alzua, B. R., and Marsh, E. N. G. (2014). Recent advances in radical SAM enzymology: New structures and mechanisms. ACS Chem. Biol. 9, 1929–1938. doi:10.1021/cb5004674. Yang, J., Kulkarni, K., Manolaridis, I., Zhang, Z., Dodd, R. B., Mas-Droux, C., et al. (2011). Mechanism of isoprenylcysteine carboxyl methylation from the crystal structure of the integral membrane methyltransferase ICMT. Mol. Cell 44, 997–1004. doi:10.1016/j.molcel.2011.10.020. Yoon, K. S., Bobst, C., Hemann, C. F., Hille, R., and Tabita, F. R. (2001). Spectroscopic and functional properties of novel 2[4Fe-4S] cluster-containing ferredoxins from the green sulfur bacterium Chlorobium tepidum. J. Biol. Chem. 276, 44027–44036. doi:10.1074/jbc.M107852200. Zhao, J. D., Warren, P. V, Li, N., Bryant, D. A., and Golbeck, J. H. (1990). Reconstitution of electron transport in Photosystem I with PsaC and PsaD proteins expressed in Escherichia coli. FEBS Lett. 276, 175–180. doi:10.1016/0014- 5793(90)80536-R. Zheng, L., Cash, V. L., Flint, D. H., and Dean, D. R. (1998). Assembly of iron-sulfur clusters: Identification of an iscSUA-hscBA-fdx gene cluster from Azotobacter vinelandii. J. Biol. Chem. 273, 13264–13272. doi:10.1074/jbc.273.21.13264.
119
Figure 3.1 Scheme showing the biosynthetic pathway leading from Chlide a to BChl c. The side chain at C-8 (R-8) can be ethyl, n-propyl, isobutyl, or neo-pentyl. The side chain at C-12 (R12) can be methyl or ethyl. The C-8 ethyl group of 3-vinyl-BChlide d is methylated by BchQ, and the C-12 methyl group of 3-vinyl-BChlide d is methylated by BchR (Gomez Maqueo Chew et al., 2007). The asterisk at C-31indicates a chiral carbon center that can have either R- or S- stereochemistry. For further details, see the Introduction (Bryant et al., 2012; Gomez Maqueo Chew and Bryant, 2007; Liu and Bryant, 2012; Bryant and Liu, 2013). 120
Figure 3.2 Organization of the gene cluster encoding bciD and cruB in brown-colored GSB. Diagram showing the organization of the gene cluster containing bciD in the genomes of five brown-colored, BChl e-producing GSB. Selected, possibly significant genes in the cluster are represented by colored arrows. Lines connect homologous genes between clusters; SDR, short- chain dehydrogenase/reductase; ICMT, isoprenylcysteine methyltransferase. 121
A B
Figure 3.3 Inactivation of bciD in Cba. limnaeum. A. Physical map of the conjugation plasmid pCLON::bciD and scheme showing the product of a single homologous recombination event with Cba. limnaeum chromosomal DNA which leads to inactivation of bciD in positive transconjugants. B. Agarose gel (1%, w/v) showing PCR amplicons derived from primers flanking the 5ʹ end of bciD and within aadA as shown (see Experimental Procedures for additional details). Lane 1, DNA ladder with sizes shown in bp; Lane 2, no amplicon is produced with chromosomal DNA from wild-type Cba. limnaeum as template; Lanes 3 and 4, 1185-bp amplicons produced with chromosomal template DNA from two DNA preparations from the bciD mutant of Cba. limnaeum.
122
Figure 3.4 Absorbance spectra of whole cells and pigment extracts. A, whole-cell absorbance spectra of the wild-type (solid line) and bciD mutant (dotted line) strains of Cba. limnaeum compared with wild-type Cba. tepidum (dashed line). B, absorbance spectra of pigments extracted with 100% methanol from the wild-type (solid line) and bciD mutant (dotted line) strains of Cba. limnaeum compared with wild-type Cba. tepidum (dashed line). To facilitate the comparison, the spectra were normalized at the Qy absorbance maxima for A and at the Soret maxima for B. AU, absorbance units.
123
Figure 3.5 SDS-PAGE analysis of fractions from the purification of recombinant BciD-His6. Lane 1, whole-cell extract of the BciD-His6 expression strain (run on a separate gel); lane 2, pellet after low speed centrifugation; lane 3, crude supernatant after low speed centrifugation to remove membranes and debris; lane 4, flow-through fraction from nickel chelation chromatography; lane 5, column wash, fraction 1; lane 6, column wash, fraction 2; lane 7, column wash, fraction 3; lane 8, purified BciD-His6; lane 9, protein mass markers. BciD-His6 has an apparent mass of about 44 kDa
124
Figure 3.6 UV-visible absorbance spectra of as-isolated and reconstituted recombinant BciD-His6. The spectra for as-isolated (solid line) and reconstituted BciD-His6 are shown. To facilitate comparison, the spectra were normalized at the UV absorbance maxima. AU, absorbance units.
125
Figure 3.7 EPR spectra of as-isolated and reconstituted recombinant BciD-His6. EPR spectra of as-isolated (solid line) and reconstituted recombinant BciD-His6 (dotted line). Each spectrum represents the difference spectrum between the reduced sample and the untreated (oxidized) sample. The samples were reduced by the addition of 5 mM sodium dithionite. The concentration of reconstituted BciD was 25 μM, and the concentration of as-isolated BciD was 55 μM. Spectrometer conditions are provided under “Experimental Procedures.” The g-values of the axial signal are indicated above the spectra. AU, absorbance units; mT, millitesla.
126
Figure 3.8 Reversed-phase HPLC analysis of BciD reaction with BChlide c. A, reversed- phase HPLC elution profile of BciD control reaction (dotted line) and the 1-h reaction (solid line) using BChlide c as the substrate. Masses of selected peaks in Da are indicated above the peaks. The numbered ranges indicated by brackets in A indicate the ranges at which 7–1-OH-BChlide c (range 1), BChlide e (range 2), BChlide c (range 3), and BPheide c (range 4) methylation homologs elute (also see Table 3.2). The arrows indicate the conversion of [Pr,Et]-BChlide c into 71-OH-BChlide c and then to BChlide e. B, comparison of the absorbance spectra of the 26.1-min substrate peak in A (dotted line) with BChl c in methanol (dashed line). C, comparison of the absorbance spectra of the 26.1-min substrate peak in A (dotted line) and the 20.0-min BChlide e product peak (solid line).D, comparison of the absorbance spectra of the 20.0-min product peak (solid line) and BChl e in methanol (dashed-dotted line). E, comparison of the 26.1-min substrate peak (dotted line) with the 15.3-min product peak (long-dashed line). For identities of compounds and additional information, see Table 3.2. 127
Figure 3.9 Reversed-phase HPLC analysis of 5-deoxyadenosine produced by BciD during reaction with BChlide c. Shown is a reversed-phase HPLC elution profile of SAM standard (dashed line), 5ʹ-deoxyadenosine standard (long-dashed line), 1-h control reaction with BChlide c (dotted line), and 1-h complete reaction with BciD (solid line) using BChlide c as the substrate. SAM (4.5 min) is consumed, and 5ʹ-deoxyadenosine (17.8 min) is produced during the reaction
128
Figure 3.10 Reversed-phase HPLC analysis of BciD reactions with BChlide d. A, reversed- phase HPLC elution profile of control reaction (dotted line) and the 1-h complete reaction (solid line) using BChlide d as the substrate for BciD. The numbered ranges indicated by brackets in A indicate the ranges at which 71-OH-BChlide d (range 1), BChlide f (range 2), BChlide d (range 3), and BPheide d (range 4) methylation homologs elute (also see Table 3.3). The arrow indicates the conversion of [Pr,Et]-BChlide d into [Pr,Et]-BChlide f. Masses of selected peaks in Da are indicated above the peaks. B, comparison of the absorbance spectra of the 21.6-min substrate peak in A (dotted line) with BChl d in methanol (dashed line). C, comparison of the absorbance spectra of the 21.6-min substrate peak in A (dashed line) and the 18.9-min product peak (solid line). D, comparison of the absorbance spectra of the 18.9-min product peak (solid line) and BChl f in methanol (dashed-dotted line). E, comparison of the absorption spectrum of the product peak at 16.5 min (71-OH-BChlide d) with the absorption spectrum of the substrate peak at 21.6 min ([Et,Et]-BChlide d). See Table 3.3 for identities of compounds and additional information. 129
Figure 3.11 Sequence alignment of BciD proteins. Sequence alignment of BciD proteins from four brown-colored, BChl e-producing GSB. The three conserved cysteines highlighted in grey at positions 124, 133, and 136 represent the proposed ligands of the non-canonical CXXXXXXXXCXXC binding motif for a [4Fe-4S] cluster. In the consensus alignment, stars (*) indicate identical residues; colons (:) represent conservative replacements between groups of residues with strongly similar properties; and periods (.) represent conservation between groups of residues with weakly similar properties.
130
Figure 3.12 Proposed reaction scheme for BciD and BchK to convert BChlide c into BChl e. BciD is proposed to act by sequential hydroxylation of the C-7 methyl group, producing 71- monohydroxy and 71-dihydroxy derivatives of BChlide c as intermediates. Spontaneous dehydration of the geminaldiol intermediate produces the 7-formyl group of BChlide e. BChlide e is esterified in vivo by BchK to produce BChl e.
131
Figure 3.13 Reversed Phase HPLC Elution Profile of 40C Experiment. Reversed-phase HPLC elution profile of BciD reaction at RT at start of the reaction (dotted line), the 2-h RT reaction (black line), and the 2-h 40 °C reaction (grey line) using BChlide c as the substrate.
132
133 Figure 3.14 Phylogenetic tree showing the relationship among chlorophyll a synthases (ChlG), bacteriochlorophyll a synthases (BchG) and bacteriochlorophyll c/d/e synthases (BchK) with selected prenyl transferases (MenA) as the outgroup. The sequences from fourteen completely sequenced GSB are included. Each GSB has one ChlG and one BchG enzyme, but there are two distinctive BchK paralogs designated BchK1 and BchK2. Some GSB strains have one paralog while others have two. All brown-colored GSB strains have at least one BchK2 paralog. For additional details, see the main text. Phylogenetic relationships were inferred by the Maximum Likelihood method based on the JTT matrix-based model (Jones et al., 1992). The green shading indicates strains that synthesize BChl c or BChl d, and the brown shading indicates strains that synthesize BChl e.
134 Table 3.1 Oligonucleotide primers used in this study.
Primer Name Sequence (5′ to 3′)1
CLbciDconSphIF ATCGATCGGCATGCTTAACAAACCACCAGCCTCCG
CLbciDconSalIR ATCGATCGGTCGACGTTCGAGCCCCATCAGGAT
CLbciDtestF TAAAGGCGCGACTGCTTTCC
aadAtestR ATCACTGTGTGGCTTCAGGC
CLbciDNdeIF GCTATACATATGAGCACAAAAAGGGTT
CLbciDEcoRIR GCTATAGAATTCCTAACAGACGGGGT
CLbciDstopLeuF GAGCTCGAATTCAAACAGACGGGGTAGGCTTC
CLbciDstopLeuR GAAGCCTACCCCGTCTGTTTGAATTCGAGCTC
1 Introduced restriction sites appear in bold and are underlined.
135 Table 3.2 Summary of properties and assignments of peaks in BciD reaction with BChlide c (see Figure 3.8).1
Control reaction, BChlide c Complete reaction, BChlide c
Peak Retentio Absorbanc Mas Proposed Retentio Absorbanc Mas Proposed numbe n time e peaks s pigment n time e peaks s pigment r (min) (nm) (Da) assignmen (min) (nm) (Da) assignmen t 1 t 7 -OH- 1 12.1 440, 666 ND BChlide c 1 2 15.3 441, 666 632 7 -OH-[Pr, Et]-
3 16.6 475, 660 ND BChlide e 1 7 -OH-[Ib, 4 18.7 441, 666 646 Et]- 5 20.0 475, 660 630 [Pr, Et] - BChlide e 6 21.6 442, 474, ND mixed peak 7 22.3 437, 671 602 [Et, Et]- 22.5 437, 671 602 [Et, Et]- BChlide c BChlide c 8 26.1 437, 671 616 [Pr, Et]- 26.3 437, 671 616 [Pr, Et]- BChlide c BChlide c 9 29 437, 671 616, BChlide c 29.2 437, 671 616, BChlide c 630 630 10 32.3 437, 671 630 [Ib, Et]- 32.5 437, 671 630 [Ib, Et]- BChlide c BChlide c 11 32.8 413, 670 580 [Et, Et]- 32.9 413, 670 580 [Et, Et]- BPheide c BPheide c 12 35 438, 671 644 [Np, Et]- 35.1 438, 671 644 [Np, Et]- BChlide c BChlide c 13 36.9 414, 670 594 [Pr, Et]- 37.0 414, 670 594 [Pr, Et]- BPheide c BPheide c 14 39.6 414, 670 594 BPheide c 39.7 414, 670 594 BPheide c 15 40.3 414, 670 608 BPheide c 40.4 413, 670 608 BPheide c 16 43 414, 670 608 [Ib, Et]- 43.1 414, 670 608 [Ib, Et]- BPheide c BPheide c 1Abbreviations: ND, Not determined; Me, Methyl; Et, ethyl, Pr, n-propyl; Ib, Isobutyl,
Np, neopentyl. 136 Table 3.3 Summary of properties and assignments of peaks in BciD reaction with BChlide d (see Figure 3.10).1
Control reaction, BChlide d Complete reaction, BChlide d
Retention Proposed Retention 2 Proposed Peak Absorbance Mass Absorbance Mass time pigment time pigment number peaks (nm) (Da) peaks (nm) (Da) (min) assignment (min) assignment 1 1 11.4 433, 653 ND 7 -OH- BChlide d 1 2 14.1 433, 653 ND 7 -OH- BChlide d 3 15.5 465, 646 ND [Et, Et]- BChlide f 1 4 16.5 433, 653 ND 7 -OH- BChlide d 5 18.0 430, 659 574 [Me, Et]- 18 430, 659 (574) [Me, Et]- BChlide d BChlide d 6 18.9 465, 645 616 [Pr, Et]- BChlide f 7 21.6 429, 658 588 [Et, Et]- 21.7 430, 658 588 [Et, Et]- BChlide d BChlide d 8 25.4 430, 658 602 [Pr,Et]- 25.5 430, 659 (602) [Pr, Et]- BChlide d BChlide d 9 26.9 430, 658 602 BChlide d 27 430, 658 (602) BChlide d 10 30.2 430, 659 616 [Ib,Et]- 30.2 430, 659 616 [Ib, Et]- BChlide d BChlide d 11 31.5 408, 659 566 [Et, Et]- 31.5 408, 659 566 [Et, Et]- BPheide d BPheide d 12 35.6 408, 660 580 [Pr, Et]- 35.6 408, 660 (580) [Pr, Et]- BPheide d BPheide d 1Abbreviations: ND, Not determined; Me, Methyl; Et, ethyl, Pr, n-propyl; Ib, Isobutyl,
Np, neopentyl.
137 Chapter 4 Bioinformatic Insights into (Bacterio)chlorophyll Biosynthesis in Green
Bacteria
Publication: Jennifer L. Thweatt, Daniel P. Canniffe, and Donald A. Bryant (2019)
Biosynthesis of chlorophylls and bacteriochlorophylls in green bacteria. Advances in
Botanical Research. 90, 35-89.
Contributions: JLT performed the analyses in this study, made all figures and wrote this chapter, some interpretations are discussed in review of chlorophyll and bacteriochlorophyll biosynthesis in green bacteria (see Chapter 2). DAB provided assistance with interpretation of results and editing of this work.
138 4.1 Abstract
Green bacteria comprise a diverse group of anoxygenic phototrophic organisms
from the phyla Chlorobi, Chloroflexi, and Acidobacteria that use chlorosomes for light
harvesting. All green bacteria can produce BChl a and either BChl c., d, e, or f, and the green Chlorobi and green Acidobacteria additionally produce Chl a. The biosynthetic pathways for (B)Chl biosynthesis in green bacteria have been nearly fully elucidated over the past two decades. The ever-increasing number of available genomes of green bacteria has made it possible to analyze the biosynthetic capabilities of new organisms and to examine the evolution of (B)Chl biosynthesis in green bacteria. In the process of writing a review of (B)Chl biosynthesis, it became necessary to perform bioinformatic analyses to identify (B)Chl biosynthetic genes in newly available genomes and to confirm or question prior annotations. In this work, special attention is paid to the enzymes involved in the following reactions: the anoxic coproporphyrinogen III oxidative decarboxylation, protoporphyrinogen (Protogen) IX oxidation, Proto IX magnesium chelation; C13 propionate methylation of Mg-Proto IX, oxidative ring cyclization of Mg-Proto IX 13- monomethyl ester to form the isocyclic E-ring of the macrocyle, C3 vinyl hydration, C8 and C12 methylation, and transesterification of (B)Chlides with alcohol pyrophosphates
139 4.2 Introduction
Green bacteria are a diverse group of bacteria which comprise some members of
the Chloroflexi and Chlorobi and the Acidobacterium, Cab. thermophilum. Green
bacteria share the common trait that they use chlorosomes for light-harvesting during phototrophic growth (see Chapter 1). Chlorosomes are made up of bacteriochlorophyll
(BChl) c, d, e, or f molecules that are self-assembled into nanotubular aggregate structures enveloped by a lipid monolayer. The baseplate of chlorosomes contains BChl a. Additionally, the type-1 reaction centers (RCs) of Cab. thermophilum and green
Chlorobi contain Chl a, BChl a and Zn-BChl a′; while the type-2 RCs of phototrophic
Chloroflexi contain BChl a and derivatives (see Chapter 1). The biosynthetic pathways
of all (B)Chls branch form a common pathway for tetrapyrrole biosynthesis, specifically
branching from heme biosynthesis at the insertion of Mg++ into protoporphyrin (Proto) IX
to form Mg-Proto IX. The biosynthetic pathways of Chl a, BChl a, and BChl c, d, e, or f
are shared from Mg-Proto IX to a branch point at chlorophyllide (Chlide) a where the
three diverge (see Chapter 2). Many of the genes encoding enzymes involved in (B)Chl
biosynthesis in green bacteria have been annotated based on comparison of available
genomes to biochemically characterized enzymes which were initially studied in purple
phototrophic bacteria and cyanobacteria (see Chapter 2). The model green-colored green
sulfur bacterium (GSB) Cba. tepidum has been used to confirm gene annotations based
on these enzymes and to identify genes specific to the BChl c, d, e, or f biosynthetic
pathway (Bröcker et al., 2008; Frigaard et al., 2002; Gomez Maqueo Chew and Bryant
2007; Gomez Maqueo Chew et al., 2009; 2008; 2007; 2004; Harada et al., 2005; Johnson 140 and Schmidt-Dannert, 2008; Liu and Bryant 2011a; Liu and Bryant 2011b; Maresca et
al., 2004; Saga et al., 2015). While the final step in the biosynthesis of BChlide e or f was
identified in the model brown-colored GSB, Cba. limnaeum (Harada et al., 2013; Thweatt
et al. 2017; see Chapter 3). The current state of knowledge of the biosynthetic pathways
of (B)Chls in green bacteria is thoroughly reviewed in Chapter 2. This chapter discusses
the bioinformatic analyses that were used to verify or question previous genome
annotations, search for genes encoding missing enzymes, and examine the phylogenetic
relationships for selected sequences.
The availability of fully sequenced genomes of green bacteria has made
bioinformatic analyses of (B)Chl biosynthesis in these organisms possible. Previous
reviews on this material have also taken advantage of available green bacterial genomes
to investigated (B)Chl biosynthesis (Bryant and Liu, 2013; Bryant et al., 2012; Liu and
Bryant, 2012). The diversity of green bacteria, availability of several genomes, and their ability to synthesize multiple types of (B)Chl make green bacteria a unique group for understanding the evolution of (B)Chl biosynthesis. In addition to reviews and autoannotations, genome papers which specifically discuss the (B)Chl biosynthetic enzymes encoded in green bacterial genomes or metagenomes are available for Cba. tepidum, Ca. T. aerophilum, Cfl. aurantiacus Cab. thermophilum, and Osc. trichoides
(Eisen et al., 2002; Frigaard et al., 2003; Garcia Costas et al., 2012; Grouzdev et al.,
2015; Liu et al., 2012; Tang et al., 2011). In the current work complete or nearly
complete permanent draft genomes or metagenome-assembled genomes (MAGs) are
analyzed for 17 Chlorobia, 7 green FAPs, and Cab. thermophilum (Table 4.1). Two 141 additional metagenome assemblies of Ca. T. aerophilum strains were used to investigate
missing genes in this Candidatus strain (Table 4.6). Since the last major reviews and phylogenetic analyses of green bacterial (B)Chl biosynthesis in 2012 and 2013, draft genomes have been completed or finalized for three Chlorobi and Osc. trichoides, and
MAGs have become available for two new assemblies of Ca. T. aerophilum and for two
green FAPs, Ca. V. mediisalina and Ca. C. asiatica. The work in this chapter adds to our
understanding of (B)Chl biosynthesis in green bacteria by incorporating these new
sequences and exploring select enzymes in depth. These selected enzymes will be
introduced below and are discussed in the context of the full (B)Chl biosynthetic pathway
in Chapter 2.
The enzymes involved in eight different reaction types are given special attention
in this chapter. They are as follows: 1) the anoxic coproporphyrinogen III oxidative
decarboxylation; 2) protoporphyrinogen (Protogen) IX oxidation; 3) Proto IX magnesium
chelation; 4) C13 propionate methylation of Mg-Proto IX; 5) oxidative ring cyclization of
Mg-Proto IX 13-monomethyl ester to form the isocyclic E-ring of the macrocyle; 6) C3
vinyl hydration; 7) C8 and C12 methylation; 8) transesterification of (B)Chlides with
alcohol pyrophosphates. The first two reactions listed here function in heme biosynthesis
as well as (B)Chl biosynthesis and may be carried out by different isofunctional enzymes
under oxic or anoxic conditions.
Oxidative decarboxylation of two of the propionate groups of
coproporphyrinogen III to form the two vinyl groups of protoporphyrinogen IX occurs
sequentially, first to the group on the A ring and then to the group on the B ring (see 142 Chapter 2) (Dailey et al., 2017). Under anoxic conditions this reaction is carried out by coproporphyrinogen III dehydrogenase, HemN, while under oxic conditions it is carried out by coproporphyrinogen decarboxylase, HemF (see Chapter 2). HemN is a radical-
SAM enzyme which binds a 4Fe-4S cluster and two molecules of SAM (Layer et al.,
2005). The 4Fe-4S cluster is ligated by a 3-cysteine motif and one of the SAM molecules.
Catalysis involves formation of a 5′-deoxyandenosyl radical which then removes a
hydrogen from the β carbon of the propionate group, resulting in a carbon radical at that
position. A vinyl group is then formed with the radical electron moving to an unknown
electron acceptor and the loss of CO2 (Layer et al., 2005, 2006). HemN is of interest to
this study for two reasons: firstly, most green bacteria contain multiple paralogs of
HemN, and secondly, a homolog of HemN called HemW has recently been characterized
as a heme chaperone protein (Abicht et al., 2012; Eisen et al., 2002; Tang et al., 2011;
Haskamp et al., 2018). The second reaction of interest is the six-electron oxidation of
Protogen IX to form Proto IX. This reaction can be performed by three different isofunctional enzymes. Two of these are oxygen-independent enzymes which are referred to as Protogen dehydrogenases (HemG and HemJ), and the third has been characterized as an oxygen-dependent enzyme referred to as Protogen oxidase (HemY) (Dailey et al.,
2017; Kato et al., 2010; Kobayashi et al., 2014; Porra et al., 1996; Sasarman et al., 1993;
Skotnicová et al., 2018) (see Chapter 2). HemY is the Protogen oxidation enzyme of interest in this study because genes annotated as hemY have been found in both oxygen- tolerant and strictly anaerobic green bacteria which lack hemG and hemJ (Eisen et al.,
2002; Tang et al., 2011; Garcia Costas et al., 2012). The full oxidation of Protogen IX to
Proto IX by HemY uses three molecules of O2 and produces three H2O2 molecules 143 (Dailey and Dailey, 1996). HemY binds one FAD molecule and so the full reaction
requires three sequential two-electron oxidations (Koch et al., 2004). However, the exact
mechanism is not currently known (Dailey et al., 2017).
The following three reactions of interest are in the portion of the (B)Chl
biosynthetic pathway leading from Proto IX to Chlide a, reactions that are shared by all
(B)Chl biosyntic pathways in green bacteria (see Chapter 2). Proto IX magnesium
chelation is carried out by a three subunit enzyme known as Proto IX magnesium
chelatase, ChlHDI/BchHDI (Willows et al., 1996). Multiple paralogs of the ChlH/BchH
subunit are found in green bacteria (Liu and Bryant, 2012. Therefore, assigning identities
to these paralogs requires phylogenetic analysis in new genome sequences. Genes
encoding known enzymes to carry out the fourth and fifth reactions discussed in this
chapter are missing from Ca. T. aerophilum (Liu et al., 2012). In other organisms, C13
propionate methylation of Mg-Proto IX to form Mg-Proto IX 13-monomethyl ester is carried out by a SAM-dependent O-methyltransferase known as Mg-Proto IX methyltransferase, ChlM/BchM (Bollivar et al., 1994a; Gibson and Hunter, 1994). The methyl group of the resulting monomethyl ester is derived from SAM (Bollivar et al.,
1994a). The formation of the isocyclic-E ring can be catalyzed by two isofuntional enzymes in other green bacteria (Liu and Bryant 2012). These two enzymes are known as anaerobic or aerobic oxidative magnesium-protoporphyrin IX monomethyl ester cyclases,
BchE and AcsF, respectively. Additionally, in other organisms AcsF may act with the additional subunits BciE or Ycf54 (Albus et al., 2012; Chen et al., 2017, 2018; Gough et 144 al, 2000; Hollingshead et al., 2012; Pinta et al., 2002). However; none of these have been
identified in Ca. T. aerophilum (Liu et al., 2012).
The remaining three reactions discussed here occur after the pathway branches
from Chlide a (see Chapter 2). The hydration of the C3 vinyl group to form a C3 hydroxyethyl group occurs in both the BChl a and Bchl c, d, e, or f pathways. Some
green bacteria have multiple paralogs of the C-3 hydratase enzyme, BchF, that catalyzes
this reaction (Liu and Bryant, 2012). As with the ChlH/BchH subunit, phylogentic
analysis is required to assign names to the genes in newly sequenced genomes. The
methylation of C8 and C12 occurs only in the BChl c, d, e, or f biosynthetic pathway of
some green bacteria (Bryant et al., 2012). These reactions are performed by the C8 and
C12 methyltranferase enzymes designated BchQ and BchR, respectively (Gomez
Maqueo Chew et al., 2007). BchQ and BchR are radical-SAM methyltransferases that
belong to the same protein family as BchE and P-methyltransferase, which are
cobalamin-dependent, radical-SAM enzymes (Booker, 2009; Gomez Maqueo Chew et
al., 2007; Huster and Smith 1990; Wang, 2018). Green bacteria have also been found to
encode other enzymes in this family, so additional phylogenetic analyses were used to
examine annotations of bchQ and bchR (Gomez Maqueo Chew et al., 2007). The final
reaction of interest to this study is one of the final steps of (B)Chl biosynthesis and
functions in the biosynthetic pathways of all (B)Chls in green bacteria. This reaction is
the trans-esterification of (B)Chlide with an alcohol pyrophosphate to produce (B)Chl. It is catalyzed by enzymes known as Chl a synthase (ChlG), BChl a synthase (BchG), and
BChl c synthase (BchK) (Bollivar et al., 1994b; Frigaard et al 2002; Gaubier et al., 1995; 145 Oster et al., 1997; Rudiger et al., 1980; Saga et al., 2015). Because these enzymes are
similar and each green bacterium has more than one of type of (B)Chl synthase,
additional phylogentic analysis was required to annotate genes encoding the (B)Chl
synthases in new genome sequences (Liu and Bryant, 2012).
4.3 Experimental Procedures
4.3.1 Genomes and Metagenomes Used in This Work
The genomes used in this chapter were retrieved from the Joint Genome Institute
Integrated Microbial Genomes (JGI-IMG) database while the metagenome assemblies were retrieved from the National Center for Biotechnology Information (NCBI) databases
(Chen et al. 2018). The genomes are as follows (see Table 4.1 for accession numbers):
Brown-colored Chlorobiaceae (6): Cba. limnaeum DSM 1677T, Cba. limnaeum Rk-j-1,
Prosthecochloris sp. BS-1, Chl. clathratiforme DSM5477T, Chl. phaeobacteroides DSM
266, Prosthecochloris sp. CIB2401; Green-colored Chlorobiaceae (9): Cba. tepidum
TLS, Chl. limicola DSM245, Chl. chlorochromatii CaD3, Chl. phaeovibriodes DSM 265,
Ptc. bathyomarina GSB1, Chl. luteolum DSM273, Chl. ferrooxidans DSM 13031, Ptc.
aestuarii DSM 271, Cba. parvum DSM 263; Chp. thalassium ATCC 35110 (1); ‘Ca. T.
aerophilum’* (1); Cab. thermophilum B (1); Deep Branching Chloroflexineae (3);
Osc. trichoides DG-6, ‘Ca. Chloroploca asiatica’ B7–9*, ‘Ca. Viridilinea mediisalina’
Kir15-3F*; Chloroflexus spp. (4): Cfl. aurantiacus J-10-fl, Cfl. sp. Y-400-fl, Cfl. sp. Y-
396-1, Cfl. aggregans DSM 9485. * in this list indicate organisms whose draft genomes
were assembled from metagenomic data. 146 4.3.2 Phylogenetic Tree Building
Multiple sequence alignments were performed using MUSCLE aligner from
EMBL-EBI (Madeira et al., 2019). Molecular phylogenetic analyses were performed using MEGA7 (Kumar et al., 2016). The phyolgenetic relationships were inferred by using the Maximum Likelihood method based on the JTT matrix-based model (Jones et al., 1992). The trees with the highest log-likelihood are shown in each figure and the percentage of trees in which the associated taxa clustered together is shown next to the branches for values above 50%. Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Joining and BioNJ algorithms to a matrix of pairwise distances estimated using a JTT model, and then selecting the topology with superior log-likelihood value. The trees are drawn to scale, with branch lengths measured in the number of substitutions per site. Figure legends specify the number of amino acid sequences, the total number of positions in the final dataset and the treatment of gaps and missing data for the dataset.
4.3.3 Sequence Similarity Networks
Sequence similarity networks (SSN) were generated for selected protein sequences using the Enzyme Function Initiative-Enzyme Similarity Tool (EFI-EST) web tool (Gerlt et al., 2015). SSNs were visualized, analyzed and figures generated using
Cytoscape V3.3.0 (Shannon et al., 2003). Individual figure legends specify the unique parameters used to generate and analyze each SSN.
.
4.4 Results 147 4.4.1 Identification of Genes Encoding (B)Chl Biosynthetic Enzymes in Green
Bacteria
Previous biochemical and bioinformatic analyses have identified many of the
genes required for the (B)Chl biosynthesis in green bacteria (see Chapter 2). In the
process of writing a thorough review of the (B)Chl biosynthetic pathways in green
bacteria it became necessary to search individual genomes and metagenomes to confirm
auto-annotations or identify previously unidentified genes. For this purpose, searches
using blastp or tblastn with a known enzyme query sequence were used to find proteins
of interest and their gene loci. Table 4.1 lists the genomes and metagenomes used
throughout this chapter. Tables 4.2, 4.3, 4.4, and 4.5 list gene identifiers associated with
each protein from the JGI database or protein accession numbers from the NCBI
database. Figure 2.1 in Chapter 2 summarizes the enzymes encoded by different groups
of green bacteria. Unambiguous assignment of the genes encoding HemF, ChlD/BchD,
ChlI/BchI, BciA, BciB, BchNLB. BchXYZ, BchC, BciC, BchU, BchP, and BciD was
based on blast searches and/or previous work; genes encoding the remaining enzymes
required further phylogenetic analysis to assign putative functions in some organisms.
Genes encoding HemF are present in the genomes of Chloroflexus spp., Cab. thermophilum, and Ca. T aerophilum. The strict anaerobes of the GSB, Osc. trichoides,
‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ do not contain hemF genes. All green bacterial genomes contain genes encoding homolog(s) of HemN (Table 4.2), which will be discussed further in the next section. A single copy of the gene encoding ChlD/BchD and
ChlI/BchI was found in the genomes and metagenomes of all green bacteria used in this 148 study. Additionally, all had at least one gene encoding BciA or BciB, with BciA limited
to the Chlorobi. Some Chlorobi have copies of both genes and/or multiple copies of BciB
(Table 4.3) (Liu and Bryant, 2011a). All green bacteria have single copies of the genes
encoding BchNLB, BchXYZ, BchC and BciC (Table 4.3 and 4.4). BchU, is found in all
green bacteria that contain BChl c or BChl e, but Ca. T. aerophilum, which synthesizes
BChl d, lacks a gene encoding BchU (Table 4.4). All green bacteria have at least one gene encoding BchP, while some have additional paralogs of unknown function or that have been characterized as CruI (Table 4.5) (Canniffe et al., 2018). Finally, all brown- colored GSB contain a single copy of the gene encoding BciD (Thweatt et al., 2017; see
Chapter 3).
4.4.2 Decarboxylation of Coproporphyrinogen III
Previous analyses reported that Chloroflexus spp. and Cab. thermophilum genomes contain homologs encoding HemF and two homologs encoding of HemN, while
GSB contain two homologs encoding HemN (Eisen et al., 2002; Garcia Costas et al.,
2012; Tang et al., 2011). In order to determine which enzyme is responsible for this reaction in the anaerobic organisms ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ a tblastn search of their metagenomes was performed using HemF and HemN sequences identified in other green bacteria. As expected for anaerobic organisms, open reading frames encoding HemN sequences were identified but none encoding HemF were identified
(Table 4.2). As stated above the genomes of all green bacteria encode homologs of
hemN. All genomes analyzed in this study encode two homologs of HemN with the
exception of Ca. T. aerophilum which encodes only one. A maximum likelihood tree of 149 the HemN proteins of these green bacteria revealed two distinct clades of HemN
homologs (Figure 4.1). A recent study has shown that some homologs of HemN are
heme-binding proteins that lack dehydrogenase activity, and these homologs have been
renamed HemW (Abicht et al., 2012; Haskamp et al., 2018). Biochemically characterized
HemN enzymes have a conserved catalytic motif CXXXCXXCXC while the
characterized HemW proteins have a conserved CXXXCXXCXF motif where the final
cysteine, which is required for catalysis, is replaced by a phenylalanine (Haskamp et al.,
2018; Layer et al., 2002). Comparing the sequence motifs of green bacterial HemN
homologs to biochemically characterized HemN and HemW, it is apparent that Ca. T.
aerophilum encodes HemW but not HemN and all other green bacterial genomes encode
both HemN and HemW (Figure 4.2).
4.4.3 Oxidation of Protoporphyrinogen IX
In the genomes of both Chloroflexi and Ca. T. aerophilum, genes encoding HemY
are easily identifiable, although it should be noted that the gene in Ca. T. aerophilum was
misannotated as hemG. The literature has conflicting information on genes related to
protoporphyrinogen IX oxidation in GSB. Kato et al. (2010) did not identify any copies
of hemG or hemY in GSB (Kato et al., 2010). However, other more recent analyses found
hemY-related genes in GSB, and hemG in Prosthecochloris (Ptc.) sp. BS-1 (Dailey et al.,
2017; Kobayashi et al., 2014). Additionally, individual GSB genomes have annotated
hemY genes despite their status as strict anaerobes. In order to more fully understand
these annotations, a blastp search was run using the proteins encoded by open reading
frames annotated as hemY in Cba. tepidum and Chloroflexus aggregans. These sequences 150 were combined with those from biochemically characterized HemY enzymes from
Bacillus subtilus, Myxoccocus xanthus, Homo sapiens, Arabadopsis thalinia, Nicotiana
tabacum, and Plasmodium falciparum and used to create an SSN (Figure 4.3).
Subsequently, sequences chosen from the HemY, “HemY”, and phytoene desaturase
clusters were used to create a maximum likelihood tree. Together these analyses showed
that the anaerobic GSB and the microaerophilic Cab. thermophilum, have HemY
homologs that are more closely related to each other than to other HemY homologs.
Surprisingly, the anaerobic Osc. trichoides, ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’
have a HemY homolog or homologs that is more closely related to those of the
facultatively aerobic Chloroflexus spp. and aerobic Ca. T. aerophilum and other
biochemically characterized HemY enzymes including CgoX from B. subtilis (Figure 4.3
and 4.4). Interestingly, the HemY enzyme from P. falciparum also clusters more closely
with the other characterized HemY enzymes than with the enzymes found in Cab.
thermophilum and GSB. However, it is apparent from both the SSN and the maximum
likelihood tree that this enzyme is not closely related to other characterized HemY
enzymes.
4.4.4 Magnesium Chelation
Prior biochemical and bioinformatic analyses have identified the subunits of the
MgCH in most Chlorobi, Chloroflexi, and Cab. thermophilum (Gomez Maqueo Chew et
al., 2009; Johnson and Schmidt-Dannert 2008; Liu and Bryant, 2012). Generally, members of the phylum Chlorobi have three MgCHs, made up of BchHDI, BchSDI, and
BchTDI; Chloroflexi have two MgCHs, BchHDI and BchS/T-DI; and Cab. thermophilum 151 has only BchHDI. In order to assign the subunits of the ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ strains, a tblastn search was performed on the metagenomes, and three genes encoding homologs of BchH were identified in each organism. The protein sequences of all BchH homologs were used to construct a maximum likelihood tree (Figure 4.5). This phylogentic analysis showed that like other members of the phylum Chloroflexi, ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ have two genes encoding BchH and one encoding a
BchS/BchT-type subunit. It can be inferred from this phylogeny that the BchS/BchT and the BchH clades emerged from a duplication event, and the BchS and BchT clades of
Chlorobi then later diverged via a duplication event within the BchS/BchT clade. Initial searches of Ca. T. aerophilum revealed three partial sequences, all of which are found within the BchS/T clade. The three partial sequences can be concatenated to form a single, nearly full-length BchS/T sequence; it is therefore likely that Ca. T. aerophilum contains a single copy of BchS and lacks BchH, similar to Chl. chlorochromatii, which has BchS and BchT but lacks BchH. Finally, the Cab. thermophilum genome encodes one
BchH homolog that belongs to the BchH clade.
4.4.5 C13 Propionate Methylation
A single homolog of the gene encoding the Mg-protoporphyrin IX
methytransferase BchM was found in the genomes of all green bacteria with the
exception of Ca. T. aerophilum (Table 4.3) (Gibson and Hunter, 1994). When the draft
metagenome of ‘Ca. T. aerophilum’ OS was published it was the only metagenome
available for this Candidatus species (Liu et al., 2012). However; two new metagenome
assemblies have become available more recently (Table 4.6) (Roy et al., 2019; Stamps et 152 al., 2014). Previous work found that the metagenome of ‘Ca. T. aerophilum’ OS was missing bchM (Liu et al., 2012). In order to check for the presence of bchM, all available metagenomes for ‘Ca. T. aerophilum’ were subjected to tblastn searches for sequences encoding BchM. All three metagenomes lacked homologs of bchM.
In order to identify other possible candidates for C13 propionate methylation, a list of possible O-methyltransferases was compiled for the Ca. T. aerophilum OS metagenome by searching the NCBI Conserved Domain Database (CDD) with all annotated proteins from the metagenome. From this search 162 hits were found in super families containing O-methyltransferases, and of those hits, 128 had an e-value of less than 1 × 10-5, and from within that group, 58 hits had unique accession numbers (Table
4.7). Of the 58 unique accession numbers, two duplicated sets of sequences were found,
so a total of 56 unique candidates were identified. The 56 unique candidates were then
used to perform a blastP against the NCBI nr database. This resulted in 15 sequences that,
excluding Ca. T. aerophilum, had best hits within Chlorobi; 13 of these had e-values <1 ×
10–60, one had an e-value on the order of 1 × 10–35, and one had an e-value on the order of
1 × 10–16. An additional three sequences had best hits in Chp. thallasium but did not have
hits in other GSB (Table 4.8). Upon a second blastP search of all Chlorobi only one of these three sequence produced a hit other than to Ca. T. aerophilum and Chp. thalassium.
For the purpose of the candidate list, the thirteen sequences with best hits in GSB were considered very unlikely and removed from subsequent analyses, while the three with best hits in Chp. thalassium only were retained, for a total of 41 candidates. Investigation of the metagenome encoding these protein sequences revealed 15 sequences which were 153 within 22 kb of genes related to (B)Chl or heme biosynthesis, and of those, seven
sequences were within 10 kb of genes related to (B)Chl or heme biosynthesis and one
was directly adjacent to the gene encoding HemY. Additionally, 13 of the sequences
were within 10 kb of the edge of a contig or a run of N’s in the middle of a contig. The
candidate genes and these data are summarized in Table 4.8.
4.4.6 Isocyclic E-Ring Formation
Formation of the isocyclic E-ring in (B)Chl biosynthesis may be accomplished by
BchE, AcsF, AcsF with Ycf54 or with BciE (Albus et al., 2012; Chen et al., 2017, 2018;
Gough et al., 2000; Hollingshead et al., 2012; Pinta et al., 2002). The genomes of green
bacteria were searched for sequences encoding all four of these enzymes. No sequences
encoding Ycf54 or BciE were found. As seen in other studies, a gene encoding AcsF was
absent from the strictly anaerobic GSB and Osc. trichoides; additionally, this analysis
showed that ‘Ca. C. asiatica’ also lacked this gene as expected for an anaerobic organism.
Surprisingly, the anaerobe, ‘Ca. V. mediisalina,’ did contain a gene encoding AcsF as did
Cab. thermophilum and the facultatively aerobic or microaerophilic Chloroflexus spp.
(Table 4.3) (Bryant et al., 2012). A copy of the gene encoding BchE was found in all green bacteria with the exception of the metagenomic assembly of ‘Ca. T. aerophilum
OS’ (Table 4.3) (Liu and Bryant, 2012). This metagenome also lacked bchM, and acsF
(see above). As with bchM, newly available metagenomes (see Table 4.6) for ‘Ca. T.
aerophilum’ were searched, and they did not contain genes encoding BchE, AcsF, Ycf54,
or BciE. Blast hits to BchE were not a close match in ‘Ca. T. aerophilum’ and were
instead likely due to BchQ, BchR or one of the other BchE/P-methyltransferase family 154 enzymes found in other green bacteria (Gomez Maqueo Chew et al., 2007). These sequences were analyzed by SSN and maximum likelihood trees to test this idea further.
None of the BchE/P-methyltransferase family enzymes found encoded in the metagenome of ‘Ca. T. aerophilum’ were closely related to BchE in either of these analyses (Figure 4.7, Figure 4.8) (see discussion of methyltransferases below).
4.4.7 Hydration of the C3 vinyl group
Hydration of the C3 vinyl group must occur during the biosynthesis of BChl a and
BChl c, d, e, or f. Previous work has shown that Chloroflexi, and Cab. thermophilum have a single copy of the gene encoding the C3 hydratase known as BchF, while green- colored GSB mostly have two paralogs of the gene and brown-colored GSB mostly have three copies with the second and third copy encoding C3 hydratases known as BchV and
BchF3 respectively (Bryant and Liu, 2013; Eisen et al., 2002; Garcia Costas et al., 2012;
Tang et al., 2011; Tank et al., 2017). As with other Chloroflexi, searches of ‘Ca. V. mediisalina’ and ‘Ca. C. asiatica’ metagenomes determined that each had one copy of the bchF gene, encoding the C3-hydratase. In order to investigate the phylogenetic relationships between BchF, BchV, and BchF3 a maxium likelihood tree was constructed using the sequences for the C3 hydratases from all green bacteria used in the chapter and
BchF encoded by Rhodobacter sphaeroides (Burke et al., 1993). This analysis showed that there is a clear divergence between the BchF clade containing sequences of R. sphaeroides BchF and BchF from all green bacteria and the BchV/F3 clade found only in members of the phylum Chlorobi. Additionally, there is a clade within the BchV/F3 clade which contains all of the C3-hydratase paralogs of brown-colored GSB which are found 155 within the brown-colored GSB gene cluster; this includes the single BchV/F3 C3
hydratase encoded by Prosthecochloris sp. BS-1. Here the sequences in this clade will be
designated as BchF3 and the remaining sequences in the BchV/F3 clade as BchV.
4.4.8 Methylation at C8 and C12
BchQ and BchR have previously been found in Chlorobia, Cab. thermophilum,
and Osc. trichoides, but not in Chloroflexus spp. (Bryant et al., 2012). Additionally, most
brown-colored GSB encode a second paralog of BchQ, denoted BchQ2, just upstream
from a conserved gene cluster found in all brown-colored GSB (see Chapter 3) (Thweatt
et al., 2017). In order to identify homologs of BchQ and BchR in ‘Ca. C. asiatica’ and
‘Ca. V. mediisalina’, green bacterial genomes were searched for homologs of BchR,
BchQ, and BchE. The resulting sequences were then used to create an SSN which incorporated 1/50 of the Interpro class B methyltransferase family IPR034466 family.
Clusters containing more than two of the blast hits from green bacteria are shown in
Figure 4.7. Each of the seven BchE/P-methyltransferase family enzymes encoded by
Cba. tepidum is represented in Figure 4.7 (Gomez Maqueo Chew et al., 2007). Unique sub-clusters were identified containing BchQ, BchR, BchE, and CT1903. The sequences designated CT1502 and CT0072 were found in a shared subcluster and CT1697 formed its own separate cluster. The four sequences from Cba. tepidum which are designated by their locus tag were used to search the Structure Function Linkage Database (SFLD) and
CT1903 was identified as being in the HpnP-like enzyme family with an e-value on the order of 10–213. CT1502 and CT0072 also had closest hits to the HpnP-like family but
with e-values on the order of 10–70 and 10–72 respectively., and CT1697 has a closest 156 match to the BchE family with an e-value on the order of 10–70. Based on these data the
(sub)clusters containing CT1697, CT1502, and CT0072 continued to be designated by the Cba. tepidum locus tags and the sub-cluster containing CT1903 was designated
HpnP-like. HpnP is a C-2 methyltransferase which uses hopanoids as a substrate. The
BchQ, BchR, CT1502/CT0072 and HpnP-like subclusters are all connected to the same main cluster. This main cluster will be referred to as the methyltransferase main cluster.
The sequence corresponding to locus Cabther_B0251 was found in a subcluster which lacked a homolog in Cba. tepidum. Cabther_B0251 was used to search the SFLD and had a best hit to the hopanetetrol cyclitol ether synthase family with and e-value on the order
of of 10–213. Cab. thermophilum has been shown to produce bacteriohopanetetrol cyclitol
ether and so this subcluster was identified as hopanetetrol cyclitol ether synthase (Tank
and Bryant, 2015). Hopanetetrol cyclitol ether synthase is responsible for a ring
rearrangement reaction needed to produce hopanetetrol cyclitol ether. The hopanetetrol
cyclitol ether synthase subcluster is connected to the same main cluster as BchE. This
main cluster will be referred to as the cyclization associated cluster.
As expected, Osc. trichoides, and members of the Chlorobi each had sequence(s)
in the BchQ, BchR, and BchE subclusters, with the exception of Ca. T. aerophilum which
lacks BchE, while the Chloroflexus spp. all had a sequence in the BchE subcluster. The
blast hits from ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ were identified within the SSN.
Both strains were shown to encode BchQ and ‘Ca. V. mediisalina’ also encodes BchR.
‘Ca. V. mediisalina’ also contains two additional sequences within the methyltransferase
main cluster. Finally, each strain has a sequence attached to the cyclization associated 157 main cluster, which is mostly closely clustered with blast his from other members of the phylum Chloroflexi.
Cab. thermophilum had one sequence in the BchE subcluster; however, it did not
have a sequence in the BchQ or BchR subclusters. This was surprising since the
sequences encoded by Cabther_B0082 and Cabther_B0083 were previously annotated as
BchQ and BchR. Cabther_B0083 is found in the methyltransferase main cluster, while
Cabther_B0082 is found within the cyclization associated main cluster (Garcia Costas et
al., 2012). Cabther_B0083 and Cabther_B0082 did not belong as clearly to one family in
the SFLD database as sequences that clustered closely with one of the named subclusters.
Cabther_B0083 had hits in the BchQ, BchR, and HpnP-like families with e-values on the order of 10–75, 10–72, and 10–72, respectively. Cabther_B0082 had its best hit to the BchE
family with an e-value on the order of 10–54. These results agree with the placement of
the nodes representing Cabther_B0082 and Cabther_B0083 in the SSN, indicated by red
arrows. Based on this SSN neither Cabther_B0082 nor Cabther_B0083 could confidently
be assigned to a subcluster.
In order to investigate phylogentic relationships among these sequences, a
maximum likelihood tree was constructed on the basis of the SSN clusters. All sequences
from the BchQ. BchR, and BchE subclusters were used. Sequences from the
CT1502/CT0072 subcluster were chosen from Ptc. aestuari, Cba. tepdium, Chl. limicola,
Chp. thalassium, and Ca. T. aerophilum. Additionally, the sequences associated with
Cabther_B0082, Cabther_B0083 and the two ‘Ca. V. mediisalina’ sequences found
within the methyltransferase main cluster were included. From this phylogenetic tree it 158 appears that the two ‘Ca. V. mediisalina’ sequences found within the methyltransferase main cluster are most closely related to the CT1502/CT0072 sequences, while
Cabther_B0083 is the most distantly related sequence in the clade containing BchQ,
BchR, and CT1502/CT0072. The Cabther_B0082 sequence, like in the SSN, is more closely related to BchE than to the clade containing BchQ, BchR, and CT1502/CT0072.
4.4.9 Esterification of (B)Chlide
The sequences of (B)Chl synthases in green bacteria were compiled and used to make a maximum likelihood tree to investigate their phylogenetic relationships (Table
4.4 and Figure 4.9). As expected, this phylogenetic analysis confirmed that all green bacteria encoded BchG and BchK while Cab. thermophilum and members of the
Chlorobi, which make Chl a, additionally have ChlG. It also showed that like other
Chloroflexi, ‘Ca. C. asiatica’ and ‘Ca. V. mediisalina’ each had homologs of BchG and
BchK. Additionally, both ‘Ca. V. mediisalina’ and Osc. trichoides each had two copies of
BchK. Comparison of the clades containing BchG, ChlG and BchK show that the clade containing all (B)Chl synthases branches from a clade containing other prenyltransferase enzymes. It also shows that the bacteriochlorin synthase clade, BchG, branches from within a preexisting chlorin synthase clade. Finally, this analysis showed that the BchK sequences of Chloroflexi and Cab. thermophilum form separate clades, which are not associated with the BchK1 and BchK2 clades previously designated in GSB (see
Chapter 3).
4.5 Discussion 159 4.5.1 Early Steps
When the genomes of Cab. thermophilum and Cfl. aurantiacus were first sequenced and found to have multiple paralogs of hemN, it was assumed that both encoded active HemN enzymes. This led to hypotheses that the two copies of HemN might be differentially regulated based on environmental factors like light or oxygen tension in GSB and Chloroflexi, respectively (Eisen et al., 2002; Tang et al., 2011).
Characterized HemN enzymes possess a CXXXCXXCXC motif which is required for catalysis (Layer et al., 2005). The first three cysteines in this motif bind a radical-SAM
4Fe-4S cluster, while the fourth cysteine is not required for binding of this cluster but is required for catalysis (Layer et al., 2002). In contrast, characterized HemW heme binding proteins have a CXXXCXXCXF motif that also binds a radical-SAM 4Fe-4S cluster but does not catalyze the decarboxylation of coproporphyrinogen III (Abicht et al., 2012;
Haskamp et al., 2018). Analysis of the HemN homologs encoded by green bacteria clearly show that all green bacteria contain one HemW homolog and all except Ca. T. aerophilum have one HemN homolog (Figure 4.1, 4.2, Table 4.2) The absence of HemN in Ca. T. aerophilum, while unique among green bacteria is not surprising for an organism that grows aerobically and possesses HemF. Because HemW has been characterized as a heme binding protein, it is possible, that it plays a role in heme and
(B)Chl biosynthesis in green bacteria, or as a heme chaperone for insertion of heme cofactors as has been suggested for other organisms (Haskamp et al., 2018).
Characterization of protoporphrinogen oxidase enzymes has shown that HemY uses molecular oxygen as the electron acceptor while HemG uses acceptors associated 160 with the electron transport chain; ubiquinone and menaquinone have been used as
electron acceptors various electron acceptors including ubiquinone and menaquinone in
vitro (Boynton et al., 2009; Daily and Daily, 1996a; 1996b; Daily et al., 2017; Mobius et
al., 2010; Sasarman et al., 1993). As a result, characterization of HemY enzymes has
been performed under oxic conditions, with the exception of the enzyme from P.
falciparum, which was only shown to be active under anoxic conditions. This work showed that the HemY enzyme from P. falciparum could use FAD, NAD+, and NADP+
as electron acceptors in vitro and that in vivo, it was dependent on the electron transport
chain (Nagaraj et al., 2010). Previous work has annotated HemY-encoding genes in all
classes of green bacteria (Eisen et al., 2002; Frigaard et al., 2006; Garcia Costas et al.,
2012; Kobayashi et al., 2014; Tang et al., 2011). This analysis found that all green
bacteria had a gene related to either the hemY gene annotated in Cfl. aggregans or the one
in Cba. tepidum (Figure 4.3, 4.4, Table 4.2). Because GSB and the deeply branching
Chloroflexineae are strict anaerobes, the annotation of an enzyme expected to require molecular oxygen as an electron acceptor and conflicting results in the literature concerning HemY in GSB promted further analyses in this study (Kato et al., 2010;
Kobayashi et al., 2014)
A previous study had produced phylogenetic analyses which showed that the non-
canonical HemY from Plasmodium falciparum mitochondria is in the same clade as
“HemY” from some GSB (Kobayashi et al., 2014). However, the analyses shown here did not replicate that observation (Figure 4.4). A HemY SSN in fact showed that the
Plasmodium falciparum mitochondrial HemY did not cluster with those from GSB 161 (Figure 4.3). Even more perplexing is the fact that all of the deeply branching
Chloroflexineae have a HemY sequence which is closely related to those of other
Chloroflexi and clusters with HemY enzymes characterized as requiring molecular
oxygen in other organisms. Interestingly, Osc. trichoides and Ca. C. asiatica both have a
second HemY sequence which appears to be more phylogenetically distant from those
characterized as requiring molecular oxygen (Table 4.2, Figure 4.3, 4.4). It is also
interesting to note that in addition to a HemY sequence closely related to that of other
Chloroflexi, Ca. V. mediisalina also has a gene encoding the oxygen requiring enzyme
AcsF. In both the SSN and phylogenetic tree of HemY related sequences it is clear that
GSB and Cab. thermophilum HemY sequences form a distinct clade and cluster distinct
from other HemY sequences. Based on this and the well-characterized, strictly anaerobic
growth of GSB, it is very likely that these sequences, denoted here as “HemY,” do not
function in the same way as canonical HemY enzymes. They are likely either a HemY- like enzyme which is capable of using other electron acceptors, as was seen for P.
falciparum or they are not protoporphrinogen oxidase enzymes and an as yet unidentified
enzyme performs this function in GSB and Cab. thermophilum. A similar case can be
made for the second sequence found in Osc. trichoides and Ca. C. asiatica. Athough they
do not form a clearly separate cluster or clade in these analyses, it is apparent that they
are not closely related to other Chloroflexi HemY sequences. Finally, the characterization
of P. falciparum HemY demonstrates that a HemY-like enzyme that does not require
molecular oxygen does exist; however, its lack of a close relationship with the enzymes
encoded by green bacteria leaves it an open question as to whether the deeply branching
Chloroflexineae, GSB, and Cab. thermophilum use such an enzyme. Future research 162 either to biochemically characterize the HemY proteins encoded by green bacteria or to genetically complement known hemY inactivation strains with “hemY” could help to shed light on the function of these sequences and protoporphyrinogen oxidation in green bacteria.
4.5.2 Proto IX to Chlide a
Phylogenetic analysis of the magnesium chelatase large subunits encoded by green bacterial genomes showed that the BchH, BchS, and BchT subunits diverge from the large subunit of cobalt chelatase, CobN (Figure 4.5). This analysis also shows that the BchS/T clade and the BchH clade diverge from each other relatively soon after branching from CobN. The BchS/T clade eventually diverged into BchS and BchT in an ancestor of modern GSB. In addition, these analyses confirmed the previous finding that most Chlorobi have three paralogs of BchH, which are denoted BchH, BchS, and BchT
(Gomez Maqueo Chew et al., 2009; Liu and Bryant, 2012). Two exceptions are Ca. T. aerophilum, which only has a single sequence within the BchS/T clade, and Chl. chlorochromatii, which has both BchS and BchT but not BchH. In this analysis the
BchS/T sequence of Ca. T aerophilum has been designated BchS, because it is the only copy in the BchS/T clade, and BchT from Cba. tepdium was shown to have very low enzyme activity compared to BchS, and likely could not produce the large amounts of
BChl c found in chlorosomes (Gomez Maqueo Chew et al., 2009; Johnson and Schmidt-
Dannert 2008). BchS encoded by Ca. T. aerophilum was located in three partial sequences, which is likely an artifact of metagenome sequencing and assembly and is not a true representation of the genome in any single organism. While the metagenome of Ca. 163 T. aerophilum appears to contain the mostly complete genome of this Candidatus strain,
the fragmentation of this sequence highlights the caution that must be used when
interpreting metagenome assemblies. The phylogentic analysis of subunits from
Chloroflexi also confirmed that all of the Chlorflexi genomes including those of Ca. V.
mediisalina and Ca. C. asiatica contain genes which encode two paralogs that fall within
the BchH clade and one paralog which falls within the BchS/T clade (Figure 4.5). In this
analysis the third Chloroflexi paralog was also designated BchS as in Ca. T. aerophilum.
Finally, it is interesting to note that the single BchH homolog of Cab. thermophilum, the
only green bacterium that encodes a single BchH paralog, appears closer to the inferred
ancestral node of the BchH clade than any other sequence in the clade.
Genes encoding BchM were identified in all green bacteria with the exception of
Ca. T aerophilum (Table 4.3). The absence of a gene encoding BchM was also noted
when the Ca. T. aerophilum metagenome was first published (Liu et al., 2012). Since
then, two additional metagenome assemblies from independent research groups have
been published (Roy et al., 2019; Stamps et al., 2014). A search of each of those
assemblies also returned no homologs of BchM. The absence of BchM suggests three
possibilities: first, that the gene encoding BchM is simply in an unsequenced region of
the genome; second, that the C13 propionate group is not methylated in Ca. T. aerophilum; and third, that another O-methyltransferase is responsible for this methylation in Ca. T. aerophilum. The first possibility will likely not be ruled out until a pure culture is obtained and the full genome sequenced. Alternatively, one of the other two possibilities might be demonstrated experimentally. The second possibility seems 164 very unlikely for two reasons. Firstly, Ca. T. aerophilum synthesizes both BChl a and Chl
a which should have a methyl group at this position; and secondly, Ca. T. aerophilum
encodes BciC which is responsible for removal of this methyl group in the biosynthesis
of BChl c, d, e, or f. In order to address the third possibility a list of O-methytransferase
candidates was compiled for this chapter. In total, 41 sequences are listed as candidates in
Table 4.8. Several of these genes are near other heme or (B)Chl biosynthesis genes
suggesting that they may be interesting candidates. Additionally, some are near the edge
of a contig suggesting that the full gene or gene region may not have been sequenced. A
thorough testing of these candidates could be completed in the future by
complementation of a bchM mutant in a photoheterotoph with an amenable genetic
system.
Based on this and prior analyses, isocyclic E-ring formation can be attributed to
AcsF and/or BchE in Cab. thermophilum and Chloroflexus spp, and BchE in the strictly anaerobic GSB and Osc. trichoides (Liu and Bryant, 2012). The two new members of the deeply branching Chloroflexineae were also both found to encode homologs of BchE in their genomes; however, unexpectedly Ca. V mediisalina also encoded a homolog of
AcsF. Three obvious explanations come to mind for this difference between Ca. V. mediisalina and other strict anaerobes. First, the gene encoding AcsF is not used for BChl biosynthesis in this organism but is a remnant from an aerobic ancestor; second, AcsF might serve another function as has been suggested in Cfl. aurantiacus; and third, that
Ca. V mediisalina is not in fact strictly anaerobic under all conditions in the environment
(Tang et al., 2009). Further characterization of this new strain will be required to 165 determine which of these three possibilities is most likely. Finally isocyclic E ring formation in Ca. T. aerophilum has not been assigned to any gene product to date (Table
4.3). As with BchM, the initial metagenome lacked genes encoding AcsF and BchE (Liu et al., 2012). The analyses in this study additionally checked Ca. T. aerophilum for genes encoding Ycf54 and BciE, however none were identified. All four of these enzymes were also absent in the two more recent Ca. T. aerophilum metagenome assemblies (Table
4.6). This absence of a known enzyme for formation of the isocyclic E ring is unique among green bacterial genomes but has also been reported in the genome of stramenopile algae (Matsuo and Inagaki, 2018). As with BchM it is possible that the gene encoding this function has simply not been sequenced or included in the metagenome assemblies available at this time or that an as yet unknown enzyme is responsible for this reaction.
Considering that a gene encoding BchM is missing, and that the gene encoding BchS is fragmented in the Ca. T. aerophilum metagenome it seems very possible that one of the known genes lies in an unsequenced region. However; the existence of the stramenopile algae also lacking a known enzyme for formation of the isocyclic E ring strongly suggests that at least one unknown enzyme exists that can perform this function.
4.5.3 Chlide a to BChlide a and BChlide c, d, e, and f
As in other studies the analysis of green bacterial genomes performed in this study showed that Chloroflexi and Cab. thermophilum genomes encode only one homolog of BchF while most Chlorobi genomes encode two or three paralogs of BchF.
This trend held true for the Ca. V mediisalina and Ca. C. asiatica metagenome assemblies, which each encoded a single BchF. This means that among the genomes of 166 green bacteria studied to date, only those of Chlorobi have BchV and BchF3 (Table 4.4).
Phylogentic analysis of the C3 hydratase sequences from green bacteria shows a BchF
clade which contains sequences from all three phyla of green bacteria and a BchV/F3
clade which only contains sequences from members of the Chlorobi (Figure 4.6). Within the BchV/F3 clade, a duplication event in an ancestor of modern Chlorobi gave rise to the
BchV and BchF3 clades. This is similar to the evolutionary relationship between BchH,
BchS, and BchT with the caveat that sequences in the BchF3 clade mostly belong to the brown-colored green sulfur bacteria while all GSB have both BchS and BchT. It is also important to note that all of the sequences from brown-colored GSB in the BchF3 clade are encoded in a gene cluster found in all brown-colored GSB (Thweatt et al., 2017; see
Chapter 3).
In Cba. tepidum it has been demonstrated that BchF and BchV produce differing ratios of R and S epimers at the C31 position and that their activity varies based on the
amount of methylation at the C8 and C12 positions (Gomez Maqueo Chew et al 2004;
Harada et al., 2015; Teramura et al., 2016; 2018). Generally, BchF produces mostly R
epimers while BchV produces both R and S epimers and BchV is more active than BchF
on more highly methylated BChlide species. Increased methylation skews the ratio of R
and S epimers further toward S chirality. Considering the interplay between methylation
state and hydratase activity, it is somewhat perplexing that Cab. thermophilum and the
deeply branching Chloroflexineae only have a single C-3 hydratase, because they also
produce BChl c methylated at the C8 and/or C12 positions. The duplication event which
gave rise to the BchV/F3 clade seems to have given Chlorobi a distinct evolutionary 167 advantage when it comes to fine tuning the stereochemistry of the C31 position in BChl c,
d, or e; this in turn allows them greater ability to tune the absorption characteristics of
their chlorosomes.
The genomes of all green bacteria contain genes encoding members of the
BchE/P-methyltransferase family. Analysis of the sequences in this family confirmed that
Chloroflexus spp. lacked BchQ and BchR while Chlorobi and Osc. trichoides had both.
Similar to Osc trichoides, Ca. V. mediisalina had sequences related to BchQ and BchR; however, Ca. C. asiatica only appears to encode BchQ (Figure 4.7, 4.8). Most surprisingly, sequences which were previously annotated as BchQ and BchR in Cab. thermophilum were shown by both SSN and maximum likelihood phyolgentic analysis to be distinct from the BchQ and BchR found in both the Chlorobi and the deeply branching
Chloroflexineae (Garcia Costas et al., 2012). The enzyme encoded by Cabther_B0083 does not cluster closely with other BchQ and BchR sequences from green bacteria but, based on its position in the SSN, it does likely function as a methyltransferase and is the most similar sequence encoded by Cab. thermophilum to BchQ and BchR. It seems plausible that while this would be the most distantly related C8 or C12 methyltransferase, this enzyme may still perform one or both of these functions. In contrast the enzyme encoded by Cabther_B0082 clusters more closely with BchE than BchQ or BchR; this agrees with the phylogentic tree which shows it is most closely related to the BchE clade
(Figure 4.8). Based on these analyses it seems much less likely that Cabther_B0082 functions as a C8 or C12 methyltransferase in BChl c biosynthesis. However; the fact that the Cabther_B0082 locus is in the same gene region as other (B)Chl biosynthesis genes 168 could be used as a counter argument, because it suggests this gene product may also be
involved in (B)Chl biosynthesis. The functionality of each of these enzymes could be
tested in the future by integration of the Cab thermophilum gene(s) into the genome of
Cba. tepidum strains containing mutations in bchQ and/or bchR (Gomez Maqueo Chew
et al., 2007)
4.5.4 Final Steps
All green bacteria make multiple types of (B)Chl and make use of multiple
paralogs of (B)Chl synthase (see Chapter 1 and Chapter 2). Prior work has shown that
Chloroflexi, Cab. thermophilum and Chlorobi have two paralogs, known as BchG and
BchK, which synthesize BChl a and BChl c, d, e, or f, respectively, and that Cab. thermophilum and Chlorobi have an additional paralog known as ChlG to synthesize Chl a (Bryant et al., 2012; Eisen et al., 2002; Frigaard et al., 2002, 2003; Garcia Costas et al.,
2012; Tang et al., 2011; Thweatt et al., 2017) This analysis showed that Ca. V. mediisalina and Ca. C. asiatica also contain homologs of BchG and BchK as expected for
Chloroflexi species. Interestingly, both Ca. V. mediisalina and Osc. trichoides had two sequences corresponding to BchK. This is unique within the green FAPs; however, several Chlorobi species also have multiple sequences corresponding to BchK (Liu and
Bryant, 2012). Previous phylogentic analysis of the (B)Chl synthase enzymes from GSB showed that two subclades, BchK1 and BchK2, exist within the GSB BchK clade, and that BchK2 is specifically associated with brown-colored GSB (see Chapter 3) (Thweatt et al., 2017). The analysis in this study, which included sequences from all green bacteria, showed that Chloroflexi and Cab. thermophilum BchK sequences are both outside of the 169 GSB BchK1 and BchK2 clades. This is significant because the only biochemically characterized BchK enzyme is that of Cfl. aurantiacus (Schoch et al., 1999). Finally, this analysis shows that (B)Chl synthases are derived from other prenyl transferases, and that the BChl synthase, BchG, branched from within a preexisting chlorin transferase clade.
Thus, BchG arose by a duplication from within an existing ancestral ChlG/BchK clade.
This strongly suggests that chlorin synthases existed prior to the branching of BchG in evolutionary history. This is in agreement with the idea that Chl a predated BChl a in the evolution of photosynthesis (Bryant et al., 2012; Bryant and Liu, 2013; Gomez Maqueo
Chew and Bryant, 2007).
4.5.6 Concluding Remarks
As more genome sequences of green bacteria become available, further bioinformatic analyses may shed new light on the distribution and evolution of (B)Chl biosynthesis in green bacteria. This set of analyses, carried out approximately six years after the last major review on the subject, has particularly benefited from the availability of new sequences from anaerobic green FAPs and Ca. T. aerophilum. Further analysis of the currently sequenced anaerobic green FAPs in addition to sequencing of new organisms may help to answer some remaining questions about (B)Chl biosynthesis in green FAPs. For example, the presence of AcsF in Ca. V. mediisalina could support the idea that AcsF may serve another function in Chloroflexi spp. or may challenge the characterization of Ca. V. mediisalina as a strict anaerobe. It will also be interesting to see if new genomes also show the presence of multiple homologs of BchK in the deeply 170 branching Chloroflexineae. Green bacteria in general, but in particular those within the
Chlorobi, have benefitted from duplication events within the genes related to (B)Chl biosynthesis. Further biochemical and mutagenesis studies will be needed to demonstrate the functions of these duplicates in vivo. Similarly, further studies of Ca. T. aerophilum
will be required to assign the enzymes responsible for C13 propionate group methylation
and formation of the isocyclic E ring.
171 References Abicht, H. K., Martinez, J., Layer, G., Jahn, D., and Solioz, M. (2012). Lactococcus lactis HemW (HemN) is a haem-binding protein with a putative role in haem trafficking. Biochem. J. 442, 335–343. doi:10.1042/BJ20111618. Albus, C. A., Salinas, A., Czarnecki, O., Kahlau, S., Rothbart, M., Thiele, W., et al. (2012). LCAA, a novel factor required for magnesium protoporphyrin monomethylester cyclase accumulation and feedback control of aminolevulinic acid biosynthesis in tobacco. Plant Physiol. 160, 1923–1936. doi:10.1104/pp.112.206045. Bollivar, D. W., Suzuki, J. Y., Beatty, J. T., Dobrowolski, J. M., and Bauer, C. E. (1994a). Directed mutational analysis of bacteriochlorophyll a biosynthesis in Rhodobacter capsulatus. J. Mol. Biol. 237, 622–640. doi:10.1006/jmbi.1994.1260. Bollivar, D. W., Wang, S., Allen, J. P., and Bauer, C. E. (1994b). Molecular genetic analysis of terminal steps in bacteriochlorophyll a biosynthesis: Characterization of a Rhodobacter capsulatus strain that synthesizes geranylgeraniol-esterified bacteriochlorophyll a. Biochemistry 33, 12763–12768. doi:10.1021/bi00209a006. Boynton, T. O., Daugherty, L. E., Dailey, T. A., and Dailey, H. A. (2009). Identification of Escherichia coli HemG as a novel, menadione-dependent flavodoxin with protoporphyrinogen oxidase activity. Biochemistry 48, 6705–6711. doi:10.1021/bi900850y. Bryant, D. A., and Liu, Z. (2013). Green bacteria: insights into green bacterial evolution through genomic analyses. Elsevier doi:10.1016/B978-0-12-397923-0.00004-7. Bryant, D. A., Liu, Z., Li, T., Zhao, F., Garcia Costas, A. M., Klatt, C. G., et al. (2012). “Comparative and functional genomics of anoxygenic green bacteria from the taxa Chlorobi, Chloroflexi, and Acidobacteria,” in Advances in Photosynthesis and Respiration, Vol. 35 Functional Genomics and Evolution of Photosynthetic Systems, eds. R. Burnap and W. Vermaas (Dordrecht: Springer Netherlands), 47–102. doi:10.1007/978-94-007-1533-2_3. Booker, S.J. (2009). Anaerobic functionalization of unactivated C–H bonds. Curr. Opin. Chem. Biol.13, 58-73. doi: 10.1016/j.cbpa.2009.02.036. Burke, D. H., Alberti, M., and Hearst, J. E. (1993). bchFNBH bacteriochlorophyll synthesis genes of Rhodobacter capsulatus and identification of the third subunit of light-independent protochlorophyllide reductase in bacteria and plants. J. Bacteriol. 175, 2414–2422. doi:10.1128/jb.175.8.2414-2422.1993. Canniffe, D. P., Thweatt, J. L., Chew, A. G. M., Hunter, C. N., and Bryant, D. A. (2018). A paralog of a bacteriochlorophyll biosynthesis enzyme catalyzes the formation of 1,2-dihydrocarotenoids in green sulfur bacteria. J. Biol. Chem. 293, 15233–15242. doi:10.1074/jbc.RA118.004672. Chen, G. E., Canniffe, D. P., and Hunter, C. N. (2017). Three classes of oxygen- dependent cyclase involved in chlorophyll and bacteriochlorophyll biosynthesis. Proc. Natl. Acad. Sci. USA 114, 6280–6285. doi:10.1073/pnas.1701687114. Chen, G. E., Canniffe, D. P., Barnett, S. F. H., Hollingshead, S., Brindley, A. A., Vasilev, C., et al. (2018). Complete enzyme set for chlorophyll biosynthesis in Escherichia coli. Sci. Adv. 4. doi:10.1126/sciadv.aaq1407. 172 Dailey, H. A., Dailey, T. A., Gerdes, S., Jahn, D., Jahn, M., O’Brian, M. R., et al. (2017). Prokaryotic heme biosynthesis: multiple pathways to a common essential product. Microbiol. Mol. Biol. Rev. 81, e00048-16. doi:10.1128/MMBR.00048-16. Dailey, T. A., and Dailey, H. A. (1996). Human protoporphyrinogen oxidase: Expression, purification, and characterization of the cloned enzyme. Protein Sci. 5, 98–105. Eisen, J. A, Nelson, K. E., Paulsen, I. T., Heidelberg, J. F., Wu, M., Dodson, R. J., et al. (2002). The complete genome sequence of Chlorobium tepidum TLS, a photosynthetic, anaerobic, green-sulfur bacterium. Proc. Natl. Acad. Sci. USA. 99, 9509–9514. doi:10.1073/pnas.132181499. Frigaard, N.U., Gomez Maqueo Chew A., Maresca J.A., and Bryant D.A. (2006) “Bacteriochlorophyll Biosynthesis in Green Bacteria.”in: Advances in Photosynthesis and Respiration, Vol. 25 Chlorophylls and Bacteriochlorophylls. ed Grimm B., Porra R.J., Rüdiger W., Scheer H. (Springer, Dordrecht), 201-221. doi:10.1007/1-4020- 4516-6_15. Frigaard, N. U., Voigt, G. D., and Bryant, D. A. (2002). Chlorobium tepidum mutant lacking bacteriochlorophyll c made by inactivation of the bchK gene, encoding bacteriochlorophyll c synthase. J. Bacteriol. 184, 3368–3376. doi:10.1128/JB.184.12.3368. Frigaard, N. U., Gomez Maqueo Chew, A., Li, H., Maresca, J. A., and Bryant, D. A. (2003). Chlorobium tepidum: Insights into the structure, physiology, and metabolism of a green sulfur bacterium derived from the complete genome sequence. Photosynth. Res. 78, 93–117. doi:10.1023/B:PRES.0000004310.96189.b4. Garcia Costas, A. M., Liu, Z., Tomsho, L. P., Schuster, S. C., Ward, D. M., and Bryant, D. A. (2012). Complete genome of Candidatus Chloracidobacterium thermophilum, a chlorophyll-based photoheterotroph belonging to the phylum Acidobacteria. Environ. Microbiol. 14, 177–190. doi:10.1111/j.1462-2920.2011.02592.x. Gaubier, P., Wu, H. J., Laudié, M., Delseny, M., and Grellet, F. (1995). A chlorophyll synthetase gene from Arabidopsis thaliana. Mol. Gen. Genet. 249, 58–64. doi:10.1007/BF00290236. Gibson, L. C. D., and Hunter, C. N. (1994). The bacteriochlorophyll biosynthesis gene, bchM, of Rhodobacter sphaeroides encodes S-adenosyl-L-methionine - Mg protoporphyrin-IX methyltransferase. FEBS Lett. 352, 127–130. doi:10.1016/0014- 5793(94)00934-1. Gomez Maqueo Chew, A., and Bryant, D. A. (2007). Chlorophyll biosynthesis in bacteria: the origins of structural and functional diversity. Annu. Rev. Microbiol. 61, 113–129. doi:10.1146/annurev.micro.61.080706.093242. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2009). Mutational analysis of three bchH paralogs in (bacterio-)chlorophyll biosynthesis in Chlorobaculum tepidum. Photosynth. Res. 101, 21–34. doi:10.1007/s11120-009-9460-0. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2008). Identification of the bchP gene, encoding geranylgeranyl reductase in Chlorobaculum tepidum. J. Bacteriol. 190, 747–749. doi:10.1128/JB.01430-07. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2007). Bacteriochlorophyllide c C-82 and C-121 methyltransferases are essential for 173 adaptation to low light in Chlorobaculum tepidumJ. Bacteriol. 189, 6176–6184. doi:10.1128/JB.00519-07. Gomez Maqueo Chew, A., Frigaard, N. U., and Bryant, D. A. (2004). “Identification of BchV, a C-31 hydratase specific for hypermethylated bacteriochlorophyll c in Chlorobium tepidum,” in Photosynthesis: Fundamental Aspects to Global Perspectives, eds. A. van der Est and D. Bruce (Lawrence, KS: Allen Press), 875– 877. Gough, S. P., Petersen, B. O., and Duus, J. Ø. (2000). Anaerobic chlorophyll isocyclic ring formation in Rhodobacter capsulatus requires a cobalamin cofactor. Proc. Natl. Acad. Sci. USA 97, 6980–6913. doi:10.1073/pnas.97.12.6908. Grouzdev, D. S., Kuznetsov, B. B., Keppen, O. I., Krasil’nikova, E. N., Lebedeva, N. V., and Ivanovsky, R. N. (2015). Reconstruction of bacteriochlorophyll biosynthesis pathways in the filamentous anoxygenic phototrophic bacterium Oscillochloris trichoides DG-6 and evolution of anoxygenic phototrophs of the order Chloroflexales. Microbiol. (United Kingdom) 161, 120–130. doi:10.1099/mic.0.082313-0. Harada, J., Mizoguchi, T., Satoh, S., Tsukatani, Y., Yokono, M., Noguchi, M., et al. (2013). Specific gene bciD for C7-methyl oxidation in bacteriochlorophyll e biosynthesis of brown-colored green sulfur bacteria. PLoS One 8, e60026. doi:10.1371/journal.pone.0060026. Harada, J., Saga, Y., Yaeda, Y., Oh-Oka, H., and Tamiaki, H. (2005). In vitro activity of C-20 methyltransferase, BchU, involved in bacteriochlorophyll c biosynthetic pathway in green sulfur bacteria. FEBS Lett. 579, 1983–1987. doi:10.1016/j.febslet.2005.01.087. Harada, J., Teramura, M., Mizoguchi, T., Tsukatani, Y., Yamamoto, K., and Tamiaki, H. (2015). Stereochemical conversion of C3-vinyl group to 1-hydroxyethyl group in bacteriochlorophyll c by the hydratases BchF and BchV: Adaptation of green sulfur bacteria to limited-light environments. Mol. Microbiol. 98, 1184–1198. doi:10.1111/mmi.13208. Haskamp, V., Karrie, S., Mingers, T., Barthels, S., Alberge, F., Magalon, A., et al. (2018). The radical SAM protein HemW is a heme chaperone. J. Biol. Chem. 293, 2558–2572. doi:10.1074/jbc.RA117.000229. Hollingshead, S., Kopecná, J., Jackson, P. J., Canniffe, D. P., Davison, P. A., Dickman, M. J., et al. (2012). Conserved chloroplast open-reading frame ycf54 is required for activity of the magnesium protoporphyrin monomethylester oxidative cyclase in Synechocystis PCC 6803. J. Biol. Chem. 287, 27823–27833. doi:10.1074/jbc.M112.352526. Huster, M. S., and Smith, K. M. (1990). Biosynthetic studies of substituent homologation in bacteriochlorophylls c and d. Biochemistry 29, 4348–4355. doi:10.1021/bi00470a013. Johnson, E. T., and Schmidt-Dannert, C. (2008). Characterization of three homologs of the large subunit of the magnesium chelatase from Chlorobaculum tepidum and interaction with the magnesium protoporphyrin IX methyltransferase. J. Biol. Chem. 283, 27776–27784. doi:10.1074/jbc.M804486200. 174 Jones, D. T., Taylor, W. R., and Thornton, J. M. (1992). The rapid generation of mutation data matrices from protein sequences. Bioinformatics 8, 275–282. doi:10.1093/bioinformatics/8.3.275. Kato, K., Tanaka, R., Sano, S., Tanaka, A., and Hosaka, H. (2010). Identification of a gene essential for protoporphyrinogen IX oxidase activity in the cyanobacterium Synechocystis sp. PCC6803. Proc. Natl. Acad. Sci. USA 107, 16649–16654. doi:10.1073/pnas.1000771107. Kobayashi, K., Masuda, T., Tajima, N., Wada, H., and Sato, N. (2014). Molecular phylogeny and intricate evolutionary history of the three isofunctional enzymes involved in the oxidation of protoporphyrinogen IX. Genome Biol. Evol. 6, 2141– 2155. doi:10.1093/gbe/evu170. Koch M., Breithaupt C., Kiefersauer R., Freigang J., Huber R., Messerschmidt A. (2004). Crystal structure of protoporphyrinogen IX oxidase: a key enzyme in haem and chlorophyll biosynthesis. EMBO J 23, 1720–1728. doi:10.1038/sj.emboj.7600189. Kumar, S., Stecher, G., and Tamura, K. (2016). MEGA7: Molecular evolutionary genetics analysis version 7.0. molecular biology and evolution. Mol. Biol. Evol. 33, 1870–1874. Layer, G., Grage, K., Teschner, T., Schünemann, V., Breckau, D., Masoumi, A., et al. (2005). Radical S-adenosylmethionine enzyme coproporphyrinogen III oxidase HemN: Functional features of the [4Fe-4S] cluster and the two bound S-adenosyl-L- methionines. J. Biol. Chem. 280, 29038–29046. doi:10.1074/jbc.M501275200. Layer G., Pierik A.J., Trost M., Rigby S.E., Leech H.K., Grage K., Breckau D., Astner I et al. (2006). The substrate radical of Escherichia coli oxygen-independent coproporphyrinogen III oxidase HemN. J Biol Chem 281,15727–15734. doi:10.1074/jbc.M512628200. Layer G., Verfürth K., Mahlitz E, Jahn D. (2002) Oxygen-independent coproporphyrinogen-III oxidase HemN from Escherichia coli. J Biol Chem. 277, 34136-42. doi:10.1074/jbc.M205247200 Liu, Z., Klatt, C. G., Ludwig, M., Rusch, D. B., Jensen, S. I., Kühl, M., et al. (2012). “Candidatus Thermochlorobacter aerophilum”: An aerobic chlorophotoheterotrophic member of the phylum Chlorobi defined by metagenomics and metatranscriptomics. ISME J 6, 1869–1882. doi:10.1038/ismej.2012.24. Liu, Z., and Bryant, D. A. (2012). “Bacteriochlorophyll biosynthesis and assembly in green chlorophototrophic bacteria: Theme and variations,” in Handbook of Porphyrin Science with Applications to Chemistry, Physics, Materials Science, Engineering, Biology and Medicine, Vol 20: Chlorophylls and Related Systems Handbook of Porphyrin Science., ed. R. Kadish, KM and Smith, KM and Guilard (Singapore: World Scientific Publishing Company Pte Ltd), 107–140. Liu, Z., and Bryant, D. A. (2011a). Multiple types of 8-vinyl reductases for (Bacterio)chlorophyll biosynthesis occur in many green sulfur bacteria. J. Bacteriol. 193, 4996–4998. doi:10.1128/JB.05520-11. Liu, Z., and Bryant, D. A. (2011b). Identification of a gene essential for the first committed step in the biosynthesis of bacteriochlorophyll c. J. Biol. Chem. 286, 22393–22402. doi:10.1074/jbc.M111.249433. 175 Möbius, K., Arias-Cartin, R., Breckau, D., Haennig, A.L., Riedmann, K., Biedendieck, R., et al. (2010). Heme biosynthesis is coupled to electron transport chains for energy generation. Proc. Natl. Acad. Sci. U. S. A. 107, 10436–10441. doi:10.1073/pnas.1000956107. Maresca, J. A., Gomez Maqueo Chew, A., Ponsatí, M. R., Frigaard, N. U., Ormerod, J. G., and Bryant, D. A. (2004). The bchU gene of Chlorobium tepidum encodes the C- 20 methyltransferase in bacteriochlorophyll c biosynthesis. J. Bacteriol. 186, 2558– 2566. doi:10.1128/JB.186.9.2558-2566.2004. Matsuo, E., and Inagaki, Y. (2018). Patterns in evolutionary origins of heme, chlorophyll a and isopentenyl diphosphate biosynthetic pathways suggest non photosynthetic periods prior to plastid replacements in dinoflagellates. PEER J 6, e5345. doi:10.7717/peerj.5345. Nagaraj, V. A., Arumugam, R., Prasad, D., Rangarajan, P. N., and Padmanaban, G. (2010). Protoporphyrinogen IX oxidase from Plasmodium falciparum is anaerobic and is localized to the mitochondrion. Mol. Biochem. Parasitol. 174, 44–52. doi:10.1016/j.molbiopara.2010.06.012. Oster, U., Bauer, C. E., and Rüdiger, W. (1997). Characterization of chlorophyll a and bacteriochlorophyll a synthases by heterologous expression in Escherichia coli. J. Biol. Chem. 272, 9671–9676. doi:10.1074/jbc.272.15.9671. Pinta, V., Picaud, M., Reiss-Husson, F., and Astier, C. (2002). Rubrivivax gelatinosus acsF (previously orf358) codes for a conserved, putative binuclear-iron-cluster- containing protein involved in aerobic oxidative cyclization of Mg-protoporphyrin IX monomethylester. J. Bacteriol. 184, 746–753. doi:10.1128/JB.184.3.746-753.2002. Porra, R. J., Schafer, W., Gadon, N., Katheder, I., Drews, G., and Scheer, H. (1996). Origin of the two carbonyl oxygens of bacteriochlorophyll alpha - Demonstration of two different pathways for the formation of ring E in Rhodobacter sphaeroides and Roseobacter denitrificans, and a common hydratase mechanism for 3-acetyl group formation. Eur. J. Biochem. 239, 85–92. doi:10.1111/j.1432-1033.1996.0085u.x. Roy, C., Bakshi, U., Rameez, M.J., Mandal, S., Haldar, P.K., Pyne, P. and Ghosh, W. (2019). Phylogenomics of an uncultivated, aerobic and thermophilic, photoheterotrophic member of Chlorobia sheds light into the evolution of the phylum Chlorobi. Comp. Biol.& Chem. 80, 206-216. doi: 10.1016/j.compbiolchem.2019.04.001 Rüdiger, W., Benz, J., and Guthoff, C. (1980). Detection and partial characterization of activity of chlorophyll synthetase in etioplast membranes. Eur. J. Biochem. 109, 193– 200. doi:10.1111/j.1432-1033.1980.tb04784.x. Saga, Y., Hirota, K., Harada, J., and Tamiaki, H. (2015). In vitro enzymatic activities of bacteriochlorophyll a synthase derived from the green sulfur photosynthetic bacterium Chlorobaculum tepidum. Biochemistry 54, 4998–5005. doi:10.1021/acs.biochem.5b00311. Sasarman, A., Letowski, J., Czaika, G., Ramirez, V., Nead, M. A., Jacobs, J. M., et al. (1993). Nucleotide-sequence of the hemg gene involved in the protoporphyrinogen oxidase activity of Escherichia-coli K12. Can. J. Microbiol. 39, 1155–1161. doi:10.1139/m93-174. 176 Schoch, S., Oster, U., Mayer, K., Feick, R., and Rudiger, W. (1999). Subtrate specificity of overexpressed bacteriochlorophyll synthase from Chloroflexus aurantiacus. in Chloroplast: From Molecular Biology to Biotechnology ., ed. H. Argyroudi Akoyunoglou, JH and Senger (Netherlands: Springer), 213–216. Skotnicová, P., Sobotka, R., Shepherd, M., Hájek, J., Hrouzek, P., and Tichý, M. (2018). The cyanobacterial protoporphyrinogen oxidase HemJ is a new b-type heme protein functionally coupled with coproporphyrinogen III oxidase. J. Biol. Chem. 293, 12394–12404. doi:10.1074/jbc.RA118.003441. Stamps, B.W., Corsetti, F.A., Spear, J.R., Stevenson, B.S. (2014) Draft genome of a novel Chlorobi member assembled by tetranucleotide binning of a hot spring metagenome. Genome Announc. 2, e00897-14. doi:10.1128/genomeA.00897-14 Tank, M., and Bryant, D. A. (2015). Chloracidobacterium thermophilum gen. nov., sp. nov.: An anoxygenic microaerophilic chlorophotoheterotrophic acidobacterium. Int. J. Syst. Evol. Microbiol. 65, 1426–1430. doi:10.1099/ijs.0.000113. Tank, M., Liu, Z., Frigaard, N. U., Tomsho, L. P., Schuster, S. C., and Bryant, D. A. (2017). Complete genome sequence of the photoautotrophic and bacteriochlorophyll e-synthesizing green sulfur bacterium Chlorobaculum limnaeum DSM 1677T. Genome Announc. 5, e00529-17. doi:10.1128/genomeA.00529-17. Tang, K. H., Barry, K., Chertkov, O., Dalin, E., Han, C. S., Hauser, L. J., et al. (2011). Complete genome sequence of the filamentous anoxygenic phototrophic bacterium Chloroflexus aurantiacus. BMC Genomics 12, 334. doi:10.1186/1471-2164-12-334. Tang, K. H., Wen, J., Li, X., and Blankenship, R. E. (2009). Role of the AcsF protein in Chloroflexus aurantiacus. J. Bacteriol. 191, 3580–3587. doi:10.1128/JB.00110-09. Teramura, M., Harada, J., and Tamiaki, H. (2018). In vitro enzymatic assays of photosynthetic bacterial 3-vinyl hydratases for bacteriochlorophyll biosyntheses. Photosynth. Res. 135, 319–328. doi:10.1007/s11120-017-0415-6. Teramura, M., Harada, J., and Tamiaki, H. (2016). In vitro stereospecific hydration activities of the 3-vinyl group of chlorophyll derivatives by BchF and BchV enzymes involved in bacteriochlorophyll c biosynthesis of green sulfur bacteria. Photosynth. Res. 130, 33–45. doi:10.1007/s11120-016-0220-7. Thweatt, J. L., Ferlez, B. H., Golbeck, J. H., and Bryant, D. A. (2017). BciD is a radical S-adenosyl-L-methionine (SAM) enzyme that completes bacteriochlorophyllide e biosynthesis by oxidizing a methyl group into a formyl group at C-7. J. Biol. Chem. 292, 1361–1373. doi:10.1074/jbc.M116.767665. Wang, S.C. (2018). Cobalamin-dependent radical S-adenosyl-L-methionine enzymes in natural product biosynthesis. Nat. Prod. Rep. 35, 707-720. doi: 10.1039/C7NP00059F Willows, R. D., Gibson, L. C. D., Kanangara, C. G., Hunter, C. N., and von Wettstein, D. (1996). Three separate proteins constitute the magnesium chelatase of Rhodobacter sphaeroides. Eur. J. Biochem. 235, 438–443. doi:10.1111/j.1432-1033.1996.00438.x.
177
Figure 4.1- Phylogeny of HemN Homologs in Green Bacteria Maximum likelihood tree showing protein sequences of all HemN homologs in green bacteria, the biochemically characterized HemW from Lactobacillus lactis, and NifB from Cba. tepidum as the outgroup. The tree with the highest log likelihood (-17085.9738) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 45 amino acid sequences. All positions containing gaps and missing data were eliminated. There were a total of 351 positions in the final dataset. 178
Figure 4.2 Multiple Sequence Alignment of Green Bacterial HemN Paralogs Mulitple sequence alignment of the N-terminal portion of the HemN paralogs found in green bacteria (See also Figure 2.1) showing the conserved motifs found in HemN and HemW. The cysteines that correspond to the HemN or HemW motif are highlighted in blue while the phenylalanine found in HemW (replacing a cysteine found in HemN) is highlighted in red. 179 -
Figure 4.3 Sequence Similarity Network of HemY Related Sequences in Green Bacteria SSN of HemY related sequences from green bacteria, and biochemically characterized HemY enzymes. The network contains all edges with an alignment score of greater than 13. Each node represents sequences of 100% identity.
180
Figure 4.4- Phylogeny of HemY Related Protein in Green Bacteria Maximum likelihood tree showing all protein sequences from clusters HemY and “HemY” represented in the SSN in Figure 4.3 Additional select sequences from the phytoene desaturase (CrtI) clutser and the CrtI enzyme from Rhodobacter sphaeroides were included as an outgroup. The tree with the highest log likelihood (-26906.6004) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 38 amino acid sequences. There were 739 positions in the final dataset. Bootstrap values are based on 500 replicates. 181
182 Figure 4.5- Phylogeny of BchH Paralogs in Green Bacteria Maximum likelihood tree showing all protein sequences of BchH, BchS, and BchT in green bacteria along with select sequences of CobN used as the outgroup. The tree with the highest log likelihood (-84649.1733) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 76 amino acid sequences. There were 1702 positions in the final dataset. Bootstrap values are based on 100 replicates. 183
Figure 4.6- Phylogeny of BchF Paralogs in Green Bacteria Maximum likelihood tree of all BchF paralogs in green bacteria and the BchF enzyme of Rhodobacter sphaeroides. Sequences from brown-colored GSB are highlighted in brown. The * here is to clarify that in the case of the two green-colored GSB in the BchF3 clade each only has two paralogs of BchF, and these sequences are likely referred to as BchV in other literature. The tree with the highest log likelihood (-6959.5368) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 48 amino acid sequences. There were 241 positions in the final dataset. Bootstrap values are based on 500 replicates. 184
Figure 4.7 Sequence Similarity Network of BchQ, BchR and BchE Related Sequences in Green Bacteria SSN of blast hits to BchQ, BchR, and BchE from green bacteria combined with sequences from 1/50 of the IPR034466 Interpro family. Sequences were filtered for proteins between 270 and 950 amino acids in length. The network was constructed with 418 sequences from the Interpro family and 158 sequences from blast hits in green bacteria for a total of 576 sequences. For visualization here, only clusters with more than 2 sequences from green bacteria were included. The network contains all edges with an alignment score of greater than 63. Each node represents sequences of 90% identity.
185
186 Figure 4.8- Phylogeny of BchQ, BchR, and BchE in Green Bacteria Maximum likelihood tree of all sequences from green bacteria represented in the BchQ, BchR, and BchE cluster in the SSN in Figure 4.5 in addition to select sequences from the CT1502/CT0072 cluster, two sequences from Ca. V m that cluster nearby, and sequences encoded by CabtherB0082 and CabtherB0083. The tree with the highest log likelihood (- 38662.0371) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 85 amino acid sequences. All positions with less than 5% site coverage were eliminated. That is, fewer than 95% alignment gaps, missing data, and ambiguous bases were allowed at any position. There are 638 positions in the final dataset. Bootstrap values are based on 100 replicates.
187
188 Figure 4.9- Phylogeny of ChlG, BchG and BchK in Green Bacteria Maximum likelihood tree of (B)Chl synthase related proteins from green bacteria. Sequences associated with brown-colored GSB in the BchK clade are highlighted in brown. The tree with the highest log likelihood (-34860.0980) is shown. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 96 amino acid sequences. There are 482 positions in the final dataset. Bootstrap values are based on 100 replicates.
189 Table 4.1-Green Bacterial Genomes Analyzed in this Work Categories correspond to those used in Figure 2.1. For further information on green bacteria, see Chapter 1. Accessions for genomes found in public databases are listed either : a-JGI (IMG Genome ID) or b-NCBI (Genbank). The Seq Prefix column lists either the locus tag prefix found in the JGI database or the first three letters of the NCBI protein accession numbers, these prefixes are used in the phylogenetic analyses in this chapter.*Indicates organisms in this list whose draft genomes were assembled from metagenomic data. Category Organism Genome Seq Prefix Brown Cba. limnaeum DSM 1677T 2501846309b ClimDRAFT Chlorobiaceae Cba. limnaeum Rk-j-1 2511231051b ClimR Prosthecochloris sp. BS-1 642555122b Cphamn1 Chl. clathratiforme 642555146b Ppha DSM5477T Chl. phaeobacteroides DSM 639633020b Cpha266 266 Prosthecochloris sp. CIB2401 2721755577b Ga0175537 Green Cba. tepidum TLS 637000073b CT Chlorobiaceae Chl. limicola DSM245 642555121b Clim Chl. chlorochromatii CaD3 637000072b Cag Chl. phaeovibriodes DSM 265 640427130b Cvib Ptc. bathyomarina GSB1 2757320899b Ga0226291 Chl. luteolum DSM273 637000205b Plut Chl. ferrooxidans DSM 13031 638341060b CferDRAFT Ptc. aestuarii DSM 271 642555149b Paes Cba. parvum DSM 263 642555120b Cpar Chp. thalassium Chp. thalassium ATCC 35110 642555123b Ctha Ca. T. aerophilum ‘Ca. T. aerophilum’* PHFL00000000.1a RFM Cab. thermophilum Cab. thermophilum B 2512047033b Cabther Deeply Branching Osc. trichoides DG-6 649989977b OSCT Chloroflexineae ‘Ca. Chloroploca asiatica’ LYXE00000000.1a PDV, PDW B7–9* ‘Ca. Viridilinea mediisalina NQWI00000000.1a PDV, PDW Kir15-3F* Chloroflexus spp. Cfl. aurantiacus J-10-fl 641228485b Caur Chloroflexus. sp. Y-400-fl 643692015b Chy400 Chloroflexus sp. Y-396-1 2506520040b ChY396 Cfl. aggregans DSM 9485 643348527b Cagg
190 Table 4.2-(B)Chl Genes in Green Bacteria-Early Steps
Organism HemF HemN HemW HemY "HemY" HemG Cba. limnaeum T DSM 1677 none 2501893722 2501895557 none 2501893827 none Cba. limnaeum Rk-j-1 none 2511494694 2511494899 none 2511494797 none Prosthecochloris sp. BS-1 none 642683170 642684731 none 642684627 642683905 Chl. clathratiforme DSM5477T none 642726997 642726624 none 642728697 none Chl. phaeobacteroides DSM 266 none 639765212 639766933 none 639766831 none Prosthecochloris sp. CIB2401 none 2723115006 2723115321 none 2723115259 none Cba. tepidum TLS none 637115309 637116948 none 637116837 none Chl. limicola DSM245 none 642669550 642669778 none 642669706 none Chl. chlorochromatii CaD3 none 637772811 637774344 none 637772773 none Chl. phaeovibriodes DSM 265 none 640453793 640452692 none 640452760 none Ptc. bathyomarina GSB1 none 2758495937 2758496329 none 2758496259 none Chl. luteolum DSM273 none 637769612 637768260 none 637768329 none Chl. ferrooxidans DSM 13031 none 639204976 639206099 none 639205807 none Ptc. aestuarii DSM 271 none 642724597 642726001 none 642725919 none Cba. parvum DSM 263 none 642720152 642718876 none 642718950 none Chp. thalassium ATCC 35110 none 642716769 642716658 none 642718950 none ‘Ca. T. aerophilum’ RFM23036.1 none RFM24189.1 RFM23380.1 none none Cab. thermophilum B 2512358972 2512358194 2512360066 2512358728 none none 191
Osc. trichoides 650113407; DG-6 none 650112131 650114733 650112931 none none ‘Ca. Chloroploca PDV98931.1; asiatica’ B7–9 none PDV98935.1 PDV97111.1 PDV9926.1 none none ‘Ca. Viridilinea mediisalina Kir15-3F none PDW02979.1 PDW03555.1 PDW02975.1 none none Cfl. aurantiacus J-10-fl 641373281 641371323 641370895 641371324 none none Chloroflexus sp. Y-400-fl 643712989 643710881 643710413 643710882 none none Chloroflexus sp. Y-396-1 2506717593 2506716614 2506716854 2506716615 none none Cfl. aggregans DSM 9485 643566136 643568177 643568366 643568176 none none
192 Table 4.3 (B)Chl Genes in Green Bacteria Leading from ProtoIX to Chlide a. # Organism BchH BchH BchS BchT BchD BchI Cba. limnaeum DSM 1 T 1677 2501893909 na 2501893911 2501893947 2501893946 2501893945 2 Cba. limnaeum Rk-j-1 2511496533 na 2511496535 2511495527 2511495528 2511495529 3 Prosthecochloris sp. BS-1 642684694 na 642684692 642683549 642683548 642683547 Chl. clathratiforme 4 T DSM5477 642728740 na 642728738 642728009 642728010 642728011 Chl. phaeobacteroides 5 DSM 266 639766874 na 639766872 639766213 639766214 639766215 Prosthecochloris sp. 6 CIB2401 2723115293 na 2723115292 2723114000 272311399 2723113998
7 Cba. tepidum TLS 637116896 na 637116894 637116241 637116242 637116243
8 Chl. limicola DSM245 642669747 na 642669745 642924245 642669098 642669099 Chl. chlorochromatii 9 CaD3 none na 637772754 637773513 637774159 637774160 Chl. phaeovibriodes DSM 10 265 640452722 na 640452724 640453470 640453471 640453472
11 Ptc. bathyomarina GSB1 2758496294 na 2758496292 2758495051 2758495050 2758495049
12 Chl. luteolum DSM273 637768291 na 637768293 637768828 637768827 637768826 Chl. ferrooxidans DSM 13 13031 639206222 na 639206223 639205844 639205845 639205846
14 Ptc. aestuarii DSM 271 642725959 na 642725954 642725423 642725424 642725425
15 Cba. parvum DSM 263 642718907 na 642718909 642719378 642719376 642719375 Chp. thalassium ATCC 16 35110 642717178 na 642717180 642717595 642717970 642717969 RFM23248.1 17 ‘Ca. T. aerophilum’ +23247.1 none na +23035.1 none RFM23270.1 RFM24577.1
18 Cab. thermophilum B 2512358129 na none none 2512359281 2512359649
19 Osc. trichoides DG-6 650112392 650113406 650113678 none 650114405 650114406 ‘Ca. Chloroploca asiatica’ PDV98936. 20 B7–9 PDV96852.1 1 PDV98441.1 none PDV99933.1 PDV99761.1 ‘Ca. Viridilinea PDW02980 PDW02570. 21 mediisalina Kir15-3F PDW01774.1 .1 PDW03164.1 none 1 PDW02569.1
22 Cfl. aurantiacus J-10-fl 641373831 641373273 641374039 none 641371105 641371104
23 Chloroflexus sp. Y-400-fl 643713580 643712981 643713803 none 643710641 643710640 250671758 24 Chloroflexus sp. Y-396-1 2506717805 5 2506717040 none 2506714574 2506714573 25 Cfl. aggregans DSM 9485 643565438 643566128 643565108 none 643567943 643567944 193
# Chl/BchM BchE AcsF BciA BciB Chl/BchL Chl/BchN Chl/BchB 1 2501893908 2501893907 none 2501894594 none 2501894095 2501894093 2501894094 2 2511496532 2511496531 none 2511494419 none 2511495211 2511495213 2511495212 3 642684695 642684696 none 642683875 none 642684744 642684746 642684745
4 642728741 642728742 none 642727411 642728895 642726618 642726616 642726617 640068628; 5 639764788; 639766875 639766876 none none 639765294 639766944 639766946 639766945
6 2723115294 2723115295 none 2723114566 none 2723115327 2723115329 2723115328
7 637116897 637116898 none 637116007 none 637117093 637117095 637117094 642669394; 8 642669748 642669749 none none 642668576 642669783 642669785 642669784
9 637772755 637772756 none 637773590 none 637774351 637774353 637774352
10 640452721 640452720 none 640452933 none 640452687 640452685 640452686 275849593 11 2758496295 2758496296 none 2758495378 2 2758496337 2758496339 2758496338
12 637768290 637768289 none 637768511 none 637768255 6377768253 637768254
13 639206221 639206220 none 639206137 639205748 639206090 639206088 639206089
14 642725960 642725961 none 642725207 642724602 642726010 642726012 642726011
15 642718906 642718905 none 642719666 none 642718863 642718861 642718862
16 642717177 642716695 none none 642717102 642717264 642717266 642717265 RFM23238. RFM23834.1/ RFM23832.1/ RFM23833.1/ 17 see text none none none 1 RFM25128.1 RFM25126.1 RFM25127.1 251235813 18 2512359774 2512358195 2512360263 none 2 2512358131 2512359929 2512359928
19 650113405 650112132 none none 650113418 650113419 650113416 650113417
20 PDV98937.1 PDV99400.1 none none PDV96621.1 PDV96620.1 PDV96623.1 PDV96622.1 PDW02983. PDW01131. PDW02981. PDW00547. 21 1 1 1 none 1 PDW00548.1 PDW02967.1 PDW02966.1
22 641373270 641374348 641373272 none 641373237 641373236 641373239 641373238
23 643712978 643714133 643712980 none 643712941 643712940 643712943 643712942 250671755 24 2506717582 2506716979 2506717584 none 7 2506717556 2506717559 2506717558 25 643566125 643565184 643566127 none 643568288 643568287 643568290 643568289
194 Table 4.4 (B)Chl Genes in Green Bacteria Leading from Chlide a to BChlide c, d, e, or f # Organism BchX BchY BchZ BchF BchV BchF3 Cba. limnaeum DSM 1 T 1677 2501895001 2501893732 2501894355 2501895003 2501895209 2501895604 2 Cba. limnaeum Rk-j-1 2511495911 2511494704 2511496062 2511495913 2511495336 2511494852 3 hloris sp. BS-1 642683439 642684553 642682692 642683441 none 642682782 Chl. clathratiforme 4 T DSM5477 642727230 642728593 642726472 642727232 642728527 642728905 Chl. phaeobacteroides 5 DSM 266 639766375 639766656 639767060 639766373 639766587 639764798 Prosthecochloris sp. 6 CIB2401 2723113917 2723115199 2723113380 2723113919 2723115149 2723113463 7 Cba. tepidum TLS 637116370 637116766 637117063 637116368 637116716 none 642668745; 8 Chl. limicola DSM245 642668524 642669630 642667784 642668526 642669572 none Chl. chlorochromatii 9 CaD3 637772928 637772851 637774253 637772930 637774182 none Chl. phaeovibriodes 10 DSM 265 640453665 640452812 640452594 640453663 640452774! none Ptc. bathyomarina 11 GSB1 2758494925 2758496185 2758494372 2758494927 2758494449! none Chl. luteolum 12 DSM273 637769461 637768381 637768162 637769459 637768447 none Chl. ferrooxidans 13 DSM 13031 639206670 639204912 639206643 639206672 639204859 none Ptc. aestuarii DSM 14 271 642725525 642725847 642726011 642725523 642725789 none Cba. parvum DSM 15 263 642719392 642719020 642718760 642719394 642719073 none Chp. thalassium 16 ATCC 35110 642718622 642716075 642716701 none 642718620 none RFM24534.1; 17 ‘Ca. T. aerophilum’ RFM24537.1 RFM23516.1 RFM24144.1 none RFM24644.1 none 18 Cab. thermophilum B 2512359818 2512360455 2512360411 2512358133 none none 19 Osc. trichoides DG-6 650114407 650114526 650114525 650114409 none none ‘Ca. Chloroploca 20 asiatica’ B7–9 PDV99759.1 PDV98336.1 PDV98335.1 PDV99932.1 none none ‘Ca. Viridilinea 21 mediisalina Kir15-3F PDW02585.1 PDV99952.1 PDV99953.1 PDW02566.1 none none Cfl. aurantiacus J-10- 22 fl 641371102 641374474 641374475 641371100 none none Chloroflexus sp. Y- 23 400-fl 643710636 643714276 643714277 643710634 none none Chloroflexus. sp. Y- 24 396-1 2506714569 2506717307 2506717306 2506714567 none none Cfl. aggregans DSM 25 9485 643567948 643565698 643565699 643567950 none none
195
# BchC BciC BchQ BchQ2 BchR BchU BciD 1 2501895002 2501894618 2501895207 2501895606 2501895223 2501893505 2501895602 2511495322; 2 2511495912 2511494441 2511495337 2511494851 2511495873 2511496592 2511494854 3 642683440 642683867 none 642682781 642683522 642684990 642682784 642728906; 4 642727231 642727556 642728891 642728911 642727834 642728912 642728903 5 639766374 639765867 truncated 639764800 639766230 639767282 639764796 6 2723113918 2723114404 2723115150 2723113462 2723113982 2723115512 2723113465 7 637116369 637116021 637116717 none 637116268 637114950 none 642669573; 8 642668525 642668828 642668780 none 642669123 642670084 none 637774077; 9 637772929 637773600 637772642 none 637773461 637772641 none 10 640453664 640453262 640452773 none 640453461 640454179 none 2758494953; 11 2758494926 2758495386 2758494448 none 2758495024 2758496533 none 637768974; 12 637769460 637769120 637768446 none 637769356 637770135 none 639206460; 13 639206671 639206303 639206016 none 639206190 639206015 none 14 642725524 642725141 642725790 none 642725455 642726229 none 15 642719393 642719688 642719072 none 642719985 642923841 none 16 642718621 642718096 642717185 none 642718202 642717390 none RFM25204.1/ 17 RFM24535.1 RFM24422.1 RFM23760.1 none RFM23847.1 none none 18 2512360410 2512358134 see text none see text 2512358635 none 19 650114408 650114545 650113564 none 650113563 650113124 none 20 PDV99758.1 PDV98734.1 PDV97109.1 none none PDW01266.1 none 21 PDW02567.1 PDW03483.1 PDW04312.1 none PDW04696.1 PDW04110.1 none 22 641371101 641371590 none none none 641370823 none 23 643710635 643711184 none none none 643710337 none 24 2506714568 2506716021 none none none 2506717180 none 25 643567949 643567067 none none none 643566054 none
196 Table 4.5 (B)Chl Genes in Green Bacteria Final Steps m ChlG BchG BchK BchK1 BchK2 BchP Cba. limnaeum DSM 1677T 2501893985 2501894297 none 2501895222 2501895540 2501895761 Cba. limnaeum Rk-j-1 2511495489 2511495004 none 2511495323 2511494917 2511496189 Prosthecochloris sp. 642684714;642 BS-1 642683590 642683115 none none 684713 642682544 Chl. clathratiforme DSM5477T 642727393 642726965 none none 642726656 642726331 Chl. phaeobacteroides DSM 266 639766035 639765181 none none 639766918 639764645 Prosthecochloris sp. CIB2401 2723114013 2723113816 none none 2723115305 2723113272 Cba. tepidum TLS 637116212 637116554 none 637116930 none 637117200 Chl. limicola DSM245 642668772 642668261 none none 642669763 642667646 Chl. chlorochromatii CaD3 637773636 637774279 none 637774078 637772736 637772561 Chl. phaeovibriodes DSM 265 640453150 640453818 none none 640452708 640452422 Ptc. bathyomarina GSB1 2758495102 2758495995 none 2758494954 2758496310 2758494229 Chl. luteolum DSM273 637768945 637769638 none 637768975 637768276 637768044 Chl. ferrooxidans DSM 13031 639206018 639205005 none none 639206122 639206270 Ptc. aestuarii DSM 271 642725375 642724544 none none 642725982 642724015 Cba. parvum DSM 263 642719913 642719217 none 642718891 none 642718656 Chp. thalassium ATCC 35110 642718508 642717930 none 642716462 none 642717164 ‘Ca. T. aerophilum’ RFM24816.1 RFM23684.1 none RFM23284.1 none RFM24125.1 Cab. thermophilum B 2512358092 2512360647 2512360120 none none 2512359034 Osc. trichoides DG-6 650114433; none 650115175 650113123 none none 650115174 ‘Ca. Chloroploca asiatica’ B7–9 none PDW01241.1 PDW01265.1 none none PDW01240.1 ‘Ca. Viridilinea PDW04109.1; mediisalina Kir15-3F none PDW03983.1 PDW03495.1 none none PDW03982.1 Cfl. aurantiacus J-10- fl none 641372762 641370824 none none 641372761 Chloroflexus sp. Y- 400-fl none 643712428 643710338 none none 643712427 Chloroflexus sp. Y- 396-1 none 2506716035 2506717179 none none 2506716036 Cfl. aggregans DSM 9485 none 643567458 643566053 none none 643567459 197 Table 4.6 Metagenome-assembled genomes of ‘Candidatus Thermochlorobacter aerophilum’ Information on the three currently available metagenome assemblies for strains of Candidatus Thermochlorobacter aerophilum used to search for genes encoding C13 propionate methyltransferase and the oxidative isocyclic E ring cyclase. Organism WGS Biosample Reference
Candidatus Thermochlorobacter PHFL00000000 SAMN07635172 Liu et al., 2012 aerophilum OS
Candidatus JPGV00000000 SAMN02899251 Stamps et al., 2014 Thermochlorobacteriaceae bacterium GBChlB
Chlorobium sp. 445 NSLH00000000 SAMN07581424 Roy et al., 2019
198 Table 4.7 Summary of O-methyltransferase Hits in the CDD from Ca. T. aerophilum
Super Family Unique CDD Super Family Hits e < 10-5 Name Identifier
Cl28429, Cl06870, SpoU rel 5 5 5 Cl21505, UbiG Cl28104 26 25 24 Class I MTase Cl17173 49 46 22 Hen1 like Cl25638 4 1 0 SmtA Cl28099 30 23 0 PEMT Cl21511 3 2 2 ATase Cl27615 2 2 2 PCMT Cl28095 9 2 1 YrrM Cl28097 8 3 0 TrmJ Cl28430 2 1 0 TP MTase Cl00304 2 2 2 CMAS Cl28102 22 16 0 Total 14 162 128 58
199 Table 4.8-List C13 Propionate Methyltransferase Candidates in Ca. T. aerophilum OS Length best hit Accession Description Nearby in Genome Best Hit Best Hit Description (AA) E value isoprenylcysteine carboxyl RFM24429.1 167 methyltransferase hemA 16kb WP_090989385.1 1.1E-57 same as query[Bacillus sp. OV322] same as query[Porphyromonadaceae RFM24486.1 321 class I SAM-dependent methyltransferase 0 WP_068906906.1 1.1E-54 bacterium H1] bchF, bchC, bchX ~20kb; RFM24553.1 245 class I SAM-dependent methyltransferase bchI ~20 kb PZU96169.1 2.8E-86 same as query [Pseudanabaena sp.] hypothetical protein DME24_15855 RFM24203.1 304 class I SAM-dependent methyltransferase hemW ~16kb PYJ58549.1 6E-148 [Verrucomicrobia bacterium] bchZ ~20kb; bchP ~25 kb; RFM24102.1 419 class I SAM-dependent methyltransferase ~3kb from start of contig TMI62435.1 0 same as query [Bacteroidetes bacterium] same as query [Chitinophagaceae RFM24103.1 262 FkbM family methyltransferase same as 102 WP_054279951.1 2.7E-88 bacterium PMP191F] Methylase involved in RFM25416.1 235 class I SAM-dependent methyltransferase on edge of sequence KPQ39534.1 5.8E-62 ubiquinone/menaquinone biosynthesis SDR family NAD(P)-dependent TPA: short-chain dehydrogenase RFM23890.1 153 oxidoreductase on edge of contig HBH06518.1 1.2E-48 [Flavobacteriales bacterium] bchNLB ~3.5 kb; bchJ ~1.5kb; RFM23838.1 282 class I SAM-dependent methyltransferase ferrochelatase ~10 kb WP_011956646.1 1.9E-63 same as query [Roseiflexus sp. RS-1] ferrocheletase ~ 3kb; bchJ ribosomal small subunit Rsm22 RFM23850.1 318 methyltransferase ~17 kb; bchNLB ~19 kb WP_012500121.1 6E-86 [Chloroherpeton thalassium] RFM23801.1 284 3-hydroxybutyryl-CoA dehydrogenase 0 KXK48806.1 1E-143 same as query [Chlorobi bacterium OLB5] isoprenylcysteine same as query [Nostoc sp. 'Peltigera RFM23811.1 223 carboxylmethyltransferase family protein 0 WP_094342724.1 1.9E-56 membranacea cyanobiont' 232] RFM23700.1 131 MGMT family protein bchG ~20kb RJP64139.1 2.1E-39 same as query [Ignavibacteriales bacterium] RFM23595.1 276 FkbM family methyltransferase 0 WP_133794891.1 3.8E-32 same as query [Prosthecobacter fusiformis] RFM23572.1 275 class I SAM-dependent methyltransferase 0 REJ81906.1 7E-131 same as query[Bacteroidetes bacterium] TPA: hypothetical protein DCY06_01755 RFM23520.1 252 class I SAM-dependent methyltransferase ~2kb from contig edge HAY32834.1 4.4E-78 [Bacteroidetes bacterium] RFM23539.1 142 AraC family transcriptional regulator ~1kb from contig edge WP_109744839.1 1.6E-27 same as query [Arcicella aurantiaca] RFM23509.1 264 class I SAM-dependent methyltransferase bchY ~10kb down TAH40367.1 6E-112 same as query[Bacteroidetes bacterium] RFM23465.1 316 site-specific DNA-methyltransferase 0 WP_069965860.1 3E-157 1220] TPA: class I SAM-dependent RFM23381.1 275 class I SAM-dependent methyltransferase adjacent hemY HAQ20956.1 6.1E-34 methyltransferase [Prolixibacteraceae protoheme IX farnesyl RFM25266.1 201 class I SAM-dependent methyltransferase transferase ~22kb RQW02787.1 2.5E-31 same as query[Calditrichaeota bacterium] MULTISPECIES: same as query RFM23188.1 199 class I SAM-dependent methyltransferase 0 WP_018139937.1 3.5E-79 [Thioalkalivibrio] RFM23159.1 308 site-specific DNA-methyltransferase starting edge of contig WP_103078433.1 2E-127 same as query [Petrotoga miotherma] 23S rRNA (guanosine(2251)-2'-O)- RFM23079.1 155 RNA methyltransferase ~1kb from contig edge WP_084578310.1 4.4E-44 methyltransferase RlmB [Sporomusa methyltransferase domain-containing class I SAM-dependent methyltransferase RFM23081.1 244 protein ~2kb from edge of contig TAE29545.1 7E-113 [Candidatus Kapabacteria bacterium] RFM23087.1 253 class I SAM-dependent methyltransferase 0 WP_092481993.1 7.7E-91 same as query [Desulfallas geothermicus] hypothetical protein A2V82_09620 RFM23090.1 339 class I SAM-dependent methyltransferase 0 OGC01799.1 3.2E-72 [candidate division KSB1 bacterium methyltransferase domain-containing same as query [Hymenobacter sp. RFM23099.1 207 protein 0 WP_086596522.1 2E-99 MIMBbqt21]
RFM23104.1 268 class I SAM-dependent methyltransferase 0 WP_006508154.1 1.2E-60 same as query [Xenococcus sp. PCC 7305] same as query [Nostoc sp. 'Peltigera RFM23049.1 285 class I SAM-dependent methyltransferase hemF ~10kb WP_094340588.1 1.4E-63 membranacea cyanobiont' 232] TPA: hypothetical protein DCL61_16730 RFM23051.1 235 class I SAM-dependent methyltransferase same as above HAG82756.1 4.5E-96 [Cyanobacteria bacterium UBA12227] TPA: hypothetical protein DCM10_08675 RFM23060.1 84 hypothetical protein D0433_13570 hemF ~22kb HAI18071.1 1E-07 [Xanthomarina gelatinilytica] hypothetical protein UU55_C0016G0009 RFM23066.1 95 hypothetical protein D0433_13605 ~5kb from edge of contig KKS02248.1 2.8E-21 [candidate division WWE3 bacterium methyltransferase domain-containing RFM23071.1 217 protein ~200bp from edge of contig WP_044513364.1 2.7E-90 same as query [Hymenobacter sp. DG25B] ferrochelatase ~2kb down; ribosomal small subunit Rsm22 RFM25201.1 335 methyltransferase ~5 kb from edge of contig WP_012500121.1 1E-86 [Chloroherpeton thalassium] methyltransferase domain-containing RFM25014.1 152 protein ~8kb from edge of contig WP_096574832.1 1.9E-60 MULTISPECIES: same as query [Nostocales] putative methyltransferase RFM25075.1 206 class I SAM-dependent methyltransferase 0 BAH40161.1 5.6E-68 [Gemmatimonas aurantiaca T-27] methyltransferase domain-containing ~1.5kb from edge of contig; TPA: methyltransferase type 11 RFM24879.1 271 protein 0.5 kb from sequence edge HCA81768.1 5E-108 [Bacteroidetes bacterium] TPA: hypothetical protein DCQ29_08645 RFM24759.1 252 FkbM family methyltransferase chlG ~8kb HAN38955.1 7E-118 [Chitinophagaceae bacterium] methyltransferase domain-containing hypothetical protein DMF72_05545 RFM24656.1 270 protein 0 PYS24285.1 1.9E-44 [Acidobacteria bacterium] RsmB/NOP family class I SAM-dependent RFM24344.1 446 RNA methyltransferase 0 WP_041468432.1 2E-139 same as query [Chloroherpeton thalassium] 200 Chapter 5 Light Harvesting Apparatus and Pigments of Thiohalocapsa MS
Unpublished data.
Contributions: The characterization of Thiohalocapsa MS and experimental design were
carried out by JLT and this chapter written and all figures made by JLT. Dr. Marcus Tank
collected and isolated Thiohalocapsa MS and provided the isolate for characterization.
DAB provided help with data analysis, and editing of this work.
201 5.1 Abstract
The photosynthetic pigments and light harvesting complexes of a newly isolated
Thiohalocapsa sp. are described in this work. This strain was isolated from the alkaline
silaceous hot spring Mushroom Spring Yellowstone National Park. Thiohalocapsa MS,
as this strain is referred to here, is the first thermophilic member of the genus
Thiohalocapsa and the third thermophilic species of the family Chromatiaceae. The main light-harvesting pigments are identified as bacteriochlorophyll a, spirilloxanthin, and anhydro-rhodovibrin. Two spectral variants of LH2, denoted B800-B830 and B800-
B855, were identified and co-purified from this organism. In addition, a B800-B900
LH1-RC was purified from Thiohalocapsa MS. Like other thermophilic purple sulfur bacteria (PSB) the near infrared absorbance maximum of the Thiohalocapsa MS LH1-RC was above 890 nm. The near-IR absorbance of this LH1-RC complex in response to the addition or chelation of Ca2+ was investigated and shown to be similar to other thermophilic PSB. In the presence of Ca2+ ions LH1-RC had a near-IR absorbance maximum ~895-899 nm while under Ca2+ chelating conditions the maximum was shift to
~888-892 nm. Based on these observations it is likely that like other thermophilic PSB
the LH1-RC complex of Thiohalocapsa MS binds Ca2+ ions that enhance its thermal
stability.
202 5.2 Introduction
5.2.1 Purple Sulfur Bacteria
Purple sulfur bacteria (PSB) are anoxygenic chlorophototrophic members of the
Gammaproteobacteria order Chromatiales. They are represented by two families,
Chromatiaceae and Ectothiorhodospiraceae, which together contain 29 described genera with 74 described species (Thiel et al., 2018). They are typically photoautotrophs which use reduced sulfur compounds as their electron sources and fix carbon via the Calvin
Benson Bassham cycle (Imhoff, 2015a, 2015b). PSB store elemental sulfur in polysulfur globules, with the Chromatiaceae exhibiting internal sulfur globules and the
Ectothiorhodospiraceae external sulfur globules (Imhoff, 2015a, 2015b; Thiel et al.,
2018).
PSB are found in a wide variety of environments in which both light and sulfide are present. They are often found in the anoxic or microoxic zones at or below the chemocline in stratified sulfidic lakes or sediments (Thiel et al., 2018). Unlike the green sulfur bacteria (GSB) (see Chapter 1), which reside strictly in the anoxic zones of such environments, many PSB are oxygen-tolerant and may even be found in the oxic zones above the chemocline, though they require anaerobic or microaerophilic conditions to synthesize bacteriochlorophyll (BChl) and grow phototrophically (Boldareva-Nuianzina et al., 2013; Thiel et al., 2018). PSB typically have a lower affinity for sulfide than GSB, and they often absorb longer wavelengths of light than GSB but require higher irradiance levels due to smaller antenna sizes (Bryant and Canniffe, 2018; Overmann and Garcia-
Pichel, 2013; Thiel et al., 2018; van Gemerden and Mas, 1995). Taken together, these 203 factors typically lead to stratified environments where PSB bacteria are found in the
upper sulfidic layers near the chemocline while GSB, which are well adapted to low-light
conditions, are found in the strictly anoxic lower layers (Overmann and Garcia-Pichel,
2013; van Gemerden and Mas, 1995).
Like the Chloroflexi discussed in Chapter 1, all purple phototrophic bacteria have
type-2 reaction centers. In PSB these reaction centers contain either BChl a (see
Chapters 1 and 2) or BChl b (and BPhe a or b) in addition to various carotenoids
(Bullough et al., 2009; Imhoff, 2017). While two types of BChl are found in PSB, at least
50 different carotenoids have been characterized in anaerobic chlorophototrophic purple
bacteria (Takaichi, 2009). Commonly found carotenoids from PSB include
spirilloxanthin, okenone, lycopene, and rhodopinal in addition to intermediates in the
biosynthetic pathways of these carotenoids (Maresca et al., 2008; Takaichi, 2009; Vogl
and Bryant, 2011). The carotenoid content of PSB is responsible for their typical pinkish
to reddish-purple color (Thiel et al., 2018). As mentioned above, PSB absorb light into
the near-IR range. Monomeric BChl a has a Qy absorbance maximum of 770 nm in
organic solvents while monomeric BChl b has a Qy absorbance maximum of 800 nm
(Robert, 2009). In vivo the absorbance of BChls in the PSB light harvesting apparatus are
shifted to higher wavelengths by pigment-pigment and pigment-protein interactions
(Bryant and Canniffe, 2018; Robert, 2009). BChl a-containing species typically have Qy
absorbance maxima around 800 nm, 820-850 nm, and 870-920 nm. BChl b-containing species have absorbance maxima even further into the near-IR range with Qy peaks
beyond 1000 nm (Robert, 2009). As a result, BChl b-containing species are particularly 204 well suited to sandy soils in which far-red and near infrared wavelengths of light
penetrate to the sulfidic layer (Imhoff, 2017).
The light-harvesting apparatus of purple bacteria is made up of two antenna types
known as the light-harvesting 1 (LH1) complex and the light-harvesting 2 (LH2)
complex. Both LH1 and LH2 complexes are toroidal, integral membrane complexes that
are formed from oligomerization of alpha and beta heterodimers. Each heterodimer and
its associated pigments are known as a protomer. LH1 is closely associated with the RC
in what is typically referred to as the LH1-RC core complex, while LH2 is a peripheral antenna complex which absorbs light and transfers energy to the LH1-RC complex (see
Figure 5.1) (Bryant and Canniffe, 2018; Bullough et al., 2009; Gabrielsen et al., 2009).
The LH1-RC complex is comprised of 14-16 LH1 protomers per RC which encircle the
RC (Bryant and Canniffe, 2018). The LH1 protomer consists of an alpha subunit and a
beta subunit, each a single transmembrane alpha-helix around 50 amino acids in length,
two BChl a (or b) molecules and one carotenoid molecule (Figure 5.1B). In the LH1 complex the BChl a (or b) molecules are tightly coupled and in BChl a-containing strains
these BChl a molecules collectively have a Qy absorbance band around 870-890 nm. LH1
complexes exist in either monomeric or dimeric forms which yield circular, ovate, or S-
shaped complexes, respectively (Bullough et al., 2009; Robert, 2009).
LH2 is a toroidal complex comprised of 8 or 9 protomer units, depending on the
species, although it is important to note that some reports have suggested that a single
species may contain LH2 complexes of both 8 and 9 protomer units (Bryant and
Canniffe, 2018; Gabrielsen et al., 2009). The LH2 protomer consists of alpha and beta 205 subunits, three BChl a molecules and one carotenoid. Two of the BChl a molecules
within the LH2 are perpendicular to the plane of the membrane and are very tightly
coupled (Figure 5.1C). Depending on species, these BChls give rise to a Qy absorbance
band of ~820, 830, or 850 nm. These BChls are collectively referred to as B820, B830, or
B850, respectively. The additional BChl a molecule in the protomer is oriented nearly
parallel to the plane of the membrane and gives rise to a Qy absorbance band around 800
nm—these are the B800 BChls (Robert, 2009). Collectively, each LH2 complex has two spectral types of BChl a and the complexes are named based on these characteristics as follows: B800-B820, B800-B830, B800-B850. Initially, the B800-B850 complex was referred to as LH2 and the B800-B20 complex was called LH3. However, as more complexes have been characterized, it was recognized that peripheral antenna complexes exist with a variety of Qy absorbance peaks, and it has been suggested that all such
complexes be referred to as LH2 and designated by their absorbance features (Gabrielsen
et al., 2009). This convention will be used throughout this chapter.
5.2.2 Newly Isolated Thiohalocapsa sp. from Mushroom Spring, Yellowstone
National Park.
Mushroom Spring in Yellowstone National Park has been used as a model system for the study of chlorophototrophic microbial mat communities for over a decade.
Mushroom Spring is an alkaline, siliceous hot spring with effluent channels ranging from about 69 to 40 °C (Tank et al., 2017). Recently, a Thiohalocapsa species has been isolated from Mushroom Spring. Organisms from the genus Thiohalocapsa are PSB of 206 the family Chromatiaceae which typically grow around 20-30 °C (Imhoff et al., 1998).
This organism will become only the second fully described thermophilic member of the
PSB. Thermochromatium (Tch.) tepidum, which grows optimally at 50°C, is the only
other described species of thermophilic PSB. However, the LH1-RC complex has
recently been characterized from Allochromatium (Alc.) tepidum, which grows optimally
around 45°C. Additionally, a thermotolerant strain related to Marichromatium (Mch.)
gracile has been reported which can grow up to 44°C (Imhoff, 2017; Kimura et al., 2017;
Madigan, 1986; Serrano et al., 2009). This will be only the third Thiohalocapsa spp. described and the first that is thermophilic (Figure 5.2) (Anil Kumar et al., 2009;
Caumette et al., 1991). Here I will describe the light-harvesting pigments and properties of the photosynthetic apparatus of the newly isolated Thiohalocapsa sp., which will be referred to as Thiohalocapsa MS throughout this chapter. I will compare these data with the published descriptions of the pigments and photosynthetic apparatus of the other
Thiohalocapsa spp., Tch. tepidum, and to Alc. tepidum.
5.3 Experimental Procedures
5.3.1 Strains used in this study
Thiohalocapsa MS was isolated from Mushroom Spring, Yellowstone National
Park, WY, USA, by Dr. Marcus Tank and provided for these analyses. Thiohalocapsa
MS was grown at ca. 45 °C in Pfennig’s medium under illumination from incandescent bulbs (Eichler and Pfennig, 1988; Imhoff, 2015). Sulfide levels were monitored by use of lead acetate paper (GE Healthcare Whatman/Fischer Scientific, Göteborg, Sweden) and
cultures were fed with a filter-sterilized sulfide feeding solution as needed to increase the 207 growth yield. The sulfide feeding solution was based upon protocols used in the Imhoff
laboratory. It was made by dissolving 7 g of Na2S • 9H2O and 2.65 g of Na2CO3 in 250
mL H2O and adjusting the pH to 7.3 by addition of H2SO4, the solution was stored in
crimp top bottles with thick butyl rubber stoppers (Imhoff, 2015).
5.3.2 Phylogeny of Purple Sulfur Bacteria
The 16S ribosomal RNA (rRNA) genes of all type strains of the order
Chromatiales based on EMBL taxonomy were downloaded from the Silva rRNA
database SSU release 132 (Quast et al., 2013). Additionally, the 16S rRNA gene
sequence of Tch. tepidum was downloaded from the NCBI database. Sequences were
aligned with the SINA aligner v1.2.11, and the phylogenetic tree was computed using
FastTree (Price et al., 2009; Pruesse et al., 2012). The tree was visualized and modified
using iTol (Letunic and Bork, 2016).
5.3.3 Microscopy
Microscopy was performed on a Nikon Eclipse E400 microscope (Nikon
Corporation, Tokyo, Japan) equipped with a Nikon Digital Sight DS-QiMc camera.
Images were captured using NIS-Elements program version D 4.11.00 from Nikon.
Phase-contrast images were taken using the 100× phase contrast oil immersion objective
and phase contrast wheel set to the Ph3 position. Fluorescence images were obtained by
using a Nikon 100W mercury lamp (LH-M100CB-1) and super high pressure mercury lamp power source (C-SHG1) combined with a custom optical filter set from Chroma
(Chroma Technology Corporation, Bellows Falls, VT) designed for BChl excitation and the subsequent detection of emission by the camera. The excitation filter transmits 550- 208 350 nm light while the emission filter transmits 850-1100 nm light. Fluorescence images
were taken using the same 100× objective, with the phase-contrast wheel set to the closed
position to block out light from the light microscopy source.
5.3.4 Spectroscopic Analysis
UV-visible absorbance spectra of samples were collected on either GENESYS 10
spectrophotometer or a GENESYS 50 spectrophotometer (ThermoFisher Scientific Corp.,
Waltham, MA). Whole cell measurements were performed in a 40% sucrose solution. For pigments in methanol, pigments were extracted in 100% methanol from whole cells prior to measurements. Chromatophore samples were measured in isolation buffer 20 mM 4-
(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 7.5. The LH1 and LH2
samples were measured in isolation buffer with 0.02% n-dodecyl-β-D-malotside (DDM)
(Chem-Impex International Inc. Wood Dale, IL) detergent present with or without the addition of CaCl2, EDTA, or EGTA as indicated in the text.
5.3.5 Pigment Separation and Analysis
Pigments were separated and analyzed by reversed-phase HPLC using a 25 cm ×
4.6-mm Discovery 5-μm C-18 column (Supelco, Bellefonte, PA) and an Agilent series
1100 HPLC system equipped with a diode array detector (Agilent Technologies, Palo
Alto, CA) (Frigaard et al., 1997). The data were analyzed using Agilent ChemStation
software (revision B.02.01-SR1 6100 series). Pigment samples were extracted from
whole cells in 7:2 acetone:methanol and filtered through a 0.2-μm syringe filter, and 10
mM ammonium acetate was added before injection onto the column. A previously
described separation method was used for analysis (Frigaard et al., 1997; Vogl et al. 209 2012). Breifly solvent A was composed of methanol:acetonitrile: water, 42:33:25 v/v/v
and solvent B was composed of methanol:acetonitrile:ethyl acetate, 50:20:30 v/v/v. At 0
min the solvent was 30% B, from 0-52 mins it was linearly increased to 100% B and held
constant for 6 min before washing and subsequent samples.
5.3.6 Isolation and Purification of Light Harvesting Complexes
For LH complex purification, batch cultures (1 L) were grown in Pyrex bottles
with minimal headspace to reduce oxygen exposure and with stirring to provide even
illumination. Cells were harvested by centrifugation at 8,000 × g and resuspended in isolation buffer. Cells were lysed by passage through a chilled French pressure cell, and cell debris was removed by centrifugation at ~15,000 × g for 15 min. The supernatant was collected and chromatophores were harvested by ultracentrifugation with a Beckman
L8-80M ultracentrifuge in a Ti70 rotor at 45,000 rpm (~149,000 × g) for 90 min at 4°C and resuspended in a minimal volume of isolation buffer. LH complexes were solubilized from chromatophore preps by incubation with 2% DDM at room temperature for 1.5 hours in the dark. Unsolubilized material in chromatophore membranes was pelleted in a microcentrifuge at 14,000 rpm (~20,000 × g). The LH1 and LH2 complexes were separated on a sucrose density gradient ranging from 5% to 25% sucrose prepared with
0.02% DDM in the isolation buffer. The density gradients were subjected to ultracentrifugation with a Beckman L8-80M Ultracentrifuge at 29,000 rpm in a Ti70 rotor (~62,000 × g) for 12 h at 4°C. The two pink fractions were collected, dialyzed against isolation buffer, and concentrated using a spin concentrator device with a 3,000 kDa molecular weight cut-off membrane. The concentrated samples were then subjected 210 to a second sucrose gradient from 10% - 22% sucrose in 0.02% DDM in isolation buffer.
The pink fractions were collected, dialyzed and concentrated as above and stored at 4 °C.
Analyses were performed shortly after purification.
5.4 Results
5.4.1 Identification, Growth, and Absorbance Spectra
Liquid cultures of the Thiohalocapsa MS grown in Pfennig’s media at 45 °C are pink in color (Figure 5.3A). The cell density of these cultures increased with regular feeding with a sulfide feeding solution. These observations are consistent with its identification as a PSB. The 16S ribosomal RNA gene was partially sequenced using universal primers 8F and 1492R (Turner et al., 1999). The resulting sequence showed
~96% sequence identity to Thiohalocapsa halophila strain 4270T, and was predicted to be a Thiohalocapsa sp. using the RDP classifier (Wang et al., 2007). Taken together this information led to the identification of Thiohalocapsa MS as a member of the genus
Thiohalocapsa, but did not identify it as belonging to a previously known species of
Thiohalocapsa. When cultures were examined under a microscope, small coccoid to
ovoid cells approximately 2.5-3 μm in diameter were observed; phase-bright sulfide
globules were visible within cells as expected for a Thiohalocapsa sp. (Figure 5.3B).
Additionally, cells were fluorescent when using filters designed to detect BChl- containing organisms, consistent with the presence of BChl a (or BChl b) in LH complexes (Figure 5.3C). Some heterogeneity can be seen in the fluorescence image which is likely due to the plane of focus on the cells and to differences in cell growth stage. 211 To characterize the Thiohalocapsa cultures further, UV-Visible absorbance
spectra were measured for whole cells in 40% sucrose and methanol extracted pigments
(Figure 5.4). The whole-cell spectrum showed peaks at 374, 799, 831, 861, and a shoulder from 893-898 nm consistent with the presence of an LH1-RC and one or two
different LH2 complexes, all containing BChl a (Figure 5.4A). Additionally, a prominent triple peaked feature between 450 and 550 nm was present consistent with the presence of carotenoids. The spectrum of the methanol-extracted pigments showed a Soret peak at
363 nm and a Qy peak at 772 nm which matches the expected absorbance peaks of BChl
a (Figure 5.4B) (Scheer, 2006).
5.4.2 Pigment Analysis
Reversed-phase HPLC was used to analyze pigment extracts in order to confirm the presence of BChl a and identify the carotenoids present in Thiohalocapsa MS and
allow comparison to Rhodospirillum (Rsp). rubrum, a purple non-sulfur bacterium with
previously characterized pigments (Schwerzmann and Bachofen, 1989). Figure 5.5A
shows the elution profile of pigments extracted from Thiohalocapsa MS monitored at 770
nm to detect BChl a, and Figure 5.6A shows the elution profile monitored at 491 nm to
detect carotenoids. In Figure 5.5A a prominent peak can be seen at 35.8 min. The
absorbance spectrum for the 35.8 min peak is shown in Figure 5.5B. Based on the elution
time and absorbance spectrum, this peak was identified as BChl a with phytol as the
esterifying alcohol. 212 In the 491 nm elution profile of Thiohalocapsa MS (Figure 5.6A) prominent
peaks can be seen at 38.8 and 44.8 min. The two major peaks from Thiohalocapsa MS elute at the same time as sprilloxanthin and anhydro-rhodovibrin from Rsp. rubrum. The
absorbance spectra of the 38.8 min peak from Thiohalocapsa MS and the 39.1 min peak
from Rsp. rubrum are compared in Figure 5.6B and the spectra of the 44.8 min peak
from Thiohalocapsa MS and the 45.1 min peak from Rsp. rubrum are compared in
Figure 5.6C. Based on the elution times and in-line absorbance spectra, the 38.8 min peak was identified as spirilloxanthin and the 44.8 min peak was identified as anhydro- rhodovibrin. These pigment analyses show that the major pigments present in
Thiohalocapsa MS are BChl a, spirilloxanthin, and anhydro-rhodovibrin.
5.4.3 Purification and Characterization of LH Complexes
In order to characterize the features of the different LH complexes present in
Thiohalocapsa MS, the LH1-RC and LH2 complexes were solubilized from crude chromatophore preparations and purified via sucrose density gradient. Treatment of the crude chromatophore preparation with 2% (w/v) DDM resulted in solubilization of both the LH1-RC and LH2 complexes as monitored by UV-Vis absorbance. Figure 5.7A shows the UV-Vis absorbance spectrum of the crude chromatophore preparation which has four absorbance features in the far-red and near infrared region at 798, 830, 861, and
892-897 nm attributed to the BChl a molecules within the LH1-RC and LH2 complexes.
The solubilized complexes were separated by sucrose density gradients which showed two prominent pink bands (Figure 5.7B). Based on UV-Vis absorbance spectra, 213 the upper pink band consisted of two spectrally distinct LH2 complexes assigned as
B800-B830 and B800-B855 based on the three absorbance features present at 795 nm,
829 nm and 855 nm (Figure 5.7B). Similarly, the lower pink band was assigned as the
LH1-RC complex based on UV-Vis absorbance features at 798 and 899 nm. Interestingly the carotenoid absorbance features seen between 450-600 nm appear differ slightly for the LH1-RC and the LH2 complexes.
5.4.4 Effect of Ca2+ on the LH1-RC Complex
To determine if the LH1-RC complex of Thiohalocapsa MS was affected by the
2+ presence of Ca ions, purified complexes were treated with chelators or CaCl2 and their
absorbances were monitored by UV-Vis spectroscopy. The UV-Vis absorbance spectra of
LH1-RC complexes showed that Ca2+ ions caused a change in the maxima of the longest
wavelength absorbance feature. (Figure 5.8). In native LH1-RC samples treated with
CaCl2 for three hours, no change was seen in the absorbance maxima of the 898 nm peak
compared to untreated complexes. In contrast, the sample treated with EDTA for three
hours demonstrated a shift of this absorbance peak to 892 nm (Figure 5.8A). When the
incubation was allowed to continue overnight at 4 °C, the native LH1-RC and CaCl2
treated samples showed an absorbance peak at 896 nm while the EDTA treated sample
showed a peak at 891 nm (Figure 5.8B). An additional experiment was carried out with the EDTA-treated sample in which the sample was desalted using a PD-10 column and
CaCl2 added back to this sample. The EDTA-treated and desalted sample showed an absorbance peak at 890 nm while the CaCl2 “rescued” sample showed peak at 895 nm
both in the 5 min and overnight treatments (Figure 5.8C). Finally, a similar experiment 214 was performed that compared native LH1-RC to EGTA-treated LH1-RC. In this experiment the native LH1-RC sample had an absorbance peak at 896 nm, while the sample after 20-min EGTA treatment had an absorbance peak at 888 nm; the sample treated overnight had an absorbance peak at 890 nm (Figure 5.8D) (Table 5.1).
5.5 Discussion
Thiohalocapsa MS shares some physiological features with other Thiohalocapsa spp. For example, Thiohalocapsa are coccoid in shape, produce BChl a, and can have internal sulfur globules (Imhoff et al., 1998). It is also unique in several ways. Firstly, it is the only Thiohalocapsa currently known to grow under thermophilic conditions.
Secondly, Thiohalocapsa MS has as its two main carotenoids spirilloxanthin and anhydro-rhodovibrin while Thc. halophila and Thc. marina are described as producing mainly carotenoids of the okenone series (Caumette et al., 1991; Kumar et al., 2009).
Finally, Thc. halophila and Thc. marina both have absorbance features around 800 nm and 880 nm corresponding to LH1-RC, and each only appears to produce one type of
LH2, B800-830 or B800-845 for Thc. halophile and Thc. marina respectively (Caumette et al., 1991; Kumar et al., 2009). In contrast Thiohalocapsa MS LH1 has absorbance features at 795 nm and 899 nm, the latter significantly red-shifted from other characterized Thiohalocapsa sp. Additionally, Thiohalocapsa MS was shown to have two spectral types of LH2, here designated as B800-B830 and B800-B855. While this has not been seen in other Thiohalocapsa spp., it is not unique to Thiohalocapsa MS. In the PSB
Alc. vinosum, at least three spectral types of LH2 are produced (Carey et al., 2014;
Thornber and Sokoloff, 1970). Carey et al. were able to purify three different LH2 and 215 showed that the relative amounts of each LH2 type changed in response to sulfide, light
intensity, and growth temperature. Considering that Thiohalocapsa MS was isolated from
a microbial mat environment known to contain a diverse ecosystem of phototrophs, it
would be evolutionarily advantageous to have the ability to adjust the absorbance of its
light harvesting apparatus in response to redox state, light availability, and temperature
(Tank et al., 2017). Future experiments comparing the UV-Vis absorbance of
Thiohalocapsa MS under varied growth conditions should expand our understanding of
the regulation of Thiohalocapsa MS LH2 spectral types. Additionally, the genome of Alc.
vinosum revealed that there were six different sequences encoding both the alpha (PucA)
and beta (PucB) subunits of LH2 (Weissgerber et al., 2011). Interestingly, peptide
analysis of purified LH2 complexes from Alc. vinosum showed that each LH2 type
contained a heterogeneous mixture of PucA and PucB paralogs, raising questions about
whether the full array of LH2 types have been studied in this organism and what effect
each polypeptide has on the absorbance features of the LH2 complex (Carey et al., 2014).
It will be interesting to see if the genome of Thiohalocapsa MS also has more copies of
pucA and pucB than the organism has observed LH2 complexes. With the availability of the genome sequence, and the development of a protocol to fully separate B800-B830 and B800-B855, Thiohalocapsa MS may, like Alc. vinosum, become a useful organism for studying the diversity and regulation of LH2 complexes in PSB.
Thiohalocapsa MS differs from other Thiohalocapsa spp. in its carotenoid content and LH1-RC complexes; however, it is similar to other thermophilic or thermotolerant
PSB in both respects. Mch. gracile strain SW26 was characterized as having carotenoids 216 in the spirilloxanthin series, although its exact carotenoid content was not determined
(Serrano et al., 2009). Tch. tepidum and Alc. tepidum also both contain carotenoids of the
spirilloxanthin series with their two main carotenoids being spirilloxanthin and rhodopin
or spirilloxanthin (Kimura et al., 2017; Suzuki et al., 2007). Both organisms also
contained smaller amounts of lycopene, anhydrorhodovibrin, and other intermediates in
the spirilloxanthin pathway. The crystal structure of Tch. tepidum LH1-RC shows the
presence of spirilloxanthin in the LH1-RC, and in Alc. tepidum spirilloxanthin was the carotenoid found in the highest amount in LH1-RC. (Kimura et al., 2017; Niwa et al.,
2014; Yu et al., 2016, 2018) With this in mind, it is interesting that Thiohalocapsa MS also produces spirilloxanthin as one of its main carotenoids. In addition to spirilloxanthin series carotenoids, all thermophilic PSB characterized so far have an LH1-RC which absorbs beyond 890 nm (Kimura et al., 2017, Madigan, 1986). It is important to note that the whole cell spectrum of Mch. gracile did not possess a feature related to LH1 BChl a, likely the LH1-RC complex was produced at low amounts under the growth conditions employed and was not obvious in the absorbance spectra (Serrano et al., 2009). The native LH1-RC complexes of Tch. tepidum and Alc. tepidum have been reported to be
B915 and B890 complexes respectively. This work shows that Thiohalocapsa MS native
LH1-RC has near-IR features at 798 nm and 899 nm, and should be designated as B900.
Similar to the relationship between Thiohalocapsa MS and other Thiohalocapsa sp. LH1-
RC, which have B900 and B880 LH1-RC complexes respectively, Alc. vinosum has LH1 complex with a BChl a absorbance maximum at 884 nm while Alc. tepidum has an absorbance maximum at 890 nm (Caumette et al., 1991; Kimura et al., 2017; Kumar et al., 2009). Perhaps the most interesting feature of the two previously characterized LH1- 217 RC complexes of thermophilic PSB is their relationship to Ca2+ ions. This relationship was first hinted at in Tch. tepidum when it was shown that the LH1 BChl a feature was quite different in LH1-RC complexes purified via anion exchange chromatography in comparison to purification by sucrose density gradient centrifugation (Fathir et al., 1998).
The complexes purified on sucrose density gradients had an absorbance maximum at 915 nm while the complexes purified by anion-exchange chromatography had a maximum at
885 nm. In follow-up studies, LH1-RC was treated with CaCl2 or EDTA and the
absorbance maximum for the complexes shifted from ~915 nm to ~880 nm, a shift of ~
35 nm (Kimura et al., 2008). Subsequently Ca2+ was found to be bound to the LH1-RC
complexes in the crystal structure from Tch. tepidum near the BChl a binding site (Niwa
et al., 2014). Recently the Alc. tepidum LH1-RC has been shown to have a similar
response to CaCl2 and EDTA treatments with a shift in complex absorbance maxima
from 890 nm to 882 nm a shift of 8 nm (Kimura et al., 2017). Here it is reported that
Thiohalocapsa MS LH1-RC shows a similar response to Ca2+ ions. Treatment with
EDTA showed a shift of blue-shift of 5-6 nm which could be rescued by adding CaCl2
back to the complex, while treatment with EGTA showed a blue-shift of 6-8 nm.
Additionally, Ca2+ ions have been shown to increase the thermal stability of LH1-RC
from Alc. tepidum and Thc. tepdium (Kimura et al., 2017). Based on the change in the
near-IR absorbance maximum of Thiohalocapsa MS LH1-RC and the analyses of LH1-
RC from Tch. tepdium and Alc. tepdium, it seems likely that Ca2+ ions are bound to LH1-
RC from Thiohalocapsa MS and that they enhance the thermally stability of the LH1-RC complex. 218 Thiohalocapsa MS is the first member of the Thiohalocapsa genus that grows
under thermophilic conditions. Based on characterization of the pigments and light
harvesting complexes of this newly isolated organism and upon comparison to
mesophilic Thiohalocapsa and thermophilic PSB, it is likely that several features of the
photosynthetic apparatus in Thiohalocapsa MS are related to growth at elevated
temperatures. These include the predominance of spirilloxanthin series carotenoids, an
LH1 BChl a absorbance maximum beyond 890 nm that might be beneficial in the
microbial mats in which it occurs, and a shift in that absorbance maximum in response to
the loss of Ca2+ ions. Further insights into the relationship between LH1-RC of
Thiohalocapsa MS and other themophilic PSB will also likely be gained from the upcoming genome sequence of Thiohalocapsa MS. Additionally, a full physiological description of Thiohalocapsa MS will reveal whether other aspects of the Thiohalocapsa
MS physiology are similar to other Thiohalocapsa spp. or to other thermophilic PSB. 219 References Boldareva-Nuianzina, E. N., Bláhová, Z., Sobotka, R., and Koblízek, M. (2013). Distribution and origin of oxygen-dependent and oxygen-independent forms of Mg- protoporphyrin monomethylester cyclase among phototrophic proteobacteria. Appl. Environ. Microbiol. 79, 2596–2604. doi:10.1128/AEM.00104-13. Bryant, D. A., and Canniffe, D. P. (2018). How nature designs light-harvesting antenna systems: design principles and functional realization in chlorophototrophic prokaryotes. J. Phys. B At. Mol. Opt. Phys. 51, 033001. doi:10.1088/1361- 6455/aa9c3c. Bullough, P. A., Qian, P., and Hunter, C. N. (2009). Reaction center-light-harvesting core complexes of purple bacteria, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), pp. 155–179. doi:10.1007/978-1-4020-8815-5_9. Carey, A.M., Hacking, K., Picken, N., Honkanen, S., Kelly, S., Niedzwiedzki, D.M. et al. (2014). Characterisation of the LH2 spectral variants produced by the photosynthetic purple sulphur bacterium Allochromatium vinosum. Biochimica et Biophysica Acta. 1837, 1849-1860. doi:10.1016/j.bbabio.2014.07.022. Caumette, P., Baulaigue, R., and Matheron, R. (1991). Thiocapsa halophila sp. nov., a new halophilic phototrophic purple sulfur bacterium. Arch. Microbiol. doi:10.1007/BF00248613. Eichler B., and Pfennig N. (1988). A new green sulfur bacterium from a freshwater pond. In: Olson JM, Stackebrandt E, Trüper H (eds) Green photosynthetic bacteria. Plenum, New York, pp. 233–235 Frigaard, N. U., Takaichi, S., Hirota, M., Shimada, K., and Matsuura, K. (1997). Quinones in chlorosomes of green sulfur bacteria and their role in the redox- dependent fluorescence studied in chlorosome-like bacteriochlorophyll c aggregates. Arch. Microbiol. 167, 343–349. doi:10.1007/s002030050453. Gabrielsen, M., Gardiner, A. T., and Cogdell, R. J. (2009). Peripheral complexes of purple bacteria, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), pp. 135–153. doi:10.1007/978-1-4020-8815-5_8. Imhoff, J. F. (2015a). “Chromatiaceae,” in Bergey’s Manual of Systematics of Archaea and Bacteria (American Cancer Society), 1–12. doi:10.1002/9781118960608.fbm00219. Imhoff, J. F. (2015b). Ectothiorhodospiraceae, in Bergey’s Manual of Systematics of Archaea and Bacteria (American Cancer Society), 1–5. doi:10.1002/9781118960608.fbm00220. Imhoff, J. F. (2017). Diversity of anaerobic anoxygenic phototrophic purple bacteria, in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, ed. P. C. Hallenbeck (Cham: Springer International Publishing), pp. 47–85. doi:10.1007/978-3-319-46261-5_2. Imhoff, J. F., Suling, J., and Petri, R. (1998). Phylogenetic relationships among the Chromatiaceae, their taxonomic reclassification and description of the new genera Allochromatium, Halochromatium, Isochromatium, Marichromatium, Thiococcus, 220 Thiohalocapsa and Thermochromatium. Int. J. Syst. Evol. Microbiol. 48, 1129–1143. doi:10.1099/00207713-48-4-1129. Kimura, Y., Lyu, S., Okoshi, A., Okazaki, K., Nakamura, N., Ohashi, A., et al. (2017). Effects of calcium ions on the thermostability and spectroscopic properties of the LH1-RC complex from a new thermophilic purple bacterium Allochromatium tepidum. J. Phys. Chem. B. doi:10.1021/acs.jpcb.7b03341. Kumar, A. P., Srinivas, T. N. R., Thiel, V., Tank, M., Sasikala, C., Ramana, C. V, et al. (2009). Thiohalocapsa marina sp. nov., from an Indian marine aquaculture pond. Int. J. Syst. Evol. Microbiol. 59, 2333–2338. doi:10.1099/ijs.0.003053-0. Letunic, I., and Bork, P. (2016). Interactive tree of life (iTOL) v3: an online tool for the display and annotation of phylogenetic and other trees. Nucleic Acids Res. 44, W242–W245. doi:10.1093/nar/gkw290. Madigan, M. T. (1986). Chromatium tepidum sp. nov., a thermophilic photosynthetic bacterium of the family Chromatiaceae. Int. J. Syst. Evol. Microbiol. 36, 222–227. doi:10.1099/00207713-36-2-222. Maresca, J. A., Graham, J. E., and Bryant, D. A. (2008). The biochemical basis for structural diversity in the carotenoids of chlorophototrophic bacteria. Photosynth. Res. 97, 121–140. doi:10.1007/s11120-008-9312-3. Overmann, J., and Garcia-Pichel, F. (2013). The phototrophic way of life, in The Prokaryotes: Prokaryotic Communities and Ecophysiology. doi:10.1007/978-3-642- 30123-0_51. Papiz, M.Z., Prince, S.M., Howard, T., Cogdell, R.J., Isaacs, N.W. (2003) The structure and thermal motion of the B800-850 LH2 complex from Rps. acidophila at 2.0A resolution and 100K: new structural features and functionally relevant motions. J Mol Biol. 326, 1523-1538.doi: 10.1016/s0022-2836(03)00024-x Price, M. N., Dehal, P. S., and Arkin, A. P. (2009). Fasttree: Computing large minimum evolution trees with profiles instead of a distance matrix. Mol. Biol. Evol. 26, 1641– 1650. doi:10.1093/molbev/msp077. Pruesse, E., Peplies, J., and Glöckner, F. O. (2012). SINA: Accurate high-throughput multiple sequence alignment of ribosomal RNA genes. Bioinformatics 28, 1823– 1829. doi:10.1093/bioinformatics/bts252. Quast C., Pruesse E., Yilmaz P., Gerken J., Schweer T., Yarza P., Peplies J., Glöckner F.O. (2013). The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucl. Acids Res. 41, D590-D596. Robert, B. (2009). Spectroscopic properties of antenna complexes from purple bacteria, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), 199–212. doi:10.1007/978-1- 4020-8815-5_11. Schwerzmann, R.U., and Bachofen R. (1989) Carotenoid profiles in pigment-protein complexes of Rhodospirillum rubrum. Plant Cell Physio. 30, 497-504. Serrano W., Amann R., Fischer U. (2009). A new moderately thermophilic and high sulfide tolerant biotype of Marichromatium gracile, isolated from tidal sediments of the German Wadden Sea: Marichromatium gracile biotype thermosulfidiphilum. Syst. Appl. Microbiol. 32, 1-7. doi:10.1016/j.syapm.2008.11.002. 221 Scheer, H. (2006). “An overview of chlorophylls and bacteriochlorophylls: Biochemistry, biophysics, functions and applications,” in Advances in Photosynthesis and Respiration, Vol. 25, eds. B. Grimm, R. J. Porra, W. Rudiger, and H. Scheer (Dordrecht, Netherlands: Springer), 1–26. Suzuki H., Hirano Y., Kimura Y., Takaichi S., Kobayashi M., Miki K., Wang Z.Y. (2007). Purification, characterization and crystallization of the core complex from thermophilic purple sulfur bacterium Thermochromatium tepidum. Biochim Biophys Acta. 1767,1057-63. doi: 10.1016/j.bbabio.2007.06.002 Takaichi, S. (2009). Distribution and biosynthesis of carotenoids, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), pp. 97–117. doi:10.1007/978-1-4020- 8815-5_6. Tank, M., Thiel, V., Ward, D. M., and Bryant, D. A. (2017). A panoply of phototrophs: An overview of the thermophilic chlorophototrophs of the microbial mats of alkaline siliceous hot springs in Yellowstone National Park, WY, USA, in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, ed. P. C. Hallenbeck (Berlin: Springer), pp. 87–137. doi:10.1007/978-3-319-46261-5_3. Thiel, V., Tank, M., and Bryant, D. A. (2018). Diversity of chlorophototrophic bacteria revealed in the omics era. Annu. Rev. Plant Biol. 69, 21–49. doi:10.1146/annurev- arplant-042817-040500. Thornber, J.P. and Sokoloff, M.K. (1970). Photochemical reactions of purple bacteria as revealed by studies of 3 spectrally different carotenobacteriochlorophyll-protein complexes isolated from Chromatium strain D. Biochemistry 9, 2688-2698. doi:10.1021/bi00815a017. Turner, S., Pryer, K.M., Miao, V.P.W., and Palmer, J.D. (1999). Investigating deep phylogenetic relationships among cyanobacteria and plastids by small subunit rRNA sequence analysis. Journal of Eukaryotic Microbiol. 46, 327–338. van Gemerden, H., and Mas, J. (1995). Ecology of phototrophic sulfur bacteria, in Advances in Photosynthesis and Respiration Vol. 2 Anoxygenic photosynthetic bacteria, eds. R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Dordrecht, Netherlands: Springer), pp. 49–85. Vogl, K., and Bryant, D. A. (2011). Elucidation of the Biosynthetic Pathway for Okenone in Thiodictyon sp. CAD16 Leads to the Discovery of Two Novel Carotene Ketolases. J. Biol. Chem. 286, 38521–38532. doi:10.1074/jbc.M111.280131. Vogl, K., Tank, M., Orf, G. S., Blankenship, R. E., and Bryant, D. A. (2012). Bacteriochlorophyll f: properties of chlorosomes containing the “forbidden chlorophyll.” Front. Microbiol. 3, 1–12. doi:10.3389/fmicb.2012.00298. Wang Q., Garrity G.M., Tiedje J.M., Cole J.R. (2007). Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 73, 5261-5267. doi:10.1128/AEM.00062-07. Weissgerber, T., Zigann, R., Bruce, D., Chang, Y.J., Detter, J.C., Han, C., Hauser, L., et al., (2011). Complete genome sequence of Allochromatium vinosum DSM 180(T). Stand. Genomic Sci. 5, 311-330. Yu, L.J., Kawakami, T., Kimura, Y., Wang-Otomo, Z.Y. (2016). Structural basis for the unusual Qy red-shift and enhanced thermostability of the LH1 Complex from 222 Thermochromatium tepidum. Biochemistry. 55, 6495-6504. doi: 10.1021/acs.biochem.6b00742 Yu, L.J., Suga, M., Wang-Otomo, ZY., Shen, J.R. (2018) Structure of photosynthetic LH1-RC supercomplex at 1.9 Å resolution. Nature. 556, 209-213. doi: 10.1038/s41586-018-0002-9.
223
Figure 5.1 Structures of Light-Harvesting and Reaction Complexes of a PSB. A. Representation of the arrangement of LH1-RC and LH2 in the membrane of a hypothetical purple bacterium. LH1-RC is represented by the crystal structure from Thc. tepidum (PDB:5Y5S) and LH2 is represented by the crystal structure of Rps. acidophila (PDB1NKZ) (Papiz et al., 2003; Yu et al., 2018). Subunits and their ligands are labled in the same color as they appear in the figure. BChl a in LH1-RC and B850 BChl a in LH2 are colored in cyan, B800 BChl a in LH2 is colored in magenta. B. Top and side views of LH1stucture with the RC and cytochrome subunits removed so that BChl a and spirilloxanthin bound to the LH1 protomers can be seen. C. Top and side view of LH1 so that orientation of both spectral types of BChl a can be viewed. 224
Figure 5.2 16S rRNA Phylogenetic Tree of the Order Chromatiales Phylogeny of the order Chromatiales. Members of the family Chromatiaceae are colored purple based on EMBL taxonomy, while members of the Ectothiorhodospiraceae are colored in turquoise. Branches containing PSB are highlighted in salmon colored boxes. * marks thermophilic or thermotolerant PSB or their closest relatives. 225
B C
A
Figure 5.3 Appearance of Thiohalocapsa MS A. Culture of Thiohalocapsa MS in crimp top vial. B. Phase-contrast micrograph of Thiohalocapsa MS. Red arrow points to phase-bright internal sulfide globule.C-Fluorescence image of the same cells as in panel B showing cells containing BChl a. Image taken using a custom optical filter set designed for BChl excitation and emission. The excitation filter transmits 550-350 nm light while the emission filter transmits 850-1100 nm light. 226
Figure 5.4 UV-Vis Absorbance Spectra A. Spectrum of whole cells suspended in 40% sucrose solution. B. Spectrum of pigments extracted in 100% methanol from whole cells.
227
Figure 5.5 Elution profile from reversed-phase HPLC monitored at 770 nm. A. The elution profile of pigments extracted from Thiohalocapsa MS monitored at 770 nm to detect BChl a. B. The in-line absorbance spectrum of the 35.8-min peak shown in the elution profile from panel A. 228
Figure 5.6 Elution profiles from reversed-phase HPLC monitored at 491 nm. A. The elution profile of pigments extracted from Thiohalocapsa MS and Rsp. rubrum monitored at 491 nm to detect carotenoids. B. The in-line absorbance spectra of the ~39-min peaks shown in the elution profile from panel A. C. The in-line absorbance spectra of the ~45-min peaks shown in panel A. 229
Figure 5.7 Chromatophores and LH complexes of Thiohalocapsa MS. A. Image of chromatophore pellet after centrifugation and the UV-Vis absorbance spectrum of the chromatophore preparation. B. Image of primary sucrose density gradient showing pink bands corresponding to LH1-RC and LH2 and UV-Vis absorbance spectra of LH1 and LH2 purified complexes after second sucrose density gradient.
230
Figure 5.8 LH1-RC response to CaCl2 and chelators A. Comparison of absorbance spectra of native LH1-RC, treated for 3 h with CaCl2, and LH1-RC treated for 3 h with EDTA. B. Comparison of the absorbance spectra of the same experiment in panel A allowed to run overnight (O/N; ~18 h). C. Comparison of the absorbance spectra of EDTA-treated and desalted LH1-RC when CaCl2 was added back. Measurements were taken at 5 min and after incubation overnight. D.Comparison of UV-vis absorbance spectra of native LH1-RC and EGTA-treated LH1-RC. Measurements were taken at 20-min and overnight (O/N) time points.
231 Table 5.1 Effect of Ca2+ on LH1-RC from Thermophillic PSB Organism Near-IR Maximum Near-IR Maximum Shift of Near-IR with Ca2+ (nm) without Ca2+ (nm) Maximum (nm) Tch. tepidum 915a; 913b 880a; 876b 35a; 37b Alc. tepiduma 890 882 8 Thiohalocapsa MSc 895-899 888-892 5-8 aKimura et al., 2017 bKimura et al., 2008 cThis work.
232 Chapter 6 Concluding Remarks
6.1 Overview and Discussion
Anoxygenic phototrophic bacteria have been confirmed in six bacterial phyla, and recent genome sequence data points to their presence in two more phyla (Tahon and Willems, 2017;
Thiel et al., 2018; Ward et al., 2019). These organisms are important to our understanding of the evolution of photosynthesis, primary productivity on both modern Earth and in the past, the limits of phototrophy on Earth, and alternative energy sources (Imhoff, 2017; Judson, 2017;
Martin et al., 2017; Saer and Blankenship 2017). In this work anoxygenic phototrophy is explored through looking at pigment biosynthesis in green bacteria, and characterization of the pigments and light-harvesting apparatus of a newly isolated purple sulfur bacterium.
Green bacteria produce chlorosomes, which are the most efficient light harvesting antenna produced by any phototrophic organism (Saer and Blankenship, 2017). In fact, the phototroph currently known for living at the lowest irradiance on Earth is a brown-colored green sulfur bacterium (Manske, et al., 2005). All green bacteria produce at least BChl a and BChl c, d, e, or f, and Cab. thermophilum and the green Chlorobi additionally produce Chl a (Chapter 1).
The availability of sequenced genomes along with information on pigment composition of organisms has made it possible to perform useful bioinformatic analyses on the (B)Chl biosynthetic pathways in green bacteria (Chapter 2 and Chapter 4) (Thweatt et al., 2019). In this work these analyses showed that most green bacteria, previously thought to have two functional copies of HemN that were differentially regulated based on growth conditions, in fact have one copy of HemN and one copy of a heme chaperone known as HemW (Abicht et al.,
2012; Eisen et al., 2002; Haskamp et al., 2018; Tang et al., 2011). It also revealed that 233 surprisingly, newly characterized anaerobic green Chloroflexi, which encode BchQ and BchR,
only possess a single BchF homolog unlike the green Chlorobi that have multiple BchF
homologs (Bryant and Liu 2013; Gomez Maqueo Chew et al., 2007; Grouzdev et al., 2018).
Phylogentic analysis of the three different types of (B)Chl synthases in green bacteria also
revealed that a chlorin synthase clade appears to predate the bacteriochlorin synthase clade,
which agrees with the hypothesis that Chls predated BChls in the evolution of phototrophy
(Bryant et al., 2012; Bryant and Liu, 2013; Gomez Maqueo Chew and Bryant, 2007). New
questions were also raised through this work, like the nature of the HemY enzymes in green
bacteria and the function of genes previously annotated as encoding BchQ and BchR in Cab.
thermophilum (Garcia Costas et al., 2012). Another significant finding was that even with the publication of three metagenome assembled genomes, no gene encoding a known enzyme for formation of the isocyclic-E ring has been identified in Ca. T aerophilum. This suggests that a
novel enzyme may yet be identified in this organism. These questions will likely only be
answered by biochemical characterization and careful genetic manipulations like those used in
Chapter 3 to confirm the function of BciD (Thweatt et al., 2017).
Prior to this work, bioinformatic analysis had determined that BciD was a putative radical-SAM enzyme and a candidate for catalyzing the reaction converting BChlide c or d to
BChlide e or f, while genetic manipulation had confirmed that it was required for production of
BChl e in vivo (Harada et al., 2013). In this work the in vivo phenotype was confirmed and
biochemical characterization determined that BciD is a radical-SAM enzyme containing a 4Fe-
4S cluster that is sufficient to convert BChlide c or d to BChlide e or f. It is proposed that this
reaction progresses via consecutive hydroxylation reactions which form a geminal-diol that spontneously dehdrates to form a formyl group (Thweatt et al., 2017). Further biochemical 234 analyses will be needed to confirm this hypothesis and determine the biochemical mechanism for this reaction. Additionally, confirmation of the function of this enzyme should inform future genetic manipulations in GSB which aim to convert green-colored GSB to brown-colored GSB in order to better understand the different properties of chlorosome possessing these pigments.
As mentioned above brown-colored green sulfur bacteria can live in very low light irradiances, and they are also the only organisms which produce BChl e. Continued research into these organisms is of interest to our understanding of the lower light limit of phototrophy on Earth and has implications for the design of artificial light harvesting systems in the field of alternative energy research. Additionally, these studies help to define the habitats we might consider capable of supporting phototrophy elsewhere in the universe (Saer and Blankenship, 2017).
The final chapter in this dissertation describes the characterization of a newly isolated purple sulfur bacterium (PSB) from Mushroom Spring in Yellowstone National Park (Chapter
5). Mushroom Spring has microbial mats that contain a great diversity of anoxygenic phototrophs (Tank et al., 2017). The newly isolated PSB, Thiohalocapsa MS, is the first thermophilic organism identified in the genus Thiohalocapsa and only the third thermophile identified among PSB (Caumette et al., 1991; Kumar et al., 2009). Like other Thiohalocapsa spp., it is an ovoid or coccoid organism that has internal sulfur globules and produces BChl a as its main light-harvesting pigment (Imhoff et al., 1998). However, unlike other Thiohalocapsa spp. it grows well at 45°C, uses carotenoids of the spirilloxanthin series, produces two spectral forms of LH2, and has an LH1-RC complex which absorbs light beyond 890 nm. In light of these differences, its carotenoids and the properties of its LH1-RC were compared to the other thermophilic and thermotolerant PSB (Madigan, 1986; Kimura et al., 2017, Serrano et al., 2009).
It was similar to other thermophilic PSB that also produce carotenoids of the spirilloxanthin 235 series and produce LH1-RC complexes that absorb beyond 890 nm (Kimura et al., 2017; Serrano
et al., 2009; Suzuki et al., 2007). Additionally, it was demonstrated that like other thermophilic
PSB, the LH1-RC complex near-IR absorbance maximum experiences a blue-shift in response to the loss of Ca2+ ions (Kimura et al., 2008; Kimura et al., 2017). Based on this information it is
hypothesized that, like other thermophilic PSB, the LH1-RC complex binds Ca2+ ions, which
enhance its thermostability and thus allow the organism to grow under thermophilic conditions.
Future work will be needed to test this hypothesis. Additional work characterizing the full range
of growth physiology and the forthcoming genome sequence of Thiohalocapsa MS will provide
more information on how Thiohalocapsa MS compares to other Thiohalocapsa spp. and other thermophilic PSB. Currently Tch. tepidum is the most thermophilic PSB and has an LH1-RC
absorbance maximum that is farthest shifted to the near-IR range, around 915 nm (Kimura et al.,
2017). Characterization of more thermophilic PSB and studies of their LH1-RC complexes will
help to provide more information on what limits the temperature range of these organisms.
Finally, it is interesting to note that the phototrophic microbial mat in Mushroom Spring has been
extensively studied for decades, first by classical microbial ecology and physiology and more
recently by metagenomic methods, and yet new chlorophototrophs with interesting properties are
still being discovered and characterized from the mat community of this hot spring(Tank et al.,
2017).
In summary, the studies presented here add to our understanding of specific anoxygenic
chlorophototrophs which live in low-light or high-temperature environments and present new questions for future work on these subjects. They also highlight the importance of combining biochemical characterization, microbial physiology, and new genomic and bioinformatic tools in the study of anoxygenic chlorophototrophy. 236 References
Abicht, H. K., Martinez, J., Layer, G., Jahn, D., and Solioz, M. (2012). Lactococcus lactis HemW (HemN) is a haem-binding protein with a putative role in haem trafficking. Biochem. J. 442, 335–343. doi:10.1042/BJ20111618 Caumette, P., Baulaigue, R., and Matheron, R. (1991). Thiocapsa halophila sp. nov., a new halophilic phototrophic purple sulfur bacterium. Arch. Microbiol. 155, 170–176. doi:10.1007/BF00248613. Eisen, J. A, Nelson, K. E., Paulsen, I. T., Heidelberg, J. F., Wu, M., Dodson, R. J., et al. (2002). The complete genome sequence of Chlorobium tepidum TLS, a photosynthetic, anaerobic, green-sulfur bacterium. Proc. Natl. Acad. Sci. USA 99, 9509–9514. doi:10.1073/pnas.132181499. Harada, J., Mizoguchi, T., Satoh, S., Tsukatani, Y., Yokono, M., Noguchi, M., et al. (2013). Specific gene bciD for C7-methyl oxidation in bacteriochlorophyll e biosynthesis of brown-colored green sulfur bacteria. PLoS One 8, e60026. doi:10.1371/journal.pone.0060026 Haskamp, V., Karrie, S., Mingers, T., Barthels, S., Alberge, F., Magalon, A., et al. (2018). The radical SAM protein HemW is a heme chaperone. J. Biol. Chem. 293, 2558–2572. doi:10.1074/jbc.RA117.000229. Imhoff, J. F. (2017). “Anoxygenic phototrophic bacteria from extreme environments,” in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, P.C. Hallenbeck, ed. (Berlin: Springer), pp. 427–480. doi:10.1007/978-3-319-46261- 5_13. Imhoff, J. F., Suling, J., and Petri, R. (1998). Phylogenetic relationships among the Chromatiaceae, their taxonomic reclassification and description of the new genera Allochromatium, Halochromatium, Isochromatium, Marichromatium, Thiococcus, Thiohalocapsa and Thermochromatium. Int. J. Syst. Evol. Microbiol. 48, 1129–1143. doi:10.1099/00207713-48-4-1129. Judson, O. P. (2017) The energy expansions of evolution. Nature Ecology & Evolution. 1, 0138 Kimura, Y., Hirano, Y., Yu, L.J., Suzuki, H., Kobayashi, M., Wang, Z.Y. (2008) Calcium ions are involved in the unusual red shift of the light-harvesting 1 Qy transition of the core complex in thermophilic purple sulfur bacterium Thermochromatium tepidum. J. Biol. Chem. 283,13867–13873. doi: 10.1074/jbc.M800256200. Kimura, Y., Lyu, S., Okoshi, A., Okazaki, K., Nakamura, N., Ohashi, A., et al. (2017). Effects of calcium ions on the thermostability and spectroscopic properties of the LH1-RC complex from a new thermophilic purple bacterium Allochromatium tepidum. J. Phys. Chem. B. 121, 5025-5032. doi:10.1021/acs.jpcb.7b03341. Kumar, A. P., Srinivas, T. N. R., Thiel, V., Tank, M., Sasikala, C., Ramana, C. V, et al. (2009). Thiohalocapsa marina sp. nov., from an Indian marine aquaculture pond. Int. J. Syst. Evol. Microbiol. 59, 2333–2338. doi:10.1099/ijs.0.003053-0. Madigan, M. T. (1986). Chromatium tepidum sp. nov., a thermophilic photosynthetic bacterium of the family Chromatiaceae. Int. J. Syst. Evol. Microbiol. 36, 222–227. doi:10.1099/00207713-36-2-222. 237 Manske, A. K., Glaeser, J., Kuypers, M. M. M., and Overmann, J. (2005). Physiology and phylogeny of green sulfur bacteria forming a monospecific phototrophic assemblage at a depth of 100 meters in the Black Sea. Appl. Environ. Microbiol. 71, 8049–8060. doi:10.1128/AEM.71.12.8049-8060.2005. Martin, W. F.., Bryant, D. A., Beatty, J. T. (2018) A physiological perspective on the origin and evolution of photosynthesis FEMS Microbiol. Rev. 42, 205–231, doi:10.1093/femsre/fux056 Saer, R. G., and Blankenship, R. E. (2017). Light harvesting in phototrophic bacteria: structure and function. Biochem. J. 474, 2107–2131. doi:10.1042/BCJ20160753. Serrano W., Amann R., Fischer U. (2009). A new moderately thermophilic and high sulfide tolerant biotype of Marichromatium gracile, isolated from tidal sediments of the German Wadden Sea: Marichromatium gracile biotype thermosulfidiphilum. Syst. Appl. Microbiol. 32, 1–7. doi:10.1016/j.syapm.2008.11.002 Suzuki H., Hirano Y., Kimura Y., Takaichi S., Kobayashi M., Miki K., Wang Z.Y. (2007). Purification, characterization and crystallization of the core complex from thermophilic purple sulfur bacterium Thermochromatium tepidum. Biochim Biophys Acta 1767,1057–63. doi: 10.1016/j.bbabio.2007.06.002 Tahon, G., and Willems, A. (2017). Isolation and characterization of aerobic anoxygenic phototrophs from exposed soils from the Sør Rondane Mountains, East Antarctica. Syst. Appl. Microbiol. 40, 357–369. doi: 10.1016/j.syapm.2017.05.007 Tang, K. H., Barry, K., Chertkov, O., Dalin, E., Han, C. S., Hauser, L. J., et al. (2011). Complete genome sequence of the filamentous anoxygenic phototrophic bacterium Chloroflexus aurantiacus. BMC Genomics 12, 334. doi:10.1186/1471-2164-12-334. Tank, M., Thiel, V., Ward, D. M., and Bryant, D. A. (2017). A panoply of phototrophs: An overview of the thermophilic chlorophototrophs of the microbial mats of alkaline siliceous hot springs in Yellowstone National Park, WY, USA, in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, ed. P. C. Hallenbeck (Berlin: Springer), pp. 87–137. doi:10.1007/978-3-319-46261-5_3. Thiel, V., Tank, M., and Bryant, D. A. (2018). Diversity of chlorophototrophic bacteria revealed in the omics era. Annu. Rev. Plant Biol. 69, 21–49. doi:10.1146/annurev- arplant-042817-040500. Thweatt, J.L., Canniffe, D.P., and Bryant, D.A. (2019). Biosynthesis of chlorophylls and bacteriochlorophylls in green bacteria. In: Advances in Botanical Research Vol. 90 Metabolism, Structure and Function of Plant Tetrapyrroles Introduction, Microbial and Eukaryotic Chlorophyll Synthesis and Catabolism: Bernhard Grimm, Ed. (Academic Press, London, UK), pp. 35–89. Thweatt, J. L., Ferlez, B. H., Golbeck, J. H., and Bryant, D. A. (2017). BciD is a radical S-adenosyl-L-methionine (SAM) enzyme that completes bacteriochlorophyllide e biosynthesis by oxidizing a methyl group into a formyl group at C-7. J. Biol. Chem. 292, 1361–1373. doi:10.1074/jbc.M116.767665. Ward, L.M., Cardona, T., and Holland-Moritz H. (2019). Evolutionary implications of anoxygenic phototrophy in the bacterial phylum Candidatus Eremiobacterota (WPS- 2). Front. Microbiol. 10:1658. doi: 10.3389/fmicb.2019.01658 Yu, L.J., Kawakami, T., Kimura, Y., Wang-Otomo, Z.Y. (2016). Structural basis for the unusual Qy red-shift and enhanced thermostability of the LH1 Complex from 238 Thermochromatium tepidum. Biochemistry 55, 6495–6504. doi: 10.1021/acs.biochem.6b00742 Yu, L.J., Suga, M., Wang-Otomo, ZY., Shen, J.R. (2018) Structure of photosynthetic LH1-RC supercomplex at 1.9 Å resolution. Nature 556, 209–213. doi: 10.1038/s41586-018-0002-9.
239
Appendix A. Co-culture Growth Studies of Prosthecochloris sp. HL-130-GSB and
Geobacter sulfurreducens PCA
A.1 Abstract
A strain of the green sulfur bacterium Prosthecochloris (Ptc.) aestuarii 728, which was isolated from a microbial mat from Hot Lake, WA, USA, was recently shown to grow syntrophically via direct electron transfer from Geobacter sulfurreducens.
Concurrently, another strain of Prosthecochloris sp., designated Prosthecochloris sp. HL-
130-GSB, was isolated from the same microbial mat. Prosthecochloris sp. HL-130-GSB has had its physiology characterized and its genome sequenced. In order to determine if
Prosthecochloris sp. HL-130-GSB was capable of syntrophic growth with G. sulfurreducens, this study tested co-cultivation of these two strains under the same growth conditions used in the previous syntrophy study. The data presented here show that Prosthecochloris sp. HL-130-GSB is also capable of syntrophic growth with G. sulfurreducens.
240 A.2 Introduction
Green sulfur bacteria (GSB) are obligately anaerobic photolithoautotrophs that fix
CO2 via the reverse TCA cycle (Wahlund and Madigan, 1993; Wahlund and Tabita,
2+ 1997). GSB have been shown to use reduced sulfur sources, H2, and Fe as electron
donors for photosynthesis (Frigarrd and Dahl, 2009; Heising et al., 1999). A recent study
showed that one GSB isolate, designated Prosthecochloris (Ptc.) aestuarii strain 728,
could also grow using electrons supplied via an electrode or by direct electron transfer
from Geobacter sulfurreducens (Ha et al. 2017). Ptc. aestuarii 728 was isolated from a
phototrophic microbial mat from Hot Lake, a hypersaline, epsomitic stratified lake near
Oroville, Washington (Lindemann et al., 2013). Rhodopsuedomonas palustris TE-1, a
purple non-sulfur bacterium, is the only other phototroph that has been shown to grow
using electrons from an electrode (Bose et al. 2014). Ptc. aestuarii 728 is the only phototroph that has been shown to grow using direct electron transfer from another organism as its source of electrons. The authors termed this new mode of growth, syntrophic anaerobic photosynthesis. Simultaneously with the isolation of Ptc. aestuarii
728, another Prosthecochloris sp. strain from the Hot Lake microbial mat was isolated.
This strain, designated Prosthecochloris sp. HL-130-GSB, has been characterized with
respect to its growth physiology and pigment composition, and additionally its genome
has been sequenced (Genebank accession: CP020873; Vera Theil, personal
communication). Since Ptc. aestuarii 728 and Prosthecochloris sp. HL-130-GSB were
isolated from the same microbial mat but under different conditions, it was unclear
whether these related strains would both be capable of syntrophic anaerobic
photosynthesis. In this study Prosthecochloris sp. HL-130-GSB and Geobacter 241 sulfurreducens PCA were co-cultured following the procedures used by Ha et al. (2017) in order to determine if Prosthecochloris sp. HL-130-GSB is capable of syntrophic anaerobic photosynthesis.
A.3 Experimental Procedures
A.3.1 Strains Used in This Work
Prosthecochloris sp. HL-130-GSB was isolated from a microbial mat in Hot Lake in WA, USA. The original mat sample was obtained from Stephen Lindemann at PNNL and strain HL-130-GSB was isolated by Vera Thiel and co-workers. Vera Thiel provided this isolate which was maintained in HL-CL media. G. sulfurreducens strain PCA (ATCC
51573) was obtained from the lab of Stephen Lindemann at PNNL and maintained in GB media.
A.3.2 Growth Media
Three different anaerobic growth media were used in this study; they are referred to as HL-CL, GB, and mGB. HL-CL is a modification of the CL (Chlorbi liquid) medium routinely used to grow GSB (Frigaard and Bryant 2001; Wahlund and Madigan, 1995).
To make 1L of HL-CL medium, 200 ml of 5× HL (hot lake) salts was mixed with the components of CL medium and brought to a final volume of 1L before autoclaving. To make 1L of 5× HL salts 493 g of MgSO4 • 7H2O, 57 g Na2SO4 and 7.5 g KCl were added
to a final volume of 1L and autoclaved. GB (Geobacter) medium was made according to
the recipe of Srikanth et al., 2008 with the addition of the vitamin mix describe in Table 1
of Balch et al., 1979, and supplemented with 20 mM sodium acetate and 40 mM sodium 242 fumartate. The basal mGB media (modified GB) was made according to the recipe found in Ha et al. 2017.
A.3.3 Growth Studies
Stock cultures of Prosthecochloris sp. HL-130-GSB were maintained in HL-CL media. Stock cultures of G. sulferreducens PCA were maintained in GB media with acetate and fumarate. All manipulations and transfers of these cultures were performed in a Coy anoxic chamber maintained with an atmosphere of 10% H2, 10% CO2, 80% nitrogen. Each culture was maintained in crimp top bottles with rubber stoppers and subcultures were transferred to crimp top tubes with thick butyl rubber stoppers. Cultures were grown in an incubator maintained at 29°C with illumination from a halogen light bulb. For syntrophic growth experiments, starter cultures were transferred in mGB with appropriate supplements several times in order to obtain sufficient starting material.
Starter cultures of Prosthecochloris sp. HL-130-GSB were grown in mGB supplemented with 3.5 mM Na2S•9H2O. Starter cultures of G. sulfurreducens were grown in mGB supplemented with 20mM sodium fumarate and 10mM sodium acetate. For co-culture experiments cells from the appropriate starter culture were pelleted and washed with mGB medium without any additives. The appropriate cells were then inoculated into mGB with 10mM acetate and 0.5mM thiosulfate and growth was monitored via optical density at 600 nm and 650 nm in sealed culture tubes using a Genesys 50 UV-Vis spectrophotomer (Thermo Scientific, Waltham, MA).
A.4 Results 243 In order to see if Prostecochloris sp. HL-130-GSB is capable of syntrophic
anaerobic photosynthesis co-culture growth studies were conducted with G.
sulfurreducens. In these experiments axenic cultures of G. sulfurreducens, and
Prosthecochloris sp. HL-130-GSB were compared to co-cultures containing both strains.
These cultures were grown in mGB with 10 mM acetate as the only electron source present and a small amount of thiosulfate as the sulfur source (mGB AT). Growth studies showed that while neither axenic strain can grow in this medium, the co-cultures can grow (Figure A.1). In a follow-up study the growth of G. sulfurreducens in mGB with acetate and fumarate (mGB AF) and Prosthecochloris sp. HL-130-GSB in mGB with sulfide (mGB S) were compared to the mGB AT single and co-cultures. This experiment showed that co-cultures in mGB AT grew similarly to axenic cultures with appropriate electron donors and acceptors present while axenic cultures in mGB AT again showed no growth (Figure A.2). Together these data show that Prostecochloris sp. HL-130-GSB can grow syntrophically with G. sulfurreducens with acetate as the sole electron source.
A.5 Discussion
The data presented here show that Prosthecochloris sp. HL-130-GSB is capable of growth in co-culture with G. sulfurreducens when acetate is provided as the sole electron source. Previous work showed that Ptc. aestuarii 728 syntrophic growth under these conditions was light-dependent and resulted in the concomitant usage of acetate (Ha et al., 2017). They proposed a new mode of syntrophic growth designated syntrophic anaerobic photosynthesis which is based on direct electron sharing between G. sulfurreducens and Ptc. aestuarii 728. Based on the results presented here, 244 Prosthecochloris sp. HL-130-GSB also appears to be capable of syntrophic anaerobic
photosynthesis. The complete genome sequence of HL-130-GSB is available in NCBI
Genebank database accession: CP020873. The availability of this genome will allow
future research on the genetic and biochemical basis of syntrophic anaerobic
photosynthesis.
These experiments were able to repeat the co-culture growth seen by Ha et al.
(2017); however, some caveats to working with this strain under these conditions should
be noted. Firstly, Prosthecochloris sp. HL-130-GSB did not grow to high ODs in mGB
medium even when supplemented with sulfide (mGB S). This is likely at least partially
due to the fact that the salt composition of mGB S medium is quite different than that of
the HL-CL medium in which it was routinely grown. Notably, mGB has 100× less
MgSO4 and lacks Na2SO4 (Table A.1, Table A.2). Growth in mGB S was somewhat
improved by successive sub-culturing of the strain in mGB S media. In fact, initial
attempts at syntrophic co-cultures transferred from starters grown in HL-CL medium failed repeatedly. It is therefore recommended that if this strain is to be used for further study of syntrophic growth with Geobacter sp. that the strain first be adapted to the medium for co-culture via successive transfers. Secondly, the Ptc. aestuarrii 728 strain may have been subjected to additional selection pressure during experiments using an electrode as the electron source. Selecting Prosthecochloris sp. HL-130-GSB via growth in a bioelectrochemical system using an electrode as the electron source may enhance its ability to grow in co-culture with Geobacter sp. Additionally, the availability of sequenced genomes for both G. sulfurreducens PCA and Prosthecochloris sp. HL-130- 245 GSB may allow for further optimization of the syntrophic growth media to allow
increased growth yields in future experiments. It would also be useful to test if the type
strain of Ptc. aestuarii is capable of syntrophic anaerobic photosynthesis since this strain does not come from an environment rich in MgSO4 and therefore may grow more easily
under low sulfate salt conditions (Gorlenko, 1970). Thirdly, it is important to note that
HL-CL and mGB S media are reduced by the addition of Na2S which scavenges any
oxygen present as a result of incidental oxygen exposure. In contrast, mGB AT does not
contain any reductants other than a small amount of thiosulfate. As a result, the co-culture
experiments are very sensitive to incidental oxygen exposure during handling, which is
possibly responsible for variable growth rates seen in the replicates presented here.
Finally, growth of the co-cultures, and to a lesser extent the axenic cultures, was vastly
improved by allowing more time between measurements. This is likely due to the cells
being allowed to settle at the bottom of the tube and form close contacts required for
direct electron transfer between cells. Settling may also provide protection from small
amounts of oxygen exposure.
Ha et al. 2017 also showed that syntrophic growth was dependent on the presence
of ombB-omaB-omcB gene cluster in Geobacter sulfurreducens. These genes are part of a
cluster encoding a porin-cytochrome protein complex which is required in G.
sulfurreducens for growth using external electron transfer (Liu et al., 2014). Currently, no
information is available on what mechanisms might allow Ptc. aestuarii 728 and
Prosthecochloris sp. HL-130-GSB to accept electrons from an electrode or Geobacter sp.
Development of a genetic system in Prosthecochloris sp. HL-130-GSB or demonstration 246 of syntrophic anaerobic photosynthesis in a GSB with an established genetic system will
allow for further investigation of these mechanisms. Interestingly, in Rps. palustris TIE-1
an operon required for growth on Fe2+, pioABC, was also shown to affect the efficiency of electron uptake from an electrode (Bose et al., 2014; Jiao and Newman, 2007). By analogy future studies focusing on known electron transport mechanisms in GSB may help to shed light on the mechanisms that allow syntrophic anaerobic photosynthesis in
GSB.
247 References
Balch, W.E., Fox, G.E., Magrum, L.J., Woese, C.R., Wolfe, R.S. (1979). Methanogens: Reevaluation of a unique biological group. Microbiol. Rev. 4, 260-296. Bose, A., Gardel, E.J., Vidoudez, C., Parra, E.A., Girguis, P.R. (2014) Electron uptake by iron-oxidizing phototrophic bacteria. Nature Com. 5, 3391. doi:10.1038/ncomms4391 Frigaard, N. U., and Bryant, D. A. (2001). Chromosomal gene inactivation in the green sulfur bacterium Chlorobium tepidum by natural transformation. Appl Env. Microbiol 67, 2538– 2544. doi:10.1128/AEM.67.6.2538. Frigaard, N., and Dahl, C. (2009). “Sulfur Metabolism in Phototrophic Sulfur Bacteria.” In Advances in Microbial Physiology Vol 54, ed. R. K. Poole, (Academic Press) pp. 103- 200 doi:10.1016/S0065-2911(08)00002-7. Gorlenko, V.M. (1970). A new phototrophic green sulphur bacterium- Prosthecochloris aestuarii nov. gen. nov. spec. Z. Allg. Mikrobiol. 10, 147-149. doi: 10.1002/jobm.19700100207 Ha, P.T., Lindemann, R.S., Shi, L., Dohnalkova A.C., Fredrickson, J.K., Madigan, M.T., and Beynal, H. (2017). Syntrophic anaerobic photosynthesis via direct interspecies electron transfer. Nature Com. 8, 13924. doi:10.1038/ncomms13924. Heising, S., Richter, L., Ludwig, W., and Schink, B. (1999). Chlorobium ferrooxidans sp. nov., a phototrophic green sulfur bacterium that oxidizes ferrous iron in coculture with a “Geospirillum” sp. strain. Arch. Microbiol. 172, 116–124. doi:10.1007/s002030050748. Jiao, Y. and Newman, D.K. (2007). The pio operon is essential for phototrophic Fe(II) oxidation in Rhodopseudomonas palustris TIE-1. J. Bacteriol. 189, 1765–1773. doi: 10.1128/JB.00776-06 Lindemann, S.R., Moran, J.J., Stegen, J.C., Renslow, R.S., Hutchison, J.R., Cole, J.K., Dohnalkova, A.C. et al. (2013). The epsomitic phototrophic microbial mat of Hot Lake, Washington: community structural responses to seasonal cycling. Front. Microbiol.4, 323. doi:10.3389/fmicb.2013.00323. Liu, Y., Wang Z., Liu J., Levar C., Edwards M.J., Babauta J.T., Kennedy D.W., Shi Z., Beyenal H., et al. (2014). A trans-outer membrane porin-cytochrome protein complex for extracellular electron transfer by Geobacter sulfurreducens PCA. Environ. Microbial. Rep. 6, 776-785. doi: 10.1111/1758-2229.12204. Srikanth, S., Marsili, E., Flickinger M.C., Bond, D.R. (2008). Electrochemical characterization of Geobacter sulfurreducens cells immobilized on graphite paper electrodes. Biotech. Bioeng. 99, 1065-1073. doi: 10.1002/bit.21671 Wahlund, T. M., and Madigan, M. T. (1993). Nitrogen fixation by the thermophilic green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 175, 474–478. doi:10.1128/jb.175.2.474-478.1993. Wahlund, T. M., and Madigan, M. T. (1995). Genetic transfer by conjugation in the thermophilic green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 177, 2583–2588. Wahlund, T. M., and Tabita, F. R. (1997). The reductive tricarboxylic acid cycle of carbon dioxide assimilation: Initial studies and purification of ATP-citrate lyase from the green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 179, 4859–4867. doi:10.1128/jb.179.15.4859-4867.1997.
248
Figure A.1 Co-culture Growth Experiment
Co-cultures and single cultures grown in mGB media with acetate and thiosulfate added. Each condition was run in triplicate tubes. Co-culture growth curves are represented by the solid bright green line. Prosthecochloris sp. single cultures are represented by a dotted light green line and G. sulfurreducens PCA cultures are represented by a dotted orange line.
249
Figure A.2 Growth Experiment with Varied Additives
Co-cultures and axenic cultures grown in mGB media with various additives. Co-culture growth curves are represented by the solid bright green line. Prosthecochloris sp. axenic cultures are represented by a light green line and G. sulfurreducens axenic cultures are represented by an orange line. The solid and dotted lines represent cultures grown in mGB medium supplemented with acetate and thiosulfate. The dashed light green line represents the Prosthecochloris sp. grown in mGB supplemented with sodium sulfide. The dashed orange line represents G. sulfurreducens grown in mGB supplemented with acetate and fumarate.
250 Table A.1: Comparison of Basic Media Components of HL-Cl and mGB.
Component HL-CL mM mGB mM HL-CL:mGB
NaCl 6.84 150.24 1:22
CaCl2 •2H2O 0.34 0.48 1:1.4
MgSO4 •7H2O 400.86 4.06 99:1
Na2EDTA •2H2O 0.048 0 NA
Na2SO4 80.26 0 NA
2- Total SO4 ions 481.12 4.06 119:1
KCl 20.12 50.97 1:2.5
NH4Cl 7.48 4.67 1.6:1
– NH4CH3OO 6.49 0 NA
– NaCH3OO * 0 0; 10 NA; NA
+ Total NH4 ions 13.97 4.67 3:1
– Total CH3OO ions 6.49 0; 10 NA; 1:1.5
Na2S2O3 •5H2O* 9.27 0; 0.5 NA; 19:1
KH2PO4 3.67 3.67 1:1
MOPS 10.04 2.39 4.2:1 251
NaHCO3 23.8 29.76 1:1.3
Na2S •9H2O* 2.5 0; 3.5 NA; 1:1.4
Na-fumarate* 0 0; 20 NA; NA
* indicates that in the mGB medium this component is only added when indicated in the text
252 Table A.2: Comparison of Vitamin and Trace Element Components HL-CL and mGB mGB medium additionally contained 0.02 mg/L of biotin and folic acid, 0.1 mg/L pyridoyine HCl, and 0.05 mg/L riboflavin, thiamin HCl, nicitinic acid, calcium pantothenate, amino benxoic acid and lipoic acid.
Component HL-CL (mM)* mGB (mM)* HL-CL:mGB
Vitamin B12 0.02 (mg/L) 0.001 (mg/L) 20:1
Nitrilotriacetic acid 0 0.079 NA
MnCl2 •4H2O 0.0005 0.005 1:10
FeSO4 •7H2O 0 0.011 NA
FeCl2 •4H2O 0.0075 0 NA
Total Fe2+ ions 0.0075 0.011 1:1.5
CoCl2 •6H2O 0.0008 0.0071 1:8.9
ZnCl2 0.0005 0.007 1:14
CuSO4 •5H2O 0 0.002 NA
CuCl2 •2H2O 0.0001 0 NA
Total Cu2+ ions 0.0001 0.002 1:20
AlK(SO4)2 •12H2O 0 0.00011 NA
H3BO3 0.0001 0.0008 1:8
Na2MoO4 •2H2O 0.0008 0.004 1:5 253
NiCl2 •6H2O 0.0001 0.009 1:90
NaWO4 •2H2O 0.000006 0.0006 1:1000
Na2SeO4 0 0.005 NA
NaHSeO3 0.00001 0 NA
2- Total SeOx ions 0.00001 0.005 1:500
VOSO4 2H2O 0.00015 0 NA
*concentrations in this table are in mM unless otherwise indicated
254 Appendix B.Genome Sequence of Thiohalocapsa MS
B.1 Introduction
Thiohalocapsa MS is a purple sulfur bacterium (PSB) which was recently isolated from Mushroom Spring, Yellowstone National Park, WY (Dr. Marcus Tank, personal communication). Thiohalocapsa MS is one of only a few PSB that have been shown to
grow above 45 °C. It is currently the only Thiohalocapsa sp. that grows at elevated
temperatures. This work describes the sequencing and analysis of the genome of
Thiohalocapsa MS, which was carried out in order to learn more about the biosynthetic
capabilities of this new organism.
B.2 Experimental Procedures
B.2.1 Strains Used in This Study
Thiohalocapsa MS was isolated from Mushroom Spring, Yellowstone National
Park, WY, USA, by Dr. Marcus Tank. Thiohalocapsa MS was grown at 45 °C in
Pfennig’s medium under illumination from incandescent bulbs (Eichler and Pfennig,
1988; Imhoff, 2015). Sulfide levels were monitored by using lead acetate paper (GE
Healthcare Whatman/Fischer Scientific, Göteborg, Sweden), and cultures were fed with a
filter-sterilized sulfide solution. The sulfide feeding solution was made by dissolving 7 g
of Na2S • 9H2O and 2.65 g of Na2CO3 in 250 mL H2O and by adjusting the pH to 7.3 by
addition of H2SO4; the solution was stored in crimp-top bottles with thick butyl rubber
stoppers (Imhoff, 2015). 255 B.2.2 Genomic DNA (gDNA) Preparation
Cells were harvested during stationary growth phase and gDNA was prepared using a
Qiagen Genomic-tip kit (Qiagen, Germantown, MD). The protocol for preparation of
Gram-negative bacterial gDNA found in the Qiagen genomic DNA Handbook was used with minor modifications (Qiagen, Germantown, MD). All steps were performed with wide bore pipette tips in order to avoid DNA shearing. The lysis and protease incubation at 37 °C was extended up to 1 hour. Additionally, the 50 °C incubation following addition of buffer B2 was extended up to 1 hour. If samples still appeared pink, due to intact protein-rich, light-harvesting complexes, then the incubation was extended further.
Additional wash steps were performed after the sample was added to the Genomic-tip column and during ethanol precipitation. The purified gDNA was resuspended in freshly made DNAse free 10mM Tris-HCl pH 8.5 and stored at 4°C for sequencing.
B.2.3 gDNA Sequenceing
Sequencing was performed at the Penn State Genomics Core Facility (University
Park, PA) using a PacBio Sequel sequencer. Template DNA was sheared to circa 10 Kb, and the sample library was prepared using the SMRTbell Express template from PacBio.
Data was collected for circular consensus sequences (CCS) and processed by the
Genomic Core Facility in conjunction with the lab of Dr. Istvan Albert at Penn State.
B.2.4 Bioinformatic Analyses
The genome was assembled de novo from CCS reads using Unicycler, Galaxy version 0.4.8.0, (Wick et al., 2016) on the Galaxy platform via the usegalaxy.org server
(Afgan et al., 2018). The assembler was run using default settings. The assembly was 256 uploaded to the RAST server and initial annotation was performed using RASTtk (Aziz
et al., 2008; Brettin et al 2015; Overbeek et al., 2014). Manual annotation was performed
for missing genes using BlastP searches of known enzymes from the NCBI database
(Altschul et al., 1997).
B.3 Results and Discussion
B.3.1 Overview of Genome
The genome assembly for Thiohalocapsa MS consists of a single contig of 4,697,377
bp in length with a GC content of 68.8%. The RASTtk annotated genome contains 4167
protein-coding genes, and 48 RNA genes, comprising 45 tRNAs and 3 rRNAs. Genes encoding putative enzymes for the Calvin Benson Bassham cycle, nitrogen fixation and oxidation of sulfide were identified. Additionally, genes encoding pigment biosynthesis enzymes were identified for production of BChl a and carotenoids in the spirilloxanthin series. Genes encoding the proteins which make up LH1, LH2, and the type-2 RC found in purple phototrophic bacteria were also identified. Together these data are consistent with classification of Thiohalocapsa MS as a nitrogen-fixing photoautotroph which can use sulfide and thiosulfate as an electron source.
B.3.2 Central Metabolism
B.3.2.1 Carbon metabolism
The genome of Thiohalocapsa MS encodes enzymes for the complete oxidative TCA cycle, the glycolysis pathway, and the pentose phosphate pathway (Table B.1). Several genes in the glycolysis pathway form a gene cluster comprising fructose-bisphosphate aldolase, pyruvate kinase, phosphoglycerate kinase, and the NAD+-dependent 257
glyceraldehyde-3-phosphate dehydrogenase. The fixation of CO2 in purple bacteria
proceeds via the Calvin-Benson-Bashamm cycle, for which a full set of genes is present
in Thiohalocapsa MS (Table B.2). The genes rbcS/rbcL which encode ribulose-1,5,-
bisphosphate (RuBisCo) subunits appear in a gene cluster with several other genes
annotated as encoding carboxysome proteins. The genome of Alc. vinosum also has carboxysome genes, although carboxysomes have not been observed in this organism
(Weissgerber, 2011). In contrast, the carboxysome of the Gammaproteobacterium
Halothiobacillus neapolitan has been studied for decades (Shively et al., 1973). Further work will be required to determine if carboxysomes are produced in vivo by
Thiohalocapsa MS.
B.3.2.2 Nitrogen and Amino Acid Biosynthesis
Genes encoding all three nitrogenase subunits were identified in the genome assembly from Thiohalocapsa MS. These genes have been annotated as nifD, nifK, and
nifH, which are predicted to encode the subunits of a molybdenum-containing
nitrogenase capable of converting dinitrogen into ammonia (Table B.3). The genes
required for conversion of ammonia into glutamine, aspartate, and glycine were
identified. From glutamine the genome encodes the capacity to produce glutamate and
subsequently proline. From aspartate the genome encodes the ability to produce
asparagine and lysine. From glycine the genome encodes the ability to produce serine and subsequently tryptophan, and cysteine. Alanine biosynthesis is encoded via the pathway from cysteine using cysteine desaturase and alanine racemase. Valine, leucine and isoleucine biosynthesis from threonine is encoded in the genome. Additionally the 258 genome encodes biosynthesis of histidine, tyrosine, phenylalanine and tryptophan from
precursors in the pentose phosphate pathway. Genes encoding enzymes for arginine
biosynthesis were identified proceeding from aspartate or glutamate via the urea cycle.
The biosynthetic pathways to threonine and methionine are both missing one enzyme, but
the organism grows without the addition of amino acids, so these functions must be encoded in the genome.
B.3.2.3 Sulfur Metabolism
The genome of Thiohalocapsa MS encodes genes for the oxidation of sulfur compounds. Thiohalocapsa MS encodes both a sulfide/quinone oxidoreductase (SQR), encoded by sqrD and sqrF, and a flavocytochrome c sulfide dehydrogenase, encoded by fccA and fccB similar to Allochromatium (Alc.) tepidum (Dahl, 2017) (Table B.4).
Thiohalocapsa MS also has several dsr genes, like Alc. tepidum, which encode Dsr enzymes that oxidize sulfide to sulfite in sulfur-oxidizing organisms (Weissgerber et al.,
2011). The reverse dissimilatory sulfite reductase is encoded by dsrA and dsrB. The dsrAB genes are found in a cluster in the genome with dsrMKJOP which encode a sulfite reductase-associated complex that is present in other purple sulfur bacteria (Dahl et al.,
2005; Grimm et al., 2010; Grein et al., 2010). Further, oxidation of sulfite to sulfate is encoded by the aprA, aprB, aprM, and sat genes or the soeA, soeB, and soeC genes. Like the dsr genes, the apr and sat genes are homologs of genes used in dissimilatory sulfate reduction. In sulfur oxidizers AprAB oxidizes sulfite to adenyl sulfate (APS) and sulfate adenylytransferase, encoded by sat, oxidizes APS to sulfate. In contrast SoeABC oxidizes
sulfite directly to sulfate (Dahl, 2017). Thiosulfate oxidation appears to occur via the Sox 259 system with genes present encoding SoxA, SoxB, SoxX, SoxY, SoxZ, and SoxW (Table
B.4). This is significant because thiosulfate utilization is important for the ability of
sulfide oxidizers to grow on solid media. Growth on solid media allows for easier colony
isolation for mutagenesis and maintaining stocks. Future experiments should be carried
out to test the growth of Thiohalocapsa MS on solid media containing thiosulfate.
B.3.3 Photosynthesis Related Genes
B.3.3.1 Pigment Biosynthesis
All of the genes required for biosynthesis of spirilloxanthin from geranylgeranyl
pyrophosphate were identified in the genome assembly. These include crtB, crtI, crtC,
crtD, and crtF. In the genome crtB and crtD are adjacent to each other as are crtI and
crtC. The crtF gene is located directly adjacent to the geranylgeranyl pyrophosphate
synthetase (Table B.5). The presence of this set of genes is expected based on pigment
analyses performed previously, which identified spirilloxanthin and anhydrorhodovibrin
as the main carotenoids in Thiohalocapsa MS (see Chapter 5).
As expected based on prior pigment analysis, the genome of Thiohalocapsa MS
encodes all of the enzymes required for biosynthesis of BChl a from 5-aminolevulinic acid (5-ALA). 5-ALA is synthesized from glutamate via L-glutamyl-tRNA. The genome encodes both the oxygen-dependent coproporphyrinogen III oxidase, HemF, and the oxygen-independent enzyme, HemN. In contrast, only one isoenzyme of protoporphyrinogen oxidase was identified and was annotated as HemY, the oxygen- dependent enzyme (see Chapter 2). The genes encoding the biosynthetic pathway from protoporphyrin IX to BChl a are found in two main gene clusters in the genome. One 260 cluster contains bchH, bchM, bchNLB, and bchF while the other has bchXYZ and bchC.
Additionally, the bchG and bchP genes are adjacent to one another and the bciA and bchE
genes are not clustered with other photosynthetic genes (Table B.5).
B.3.3.2 Light-Harvesting Apparatus
As in other purple bacteria Thiohalocapsa MS encodes a type-2 reaction center with
H, L, and M subunits in addition to a cytochrome c subunit (Bullough et al., 2009). The
genome also encodes the alpha and beta subunits of LH1 and LH2 light harvesting
complexes found in purple bacteria (Gabrielsen et al., 2009). The genes encoding the
reaction center H subunit and PucC, which is involved in assembly of the light harvesting
apparatus, are directly downstream of the gene cluster containing bchH, bchM, bchNLB,
and bchF (Mothersole et al., 2016). The genes encoding the L and M RC subunits, the cytochrome c subunit and 3 pairs of alpha/beta subunits belonging to LH1 and LH2 are encoded adjacent to bchXYZ and bchC. An additional gene cluster with homologs of four pairs of LH1 or LH2 alpha/beta subunits is found elsewhere in the genome for a total of seven alpha/beta subunit pairs (Table B.6). Thiohalocapsa MS does not appear to encode the PufX subunit which is found in the type-2 RC of some purple bacteria (Bullough et al., 2009). The availability of sequences for the genes encoding the light-harvesting apparatus of Thiohalocapsa MS may provide an avenue to better understand the proteins involved in the LH1 and LH2 complexes of this organism. Of particular interest is the relationship of temperature and Ca2+ with the LH1 complex, and the peptide composition
of the different spectral forms of LH2 (see Chapter 5).
261 B.4 References
Afgan, E., Baker, D., Batut, B., van den Beek, M., Bouvier, D., Čech, M., Chilton, J., Clements, D., Coraor, N. et al., (2018). The Galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2018 update, Nucl. Acids Res. 46, W537–W544. doi:10.1093/nar/gky379. Altschul, S.F., Madden, T. L., Schäffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25,3389-3402. Aziz, R.K., Bartels, D., Best, A.A., DeJongh, M., Disz, T., Edwards, R.A., Formsma, K., Gerdes, S., Glass, E.M., et al., (2008). The RAST Server: Rapid Annotations using Subsystems Technology.BMC Genomics. 8,75. doi: 10.1186/1471-2164-9-75. Brettin T, Davis JJ, Disz T, Edwards RA, Gerdes S, Olsen GJ, Olson R, Overbeek R, Parrello B, Pusch GD, et al., (2015). RASTtk: A modular and extensible implementation of the RAST algorithm for building custom annotation pipelines and annotating batches of genomes. Sci. Rep. 5,8365. doi: 10.1038/srep08365. Bullough, P. A., Qian, P., and Hunter, C. N. (2009). Reaction center-light-harvesting core complexes of purple bacteria, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), pp. 155–179. doi:10.1007/978-1-4020-8815-5_9. Dahl, C. (2017). Sulfur metabolism in phototrophic Bacteria, in Modern Topics in the Phototrophic Prokaryotes: Environmental and Applied Aspects, ed. P. C. Hallenbeck (Cham: Springer International Publishing), pp. 27–66. doi: 10.1007/978-3-319- 51365-2_2. Dahl, C., Engels S., Pott-Sperling A.S., Schulte A., Sander J., Lübbe Y., Deuster O., Brune D.C. (2005) Novel genes of the dsr gene cluster and evidence for close interaction of Dsr proteins during sulfur oxidation in the phototrophic sulfur bacterium Allochromatium vinosum. J. Bacteriol. 187,1392-1404. doi: 10.1128/JB.187.4.1392-1404.2005 Eichler B., and Pfennig N. (1988). A new green sulfur bacterium from a freshwater pond. In: Olson JM, Stackebrandt E, Trüper H (eds.) Green photosynthetic bacteria. Plenum, New York, pp. 233–235 Gabrielsen, M., Gardiner, A. T., and Cogdell, R. J. (2009). Peripheral complexes of purple bacteria, in The Purple Phototrophic Bacteria, eds. C. N. Hunter, F. Daldal, M. C. Thurnauer, and J. T. Beatty (Dordrecht: Springer Netherlands), pp. 135–153. doi:10.1007/978-1-4020-8815-5_8. Grein, F., Pereira, I.A., Dahl, C. (2010). Biochemical characterization of individual components of the Allochromatium vinosum DsrMKJOP transmembrane complex aids understanding of complex function in vivo. J. Bacteriol. 192, 6369-6377. doi: 262 10.1128/JB.00849-10. Grimm F., Cort J.R., Dahl C. (2010). DsrR, a novel IscA-like protein lacking iron- and Fe-S-binding functions, involved in the regulation of sulfur oxidation in Allochromatium vinosum. J. Bacteriol. 192,1652-1661. doi: 10.1128/JB.01269-09. Imhoff, J. F. (2015). “Chromatiaceae,” in Bergey’s Manual of Systematics of Archaea and Bacteria (American Cancer Society), 1–12. doi:10.1002/9781118960608.fbm00219. Mothersole, D. J., Jackson, P.J., Vasilev, C., Tucker, J.D., Brindley, A.A., Dickman, M. J., and Hunter, C. N. (2016) PucC and LhaA direct efficient assembly of the light‐ harvesting complexes in Rhodobacter sphaeroides. Mol. Microbiol. 99, 307–327. doi: 10.1111/mmi.13235 Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA, Gerdes S, Parrello B, et al., (2014). The SEED and the Rapid Annotation of microbial genomes using Subsystems Technology (RAST). Nucleic Acids Res. 42, D206-214. doi:10.1093/nar/gkt1226. Shively, J.M., Ball F.L., Kline B.W. (1973) Electron Microscopy of the Carboxysomes (Polyhedral Bodies) of Thiobacillus neapolitanus. J. of Bacteriol. 116, 1405–1411. Weissgerber T, Zigann R, Bruce D, Chang YJ, Detter JC, Han C, Hauser L, Jeffries CD, Land M, Munk AC, Tapia R, Dahl C. (2011). Complete genome sequence of Allochromatium vinosum DSM 180(T). Stand. Genomic Sci. 31, 311-330. doi: 10.4056/sigs.2335270. Wick, R. R., Judd, L. M., Gorrie, C. L., and Holt, K. E. (2017). Unicycler: resolving bacterial genome assemblies from short and long sequencing reads. PLoS Comp. Bio. 13, e1005595. doi:10.1371/journal.pcbi.1005595
263 Table B.1 Carbon Metabolism Genes *indicates RAST abbreviation with multiple enzyme types Enzyme RAST Abbreviation CDS Number(s)
Glucose-6-phosphate 1-dehydrogenase GPDH 3203 (EC 1.1.1.49)
6-phosphogluconolactonase (EC *PGL 3615 3.1.1.31)
6-phosphogluconate dehydrogenase, PglDH 3785 decarboxylating (EC 1.1.1.44)
Ribose 5-phosphate isomerase A (EC *Ris 712 5.3.1.6)
Ribulose-phosphate 3-epimerase (EC Repi 3337 5.1.3.1)
Transketolase (EC 2.2.1.1) *TK 1528;2531;2589;549;977
Transaldolase (EC 2.2.1.2) TA 963
Fructose-6-phosphate phosphoketolase FPK 1964; 932 (EC 4.1.2.22)
Xylulose-5-phosphate phosphoketolase XPK 1964; 932 (EC 4.1.2.9)
Ribose-phosphate pyrophosphokinase PRPPS 2633 (EC 2.7.6.1)
Citrate synthase (si) (EC 2.3.3.1) gltA 2707; 3510
Aconitate hydratase (EC 4.2.1.3) *AcoH 711
2-oxoglutarate dehydrogenase E1 sucA 1915 component (EC 1.2.4.2)
2-oxoglutarate dehydrogenase E2 *sucB 3477+ component (EC 2.3.1.61)
Succinyl-CoA ligase [ADP-forming] *sucD 2642 alpha chain (EC 6.2.1.5)
Succinyl-CoA ligase [ADP-forming] *sucS 2641 beta chain (EC 6.2.1.5)
Succinate dehydrogenase iron-sulfur sdhB 3856 protein (EC 1.3.99.1) 264 Succinate dehydrogenase flavoprotein sdhA 3857 subunit (EC 1.3.99.1)
Fumarate hydratase class I, aerobic (EC *fum 924 4.2.1.2)
Malate dehydrogenase (EC 1.1.1.37) *MD 1799
Isocitrate dehydrogenase [NADP] (EC *icd 3313 1.1.1.42)
Dihydrolipoamide dehydrogenase of lpdAp 2475 pyruvate dehydrogenase complex (EC 1.8.1.4)
Glucokinase (EC 2.7.1.2) *glk 605
Glucose-6-phosphate isomerase (EC *pgi 3083 5.3.1.9)
6-phosphofructokinase (EC 2.7.1.11) *pfk 3491
Fructose-1,6-bisphosphatase, type I (EC *fbp 1809 3.1.3.11)
Fructose-bisphosphate aldolase class II *fba 973 (EC 4.1.2.13)
Triosephosphate isomerase (EC 5.3.1.1) TpI 1854
NAD-dependent glyceraldehyde-3- *gap 316;976 phosphate dehydrogenase (EC 1.2.1.12)
Phosphoglycerate kinase (EC 2.7.2.3) PgK 975
Phosphoglycerate mutase (EC 5.4.2.1) *pgm 404
Enolase (EC 4.2.1.11) EnO 515;558
Pyruvate kinase (EC 2.7.1.40) PyK 974
Phosphoenolpyruvate synthase (EC *pps 2940;879 2.7.9.2)
265 Table B.2 Calvin Benson Basham Cycle Genes
*indicates RAST abbreviation with multiple enzyme types
Enzyme RAST Abbreviation CDS Number(s)
Phosphoribulokinase (EC 2.7.1.19) PRK 3519
Ribulose bisphosphate carboxylase *RuBisCo 303;304 large chain (EC 4.1.1.39); Ribulose bisphosphate carboxylase small chain (EC 4.1.1.39)
Phosphoglycerate kinase (EC PGK 975 2.7.2.3)
NAD-dependent glyceraldehyde-3- *GAPDH 316;976 phosphate dehydrogenase (EC 1.2.1.12)
Triosephosphate isomerase (EC TPI 1854 5.3.1.1)
Fructose-bisphosphate aldolase class *FBA 973 II (EC 4.1.2.13)
Fructose-1,6-bisphosphatase, type I *FBP 1809 (EC 3.1.3.11)
Transketolase (EC 2.2.1.1) *TK 1528;2531;2589;549;977
Ribulose-phosphate 3-epimerase RPE 3337 (EC 5.1.3.1)
Ribose 5-phosphate isomerase A *Ris 712 (EC 5.3.1.6)
266 Table B.3 Nitrogen Fixation Genes
Enzyme RAST Abbreviation CDS Number(s)
Nitrogenase (molybdenum-iron)-specific NifA 2163 transcriptional regulator NifA
Cysteine desulfurase (EC 2.8.1.7), NifS NifS 3051 subfamily
Iron-sulfur cluster assembly scaffold protein NifU 3050 NifU
Nitrogenase FeMo-cofactor synthesis FeS NifB 1061 core scaffold and assembly protein NifB
4Fe-4S ferredoxin, nitrogenase-associated frdN 1062;339;761
Nitrogenase FeMo-cofactor carrier protein NifX 343 NifX
NifX-associated protein NifX2 341
Nitrogenase cofactor carrier protein NafY NafY 758
Nitrogenase FeMo-cofactor scaffold and NifE 345 assembly protein NifE
Nitrogenase FeMo-cofactor scaffold and NifN 344 assembly protein NifN
Nitrogenase FeMo-cofactor synthesis NifQ 1067 molybdenum delivery protein NifQ
Homocitrate synthase (EC 2.3.3.14) NifV 328
Nitrogenase stabilizing/protective protein NifW 330 NifW
NifM protein NifM 2200
Nitrogenase (molybdenum-iron) reductase NifH 765 and maturation protein NifH
Nitrogenase (molybdenum-iron) alpha chain NifD 764 (EC 1.18.6.1) 267 Nitrogenase (molybdenum-iron) beta chain NifK 763 (EC 1.18.6.1)
NifZ protein NifZ 331
NifT protein NifT 762
Nitrogenase-associated protein NifO NifO 1063
268 Table B.4 Sulfur Metabolism Genes
*indicates RAST abbreviation with multiple enzyme types, **indicates that a gene was putatively identified via BLAST, square brackets indicate original RAST annotation when BLAST was used for identification
Enzyme RAST Abbreviation CDS Number(s)
Dissimilatory sulfite reductase, alpha DsrA 3187 subunit (EC 1.8.99.3)
Dissimilatory sulfite reductase, beta DsrB 3188 subunit (EC 1.8.99.3)
Dissimilatory sulfite reductase, DsrC 2438;2725;3617 gamma subunit (EC 1.8.99.3)
Sulfite reduction-associated complex DsrM 3193 DsrMKJOP protein DsrM (= HmeC)
Sulfite reduction-associated complex DsrK 2022, DsrMKJOP protein DsrK (=HmeD)
Protein similar to glutamate synthase DsrL 3195 [NADPH] small chain, clustered with sulfite reductase
Sulfite reduction-associated complex DsrJ 3196 DsrMKJOP multiheme protein DsrJ (=HmeF)
Sulfite reduction-associated complex DsrO 3197 DsrMKJOP iron-sulfur protein DsrO (=HmeA)
Sulfite reduction-associated complex DsrP 3198 DsrMKJOP protein DsrP (= HmeB)
IscA-like protein, DsrR DsrR 3200 sulfur oxidation protein SoxA SoxA 3768
Sulfur oxidation protein SoxB SoxB 1089
SoxK** [hypothetical protein] NA 3769
SoxL** [hypothetical protein] NA 3770 269 Sulfur oxidation protein SoxX SoxX 3767
Sulfur oxidation protein SoxY SoxY 3033
Sulfur oxidation protein SoxZ SoxZ 3032 thioredoxin SoxW SoxW 231;2522;3365;3545
SqrD** [sulfide quinone NA 1254 oxidoreductase]
SqrF** [FAD-dependent pyridine NA 3240 nucleotide-disulphide oxidoreductase]
FccA** [Cytochrome c553] NA 3397
FccB**[Sulfide dehydrogenase [SoxF] 3396 (flavocytochrome C) flavoprotein chain precursor (EC 1.8.2.-)]
Sat** [Sulfate adenylyltransferase NA 826 (EC 2.7.7.4)]
Adenylylsulfate reductase alpha- AprA 823 subunit (EC 1.8.99.2)
Adenylylsulfate reductase beta- AprB 824 subunit (EC 1.8.99.2)
AprM** [Adenylylsulfate reductase [R1] 825 membrane anchor]
SoeA** [Anaerobic dimethyl NA 891 sulfoxide reductase chain A (EC 1.8.5.3)]
SoeB** [Anaerobic dimethyl NA 893 sulfoxide reductase chain B (EC 1.8.5.3]
SoeC** [Anaerobic dimethyl NA 894 sulfoxide reductase chain C (EC 1.8.5.3)]
270 Table B.5 Pigment Biosynthesis Genes
*indicates RAST abbreviation with multiple enzyme types, **indicates that a gene was putatively identified via BLAST
Enzyme RAST Abbreviation CDS Number(s)
Protoporphyrin IX Mg-chelatase subunits *PMgC 1112;2464;2465 HDI (EC 6.6.1.1)
Mg-protoporphyrin O-methyltransferase PMgMT 1114 (EC 2.1.1.11)
Mg-protoporphyrin IX monomethyl ester ChlEAn 59 oxidative cyclase (anaerobic) (EC 1.14.13.81)
Divinyl protochlorophyllide a 8-vinyl- *DVR 665** reductase (EC 1.3.1.75)
Protein BchJ, involved in reduction of C-8 *DVR 3393 vinyl of divinyl protochlorophyllide
Light-independent protochlorophyllide *pChl_R 1110;1111;1113 reductase ChlLNB (EC 1.18.-.-)
Chlorophyll a synthase ChlG (EC ChlG 1791 2.5.1.62)
Geranylgeranyl hydrogenase BchP BchP 1792
Chlorophyllide reductase subunit *Chld_R 3098;3099;3100 BchXYZ (EC 1.18.-.-)
2-vinyl bacteriochlorophyllide hydratase BchF 1109 BchF (EC 4.2.1.-)
2-desacetyl-2-hydroxyethyl BchC 3101 bacteriochlorophyllide a dehydrogenase BchC
Phytoene synthase CrtB (EC 2.5.1.32) *P/DSs 1298;2474
Phytoene dehydrogenase (EC 1.14.99.-) *PD 1299;2287
Geranylgeranyl diphosphate synthase (EC GPS 953 2.5.1.29) 271 CrtT-methyltransferase-like protein TM 3347;3356
Regulator of carotenoid biosynthesis RCB 1108
Methoxyneurosporene dehydrogenase (EC MND 3222 1.14.99.-)[CrtI**]
Hydroxyneurosporene dehydrogenase (EC HNSD 3223 1.-.-.-)[CrtC**]
(2E,6E)-farnesyl diphosphate synthase FPS 2105;953 (EC 2.5.1.10)
[CrtF**] NA 3452
272 Table B.6 Light Harvesting Apparatus Genes
**indicates that a gene was putatively identified via BLAST
Enzyme RAST Abbreviation CDS Number(s)
Light-harvesting LHI, alpha subunit LHIA 3089; 3096
Light-harvesting LHI, beta subunit LHIB 3092
Light-harvesting LHII, beta subunit LHIIB 3090; 3097 B light-harvesting alpha subunits ** NA 3061; 3063; 3065; 3067; 3091; light-harvesting beta subunits ** NA 3062; 3064; 3066; 3068
Photosynthetic reaction center L pufL 3095 subunit
Photosynthetic reaction center M pufM 3094 subunit
Photosynthetic reaction center puf2C 3093 cytochrome c subunit
Photosynthetic reaction center H puhA 1116 subunit
Putative photosynthetic complex Hyp1 1117 assembly protein
Vita
Jennifer L. Thweatt
EDUCATION Ph.D. Biochemistry, Microbiology, and Molecular Biology with Dual-Title Ph.D in Astrobiology December 2019 Thesis: Characterization of Pigment Biosynthesis and Light-Harvesting Complexes of Selected Anoxygenic Phototrophic Bacteria. Advisor: Dr. Donald A. Bryant B.Sc. Microbiology; University of California, Santa Barbara June 2008
RESEARCH EXPERIENCE Graduate Research Assistant January 2012-Present The Pennsylvania State University, Laboratory of Dr. Donald A. Bryant Staff Research Associate Children’s Hospital Oakland Research Institute Laboratory of Dr. Peter Beernink December 2010-July 2011 Laboratory of Dr. Stuart Smith March 2009-August 2010
AWARDS AbSciCon 2017 Student Travel Grant April 2017 Astrobiology Science Conference 2017 NASA Space Grant Graduate Fellowship Fall 2016-Spring 2017 Pennsylvania Space Grant Consortium
PUBLICATIONS JL Thweatt, DP Canniffe, DA Bryant (2019) Bacteriochlorophyll Biosynthesis in Green Bacteria, Metabolism, Structure and Function of Chlorophylls In: Advances in Botanical Research Vol. 90 Metabolism, Structure and Function of Plant Tetrapyrroles Introduction, Microbial and Eukaryotic Chlorophyll Synthesis and Catabolism. Bernhard Grimm, ed. (Academic Press, London, UK), pp. 35–89. DP Canniffe, JL Thweatt, AGM Chew, C N Hunter, and D A Bryant (2018) A paralog of a bacteriochlorophyll biosynthesis enzyme catalyzes the formation of 1,2-dihydrocarotenoids in green sulfur bacteria. The Journal of Biological Chemistry 293, 15233–15242. JL Thweatt, BH Ferlez, JH Golbeck, DA. Bryant (2017) BciD is a Radical-SAM Enzyme That Completes Bacteriochlorophyllide e Biosynthesis by Oxidizing a Methyl Group into a Formyl Group at C-7, The Journal of Biological Chemistry 292:1361-1373. Y Tsukatani, T Mizoguchi, J Thweatt, M Tank, DA Bryant, H Tamiaki (2016) Glycolipid Analyses of Light-harvesting Chlorosomes from Envelope Protein Mutants of Chlorobaculum tepidum, Photosynthesis Research 128:235-241.