ENVIRONMENTAL DETERMINATION AND DYNAMIC REGULATION OF THE CARBON CONCENTRATING MECHANISM IN FREMYELLA DIPLOSIPHON By Brandon Rohnke A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Biochemistry and Molecular Biology – Doctor of Philosophy 2019 PUBLIC ABSTRACT ENVIRONMENTAL DETERMINATION AND DYNAMIC REGULATION OF THE CARBON CONCENTRATING MECHANISM IN FREMYELLA DIPLOSIPHON By Brandon Rohnke Cyanobacteria are single-celled organisms that perform oxygenic photosynthesis. They are major contributors to Earth’s carbon cycle and are a rich source of biological energy. Since they inhabit many diverse habitats, they must regulate photosynthesis based on resource availability; important resources for photosynthesis include light and carbon dioxide (CO2) levels. One feature that cyanobacteria use to maximize photosynthetic efficiency involves the concentration of carbon dioxide around the carbon-fixing rubisco enzyme by trapping it in sub-cellular compartments, which allows the organisms to grow even at low CO2 levels. Investigations in this study explore how a specific cyanobacterium, Fremyella diplosiphon, controls the carbon concentrating mechanism (CCM) in response to the color and intensity of light and in response to CO2 levels. I present evidence that the ability of F. diplosiphon to detect light color and light intensity is involved in the regulation of the CCM. Furthermore, I contribute to the study of an underexplored protein that interacts with rubisco and with a possible role in responding to environmental conditions. I then analyze how the response of F. diplosiphon to resource availability impacts its ability to fix carbon. These investigations provide a picture of the many factors that contribute to the regulation of photosynthesis in cyanobacteria, with insight into the multiple components that could be leveraged for improved biofuel and biotechnology production. ABSTRACT ENVIRONMENTAL DETERMINATION AND DYNAMIC REGULATION OF THE CARBON CONCENTRATING MECHANISM IN FREMYELLA DIPLOSIPHON By Brandon Rohnke Cyanobacteria are single-celled photoautotrophic organisms that are major contributors to global carbon fixation. Since the accumulation of significant amounts of oxygen (O2) in the atmosphere, they have contended with decreased amounts of available inorganic carbon (Ci) with which to supply photosynthesis. This has led to the evolution of a carbon concentrating mechanism (CCM) that involves increasing the Ci uptake into the cell and utilizing a specialized bacterial microcompartment (BMC) called the carboxysome to trap carbon dioxide (CO2) around the carbon-fixing enzyme, rubisco. Since the CCM is an integral part of cyanobacterial photosynthesis, it is expected to be regulated by many environmental factors that affect photosynthesis. Here, I present findings from studies on Fremyella diplosiphon, a model cyanobacterium that exhibits complementary chromatic acclimation (CCA), wherein photosynthetic pigmentation and efficiency are tuned in response to environmental light cues. A photoreceptor, RcaE, that controls CCA was shown to be important for determining the abundance and stoichiometry of CCM components. The size and abundance of carboxysomes were found to correlate with the ratio of carboxysomal cargo:shell in a ΔrcaE mutant strain, suggesting a role for RcaE in regulating carboxysome morphology. Additionally, F. diplosiphon is one of many cyanobacteria that express an activase-like cyanobacterial (ALC) protein, a homologue to rubisco activase that is an essential protein in plants for rubisco activity. The ALC protein is not coded for in the genomes of many model cyanobacteria but was predicted to be targeted to the carboxysome when present, thus representing an important factor in the nuanced regulation of the CCM. Through contributions to a study highlighting the carboxysomal localization and enzymatic activity of the ALC protein, I provide evidence that ALC is involved in the cellular response to Ci availability. Additionally, computational modeling of the interaction between ALC and rubisco contrasted two potential binding sites and suggested that the interaction could depend on species of origin and post-transcriptional modification. Given the myriad of factors impacting carboxysome regulation, I then analyzed the carbon fixation capabilities of F. diplosiphon strains under variations in light quality, light quantity, and Ci availability. Assessment of carbon fixation behavior utilizing a novel application of carbon response curves to cyanobacteria was consistent with expectations and provided additional insight into which components of the CCM respond to environmental cues. The ΔrcaE mutant exhibited a noteworthy green-light-dependent limitation in carbon fixation, and further analyses suggested that this depended on rubisco levels and the expression of Ci-uptake genes. These findings, alongside the behavior of other cultivation conditions and F. diplosiphon strains, were used to distinguish at a preliminary level the components of carbon response curves in cyanobacteria as a method to assess carbon fixation behavior and the functional impacts of CCM regulation. To Mom, for her love and support throughout this journey. You mean the world to me. To Mr. Brian Cox, in whose biochemistry class I fell in love with the beauty of photosynthesis. and In loving memory of Will Classen, my Opa, who encouraged me to stay curious through research. iv ACKNOWLEDGEMENTS I would first like to express my deep thankfulness to Dr. Beronda Montgomery, my amazing mentor who lives up to the word ‘mentor’ more than I could have ever hoped for. She is a blessing to so many people and does so with such kindness and enthusiasm that it is contagious. I have grown as a scientist and as a person under her care, and I hope to show the world what a treasure we have in her. I am also truly grateful to Dr. Cheryl Kerfeld, who has assisted my work through exciting collaboration and expert discussion. Thanks goes out to Drs. Thomas Sharkey, Claire Vieille, and Kristin Parent, who have served on my committee and have encouraged strong scientific thought through their careful and helpful insight into my work. I am indebted to Alicia Withrow who taught me how to use a transmission electron microscope and supported the many projects with materials, troubleshooting advice, and a sense of wonder at the beauty of cells. I am grateful to Sigal Lechno-Yossef, Shailendra Singh, and Bagmi Pattanaik for their collaborations found in this dissertation. A huge shout out goes to the skilled and enthusiastic undergraduate students I had the opportunity to mentor, Tim Rapp and Kiara Rodriguez-Perez, who both performed amazing work that contributed to this project as well. I would also like to thank the entirety of the Plant Research Laboratory at Michigan State University, who foster a fun and scientific community full of promising discussion and exciting ideas. Lastly, I would like to express my thanks to fellow members of the Montgomery Lab past and present who I have crossed paths with. This work was supported by the U.S. Department of Energy (Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, grant no. DE-FG02-91ER20021 to B.L.M.). v TABLE OF CONTENTS LIST OF TABLES ......................................................................................................................... x LIST OF FIGURES ...................................................................................................................... xi KEY TO ABBREVIATIONS ..................................................................................................... xiv CHAPTER 1 ................................................................................................................................... 1 Balancing Photosynthesis and the Carbon Concentrating Mechanism in Cyanobacteria ...... 1 CHAPTER 2 ................................................................................................................................... 7 RcaE-Dependent Regulation of Carboxysome Structural Proteins Has a Central Role in Environmental Determination of Carboxysome Morphology and Abundance in Fremyella diplosiphon ............................................................................................................................. 7 2.1 Abstract ..................................................................................................................... 8 2.2 Importance ................................................................................................................ 8 2.3 Introduction .............................................................................................................. 9 2.4 Results .................................................................................................................... 12 2.4.1 RcaE regulates carboxysome size and abundance in F. diplosiphon ........... 12 2.4.2 RcaE regulates carboxysome-associated gene expression and protein accumulation in F. diplosiphon .............................................................................. 13 2.4.3 RcaE regulates the response of carboxysome structure to changes in light quality and intensity in F. diplosiphon ................................................................... 20 2.4.4 RcaF and RcaC do not function with RcaE in the regulation of carboxysome-associated gene expression in F. diplosiphon .................................. 21 2.4.5 RcaE-dependent regulation of cell shape and intracellular reactive oxygen species levels are not correlated with the regulation of carboxysome structure in F. diplosiphon ..................................................................................... 22 2.4.6 The structures of polyphosphate bodies are also regulated by RcaE in F. diplosiphon ............................................................................................................. 26 2.4.7 Total carboxysome population size or volume is regulated by RcaE in F. diplosiphon ............................................................................................................. 28 2.5 Discussion ............................................................................................................... 29 2.6 Materials and methods ............................................................................................ 34 2.6.1 Culture conditions ........................................................................................ 34 2.6.2 TEM and EDX analysis ................................................................................ 35 2.6.2.1 TEM analysis of sectioned cells ......................................................... 35 2.6.2.2 Negative-staining TEM analysis of whole cells ................................ 36 2.6.2.3 EDX STEM analysis .......................................................................... 37 2.6.3 Carboxysome and PPB size and number quantification .............................. 37 2.6.4 qPCR analyses .............................................................................................. 38 2.6.5 Measurement of ROS ................................................................................... 39 2.6.6 Protein extraction ......................................................................................... 40 vi 2.6.7 Quantitative western blot analysis ................................................................ 41 2.6.8 Densitometry analysis .................................................................................. 42 2.6.9 Purification of FdCcmM after expression in E. coli .................................... 42 2.6.10 Protein sequencing analysis of ~30 kDa band identified from anti-CcmM immunoblot of FdCcmM ....................................................................................... 43 2.6.11 Statistical analysis ...................................................................................... 44 2.7 Acknowledgements ................................................................................................ 45 CHAPTER 3 ................................................................................................................................. 46 Cyanobacterial Carboxysomes Contain a Unique Rubisco-Activase-Like Protein .............. 46 3.1 Introduction ............................................................................................................ 47 3.2 Results .................................................................................................................... 50 3.2.1 Bioinformatic analysis of the ALC gene family .......................................... 50 3.2.2 Structural modeling of the ALC ................................................................... 55 3.2.3 Cellular localization of the ALC from F. diplosiphon ................................. 59 3.2.4 The F. diplosiphon ALC has ATPase activity and interacts with rubisco but does not function as rubisco activase ............................................................... 60 3.2.5 F. diplosiphon ALC is upregulated under CO2 enrichment ......................... 64 3.2.6 Ultrastructural characterization of a F. diplosiphon ALC deletion mutant shows misregulated response to CO2 availability .................................................. 65 3.3 Discussion ............................................................................................................... 68 3.4 Materials and methods ............................................................................................ 74 3.4.1 Bioinformatic analyses ................................................................................. 74 3.4.2 Protein homology modeling ......................................................................... 76 3.4.3 Cloning and growth conditions .................................................................... 77 3.4.4 Construction of a methylating plasmid ........................................................ 78 3.4.5 Construction of mutant strains and strains expressing fluorescent fusion proteins ................................................................................................................... 78 3.4.6 Confocal scanning laser microscopy ............................................................ 79 3.4.7 Protein expression and purification from E. coli .......................................... 80 3.4.8 ATPase activity assay ................................................................................... 83 3.4.9 Rubisco and activity assays........................................................................... 83 3.4.10 Turbidity assays........................................................................................... 84 3.4.11 qPCR analysis............................................................................................. 85 3.4.12 TEM analysis............................................................................................... 86 3.5 Acknowledgements ................................................................................................ 86 3.6 Author contribution ................................................................................................ 87 CHAPTER 4 ................................................................................................................................. 88 Binding Options for the Small Subunit-Like Domains of Cyanobacteria to Rubisco .......... 88 4.1 Abstract ................................................................................................................... 89 4.2 Introduction ............................................................................................................ 89 4.3 Methods .................................................................................................................. 92 4.3.1 Protein homology modeling ......................................................................... 92 4.3.2 MSA of ALC SSLDs .................................................................................... 92 4.3.3 Analysis of protein-protein interactions ....................................................... 93 vii 4.4 Binding at the RbcS1 position ................................................................................. 94 4.5 Binding at the equatorial (M) position ................................................................... 98 4.6 Discussion ............................................................................................................... 98 4.7 Acknowledgments ................................................................................................ 100 4.8 Author contribution .............................................................................................. 101 CHAPTER 5 ............................................................................................................................... 102 Linking the Dynamic Response of the Carbon Concentrating Mechanism to Carbon Assimilation Behavior in Fremyella diplosiphon ............................................................... 102 5.1 Abstract ................................................................................................................. 103 5.2 Introduction .......................................................................................................... 103 5.3 Results .................................................................................................................. 108 5.3.1 Carbon assimilation measurements in F. diplosiphon respond to light, inorganic carbon availability, and physiological state ......................................... 108 5.3.2 Effect of non-saturating light on carbon assimilation ................................ 112 5.3.3 Effect of different light intensities during growth on carbon assimilation potential ................................................................................................................ 112 5.3.4 Effect of inorganic carbon availability during growth on carbon assimilation ........................................................................................................... 114 5.3.5 Rates of O2 evolution in F. diplosiphon strains under red and green light ....................................................................................................................... 117 5.3.6 TEM analysis of carboxysome morphology in response to light conditions and carbon availability ........................................................................ 119 5.3.7 Transcriptional regulation of carbon concentrating mechanism components measured by qPCR analysis ............................................................. 123 5.4 Discussion ............................................................................................................. 128 5.4.1 Use of the carbon response curve in cyanobacteria ................................... 128 5.4.2 The low Ci phase of the carbon response curve (<100 ppm CO2s) is Ci-uptake driven ................................................................................................... 130 5.4.3 The high Ci phase of the carbon response curve (>100 ppm CO2s) is responsive to multiple photosynthetic parameters ............................................... 132 5.4.4 Regulation of the ccm operon in F. diplosiphon ........................................ 134 5.4.5 Impact ......................................................................................................... 135 5.5 Materials and methods .......................................................................................... 135 5.5.1 Growth conditions ...................................................................................... 135 5.5.2 Carbon response curve analysis using F. diplosiphon discs ...................... 137 5.5.3 O2 evolution analysis .................................................................................. 138 5.5.4 TEM analysis .............................................................................................. 139 5.5.5 qPCR analysis ............................................................................................ 139 5.5.6 Chlorophyll extraction ................................................................................ 140 5.5.7 Statistical analysis ...................................................................................... 140 5.6 Acknowledgements .............................................................................................. 140 CHAPTER 6 ............................................................................................................................... 142 Conclusions and Perspectives ............................................................................................. 142 viii BIBLIOGRAPHY....................................................................................................................... 148 ix LIST OF TABLES Table 2.1: Quantification of average carboxysome size and average carboxysome number per cell section in WT and ΔrcaE strains of F. diplosiphon from Figure 2.2 .................................... 15 Table 2.2: RNA sequencing data for carboxysome genes from F. diplosiphon SF33 WT and ΔrcaE mutant strains grown under GL or RL conditions ............................................................ 18 Table 2.3: RNA sequencing data for polyphosphate synthesis and degradation genes from F. diplosiphon SF33 WT and ΔrcaE mutant strains grown under GL or RL conditions ................ 29 Table 2.4: qPCR primers used in chapter 2 ................................................................................. 39 Table 3.1: Correlation between ALC subtype and taxonomic subclade ...................................... 53 Table 3.2: Pairwise sequence alignment scores of ALC and RbcL sequences ............................ 53 Table 3.3: Quantification of average carboxysome sizes and average numbers of carboxysomes per cell section in WT and Δalc F. diplosiphon from Figure 3.11 .............................................. 67 Table 4.1: Predicted small subunit interactions of rubisco from F. diplosiphon and Syn7942 ... 96 Table 4.2: Comparison of predicted salt bridges between RbcL and RbcS or the SSLD ............ 97 Table 5.1: Quantification of average carboxysome sizes and numbers per cell section in Figure 5.11 ............................................................................................................................................. 122 Table 5.2: Primers used for qPCR probes in chapter 5 .............................................................. 124 Table 5.3: Relative expression of CCM genes in red versus green light conditions in F. diplosiphon strains ..................................................................................................................... 125 Table 5.4: Relative expression of CCM genes under increasing light intensity ........................ 126 Table 5.5: Relative expression of CCM genes under decreasing carbon availability ............... 128 x LIST OF FIGURES Figure 1.1: Protein-level schematics of β-carboxysomes .............................................................. 5 Figure 2.1: Growth curve of F. diplosiphon WT and ΔrcaE strains under WL .......................... 12 Figure 2.2: Carboxysome structure, size, and abundance determination in F. diplosiphon strains under GL and RL conditions ........................................................................................................ 14 Figure 2.3: Carboxysome operons and qPCR-based gene expression analyses in F. diplosiphon ................................................................................................................................... 16 Figure 2.4: Immunoblot analyses of carboxysome protein accumulation in F. diplosiphon ....... 19 Figure 2.5: Protein sequencing of M* band ................................................................................. 20 Figure 2.6: TEM analysis of cellular morphology of F. diplosiphon strains under GL and RL conditions ..................................................................................................................................... 22 Figure 2.7: qPCR-based gene expression analyses in WT, ΔrcaF, and ΔrcaC strains of F. diplosiphon ................................................................................................................................... 23 Figure 2.8: TEM analysis of ultrastructure of ∆bolA strain of F. diplosiphon ............................ 24 Figure 2.9: Accumulation of ROS and carboxysome structure in F. diplosiphon ....................... 25 Figure 2.10: Assessment of PPB structure via TEM of whole cells of F. diplosiphon strains .... 27 Figure 2.11: Negative staining via TEM of whole cells of a Syn7942 WT strain and a carboxysome-deficient ΔccmK2-ccmO strain grown under WL conditions ............................... 27 Figure 2.12: EDX spectroscopy elemental analysis in STEM mode of negative-stained whole- cell F. diplosiphon ∆rcaE mutant strain grown under RL ........................................................... 28 Figure 3.1: Phylogenetic analysis of cyanobacterial ALC and RbcL sequences ......................... 51 Figure 3.2: Primary and secondary structure comparison of RbcS and the SSLDs of FdALC and CcmM .......................................................................................................................................... 54 Figure 3.3: Structural modeling of FdALC ................................................................................. 56 Figure 3.4: The oligomeric state of purified recombinant FdALC .............................................. 57 Figure 3.5: FdALC evolutionary conservation structural models ............................................... 58 xi Figure 3.6: Co-localization of ALC and RbcL in carboxysomes of F. diplosiphon .................... 60 Figure 3.7: Biochemical activity of recombinant rubisco and FdALC ....................................... 61 Figure 3.8: Effect of RuBP concentration on activity of rubisco purified from WT F. diplosiphon ................................................................................................................................... 63 Figure 3.9: The effect of full length ALC compared to its two separate domains on rubisco aggregation ................................................................................................................................... 64 Figure 3.10: Relative expression of alc and ccmM genes under varying Ci levels ...................... 65 Figure 3.11: alc phenotype in F. diplosiphon ............................................................................ 66 Figure 3.12: Diversity of cellular morphology in the Δalc mutant strain when grown under air enriched for CO2 .......................................................................................................................... 68 Figure 3.13: Construction of Δalc mutation by homologous recombination and verification of complete segregation ................................................................................................................... 79 Figure 4.1: Structural comparison of RbcS and SSLD ................................................................ 94 Figure 5.1: Carbon assimilation response to light and Ci availability ....................................... 109 Figure 5.2: Carbon assimilation response to Ci availability for BG11/HEPES blank ............... 110 Figure 5.3: Chlorophyll a levels versus OD750 for cyanobacteria used in CRC analysis .......... 111 Figure 5.4: Carbon assimilation response to Ci availability in response to various light intensities ................................................................................................................................... 113 Figure 5.5: Growth rates of F. diplosiphon strains under increasing GL-enriched WL intensity ...................................................................................................................................... 114 Figure 5.6: Carbon assimilation response to Ci availability under non-saturating light conditions after acclimation to HL .............................................................................................................. 115 Figure 5.7: Carbon assimilation response to Ci availability after acclimation to various Ci levels .......................................................................................................................................... 116 Figure 5.8: Carbon assimilation response to Ci availability in non-saturating light after acclimation to Ci downshift ....................................................................................................... 117 Figure 5.9: Oxygen evolution of F. diplosiphon strains acclimated to red or green light ......... 118 xii Figure 5.10: TEM analysis of cellular morphology of F. diplosiphon strains under changing light or Ci availability ......................................................................................................................... 120 Figure 5.11: Carboxysome morphology under diverse physiological conditions ..................... 121 xiii KEY TO ABBREVIATIONS A ...................................................... Carbon assimilation, the net rate of CO2 uptake per unit area AA ............................................................................................................................. Ascorbic Acid AAA+ ............................................................ ATPases Associated with diverse cellular Activities ALC .................................................................... Rubisco-Activase-Like protein of Cyanobacteria Ava .................................................................................................................. Anabaena variabilis BG11/HEPES .......................................................... BG11 medium with 20 mM HEPES (pH 8.0) BMC .................................................................................................. Bacterial Microcompartment BMC-H ....................................................... Bacterial Microcompartment Hexameric shell protein BMC-P ....................................................... Bacterial Microcompartment Pentameric shell protein BMC-T ............................. Bacterial Microcompartment Trimeric (pseudo-hexamer) shell protein CA ................................................................................................................... Carbonic Anhydrase CCA ................................................................................. Complementary Chromatic Acclimation CCM .......................................................................................... Carbon Concentrating Mechanism Chla ........................................................................................................................... Chlorophyll a Ci .......................................................................................................................... Inorganic Carbon CI ...................................................................................................................... Confidence Interval CO2 ......................................................................................................................... Carbon dioxide CO2s ........................................................................................................ [CO2] in sample chamber Cq ..................................................................................................... Quantification cycle for qPCR CRC ........................................................................................................... Carbon Response Curve DCBQ ................................................................................................ 2,6-dichloro-p-benzoquinone xiv DCF ............................................................................................. 2’,7’-dichlorodihydrofluorescein DCF-DA ....................................................................... 2’,7’-dichlorodihydrofluorescein diacetate EDX ......................................................................................................... Energy Dispersive X-ray FdALC .................................................................................................... ALC from F. diplosiphon FdCcmM .............................................................................................. CcmM from F. diplosiphon FRET ....................................................................................... Förster Resonance Energy Transfer GL ................................................................................................................................. Green Light HCR ............................................................................................................ High Carbon Requiring Km ...................................................................................................................... Michaelis constant M .......................................................................... Equatorial binding position of SSLD to rubisco M* .............................................................................. 30 kDa band tagged by α-CcmM antibodies M35 .............................................................................. C-terminal fragment of CcmM, CcmM-35 M58 .................................................................................................. Full-length CcmM, CcmM-58 MGL ............................................................................................................... Medium Green Light MRL .................................................................................................................. Medium Red Light MSA ................................................................................................. Multiple Sequence Alignment mT2 ............................................................................................................................. mTurquoise2 O2 ....................................................................................................................... Molecular oxygen ODλ .......................................................................................................... Optical Density at λ (nm) PL ............................................................................................................. Photosynthetic Lamellae PPB ................................................................................................................. Polyphosphate Body PSII .......................................................................................................................... Photosystem II Q .................................................................. Light dosage, i.e. photosynthetic photon flux density xv qPCR ............................................................. Quantitative Real-Time Polymerase Chain Reaction RbcL ......................................................................................................... Large subunit of rubisco RbcS .......................................................................................................... Small subunit of rubisco Rca ....................................................................................................................... Rubisco Activase RL .................................................................................................................................... Red Light ROS ......................................................................................................... Reactive Oxygen Species Rubisco ............................................................. Ribulose-1,5-biphosphate carboxylase/oxygenase RuBP .................................................................................................... D-ribulose-1,5-biphosphate SD ..................................................................................................................... Standard Deviation SE ............................................................................................................................. Standard Error SF33 ............................................. Short-filament wild-type pigmentation strain of F. diplosiphon SSLD .................................................................................................. Small Subunit-Like Domain STEM ....................................................................... Scanning Transmission Electron Microscopy Syn6301 ................................................................................. Synechococcus elongatus PCC 6301 Syn6803 .............................................................................................. Synechocystis sp. PCC 6803 Syn7942 ................................................................................. Synechococcus elongatus PCC 7942 SynCcmM ......................................................... CcmM from Synechococcus elongatus PCC 7942 TEM ......................................................................................... Transmission Electron Microscopy Vmax .......................................................................................................................... Maximum rate WL ................................................................................................................................ White Light WT ....................................................................................................................... Wild-Type Strain YFP ...................................................................................................... Yellow Fluorescent Protein xvi 1 Balancing Photosynthesis and the Carbon Concentrating Mechanism in Cyanobacteria CHAPTER 1 1 Photosynthesis is one of the core energy sources in the global ecosystem. In a series of energy transfers and organic reactions, sunlight is converted into chemical energy and stored in high- energy organic molecules. This is the foundation of the global carbon cycle and reflects the ultimate biological source of the energy found in both food and fuels. Understandably, improving photosynthesis in general is a major area of interest for biotechnology research. Moreover, many disciplines benefit from the study of how regulatory mechanisms enable successful photosynthesis through coordinating a network of inter-dependent cellular processes under dynamic environmental conditions. The main carbon-fixing enzyme ribulose-1,5-biphosphate carboxylase/oxygenase (rubisco) is a component of photosynthesis that is tightly regulated by many mechanisms (see reviews18,84,145,194). This enzyme has two substrates: CO2 and a pentose sugar molecule, RuBP. Through the reactions of the Calvin-Benson cycle, RuBP is regenerated with a net incorporation of CO2 into organic molecules37. On one hand, this reaction is dependent on how quickly RuBP can be recycled, which is in turn dependent on how much energy has been acquired from the light reactions of photosynthesis. However, the reaction also depends heavily on CO2 availability, with atmospheric concentrations of CO2 being rate limiting in many natural conditions55,115. Discerning between these effects through modeling has driven significant advances in understanding photosynthetic rates and parameters55,115. In addition to rate limiting factors, it is also important to consider why overall flux through the light reactions and Calvin- Benson cycle should be kept in balance; rubisco reflects a significant investment of nitrogen and energy stores so should not be in excess, but insufficient rubisco activity can lead to unused light energy causing lethal damage through photo-oxidative stress, such as the formation of reactive 2 oxygen species (ROS) (see review166). Overall, rubisco must be regulated by a substantial network of factors in photosynthetic organisms for efficient and safe photosynthesis. Of the many organisms with the capacity to undergo photosynthesis, cyanobacteria are major contributors to global carbon fixation due to their abundance in diverse aquatic ecosystems58,110. Additionally, cyanobacteria hold a significant role in the evolutionary history of photosynthesis; they have an ancestral relationship with chloroplasts in green algae and higher order plants45,62,225; in addition, their dominance in the Proterozoic ecosphere drove atmospheric oxygenation53,177,220. Since O2 can also react with rubisco in an energetically wasteful side- reaction (photorespiration), the rising O2:CO2 ratio posed an issue to cyanobacteria’s continued ability to photosynthesize, driving the evolution of a carbon concentrating mechanism (CCM). The CCM can be conceptually divided into two aspects: 1) expression of systems that increase inorganic carbon (Ci) uptake capabilities of the cell and 2) the encapsulation of rubisco into a compartment with a protein shell that serves as a barrier to CO2 diffusion. Comprehensive research has established that active transport and CO2 hydration are used to increase Ci uptake into the cell (see review30,155) as HCO3 -, which is only appreciably converted to CO2 in the presence of active carbonic anhydrase (CA). Both rubisco and CA were found to be compartmentalized in proteinaceous bacterial microcompartments (BMCs) called carboxysomes60,91,210, first identified as polyhedral, electron-dense bodies during ultrastructural analysis of various cyanobacteria59,82. Unlike the cellular membrane, the protein shell of the carboxysome serves as a barrier to CO2 diffusion46. Since the high levels of HCO3 - accumulated in the cytosol can diffuse into the carboxysome but are trapped once converted to CO2, 3 encapsulated rubisco experiences a locally high [CO2], with consumption of CO2 further driving cellular Ci uptake. Thanks to this CCM, cyanobacteria are able to grow at relatively low levels of available Ci despite cyanobacterial rubisco having a very high Km for CO2 compared to the rubisco of plants11,86,204. When the CCM is compromised, cyanobacteria can no longer grow at ambient [CO2], exhibiting a high-carbon requiring (HCR) phenotype. As with the Ci-uptake system, significant progress has been made in identifying the composition and biogenesis of the carboxysome. Two types of carboxysomes have been identified depending on the form of rubisco, with cyanobacteria containing Form IA rubisco having α-carboxysomes and Form IB rubisco having β-carboxysomes. Though their components are distinct and there are differences in assembly pathways, the two carboxysomes contain strong parallels in components12,54,91,160,219. In both, the functional unit of the protein shell is a protein domain (pfam00936) that forms six-sided hexameric structures (BMC-H) which can self-assemble into the faces of the carboxysome56,92 (Figure 1.1A). Importantly, the hexameric structure defines a pore; for the major shell protein in β-carboxysomes, CcmK2, it is an ~7Å positively-charged pore92 that is expected to allow the passage of small, negatively charged ions such as HCO3 -120. However, the pore properties are variable in many paralogues of shell proteins36,192, suggesting that shell composition can be used to adjust pore permeability193, and thus associated carboxysome function. Proteins with two pfam00936 domains in tandem (BMC-T) that likely form stacked hexamers, and pentameric proteins (BMC-P) form the vertices (Figure 1.1A). In β-carboxysomes, the assembly of the carboxysome depends on protein-protein interactions and begins with the aggregation of rubisco and CcmM38,213. CcmM proteins contain 3-5 C- 4 Figure 1.1: Protein-level schematics of β-carboxysomes. (A) Carboxysome structure modeled by Raul Gonzalez, Seth Axen, Markus Sutter, Sarah Newnham, Clément Aussignargues and Cheryl Kerfeld, reproduced under a Creative Commons Attribution Share Alike 4.0 International License. Images has been modified to remove protein side views and to enlarge text. (B) Schematic of known interactions between β-carboxysome proteins. CcmK2 is the major shell protein with a close paralogue CcmK1; CcmK3, K4, & K6 are additional paralogs that can form homodimers, heterodimers, & stacked dodecamers192,193, CcmO & CcmP are BMC-T shell proteins, and CcmL is a BMC-P protein. Dark spots indicate approximate relative pore size based on available crystal structures. terminal repeats of a domain with homology to RbcS116,156 and the multiple repeats contribute to a paracrystalline network of rubisco in the cargo of carboxysomes112,113,171,213. The N-terminal domain of CcmM either acts as a γ-class CA itself6, or interacts with cyanobacterial β-class CA CcaA125, in addition to interactions with CcmN and CcmK244,94. Shell protein interactions drive the encapsulation of the cargo with CcmL serving to cap the vertices38,159 (Figure 1.1B). This self-assembly process and the morphology of carboxysomes likely depends on the stoichiometry of carboxysome components113,114,200, highlighting one area with potential for regulation that can also provide insight into control of BMCs in biotechnology. Given the tight and multi-faceted regulation of rubisco in plants, it is expected that the CCM, which enhances rubisco activity in cyanobacteria, is regulated at many levels by environmental cues and photosynthetic rates. The CCM reflects a significant metabolic investment, which is 5 evidenced by the inducible characteristics of many CCM components in response to high light61,76,79,123,199 or low Ci availability31,48,124,214, conditions which increase the need for available Ci. In stable conditions, these large changes occur rapidly (likely within minutes) in the Ci- uptake system and more slowly (on the order of days) for carboxysomes. With the importance for balancing carbon fixation rate and flux through the light reactions in dynamic conditions, cyanobacteria are expected to have nuanced methods to fine-tune factors such as carboxysome morphology, abundance, shell composition, and packing density in response to environmental cues. Fremyella diplosiphon is a freshwater, filamentous cyanobacteria that is notable for its characteristic acclimation to external light cues. This responses is known as complementary chromatic acclimation (CCA) and includes distinct pigmentation phenotypes that allow acclimated cyanobacteria to thrive in different light conditions, such as when transitioning to deeper sections of the water column where red light becomes less available129. In this dissertation, I highlight contributions made toward identifying the impact of a light-sensing photoreceptor responsible for CCA on the regulation of the stoichiometry of CCM components, exploring the potential regulatory role of a rubisco activase homologue in cyanobacteria, and examining the functional impacts of CCM regulation on carbon assimilation rates in the cyanobacterium Fremyella diplosiphon. 6 CHAPTER 2 2 RcaE-Dependent Regulation of Carboxysome Structural Proteins has a Central Role in Environmental Determination of Carboxysome Morphology and Abundance in Fremyella diplosiphon This chapter contains information published in: Rohnke, B. A., Singh, S. P., Pattanaik, B., and Montgomery, B. L. (2018) RcaE-dependent regulation of carboxysome structural proteins has a central role in environmental determination of carboxysome morphology and abundance in Fremyella diplosiphon. mSphere 3, e00617-17. doi: 10.1128/mSphere.00617-17 reproduced under the terms of the Creative Commons Attribution 4.0 International license and modified to incorporate the supplemental information into the body of the text, renumber figures, tables, and references to be consistent with the dissertation, and use abbreviations defined in the KEY TO ABBREVIATIONS. 7 2.1 Abstract Carboxysomes are central to the CCM and carbon fixation in cyanobacteria. Although the structure is well understood, roles of environmental cues in the synthesis, positioning, and functional tuning of carboxysomes have not been systematically studied. F. diplosiphon is a model cyanobacterium for assessing impacts of environmental light cues on photosynthetic pigmentation and tuning of photosynthetic efficiency during CCA, which is controlled by photoreceptor RcaE. Given the central role of carboxysomes in photosynthesis, we investigated roles of light-dependent RcaE signaling on carboxysome structure and function. A ΔrcaE mutant exhibits altered carboxysome size and number, ccm gene expression, and carboxysome protein accumulation relative to wild type (WT). Several Ccm proteins, including carboxysome shell proteins and core-nucleating factors, overaccumulate in ΔrcaE cells relative to WT. Additionally, levels of carboxysome cargo rubisco are lower to unchanged in ΔrcaE compared to WT. This shift in ratios of carboxysome shell and nucleating components to the carboxysome cargo appears to drive carboxysome morphology and abundance dynamics. Carboxysomes are also occasionally mislocalized spatially to the periphery of spherical mutants within thylakoid membranes suggesting that carboxysome positioning is impacted by cell shape. The RcaE photoreceptor links perception of external light cues to regulating carboxysome structure and function, and thus, cellular capacity for carbon fixation. 2.2 Importance Carboxysomes are proteinaceous subcellular compartments, or bacterial organelles, found in cyanobacteria that consist of a protein shell surrounding a core primarily composed of the enzyme rubisco that is central to the CCM and carbon fixation. Whereas significant insights have 8 been gained regarding the structure and synthesis of carboxysomes, limited attention has been given to how the size, abundance and protein composition is regulated to ensure optimal carbon fixation in dynamic environments. Given the centrality of carboxysomes in photosynthesis, we provide an analysis of the role of a photoreceptor RcaE, which functions in matching photosynthetic pigmentation to the external environment during CCA and thereby optimizing photosynthetic efficiency, in regulating carboxysome dynamics. Our data highlight a role for RcaE in perceiving external light cues and regulating carboxysome structure and function and, thus, cellular capacity for carbon fixation and organismal fitness. 2.3 Introduction Some cyanobacterial strains tune photosynthetic capacity to environmental cues, including changes in the availability of light. F. diplosiphon is a filamentous, freshwater cyanobacterium that exhibits CCA, which is a process to tune photosynthetic pigment type and levels to changes in the prevalent wavelengths of external light129. In F. diplosiphon, CCA-associated changes occur primarily in response to the presence and abundance of red versus green wavelengths of light19. In red-enriched light, F. diplosiphon accumulates red-absorbing, green-colored phycocyanin in light-harvesting complexes to maximize light absorption for photosynthesis. Conversely, under green-enriched conditions, F. diplosiphon accumulates green-absorbing, red- colored phycoerythrin for promoting light harvesting. In addition to pigmentation changes, cell shape and filament length also are controlled by light during CCA19. Cyanobacteriochrome (phytochrome-related) photoreceptor RcaE controls both the light-dependent regulation of pigmentation89,209 and cell and filament morphologies23,189 characteristic of CCA in F. diplosiphon. As wavelength-dependent tuning of pigmentation is linked to the maintenance of 9 optimal photosynthetic efficiency39, RcaE has a role in tuning photosynthetic potential to external light cues. In prior studies, we noted a reduction in growth of a ∆rcaE mutant strain relative to WT under ambient air in red light (RL) or green light (GL)187, and that the expression of genes associated with Ci-uptake were generally upregulated in the ∆rcaE mutant relative to WT130. Together, these phenotypes suggest an HCR phenotype associated with defects in bicarbonate uptake, with Ci-uptake, or with some part of the CCM. Apart from suggesting a potential state of Ci deficiency in ΔrcaE cells, the impact of an absence of RcaE on the expression of Ci-uptake genes is particularly significant as light has previously been reported to be required under low Ci conditions for the expression of genes impacting inducible Ci-uptake systems of the CCM in Synechocystis sp. strain PCC 6803 (hereafter Syn6803)123,124. Because light is required for this process, redox or phytochrome signals were implicated in the light-dependent cellular response to low Ci in cyanobacteria155. The regulation of inorganic carbon uptake genes involved in the CCM in a ΔrcaE mutant provided genetic evidence of involvement of the photoreceptor RcaE in responses to low Ci 130. The CCM is modular with distinct components consisting of the Ci-uptake systems at the membrane in addition to the intracellular carboxysome subcompartment14,30,86. The carboxysome is a specialized BMC containing rubisco, which functions in carbon fixation60,91. Although there have been significant insights into the structural make-up of carboxysomes91 and assembly principles of BMC shells201, there have been limited insights about the environmental inputs that regulate the synthesis, positioning, and potential functional tuning of carboxysomes. Prior studies demonstrated that regular distribution and positioning of carboxysomes along the long axis of the 10 cell is critical for maintaining carboxysome partitioning and associated cellular fitness during cell division174. Notably, carboxysomes increase in number in low carbon conditions in WT Syn680348 and Synechococcus strains127,199,219. Additionally, increased light intensity leads to an increase in the transcription of carboxysome genes76,79 and synthesis of carboxysomes in Synechococcus elongatus PCC 7942 (hereafter Syn7942)199. The increase in carboxysome number under elevated light conditions presumably increases carbon fixation capacity as a coordinate and long-term acclimation response to an increase in photosynthetic potential under increased availability of photons to drive electron transport. In addition to extended or continuous high light, the expression of carboxysome-related genes increases in the light cycle of diurnal conditions61,217 or subjective day during circadian growth81. In one proteomic study with Cyanothece, carboxysome proteins also accumulated to higher levels in the light phase of a diel cycle7. The mechanisms regulating environmental regulation of carboxysomes in cells have not received significant experimental attention. Here, we report on the investigation of the regulation of cellular responses to dynamic light conditions, including coordinate regulation of light absorption capabilities and carboxysome number, structure, and function in the CCA-capable F. diplosiphon. A ΔrcaE mutant that is incapable of normal regulation of CCA exhibits smaller and apparently more numerous carboxysomes than WT cells. Thus, we assessed ccm gene expression and protein accumulation in the red and green wavelengths critical for CCA and in WT compared to a ΔrcaE strain. Given the known phenotypes of altered cell shape23 and high accumulation of reactive oxygen species (ROS) in the ΔrcaE strain186, we also assessed whether cell shape and intracellular ROS levels have indirect impacts on carboxysome structure using cell-shape mutants and ROS-mitigating 11 compounds. Our results suggest a role for RcaE, including transcriptional regulation of ccm genes, in controlling carboxysome structure and number that may be linked to functional tuning of carboxysomes in response to external light cues. 2.4 Results 2.4.1 RcaE regulates carboxysome size and abundance in F. diplosiphon RcaE is known to control both light-dependent regulation of pigmentation89,209 and cell and filament morphologies23 in F. diplosiphon. A RcaE-deficient strain of F. diplosiphon grows slower than WT in ambient air187 (Figure 2.1) and has increased expression of Ci-uptake Figure 2.1: Growth curve of F. diplosiphon WT and ΔrcaE strains under WL. Growth rates of the WT and ΔrcaE strains in ambient air were estimated using OD750 measured once every 24 h for 7 d under continuous RL- enriched WL (~35-40 µmol m-2 s-1). Data points represent averages (± SD). *, p < 0.05 (for results of comparisons between the ΔrcaE and WT strains determined using an unpaired t test). genes130, which together suggest that the ΔrcaE mutant has an HCR phenotype. To assess subcellular differences in the ΔrcaE mutant that may underlie such an HCR phenotype, we performed detailed ultrastructure analyses of WT and ΔrcaE strains grown under both RL and GL conditions using transmission electron microscopy (TEM). WT cells were more brick shaped 12 and elongated under GL than RL (Figure 2.2A), and ΔrcaE was spherical in both RL and GL (Figure 2.2A), as previously described for confocal laser scanning microscopy-based images23. Photosynthetic lamellae (PL) were regularly arranged around the cell perimeter in WT cells grown under both RL and GL (Figure 2.2A). By contrast, PL were more irregularly arranged or dispersed in the ΔrcaE mutant cells under GL or RL. Carboxysomes were larger in size in WT cells as compared to ΔrcaE mutant cells, independent of light conditions (Figure 2.2A & 2.2B; Table 2.1). Additionally, a comparison between RL and GL showed that carboxysomes were smaller under GL than RL in both WT and ΔrcaE strains. Although smaller in size, carboxysome number per cell section was significantly greater in ΔrcaE than WT under both RL and GL conditions (Figure 2.2C; Table 2.1). 2.4.2 RcaE regulates carboxysome-associated gene expression and protein accumulation in F. diplosiphon Given the observed differences in carboxysome size, we assessed whether there were mutations in the sequences of known carboxysome genes by amplifying and sequencing targeted genomic regions. Similar to other cyanobacteria containing β carboxysomes, the key components of the carboxysome are encoded in a core ccm operon in F. diplosiphon, with additional shell proteins encoded in disparate satellite locations in the genome (Figure 2.3A). The core ccm operon encodes the shell proteins CcmK2, CcmK1, CcmL and CcmO, as well as other components, including CcmN which is essential for shell assembly and CcmM which facilitates rubisco nucleation (see review91). CcmP is encoded at a separate genomic location (Figure 2.3A). F. diplosiphon is one type of cyanobacterium that contains an expanded set of paralogs for proteins which comprise the carboxysome shell, including CcmK3 and CcmK4. These non-essential 13 Figure 2.2: Carboxysome structure, size, and abundance determination in F. diplosiphon strains under GL and RL conditions. (A) TEM analysis of cellular morphology of F. diplosiphon. Representative images of WT pigmentation strain and ∆rcaE mutant strain under GL and RL. C, carboxysomes indicated by white arrows; PL, photosynthetic lamellae indicated by gray arrowheads. Bar represents 1 µm. Carboxysome (B) size and (C) number measurements of WT and ∆rcaE strains under GL and RL. To determine size, the maximum diameter of at least 25 carboxysomes were measured from each strain under each growth condition and presented as a boxplot. Boxplots were used as they present the entire data population spread, ordered from smallest to largest. The horizontal bold line inside each box corresponds to the median, and the box covers the 2nd and 3rd quartile groups (the middle 50% of all values). The vertical line below the box corresponds to the 1st quartile group (the smallest 25% of all values) and the line above the box corresponds to the 4th quartile group (the largest 25% of all values). Presenting the entire spread of data allows for visualization of differences between population spreads. Corresponding averages (± SE) can be found in Table 2.1. Statistical analyses were conducted using a Welch two sample t test performed in R. Identical letters over bars represent a homogenous mean group (p > 0.05), different symbols indicate a significant difference (p < 0.05) from others. 14 Table 2.1: Quantification of average carboxysome size and average carboxysome number per cell section in WT and ΔrcaE strains of F. diplosiphon from Figure 2.2. GL MRL MGL RL + AA RLa *,# 22 3.0 ± 0.3 6.2 ± 0.3 * 3.8 ± 0.2 27 43 3.5 ± 0.2 6.7 ± 0.3 * 7.1 ± 0.3 * 3.5 ± 0.2 7.2 ± 0.3 * 328 ± 14 340 ± 19 45 106 35 6.4 ± 0.2 * 227 ± 19 # 224 ± 12 * 214 ± 11 * 398 ± 28 ** 250 ± 13 *,**,# 2.8 ± 0.2 ** 174 ± 5 380 ± 317 ± 18 *,** WT ΔrcaEc WT ΔrcaE WT ΔrcaE WT ΔrcaE WT ΔrcaE Carboxy- some size (nm)b Carboxy- somes/cell section Sample size (n) for carboxy- some size measurem ents Sample size (n) for carboxy- somes/ cell section measure- ments a Indicates light conditions under which WT and ΔrcaE cells are grown, i.e., RL, red light at ~10 to 12 µmol m-2 s-1; GL, green light at ~10 to 12 µmol m-2 s-1; MRL, medium red light at ~30 µmol m-2 s-1; MGL, medium green light at ~30 µmol m-2 s-1; RL + AA, red light at ~10 to 12 µmol m-2 s-1 with added ascorbic acid (AA) at 2 mM. b Numbers for carboxysome size and carboxysome/cell section are represented as average ± SE. c Statistical analyses, p < 0.05 indicated as follows: *, WT vs. ΔrcaE in same condition; **, low light vs. medium light for same light quality for the same strain; #, GL vs. RL for same light intensity for the same strain. 186 114 215 105 178 28 47 35 64 71 28 47 29 51 91 paralogs are often at disparate locations from the core ccm operon and provide an expanded set of carboxysome shell subunits that have been hypothesized to afford selective advantages by altering carboxysome shell permeability, and thus function, under dynamic growth conditions159,192. Based on Sanger sequencing performed on PCR-amplified, ccm gene- containing regions of the genome, we identified no mutations in the sequences of known ccm or carboxysome genes (data not shown) in the ΔrcaE strain relative to WT. 15 Figure 2.3: Carboxysome operons and qPCR-based gene expression analyses in F. diplosiphon. (A) Carboxysome-associated genes and operons found in F. diplosiphon. (B to I) Data represent levels of expression of ccm genes, including (B) ccmK1, (C) ccmK2, (D) ccmL, (E) ccmM, (F) ccmN, (G) ccmO, (H) ccmK3, and (I) ccmK4, in WT and ΔrcaE strains grown under GL or RL. Levels of expression of genes are presented relative to the results determined for the internal control orf10B, and the data in each panel are shown relative to the expression level of the gene of interest in WT cells in GL. Bars represent averages (± SD) of data from three independent biological replicates. Identical letters over bars represent homogenous mean groups (p > 0.05); different symbols indicate a statistically significant difference (p < 0.05) from others. 16 We proceeded to assess differences in the expression of ccm genes using data from a prior RNA- sequencing analysis comparing WT and ΔrcaE strains146. All ccm genes, with the exception of ccmO, ccmK3, and ccmP, exhibited significantly increased mRNA levels in the ΔrcaE mutant relative to WT in RNA-seq analysis (Table 2.2). Notably, rbc genes were largely downregulated in the ΔrcaE mutant compared to WT (Table 2.2). We confirmed differences for select ccm genes by quantitative real-time polymerase chain reaction (qPCR) (Figure 2.3B). To assess whether the observed transcriptional responses were also apparent at the protein level, CcmM, CcmK2, and RbcL proteins were examined using immunoblot analyses. CcmM and CcmK2 proteins accumulated to higher levels in ΔrcaE relative to WT (Figure 2.4), reflecting that these factors are regulated at the transcriptional level. CcmM exhibits two forms in cells, due to an internal ribosome binding site on the transcript114. This results in two distinct forms of CcmM, i.e., ~58 kDa (CcmM-58 or M58) and ~35 kDa (CcmM-35 or M35), that accumulate in cyanobacteria. CcmM-58 levels especially were elevated in ΔrcaE relative to WT, although CcmM-35 levels were somewhat elevated particularly in GL (Figure 2.4A). Thus, the ratio of CcmM-58 to CcmM-35 changed in WT relative to the ΔrcaE mutant, in addition to total CcmM levels. Prior analyses in which the ratio of M58:M35 was altered by overexpressing the full length CcmM or CcmM-35 version resulted in an alteration in carboxysome size in Syn7942114. In these studies, a reduction in CcmM-58 relative to CcmM-35 levels resulted in larger carboxysomes, whereas increased CcmM-58 relative to Ccm-35 levels were correlated with smaller carboxysomes114. Notably, another band at approximately 30 kDa was detected by the anti-CcmM antibody (i.e., M* in Figure 2.4A) and accumulated to higher levels in the ΔrcaE mutant relative to WT. We conducted protein sequence analysis to determine the identity of this 17 Table 2.2: RNA sequencing data for carboxysome genes from F. diplosiphon SF33 WT and ΔrcaE mutant strains grown under GL or RL conditions. Genea ccmK2 ccmK1 ccmL ccmM ccmN ccmO lysR rbcL rbcX rbcS ccmK3 ccmK4 ccmP ccaA Avab homo -log Ava_ 4472 Ava_ 4471 Ava_ 4470 Ava_ 4469 Ava_ 4468 Ava_ 4467 Ava_ 4466 Ava_ 3907 Ava_ 3906 Ava_ 3905 Ava_ 4709 Ava_ 4710 Ava_ 4911 Ava_ 2165 No. Reads WT ΔrcaE Fold changec RL vs. GL Fold changed ΔrcaE vs. WT WT ΔrcaE GL RL GL 1439 RL 775 GL RL 2954 2406 0.5* 0.8 2.1** 3.1** 791 696 1612 2592 0.9 1.6* 2.0** 3.7** 362 391 776 1336 1.1 1.7* 2.1** 3.4** 1631 1668 2934 6722 1.0 2.3** 1.8* 4.0** 1061 1409 2073 3137 1.3 1.5 2.0** 2.2* 951 6310 639 6768 6.6** 10.6** 0.7** 1.1 366 397 344 452 1.1 1.3 0.9 1.1 11568 13245 8840 29880 1.1 3.4** 0.8* 2.3 4522 5419 1249 2586 1.2 2.1** 0.3** 0.5 3828 6487 1390 3087 1.7 2.2** 0.4** 0.5 451 371 396 471 0.8 1.2 0.9 1.3 329 153 519 428 0.5** 0.8 1.6 2.8** 32 70 37 61 48 51 34 1.1 0.7 1.5 0.9 178 0.9 3.5** 0.7* 2.9** a Colors indicate which genes are near each other in the genome (black are isolated from others in distinct regions of genome for each. b ORFs were compared against Anabaena variabilis (Ava) ATCC 29413 annotated proteins using BlastX with a cut- off e-value of 0.0001 to determine Ava homolog. c Fold change, differential expression analysis between two light treatments was calculated for each strain. * p < 0.5 or ** p <0.01 is significance value calculated for RL vs. GL counts for each strain. d Fold change, differential expression analysis between two strains was calculated for each light condition. * p < 0.5 or ** p <0.01 is significance value calculated for ΔrcaE vs. WT counts for each light condition. 18 Figure 2.4: Immunoblot analyses of carboxysome protein accumulation in F. diplosiphon. Ccm protein accumulation in SF33 WT strain and ΔrcaE strains under GL and RL conditions are shown in representative blots. Protein extracts (μg of undiluted total protein extract indicated in parentheses) and two-fold dilutions as indicated by dilution factor (df, numbers above lanes) were loaded for assessment of (A) CcmM (75 μg) and (B) CcmK2 (75 μg) and RbcL (20 μg). After blotting, proteins were detected using anti-CcmM (1:5000 dilution, 3 min exposure), anti- CcmK2 (1:3000 dilution, 1 min exposure), or anti-RbcL (1:20000 dilution, 4 min exposure) antibodies. For panel A, distinct CcmM variants are indicated, which include F. diplosiphon versions of full-length CcmM-58 (M58), Ccm-35 (M35) that is derived from an internal ribosome entry site114, and a reproducibly observed ~30 kDa band that we designated M* which is also observed in Escherichia coli expressed, N-terminal or C-terminal His-tagged versions of CcmM purified via Ni-NTA affinity chromatography (Panel A, lower right). M* protein was sequenced and determined to contain peptides which map to regions throughout the full-length Ccm protein (see Figure 2.5). *, indicated a non-specific band detected with the anti-CcmM antibodies. The lower portion of panel A with increased dilutions of soluble protein from the ΔrcaE strain was included to allow comparison of WT and ΔrcaE in a range of protein concentrations that was not saturating for ΔrcaE, given its significantly higher accumulation of CcmM-reactive bands in the same dilution range shown in upper portion of the panel. 19 band and determined that peptides for this protein map throughout the full length CcmM sequence (Figure 2.5), indicating overaccumulation of CcmM-derived bands in ΔrcaE. Figure 2.5: Protein sequencing of M* band. Highlighted regions represent exclusive peptides of the 30-kDa band identified in anti-CcmM immunoblots which map to the F. diplosiphon CcmM sequence. Nine exclusive unique peptides and 12 exclusive unique spectra were identified among 86 total spectra. The identified peptides map 152/552 amino acids (i.e., 28% coverage) of full-length CcmM. As expected based on transcriptional downregulation of rbcL in GL, RbcL protein levels were significantly lower in the ΔrcaE mutant under these conditions (Figure 2.4B). Based on densitometry analysis, RbcL levels were reduced by 64% (SD = 0.07, n = 5) in ΔrcaE compared to WT in GL conditions. Under RL, RbcL protein levels may either decrease slightly or remain roughly constant, in contrast to the observed transcriptional upregulation. We observed a 31% reduction (SD = 0.31, n = 6) in RbcL levels in ΔrcaE compared to WT in RL. Together, these findings suggest an overaccumulation of carboxysome shell proteins and the CcmM protein which functions in nucleating the cargo relative to the levels of the carboxysome rubisco cargo, which are lower in ΔrcaE mutant compared to WT. 2.4.3 RcaE regulates the response of carboxysome structure to changes in light quality and intensity in F. diplosiphon Due to the ability of photoreceptors such as RcaE to respond to light quantity in addition to light quality15,161,164 and the prior correlation of increased carboxysome numbers under increased light 20 intensity in a cyanobacterium199, we assessed carboxysome structure in WT and ΔrcaE strains under a variety of light conditions. Using TEM-based analyses, we measured carboxysome diameter and number/cell section for both strains grown under ~30 µmol m-2 s-1 medium RL (MRL) or GL (MGL). The ΔrcaE strain retained a small carboxysome phenotype relative to WT under both MRL and MGL (Table 2.1). However, the higher light intensity resulted in statistically larger carboxysomes in ΔrcaE under both MRL and MGL conditions, as well as in WT under MGL, when compared to standard (i.e., 10-15 µmol m-2 s-1) light conditions (Table 2.1). However, no difference in size was noted when comparing WT grown under MGL to those grown under MRL or in WT under MRL compared to standard RL (Table 2.1). Thus, the loss of RcaE leads to light-dependent changes in all conditions, even if no changes are observed in WT. 2.4.4 RcaF and RcaC do not function with RcaE in the regulation of carboxysome-associated gene expression in F. diplosiphon RcaF and RcaC function downstream of RcaE in the regulation of pigmentation90. Although, not required for RcaE-dependent regulation of morphology in GL, RcaF and RcaC contribute to morphology regulation under RL conditions24. To determine whether these effectors function downstream of RcaE in the regulation of carboxysome structure in cells, we assessed carboxysomes in ΔrcaF and ΔrcaC mutants. However, carboxysomes were very similar in appearance in these mutants to those in WT cells (Figure 2.6). Thus, RcaE-dependent regulation of carboxysomes does not occur through known response regulators RcaF or RcaC, as carboxysomes do not differ significantly from WT in either ΔrcaF or ΔrcaC mutants. In additional support of this TEM-based observation, the expression of ccm genes is not altered 21 Figure 2.6: TEM analysis of cellular morphology of F. diplosiphon strains under GL and RL conditions. Representative images of WT pigmentation strain (top), ∆rcaF mutant strain (middle), and ∆rcaC mutant strain (bottom). C, carboxysomes indicated by white arrows. Bar represents 1 µm. significantly in either ΔrcaF or ΔrcaC (Figure 2.7). Thus, RcaE appears to function primarily through unknown effectors to regulate carboxysome structure. 2.4.5 RcaE-dependent regulation of cell shape and intracellular reactive oxygen species levels are not correlated with the regulation of carboxysome structure in F. diplosiphon Initial assays indicated that, in addition to being smaller and more numerous in ΔrcaE relative to WT, carboxysomes occasionally were mislocalized among thylakoid membranes rather than the expected location in the cytosol in ΔrcaE mutant cells (Figure 2.2A). Prior studies indicated 22 Figure 2.7: qPCR-based gene expression analyses in WT, ΔrcaF, and ΔrcaC strains of F. diplosiphon. Expression levels of (A) ccmK2 and (B) ccmM genes in WT, ΔrcaF, and ΔrcaC strains grown under GL or RL are shown. Expression data for genes are presented relative to the internal control orf10B, and the data in each panel are shown relative to expression of the gene of interest in WT cells under GL conditions. Bars represent averages (± SD) of data from three independent biological replicates. Identical letters over bars represent homogenous mean groups (p > 0.05); different symbols indicate a statistically significant difference (p < 0.05) from others. movement of carboxysomes from the central cytoplasm to the cell periphery under conditions of low inorganic carbon levels127. To determine whether this mislocalization phenotype or the observed carboxysome structural defect phenotype were primarily correlated with the spherical cell shape of the ΔrcaE mutant or other parameters, we assessed another spherical mutant of F. diplosiphon, i.e., ΔbolA189. BolA is a morphogene shown to be involved in regulation of cell shape in a number of bacteria3,173. The deletion of bolA is associated with large spherical cell shape in a ΔbolA mutant of F. diplosiphon189 and its overexpression induces spherical cell morphology190. The ΔbolA mutant exhibited WT-sized carboxysomes; yet, these structures occasionally were mislocalized and found closer to the periphery of the thylakoid membranes, rather than centrally in the cytosol, in cells (Figure 2.8). Given the prior recognition that redox state of the cell may impact CcmM activity148, we investigated whether high ROS levels characteristic of ΔrcaE cells may contribute to the observed disruptions in CcmM levels and carboxysome phenotypes observed in this strain. As previously reported ΔrcaE cells accumulated elevated levels of ROS186,188 (Figure 2.9). Thus, we 23 Figure 2.8: TEM analysis of ultrastructure of ∆bolA strain of F. diplosiphon. One representative image is shown. C, carboxysomes indicated by white arrows. Bar represents 1 µm. investigated whether intracellular ROS accumulation is correlated with the smaller carboxysomes apparent in the ΔrcaE strain. To investigate the potential role of ROS in regulating carboxysome size, we treated ΔrcaE and WT cells with the ROS-mitigating antioxidant ascorbic acid (AA)186. AA-treated ΔrcaE cells exhibited reduced intracellular ROS levels compared to the untreated parental ΔrcaE strain186 (Figure 2.9A). However, there were no significant differences between the size of carboxysomes in ΔrcaE in the presence or absence of AA, and carboxysomes were significantly smaller than WT controls in each case (Figure 2.9B & 2.9C). Furthermore, the aforementioned ΔbolA strain also has elevated intracellular ROS levels189,190, which are not correlated with a change in carboxysome size in this strain relative to WT (Figure 2.8). 24 Figure 2.9: Accumulation of ROS and carboxysome structure in F. diplosiphon. (A) ROS-dependent DCF fluorescence and cell component fluorescence in F. diplosiphon under RL after 3 d treatment with (+AA) or without (-AA) ascorbic acid (2 mM) added to the growth medium in SF33 and ΔrcaE strains. The control represents BG-11 medium + DCFH-DA fluorescence without cells. Bars represent the average of FU, arbitrary fluorescent units at 520 nm relative to OD750 (FU520/OD750). Error bars indicate SD. (B) TEM analysis of strains under RL grown in presence of AA. Bars represent 1 µm. (C) Carboxysome size measurements of WT and ∆rcaE strains under RL, with and without AA. To determine size, the maximum diameters of at least 25 carboxysomes were measured from each strain under each growth condition and presented as a boxplot, with the bold line signifying the median diameter, the box representing the 2nd and 3rd quartile groups (the middle 50% of all values), and the lower vertical line corresponding to the 1st quartile group (the smallest 25% of all values) and the upper line corresponding to the 4th quartile group (the largest 25% of all values). Corresponding averages (± SE) can be found in Table 2.1. Statistical analyses were conducted using a Welch two sample t test performed in R. Identical letters over bars represent a homogenous mean group (p > 0.05), different symbols indicate a significant difference (p < 0.05) from others. 25 2.4.6 The structures of polyphosphate bodies are also regulated by RcaE in F. diplosiphon To independently assess whether the smaller and greater number of carboxysomes per cell section of a ΔrcaE mutant observed in thin-section TEM analysis represent a smaller size, yet larger number of total carboxysomes in cells or an alteration in total carboxysome volume, we attempted to assess the whole cell population of carboxysomes. We used negative-staining of whole cyanobacterial cells with TEM analysis97. Results from these analyses indicated a larger number of smaller electron-dense bodies that appeared to have the potential shape of carboxysomes in ΔrcaE cells compared to WT, especially in RL (Figure 2.10). To confirm the identity of these structures, we used negative whole cell staining of Syn7942 WT cells and a carboxysome-deficient strain as potential controls (Figure 2.11). Upon observing similar electron-dense bodies in both of these lines, we conducted combined Scanning TEM (STEM) and energy-dispersive X-ray spectroscopy (EDX) elemental analysis methods to identify the smaller, more numerous structures apparent in ΔrcaE cells. EDX analyses indicated that the bodies observed were polyphosphate bodies (PPB) (Figure 2.12). This finding indicated an unexpected role for RcaE in regulating both carboxysome and PPB size and abundance. In bacteria, ppk (encoding polyphosphate kinase 1) and ppx (encoding exopolyphosphatase) are involved in synthesis and degradation of polyphosphate (see review29). Notably, a Δppk mutant in Synechococcus lacked similar dark bodies in negative-staining TEM analysis to those we observed in F. diplosiphon63. Additionally, the Δppk had altered regulation of several ccm genes compared to WT, indicating a potential functional correlation between disruptions in PPB formation and carboxysome synthesis63. Given these observations, we assessed whether ppk and ppx mRNA levels were altered in our RNA-seq data. However, there were no significant 26 Figure 2.10: Assessment of PPB structure via TEM of whole cells of F. diplosiphon strains. (A) Representative TEM images of SF33 WT strain and ∆rcaE mutant strain grown under RL and GL. Dark spots are electron-dense bodies. (B) Quantification of diameter of PPB. To determine size, the diameters of at least 25 PPBs were measured from each strain under each growth condition and presented as a boxplot, with the bold line signifying the median diameter, the box representing the 2nd and 3rd quartile groups (the middle 50% of all values), and the lower vertical line corresponding to the 1st quartile group (the smallest 25% of all values) and the upper line corresponding to the 4th quartile group (the largest 25% of all values). Statistical analyses were conducted using a Welch two sample t test performed in R. Identical letters over bars represent a homogenous mean group (p > 0.05), different symbols indicate a significant difference (p < 0.05) from others. Figure 2.11: Negative staining via TEM of whole cells of a Syn7942 WT strain and a carboxysome-deficient ΔccmK2-ccmO strain grown under WL conditions. To assess whether the electron-dense bodies observed in negative-stained whole cells were carboxysomes or other subcellular structures, we compared TEM images of (A) the WT strain, which was grown in ambient air, and (B) a carboxysome-deficient ΔccmK2-ccmO strain, which has a high-carbon-requiring growth phenotype and thus was grown in 3% CO2. 27 Figure 2.12: EDX spectroscopy elemental analysis in STEM mode of negative-stained whole-cell F. diplosiphon ∆rcaE mutant strain grown under RL. Multiple elements were analyzed for colocalization with electron-dense bodies. (A and C) Phosphate (P). (B) Electron-dense particles seen in STEM analysis. (D) Magnesium (Mg). (E) Potassium (K). The bar represents 2 µm and is applicable to panel A only. differences in the mRNA levels of these genes accumulating in ΔrcaE vs. WT (Table 2.3). Thus, RcaE appears to regulate carboxysomes through transcriptional control of carboxysome genes; yet, the disruption in PPBs in ΔrcaE occurs without significant regulation of expression of genes known to impact PPB formation. 2.4.7 Total carboxysome population size or volume is regulated by RcaE in F. diplosiphon As an alternative method to negative-stain TEM analysis to determine whether the smaller carboxysomes of ΔrcaE represent a smaller size, yet larger total population of carboxysomes in cells, we counted the number of carboxysomes observed in series of TEM thin sections and estimated total number per cell based on prior methods35. These analyses indicated that indeed the ΔrcaE mutant has a larger number of smaller carboxysomes per cell section than does WT under both GL and RL conditions (Figure 2.2C, Table 2.1). 28 Table 2.3: RNA sequencing data for polyphosphate synthesis and degradation genes from F. diplosiphon SF33 WT and ΔrcaE mutant strains grown under GL or RL conditions. Gene ppk ppx Avaa homo -log Ava_ 3165 Ava_ 3530 No. Reads WT ΔrcaE GL 474 RL 349 GL 559 RL 536 Fold changeb RL vs. GL Fold changec ΔrcaE vs. WT WT ΔrcaE GL RL 0.7 1.0 1.2 1.5 410 298 408 427 0.7 1.0 1.0 1.4 a ORFs were compared against Ava ATCC 29413 annotated proteins using BlastX with a cut-off e-value of 0.0001 to determine Ava homolog. b Fold change, differential expression analysis between two light treatments was calculated for each strain. Note: no significant differences were detected using t-test. c Fold change, differential expression analysis between two strains was calculated for each light condition. Note: no significant differences were detected using t-test. Given the correlation between changes in light intensity with changes in carboxysome size, we also assessed whether the number of carboxysomes increased under increased light intensity. Light intensity typically did not alter the average carboxysome number per cell in either strain, except for a slight decrease in WT under MGL (Table 2.1). Similarly, there were no significant differences in carboxysome number when comparing standard RL and GL in either strain. This suggests that the number of carboxysomes per cell is well-maintained in WT, that the number is dependent on the presence of RcaE in F. diplosiphon, and that carboxysome size is primarily sensitive to dynamic photoenvironments. 2.5 Discussion Here, we report a regulatory role for RcaE in maintaining carboxysome size, quantity per cell, and contributing to carboxysome subcellular localization in F. diplosiphon. The ΔrcaE mutant, which lacks cyanobacteriochrome RcaE photoreceptor89, has smaller and more numerous 29 carboxysomes than the parental WT line. These observations provide evidence that RcaE contributes to the regulation of carboxysome size and quantity in F. diplosiphon. Carboxysomes also are mislocalized occasionally within the thylakoid membranes of this strain. Notably, prior studies have reported a shift in location of carboxysomes from the central cytoplasm to the periphery of the cell under levels of reduced Ci availability127. PPB morphology also is disrupted in the ΔrcaE strain, with more numerous and smaller PPBs than observed for WT cells. This is notable given several prior recognized associations between phosphate-rich PPBs and carboxysomes in bacteria. In one study in a proteobacterium, the position of PPBs correlated with the positioning and structure of carboxysomes80. More closely related to the work here, carboxysomes have been previously reported to be closely associated or grouped with PPBs in some cyanobacterial strains109,138. Whether these associations point to functional interaction remains to be definitively determined; however, misregulation of ccm genes in a Synechococcus mutant lacking PPBs hints at a functional association63. Given the noted association of PPBs with DNA in the cytoplasm134,140,181, the physical co-localization may indicate that carboxysomes are also nearby or associated with DNA. This association allows for a number of possible connections between chromosome condensation/de-condensation dynamics, gene expression regulation, and subcellular structures to be explored, especially since the ΔrcaE mutant exhibits disruptions to both carboxysome and PPB morphology. Our observations that only some ccm genes are significantly misregulated, that ccm genes in disparate regions of the genome are misregulated, and that there is no apparent change in expression of ppk and ppx genes which are associated with PPB synthesis in the ΔrcaE mutant, suggest that disruptions to carboxysome and PPB structures do not arise from non-specific changes to 30 chromosome accessibility in the nucleoplasm. However, the shared structural phenotypes of carboxysomes and PPBs, alongside their previously reported associations, most likely highlight a robust interconnectivity (perhaps functional) between these subcellular structures. Taken together, we hypothesize that RcaE could have a role in multiple aspects of carboxysome regulation, including interactions with PPBs, that are likely critical for carboxysome dynamics and function in carbon fixation in a cyanobacterium. RcaE was previously described as the photosensory receptor that controls pigmentation and cellular morphology in F. diplosiphon23,89,209. RcaE works through two known response regulators RcaF and DNA-binding transcriptional regulator RcaC in regulating pigmentation4,21,40,89,90,104–106 and red-light-dependent regulation of cellular morphology24. Notably, however, RcaE does not appear to function through RcaF and RcaC in the regulation of carboxysomes, as ΔrcaF and ΔrcaC mutants have no apparent defects in the regulation of carboxysome size or positioning, nor significant misregulation of expression of major ccm genes, ccmM or ccmK2. In the regulation of cellular morphology, RcaE controls expression of the bolA morphogene189,190. Yet, RcaE also does not impact carboxysome morphology via BolA regulation as a ΔbolA mutant has WT-like carboxysomes. Thus, although RcaE impacts expression of carboxysome genes and carboxysome structure and number, the effectors through which it functions to do so appear independent of known RcaE-regulated effectors controlling pigmentation and cell shape phenotypes characteristic of CCA. Of note, localization of carboxysomes generally may be correlated with cell shape, as carboxysomes are mislocalized to the periphery of cells in mutants with a constitutive spherical 31 morphology, including both the ΔbolA mutant and the ΔrcaE strain. Previously, there have been additional correlations made between carboxysomes and cell shape. Elongated cell division mutants exhibit decreased carboxysome numbers per cell and carboxysome structural defects65. Notably, these mutants also have reduced levels of carboxysome-associated proteins97. Additionally, impairments in cell morphology due to cytoskeleton defects were correlated with altered spatial distribution or mislocalization of carboxysomes in cells174. In addition to its spherical morphology, the ΔrcaE mutant has elevated intracellular ROS levels23,186. Despite both ΔrcaE and ΔbolA strains having elevated ROS186,189, ΔrcaE mutant cells have smaller carboxysomes and ΔbolA mutant cells have WT-sized carboxysomes. Additionally, even when intracellular ROS levels were reduced in ΔrcaE mutant cells treated with an antioxidant, carboxysomes were smaller in cells lacking RcaE. Thus, RcaE appears to have a direct regulatory role in controlling carboxysome morphology and dynamics, rather than indirectly impacting carboxysomes through altering intracellular ROS accumulation. The regulatory role of RcaE related to carboxysome structure and function is linked to transcriptional regulation of ccm and carboxysome-associated genes. In a ΔrcaE mutant, Ccm structural proteins overaccumulate and carboxysome cargo protein, rubisco, underaccumulates relative to levels in WT. This observed shift in the carboxysome protein profile results in reduced cargo and simultaneous elevated levels of the shell and CcmM-58, which may contribute to the generation of smaller, more numerous carboxysomes. 32 F. diplosiphon is able to adjust carboxysome size to a number of changes in its photoenvironment. Carboxysomes in the WT strain respond to increased light availability through an increase in size, with this effect being more pronounced under GL. Carboxysomes also appear to be larger under red light compared to GL at low light levels, but this effect is lost at higher light intensities. These data are consistent with a higher linear electron flow driving a larger need for carbon fixation. Since these general behaviors are not entirely lost in the ΔrcaE strain, more cellular factors are implicated in the light-dependent regulation of carboxysome structure. However, the loss of RcaE severely limits the maximum size of carboxysomes while increasing their number in all light conditions studied. Moreover, more light-dependent differences in carboxysomes are observed in the ΔrcaE strain where WT shows limited light- dependence, suggesting that RcaE is required to maintain carboxysome homeostasis under dynamic photoenvironments. This tendency to regulate carboxysome structure encourages future analyses of the extent to which RcaE-dependent alterations to carboxysome size and distribution can specifically impact carbon fixation. Together, these results suggest that RcaE has a critical role in regulating carboxysome structure, which likely serves to match carbon fixation potential with external light cues. Given the light- dependent regulation of expression of ccm genes in RL vs. GL and altered levels of expression of ccm and carboxysome-associated genes in the RcaE-deficient strain relative to WT, both the structure (i.e., size and quantity) and composition (e.g., elevated ccmL, ccmM, ccmO and ccmN in RL vs. GL, and reduced levels of RbcL in GL vs. RL) of carboxysomes appear to be regulated and, indeed, fine-tuned in response to external light cues. Such a role for RcaE provides a key mechanism for matching carbon-fixation capacity and photosynthetic potential of cells to 33 available light. Given the prior observation that light intensity also regulates carboxysome structure and dynamics76,79,199 and that phytochrome-related photoreceptors respond to light intensity in addition to light quality168,169,211, we propose that RcaE plays a central role in tuning the structure and function of carboxysomes in response to a dynamic photoenvironment to optimize organismal fitness in F. diplosiphon. 2.6 Materials and methods 2.6.1 Culture conditions Two strains of F. diplosiphon were compared in this study: a short-filament wild-type pigment strain (hereafter WT), which was identified as SF3342, and a RcaE-deficient mutant strain (ΔrcaE)89. Strains were grown in BG-11 medium (Fluka, Buchs, Switzerland) with 20 mM HEPES (pH 8.0) (hereafter referred to as BG-11/HEPES) at 28°C with continuous shaking at 175 rpm under continuous light conditions. Liquid starter cultures were inoculated from strains maintained on solidified BG-11/HEPES media (BG-11/HEPES containing 1.5% [w/v] agar) and grown under continuous white fluorescent light (General Electric; model no. F20T12/PL/AQ/WS) at ~15 µmol m-2 s-1. Exponentially growing cultures were diluted to an initial optical density at 750 nm (OD750) of ~0.05 and were transferred to the experimental culture condition as indicated. Absorbance measurements were made with a SpectraMax M2 spectrophotometer (Molecular Devices, Sunnyvale, CA). RL and GL conditions were obtained using monochromatic growth chambers at an intensity of ~10 to 12 µmol m-2 s-1 continuous broad-band RL (CVG sleeved Rosco red 24 fluorescent tubes, General Electric; model no. F20T12/R24) or continuous broad-band GL (CVG sleeved Rosco 34 green 89 fluorescent tubes, General Electric; model no. 20T12/G78) as previously described23. Growth under medium light intensity utilized ~30 µmol m-2 s-1 continuous RL (λmax 660 nm; model no. 2506RD LED Grow Light, LED wholesalers, Hayward, CA) or GL (λmax 530 nm; Sunbow model no. SN 1320001-004, Geneva Scientific LLC). For cultures grown to test supplemental carbon dioxide conditions, we used white light (WL) conditions in a Percival I- 41LL incubator equipped with Phillips Alto II fluorescent lights (Model no. F17T81TL841) under either air or 3% CO2. Light intensities were measured with a LI-250 Light meter (LI-COR, Lincoln, NE) equipped with a quantum sensor (model LI-190SA). 2.6.2 TEM and EDX analysis 2.6.2.1 TEM analysis of sectioned cells For conventional TEM, ~10 mL of cells were harvested from exponentially growing cultures (i.e., OD750 at 0.6 – 0.8) by centrifugation at 5125 x g at room temperature for 6 min. Spent media was decanted, pellets were resuspended in remaining media (~ 200 µL), and transferred to 1.5 mL microfuge tubes. Cells were then centrifuged at 16,000 x g at room temperature for 5 min and remaining medium was removed. Cells were prefixed via resuspension with 1 mL of 2.5% (w/v) glutaraldehyde and 2.5% (w/v) paraformaldehyde in 0.1 M cacodylate buffer and incubated for 5 min at 33°C at 35% power in a Precision Pulsed Laboratory Microwave 9000 (Electron Microscopy Sciences, Hatfield, PA). After 3 x 10 min washes with 1 mL of 0.1 M cacodylate buffer, the cell pellets were resuspended in 2% (w/v) molten agarose in dH2O and then centrifuged for 1 min. The solidified agarose plug was removed from the microfuge tube and the dense, embedded cell pellet was cut into ~1-4 mm cubes. Embedded cells were washed 3 x 10 min with cacodylate buffer and then postfixed with 2% (w/v) osmium tetroxide in 35 cacodylate buffer for 5 min at microwaved at 33°C, 35% power. Cells were then washed 3 x 10 min using 0.1 M cacodylate buffer followed by 3 x 10 min washes using dH2O. Postfixed cells were blocked with 2% (w/v) uranyl acetate (Electron Microscopy Sciences, Hatfield, PA) in dH2O and microwaved for 5 min at 33°C, 35% power, which has been reported to enhance contrast of carboxysomes64. Following 3 x 15 min washes with dH2O, fixed cells were dehydrated in an acetone series (30%, 50%, 70%, 80%, 90%, 95%, 100%, 100%, 100%) in 20 min intervals using either an EMP5160 tissue processor (Boeckeler Instruments, Inc., Tucson, AZ) or manually. Dehydrated samples were infiltrated with Spurr resin (Firm Standard; Electron Microscopy Sciences, Hatfield, PA) in a 3:1, 2:2, 1:3 series of acetone:Spurr for 2-3 h at room temperature or overnight at 4°C at each step. Infiltrated cells were soaked in Spurr resin for 48 h with 3 exchanges of resin, then blocks were cured at 60°C for 48 h. Thin sections were prepared using a PowerTome XL ultramicrotome (Boeckeler Instruments, Inc., Tucson, AZ), and 70-90 nm sections (estimated from silver to gold interference color) were placed on 200 mesh Cu grids (Electron Microscopy Sciences, Hatfield, PA). Grids were stained with 4% (w/v) osmium tetroxide in dH2O for 30 min followed by Reynold’s formula (lead citrate, comprised of lead nitrate and sodium acetate; Electron Microscopy Sciences, Hatfield, PA) for 15 min while covered alongside NaOH pellets. Sections were imaged using a JEOL 100CX TEM (JEOL USA Inc., Peabody, MA) equipped with a MegaViewIII digital camera at an operating voltage of 100 V. 2.6.2.2 Negative-staining TEM analysis of whole cells. For negative-staining TEM analysis of whole cells, 2 mL of cells at an OD750 > 0.2 or 5 mL of cells at an OD750 ≤ 0.2 were harvested by centrifugation at 5125 x g at room temperature for 6 36 min. Spent media was decanted, pellets were resuspended in 15 mL of dH2O, and then cells were centrifuged at 5125 x g at room temperature for 6 min. The supernatant was discarded by aspiration and the pellets were resuspended in 0.5 mL dH2O. A 5-µL aliquot of resuspended cell pellet was placed on a 200 mesh Cu grid coated with Formvar (Electron Microscopy Sciences, Hatfield, PA) and incubated for 2 min at room temperature. The grid was blotted nearly dry with Whatman filter paper. Either 5 µL of 0.1% (w/v) uranyl acetate in dH2O (stained) or 5 µL of dH2O (unstained) was added to the grid and blotted away after 5 sec. Grids were then washed once with 5 µL of dH2O for 5 sec, then blotted nearly dry. Grids were imaged using a JEOL 100CX TEM equipped with a MegaViewIII digital camera at an operating voltage of 100V. 2.6.2.3 EDX STEM analysis EDX analysis was performed both on conventional TEM sections and negative-stained samples using a JEM-2200FS TEM (JEOL USA Inc., Peabody, MA) with an in-column energy filter operated in STEM mode at 200 kV. The analytical work was done with the attached Oxford Instrument INCA system with energy resolution of 140 eV. The images were collected with a Gatan Multiscan camera at 1024x1024 resolution. 2.6.3 Carboxysome and PPB size and number quantification To determine size of carboxysomes and polyphosphate bodies (PPBs), the diameter of at least 25 of each were measured from each strain under each growth condition in TEM images. Analysis was done in the image editing software Paint.net, and we selected the maximum diameter for consistency between irregular shapes. The number of carboxysomes and PPBs were determined by counting positively identified structures in ~30 cell sections using TEM sections (for 37 carboxysomes) or ~ 10 cells using negative-staining TEM (for PPBs) for each strain in each condition. Positive identification of a carboxysome structure (for both size and number) satisfied three criteria: 1) appearance of some sharp edges, 2) moderate electron density in contrast to the cytosol, 3) regular, paracrystalline distribution of electron density within the carboxysome. The negative-staining technique highlighted cell outlines and allowed visualization of the naturally electron-dense PPBs. For both carboxysome and polyphosphate quantification, we used boxplots to display data. Boxplots were used as they present the entire data population spread, ordered from smallest to largest. The horizontal bold line inside each boxplot graph corresponds to the median, and the box covers the 2nd and 3rd quartile groups (the middle 50% of all values). The vertical line below the box corresponds to the 1st quartile group (the smallest 25% of all values) and the line above the box corresponds to the 4th quartile group (the largest 25% of all values). Presenting the entire spread of data allows for visualization of differences between population spreads. Averages (± SE) are also presented for carboxysome size and number in tabulated format. 2.6.4 qPCR analyses The abundance of ccmK1, ccmK2, ccmL, ccmM, ccmN, ccmO, ccmK3, and ccmK4 transcripts in total RNA extracted from GL- and RL-grown WT or ∆rcaE strains of F. diplosiphon were analyzed using the delta delta Cq (ΔΔCq) method as detailed previously189. In brief, total RNA was extracted as described180,188 and reverse transcribed (0.5 μg in a 20 μL reaction mixture) with random primers using the Promega reverse transcription kit according to the manufacturer’s instructions. No reverse transcription control reactions, which lacked reverse transcriptase enzyme in the reverse transcription reaction mixture, were also performed for all samples. After 38 reverse transcription, the reaction mixture was diluted with 30 μL of nuclease-free water and 3 μL of this reaction was used in a 10 μL total qPCR reaction according to manufacturer’s instructions using the Microamp® fast optical 96-well reaction plate with barcode and ABI FAST 7500 Real-Time PCR System (Applied Biosystems) in FAST mode with Fast SYBR Green Master Mix (Applied Biosystems). Primers sets used for each gene and orf10B internal control, the latter expressed equally under GL and RL conditions197, are listed in Table 2.4. The annealing/extension temperature for all primer sets was 60°C, and all primers were verified to produce a single product by melting curve analysis. The abundance of transcripts was determined based on relative quantification with normalization to the reference transcript orf10B. All qPCR experiments were performed with three independent biological replicates and three technical replicates for each biological replicate. All qPCR procedures and analyses were performed according to the MIQE guidelines32. Table 2.4: qPCR primers used in chapter 2. Primer name ccmK1 ccmK2 ccmL ccmM ccmN ccmO ccmK3 ccmK4 orf10B Forward primer/FP (5’-3’) AACGAATTGGCAGGACATACT Reverse primer/RP (5’-3’) GCAGGCGTAGAATCTGTGAA AGGCTTGCACTTCCGATAC TGCTGATGCGATGGTGAA GTCTACTCCTGCACCTACGATA GTCTTCGAGGTGTGAAACTACTG GATTGCTCCCGAAGGTACATATT GGCTTTCGCTCTACGGTATTT TGGCACTCAGATTTATGGTACAG GTCCGAGATGGGTTCATTTAGAG CCATTACCTCCAAGCTCAGTAAA CTCCTACCATCGCTGGAAATC TGCTGCTGGAGAACAAGTAAA GTAAAGTGGATCGGAAGGATGG CAGGCAGTTGGAGCATTAGA TCAGAAACATCGCCACGAATA AGAACTACAGCGTCAGCTTAAT CTGCTTCGCTTTCAGCATTT 2.6.5 Measurement of ROS Reactive oxygen species (ROS) and other peroxide levels were measured using the fluorescent dye 2′,7′-dichlorodihydrofluorescein diacetate (DCF-DA; EMD chemicals, Gibbstown, NJ) according to previously described methods74,186. In brief, aliquots of cells were collected immediately after 39 dilution to a starting OD750 of 0.05 (Day 0) and after 72 h (Day 3) under desired growth conditions. In a dark room, aliquots were incubated with 10 µM (final concentration) of DCF-DA for 1 h at room temperature with rocking. Fluorescence measurements were then taken at 520 nm with excitation at 485 nm, using water to blank. The measurements were normalized by the OD750 of the culture and are directly proportional to total hydroxyl groups in the sample. 2.6.6 Protein extraction After 7 d of growth in the desired condition, cells were harvested by centrifugation at 5125 x g at 4°C for 10 min. Spent media was decanted, then pellets were resuspended in remaining media (~ 200 µL) and transferred to 1.5 mL microfuge tubes. Cells were then centrifuged at 16,000 x g at 4°C for 5 min, the remaining media was aspirated, and the cell pellet mass was recorded using a Mettler Toledo XS104 Analytical Balance (Mettler Toledo, Columbus, OH). The pellets were resuspended in 20 mM Tris-HCl (pH 7.5) with 0.6 M sucrose114, 0.2 mg/mL (w/v) lysozyme, 1X Protease Arrest (G Biosciences, St. Louis, MO), and 5 mM EDTA, at a ratio of 6 mL buffer per gram cell paste and transferred to 15 mL Falcon tubes. Samples were passed through a pre- chilled French Pressure Cell Press (SLM Instruments, Urbana, IL) at 500 PSI a total of three times per sample. Each sample was collected in a 15 mL Falcon tube and then transferred into 1.5 mL microfuge tubes and centrifuged at 16,000 x g at 4°C for 5 min. Following collection of the soluble fraction, the cell pellet was resuspended to the original volume using 20 mM Tris buffer to obtain a resuspended insoluble fraction of nearly equal concentration as the obtained soluble fraction. 40 2.6.7 Quantitative western blot analysis Prior to SDS-PAGE, total protein concentration of soluble lysates was measured using the bicinchoninic acid (BCA) assay (Pierce™ BCA Protein Assay Kit; Thermo Fisher Scientific, Waltham, MA) following the manufacturer’s recommendations. Lysates were then normalized by total protein, with addition of 20 mM Tris (pH 7.5) containing 0.6 M sucrose where needed. Samples normalized by total protein were then diluted with 5X SDS sample buffer, and then a 2- fold dilution series, up to a 32-fold dilution, was conducted using 1X SDS sample buffer. Insoluble fractions and whole cell pellets were resuspended in 1X SDS before loading. Samples were denatured at 95°C for either 1 min (soluble fractions) or 5-10 min (insoluble fractions). Proteins (with expected kilodalton values for monomers shown in brackets) were separated on Tris-HCl gels with 10% acrylamide (CcmM [35 or 60 kDa for the short and long isoforms, respectively] or RbcL [53 kDa]) or 15% acrylamide (CcmK2 [11 kDa]) using Tris-Glycine SDS Running Buffer. After separation by electrophoresis, proteins were transferred to Immobilon-P polyvinylidene difloride membrane (EMD Millipore, Billerica, MA) using a semidry TransBlot Turbo Transfer System (Bio-Rad, Hercules, CA) at 25V (1.0 A max) for 40 min. PVDF membranes were blocked for 1 h at room temperature using 5% (w/v) dry milk in Tris-buffered saline with 0.5% (v/v) Tween-20. After blocking, membranes were probed using polyclonal rabbit antiserum raised against Syn7942 CcmK233 or CcmM64 (α-Ccm antibodies provided by Dr. Cheryl Kerfeld), as well as antiserum raised against Spinacia oleracea RbcL (AS07 218, Lot 1004, Agrisera, Vännäs, Sweden). Primary antibody incubation was performed up to overnight at 4°C. Blots were washed 4 x 10 min in Tris-buffered saline with 0.1% (v/v) Tween before addition of goat-anti-rabbit, HRP-conjugated secondary antibody at a dilution of 1:20,000 for 1 h 41 at room temperature. Following four 10 min washes in Tris-buffered saline with 0.1% (v/v) Tween and two 5 min washes in Tris-buffered saline, HRP signal was detected using FEMTOGLOW™ Western PLUS HRP substrate (Michigan Diagnostics, Royal Oak, MI) on a ChemiDoc XP (Bio-Rad, Hercules, CA) imaging system. 2.6.8 Densitometry analysis Densitometry was performed using ImageLab (Bio-Rad, Hercules, CA) software. Lanes were manually selected and bands were detected using high sensitivity, discarding bands that were clearly staining artifacts. Disc size, which determines the baseline, was set such that it reliably connected the bases of non-overlapping peaks (typically this was a disc size of 10 - 20). Using the same method, total protein was analyzed using Coomassie-stained gels run in parallel to western blots. The ratio of total protein in WT relative to ΔrcaE was analyzed for each dilution factor and found to be nearly 1 in the linear range. 2.6.9 Purification of FdCcmM after expression in E. coli Primers for ccmM from F. diplosiphon were designed with overhanging restriction sites such that the PCR fragment could be introduced into pET28a to add either a N-terminal (using restriction sites for NheI and XhoI) or C-terminal (using restriction sites for NcoI and XhoI) 6x His tag. After standard cloning methods and bacterial transformation, E.coli BL21strains containing each of the two constructs were analyzed to confirm fragment insertion. The expression of ccmM was induced overnight at 30°C using 0.5 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG). Cell pellets were harvested, resuspended in 15mL of native binding buffer (50 mM NaH2PO4, 0.5 M NaCl, pH 8.0) and passed two times through a CF Range Cell Disruptor (Constant Systems Ltd., 42 Daventry, Northants, UK) operated at 15 kPSI in a 4°C cold room. Lysate was then spun at 5125 x g for 15 min at 4°C and the soluble fraction was extracted and incubated with Ni-NTA for 1 h in a purification column (Invitrogen Life Technologies, Carlsbad, CA). Affinity chromatography was performed according to manufacturer’s instructions and bound protein was eluted using native binding buffer containing 250 mM Imidazole. SDS-PAGE analysis was used to identify elution fractions containing purified CcmM. 2.6.10 Protein sequencing analysis of ~30 kDa band identified from anti-CcmM immunoblot of FdCcmM. After electrophoresis of an elution fraction containing purified F. diplosiphon CcmM protein expressed in E. coli with an N-terminal His tag, gel bands ranging from 28-35 kDa were digested as previously described182, with modifications, at the Proteomics Research Technology Support Facility at Michigan State University. Briefly, gel bands were dehydrated using 100% acetonitrile and incubated with 10 mM dithiothreitol in 100 mM ammonium bicarbonate (pH ~8) at 56°C for 45 min, dehydrated again and incubated in the dark with 50 mM iodoacetamide in 100 mM ammonium bicarbonate for 20 min. Gel bands were then washed with ammonium bicarbonate and dehydrated again. Sequencing grade, modified trypsin was prepared to 0.01 μg/µL (w/v) in 50 mM ammonium bicarbonate and ~50 µL of this was added to each gel band so that the gel was completely submerged. Bands were then incubated at 37°C overnight. Peptides were extracted from the gel by water bath sonication in a solution of 60%ACN/1%TCA (v/v) and vacuum dried to ~2 µL. Peptides were then re-suspended in 2% acetonitrile/0.1%TFA to 20 μL. From this, 5 µL were automatically injected by a Thermo EASYnLC 1000 (Thermo Fisher Scientific, Waltham, MA) onto a Thermo Acclaim PepMap RSLC 0.075 mm x 250 mm C18 43 column and eluted over 16 min with a gradient of 5% Buffer B (i.e., B; Buffer B = 99.9% Acetonitrile/0.1% Formic Acid) to 30% B in 1 min, ramping to 90% B at 2 min and held at 90% B for the duration of the run at a constant flow rate of 300 nL/min. Buffer A = 99.9% Water/0.1% Formic Acid. Eluted peptides were sprayed into a ThermoFisher Q-Exactive mass spectrometer (Thermo Fisher Scientific, Waltham, MA) using a FlexSpray spray ion source. Survey scans 2 were taken in the Orbi trap (35000 resolution, determined at m/z 200) and the top ten ions in each survey scan are then subjected to automatic higher energy collision induced dissociation (HCD) with fragment spectra acquired at 17,500 resolution. The resulting MS/MS spectra are converted to peak lists using Mascot Distiller, v2.6 (Matrix Scientific, Boston, MA) and searched against a database containing all cyanobacteria protein sequences and all E.coli protein sequences available from NCBInr (downloaded 2017-07-07 from www.ncbi.nlm.nih.gov) appended with common laboratory contaminants (downloaded from www.thegpm.org, cRAP project) using the Mascot searching algorithm, v 2.5. The Mascot output was then analyzed using Scaffold, v4.8.2 (www.proteomesoftware.com) to probabilistically validate protein identifications. Assignments validated using the Scaffold 1% FDR confidence filter are considered true. 2.6.11 Statistical analysis All experiments include at least three independent biological replicates and results are presented as the mean value (± SD). Statistical analyses were conducted using a Welch two sample t-test performed in R158. The significance level was set at 0.05 for all statistical analyses. 44 2.7 Acknowledgments We are grateful to Dr. Alicia Withrow of the MSU Center for Advanced Microscopy for her extensive assistance with the Transmission Electron Microscope and Dr. Xudong Fan for assistance with EDX analyses. The Ccm antibodies used in this study were graciously provided by Dr. Cheryl Kerfeld. We also thank Dr. Kerfeld and members of her research group, as well as Dr. Danny Ducat and members of his research group, for detailed discussions about experimental approaches and results from our analyses. This work was supported by the U.S. Department of Energy (Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, grant no. DE-FG02-91ER20021 to B.L.M.) and support to Shailendra Singh from the National Science Foundation (grant no. MCB-1243983 to B.L.M.). We also thank Melissa Whitaker (supported by National Science Foundation grant no. MCB-1243983 to B.L.M.) for strain maintenance and culture production. 45 3 Cyanobactarial Carboxysomes Contain a Unique Rubisco-Activase-Like Protein CHAPTER 3 This chapter contains information published in: Lechno-Yossef, S., Rohnke, B. A., Belza, A. C., Melnicki, M. R., Montgomery, B. L., and Kerfeld, C. A. (2019) Cyanobacterial carboxysomes contain a unique rubisco-activase-like protein. New Phytol. early view. doi: 10.1111/nph.16195 reproduced with permission and modified to incorporate the supplemental information into the body of the text, reorder subsection 3.4 (Materials and methods) to be after subsection 3.3 (Discussion), renumber figures, tables, and references to be consistent with the dissertation, and use abbreviations defined in the KEY TO ABBREVIATIONS. Brandon Rohnke was the primary contributor to sections 3.2.2, 3.2.5, 3.2.6, 3.4.2, 3.4.11, & 3.4.12, Tables 3.2 & 3.3, and Figures 3.3, 3.5, 3.10, 3.11, & 3.12. He contributed in part to sections 3.1, 3.2.1, 3.3, & 3.4.1, Figure 3.2, and editing of the overall manuscript. 46 3.1 Introduction Rubisco is the most abundant enzyme in nature and the major enzyme responsible for primary productivity on earth52. The most prevalent type, Form I, is composed of large and small subunits arranged in an L8S8 configuration, and is found in all photosynthetic eukaryotes and β- cyanobacteria (Form IB), as well as in all α-cyanobacteria and certain anoxygenic phototrophs (Form IA). The enzyme has a low catalytic turnover rate, suffers from energetically wasteful side-reactions with O2, and is prone to forming inactive complexes upon binding sugar phosphates, including its own substrate, RuBP. In order to relieve rubisco from these inactive states, plants utilize the molecular chaperone rubisco activase (Rca)152. This enzyme has been the focus of much study, including as part of biotechnological efforts to develop strategies to improve rubisco performance22,131. Rca belongs to the AAA+ family of proteins (ATPases associated with diverse cellular activities), of which many use the energy from ATP hydrolysis to remodel the conformation of another protein136. Accordingly, Rca removes inhibitory sugar phosphate effectors from rubisco172 by coupling ATP hydrolysis with destabilization of the rubisco active site, in a yet- unknown mechanism27. Crystal structures of Rca from tobacco (Nicotiana tabacum) and Arabidopsis thaliana exhibit critical features of the AAA+ fold, including its tendency to oligomerize in hexameric and dynamic helical states73,196. [ATP]-dependent variability of oligomeric state99,149,215 and presence of a redox-sensitive C-terminal extension contribute to the regulation of plant Rca activity by conditions such as light intensity and CO2 concentration152,223. Rca deletion in plants results in an HCR phenotype, which suggests a critical role for the activase in maintaining plants’ capacity for carbon fixation191. 47 By contrast, inactivation of rubisco by sugar phosphates is less clear in cyanobacteria, with most information obtained from organisms that lack the activase-like-protein of cyanobacteria (ALC). Unlike the plant enzyme, cyanobacterial rubisco exhibits a lower affinity for CO2 and RuBP, lower CO2/O2 specificity, and higher maximal carboxylation rate5,147,163,207. However, differences in inhibition of the uncarbamylated enzyme by RuBP have been noted between species. No inhibition by RuBP was observed in unicellular cyanobacteria5,147,163, whereas inhibition with RuBP was reported in the filamentous heterocyst-forming Anabaena variabilis, but with a Ki value curiously similar to the Km measured for RuBP carboxylation9 and an absence of significant inhibition when testing recombinant rubisco in vitro102. From these disparate studies, it is difficult to characterize whether cyanobacterial rubisco is indeed prone to inhibition by RuBP or other sugar phosphates, and moreover whether an activase is required. Indeed, until genome sequences from diverse cyanobacteria became available, it was believed that only very few cyanobacteria contained an ALC107. The lower specificity of the cyanobacterial rubisco to CO2 is likely the result of its compartmentalization in carboxysomes as part of a CCM14,86,91. These bacterial microcompartments encapsulate rubisco and CA within a polyhedral protein shell, which is believed to maintain an internally-elevated CO2 concentration34. The eco-physiologically diverse β-cyanobacteria employ β-carboxysomes, encapsulating Form IB rubisco and an absolutely- conserved γ-class CA named CcmM160,192. CcmM was shown to aggregate rubisco for β- carboxysome assembly; it contains 3-5 copies of SSLDs, accessory domains with sequence homology to RbcS38,112,142. CcmM has two alternative start sites resulting in two versions of the 48 protein: the full-length M58 contains both the CA domain and the SSLDs, while a shorter version – M35 – contains only the SSLDs114. The structure of a single SSLD derived from a CcmM contains an additional alpha helix (helix H1A) not found in RbcS171,213. Additionally, the corresponding M35 was shown to be involved in rubisco nucleation and network formation in a redox-sensitive manner, interacting with the rubisco L8S8 complex equatorially rather than by replacement of an RbcS subunit with one of its SSLDs, as previously predicted213. Given their absence from the most commonly used cyanobacterial laboratory strains, ALCs have received scant attention. Moreover, almost twenty years ago, an ALC deletion mutant in Anabaena variabilis showed only a mild reduction of growth rate in air103. However, with the increased availability of cyanobacterial genomic sequence data, it became apparent that many β- cyanobacteria encode Rca-like genes, consisting of the AAA+ ATPase domain fused to a single C-terminal SSLD222. These ALCs have been proposed to be ancestral to the plant Rca135. Here, we phylogenomically survey ALCs and reveal that they are widespread among β- cyanobacteria but are absent from one major taxonomic subclade of cyanobacteria which is predominated by unicellular species and includes several common lab strains. We found a correlation between ALC and RbcL phylogenetic subtypes encoded in the same genome, suggesting co-evolution between the ALC and RbcL. Structural modeling also suggests conservation of ATPase activity, hexameric assembly, and potential interaction between ALC and rubisco. We biochemically and physiologically characterized the ALC from F. diplosiphon UTEX 481 (also known as Tolypothrix sp. PCC 7601 and Calothrix sp. PCC 7601), demonstrating ALC localization to carboxysomes and its close physical proximity to rubisco 49 encapsulated there. We establish that F. diplosiphon ALC (FdALC) is active as an ATPase, but does not relieve inhibition by RuBP, nor does alc deletion result in obvious growth defect. Instead we find that ALC induces rubisco aggregation in an ATP-dependent manner, and that its deletion affects regulation of carboxysome biogenesis and cell morphology in response to CO2 levels. ALC is not a canonical activase, instead exerting its effect on CO2 fixation at the level of a metabolic module, the carboxysome. 3.2 Results 3.2.1 Bioinformatic analysis of the ALC gene family The ALC protein sequences were identified in cyanobacterial genomes in the Integrated Microbial Genomes database (IMG; https://img.jgi.doe.gov/) (Table S1) based on presence of the AAA+ ATPase domain (pfam00004) and each of the three highly-conserved “non-canonical pore loop” regions defined previously for Rca proteins196. The AAA+ domains of the resulting 133 sequences (Table S1) were aligned together with AAA+ domains from reference rca sequences from plants and algae and subsequently used to build a phylogenetic tree (Figure 3.1), within which six clusters could be manually identified, representing groups with shared sequence similarity. Eight ALC sequences could not be assigned to any of the six clusters (Marked ALC-X in Table S1). All but 10 sequences also contained a C-terminal extension encoding an SSLD (Table S1). The comparison between the predicted cyanobacterial ALC structure and that of plant Rca is discussed in subsection 3.2.2. No genome encodes more than one ALC, thus suggesting a lack of duplication and therefore orthology among the ALCs. 50 Figure 3.1: Phylogenetic analysis of cyanobacterial ALC and RbcL sequences. (A) Unrooted phylogeny of AAA+ domains of ALCs and Rca. The AAA+ domain of 133 ALC sequences were identified in 374 cyanobacterial genomes, aligned with 10 plant and algal Rca reference sequences (Table S1), and used for phylogenetic tree construction. (B) Phylogeny of Form IB RbcL and correspondence with ALC subtypes. RbcL sequences from 335 cyanobacterial genomes were aligned with 13 plant and algal representatives (Table S1), used for phylogenetic tree construction, and were rooted by the Form IA rubisco sequences from α-cyanobacteria (subclade C1, shown collapsed). Vertical bars depict the phylogenetic subclades of the host genomes (based on Shih et al.185). Columns at right show correspondence with ALC subtypes when present in the same genome. (C) Unrooted phylogenetic tree comparing SSLDs from CcmM (179 domains in 51 CcmM sequences) and ALC (56 representative sequences) with RbcS (56 representative sequences). 51 In order to examine the potential interaction and coevolution between rubisco and ALC, cyanobacterial RbcL sequences were aligned with the RbcL sequences from the 13 reference plant and algal species (Table S1). Phylogenetic analysis shows that RbcL sequences generally cluster together by the taxonomic groups (“phylogenetic subclades”) previously determined phylogenomically by Shih et al.16,185, although the branching order between the subclades is different, and some subclades could not be resolved based on RbcL alone (Figure 3.1B). When ALC clade IDs are mapped to the branches of RbcL sequences corresponding to the same genome, it can be seen that all species from subclade B2 lack an ALC. The organisms in this subclade are dominated by the unicellular Chroococcales, and include several model organisms such as Syn6803, Synechococcus PCC 7002, and Cyanothece ATCC 51142, as well as other well-studied strains belonging to Microcystis and Crocosphaera. The ALCs are otherwise distributed widely across all other β-cyanobacteria, with a general correlation between taxonomic subclade, RbcL sequence, and the type of ALC present (Table 3.1). Most notably, the RbcL sequences of heterocyst-forming cyanobacteria from morphological subsections IV and V165 – or subclade B1 – cluster together in one region of the tree, and mostly contain ALC-1. Similarly, the filamentous non-heterocyst-forming species (morphological subsection III) from taxonomic subclade A correspond with ALC-2. The same RbcL clade, however, was intermixed with organisms from taxonomic subclade C2, which lack an ALC, and subclade B3, which generally contain ALC-3. RbcL sequences from organisms in subclade C3 (subsections I and III) clustered in a monophyletic clade and are associated with ALC-4. RbcL sequences from organisms belonging to subclades D, E and F (subsections I and III) also clustered together and were found to contain either ALC-5 or ALC-6 (Figure 3.1B), which were the ALC types branching closest to the plant Rca cluster (Figure 3.1A); in contrast, the plant RbcL sequences are located quite 52 Table 3.1: Correlation between ALC subtype and taxonomic subclade. Reference subclade and subsection names are as published165,185. # members Shih subcladea Rippka subsectiona ALC-1 ALC-2 ALC-3 ALC-4 ALC-5 ALC-6 ALC-X 60 23 16 10 10 6 8 B1 A (B1), B3, C2 C3, (G) (A), D, E E, F IV, V, (II) III I, II, III I, III I, III I, III Various a A subclade or subsection in parentheses denotes a single sequence belonging to that group. Various distant from the D/E/F RbcL subcluster. Overall, ALCs show strong sequence similarity to Rca from green-lineage photosynthetic organisms (62% pairwise identity between consensus sequences for the ALC vs Rca AAA+ domains (Table 3.2). Table 3.2: Pairwise sequence alignment scores of ALC and RbcL sequences. Scores were generated using the LAlign webserver. Consensus sequences were generated in CLC Sequence Viewer from the respective MSA and had no gaps. Protein Sequence 1 Sequence 2 ALC Consensus of Cyanobacteria F. diplosiphon F. diplosiphon N. tabacum Consensus of Cyanobacteria with ALC Consensus of Consensus of Higher- Order Green-Lineage A. thaliana N. tabacum A. thaliana Consensus of Higher- Order Green-Lineage Consensus of Percent Sequence Percent Sequence Identity (%) Similarity (%) 62.2 62.5 61.0 88.1 86.7 85.1 85.1 86.4 95.8 96.6 Cyanobacteria with Cyanobacteria without 90.8 97.6 Form IB RbcL ALC Consensus of Cyanobacteria without ALC F. diplosiphon F. diplosiphon N. tabacum F. diplosiphon ALC Consensus of Higher- Order Green-Lineage A. thaliana N. tabacum A. thaliana S. elongatus 53 85.4 83.2 83.3 94.3 85.2 95.9 95.3 95.3 98.1 96.6 A phylogenetic tree built with RbcS and SSLDs (from either ALC or CcmM) retains the same ALC subclustering and branch ordering as the AAA+-based tree (Figure 3.1C). However, the CcmM-derived SSLDs are interspersed among those from ALC-3, representing a high degree of sequence similarity among all SSLDs, supported by conservation of characteristic primary and secondary structural features (Figure 3.2) previously described for CcmM116. Because SSLDs from CcmM have been shown to interact with rubisco38,44,112,213, the similarity shown here among all SSLDs suggests that those derived from the ALC are likely to interact with rubisco in the same manner. Figure 3.2: Primary and secondary structure comparison of RbcS and the SSLDs of FdALC and CcmM. (A) Schematic of the secondary structure for RbcS from Synechococcus elongatus PCC 6301 (hereafter Syn6301) (blue, PDB: 1RBL), CcmM-SSLD from T. elongatus (turquoise, PDB: 6MR1), or a homology model of FdALC-SSLD (red) generated in PDBsum. Helices are labeled H1-H2, β-strands are labeled B0-4. β, β-turns; γ, γ-turns; red horseshoe, β-hairpins. (B) MSA of F. diplosiphon RbcS, FdALC-SSLD (residues 334 – 424), and three F. diplosiphon CcmM SSLDs (residues 231 – 324, 353 – 447 and 353 – 447). Black highlights, identical residues in all sequences; grey highlights, similar residues; pale blue highlight, conserved RbcS N-terminal domain with the motif ERRYET, has high sequence similarity to a motif found in the helix H1A of the SSLD (E/QRRFRT); green highlights, conserved β-strand 3, pink highlights, a loop region which is conserved only in SSLDs and not in RbcS. (C) Sequence logos of RbcS sequences from 223 β cyanobacteria, 111 CcmM-SSLD domains from 32 β- cyanobacteria and 108 ALC-SSLD sequences. 54 3.2.2 Structural modeling of the ALC In order to compare the ALC to canonical plant Rca, we modeled the ALC from F. diplosiphon (hereafter: FdALC) as a monomer, using 3D crystal structures for Rca from tobacco (PDB: 3T15) and an SSLD from Thermosynechococcus elongatus CcmM (PDB: 6MR1) as templates. FdALC residues 1 – 293 were selected as the target sequence for the Rca template, and residues 334 – 424 were used for the SSLD template. Pairwise alignments of FdALC with Rca and CcmM-SSLD had 86.5% and 80.0% sequence similarity respectively, suggesting a well- conserved AAA+ domain with a flexibly-positioned SSLD extending C-terminally from helix H10 (Figure 3.3A & 3.3B). Because Rca and other AAA+ ATPase proteins function as hexamers, we also predicted the ALC quaternary structure. As shown previously with monomeric Rca196, our monomeric ALC model includes some domain shifts relative to a hexameric model for Rca (Figure 3.3C). Thus, the quaternary structure of FdALC was also modeled using the hexameric model from tobacco as a template (Figure 3.3D). A hexameric assembly of the ALC is also suggested by size exclusion chromatography and dynamic light scattering analysis of the purified protein (Figure 3.4). In order to study the structural conservation of the ALC, we mapped the conservation scores for each position in an ALC sequence alignment on to the full-length FdALC homology models. Patches of strong sequence conservation were found to cluster into sectors within the monomeric 3D model (Figure 3.5A), particularly in a region corresponding to what is conventionally deemed the top side of Rca and related AAA+ hexamers, as previously observed with plant Rca196. When the FdALC is viewed as a hexamer, the top side appears to be almost completely conserved (Figure 3.5B), whereas the bottom and perimeter regions exhibit more sequence 55 Figure 3.3: Structural modeling of FdALC. (A) Schematic representation of the tobacco Rca196 and FdALC domains. C-ext, C-extension; SSLD, Small Subunit-Like Domain. (B) Homology model of full-length FdALC using tobacco Rca and T. elongatus CcmM-SSLD as templates for the AAA+ domain and SSLD, respectively. For the AAA+ domain, helices are labeled H0-H10, β-strands are labeled 1-5. (C) Alignment of the FdALC AAA+ domain homology model (light blue), tobacco Rca (dark blue), and tobacco Rca modeled to fit a hexameric AAA+ domain (orange). (D) Homology model of FdALC hexamer using the modeled tobacco Rca hexamer as a template, viewed from the top side. Monomeric subunits alternate between grey and light blue coloration. 56 Figure 3.4: The oligomeric state of purified recombinant FdALC. (A) Size exclusion chromatography of FdALC resolved on Superdex S200 GL10/30 column. Resolution of standards of known molecular weight on the same column is shown in the inset. The FdALC is eluted in a wide non-symmetrical peak suggesting a mixture of several oligomeric states, and absence of a monomeric form. Similar profile was obtained when tobacco Rca was resolved by SEC196. (B) Dynamic Light Scattering (DLS) of purified FdALC. (DynoPro Nanostar by Wyatt Technology, Santa Barbara, CA was used for the analysis). The majority of analysed structures had an average diameter of 29.3 nm. The hexameric model (Figure 3.3D) has a diameter of 13 nm, but does not include the dynamic SSLD, hence a size of 29.3 nm, could potentially correspond to a hexamer, and is certainly larger than a monomer. variability (Figure 3.5C & 3.5D). Additionally, as the SSLD domain extends from the C- terminus of each ALC subunit, it was modeled pointing toward the bottom side of the hexamer; however, the linker is likely flexible and thus the SSLD may possibly adopt one or more alternative configurations. Nevertheless, in agreement with the nearby variability of the bottom side of the hexamer, both the linker and the SSLD show low sequence conservation, with the exception of one well-conserved region in the SSLD. In the AAA+ domain, the α/β subdomain contains much greater sequence conservation than the α-helical subdomain (Figure 3.5A), which is consistent with plant Rca196, as well as the functional role that the α/β subdomain plays in ATP binding. In particular, residues implicated in ATPase and/or activase function196 show very high conservation scores (Figure 3.5E). 57 Figure 3.5: FdALC evolutionary conservation structural models. Conservation scores based on an MSA of 133 ALC sequences mapped onto the FdALC homology model. (A) Surface representation of the monomeric form. (B) Top, (C) bottom (SSLDs were removed from the plane of view), and (D) side views of the FdALC modeled as a hexamer (Figure 3.3D). (E) Residues of known importance for activase (red labels) and ATPase (black labels, with arginine fingers labeled in blue) function as summarized by Stotz et al.196. View is indicated by the arrow pointing towards the blue colored monomer in the inset. 58 3.2.3 Cellular localization of the ALC from F. diplosiphon If the ALC functions similarly to Rca, it would require proximity to its substrate, rubisco, which is encapsulated inside carboxysomes. To test whether the ALC is localized to carboxysomes, the fluorescent protein mTurquoise2 (mT2) was translationally fused to the C-terminus of full-length FdALC. This construct was expressed in F. diplosiphon cells containing a second construct in which the Yellow Fluorescent Protein (YFP) was fused to the N-terminus of RbcL. When co- expressed, the fluorescence patterns from both FdALC-mT2 and YFP-RbcL show puncta that are characteristic of carboxysome localization38,128,174,199, with both blue and yellow fluorescence channels appearing in overlapping patterns throughout the cells, suggesting that ALC and RbcL proteins are co-localized and thus implying that the ALC is targeted to carboxysomes (Figure 3.6A). Förster resonance energy transfer (FRET)87 was used to further elucidate potential interactions between FdALC-mT2 and YFP-RbcL. After an initial image was collected (Figure 3.6A), the YFP signal in a selected region was intentionally photobleached (using the microscope laser), and a second image of the same field was taken (Figure 3.6B). An increased mT2 fluorescence after photobleaching indicates FRET between the mT2 and YFP (Figure 3.6C). This positive detection of FRET between FdALC-mT2 and YFP-RbcL suggests that the two proteins must be within 10 nm of each other inside the carboxysome, which has an average diameter of about 360 nm in F. diplosiphon, as measured by TEM (Section 3.2.6; Table 3.3). 59 Figure 3.6: Co-localization of ALC and RbcL in carboxysomes of F. diplosiphon. Confocal microscopy was performed on a strain co-expressing plasmids carrying the FdALC-mT2 fusion and another carrying the YFP-RbcL fusion. A representative field of view is presented for blue channel fluorescence (mT2), yellow channel fluorescence (YFP), overlap of the blue and yellow channels, as well as the corresponding differential interference contrast. Scale bars represent 5 m. (A) Image taken before photobleaching. (B) Blue and yellow channel fluorescence of the field shown in (A), after photobleaching of the yellow signal in a targeted region indicated by a white circle. (C) Calculated change in blue and yellow fluorescence before and after photobleaching. Averages from 10 imaged fields are shown. 3.2.4 The F. diplosiphon ALC has ATPase activity and interacts with rubisco, but does not function as rubisco activase Given that FdALC contains the residues essential for ATPase activity (Figure 3.5E), we investigated its activity upon purification as a C-terminal His tag fusion and expression in E. coli (Figure 3.7A). Using an enzyme-linked NADH oxidation assay132,202, ATPase activity in FdALC was confirmed, with a maximum ATP hydrolysis rate (Vmax) of 349 ± 60 mol mg-1 min-1 60 Figure 3.7: Biochemical activity of recombinant rubisco and FdALC. (A) Coomassie-stained SDS-PAGE of proteins purified from E. coli by affinity chromatography used for biochemical assays. Lane 1 – rubisco, Lane 2 – FdALC, Lane 3 – M35. (B) Michaelis-Menten kinetics ATP hydrolysis by FdALC expressed and purified from E. coli, using 124 g protein mL-1 with varying concentrations of ATP (n = 3). (C) Effect of RuBP concentration on rubisco activity. F. diplosiphon rubisco was pre-incubated with different concentrations of RuBP before activation with MgCl2 and bicarbonate followed by incorporation of radioactive bicarbonate (n = 3) (D) Kinetics of rubisco aggregation mediated by the FdALC and the M35. Change of turbidity at 340 nm was followed after adding rubisco with or without FdALC or M35 to the assay. Representative experiment is shown (n = 3). (E) The effect of ATP on ALC-mediated rubisco aggregation. Assay performed as described in (D), with the addition of creatine phosphate, creatine phosphokinase and ATP at stated concentrations to the reaction buffer. Representative experiment is shown (n = 3). (F) Effect of the FdALC on rubisco activity. Rubisco activity following incubation with (green bars) or without (blue bars) RuBP was assayed as in (B), activation step was conducted as described17. The presence of FdALC or bovine serum albumin (as a negative control), and/or 5 mM ATP is stated. Data from a representative biological replicate with 3 technical replicates each are shown (experiment was repeated with two more biological replicates). 61 (Figure 3.7B), which is higher than the Vmax reported for spinach Rca167, 1.5 mol mg-1 min-1. Unlike the spinach Rca, the Michaelis-Menten plot for FdALC is hyperbolic rather than sigmoidal (Figure 3.7B, compared to Figure 3 in Robinson & Portis167), suggesting that under the experimental conditions tested the ATP binding in FdALC may not be cooperative. The calculated Km for FdALC is 0.07 ± 0.02 mM ATP, comparable to the Km for spinach Rca167, implying that the ALC enzyme might operate at a similar range of physiological ATP concentrations. In order to investigate whether FdALC can affect the activity of its cognate rubisco, we first purified recombinant F. diplosiphon rubisco that was expressed in E. coli using a three-plasmid system modified from Aigner et al.2. Rubisco was purified by affinity chromatography, yielding a complex containing large and small subunits (Figure 3.7A), and activity was confirmed showing incorporation of radioactive bicarbonate. In contrast to previous studies showing a lack of RuBP inhibition for cyanobacterial rubisco from various species5,102,147,163, here we have detected inhibition of recombinant F. diplosiphon rubisco carbamylation by RuBP, reaching a calculated maximum of 85% inhibition, with half- maximum of the inhibition occurring at 0.8 ± 0.3 mM RuBP (Figure 3.7C). While variability among the biological replicates (that may be due to variability among the protein preps derived from the E. coli expression system) precludes estimating precise values for inhibition parameters, it is clear that RuBP inhibits F. diplosiphon rubisco. Moreover, rubisco purified directly from F. diplosiphon, likewise is clearly inhibited by RuBP (Figure 3.8). 62 Figure 3.8: Effect of RuBP concentration on activity of rubisco purified from WT F. diplosiphon. Rubisco was purified from F. diplosiphon by differential ammonium sulfate precipitation and ion exchange chromatography. Assay was conducted as described for Figure 3.7C. Average of 3 technical replicates is shown. Because the M35 and the ALC both contain SSLD domains, the effect of the ALC on rubisco was compared to that of recombinant M35 purified from E. coli. Similar to the recently-shown effect of M35213, addition of FdALC to rubisco causes an increase of turbidity, suggesting rubisco aggregation (Figure 3.7D). ALC-mediated rubisco aggregation took place only if the intact ALC was used, and not when either of the two separate domains (AAA+ domain or the SSLD), prepared as two synthetic proteins were mixed with rubisco, emphasizing the importance of both domains in interaction with rubisco (Figure 3.9). This ALC-mediated effect was affected by the presence of ATP (Figure 3.7E). Only a slight increase in turbidity was observed when the ALC was incubated in the absence of rubisco (Figure 3.7D). Rubisco activity was unaffected by the presence of rubisco aggregates (Figure 3.7F, blue bars), and unlike the effect of Rca on plant rubisco, the presence of recombinant FdALC with ATP did not restore activity to rubisco preparations that were pre-inhibited by 4 mM RuBP (Figure 3.7F, green bars), thus indicating that FdALC does not display canonical activase activity under the conditions tested. 63 Figure 3.9: The effect of full length ALC compared to its two separate domains on rubisco aggregation. Rubisco aggregation was tested by mixing 0.25 μM recombinant rubisco with either 2 μM FdALC (blue line) or 6 μM SSLD and 2 μM AAA+ domain (green line) purified separately from pDS2 and pSL283, respectively, as described101. Assay conditions as described for Figure 3.7. Representative result shown (n = 3). 3.2.5 F. diplosiphon ALC is upregulated under CO2 enrichment To confirm whether FdALC is expressed in vivo, we examined transcript levels using quantitative qPCR. Transcript levels of alc and ccmM in response to increasing carbon availability were measured. The expression levels of alc were an average of ~4-fold higher in WT strains grown under air enriched with 3% CO2, compared to Ci-limited cells that had been transferred from CO2 enrichment to air in order to induce Ci-stress from CO2 downshift. Additionally, expression levels also increased upon CO2-upshift for ccmM, which is known to impact carboxysome size and abundance112,170, but this response was dampened in a Δalc mutant strain that was generated to test for a functional role of FdALC (Figure 3.10). 64 Figure 3.10: Relative expression of alc and ccmM genes under varying Ci levels. qPCR data of WT and Δalc mutant strain of F. diplosiphon grown with (high Ci) or without (Air) enrichment of 3% CO2. Some samples grown with CO2 enrichment were transferred to air 18 hr prior to harvesting in order to induce low Ci stress (Ci Stress). Data are expressed based on ΔΔCT analysis using Ci Stress as the reference condition. Bars, 95% confidence interval. 3.2.6 Ultrastructural characterization of a F. diplosiphon ALC deletion mutant shows misregulated response to CO2 availability Similar to prior results for Anabaena variabilis103, a F. diplosiphon Δalc mutant exhibited limited impairments in growth under ambient CO2 and had a similar number of carboxysomes as WT cells (p = 0.372), albeit slightly enlarged (p = 0.029) (Figure 3.11, Table 3.3). Since Δrca mutant lines in plants exhibit a high-carbon requirement for growth191, a detailed ultrastructural analysis of WT and Δalc strains of F. diplosiphon under air, with or without 3% CO2 enrichment was performed, using TEM. Due to the moderately high light level of this chamber for F. diplosiphon growth, cells of both strains exhibited an irregular distribution of PL without CO2 enrichment; 65 Figure 3.11: alc phenotype in F. diplosiphon. (A) Representative images of TEM analysis of cellular ultrastructure of F. diplosiphon WT and Δalc mutant strain. Cells were grown under white light with ambient air with or without the addition of 3% CO2. C, carboxysomes (indicated by white arrows); PL, photosynthetic lamellae (indicated by black arrows). Bars, 0.5 μm. Carboxysome (B) size and (C) number measurements of WT and Δalc strains under air and 3% CO2. To determine size, the maximum diameters of carboxysomes from 20 cell sections of each strain were measured under each growth condition and are presented as a box plot. For carboxysome number, carboxysomes per cell section were counted in an additional 40 cell sections and are also presented as box plots. Carboxysomes per cell section is used as a proxy measurement for the number of carboxysomes in the whole cell and will be less than the total number per cell since it analyzes only a cross-section. Box plots were used as described in Rohnke et al.170 to highlight the population dynamics of the measurements. Corresponding averages (± SE) can be found in Table 3.3. Statistical analyses were conducted using a Welch two-sample t test performed in R. Homogenous mean groups (p > 0.05) are indicated by identical letters above bars; different letters indicate a statistically significant difference (p < 0.05) from others. 66 Table 3.3: Quantification of average carboxysome sizes and average numbers of carboxysomes per cell section in WT and Δalc F. diplosiphon from Figure 3.11. Parameter Carboxysome size (nm)b No. of carboxysomes/cell section Sample size (n) for carboxysome size measurements Sample size (n) for measurements of no. of carboxysomes/cell section Value(s) for indicated strain Aira 3% Enriched CO2 WT Δalcc WT Δalc 362 ± 15 413 ± 17* 436 ± 19** 383 ± 16* 2.1 ± 0.2 2.5 ± 0.4 1.4 ± 0.1** 2.2 ± 0.2* 66 60 77 58 42 60 58 60 a Indicates conditions under which WT and Δalc cells are grown, Air, ambient air; 3% Enriched CO2, ambient air enriched with 3% CO2 b Numbers for carboxysome size and carboxysome/cell section are represented as average ± SE. c Statistical analyses, p < 0.05 indicated as follows: *, WT vs Δalc in same condition; **, CO2 vs. Air in same strain. but a regular distribution of PL around the cell perimeter was generally recovered under 3% CO2 enrichment (Figure 3.11A), although this recovery was not always observed in the Δalc strain (Figure 3.12). Similar to Syn7942199, the WT strain of F. diplosiphon showed a significant decrease (p = 0.012) in the number of carboxysomes per cell section when grown under CO2 enrichment (Figure 3.12A & 3.12C). Additionally, there was a statistically significant increase (p = 0.003) in the carboxysomal diameter in WT under CO2 enrichment conditions (Figure 3.12A & 3.12B). In contrast, the Δalc mutant strain showed no significant differences in carboxysomes when comparing between conditions with or without CO2 enrichment (number p = 0.447, size p = 0.216); instead carboxysome size and number per cell section under CO2 enrichment were statistically indistinguishable from the WT strain grown under air (Figure 3.12B & 3.12C, Table 3.3). 67 Figure 3.12: Diversity of cellular morphology in the Δalc mutant strain when grown under air enriched for CO2. Non-representative (i.e., occasionally occurring) images of TEM analysis of F. diplosiphon Δalc mutant strain cellular morphology. Cells were grown under white light, with 3% CO2. C, carboxysomes (indicated by white arrows); PL, photosynthetic lamellae (indicated by black arrows). Bars, 0.5 μm. 3.3 Discussion Herein, a family of rubisco activase‐like homologs exclusive to beta‐cyanobacteria is characterized, which have been named activase‐like cyanobacterial proteins (ALCs). It was shown that ALCs are broadly distributed across cyanobacteria from different taxonomic groups (Figure 3.1), absent only from the α-cyanobacterial C1 subclade, which contains Form IA rubisco and α-carboxysomes, and from the B2 subclade dominated by the unicellular Chroococcales185. It was shown that the SSLD fused to the C-terminus of nearly all ALCs is extremely similar to the SSLDs of the carboxysome protein CcmM (Figure 3.1C), and that ALCs retain key structural features of plant Rca (Figure 3.3 & Figure 3.5). Using recombinant FdALC, ATPase activity was confirmed in vitro (Figure 3.7B), and its localization to carboxysomes and close proximity to RbcL was demonstrated (Figure 3.6), revealing that it is required for normal carboxysome assembly under certain growth conditions (Figure 3.11). The present study also 68 reveals some intriguing biochemical behavior suggesting that the ALC may induce rubisco network formation, similar to activity recently described for CcmM (Figure 3.7D compared to Wang et al.213). Nevertheless, no effect upon rubisco activity was observed, and thus it remains a mystery whether it functions as an activase, but with a distinctive role in rubisco maintenance. The presence of Rca-like proteins in cyanobacteria is puzzling because cyanobacterial rubisco had previously been shown to lack inactivation by RuBP5,102,147,163, and thus would not need an activase. However, for the first time, RuBP sensitivity of a cyanobacterial rubisco was clearly demonstrated (Figure 3.7B). The concentration required to reach 50% of maximum inhibition was calculated to be 0.8 mM, which is about 10-fold higher than the reported cyanobacterial rubisco Km(RuBP) 9,147,162. In comparison, the concentration required to obtain 50% inhibition of the initial activation rate of Rhodospirillum rubrum rubisco ranged from 65 and 270 M when the incubation performed at 2°C, and the lowest tested RuBP concentration of 0.4 M resulted in more than 50% inhibition of spinach rubisco83. Hence, the cyanobacterial rubisco seems to be slightly less sensitive to RuBP inhibition than the Form II bacterial rubisco, and at least 1000- fold less sensitive than the plant rubisco. If such inhibition occurs in vivo, it would set the stage for requiring an activase to restore activity. The failure herein to detect neither rubisco activity enhancement by the ALC nor restoration of activity following RuBP-inhibition (Figure 3.7F) suggests that either the ALC has some other function distinct from that known for plant Rca, or that its activase function requires an accessory component or regulatory modification that was missing from our system. At the very least, the conservation of A144 (tobacco Rca numbering; FdALC A71), K247 (FdALC K175), and non-canonical pore-loop regions, which are characteristic of Rca activity196 suggests a strong evolutionary connection to Rca function. 69 In addition to the conserved activase residues mentioned above, the ALCs show conservation of residues required for ATP hydrolysis (Figure 3.5E). Accordingly, FdALC showed ATPase activity in vitro (Figure 3.7B), similar to plant Rca, with a Km(RuBP) comparable to that of the plant enzyme (compare Figure 3.7B to report by Robinson and Portis167). The [ATP] measured in cyanobacterial cells25,26, ranging from 165-230 nmol mg chl-1, are about 10-fold higher than those measured in chloroplasts from spinach (25-40 nmol mg chl-1)75,96. However, there is currently no information about the ATP concentration inside the carboxysome, where the ALC is expected to function, moreover about how small metabolites such as ATP traverse the shell in any BMC192,193. The detection of FRET between the ALC-mT2 and the YFP-RbcL fusion proteins upon co- expression in F. diplosiphon (Figure 3.6C) implies that ALC and rubisco are located within 10 nm of each other87, although this does not prove a direct interaction. As the interior of carboxysomes is expected to be densely packed with rubisco, which measures about 10 nm per L8S8 complex22, and the average diameter of F. diplosiphon carboxysomes is about 360 nm, as measured from TEM (Figure 3.11, Table 3.3), it is very likely that the ALC is inside the carboxysome, in close proximity to rubisco. β-carboxysomes are synthesized from the inside out38, with the core forming first by aggregation of rubisco via the M35 form of CcmM consisting of three or more tandem SSLDs, followed by recruitment of the icosahedral shell. It is hypothesized that the ALC gets targeted to the carboxysome through a similar interaction involving its SSLD. 70 Recently, it was demonstrated that M35 can induce aggregation of rubisco in vitro, observable as increased turbidity and evidenced by cryo-EM as the binding of one of the three SSLDs from M35 to a peripheral interface of rubisco involving both RbcL and RbcS213. Because the increase of turbidity could only be induced if a protein with two or more SSLDs was included in the aggregation assay, the authors suggest that rubisco network formation occurs with at least two of the three SSLDs in each M35 polypeptide binding to a different L8S8 rubisco complex, serving as a crosslinker. Surprisingly, it was demonstrated herein that FdALC addition to F. diplosiphon rubisco can induce the same phenomenon of increased turbidity (Figure 3.7D). Because the ALC SSLD shares an extremely high degree of sequence conservation with the SSLDs from CcmM (Figure 3.1C & Figure 3.2), it was expected to bind rubisco L8S8 peripherally in the same manner. Because the ALC is likely to function as a hexamer, SSLDs from the six monomers could link different molecules of rubisco L8S8 complexes, in order for ALC polypeptides to serve as crosslinks. Consistent with this hypothesis, the addition of genetic truncations of ALC consisting of both the SSLD and the AAA+ domain, without the linker between them, did not induce increase in turbidity (Figure 3.9); the effect only occurs with full-length ALC. Because ALC-mediated increased turbidity depends on the absence of ATP (Figure 3.7E), it is possible that increased rubisco networking in the carboxysome at night, while the cellular ATP concentration is low and rubisco is not active, increases rubisco stability. This mechanism of protection might be the cyanobacterial parallel to the plant rubisco protection by binding of the nocturnal inhibitor 2-carboxy-D-arabinitol 1-phosphate (CA1P), and other tightly bound inhibitors which protects the molecule from proteolysis and, perhaps, from reactive oxygen species93,145. Alternatively, or additionally if carboxysome integrity is dynamic, with the degree of rubisco packaging influenced by CO2 levels200, the ALC may play a role in rubisco 71 networking in the context of carboxysome assembly dynamics in response to CO2 acclimation. This is consistent with our expression and ultrastructural data and will require detailed follow up studies. The sensitivity to ATP is a distinguishing trait of the ALC-mediated rubisco aggregation compared to the effect of M35, and might suggest different regulation of rubisco aggregation by the two, apparently redundant, systems. Although there is one study of rubisco and its ALC in Anabaena spp., most studies of cyanobacterial rubisco involved unicellular species which lack an ALC such as Synechocystis sp. PCC 6714, and Synechococcus sp. PCC 7002205, which could contribute to the reported lack of cyanobacterial rubisco inhibition. When an ALC gene was characterized in Anabaena sp. strain CA, several different cyanobacterial strains were screened for its presence107. Only heterocyst- forming cyanobacteria were found to contain an ALC in that study107. With the increasing availability of cyanobacterial genomic sequences it is now clear that many groups of cyanobacteria contain an ALC (this study and Zarzycki et al.222). It is enigmatic that ALCs are absent from the genomes of a large group of unicellular cyanobacteria (mostly subsection I165 clustered in Branch B2185). Because the organisms in this branch do not share a single habitat or environmental milieu, it is reasonable to assume that the last common ancestor to this group has lost its alc gene, and this group has evolved without an ALC. This assumption is also supported by the ability to generate alc mutants without severe growth defects (this study and Li et al.103). The correlation between the phylogenies of RbcL sequences and ALCs (Figure 3.1B) and the relationship of RbcS to the SSLD from the ALC suggest an ancient co-evolution of the two proteins. Furthermore, the nesting of CcmM-derived SSLD sequences within the clade of ALC- 3-derived SSLDs, and the closer proximity of ALC-6 to true RbcS sequences (Figure 3.1C) 72 suggests that the SSLD from ALC may be the more ancient form, predating β-carboxysomes, with CcmM recruiting the SSLD from an ALC. Thus, ALCs may now be "vestigial", explaining their absence from some cyanobacteria. This study establishes for the first time that the ALC is localized to the carboxysomes and is in close proximity to rubisco, which are encapsulated within the carboxysome microcompartment (Figure 3.6). This resembles the green alga Chlamydomonas reinhardtii, another unicellular aquatic chlorophotoautotroph which uses a functionally analogous microcompartment, the pyrenoid, for CO2 concentration around rubisco126, inside which Rca has been shown to co- localize119. Unlike higher plants, where an rca deletion causes severe growth impairment, deletion of rca in C. reinhardtii caused only a moderate decrease of growth rate under ambient CO2, prompting a suggestion that the presence of a CCM may partially compensate for the loss of Rca151. Deletion of the alc gene in our study, as well as in Anabaena variabilis103, similarly did not result in an HCR phenotype, supporting the interpretation that compartmentalization of rubisco may compensate or prevent the catalytic inefficiencies of rubisco. Upon Ci-upshift from air to high CO2, the carboxysomes of F. diplosiphon Δalc mutant failed to decrease in amount per cell nor showed an increased size, in contrast to WT199 (Figure 3.11B & 3.11C), nor did ccmM transcript abundance increase (Figure 3.10). Together with the observation that alc transcript abundance was increased in response to higher CO2 levels (Figure 3.10), these observations reinforce the idea that ALCs play an important role in metabolic response to carbon (and possibly ATP) availability by influencing rubisco packaging within the carboxysome. It is proposed that while the Δalc mutant strain is able to compensate for greater carbon stress under standard conditions, the protein is required for acclimation to help mitigate carbon stress. This 73 culminates in a distinct lack of response to the CO2 surplus. Such carbon stress would be expected if the rubisco in these cells was less active than in WT. The upregulation of alc in response to elevated CO2 further supports ALC’s role in utilizing a CO2 surplus. The ALC does not appear to function as a canonical rubisco activase, enzymatically acting on rubisco. Instead it influences the metabolic response to CO2 availability at the level of the carboxysome. Notably, the seemingly subtle benefits that could be afforded by the ALC strengthen the view that this protein serves as an evolutionary link between the fast, promiscuous carboxysomal rubisco and the Rca-requiring rubisco of higher-order plants. Understanding the involvement of the ALC in regulation of photosynthesis through its modulation of rubisco networking and carboxysome acclimation to CO2 levels could contribute to efforts to improve productivity in cyanobacteria, and potentially in plants. 3.4 Materials and methods 3.4.1 Bioinformatic analyses 353 cyanobacterial genomes retrieved from the Integrated Microbial Genomes database (IMG; https://img.jgi.doe.gov/) were profiled for the presence of the AAA+ domain (pfam00004). The resulting 4562 sequences were subsequently scored for matches to a position-specific substitution matrix (PSSM) generated for each of three conserved pore-loop regions identified in a previously-published alignment of 22 ALCs222. These regions of the conserved pore-loops were found to provide the best distinction between Rca/ALC and other unrelated AAA+ proteins. PSSM generation and scoring was performed using BioBike (http://biobike.csbc.vcu.edu:8003/biologin). Pore-loop sequences used for the generation of 74 PSSM – Loop 1 – pos. 61 – 88 (FdALC position numbering); Loop 2 – pos. 98 – 142; Loop 3 – pos. 150 – 179. Sequences which contained all 3 loops were identified as ALCs. These sequences were aligned with several previously investigated reference Rca sequences from plants and algae196. The sequences of 133 identified ALCs and 10 green-type Rca proteins (Table S1) were aligned using MUSCLE47, and resulting multiple sequence alignments (MSA) were trimmed to retain only the AAA+ domain and edited manually. Phylogenetic analysis of aligned AAA+ domain was done using PHYML69 with empirically-determined substitution rates, SH-like branch supports, and the automated method for tree improvement. Sequence similarity groups resembling subtype clades were identified manually by long internal branch lengths (ALC-1 to ALC-6); eight outliers were not assigned to any clade. The MSA of 133 ALCs was used to generate conservation scores mapped to the homology model using the ConSurf server (http://consurf.tau.ac.il/2016/). ConSurf models were rendered in PyMol. Full length RbcL sequences were retrieved from IMG for the 353 cyanobacterial genomes using BLAST queried with the F. diplosiphon RbcL. Rubisco-like protein sequences were removed based on shorter length and annotation as 2,3-diketo-5-methylthiopentyl-1-phosphate enolase. The resulting 335 RbcL sequences were aligned with 13 green-type reference RbcLs (Table S1) using MAFFT88 with iterative refinement based on local pairwise alignment (L-INS-i), and a phylogenetic tree was constructed using RaxML195 with the rapid hillclimbing search and an automatic amino acid substitution model. Phylogenetic trees were visualized using Archaeopteryx226. 75 For sequence analysis of RbcS and SSLD, proteins with pfam00101, corresponding to RbcS and SSLD were retrieved for 327 cyanobacteria genomes in IMG. A total of 107 representatives of each RbcS and CcmM, and 131 ALC sequences were used as an initial target for HMMSearch153. An HHM generated from 189 representative sequences containing pfam00101 in the RP15 database (http://pfam.xfam.org/family/PF00101), using HMMBuild was used as the HMMSearch query. The SSLDs were retrieved from CcmM and ALC sequences based on their match to the HMM, and representative sequences were aligned using MUSCLE47. The phylogenetic tree was constructed in PHYML69. HMM sequence logos were generated using Skylign.org218 using RbcS sequences from 223 β cyanobacteria, 111 CcmM-SSLD domains from 32 β-cyanobacteria and 108 ALC-SSLD sequences. 3.4.2 Protein homology modeling For ALC, the RaptorX (http://raptorx.uchicago.edu/) web server was used to generate a multi- domain homology model based on both Rca from tobacco (PDB: 3T15) and RbcS from T. elongatus (PDB: 2YBV chain B) with a flexible, linker region in between. Further homology models were made using Swiss-Model216 (https://swissmodel.expasy.org/) based on the SSLD of CcmM (PDB: 6MR1) for the final SSLD model in FdALC (residues 334 – 424), as well as based on the hexameric tobacco Rca structure (PDB: 3ZW6) for the AAA+ domain of FdALC (residues 1 – 288) and RbcS from Syn6301 (PDB: 1RBL chain M) for FdALC residues 317 – 424. In addition, alignment scores between two sequences were calculated using the LAlign webserver (https://embnet.vital-it.ch/software/LALIGN_form.html) in order to evaluate candidate template structures and to compare primary structure conservation. PDBsum 76 (https://www.ebi.ac.uk/thornton-srv/databases/cgi-bin/pdbsum/GetPage.pl?pdbcode=index.html) was used to generate schematics of protein secondary structure. 3.4.3 Cloning and growth conditions Standard molecular biology techniques were used for molecular cloning. Detailed lists of strains, plasmids and oligonucleotides used in this study are provided in Lechno-Yossef et al.101. E. coli strains were grown in LB supplemented with appropriate antibiotics at standard concentrations8. Strains DH5α or DH5αMCR were used for plasmid constructions. DH5αMCR was used to harbor methylating, conjugative and cargo plasmids for conjugation with F. diplosiphon50. Standard antibiotic concentrations of ampicillin at 100 µg mL-1, chloramphenicol at 25 µg mL-1, gentamycin at 50 µg mL-1, kanamycin at 50 µg mL-1, or spectinomycin at 100 µg mL-1 were used. SF3342 was used for strain constructions, and is treated as the WT comparison in all in vivo experiments. F. diplosiphon strains were grown in BG11/HEPES165 at 28 - 30°C in the light (ca. 25-30 µmol photons m-2 s-1) and enriched with 3% CO2, unless otherwise stated, in shaken liquid cultures. For selection of mutants on plates, BG11/HEPES was solidified with 1.2% Difco agar, and 3 g of sodium thiosulfate per liter was added to the medium. Antibiotic concentrations used for selection of F. diplosiphon mutants were kanamycin at 25-50 µg mL-1, or spectinomycin at 10 µg mL-1. 77 3.4.4 Construction of a methylating plasmid F. diplosiphon contains three known restriction enzymes – FdiI is an isoschisomer of AvaII; FdiII - isoschisomer of FspI and FdiIII - isoschisomer of SphI. The current methylating plasmid used for transformation - pJCF173, carries methylases that protect only FdiI and FdiII recognition sequences43. To include a third methylase, protecting the FdiIII recognition sequence, a new methylating construct was made. The plasmid pRL51851, containing M.Eco47II, was used as the basis for this construct. Constructs are described in Lechno-Yossef et al.101. Briefly, the chloramphenicol resistance marker in pRL518 was replaced by a gentamycin resistance cassette to allow replication and selection along with cargo plasmids containing chloramphenicol-resistance markers. Genes for M.FdiII (fdiDRAFT68880) and M.FdiIII (fdiDRAFT17080) were PCR-amplified and cloned downstream of the Amaranthus hybridus psbA promoter, a promoter that has been shown to be constitutively expressed in E. coli49 (Elhai, 1993). The genes with the promoter were then cloned in gentamycin-resistant version of pRL518, and the new construct – pSL17 was tested for protection against digestion with FspI and with SphI. 3.4.5 Construction of mutant strains and strains expressing fluorescent fusion proteins All strains were generated by di-parental mating between an E. coli DH5αMCR, containing the conjugative plasmid pRL443, methylating plasmid, pSL17 and a desired cargo plasmid as described50. Fusion fluorescent proteins were expressed from the GL-inducible promoter for phycoerythrin-coding gene cpeE, or the core phycobilisome gene promoter apcA on a replicating plasmid based on the native plasmid from F. diplosiphon, or an RSF1010-based compatible plasmid. To express the ALC-mT2 fusion, pSN7 was generated in which the coding sequence 78 was expressed from the GL-inducible promoter on a plasmid derived from pPL2.7175. The plasmid pSL198 is based on an RSF1010 replicon and contains the YFP-RbcL coding sequence from the apcA promoter. Detailed plasmid construction is presented in Lechno-Yossef et al.101. The alc knockout-mutant strain was generated by homologous recombination of pSL26, selected on spectinomycin. Complete segregation of the double recombinants was tested by PCR (Figure 3.12). . Figure 3.13: Construction of Δalc mutation by homologous recombination and verification of complete segregation. A suicide plasmid carrying a replacement of the alc gene by the resistance gene to streptomycin and spectinomycin was used to replace the gene in the chromosome of F. diplosiphon. Colony PCR was performed on 6 bacteria-free spectinomycin-resistant colonies using primers SL41 and SL43101. The expected PCR size for the fully segregated mutant is 2531 base pairs, and that of the WT is 1806 base pairs. Four out of the 6 tested colonies appear to be fully segregated deletion mutants of the ALC gene. 3.4.6 Confocal scanning laser microscopy Cell cultures were induced by growth under green light (10 – 15 µmol photons m-2 s-1) for 7 – 10 d and were immobilized on an agarose-covered microscope slide. Slides were observed using Olympus FLUOVIEW FV1000 confocal laser scanning microscope using differential interference contrast optics and fluorescence excitation and emission filters. A 60X, 1.42- 79 numeric-aperture oil immersion objective lens was used. For differential interference contrast imaging, the 488 nm laser was used. For detection of blue-channel fluorescence from the mT2, a 405 nm laser diode was used for excitation and the emission was detected with a band pass filter BA430-470. For detection of yellow-channel fluorescence from YFP, a 488 nm Argon gas laser was used for excitation and the emission was detected with a band pass filter BA535-565. FRET between FdALC-mT2 and YFP-RbcL was conducted as described by Karpova and McNally87. Briefly, an image of fluorescence in a certain field was taken in the blue and yellow channel. Then the yellow signal in a specific region of the imaged field was photobleached by the laser. The same field was imaged again. Olympus FV1000 software was used to calculate the change in fluorescence intensity in both channels in the bleached region as well as in two control regions in each visualized field. Averages from 10 different images are presented in Figure 3.7C. All images were visualized using ImageJ176. Pseudocolors were applied as follows: cyan for mT2 channel, yellow for YFP channel. 3.4.7 Protein expression and purification from E. coli The full length FdALC was cloned in a pCDFDuet1 vector (Novagene, Darmstadt), with a C- terminal 6XHisTag101, and the clone was denoted pAB2. Sequence of F. diplosiphon CcmM starting from amino acid 229 was cloned in the NdeI site of pET28a (Novagene, Darmstadt), generating pSL286 that carries N-terminally His-tagged M35. The F. diplosiphon rbcL gene was cloned in the first polylinker of pCDFDuet1 with a C-terminal StrepII tag. The two genes downstream in the genome, rbcXS, were then cloned immediately downstream with their native intergenic regions, generating pAB9. For expression of rubisco in E. coli, the F. diplosiphon 80 gene for the chaperone Raf157 was cloned in the arabinose inducible plasmid pBAD2472, generating pAB24. For expression of FdALC and M35, cultures of E. coli strain BL21 DE3 (Invitrogen, Carlsbad, CA, USA) harboring pAB2 or pSL286 were grown to optical density at 600 nm of 0.4-0.7 at 37°C, induced with 50 M IPTG, and grown for additional 18 – 24 h at 25°C. For expression of rubisco, BL21 DE3 strain containing pAB9, pAB24 and pGro7 (containing GroES and GroEL139 (TaKaRa Bio USA, Mountain View, CA). Upon inoculation, chaperones were induced with 0.2% (w/v) L-arabinose, and cultures were grown to optical density at 600 nm of 0.4-0.7 before induction with IPTG as above. Cultures were harvested by centrifugation and pellets were either processed immediately or stored at -20°C until purification. For His-tagged protein purification, cell pellets were suspended in lysis buffer (50 mM Tris, pH 8; 200 mM MgSO4; 10% glycerol) containing EDTA-free SigmaFast protease inhibitor cocktail at the recommended concentration (Sigma, St. Louis, MO) and approximately 10 g mL-1 DNAseI at 1 – 2 mL per g fresh cell weight. Cells were lysed by passing twice through a cell disruptor (Constant Systems, Aberdeenshire, UK) at 15 kPSI. Cell lysates were clarified by centrifugation at 45,000 g for 30 min, and the clear lysate was passed through a 0.45 m filter, before adding imidazole to a final concentration of 100 mM and loading on a HisTrap column (GE Healthcare, Little Chalfont, UK), equilibrated with Buffer A (50 mM Tris, pH 8; 200 mM NaCl; 10% glycerol) containing 100 mM imidazole, attached to an AKTA Pure FPLC (GE Healthcare, Little Chalfont, UK). DTT at final concentration of 1 mM was added to all buffers used for FdALC purification, but not to M35 purification buffers. For FdALC purification, column was washed with 5 column volumes of buffer A containing 100 mM imidazole and 5 column volumes of buffer A 81 containing 175 mM imidazole, the sample was eluted in buffer A containing 500 mM imidazole. For M35 purification, after washing the column with 5 column volumes of buffer A containing 100 mM imidazole, the protein was eluted with a gradient of 100 mM to 500 mM imidazole in buffer A over 10 column volumes. No further purification was done. For purification of rubisco, cell pellets were suspended in rubisco purification buffer (50 mM phosphate buffer, pH 7.6; 200 mM KCl; 1 mM EDTA; 1 mM DTT) supplemented with SigmaFast protease inhibitor cocktail at the recommended concentration (Sigma, St. Louis, MO) and approximately 10 g mL-1 DNAseI at 1 – 2 mL per g fresh cell weight. Cell suspensions were passed twice through French Press at 1100 PSI. The lysate was clarified by centrifugation as above and loaded on StrepTrap HP, 5 mL (GE Healthcare, Little Chalfont, UK), equilibrated with rubisco purification buffer. The column was washed with 5 column volumes of rubisco purification buffer and eluted in the same buffer containing 5 mM desthiobiotin (Sigma, St. Louis, MO). For results presented in Figure 3.8, rubisco was purified from F. diplosiphon cultures. F. diplosiphon cells harvested from 1 L dense culture, lysed in buffer containing 25 mM Bicine pH 7.6, 5 mM EDTA, 2 mM DTT, DNAseI and a protease inhibitor cocktail (Sigma, St. Louis, MO) by passing 3 times through French Press. The lysate was clarified by 1-hour centrifugation at 20,000 g. The supernatant was treated with 20% saturated ammonium sulfate, and precipitated proteins were discarded. Rubisco was precipitated by 50% saturation ammonium sulfate solution, suspended in lysis buffer treated with 0.5% triton X-100, and precipitated again with 20% PEG-6000. Precipitated proteins were then suspended in ion exchange loading buffer containing 100 mM phosphate and 1 mM DTT and loaded on a MonoQ HR 16/10 column (GE healthcare, Little Chalfont, UK), attached to an AKTA Pure FPLC (GE Healthcare, Little 82 Chalfont, UK). Gradient with elution buffer containing 100 mM phosphate, 1 M KCl and 1 mM DTT was used to elute rubisco. Proteins were observed on SDS-PAGE gel stained with coomassie blue, and the concentration was quantified using the BCA assay kit (Sigma, St. Louis, MO), against bovine serum albumin standard. 3.4.8 ATPase activity assay ATPase activity was performed by coupling ADP release to NADH oxidation via pyruvate kinase and lactate dehydrogenase, as described202. The reaction buffer contained 50 mM Tris- HCl pH 8.0, 20 mM KCl, 5 mM MgCl2, varying concentrations of ATP, 1 mM phosphoenolpyruvate, 0.3 mM NADH, 12 U/mL pyruvate kinase, and 12 U/mL lactate dehydrogenase. The reaction was initiated by the addition of 2.5 M recombinant ALC, in a final reaction volume of 0.5 mL. The change in absorbance was followed at 340 nm for 2 min using Agilent Cary 60 UV-Vis Spectrophotometer (Agilent, Santa Clara, CA). Activity was calculated assuming 1:1 ratio of ADP:NADH and a 6.22 mM-1cm-1 extinction coefficient. 3.4.9 Rubisco and activity assays Rubisco activity was measured by rate of incorporation of 14CO2. Reactions were carried out with purified rubisco at varied concentrations described below for individual experiments, and conducted at 25°C. To test the inhibition by RuBP (results presented in Figure 3.7C), rubisco (500 g mL-1) was first incubated for 10 minutes with varied concentrations of RuBP, and placed on ice until activation. The inhibited sample was then activated for 10 min at room temperature 83 with 20 mM MgCl2, 20 mM NaHCO3. The activity of the activated enzyme with or without pre- incubation with RuBP was measured by adding 20 L of the activated enzyme into 80 L assay buffer containing 50 mM EPPS pH 8, 5 mM MgCl2, 0.2 mM EDTA, 1 mM RuBP and 15 mM NaHCO3 and trace amount of NaH14CO3 (10 Ci / mL). The reaction was stopped after 1 minute by adding 100 L 1 M formic acid and drying on a hot plate. Each reaction was done in triplicates. Acid-stable radioactivity was measured by a scintillation counter model Tri-Carb 2800TR (Perkin Elmer, Waltham, MA). For testing activation by the ALC (Results presented in Figure 3.7F), rubisco (0.25 M) that has been pre-incubated with or without 4 mM RuBP for 10 minutes, was incubated with 50 mM EPPS pH 8, 20 mM MgCl2, 10 mM NaHCO3, 40 U mL-1 creatine phosphokinase, 4 mM creatine phosphate, 2 mM DTT, and 10% PEG-3350. Five mM ATP was included in some samples, as specified in figure legends. Either BSA (as a negative control) or recombinant ALC were included as specified in Figure 3.7. After 10 min activation 20 L of the activated rubisco were mixed with 80 L of assay buffer, incubated for 2 min and stopped as described above. 3.4.10 Turbidity assays Turbidity assays were conducted as described213. The assay was conducted in 100 L volume in buffer containing 50 mM Tris, pH 8; 10 mM Magnesium acetate; 50 mM KCl; 5 mM DTT. Rubisco (0.25 M, unless otherwise stated) was added to the assay buffer and placed in a spectrophotometric cuvette. Absorbance at 340 nm was followed using the kinetics setting on an Agilent Cary 60 UV-Vis Spectrophotometer (Agilent, Santa Clara, CA), and the additional proteins, FdALC or M35 at 2 M final concentration were mixed in the cuvette after initiation of the measurement. Change of absorbance was followed for 10 min. 84 3.4.11 qPCR Analysis F diplosiphon cells were grown with and without the enrichment of 3% CO2 for 3 d and were normalized to an OD750 of 0.40, then cells grown under 3% CO2 were split into two groups. One group (High Ci) was returned to grow under 3% CO2 enrichment, while the other (Ci Stress) was moved to grow without CO2 enrichment to simulate low Ci stress according to the methods of Wang et al.214). Cells grown without CO2 enrichment (Air) were treated as the intermediate Ci samples. Cells were then grown for 19 hrs, cooled briefly with liquid N2, then harvested by centrifugation at 5,125 x g at 4°C for 10 min. The cell pellet was transferred to a microfuge tube, centrifuged at 13,000 x g at 4°C for 5 min, decanted, and then flash frozen at -80°C. mRNA was extracted with the addition of Trizol reagent (1 mL), vortexing, incubation at 95°C for 8 min, incubation on ice for 8 min, addition of chloroform (200 μL) with brief vortexing, centrifugation at 13,000 x g for 15 min, and transfer of ~500 mL supernatant to a new tube. For RNA purification, an equal volume of isopropanol was added, and the sample was incubated at room temperature for 15 min, followed by centrifugation at 13,000 x g for 8 min. The supernatant was decanted, then 70% ethanol was added, and the sample was dried for 5-10 min following centrifugation at 13,000 x g for 8 min and decanting the supernatant. The RNA pellet was resuspended in 70 μL nuclease-free water and treated with TURBO DNA-free kit (Invitrogen, Madision, WI) according to manufacturer’s directions. mRNA was quantified using a NanoDrop ND-1000 Spectrophotometer, diluted to a concentration of 100 ng/μL, and 250 ng was used for a 25 μL reverse transcription reaction using qScript cDNA SuperMix (Quantabio, Beverly, MA) kit according to the manufacturer’s directions. No reverse transcription reactions were also performed without the addition of reverse transcriptase to evaluate genomic contamination. 85 qPCR was prepared using Fast SYBR-Green master mix (Applied Biosystems, Foster City, CA) according to the manufacturer’s directions in 384-well plates (Applied biosystems). alc, ccmM, and the endogenous control orf10B were amplified using primers specified in Lechno-Yossef et al.101. Samples were run and analyzed using the ABI QuantStudio 7 Flex PCR system (Applied Biosystems) in fast mode. Samples were run in triplicate, with 5 biological replicates from two independent experiments. Statistics were run using a Welch one-sample t test performed in Excel to generate 95% CI. 3.4.12 TEM Analysis TEM analysis and quantification of carboxysome size and number were performed according to the methods of Rohnke et al170. In total, ~60 cell sections were analyzed for carboxysome number in each strain under each condition, with carboxysome diameters measured in 20 of these cell sections. 3.5 Acknowledgements This work was supported by the Office of Science of the US Department of Energy DE-FG02- 91ER20021. The authors would like to thank Dr. Sarah E. Newnham and Dr. C. Raul Gonzalez- Esquer for help with fluorescent tag cloning, Dr. Shailendra P. Singh for assistance with structural modeling of the F. diplosiphon ALC, Dr. Dayana Muzziotti and Dr. Melinda Frame, for help with confocal microscopy, Dr. Sarathi Wijetilleke for help with rubisco assays, Mr. Damien Sheppard and Mr. Donovan Prince Smith for technical laboratory assistance, Dr. 86 Thomas D. Sharkey and Dr. Berkley Walker for assistance with ATP hydrolysis and rubisco assay experimental design, interpretation of results and critical reading of the manuscript. 3.6 Author contribution S.L. and B.A.R designed and conducted the research, analyzed and interpreted data, and wrote the article. A.C.O.B conducted biochemical assays, M.R.M assisted in bioinformatics analyses and wrote the article, C.A.K and B.L.M. designed the research, analyzed and interpreted the data, and wrote the article. 87 4 Binding Options for the Small Subunit-Like Domain of Cyanobacteria to Rubisco CHAPTER 4 This chapter contains information submitted for publication in: Rohnke, B. A., Kerfeld, C. A., and Montgomery, B. L. (2019) Binding options for the small subunit-like domain of cyanobacteria to rubisco. Front. Microbiol. Under Review. modified to incorporate the supplemental information into the body of the text, renumber figures, tables, and references to be consistent with the dissertation, and use abbreviations defined in the KEY TO ABBREVIATIONS. 88 4.1 Abstract Two proteins found in cyanobacteria contain a C-terminal domain with homology to RbcS. These SSLDs are important features of CcmM, a protein involved in the biogenesis of carboxysomes found in all β-cyanobacteria, and a rubisco activase homologue (ALC) found in over a third of sequenced cyanobacterial genomes. Interaction with rubisco is crucial to the function of CcmM and is believed to be important to ALC as well. In both cases, the SSLD aggregates rubisco, and this nucleation event may be important in regulating rubisco assembly and activity. Recently, two independent studies supported the conclusion that the SSLD of CcmM binds equatorially to L8S8 holoenzymes of rubisco rather than by displacing an RbcS, as SSLD structural homology would suggest. We use sequence analysis and homology modeling to examine whether the SSLD from the ALC could bind the large subunit of rubisco either via an equatorial interaction or in an RbcS site, if available. We suggest that the SSLD from the ALC of Fremyella diplosiphon could bind either in a vacant RbcS site or equatorially. Our homology modeling takes into account N-terminal residues not represented in available crystal structures that potentially contribute to the interface between RbcL and RbcS. Here, we suggest the perspective that binding site variability as a means of regulation is plausible and that the dynamic interaction between the RbcL, RbcS, and SSLDs may be a means of carboxysome assembly and function. 4.2 Introduction Due to the evolution of a CCM, cyanobacteria are able to significantly contribute to global carbon fixation, despite the comparatively low atmospheric CO2 levels relative to their first appearance on Earth some 3.5 billion years ago177,220. The CCM serves to significantly increase 89 the flux of inorganic carbon into proteinaceous bacterial microcompartments called carboxysomes. Carboxysomes serve to encapsulate rubisco and the shell acts as a semi- permeable barrier to CO2 escape, allowing rubisco to function under high substrate levels46. In the case of β-carboxysomes, which are present in cyanobacteria that express Form IB rubisco, synthesis occurs from the inside-out beginning with condensation of rubisco and carboxysomal protein CcmM into a liquid matrix38,137,213. The structure of CcmM is key to its nucleation of carboxysomal cargo. The C-terminus of CcmM contains 3-5 repeats of a domain that is homologous (around 60-70% similarity) to RbcS— denoted a an SSLD116,156. The SSLD repeats domain-containing portion of CcmM can also be independently transcribed through an internal ribosome entry site. Both CcmM forms—full- length M58 and truncated M35—are necessary for normal carboxysome biogenesis113,114. SSLDs were implicated in the interaction between rubisco and CcmM44,112 and were long hypothesized to bind in place of RbcS in rubisco complexes54,113,160. However, recent structural work on SSLDs demonstrates equatorial binding of CcmM to L8S8 rubisco holoenzymes171,213. This binding appears to be driven largely by electrostatic interactions and affinity for CcmM is not affected even when RbcS binding is partially compromised171. SSLDs also appear as C-terminal domains in rubisco activase homologues (ALC) found in many cyanobacteria222. Recently, the ALC was shown to localize proximal to rubisco in the carboxysome and induce rubisco aggregation, much like M35101. Together, these findings 90 provide strong evidence that the SSLD of ALC binds to rubisco and can induce liquid-liquid phase separation. As recent findings indicate that SSLDs do not displace RbcS in L8S8 rubisco holoenzymes and instead bind equatorially, it is puzzling why there would be conservation of the RbcS-like secondary and tertiary structure that facilitates interactions with RbcL. Some of the conserved residues from RbcS may fill repurposed roles in the SSLD-unique equatorial binding position, thus driving conservation of these features. Others, though, suggest that RbcS displacement may be possible. We decided to systematically homology model the SSLD and RbcL and compare the interfaces in order to evaluate the plausibility of equatorial versus RbcS substitution as a mode of binding171,213. We modeled the SSLD found in the ALC of F. diplosiphon (FdALC SSLD) and analyzed the number of predicted interactions and free energy of solvation when the SSLD binds at the RbcS site (i.e., binds an empty site or displaces RbcS) or binds equatorially. We suggest that while equatorial binding was favored for CcmM in Syn7942 which lacks an ALC homolog, the FdALC SSLD had similar interface features in both positions. We propose that the FdALC could bind either equatorially or in place of RbcS and suggest that the current models of equatorial SSLD may be a part of a larger set of possibilities depending on specific proteins, for example whether or not the cyanobacterium contains an ALC, and perhaps is reflective of the recently uncovered diversity of cyanobacterial RbcL subunits101. 91 4.3 Methods 4.3.1 Protein homology modeling The structures of the F. diplosiphon proteins were generated by homology modeling. For FdALC SSLD, the Swiss Model web server (https://swissmodel.expasy.org)20,216 was used to generate a model for amino acid residues 317 – 424 based on Syn7942 CcmM SSLD1 in the reduced (PDB: 6HBB) form as well as Syn6301 RbcS (PDB: 1RBL, Chain M). Additionally, a homology model of F. diplosiphon L8S8 rubisco was made using Syn6301 rubisco (PDB: 1RBL) as a template. Alignment scores between two sequences were calculated using the LAlign webserver (https://embnet.vital-it.ch/software/LALIGN_form.html) in order to evaluate candidate template structures and to compare primary structure conservation. 4.3.2 MSA of ALC SSLDs The MSA of 141 ALCs from cyanobacteria described in Lechno-Yossef et al.101 was trimmed to remove the ATPase domain, then the remaining regions (linker and SSLD) were re-aligned with a low gap cost at the end of sequences in CLC Sequence Viewer. This allowed for alignment of the SSLD region despite significant variations in the sizes of linkers between species, which were then trimmed to match the SSLD region identified in FdALC (residues 317 – 424, corresponding to residues 1 – 107 of the SSLD). An MSA was also generated for RbcS for each of the 128 organisms that had both a full length SSLD and an annotated RbcS sequence. MSA for RbcS and SSLD were visualized and compared using HMM logos178 generated on Skylign (http://skylign.org/). 92 4.3.3 Analysis of protein-protein interactions Using the homology model for L8S8 F. diplosiphon rubisco, the two FdALC SSLD models were aligned to RbcS1 in PyMol, and structures were generated containing each SSLD replacing RbcS1. Another structure aligned Syn7942 CcmM SSLD1 in complex with rubisco (PDB: 6HBC) to the F. diplosiphon rubisco, and then FdALC SSLD (reduced) was aligned to the CcmM SSLD resulting in a F. diplosiphon rubisco model with FdALC SSLD in the M position (Fig 4.1A). The Syn7942 CcmM SSLD1 structure was also used to replace the RbcS1 position in the Syn6301 rubisco L8S8 structure. Local refinement of structures was performed using Rosetta 3.4. Structures were subjected to the docking prepack protocol followed by the generation of 1000 decoys using the docking protocol in docking local refine mode with the SSLD as the mobile target41,68,212. Based on interface score, the top 200 structures were clustered by pairwise RMSD with a 1 Å cutoff using energy-based clustering in Rosetta 3.477. In all cases, the structure with the lowest interface score belonged to the largest cluster and was selected for use in downstream analysis. These structures, as well as the F. diplosiphon rubisco model and PDB: 6HBC, were analyzed using the Profunc web server (https://www.ebi.ac.uk/thornton-srv/databases/profunc/)100. Interactions involving the RbcS and SSLD were compared for each structure. Further analyses were performed using the Pisa webserver (http://www.ebi.ac.uk/msd-srv/prot_int/cgi- bin/piserver)98 to calculate the solvation free energy gain (ΔG) upon formation of the interfaces for each structure. 93 4.4 Binding at the RbcS1 position Each RbcS in an L8S8 assembly forms four unique protein-protein interfaces, three with the surrounding RbcL subunits and a fourth with a proximal RbcS (Figure 4.1A & 4.1B). The number of interfaces and predicted residue-level interactions were comparable to results from molecular dynamic simulations using C. reinhardtii rubisco117 and crystallographic structures95. Figure 4.1: Structural comparison of RbcS and SSLD. (A) Schematic diagram of the interactions a single RbcS (S1) or SSLD (M) has in a rubisco holoenzyme. Numbering based on van Lun et al.117. (B) Alignment of the SSLD found in ALCs (black is based on an SSLD model, burgundy on an RbcS model) to the RbcS (yellow) or SSLD (teal) binding sites of a L8S8 rubisco holoenzyme. (C) Interaction between RbcL1 (green) and RbcS (yellow) or SSLD (burgundy) in the context of the RubisCO holoenzyme. Labeled residues highlight residues predicted to interact in a salt bridge interaction (Table 4.2), with label color based on the subunit it is from. Numbering for the SSLD is based on the trimmed region beginning at residue 317 in the full length FdALC. Dashed cyan lines show the predicted interacting atoms. (D) HMM-Logo highlighting the areas of sequence conservation between RbcS and the SSLD of ALC across cyanobacteria containing both. A schematic for the secondary structure of the homology model for RbcS from F. diplosiphon is presented above the two logos. Blue squares below each residue depict regions with gaps in significant portions of the MSA. Predicted salt bridges formed by the RbcS subunit are represented by yellow diamonds, while those formed by the SSLD are represented by red arrowheads. Magenta boxes, connected by an arrow, indicate a motif found in both MSA. Yellow diamonds indicate the RbcS residues involved in salt bridge interactions in the F. diplosiphon homology model, while red triangles indicate ALC-SSLD residues involved in salt bridge interactions in the S1 position in the homology model (Table 4.2). 94 RbcS interactions with its nearest RbcL (RbcL1, Figure 4.1A) are substantial, burying ~1600 Å2 of surface area, with five predicted salt bridges, 13 hydrogen bonds, and a free energy of solvation of -5.2 kcal·mol-1 (Table 4.1). The remaining interfaces bury less area and are driven by fewer bond interactions, but favorably contribute to the overall L8S8 rubisco (Table 4.1, Column 5 – FdRbcS). When RbcS is replaced with the SSLD-modeled FdALC SSLD, most salt bridges are lost (Table 4.1, Column 3 – FdALC SSLD; Table 4.2), as are many hydrogen bonds. This is particularly true at the L1-S1 interface, where the absence of the crucial N-terminal loop (residues 3-17) of RbcS in the SSLD accounts for 3 of the 4 missing salt bridges at this interface, as well as the significant reduction of major buried surface area95,171. SSLDs have two features that may play a role in this interaction. First, SSLDs have a poorly conserved flexible linker at their N-terminus that could be involved in non-specific interactions. Additionally, a portion of the N-terminal loop in RbcS involved in L1-S1 interactions is positionally displaced in the primary structures of SSLDs. (Figure 1D, magenta box)116. Notably, the structural position of this region corresponds to a helix in SSLD structures but a loop in RbcS171,213. This ‘displaced motif’ region is conserved and resembles the important lost motif of the N-terminus of RbcS but was not noted in Ryan et al.171 nor Wang et al.213, possibly because without significant backbone rearrangement, this motif is unlikely to be positioned to bind in the same way and its conservation could be attributed to its role in binding at the SSLD equatorial interface. Overall, our modeling with the truncated SynCcmM SSLD template is consistent with the experimental observations that the SSLD structure has a significant loss of favorable binding interactions at the RbcS interface. 95 Table 4.1: Predicted small subunit interactions of rubisco from F. diplosiphon and Syn7942. Results from analyses using Profunc and Pisa webservers of protein-protein interactions in CcmM complexed with rubisco or a homology model of rubisco from F. diplosiphon. Models contained homology models of FdALC SSLD or RbcS from F. diplosiphon or CcmM SSLD1 from Syn7942, as indicated by the row labeled “Interacting Subunit”. These subunits were aligned to the position indicated in row 2 (labels based on Figure 4.1A) in rubisco L8S8 structures from F. diplosiphon (columns 3-5, 8), Syn6301 (column 6), or Syn7942 (column 7). Column 2 shows the rubisco subunit interface with the target SSLD/RbcS. Column 8, in bold, shows crystal structure data from Wang et al.213. Fd RbcS Syn 7942 CcmM SSLD1 Syn 7942 CcmM SSLD1 Fd ALC SSLD Fd ALC SSLD FdALC SSLD (RbcS model) S1 1 0 0 S1 1 1 1 S1 5 2 0 S1 0 0 0 N/A N/A N/A N/A 2 9 4 2 N/A 2 108 26 29 N/A 12 1326.5 495.1 359.4 N/A 240.7 1.3 2.6 -1.5 N/A -1.2 1 13 6 1 N/A 3 194 77 44 N/A 20 1591.5 614.9 467.9 N/A 261.4 -5.2 -1.0 -5.9 N/A -3.2 1 1 1 0 N/A 3 7 7 10 N/A 20 218.5 260.0 232.4 N/A 249.6 -0.6 0.3 -1.0 N/A -3.1 0 2 0 1 N/A 4 17 2 16 N/A 19 380.3 91.0 313.2 N/A 173.5 0.9 1.0 -1.9 N/A -3.1 96 Interacting Subunit Position L1 L2 L3 S1 S2 L1 L2 L3 S1 S2 L1 L2 L3 S1 S2 L1 L2 L3 S1 S2 L1 L2 L3 S1 S2 Number of Salt Bridges Number of Hydrogen Bonds Number of Non- bond Contacts Pisa Interface (Å2) ΔG (kcal· mol-1) M 1 0 1 1 0 1 0 4 1 0 17 0 39 22 0 236.4 0.0 511.3 240.2 0.0 -1.4 N/A 1.1 0.9 N/A M 2 0 2 1 0 4 0 3 2 0 34 0 21 21 0 448.9 0.0 479.4 239.3 0.0 0.7 N/A 3.1 -0.7 N/A Table 4.2: Comparison of predicted salt bridges between RbcL and RbcS or the SSLD. Hypothetical salt bridges in the F. diplosiphon rubisco homology model interacting with either RbcS or FdALC SSLD models at position S1 (see Figure 4.1A). The SSLD from FdALC modeled using either RbcS from Syn6301 as a template, or the CcmM SSLD1 from Syn7942 as the template. Residue numbering corresponds to F. diplosiphon sequences, with the SSLD corresponding to residues 317-424 of FdALC (e.g., residue R94 of the SSLD corresponds to residue R410 of FdALC). Interaction Rubisco:RbcS1 Rubisco:SSLD (RbcS Modeled) RbcL:SSLD (ALC SSLD Modeled) K165:E11 R168:E11 E232:K6 E352:K94 E434:K26 R188:E41 K228:E50 - K6:E44 - - - - - - - D398:R94 D397:R94 - R188:E41 - D77:K91 K6:D44 K6:E81 - - - - - - L1-S1 L2-S1 L3-S1 S2-S1 When the FdALC SSLD is modeled using RbcS as a template, part of the linker at the N- terminus of the SSLD (residues 1 – 17) is included in the model. Analysis of the FdALC SSLD in complex with rubisco suggested the potential for conservation of significantly more interactions (Table 4.1, Column 4 – FdALC SSLD [RbcS model]). Compared to the native RbcS, each interface buries slightly less area (~80% of that observed for RbcS) and is predicted to contain fewer hydrogen bonds and non-bonding contacts. Many salt bridges are potentially maintained or are similar for a total of 5 compared to the 8 found in RbcL-RbcS (Table 4.2). For example, while the SSLD model loses two salt bridges that contribute to the three structural checkpoints of the L1-S interface described in van Lun et al.117 (Figure 4.1C); this FdALC SSLD model is predicted to form a novel salt bridge with L3 and K6 forms an additional salt bridge with S2 instead of L1. Although the SSLD of ALCs shows many regions of relatively low conservation, the regions that are important for RbcS interactions are generally well conserved even in the SSLD (Figure 4.1D), with the notable uncertainty of the displaced N-terminal motif 97 (magenta box) and linker. This suggests that when the flexible linker domain is also taken into account, FdALC SSLD could occupy an empty RbcS site. 4.5 Binding at the equatorial (M) position As reported in Wang et al.213, SynCcmM SSLD1 forms favorable interactions with L8S8 rubisco in an equatorial position (we refer to this as position M). It forms a salt bridge with each rubisco subunit it contacts (L1, L3, & S1) and forms some hydrogen bonds (Table 4.1, Column 7 – Syn7942 CcmM SSLD1). The ΔG at these three interfaces are less favorable overall than those calculated for the four interfaces of FdALC SSLD (Table 1, Column 3) and SynCcmM SSLD1 in the S1 position (Table 1, Column 6 – Syn7942 CcmM SSLD1). However, considering the presence of three salt bridges and that all RbcS positions were occupied in Ryan et al.171, these data are consistent with the model that SSLDs would bind equatorially rather than displace an engaged RbcS subunit. FdALC SSLD also shows potential for interaction with the equatorial position, although the ΔG values calculated from these preliminary models are relatively less favorable. When bound at M, it is predicted to form a greater number of salt bridges and more hydrogen bonds compared to the SynCcmM SSLD1 (Table 1, Column 8 – FdALC SSLD). 4.6 Discussion Here, we present a prediction that the SSLDs found in cyanobacteria may be able to substitute for RbcS in binding RbcL; we propose that this could occur in addition to recently demonstrated equatorial binding171,213. Our analysis considers the SSLDs found in the absolutely conserved 98 carboxysomal protein CcmM and the SSLDs found in the ALC, which is present in a subset of ecophysiologically diverse cyanobacteria. In the case of the FdALC SSLD, we found that its predicted binding with RbcL when substituting for the RbcS may be more favorable than that of the SSLD of CcmM of Syn7942, an organism that lacks the ALC. Although we find that the FdALC SSLD also could engage at the equatorial site, it may be a less favorable interaction than that observed for the SSLD of CcmM171,213. We suggest that the SSLD of FdALC has the potential for both equatorial binding and binding at the RbcS position. For Syn7942, the model organism used by Wang et al.213 and more closely related to Thermosynechococcus elongatus171, which also lacks an ALC, the predicted interface appears to point more favorably towards the equatorial binding found in vitro. Thus, it is possible that both the type of SSLD and the organism could influence whether the SSLD binds exclusively equatorially, especially for interactions with L8S8 rubisco holoenzymes. Additionally, in no case did we find that the SSLD bound better than the native RbcS, supporting the view that SSLDs cannot displace RbcS, though they might bind if sites are available. In numerous quantifications of protein abundance in cyanobacteria, a shortfall of RbcS is found relative to RbcL113,200. This suggest that isoforms other than the L8S8 rubisco holoenzyme may be present in vivo. Indeed, experiments in Syn7942 report 5-6 RbcS for 8 RbcL, suggesting that RbcS binding sites could be available in vivo113,200. Additionally, we suggest it may be possible that the RbcL:RbcS ratio could be dynamically regulated with impacts on both enzyme activity and the binding position of RbcS. Moreover, rubisco undergoes numerous post-translational modifications that regulate its activity and subunit interactions (see review67). Phosphorylation can impact rubisco rate through phosphorylation sites at catalytic residues111, and/or moderate 99 the interactions between RbcL, RbcS, & rubisco activase through partially unknown sites1,70. Phosphorylation reversibly alters the electrostatic interface, potentially affecting the favorability of equatorial driven SSLD interaction. These factors suggest mechanisms by which RbcS and SSLDs could be targets of dynamic regulation. Throughout the evolution of diverse organisms containing Form I rubisco, a RbcL-RbcS fusion has never been observed. Given the importance of rubisco for survival, this is a significant clue to the potential for dynamic regulation of the RbcS:RbcL stoichiometry, potentially by the SSLDs. Though recent observations suggest that SSLDs bind primarily equatorially, we propose a potential for a dynamic relationship between multiple binding locations. Such dynamics could play a large role in the nucleation of carboxysomes, which is fascinating given the observed impact that SSLD-containing proteins have on carboxysome morphology113,170. These features would seem to depend heavily on the availability of RbcS binding locations, the flexibility of the protein structures, redox state, post-translational modifications, the species, the composition of the linker, the type of SSLD, and potentially even the subtype of RbcL101, factors to consider in future investigations on the interaction between rubisco and SSLDs. 4.7 Acknowledgments The authors would like to thank Dr. Sigal Lechno-Yossef for critical discussion of the data and interpretation of results. and Dr. Kaillathe “Pappan” Padmanabhan of the Macromolecular Computing Facility in the Michigan State University’s Department of Biochemistry and Molecular Biology for assistance with Rosetta installation and use of Linux clusters. 100 This work was supported by the U.S. Department of Energy (Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, grant no. DE-FG02-91ER20021 to B.L.M. and C.A.K.). 4.8 Author contribution B.A.R. designed and conducted the research, analyzed and interpreted data, and wrote the article. C.A.K. and B.L.M. designed the research, analyzed and interpreted the data, and wrote the article. 101 5 Linking the Dynamic Response of the Carbon Concentrating Mechanism to Carbon Assimilation CHAPTER 5 Behavior in Fremyella diplosiphon 102 5.1 Abstract Cyanobacteria utilize a CCM that enhances their carbon-fixation efficiency. Previous research has shown that the CCM is regulated by many environmental factors expected to impact photosynthesis. However, efforts to connect these findings to the functional effect on carbon assimilation rates are limited by the aqueous nature of cyanobacteria. Here, we present findings that utilize cyanobacteria in a semi-wet state on glass fiber filtration discs to establish carbon assimilation behavior using gas exchange analysis. In combination with qPCR and TEM analyses, we link the regulation of CCM components to corresponding carbon assimilation behavior in the freshwater, filamentous cyanobacterium F. diplosiphon. Inorganic carbon levels, light quantity, and light quality are all shown to influence carbon assimilation behavior. Results suggest a biphasic model of cyanobacterial carbon fixation. At low levels of CO2, behavior is driven mainly by the cyanobacterium’s Ci-uptake ability, while at higher CO2 levels the carbon assimilation behavior is multi-faceted and depends on Ci availability, carboxysome morphology, linear electron flow, and cell shape. Carbon response curves utilizing gas exchange analysis offer rapid analysis of CO2 assimilation behavior in cyanobacteria. These results provide an initial understanding important for modeling CO2 assimilation in cyanobacteria and insight into how these data correlate to the stoichiometry of CCM components. 5.2 Introduction The robust capability of cyanobacteria to fix carbon through photosynthesis is crucial to their key ecological role as one of Earth’s major primary producers. Thus, understanding the regulation of their carbon fixation mechanisms is of general biological interest and a target of applied research on autotrophic production of biofuels and bioproducts. Cyanobacteria concentrate Ci through a 103 well-established CCM (see review30), which sequesters CO2 and related enzymes and substrates into subcellular, proteinaceous, BMC called carboxysomes (see review210). As the carbon fixation steps of photosynthesis are often regulated to be kept in balance with the overall rate of photosynthesis145, components of the CCM are likely to be tuned to environmental factors that affect photosynthesis as well. Indeed, both carbon transport and carboxysome components are upregulated in conditions where there is a greater need for Ci, such as growth under low CO2 or HL31,199. Further study of the dynamic regulation of the CCM is poised to provide valuable insight into the modular components cyanobacteria utilize to coordinately control such features as the rate of photosynthesis and BMC morphology. Uptake of Ci is the first component of the CCM. Since the cellular membrane is permeable to CO2 but not HCO3 -, cyanobacteria increase the flux of Ci into the cell by using HCO3 - transporters as well as trapping CO2 as HCO3 - through the use of CO2 hydrating enzymes. Constitutively expressed, active carbon transport31,214 involves the low-affinity Na+/ HCO3 - symporter Bic in the cellular membrane157 and the hydration of cytosolic CO2 into HCO3 - by - NDH-14 (including subunits D4/F4/CupB) at the thylakoid membrane184,224, thus driving HCO3 accumulation inside the cell. A parallel set of proteins with higher substrate affinity can be induced to increase Ci-uptake and includes: SbtA, another Na+/ HCO3 - symporter183; BCT1, an ATP-dependent HCO3 - pump144; and NDH-13 (subunits D3/F3/CupA). Together, these complexes provide cyanobacteria with a high, and tunable, capacity for internal Ci influx as HCO3 -, with some measurable leakage of CO2 during Ci-uptake208. 104 The second major aspect of the CCM is comprised of the protein shell of the carboxysome which forms a sub-cellular compartment that is permeable to HCO3 - but not CO2 46. Both rubisco and CA are part of the carboxysomal cargo, which in conjunction with the high concentration of cellular HCO3 -, drives the carboxylation reaction of rubisco forward with high local concentrations of its CO2 substrate. In the case of β-carboxysomes, which are the type of carboxysomes formed in organisms with Type 1B rubisco, the ccmKMNO operon is crucial for carboxysome formation156. β-carboxysome biogenesis begins with rubisco aggregation by CcmM38, a protein which contains an N-terminal, γ-class CA domain and 3-5 C-terminal repeats of an SSLD112,114,116 that can interact with L8S8 rubisco and induce liquid-liquid phase separation171,213. Additionally, CcmM contains an internal ribosome entry site prior to the SSLD tail that results in expression of a truncated form that is crucial to carboxysome formation113,114. CcmN is then recruited to this condensate and, alongside full-length CcmM, interacts with at least CcmK2, the most abundant shell protein38,44,94. In most β-cyanobacteria, an additional β- class CA is recruited as well, though CcmM itself can fill this role in others6,148. The composition of the protein shell includes a diversity of paralogues with a potential range of substrate permeabilities. The faces of the shell are comprised of tessellating hexagonal proteins that contain a charged pore92. These hexagons result from repeats of a pfam00936 domain, either as hexamers of a single-domain monomer (BMC-H) or as trimers of monomers containing two domains in tandem (BMC-T). The BMC-H CcmK2 is present in all sequenced β-cyanobacteria and is usually joined by the highly similar CcmK1192. CcmK3 and CcmK4 always co-occur, have smaller pores, and seem to form heterohexamers and stacked dodecamers, suggesting mechanisms where they could dynamically alter the permeability of carboxysomes193. Other 105 paralogues include CcmK5 which appears to replace CcmK3/4, and CcmK6 which occurs in a heterocyst-forming clade of cyanobacteria. The BMC-T CcmO is essential for carboxysome formation38,159, while the BMC-T CcmP forms stacked hexamers which may account for transport of larger substrates36. Separate from the paralogues forming the faces, CcmL forms pentamers that serve as the vertices of the intact carboxysome structure203,206. The myriad of components that comprise the CCM are responsive to environmental conditions. Both high light and low CO2 levels tend to induce the expression of genes encoding many CCM components, especially for the high-affinity carbon transporters31,76,124. It has also been demonstrated that carboxysome morphology is dynamically responsive to light, Ci, and photosensory activity of cyanobacteriochromes170,199,200. However, many open questions remain in understanding how these changes control the carbon fixation capability of cyanobacteria. There are many available methods for assaying carbon fixation and photosynthesis that have been applied to cyanobacteria. Measuring O2 evolution, which probes linear electron flow at photosystem II (PSII) and shows reductions when CCM is compromised116,154,198, via either mass spectroscopy or with an O2 electrode is perhaps most common. Chlorophyll fluorescence can be used similarly but requires care in cyanobacteria to avoid phycobilisome absorbance or fluorescence confounding the results39. In more elaborate setups, carbon labeling can be used to determine rates of carbon assimilation and flux via either radiometry or mass spectroscopy. Due to the equilibration between CO2 and HCO3 -, both the media and cytosol can have stores of Ci separate from what is fixed, so the setup, lighting regiments, and measurement timing are used to distinguish between stores and assimilation of CO2 and HCO3 - 10,13. In general, these techniques 106 are limited to end-point assays and/or are technically challenging. For terrestrial plants, a robust method derives net gas exchange from a plot of carbon assimilation vs intracellular CO2 to establish steady state photosynthetic parameters non-destructively55. Carbon assimilation vs intracellular CO2 curves are typically modeled with three distinct regions; low intercellular Ci values are limited by the reaction rate of rubisco, higher intercellular Ci values are limited by the rate of ribulose-1,5-bisphosphate regeneration (light limited), and at the highest intercellular Ci values the curves may show a saturation due to maximum utilization of triose phosphate pools115. Due to the aqueous nature of cyanobacteria and the slow, uncatalyzed equilibration with HCO3 -, parallel methods have not been well established but are promising141. In this study, we analyze the carbon fixation characteristics of the filamentous, freshwater cyanobacterium F. diplosiphon. F. diplosiphon exhibits CCA, a process where cells respond to changes in the prevalence of light (primarily red vs. green in F. diplosiphon and many others) through altering the type and abundance of photosynthetic pigments, cell shape, and filament length19,129. Notably, cyanobacteriochrome (phytochrome-related) RcaE acts as a photoreceptor controlling CCA23,89,189,209 and contributes to photoregulation of carboxysome morphology170. In order to probe the roles of CCA, RcaE, and carboxysome regulation in carbon assimilation (A, or the net rate of CO2 uptake per unit area), we demonstrate that A can be measured using cyanobacteria in a semi-wet state with infra-red gas analysis of cyanobacterial discs. A is responsive to light quality and intensity, Ci availability, and the physiological state of cells. Generated carbon response curves (CRCs) establish that a ΔrcaE mutant strain exhibits impaired A under GL that is recovered under HL conditions. While A is only impaired in ΔrcaE relative to WT under GL, O2 evolution is impaired in the ΔrcaE mutant under both GL and RL 107 conditions. Thus, we show that informative dynamic responses can be evaluated using CRCs in cyanobacteria and, together with measurements such as O2 evolution, can be used to infer cellular propensity for Ci-uptake and active utilization in oxygenic photosynthesis. These findings are presented alongside TEM and CCM-related gene expression analyses in order to better understand how the regulation of CCM components contribute to the overall carbon fixation capabilities of cyanobacteria. 5.3 Results 5.3.1 Carbon assimilation measurements in F. diplosiphon respond to light, inorganic carbon availability, and physiological state Glass fiber-filtered F. diplosiphon strains, i.e., F. diplosiphon discs, were analyzed using a LI- COR 6800 Photosynthesis System to detect CO2 consumption. Carbon assimilation rates (A) in WT and ΔrcaE F. diplosiphon strains were responsive to light dosage (Q), showing light saturation at ~100 μmol·m-2·s-1, though this was higher (~150 μmol·m-2·s-1) in initial results with HL acclimated cultures (Figure 5.1A & Figure 5.2B). Thus, 300 μmol·m-2·s-1 was selected to serve as saturating light in further experiments. Under light saturating conditions, strains of F. diplosiphon exhibited a response in A to changing carbon levels in a standard CRC (Figure 5.1C- 5.1F). Blank glass fiber filter discs wetted with fresh cell media were used as a control and show slightly negative A values that become more negative from 600-1000 ppm (Figure 5.2). Typical A values ranged from 1-4 μmol CO2·m-2·s-1; this is in contrast to values of ~15-30 μmol CO2·m-2·s-1 seen in plants55,115. Samples were normalized by OD750 which had a roughly linear relationship with [Chla] that varied somewhat with strain and growth condition (Figure 108 Figure 5.1: Carbon assimilation response to light and Ci availability. (A and B) Response to LI-COR chamber light at 400 ppm CO2s for (A) WT and (B) ΔrcaE F. diplosiphon strains grown at low (12 μmol·m-2·s-1; white), medium (30 μmol·m-2·s-1; gray), and high (100 μmol·m-2·s-1; black) WL intensity in air, n = 3 for ML & LL and n = 1 for HL. (C to F) Response to supplied CO2 at 300 μmol·m-2·s-1 for (C) WT, (D) ΔrcaE, (E) ΔrcaC, and (F) ΔbolA F. diplosiphon strains grown at ~10-12 μmol·m-2·s-1 red (white marks) or green (black marks) light conditions. Error bars represent 95% confidence intervals for n ≥ 5 from 2 independent biological replicates. 109 Figure 5.2: Carbon assimilation response to Ci availability for BG11/HEPES blank. Response to supplied CO2 at 300 μmol·m-2·s-1 for BG11/HEPES filtered through Whatman glass fiber filter paper as a blank for the semi-wet cyanobacteria gas exchange analysis. Error bars represent a 95% confidence interval from 3 replicates. 5.3). The volume of cells used resulted in ~10-55 μg Chla that could be loaded in the gas exchange chamber. This is lower than chlorophyll levels used when assaying leaves, which range from ~120 – 180 μg in Arabidopsis thaliana108,221 (incl. unpublished lab references) and likely contributes to the lower A values in F. diplosiphon than published plant values. Additionally, as the intercellular Ci flux in cyanobacteria is complex and not precisely modeled, response curves are presented with the [CO2] levels in the sample chamber (CO2s) as the independent variable. Lastly, compensation points (roughly the point where A becomes negative) measured in cyanobacterial CRCs appear to fall between 5-25 ppm CO2s, which is likely lower than the typical values (25-100 ppm intercellular CO2) found in higher order plants28,115. This observation is consistent with the presence of a CCM in cyanobacteria. WT F. diplosiphon showed a difference in A only above 700 ppm CO2 when comparing RL and GL acclimated cultures, with GL-grown cultures reaching higher A levels (Figure 5.1C). This result is consistent with previous findings measuring O2 evolution, which show similar rates of 110 Figure 5.3: Chlorophyll a levels versus OD750 for cyanobacteria used in CRC analysis. Representative [Chla] vs. OD750 measured in extracts harvested during CRC runs. Samples include (A) WT and (B) ΔrcaE F. diplosiphon strains grown under red (RL), green (GL), low (LL), medium (ML), or high (HL) white light conditions or in RL- enriched white light under air (Air), Ci upshift (Ci Up), or Ci downshift (Ci down), as described in the materials and methods section. O2 evolution between low intensity RL and GL at ambient CO2 39. By comparison, the ΔrcaE mutant demonstrated hampered A only under GL, with max A dropping from ~4.0 to ~1.3 μmol·m-2·s-1, whereas A in ΔrcaE was statistically indistinguishable from WT under RL (Figure 5.1C & 5.1D). In order to test whether the difference in the net rate of CO2 uptake is correlated with cellular pigmentation, we measured A in a ΔrcaC mutant strain, which lacks RcaC which is a DNA- binding regulatory protein that acts downstream of RcaE. This ΔrcaC strain has a constitutive GL-like CCA phenotype71. CRC analysis indicated no difference in A values for this strain between RL and GL, with values more similar to WT under GL (Figure 5.1E). This finding suggests that the GL physiological state is partially responsible for the higher A values. In addition to pigmentation differences between RL and GL, F. diplosiphon exhibits cell shape differences controlled in part by RcaE23 and its regulation of additional genes including morphogene bolA189. To separate potential impacts of the regulation of pigmentation from the 111 regulation of cell shape, we analyzed CRCs in a ΔbolA mutant strain, which has WT pigmentation but altered cell shape. A values in a ΔbolA mutant showed no difference between RL and GL and were closer to WT under RL (Figure 5.1F). Since the ΔbolA mutant loses the characteristic transition from a spherical cell in RL to a rod-shaped cell in GL, and shows A values more similar to RL, cell shape may also play a role in CRC behavior. 5.3.2 Effect of non-saturating light on carbon assimilation In order to probe for light-limited regions of the CRC in cyanobacteria, we performed CRC analysis under non-saturating light conditions (25 and 50 μmol·m-2·s-1). We found that WT F. diplosiphon that had been grown in low light had near-saturating A values, even at as low as 50 μmol·m-2·s-1 (Figure 5.4A). However, A was severely impaired at 25 μmol·m-2·s-1 above 75 ppm CO2s (Figure 5.4C). The ΔrcaE mutant also showed a decrease in A under non-saturating light conditions and was indistinguishable from WT at 25 μmol·m-2·s-1 (Figure 5.4B & 5.4D). 5.3.3 Effect of different light intensities during growth on carbon assimilation potential Since high light is known to induce the components of CCM31,76,199, we investigated the effect of growth under increasing light intensity on CRC behavior in WT and the ΔrcaE mutant. We utilized a multi-cultivator bioreactor system with green-enriched WL at low (LL; 12 μmol·m-2·s-1), medium (ML; 30 μmol·m-2·s-1) or high (HL; 100 μmol·m-2·s-1) light intensities. Growth rate increased as light intensity increased in both strains (Figure 5.5), although cells typically exhibited chlorosis ~7 d after induction of HL, indicating light stress. CRC analysis in WT indicated that responses to LL and ML were comparable, whereas HL showed a slight decline in A levels at high CO2s (Figure 5.4E). In contrast, we observed increased A levels for 112 Figure 5.4: Carbon assimilation response to Ci availability in response to various light intensities. (A to D) Response to supplied CO2 during runs at 300 μmol·m-2·s-1 (black mark), 50 μmol·m-2·s-1 (gray marks) or 25 μmol·m-2·s-1 (white marks) for (A and C) WT and (B and D) ΔrcaE F. diplosiphon strains grown at low (12 μmol·m-2·s-1) GL-enriched WL. Panels C and D show 0-200 ppm CO2s of panels A and B, respectively. Error bars represent 95% confidence intervals for n ≥ 3 from 2 independent biological replicates. (E to F) Response to supplied CO2 at 300 μmol·m-2·s-1 for (E) WT and (F) ΔrcaE F. diplosiphon strains grown at low (12 μmol·m-2·s-1; white mark), medium (30 μmol·m-2·s-1; gray mark), and high (100 μmol·m-2·s-1; black mark) GL-enriched WL intensities in air. Error bars represent 95% confidence intervals for n ≥ 4 from 2 independent biological replicates. 113 Figure 5.5: Growth rates of F. diplosiphon strains under increasing GL-enriched WL intensity. OD720 values versus time for (A) WT and (B) ΔrcaE F. diplosiphon strains grown under WL with dominant GL wavelengths at low (12 μmol·m-2·s-1; black line), medium (30 μmol·m-2·s-1; purple line) or high (100 μmol·m-2·s-1; blue line) intensity. Shaded area represents ± SD for n ≥ 4 from at least 2 independent biological replicates. the ΔrcaE mutant in response to increased light intensity during growth, with the strain exhibiting near-WT levels under HL (Figure 5.4E & 5.4F). Also under HL acclimation, the two strains exhibited indistinguishable A values under non-saturating light conditions (Figure 5.6).Thus, it appears that WT utilizes light efficiently even under LL and that the ΔrcaE mutant is light starved under LL, with HL exposure resulting in recovery of its low A phenotype under green wavelengths. 5.3.4 Effect of inorganic carbon availability during growth on carbon assimilation We next explored the impact of Ci availability on CRC behavior. Cells were grown in air, Ci upshift (3% CO2), or Ci downshift (3 d growth in 3% CO2 followed by a transfer to air for 19 h) in chambers illuminated with 35-40 μmol·m-2·s-1 WL which induced growth consistent with growth in monochromatic RL and ambient air (Figure 5.1). Under these conditions, WT and 114 Figure 5.6: Carbon assimilation response to Ci availability under non-saturating light conditions after acclimation to HL. Response to supplied CO2 at 300 μmol·m-2·s-1 (black mark) or 25 μmol·m-2·s-1 (white mark) for (A and C) WT and (B and D) ΔrcaE F. diplosiphon strains grown at high (100 μmol·m-2·s-1) WL intensity. Panels C and D show 0-200 ppm CO2s of panels A and B, respectively. Error bars represent 95% confidence intervals for n ≥ 3 from 2 independent biological replicates. ΔrcaE strains exhibited similar A behavior under air (Figure 5.7A & 5.7B). The behavior of these two strains was similar below 200 ppm CO2s in all conditions, and as expected the compensation point appears to decrease as the cultures become more acclimated to lower Ci levels and induce high-affinity CCM systems (Figure 5.7C & 5.7D). During acclimation to Ci downshift, the two strains also performed similarly to each other in runs under non-saturating light conditions (Figure 5.8A & 5.8B); notably, non-saturating light shifted the apparent compensation point higher in both (Figure 5.8C & 5.8D). However, the ΔrcaE mutant strain 115 Figure 5.7: Carbon assimilation response to Ci availability after acclimation to various Ci levels. Response to supplied CO2 at 300 μmol·m-2·s-1for (A and C) WT and (B and D) ΔrcaE F. diplosiphon strains grown at medium (~35 μmol·m-2·s-1) RL-enriched WL in air with 3% CO2 enrichment (Ci Up; black mark), without enrichment (Air; gray mark), or under Ci downshift (Ci Down; white mark). Panels C and D show 0-200 ppm CO2s of panels A and B, respectively. Error bars represent 95% confidence intervals for n ≥ 4 from 2 independent biological replicates. exhibited a deficiency in A under Ci upshift and a less robust response to Ci downshift at higher CO2s levels relative to WT (Figure 5.7B). Thus, the ΔrcaE mutant strain appears to have a deficiency in responding to Ci levels. 116 Figure 5.8: Carbon assimilation response to Ci availability in non-saturating light after acclimation to Ci downshift. Response to supplied CO2 at 300 μmol·m-2·s-1 (black marks) or 25 μmol·m-2·s-1 (white marks) for (A and C) WT and (B and D) ΔrcaE F. diplosiphon strains grown at medium (~35 μmol·m-2·s-1) RL-enriched WL under Ci downshift. Panels C and D show 0-200 ppm CO2s of panels A and B, respectively. Error bars represent 95% confidence intervals for n ≥ 3 from 2 independent biological replicates. 5.3.5 Rates of O2 evolution in F. diplosiphon strains under red and green light To compare our findings to established methods and to compare CO2 uptake to active Ci utilization in oxygenic photosynthesis, we analyzed O2 evolution in WT and ΔrcaE strains acclimated to RL or GL (Figure 5.9, white bars). We found that WT produced O2 at higher initial rates in GL compared to cells grown under RL. O2 evolution was significantly decreased in ΔrcaE under both RL and GL. Whereas CRC analysis only uncovered a defect under GL conditions, the ΔrcaE stain showed reduced O2 evolution rates even when acclimated to RL. 117 Figure 5.9: Oxygen evolution of F. diplosiphon strains acclimated to red or green light. O2 levels measured after illumination by 250 μmol·m-2·s-1 WL in F. diplosiphon strains (WT or ΔrcaE) grown under RL or GL, with or without the addition of 1 mM of the electron acceptor DCBQ. Error bars represent 95% confidence intervals for n = 5 (-DCBQ) or n = 3 (+DCBQ) from 2 independent biological replicates. Additional experimentation utilized 1mM 2,6-dichloro-p-benzoquinone (DCBQ), which accepts electrons from PSII and acts as a photosynthetic uncoupler, in order to test for the total number of PSII centers capable of water oxidation66,133. In WT, we saw a substantial reduction in O2 evolution proportional to the rates seen without DCBQ (Figure 5.9, grey bars). In contrast, addition of 0.5mM DCBQ in Syn6803 increased O2 rates substantially133. The fact that the rates did not increase suggests that even without the addition of DCBQ, WT F. diplosiphon utilizes the majority of its PSII complexes that have sufficient excitement to split water (i.e., downstream regulation is not limiting WT). Furthermore, the decrease seen in A rates under RL (Figure 1C) may be attributable to PSII reaction rates as WT under RL exhibited lower O2 evolution rates with and without DCBQ compared to cells in GL. Lastly, with the addition of DCBQ the ΔrcaE mutant showed no difference from WT in either light condition, due to a lack of response in RL and an increase in O2 evolution in GL-acclimated ΔrcaE cultures (Figure 5.9). This finding 118 suggests that the apparent loss of photosynthetic rate in ΔrcaE under GL (as measured by both A and O2 evolution rates) is not due to a deficiency in PSII complex rates. 5.3.6 TEM analysis of carboxysome morphology in response to light conditions and carbon availability To contextualize the CRC behaviors, we analyzed carboxysome morphology in the studied conditions (Figure 5.10). We previously reported that under both RL and GL, carboxysomes were more abundant (higher number per cell section) and smaller in the ΔrcaE mutant relative to WT170. Additionally, carboxysome diameter decreased in both strains under GL and exhibited no light-quality-dependent changes in carboxysome abundance in either strain. We found that neither the ΔrcaC nor ΔbolA strains showed a difference in carboxysome size or shape between RL and GL (Figure 5.11A and 5.11B). These strains had significantly larger and fewer carboxysomes than in WT under GL, where WT exhibited a decrease in carboxysome diameter and trends towards higher carboxysome abundance. Under increasing light intensity, WT showed a gradual increase in carboxysome diameter that was significant when comparing HL to LL (p = 0.024, Figure 5.11C), with no increase in carboxysome abundance (Figure 5.11D). The ΔrcaE mutant showed a similar increasing trend in carboxysome diameter with HL acclimating cultures showing a significant increase (p < 0.001 when comparing HL to either ML or LL, Figure 5.11C). In contrast to WT, the ΔrcaE mutant exhibited substantial increases in carboxysome number in response to increasing light. The ΔrcaE mutant did not exhibit its characteristic increase in carboxysome abundance until it was acclimated to ML or HL in these WL growth conditions relative to WT (Figure 5.11D). 119 Figure 5.10: TEM analysis of cellular morphology of F. diplosiphon strains under changing light or Ci availability. Representative images of (A) WT, ΔrcaE, ΔrcaC, & ΔbolA strains under RL & GL, and WT & ΔrcaE strains under (B) increasing WL intensity or (C) decreasing CO2 availability. Bars, 0.5 μm; C, carboxysomes; PL, photosynthetic lamellae. 120 Figure 5.11: Carboxysome morphology under diverse physiological conditions. Boxplots displaying the full range of measurements of maximum carboxysome diameter and number of carboxysomes per cell section from TEM analysis for (A and B) WT, ΔrcaC, & ΔbolA strains of F. diplosiphon grown under RL and GL, and WT & ΔrcaE strains grown under increasing (C and D) WL intensity or (E and F) decreasing CO2 availability. Lowercase letters indicate statistically significant groups (p < 0.05) within a panel. Corresponding average ± SE and sample size are presented in Table 5.1. Data for WT under RL and GL are reproduced here from Rohnke et al.170 under the terms of the Creative Commons Attribution 4.0 International license, and data for WT under Air and Ci upshift are reproduced from Lechno-Yossef et al.101 with permission. 121 Table 5.1: Quantification of average carboxysome sizes and numbers per cell section in Figure 5.11. Conditiona Strain Carboxysome size (nm)b No. of carboxysomes/ cell section WT ΔrcaEc ΔrcaC ΔbolA WT ΔrcaE ΔrcaC ΔbolA WT ΔrcaE WT ΔrcaE WT ΔrcaE WT ΔrcaE WT ΔrcaE WT ΔrcaE 340 ± 19 224 ± 12* 323 ± 27 345 ± 15 227 ± 19# 174 ± 5*,# 325 ± 26* 336 ± 18* 318 ± 26 155 ± 9* 354 ± 23 166 ± 7* 404 ± 25# 236 ± 11*,# 436 ± 19 171 ± 7* 362 ± 15# 244 ± 10*,# 332 ± 27# 211 ± 14*,# RL GL LL ML HL Ci Upshift Air Ci Downshift 3.0 ± 0.4 6.2 ± 0.6* 1.9 ± 0.3* 2.5 ± 0.3 3.8 ± 0.3 7.2 ± 0.9* 2.0 ± 0.3* 2.6 ± 0.3* 1.9 ± 0.3 1.9 ± 0.4 1.8 ± 0.3 3.5 ± 0.6*,# 1.6 ± 0.3 5.2 ± 0.6*,# 1.4 ± 0.1 3.7 ± 0.4* 2.1 ± 0.2# 4.5 ± 0.5* 1.3 ± 0.2 3.4 ± 0.5* Sample size (n) for carboxysome size measurements Sections used (n) for no. of carboxysome/ cell section measurements 27 43 24 28 45 106 18 31 26 34 21 61 19 66 42 95 66 134 22 52 30 30 30 30 30 30 30 30 30 30 30 30 30 30 60 60 60 60 30 30 a Indicates conditions under which F. diplosiphon cells are grown as described in Methods and materials, i.e., RL, red light at ~10 to 12 µmol m-2 s-1; GL, green light at ~10 to 12 µmol m-2 s-1; LL, low GL-enriched WL at 12 µmol m-2 s-1; ML, medium GL-enriched WL at 30 µmol m-2 s-1; HL, high GL-enriched WL at 100 µmol m-2 s-1; Ci Upshift, air enriched with 3% CO2; Air, ambient air; Ci Downshift, growth under air enriched with 3% CO2 followed by a shift to ambient air for ~19 h. b Numbers for carboxysome size and carboxysome/cell section are represented as average ± SE. c Statistical analyses, p < 0.05 indicated as follows: *, mutant vs. WT in same condition; #, significant difference vs. reference condition (RL, LL, or Ci Upshift) for same strain. Ci availability also impacted carboxysome morphology. The WT strain exhibited the characteristic decrease in carboxysome abundance under Ci upshift (Figure 5.11F), while also showing an increase in carboxysome diameter (Figure 5.11E) (same data as reported in Lechno- Yossef et al.101). The Ci downshift conditions (transfer to air for ~19 h following growth in 122 elevated Ci) did not allow sufficient time for complete carboxysome acclimation, which is on the order of 2-4 d in Syn7942199. While the WT strain under Ci downshift showed similar carboxysome abundance to Ci upshift conditions, it had decreased size (p = 0.003), which could be part of the transition to the air-acclimated state (Figure 5.11E & 5.11F). The ΔrcaE mutant overall presented a misregulated response to Ci availability, showing a decrease in carboxysome diameter in response to Ci upshift (contrasted to the increase seen in WT, Figure 5.11E) and no significant response in carboxysome abundance (Figure 5.11F). 5.3.7 Transcriptional regulation of carbon concentrating mechanism components measured by qPCR analysis Multiple components of the CCM are expected to be controlled at the transcriptional level in response to light and Ci availability31,76,130,170,200. For the studied strains and growth conditions, we analyzed the transcriptome with probes for CCM components (Table 5.2) using qPCR analysis. These analyses included probes for the carboxysome-related genes in the ccmK1K2LMNO and ccmK3K4 operons, ccmK6, ccmP, rbcL or rbcS (rubisco subunits), ccaA1/2 (carboxysomal CA), and alc. Genes related to Ci-uptake were also probed, including low-Ci induced cmpA (BCT complex), sbtA, and ndhD4 (NDH-I4 complex), constitutively expressed ndhD3 (NDH-I3 complex) and bicA, and a LysR-type transcriptional regulator with homology to cmpR143 and ccmR214, the latter two of which are both involved in the transcriptional response to Ci availability. In an analysis of strains under RL and GL conditions (Table 5.3), ΔrcaE showed upregulation of ccmM and downregulation of rbcS under RL, whereas more significant changes were observed 123 Table 5.2: Primers used for qPCR probes in chapter 5. Target Gene ccmK1 ccmK2 ccmK3 ccmK4 ccmK6 ccmL ccmM ccmN ccmO ccmP ccaA1 ccaA2 alc rbcL rbcS LysR-Type cmpA sbtA ndhD3 ndhD4 bicA orf10B Forward Primer 5'-3' AACGAATTGGCAGGACATACT Reverse Primer 5'-3' GCAGGCGTAGAATCTGTGAA AGGCTTGCACTTCCGATAC TGCTGATGCGATGGTGAA TGCTGCTGGAGAACAAGTAAA GTAAAGTGGATCGGAAGGATGG CAGGCAGTTGGAGCATTAGA TCAGAAACATCGCCACGAATA GAAGCAGTAGGACGAGTGAATG ATTGGCGCTGCGATGAA GTCTACTCCTGCACCTACGATA GTCTTCGAGGTGTGAAACTACTG GCAACAAGCTGACCGTTTAC CTATCTGCAACGCACAAATATCC TGGCACTCAGATTTATGGTACAG GTCCGAGATGGGTTCATTTAGAG CCATTACCTCCAAGCTCAGTAAA CTCCTACCATCGCTGGAAATC TCATTCTAGCTCTCAAGGAGAAAC CTAGAAACAACCCGAGGCTTTA GCTCAAGTATACAGAGGCAACC GAGTCAGTACATTCTCCGCAATAA AACGAGCAGTTCGATTACCC ATGCGCTCCCATTGTTCT CCGGCAACTATTCCTACCTTATC TCGTGACAGGCAACGATTT GTTAGAAGGTGAGCGTGGTATC GAAGCCCAGTCTTGGGTAAA TGTTCGGCGCTAAATCTACTC GCTTGATGTTGTCAAAGCCTAC TCGGTCGGATTGCCTTTATTT GCCGACAAGTAGCAAACAATTC CTGCATTAACCGCAGAGATTTG GAGTATTGCTTTGGTGGCTTTG GTGGAACTGCGATCCGTAAT ATGTATAGCGGGCGATGAATAC TTCTCAGCGTTTCCCATCTC CAGGTACGGTTGAGAAGAATCA TGACTGCCGTGTACTTCTTAATC GTAGGCGATCGCTCCAATATAC GTTGCGGTTTGTACCGAATATG TGTGGCTGTAAACCTGTGAG AGAACTACAGCGTCAGCTTAAT CTGCTTCGCTTTCAGCATTT under GL, particularly the downregulation of ccmK3, rbcL, rbcS and the low-Ci induced Ci- uptake genes relative to WT. The regulation of ccmM, rbcL, and rbcS were consistent with prior results170, as were the downregulation of sbtA and ndhD3 130. WT showed few differences between RL and GL conditions; alc, bicA, and cmpA were downregulated under GL. For many genes, ΔrcaE also exhibited downregulation under GL but with the magnitudes of change more extreme and more frequently statistically significant. The ΔrcaC mutant showed almost no difference between RL and GL, except a failure to downregulate alc under GL. Lastly, the ΔbolA mutant exhibited downregulation of ccmK2, ccmK3, ccmK4, and sbtA under RL. 124 Table 5.3: Relative expression of CCM genes in red versus green light conditions in F. diplosiphon strains. qPCR expression data of WT, ΔrcaE, ΔrcaC, and ΔbolA F. diplosiphon strains grown in RL or GL (~10-12 μmol·m-2·s-1) conditions. n ccmK1a RL GL WT 5 ΔrcaEb ΔrcaC ΔbolA 5 6 6 WT 5 ΔrcaE ΔrcaC ΔbolA 5 6 6 6.1 ± 0.4 6.7 ± 0.8 5.9 ± 0.5 5.7 ± 0.5 5.9 ± 0.4 6.0 ± 0.4 6.3 ± 0.5 6.2 ± 0.1 ccmK2 6.1 ± 0.4 6.8 ± 0.8 6.0 ± 0.4 5.5 ± 0.5* 5.8 ± 0.4 6.0 ± 0.2 6.2 ± 0.5 5.9 ± 0.2 ccmK3 5.1 ± 0.3 5.6 ± 0.5 5.0 ± 0.4 4.5 ± 0.3* 5.0 ± 0.2 4.4 ± 0.2*,# 5.1 ± 0.6 4.9 ± 0.2# ccmK4 5.3 ± 0.3 5.5 ± 0.6 5.3 ± 0.4 4.7 ± 0.3* 5.0 ± 0.4 4.8 ± 0.4 5.6 ± 0.7 5.1 ± 0.3# ccmK6 -0.5 ± 0.3 -0.3 ± 0.4 -0.4 ± 0.4 -0.1 ± 0.6 -0.7 ± 0.3 -1.0 ± 0.4# -0.2 ± 0.6 -0.4 ± 0.1 ccmL 4.9 ± 0.6 5.5 ± 0.7 4.8 ± 0.4 4.3 ± 0.6 4.6 ± 0.3 4.4 ± 0.3# 4.9 ± 0.4 4.7 ± 0.2 ccmM 5.1 ± 0.5 6.3 ± 0.1* 5.2 ± 0.5 4.6 ± 0.6 5.2 ± 0.2 5.0 ± 0.3# 5.4 ± 0.5 5.0 ± 0.2 ccmN 3.8 ± 0.8 4.1 ± 0.5 3.4 ± 0.4 3.1 ± 0.6 3.5 ± 0.3 3.2 ± 0.2# 3.7 ± 0.3 3.5 ± 0.2 ccmO 3.8 ± 1.3 2.8 ± 0.7 3.5 ± 0.6 3.3 ± 0.8 3.5 ± 0.6 2.8 ± 0.2 3.6 ± 0.4 3.7 ± 0.1 ccmP 0.1 ± 0.5 0.5 ± 0.3 0.2 ± 0.2 0.2 ± 0.4 -0.2 ± 0.4 -0.4 ± 0.3# 0.2 ± 0.4 0.2 ± 0.1 ccaA1 -0.1 ± 0.7 -0.3 ± 0.7 0.5 ± 0.6 1.2 ± 1.2 -0.1 ± 0.2 -0.1 ± 0.4 0.0 ± 0.2 -0.1 ± 0.3# ccaA2 -1.2 ± 0.7 -1.4 ± 0.6 -0.7 ± 0.3 0.1 ± 1.1* -1.3 ± 0.4 -1.2 ± 0.5 -1.2 ± 0.3# -1.0 ± 0.3 alc 3.0 ± 0.2 3.0 ± 0.6 2.7 ± 0.4 3.3 ± 0.5 2.2 ± 0.2# 1.7 ± 0.4# 2.6 ± 0.2* 2.9 ± 0.1* rbcL 6.7 ± 0.9 6.0 ± 0.5 6.6 ± 0.7 5.5 ± 0.6* 6.4 ± 0.3 5.3 ± 0.3*,# 6.6 ± 0.5 6.0 ± 0.3 rbcS LysR- Type cmpA 7.1 ± 0.7 4.5 ± 0.1* 6.4 ± 0.6 5.8 ± 0.4* 6.8 ± 0.1 4.7 ± 0.2* 6.8 ± 0.3 6.2 ± 0.3* 1.6 ± 0.5 1.3 ± 0.5 1.6 ± 0.4 1.4 ± 0.3 1.3 ± 0.4 1.5 ± 0.3 1.7 ± 0.3 1.6 ± 0.1 1.8 ± 1.5 0.3 ± 0.5 0.0 ± 0.9 0.5 ± 0.6 -1.4 ± 0.4# -2.3 ± 0.5*,# -1.0 ± 0.6 -1.2 ± 0.3# sbtA 3.9 ± 0.7 4.3 ± 0.5 4.1 ± 1.1 2.8 ± 0.6* 4.5 ± 0.7 2.5 ± 0.4*,# 4.3 ± 0.6 4.0 ± 0.5# ndhD3 3.9 ± 0.5 3.9 ± 0.8 4.0 ± 0.6 3.5 ± 0.2 4.1 ± 0.2 2.7 ± 0.4*,# 4.3 ± 0.4 4.3 ± 0.4# ndhD4 2.7 ± 0.3 3.2 ± 0.6 2.5 ± 0.4 2.2 ± 0.5 2.6 ± 0.2 2.6 ± 0.4 2.6 ± 0.3 2.6 ± 0.3 0.4 ± 0.3 bicA 0.3 ± 0.3 a Data for each gene presented as -ΔCq ± SD relative to endogenous control gene orf10B, thus represent a log2 scale. b Statistical analyses, p < 0.05 indicated as follows: *, mutant vs. WT in same condition; #, GL vs. RL in same strain. 0.5 ± 0.6 0.4 ± 0.4 0.8 ± 0.6 0.0 ± 0.2# 0.1 ± 0.3 0.3 ± 0.3 Under increasing light intensity (Table 5.4), WT experienced significant upregulation of select HCO3 - transporters (likely due to an increased linear electron flow), ccmN, and ccmO, alongside a downregulation of rbcS (possibly from degradation following high light stress). The ΔrcaE 125 Table 5.4: Relative expression of CCM genes under increasing light intensity. qPCR expression data of WT and ΔrcaE F. diplosiphon strains grown at LL (12 μmol·m-2·s-1), ML (30 μmol·m-2·s-1), and HL (100 μmol·m-2·s-1) GL- enriched WL intensity. LL ML HL WT 4 n ΔrcaEb 4 WT 3 ΔrcaE 3 WT 5 ΔrcaE 5 ccmK1a 6.7 ± 0.4 6.9 ± 0.8 7.1 ± 0.3 8.2 ± 0.3*,# 6.1 ± 0.8 7.1 ± 0.9 ccmK2 6.6 ± 0.6 6.9 ± 0.8 6.8 ± 0.2 7.7 ± 0.4* 5.9 ± 0.8 6.9 ± 0.9 ccmK3 5.6 ± 0.7 5.2 ± 0.7 5.7 ± 0.4 5.7 ± 0.4 4.7 ± 0.5 5.2 ± 0.5 ccmK4 5.5 ± 0.2 5.3 ± 0.5 5.8 ± 0.2 5.9 ± 0.4 5.3 ± 0.5 5.7 ± 0.7 ccmK6 -0.5 ± 0.2 -0.8 ± 0.4 -0.5 ± 0.2 -0.4 ± 0.2 -0.4 ± 0.4 0.1 ± 0.3# ccmL 5.4 ± 0.5 5.3 ± 0.7 6.0 ± 0.5 6.6 ± 0.1# 4.8 ± 0.7 5.7 ± 0.8 ccmM 5.7 ± 0.4 5.7 ± 0.8 6.4 ± 1.1 7.2 ± 0.2# 5.1 ± 0.5 6.2 ± 0.5* ccmN ccmO ccmP 3.7 ± 0.2 3.9 ± 0.7 3.9 ± 0.3 4.7 ± 0.4 6.9 ± 1.1# 5.3 ± 0.8*,# 3.5 ± 0.2 3.3 ± 0.3 3.9 ± 0.1# 3.7 ± 0.2 7.8 ± 1.1# 6.4 ± 1.1# 0.4 ± 0.2 0.3 ± 0.3 0.6 ± 0.1 0.7 ± 0.2# 0.1 ± 0.5 0.5 ± 0.4 ccaA1 0.3 ± 0.6 0.0 ± 0.7 0.0 ± 0.5 -0.3 ± 0.7 0.3 ± 1.2 0.1 ± 0.5 ccaA2 -1.0 ± 0.5 -1.4 ± 0.4 -1.0 ± 0.1 -1.2 ± 0.5 -0.4 ± 1.5 -0.6 ± 0.8 alc rbcL rbcS LysR- Type cmpA sbtA 2.6 ± 0.5 2.2 ± 0.7 3.5 ± 0.4 3.5 ± 0.3# 2.8 ± 0.8 3.3 ± 0.5# 7.6 ± 0.7 6.4 ± 1.0 8.5 ± 1.7 8.0 ± 1.2 7.1 ± 1.1 7.0 ± 1.0 7.9 ± 0.2 5.0 ± 0.3* 8.0 ± 0.9 5.4 ± 0.1* 6.6 ± 0.4# 4.5 ± 1.4* 2.0 ± 0.4 1.8 ± 0.6 2.4 ± 0.4 2.5 ± 0.4 4.1 ± 0.2# 3.1 ± 0.4*,# -1.6 ± 0.5 -1.8 ± 0.8 -0.6 ± 0.3# -0.4 ± 0.5# 5.8 ± 0.3# 4.3 ± 1.1*,# 4.5 ± 0.2 3.5 ± 0.8 5.1 ± 0.6 4.3 ± 0.5 6.2 ± 0.2# 5.9 ± 0.7# ndhD3 4.0 ± 0.6 3.3 ± 1.0 5.0 ± 0.5 4.6 ± 0.3 4.5 ± 0.3 5.0 ± 1.1 ndhD4 2.6 ± 0.6 2.9 ± 0.8 3.3 ± 0.3 3.7 ± 0.4 2.7 ± 0.2 3.2 ± 0.8 bicA -0.6 ± 0.8 -0.2 ± 0.7 0.5 ± 0.2 0.3 ± 0.7 0.7 ± 0.4# 1.6 ± 0.6*,# a Data for each gene presented as -ΔCq ± SD relative to endogenous control gene orf10B, thus represent a log2 scale. b Statistical analyses, p < 0.05 indicated as follows: *, ΔrcaE vs. WT in same condition; #, significant difference vs. LL in same strain. mutant showed the characteristic downregulation of rbcS seen in other conditions. Additionally, it exhibited an upregulation of ccmK1 and ccmK2 under ML and ccmK6 under HL that correlates with the increase in carboxysome abundance (Figure 5.10B, Figure 5.11D). ΔrcaE showed a similar upregulation of HCO3 - transporters, ccmN, and ccmO, though not quite to the same extent 126 as WT. Lastly, ΔrcaE showed significant upregulation of alc in contrast to non-significant increases seen in WT. Since this protein is important for response to Ci limitation101, this upregulation could be meaningful. Both WT and ΔrcaE strains demonstrated significant differential expression of CCM components under decreasing Ci availability (Table 5.5). WT showed a general downregulation in shell proteins, rbcL, rbcS, and ccmM under Ci downshift, consistent with findings in Syn680348,214 and Syn7942179. It is interesting to consider how these data correlate with increased carboxysome abundance under Ci downshift reported in the literature48,127,199 and this study (Figure 5.11E & 5.11F; Ci upshift vs. Air). As previously reported, alc is downregulated under Ci downshift and was reported to be involved in the decreased carboxysome abundance under Ci upshift101. Consistent with expectations, WT also exhibited significant upregulation of the low-Ci induced Ci-uptake genes. Additionally, as in HL, ccmN and ccmO were upregulated alongside the low-Ci genes rather than consistent with their operon. This high divergence of ccmN and ccmO is not seen in HL nor Ci upshift conditions in Syn7942200 nor Syn680376,214; in these strains the two genes do not diverge from the rest of the ccm operon. Moreover, ccmO is even located in a satellite operon in Syn6803 yet does not show this same pattern. While WT upregulated the low-Ci induced Ci-uptake genes in both Air and Ci downshift, ΔrcaE only did so under Ci downshift. However, ccm gene transcription in the ΔrcaE strain was similar to WT under both Ci upshift and downshift overall, with the major differences being when the two strains transition from one to the other. Notably, ΔrcaE also recovered near-WT levels of rbcS under Ci downshift, possibly explaining the strain’s recovery of A seen in this condition. 127 Table 5.5. Relative expression of CCM genes under decreasing carbon availability. qPCR expression data of WT and ΔrcaE F. diplosiphon strains grown under Ci Upshift (3% CO2), Air, or Ci Downshift (19 h after a transfer from 3% CO2 to air). Ci Upshift Air Ci Downshift WT 5 n ΔrcaEb 4 WT 5 WT 5 ΔrcaEb 4 WT 5 ccmK1a 6.9 ± 0.6 7.7 ± 0.4 6.1 ± 0.3# 6.7 ± 0.6# 6.2 ± 0.3 6.8 ± 0.5# ccmK2 7.0 ± 0.5 7.6 ± 0.3 6.0 ± 0.3# 6.5 ± 0.5# 5.8 ± 0.3# 6.4 ± 0.5# ccmK3 5.4 ± 0.3 5.2 ± 0.5 4.2 ± 0.3# 4.9 ± 0.1* 4.1 ± 0.2# 4.2 ± 0.5# ccmK4 5.8 ± 0.3 5.6 ± 0.5 4.8 ± 0.2# 5.3 ± 0.2* 4.7 ± 0.2# 4.6 ± 0.4# ccmK6 0.3 ± 0.3 -0.1 ± 0.3 -0.4 ± 0.4# 0.0 ± 0.3 -0.7 ± 0.5# -0.4 ± 0.4 ccmL 5.7 ± 0.3 6.4 ± 0.4* 4.9 ± 0.2# 5.4 ± 0.5# 4.8 ± 0.2# 5.3 ± 0.4# ccmM 6.5 ± 0.2 7.1 ± 0.5 5.1 ± 0.2# 5.9 ± 0.7# 4.7 ± 0.1# 5.3 ± 0.6# ccmN ccmO ccmP 3.7 ± 0.3 4.8 ± 0.2* 5.4 ± 0.5# 3.8 ± 0.8* 7.4 ± 0.5# 7.2 ± 0.8# 2.8 ± 0.4 3.9 ± 0.3* 6.2 ± 0.5# 2.7 ± 0.8*,# 8.0 ± 0.4# 7.8 ± 0.8# 0.8 ± 0.3 1.0 ± 0.5 -0.1 ± 0.5# 0.6 ± 0.3* -0.9 ± 0.3# -0.7 ± 0.7# ccaA1 -1.1 ± 0.2 -0.8 ± 0.2 -0.5 ± 0.4# -0.3 ± 1.1 -0.9 ± 0.5 -0.4 ± 0.5 ccaA2 -1.9 ± 0.4 -1.8 ± 0.2 -1.4 ± 0.5 -1.3 ± 0.9 -2.2 ± 0.4 -1.7 ± 0.7 alc rbcL rbcS LysR- Type cmpA sbtA 3.8 ± 0.5 3.2 ± 0.6 2.5 ± 0.2# 3.3 ± 0.8 1.7 ± 0.3# 2.1 ± 0.6# 8.3 ± 0.3 7.4 ± 0.3* 7.1 ± 0.6# 5.6 ± 0.8*,# 7.5 ± 0.4# 7.5 ± 0.5 8.3 ± 0.5 5.6 ± 0.5* 7.5 ± 0.5# 4.2 ± 0.4*,# 7.6 ± 0.4 6.7 ± 0.3*,# 2.1 ± 0.5 2.7 ± 0.2* 3.4 ± 0.2# 1.9 ± 0.7* 3.8 ± 0.3# 3.4 ± 0.2# -4.3 ± 0.4 -3.3 ± 0.4* 6.2 ± 0.2# -1.9 ± 0.3*,# 6.6 ± 0.5# 6.7 ± 0.3# 0.2 ± 0.3 0.2 ± 0.6 4.5 ± 0.3# 1.8 ± 0.2*,# 5.1 ± 0.6# 5.3 ± 0.3# ndhD3 3.8 ± 0.3 3.8 ± 0.6 4.7 ± 0.3# 3.1 ± 1.1 5.4 ± 0.5# 6.1 ± 0.4*,# ndhD4 3.0 ± 0.2 3.8 ± 0.3* 2.3 ± 0.2# 2.9 ± 0.8 2.6 ± 0.3# 2.9 ± 0.6 bicA 0.7 ± 0.2 0.7 ± 0.3 0.8 ± 0.3 1.4 ± 0.2*,# 0.5 ± 0.2 1.3 ± 0.3*,# a Data for each gene presented as -ΔCq ± SD relative to endogenous control gene orf10B, thus represent a log2 scale. b Statistical analyses, p < 0.05 indicated as follows: *, ΔrcaE vs. WT in same condition; #, significant difference vs. Ci Upshift in same strain. 5.4 Discussion 5.4.1 Use of the carbon response curve in cyanobacteria The CCM found in cyanobacteria has multiple modular components that can respond to dynamic environmental conditions and impact photosynthetic capacity. This capability is critical to 128 organismal fitness in the diverse habitats that photo-autotrophic cyanobacteria can grow. Many biological responses such as the upregulation of Ci-uptake genes under low Ci conditions are well established, and there has been an effort to cohesively model how the complex photosynthetic parameters of cyanobacteria arise from regulation of the CCM121,141,200. These efforts are generally limited to simple model cyanobacteria, and often serve poorly to rapidly measure net Ci consumption due to the aqueous nature of these organisms. Our work with F. diplosiphon, a freshwater filamentous cyanobacterium which undergoes CCA in response to light quality, highlights multilayered connections between CCM components, nutrient availability, and the physiological state of the cell170. Efficiently connecting these factors to overall carbon assimilation was a crucial step toward understanding how these organisms (and humans, as bio- prospectors) can optimize photosynthesis. Utilization of gas exchange analysis to construct CRCs in cyanobacteria suggests that acclimation to dominant light quality through CCA has a nuanced impact on overall A behavior. WT F. diplosiphon cells assimilate more CO2 when acclimated to GL despite having smaller carboxysomes and not being tuned to the red-enriched light of the LI-COR system (Figure 5.1C). The disruption of CCA through the loss of the major photoreceptor RcaE added additional layers of complexity; since RcaE influences the stoichiometry of carboxysome components and carboxysome size under both RL and GL170, we expected a general decrease in net A. Instead, we found a GL-specific impairment (Figure 5.1D). While the small, more numerous carboxysomes of the ΔrcaE strain may contribute to overall A behavior, this observation cannot explain the higher A of WT under GL. These intriguing initial results prompted further exploration of the A behavior of cyanobacteria. 129 We provide evidence that physiologically relevant CRCs, similar in nature to the popular carbon assimilation vs intracellular CO2 curves in plants, can be obtained from cyanobacteria in a semi- wet state using cyanobacterial discs. Cells show a dosage response to both light (Figure 5.1A) and CO2, two major responses relevant to the development of advanced modeling of photosynthetic parameters in plants115. CRCs were also sensitive enough to show changes in apparent compensation point based on the physiological state of the cell (Figure 5.7C). Traditional O2 evolution experiments show similar trends, with WT having higher rates under GL than RL while ΔrcaE exhibited higher rates under RL compared to GL (Figure 5.9). Despite this, the two methods differed when comparing WT and ΔrcaE strains under RL; ΔrcaE exhibited a decrease in O2 evolution but not A rates, suggesting an impairment in utilization of CO2 for oxygenic photosynthesis even though the two strains had similar CO2 uptake rates under RL. Thus, CRCs of cyanobacterial discs offer novel insight into the CO2 uptake behavior in cyanobacteria under a broad range of Ci levels. This method also significantly reduces the time required for equilibration between CO2 and HCO3 - which allows for dynamic responses to be studied. Thus, it is a promising technique that can both stand alone as a quick measurement of net carbon assimilation, and in conjunction with established systems that more deeply probe the HCO3 -/CO2 flux. In particular it serves to more directly test CO2 utilization by cyanobacteria in contrast to well established procedures that test a cyanobacteria’s utilization of HCO3 -. 5.4.2 The low Ci phase of the carbon response curve (<100 ppm CO2s) is Ci-uptake driven Similar to the carbon assimilation vs intracellular CO2 curve in higher-order plants, the CRC in cyanobacteria appears to have regions with different contributing factors. Though the different 130 regions do not yet appear to be as clear cut or as amiable to modeling as in plants, we analyze which factors appear to contribute to observed CRC behaviors. The presence of a distinct region at low ppm CO2s is supported by the CRC behavior under non-saturating light (Figure 5.4A- 5.4D). At very low light levels, A is mostly lost and instead the cyanobacteria appear to respire. However, there still seems to be a robust carbon assimilating behavior present upon reaching low Ci levels. This is likely due the low-Ci induced uptake system and reflects the high affinity cyanobacteria have for even trace amounts of Ci. Notably, this region is very robust and rarely exhibits differences; the ΔrcaE mutant is always indistinguishable from WT below 100 ppm CO2s. There are only two conditions in which we observed changes to this region. The slope and compensation point were very responsive to a culture’s acclimation to different Ci availabilities with growth under Ci downshift prompting a robust A response even at minuscule Ci levels and a lowered apparent compensation point (Figure 5.7C & 5.7D). We were tempted to identify this as a ‘light-independent’ region, thus tested a hypothesis predicting that cultures acclimated to Ci downshift would not show a change in slope below ~100 ppm CO2s even when analyzed under non-saturating light. This was not the case, however; non-saturating light reduced the A slope and increased the compensation point (Figure 5.8C & 5.8D). This observation suggests that light availability can affect this region, albeit only in specific conditions still related to Ci-uptake capacity. Thus, we propose identifying the low Ci region of the cyanobacterial CRC as a Ci- uptake driven region. 131 5.4.3 The high Ci phase of the carbon response curve (>100 ppm CO2s) is responsive to multiple photosynthetic parameters The identification of a Ci-uptake driven region at low Ci, separate from a region that reaches Amax at high Ci, suggests a biphasic CRC behavior reminiscent of those seen in C4 plants85,150. However, in contrast to C4 plants, which are mainly limited by phosphoenolpyruvate regeneration rate at high Ci, our results suggest that the high Ci region of cyanobacteria depends on many variables, including Ci availability, carboxysome morphology, linear electron flow, and cell shape. Components of the CCM relating to Ci-uptake appear to have a broad effect on A behavior; upregulation of the low-Ci induced genes (Table 5.5) is correlated with an increase in A at all CO2s levels (Figure 5.7A). Since this increase occurs in Ci downshift conditions where WT carboxysomes have not had sufficient time to acclimate to Air conditions (Figure 5.11E & 5.11F), this is one case where we can neatly attribute a change in A behavior directly to one major component of CCM. However, under HL, we see a similar induction of the low-Ci induced genes (Table 5.4) without the corresponding increase in A (Figure 5.4E). In the same vein, the increase in A under GL in WT (Figure 5.1C) is the opposite of what we would expect given the downregulation of cmpA seen under GL (Table 5.3). Analysis of the ΔrcaE mutant strain provides some additional lines of inquiry that may offer insight. Unlike WT, increasing light intensity increased the mutant’s Amax (Figure 5.4F). One possible explanation is that the mutant experiences a greater increase in overall carboxysome volume in response to HL (Figure 5.11C & 5.11D), perhaps evidencing a role of carboxysomes 132 in A behavior. It is also worth considering that the ΔrcaE mutant is likely less efficient at light utilization, as evidenced by the fact that addition of DCBQ as an electron sink increased the mutant’s O2 evolution rate in GL but not the WT’s (Figure 5.9). Thus, the HL condition would be a greater boon to the mutant (as evidenced by its increase in A), but is likely stressful for the more efficient WT. This suggests that one or both of these factors (carboxysome size and linear electron flow) contribute to the determination of Amax. Secondly, the ΔrcaE mutant’s behavior yields insights into the A phenotype of WT under GL. Though cmpA is downregulated under GL in WL, the ΔrcaE mutant shows much more significant downregulation of low-Ci induced genes (Table 5.3), which perhaps contributes towards the low A phenotype of ΔrcaE under GL. If this were the case, it would suggest that inducible Ci-uptake systems do indeed contribute, but are being masked in the high A phenotype of WT under GL. Both ΔrcaC and ΔbolA mutants show few differences between RL and GL in the experiments performed in this study. Under both RL and GL, ΔrcaC that is constitutively in GL-like phenotypic state showed nearly identical A behaviors that were more similar to WT GL (Figure 5.1E), suggesting that GL acclimation may also contribute to WT’s high A phenotype. Additionally, even though RcaC is downstream of RcaE, its loss did not seem to affect ccm gene expression while RcaE has major impacts (Table 5.3). As for the ΔbolA strain, it too showed nearly identical A behavior in both RL and GL but was instead more similar to WT under RL (Figure 5.1F). Since the ΔbolA mutant has WT-like pigmentation, this seems to rule out the pigmentation component of GL acclimation as a driving force for the high A phenotype. However, the ΔbolA mutant does have an enlarged, spherical cell shape under both RL and GL, so it is possible that the rod shape of WT F. diplosiphon cells under GL enhances Ci-uptake. As a 133 last point of consideration on the topic, WT has smaller, more numerous carboxysomes under GL than either the ΔrcaC or ΔbolA mutants, though it is difficult to conceive how the carboxysome data correlates with a high A phenotype in both WT GL and ΔrcaC under RL and GL. Thus, we would conclude that Amax is a multifaceted region of the CRC in cyanobacteria that depends on Ci-uptake capabilities but is also impacted by at least one non-pigmentation factor related to GL acclimation such as higher PSII rates under GL (Figure 5.9), overall rate of linear electron flow, or cell shape. Additional factors may include temperature or metabolomics; it would be interesting to investigate both the flux through the Calvin-Benson cycle and triose- phosphate utilization in RL versus GL. 5.4.4 Regulation of the ccm operon in F. diplosiphon Our findings that differential gene expression patterns of ccmN and ccmO are more similar to low-Ci induced genes than the rest of the ccm operon hold interesting implications. Though ccmO is only present in this operon in ~60% of cyanobacteria192, this is still strong conservation, it is present in the ccm operon in F. diplosiphon, and ccmN is always conserved in this operon. Thus, it is noteworthy that we have identified an organism where these two genes are not co- transcribed with the rest of the operon. We have identified a LysR-type regulator immediately upstream of the ccm operon that is roughly co-transcribed with low-Ci inducible genes (Table 5.4 and Table 5.5). The predicted protein, fdiDRAFT81170, is the closest BLAST match for two notable low-Ci-induction related LysR-type regulators found in Syn6803: cmpR, a low-Ci induced regulator of cmp143 and ccmR214. CmpR is particularly interesting, as it is found 134 upstream of cmp in Syn6803 and there is no LysR-type regulator near the ccm operon. As F. diplosiphon is missing a LysR-type regulator near cmp, it is possible that fdiDRAFT81170 fills a similar role. While we have not found a promoter region near ccmN or ccmO, it is possible that the co-regulation of these genes and the high affinity carbon transporters is due to the LysR-type regulator’s novel location and associated function. 5.4.5 Impact Through this study, we have integrated physiological analysis of the cyanobacterium F. diplosiphon with the novel application of gas exchange analysis to cyanobacteria. Like many cyanobacteria, F. diplosiphon performs CCA, which offers a useful system for studying the impact of light regulation, especially as it relates to photosynthesis. We have thoroughly explored the connection between the loss of RcaE, a cyanobacteriochrome that controls the CCA pathway, and the CCM. Analysis of CRCs provides a simple method to assay the carbon assimilation phenotype of cyanobacteria, connecting findings on how the stoichiometry of CCM components impact the structure and function of carboxysomes and Ci-uptake systems. Lastly, preliminary work to identify photosynthetic parameters identifiable through CRCs could contribute valuable insight into modeling and understanding the dynamic regulation of photosynthesis in cyanobacteria. 5.5 Materials and methods 5.5.1 Growth conditions General culture inoculation and growth under RL and GL were performed as described in Rohnke et al.170. In brief, we utilized a short-filament strain of F. diplosiphon with wild-type 135 pigmentation identified as SF33 (WT)42, an RcaE-deficient mutant strain (ΔrcaE) characterized by Kehoe and Grossman89, an RcaC-deficient mutant strain (ΔrcaC) identified in our lab through a forward genetics screen, and a BolA-deficient mutant strain (ΔbolA) described in Singh and Montgomery189. Liquid cultures were inoculated from plated cultures and grown at 28°C under WL in BG-11/HEPES until they were diluted to an initial OD750 of 0.05 and transferred to experimental conditions. The effect of light intensity was tested in a Multi-Cultivator MC 1000-OD system (Photon Systems Instruments, Drasov, Czech Republic) equipped with LED WL and autonomous monitoring of OD680 and OD720 according to the manufacturer’s directions. Since the LED WL was GL dominant, starter cultures grown under GL were used for experiments involving the multi-cultivator to avoid WT showing a growth lag as it underwent CCA. Light conditions were set at a constant value of 12 μmol·m-2·s-1 (LL), 30 μmol·m-2·s-1 (ML), or 100 μmol·m-2·s-1 (HL). Since sustained HL conditions ultimately caused chlorosis, when high ODs were needed for harvesting for TEM and RNA extraction, ML and HL cultures were first grown at 12 μmol·m-2·s-1 for 1-2 d prior to the onset of ML and HL conditions. Cultures grown this way were allowed to acclimate to the higher light intensity for at least 3 d prior to harvesting. All experiments involving HL grown cultures were harvested prior (within 6 d of HL onset) to the plateauing of OD that preceded substantial cell death. The effect of carbon availability was tested in Multitron growth chambers (Infors HT, Bottmingen-Basel, Switzerland) at 30°C under WL (~35-40 μmol·m-2·s-1, RL-enriched) gassed with air either enriched with 3% CO2 (Ci upshift) or unenriched (Air). To achieve Ci downshift, 136 we shifted cultures from Ci upshift to Air conditions after 3 d of growth and resuspension in BG11/HEPES lacking sodium bicarbonate, as described in Lechno-Yossef et al.101 based on Wang et al.214. Cells were harvested for CRC, TEM, or qPCR analysis ~19 h after transfer to air (Ci downshift). 5.5.2 Carbon response curve analysis using F. diplosiphon discs OD750 was measured in triplicate for cultures growing under the desired experimental conditions and were harvested between ODs of 0.6-1.2. A total volume equal to 11.8 absorbance units (V = 11.8/OD750) was vacuum filtered through glass fiber filters with a pore size sufficiently small enough to capture >99% of F. diplosiphon cells (Whatman GF/A 47cm diameter, Sigma-Aldrich, St. Louis, MO), with a second layer of Whatman grade 1 filter paper to diffuse the filtrate more evenly. Disc diameter was selected to minimize extra surface area not necessary for the gas exchange chamber; about 47% of the disc surface area was exposed to the 6cm2 chamber and barely extended past the gaskets. Cyanobacterial discs were handled carefully with forceps, briefly dabbed on filter paper to remove excess wetness, kept on BG11/HEPES agar plates, and swiftly analyzed to minimize environmental perturbance. CO2 levels were measured with infra- red gas analysis in a LI-COR Photosynthesis System 6800 (LI-COR, Lincoln, NE), with one end of a strip of damp Whatman grade 1 filter paper placed underneath the disc as a wick. The other end was submerged in ddH2O to maintain disc dampness for the duration of the experiment, which was found to greatly increase the duration that steady state could be maintained to ~45 min (data not shown). 137 The chamber was illuminated by the standard “Sun+Sky” (RL-dominant) regime with a leaf temperature of 28°C, flow rate of 500 µmol s⁻¹, and source air with 12 ppm H2O. For the standard CRC, the initial CO2 supplied was 1000 ppm and the sample was allowed to equilibrate for at least 5m and until a steady state had been maintained for at least 3m.The CRC followed a gradient of 1000, 850, 700, 550, 400, 300, 200, 150, 100, 75, 50, 25, and 5 ppm followed by a return to 400 ppm with automatic infra-red gas analysis matching. The sample was allowed to equilibrate for ~2-3 min at each time point for a total run time of ~25 min after initial equilibration. 5.5.3 O2 evolution analysis O2 evolution was measured using an Oxytrace+ O2 electrode (Hansatech Instruments Ldt, Norfolk, England) illuminated by an acrylic projector bulb. Illumination was maintained at ~250 μmol·m-2·s-1 measured with a LI-250 Light meter (LI-COR) equipped with a quantum sensor (model US-SQS/L, Heinz Walz CmbH, Effeltrich, Germany). Cells containing ~10 μg Chla (based on OD750 extinction coefficients [Figure 5.3]) were harvested, washed 2x in 3mL BG11/HEPES lacking sodium bicarbonate, and ultimately resuspended in 1mL BG11/HEPES lacking sodium bicarbonate. Cyanobacteria were placed in the chamber and spiked with 1M sodium bicarbonate (Sigma-Aldrich) to a final concentration of 2mM prior to illumination. If applicable, 2,6-dichloro-p-benzoquinone (DCBQ; Sigma-Aldrich) was then added to a final concentration of 1mM. Cells were allowed to equilibrate at ambient light for ~1.5m then illuminated. The O2 evolution Vmax was recorded as the peak rate reached within 10m of illumination. 138 5.5.4 TEM analysis TEM analysis was performed according to the methods of Rohnke et al.170 for all experimental conditions. For Ci upshift and air conditions, 60 cell sections were randomly selected and analyzed for carboxysome number in WT and ΔrcaE with carboxysome diameters measured in 20 of these sections. In all other strains and conditions, 30 cell sections were analyzed with 10 analyzed for carboxysome diameter as well. Samples were prepared from at least two independent biological replicates. As a modification to the original method, some samples were analyzed using a JEM 1400 Flash TEM (JEOL USA Inc., Peabody, MA), still at an operating voltage of 100V. 5.5.5 qPCR analysis The abundance of ccmK1, ccmK2, ccmK3, ccmK4, ccmK6, ccmL, ccmM, ccmN, ccmO, ccmP, ccaA1, ccaA2, alc, rbcL, rbcS, fdiDRAFT81170 (a LysR-type transcriptional regulator), cmpA, sbtA, ndhD3, ndhD4, and bicA transcripts were measured in relation to the internal control orf10B in total RNA extracts from F. diplosiphon strains under various experimental conditions as described previously101,170 in accordance with MIQE guidelines 32. In brief, this involved harvesting ~20 mL of exponentially growing cells upon reaching the target OD750 (~0.5-0.6), handling the samples on ice and flash freezing the cell pellet within 1 h of harvesting, extracting with Trizol reagent incubated at 95°C followed by wash steps, DNase treatment (TURBO DNA- free kit, Invitrogen, Madison, WI), and RNA quantification using a NanoDrop ND-1000 Spectrophotometer. Reverse transcription was performed using qScript cDNA SuperMix (Quantabio, Beverly, MA) kit and qPCR was prepared using Fast SYBR-Green master mix (Applied Biosystems, Foster City, CA) in 384-well plates (Applied biosystems) with a 10 μL 139 reaction volume, both according to the manufacturer’s directions. Probe sequences are provided in Table 5.2. RNA quality was assayed using gel electrophoresis and genomic contamination was controlled for by verifying that no-template-control samples had Cq values greater than 5 cycles higher than the respective unknowns. Data reflect three technical replicates for each of at least three independent biological and are presented using the delta Cq method (ΔCq) in order to foster analysis between several strains and conditions. 5.5.6 Chlorophyll extraction Chla was measured spectrophotometrically according to the methods of de Marsac and Houmard122 as described for analysis in F. diplosiphon187. Samples were harvested in parallel with CRC analysis as a secondary validation of normalization by OD750 and at least three independent biological replicates were analyzed. 5.5.7 Statistical Analysis Experiments were performed with n ≥ 3 from at least two biological replicates for all experiments. 5.6 Acknowledgements We are grateful to Dr. David T. Hanson of the Department of Biology at The University of New Mexico for innovating the utilization of filtered liquid cultures in photosynthetic gas exchange analysis and for his kind guidance during our troubleshooting of the method. We are also grateful to Dr. Thomas D. Sharkey and Dr. Berkley Walker of the Michigan State University (MSU) Plant Biology Laboratory for providing LI-COR 6800 availability and detailed discussion of 140 methodology and results. This study is indebted to the hard work of Kiara Rodriguez of the University of Puerto Rico, who we were fortunate enough to host for a productive Research Experience for Undergraduates opportunity. Her contributions appear in Figure 5.1A-5.1D and Figure 5.4. We are also thankful to Alicia Withrow of the MSU Center for Advanced Microscopy for her extensive assistance with the transmission electron microscope and providing a diamond knife to use for this study. This work was supported by the U.S. Department of Energy (Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, grant no. DE-FG02-91ER20021 to B.L.M.). We also thank Melissa Whitaker (supported by National Science Foundation grant no. MCB-1243983 to B.L.M.) for strain maintenance and culture production. 141 CHAPTER 6 6 Conclusions and Perspectives 142 The work described in this dissertation has centered on the characterization of the regulation of the cyanobacterial CCM in response to dynamic environmental conditions expected to influence photosynthetic rate. I successfully characterized the tuning of the CCM and related carbon fixation potential to Ci and light availability in F. diplosiphon, which are trends also found in other cyanobacteria. I also have made significant contributions to understanding the regulatory roles of CCA and the rubisco activase homologue, ALC, in the CCM. These results support the view that the CCM of cyanobacteria can be finely tuned as a part of the regulation of photosynthesis. This regulation impacts the carbon assimilation behavior of F. diplosiphon, highlighting the functional impacts of regulating Ci-uptake and carboxysome morphology. RcaE, a cyanobacteriochrome that controls the RL vs. GL response of CCA in F. diplosiphon, has an important impact on CCM components and assimilation behavior. Its loss led to a complicated change in the photosynthetic behavior of F. diplosiphon, including changes in pigmentation and reductions to O2 evolution rates (Figure 5.9), demonstrating an impaired photosynthetic efficiency of a ΔrcaE mutant. This loss also resulted in a perturbed stoichiometry of CCM components related to the impaired photosynthesis and resulted in a distinct misregulation of carboxysome abundance and morphology. Carboxysomes were smaller and more numerous in a ΔrcaE mutant strain (Figure 2.2). Interestingly, this effect was not correlated with the function of known components of CCA regulation downstream of RcaE, with ΔrcaC and ΔrcaF mutants showing no changes to carboxysome morphology under RL, limited changes under GL (Figure 2.6, Figure 5.10A, Figure 5.11E & 5.11F), and only a few changes in ccm gene regulation in RL and GL (Figure 2.7, Table 5.3). These findings suggested that RcaE works through distinct effectors, or through effects on the physiological state of the cell, to regulate 143 carboxysome morphology by altering the ratio of carboxysome shell components to cargo components (Figure 2.3, Figure 2.4, Table 5.3). These changes were correlated with a decrease in carbon assimilation potential under GL (Figure 5.1C & 5.1D), showing a functional link between the detection of light quality, tuning of the CCM, and carbon assimilation potential. Our increased understanding of the role of RcaE supports the rising view that regulation of the ratio of shell components to cargo components is a major driving factor of carboxysome dynamics. Larger ratios of shell:core generally lead to smaller and/or more numerous carboxysomes, and vice versa113,200, and this includes an important consideration of the two forms of CcmM, with M35 (containing only SSLDs) being part of the core components while M58 recruits carboxysome shell components114 (Figure 2.4). This understanding of the importance of carboxysome component stoichiometry on structure and abundance provides a general understanding that the expression levels of cargo intended for incorporation into a BMC must be controlled and offers a mechanism with which to regulate BMC morphology. This research also provided evidence that ALC, the rubisco activase homologue found in cyanobacteria, should be considered as an important part of the carboxysomal cargo. In addition to being localized to carboxysomes (Figure 3.6), expression of the alc gene was found to correlate with carboxysome morphology across multiple conditions. alc was upregulated in response to Ci upshift (Figure 3.10), conditions under which carboxysomes are larger and less numerous (Figure 3.11), which is consistent with alc participating in the determination of the carboxysomal shell to cargo ratio as part of the cargo. An Δalc mutant strain of F. diplosiphon lost the response to Ci upshift (Figure 3.11), implying a causal effect. In support of this view, alc 144 expression was correlated with carboxysome abundance under GL; WT under GL had smaller, more numerous carboxysomes than either ΔrcaC or ΔbolA mutant strains (Figure 5.11B, Table 5.1), with an upregulation of alc in both mutants relative to WT (Table 5.3). Compared to RL, WT under GL had smaller carboxysomes (Figure 5.11A, Table 5.1) and the downregulation of alc in GL was the only differential expression of the carboxysome-related genes apparent. Thus, while the function of ALC remains undetermined (Figure 3.7F), it appears to have an interesting role as carboxysomal cargo interacting with rubisco, potentially mediating rubisco network formation (Figure 3.7D, Figure 3.9) alongside CcmM-35171,213, with potential impacts on rubisco packing density. In addition to expression level, ALC’s interaction with rubisco appeared to be ATP-dependent in F. diplosiphon (Figure 3.7E), suggesting that ALC could contribute to regulation of carboxysome morphology in response to multiple factors. These findings support further investigations into the regulation of carboxysome morphology in response to the cellular energy status. The misregulation of PPB in the ΔrcaE mutant (Figure 2.10) in parallel with the misregulation of carboxysomes implicates a potential functional connection between phosphate availability and carboxysomes. PPB abundance is inversely correlated with ATP levels, with PPB diameter increasing during dark periods in Syn7942 under light/dark cycles181 when ATP levels are low25,78. Assessing ATP levels in WT F. diplosiphon under RL vs GL and contrasting these with levels in the ΔrcaE null mutant would test whether the smaller, more numerous PPB in ΔrcaE and WT under GL reflect changes in the ATP status of cells, which could draw further correlations to CCM regulation. If so, the role of ATP in determining carboxysome morphology could then be tested by experimentally reducing ATP levels through the use of energy transfer inhibitors25. 145 Regulatory mechanisms controlling carboxysome morphology must also be considered alongside overall cell morphology. Recently, it was shown that carboxysome positioning can be driven by the intercellular oscillations patterns of McdA and McdB concentrations, which are also dependent on cell shape118. In light of these findings, it seems possible that localization of both PPB and carboxysomes are interrelated, potentially by this positioning system. Moreover, the impact of CCA’s regulation of cell shape is thus a crucial point of consideration. Misregulation of PL also appears to impact carboxysome positioning, seen especially in the frequently mislocalized carboxysomes of the ΔbolA spherical mutant of F. diplosiphon (Figure 2.8 & Figure 5.10A). The functional importance of this is substantial, as cell shape was implicated in the determination of A in WT under GL (Figure 5.1C) and ΔbolA (Figure 5.1F). Our understanding of A behavior in cyanobacteria has been expanded through the use of semi- wet F. diplosiphon discs. This technique allowed us to make CRC measurements comparable to the informative carbon assimilation vs intracellular CO2 curves used in higher-order plants. Two major phases of CRCs were identified in F. diplosiphon: a Ci-uptake driven region and an upper region that includes Amax and is dependent on Ci-uptake, PSII rates, overall rate of linear electron flow, cell shape, and/or carboxysome morphology. These data highlight multiple responses by the CCM to environmental factors such as light intensity, light wavelength, and Ci availability. These responses were shown to influence A behavior, elucidating a functional role for fine-tuned CCM regulation on organismal fitness in cyanobacteria. 146 Ultimately, my studies draw strong correlation between the detection of changes in the external environment and tuning of cellular capacity to produce sustaining energy through photosynthesis. The multiple points of regulation of CCM, the investment in a regulatory system to drive fine-tuning, and the direct impact of these on organismal fitness provide strong evidence of the importance of regulating multiple points of photosynthesis to thrive in dynamic environments. 147 BIBLIOGRAPHY 148 BIBLIOGRAPHY (1) Aggarwal, K. K., Saluja, D., and Sachar, R. C. (1993) Phosphorylation of rubisco in Cicer rietinum: Non-phosphoprotein nature of rubisco in Nicotiana tabacun. Phytochemistry 34, 329– 335. doi: 10.1016/0031-9422(93)80004-C (2) Aigner, H., Wilson, R. H., Bracher, A., Calisse, L., Bhat, J. Y., Hartl, F. U., and Hayer-Hartl, M. (2017) Plant RuBisCo assembly in E. coli with five chloroplast chaperones including BSD2. Science 358, 1272–1278. doi: 10.1126/science.aap9221 (3) Aldea, M., Garrido, T., Hernández-Chico, C., Vicente, M., and Kushner, S. R. (1989) Induction of a growth-phase-dependent promoter triggers transcription of bolA, an Escherichia coli morphogene. EMBO J. 8, 3923–3931. (4) Alvey, R. M., Bezy, R. P., Frankenberg‐Dinkel, N., and Kehoe, D. M. (2007) A light regulated OmpR-class promoter element co-ordinates light-harvesting protein and chromophore biosynthetic enzyme gene expression. Mol. Microbiol. 64, 319–332. doi: 10.1111/j.1365- 2958.2007.05656.x (5) Andrews, T. J., and Abel, K. M. (1981) Kinetics and subunit interactions of ribulose bisphosphate carboxylase-oxygenase from the cyanobacterium, Synechococcus sp. J. Biol. Chem. 256, 8445–8451. (6) de Araujo, C., Arefeen, D., Tadesse, Y., Long, B. M., Price, G. D., Rowlett, R. S., Kimber, M. S., and Espie, G. S. (2014) Identification and characterization of a carboxysomal γ-carbonic anhydrase from the cyanobacterium Nostoc sp. PCC 7120. Photosynth. Res. doi: 10.1007/s11120-014-0018-4 (7) Aryal, U. K., Stöckel, J., Krovvidi, R. K., Gritsenko, M. A., Monroe, M. E., Moore, R. J., Koppenaal, D. W., Smith, R. D., Pakrasi, H. B., and Jacobs, J. M. (2011) Dynamic proteomic profiling of a unicellular cyanobacterium Cyanothece ATCC51142 across light-dark diurnal cycles. BMC Syst. Biol. 5, 194. doi: 10.1186/1752-0509-5-194 (8) Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (2014) Current Protocols in Molecular Biology. Greene Publishing and Wiley Interscience. (9) Badger, M. R. (1980) Kinetic properties of ribulose 1,5-bisphosphate carboxylase/oxygenase from Anabaena variabilis. Arch. Biochem. Biophys. 201, 247–254. doi: 10.1016/0003- 9861(80)90509-3 (10) Badger, M. R., and Andrews, T. J. (1982) Photosynthesis and inorganic carbon usage by the marine cyanobacterium, Synechococcus sp. Plant Physiol. 70, 517–523. (11) Badger, M. R., Andrews, T. J., Whitney, S. M., Ludwig, M., Yellowlees, D. C., Leggat, W., and Price, G. D. (1998) The diversity and coevolution of Rubisco, plastids, pyrenoids, and 149 chloroplast-based CO2-concentrating mechanisms in algae. Can. J. Bot. 76, 1052–1071. doi: 10.1139/b98-074 (12) Badger, M. R., Hanson, D., and Price, G. D. (2002) Evolution and diversity of CO2 concentrating mechanisms in cyanobacteria. Funct. Plant Biol. 29, 161–173. doi: 10.1071/pp01213 − fluxes (13) Badger, M. R., Palmqvist, K., and Yu, J.-W. (1994) Measurement of CO2 and HCO3 in cyanobacteria and microalgae during steady-state photosynthesis. Physiol. Plant. 90, 529–536. doi: 10.1111/j.1399-3054.1994.tb08811.x (14) Badger, M. R., and Price, G. (1992) The CO2 concentrating mechanism in cyanobacteria and microalgae. Physiol. Plant. 84, 606–615. doi: 10.1034/j.1399-3054.1992.840416.x (15) Ballaré, C. L., Scopel, A. L., and Sanchez, R. A. (1991) Photocontrol of stem elongation in plant neighbourhoods: effects of photon fluence rate under natural conditions of radiation. Plant Cell Environ. 14, 57–65. doi: 10.1111/j.1365-3040.1991.tb01371.x (16) Bao, H., Melnicki, M. R., Pawlowski, E. G., Sutter, M., Agostoni, M., Lechno-Yossef, S., Cai, F., Montgomery, B. L., and Kerfeld, C. A. (2017) Additional families of orange carotenoid proteins in the photoprotective system of cyanobacteria. Nat. Plants 3, 17089. doi: 10.1038/nplants.2017.89 (17) Barta, C., Carmo-Silva, A. E., and Salvucci, M. E. (2011) Rubisco activase activity assays, in Photosynthesis Research Protocols (Carpentier, R., Ed.), pp 375–382. Humana Press, Totowa, NJ. doi: 10.1007/978-1-60761-925-3_29 (18) Bassham, J. A., Krohne, S., and Lendzian, K. (1978) In vivo control mechanism of the carboxylation reaction, in Photosynthetic Carbon Assimilation (Siegelman, H. W., and Hind, G., Eds.), pp 77–93. Springer US, Boston, MA. doi: 10.1007/978-1-4684-8106-8_6 (19) Bennett, A., and Bogorad, L. (1973) Complementary chromatic adaptation in a filamentous blue-green alga. J. Cell Biol. 58, 419–435. doi: 10.1083/jcb.58.2.419 (20) Bertoni, M., Kiefer, F., Biasini, M., Bordoli, L., and Schwede, T. (2017) Modeling protein quaternary structure of homo- and hetero-oligomers beyond binary interactions by homology. Sci. Rep. 7. doi: 10.1038/s41598-017-09654-8 (21) Bezy, R. P., and Kehoe, D. M. (2010) Functional characterization of a cyanobacterial OmpR/PhoB class transcription factor binding site controlling light color responses. J. Bacteriol. 192, 5923–5933. doi: 10.1128/JB.00602-10 (22) Bhat, J. Y., Miličić, G., Thieulin-Pardo, G., Bracher, A., Maxwell, A., Ciniawsky, S., Mueller-Cajar, O., Engen, J. R., Hartl, F. U., Wendler, P., and Hayer-Hartl, M. (2017) Mechanism of enzyme repair by the AAA+ chaperone rubisco activase. Mol. Cell 67, 744- 756.e6. doi: 10.1016/j.molcel.2017.07.004 150 (23) Bordowitz, J. R., and Montgomery, B. L. (2008) Photoregulation of cellular morphology during complementary chromatic adaptation requires sensor-kinase-class protein RcaE in Fremyella diplosiphon. J. Bacteriol. 190, 4069–4074. doi: 10.1128/JB.00018-08 (24) Bordowitz, J. R., Whitaker, M. J., and Montgomery, B. L. (2010) Independence and interdependence of the photoregulation of pigmentation and development in Fremyella diplosiphon. Commun. Integr. Biol. 3, 151–153. (25) Bornefeld, T., and Simonis, W. (1974) Effects of light, temperature, pH, and inhibitors on the ATP level of the blue-green alga Anacystic nidulans. Planta 115, 309–318. doi: 10.1007/BF00388613 (26) Bottomley, P. J., and Stewart, W. D. P. (1976) ATP pools and transients in the blue-green alga, Anabaena cylindrica. Arch. Microbiol. 108, 249–258. doi: 10.1007/BF00454849 (27) Bracher, A., Whitney, S. M., Hartl, F. U., and Hayer-Hartl, M. (2017) Biogenesis and metabolic maintenance of rubisco. Annu. Rev. Plant Biol. 68, 29–60. doi: 10.1146/annurev- arplant-043015-111633 (28) Brooks, A., and Farquhar, G. D. (1985) Effect of temperature on the CO2 / O2 specificity of ribulose-1,5-bisphosphate carboxylase/oxygenase and the rate of respiration in the light. Planta 165, 397–406. doi: 10.1007/BF00392238 (29) Brown, M. R. W., and Kornberg, A. (2008) The long and short of it – polyphosphate, PPK and bacterial survival. Trends Biochem. Sci. 33, 284–290. doi: 10.1016/j.tibs.2008.04.005 (30) Burnap, R. L., Hagemann, M., and Kaplan, A. (2015) Regulation of CO2 concentrating mechanism in cyanobacteria. Life 5, 348–371. doi: 10.3390/life5010348 (31) Burnap, R. L., Nambudiri, R., and Holland, S. (2013) Regulation of the carbon- concentrating mechanism in the cyanobacterium Synechocystis sp. PCC6803 in response to changing light intensity and inorganic carbon availability. Photosynth. Res. 118, 115–124. doi: 10.1007/s11120-013-9912-4 (32) Bustin, S. A., Benes, V., Garson, J. A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M. W., Shipley, G. L., Vandesompele, J., and Wittwer, C. T. (2009) The MIQE guidelines: Minimum information for publication of quantitative real-time PCR experiments. Clin. Chem. 55, 611–622. doi: 10.1373/clinchem.2008.112797 (33) Cai, F., Bernstein, S. L., Wilson, S. C., and Kerfeld, C. A. (2016) Production and characterization of synthetic carboxysome shells with incorporated luminal proteins. Plant Physiol. 170, 1868–1877. doi: 10.1104/pp.15.01822 (34) Cai, F., Menon, B. B., Cannon, G. C., Curry, K. J., Shively, J. M., and Heinhorst, S. (2009) The pentameric vertex proteins are necessary for the icosahedral carboxysome shell to function as a CO2 leakage barrier. PLoS ONE 4. doi: 10.1371/journal.pone.0007521 151 (35) Cai, F., Sutter, M., Bernstein, S. L., Kinney, J. N., and Kerfeld, C. A. (2015) Engineering bacterial microcompartment shells: Chimeric shell proteins and chimeric carboxysome shells. ACS Synth. Biol. 4, 444–453. doi: 10.1021/sb500226j (36) Cai, F., Sutter, M., Cameron, J. C., Stanley, D. N., Kinney, J. N., and Kerfeld, C. A. (2013) The structure of CcmP, a tandem bacterial microcompartment domain protein from the β- carboxysome, forms a subcompartment within a microcompartment. J. Biol. Chem. 288, 16055– 16063. doi: 10.1074/jbc.M113.456897 (37) Calvin, M., and Benson, A. A. (1948) The path of carbon in photosynthesis. Science 107, 476–480. (38) Cameron, J. C., Wilson, S. C., Bernstein, S. L., and Kerfeld, C. A. (2013) Biogenesis of a bacterial organelle: The carboxysome assembly pathway. Cell 155, 1131–1140. doi: 10.1016/j.cell.2013.10.044 (39) Campbell, D. (1996) Complementary chromatic adaptation alters photosynthetic strategies in the cyanobacterium Calothrix. Microbiology 142, 1255–1263. doi: 10.1099/13500872-142-5- 1255 (40) Casey, E. S., Kehoe, D. M., and Grossman, A. R. (1997) Suppression of mutants aberrant in light intensity responses of complementary chromatic adaptation. J. Bacteriol. 179, 4599–4606. (41) Chaudhury, S., Berrondo, M., Weitzner, B. D., Muthu, P., Bergman, H., and Gray, J. J. (2011) Benchmarking and analysis of protein docking performance in Rosetta v3.2. PLoS ONE 6. doi: 10.1371/journal.pone.0022477 (42) Cobley, J. G., Zerweck, E., Reyes, R., Mody, A., Seludo-Unson, J. R., Jaeger, H., Weerasuriya, S., and Navankasattusas, S. (1993) Construction of shuttle plasmids which can be efficiently mobilized from Escherichia coli into the chromatically adapting cyanobacterium, Fremyella diplosiphon. Plasmid 30, 90–105. doi: 10.1006/plas.1993.1037 (43) Cobley, J., Seneviratne, L., Drong, L., Thounaojam, M., Oda, J. F., and Carroll, J. (1999) Transposition of Tn5 derivatives in the chromatically adapting cyanobacterium, Fremyella Diplosiphon, in The Phototrophic Prokaryotes (Peschek, G. A., Löffelhardt, W., and Schmetterer, G., Eds.), pp 443–451. Springer US, Boston, MA. doi: 10.1007/978-1-4615-4827- 0_52 (44) Cot, S. S.-W., So, A. K.-C., and Espie, G. S. (2008) A multiprotein bicarbonate dehydration complex essential to carboxysome function in cyanobacteria. J. Bacteriol. 190, 936–945. doi: 10.1128/JB.01283-07 (45) Delwiche, C. F. (1999) Tracing the thread of plastid diversity through the tapestry of life. Am. Nat. 154, S164–S177. doi: 10.1086/303291 (46) Dou, Z., Heinhorst, S., Williams, E. B., Murin, C. D., Shively, J. M., and Cannon, G. C. (2008) CO2 fixation kinetics of Halothiobacillus neapolitanus mutant carboxysomes lacking 152 carbonic anhydrase suggest the shell acts as a diffusional barrier for CO2. J. Biol. Chem. 283, 10377–10384. doi: 10.1074/jbc.M709285200 (47) Edgar, R. C. (2004) MUSCLE: Multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797. doi: 10.1093/nar/gkh340 (48) Eisenhut, M., von Wobeser, E. A., Jonas, L., Schubert, H., Ibelings, B. W., Bauwe, H., Matthijs, H. C. P., and Hagemann, M. (2007) Long-term response toward inorganic carbon limitation in wild type and glycolate turnover mutants of the cyanobacterium Synechocystis sp. strain PCC 6803. Plant Physiol. 144, 1946–1959. doi: 10.1104/pp.107.103341 (49) Elhai, J. (1993) Strong and regulated promoters in the cyanobacterium Anabaena PCC 7120. FEMS Microbiol. Lett. 114, 179–184. doi: 10.1111/j.1574-6968.1993.tb06570.x (50) Elhai, J., Vepritskiy, A., Muro-Pastor, A. M., Flores, E., and Wolk, C. P. (1997) Reduction of conjugal transfer efficiency by three restriction activities of Anabaena sp. strain PCC 7120. J. Bacteriol. 179, 1998–2005. (51) Elhai, J., and Wolk, C. P. (1988) [83] Conjugal transfer of DNA to cyanobacteria, in Methods in Enzymology, pp 747–754. Academic Press. doi: 10.1016/0076-6879(88)67086-8 (52) Ellis, R. J. (1979) The most abundant protein in the world. Trends Biochem. Sci. 4, 241– 244. doi: 10.1016/0968-0004(79)90212-3 (53) Erb, T. J., and Zarzycki, J. (2018) A short history of rubisco: the rise and fall (?) of nature’s predominant CO2 fixing enzyme. Curr. Opin. Biotechnol. 49, 100–107. doi: 10.1016/j.copbio.2017.07.017 (54) Espie, G. S., and Kimber, M. S. (2011) Carboxysomes: Cyanobacterial rubisco comes in small packages. Photosynth. Res. 109, 7–20. doi: 10.1007/s11120-011-9656-y (55) Farquhar, G. D., von Caemmerer, S., and Berry, J. A. (1980) A biochemical model of photosynthetic CO2 assimilation in leaves of C3 species. Planta 149, 78–90. (56) Faulkner, M., Rodriguez-Ramos, J., Dykes, G. F., Owen, S. V., Casella, S., Simpson, D. M., Beynon, R. J., and Liu, L.-N. (2017) Direct characterization of the native structure and mechanics of cyanobacterial carboxysomes. Nanoscale 9, 10662–10673. doi: 10.1039/C7NR02524F (57) Feiz, L., Williams-Carrier, R., Wostrikoff, K., Belcher, S., Barkan, A., and Stern, D. B. (2012) Ribulose-1,5-bis-phosphate carboxylase/oxygenase accumulation factor1 is required for holoenzyme assembly in Maize. Plant Cell 24, 3435–3446. doi: 10.1105/tpc.112.102012 (58) Field, C. B., Behrenfeld, M. J., Randerson, J. T., and Falkowski, P. (1998) Primary production of the biosphere: Integrating terrestrial and oceanic components. Science 281, 237– 240. doi: 10.1126/science.281.5374.237 153 (59) Gantt, E., and Conti, S. F. (1969) Ultrastructure of blue-green algae. J. Bacteriol. 97, 1486– 1493. (60) Gaudana, S. B., Zarzycki, J., Moparthi, V. K., and Kerfeld, C. A. (2015) Bioinformatic analysis of the distribution of inorganic carbon transporters and prospective targets for bioengineering to increase Ci uptake by cyanobacteria. Photosynth. Res. 126, 99–109. doi: 10.1007/s11120-014-0059-8 (61) Gill, R. T., Katsoulakis, E., Schmitt, W., Taroncher-Oldenburg, G., Misra, J., and Stephanopoulos, G. (2002) Genome-wide dynamic transcriptional profiling of the light-to-dark transition in Synechocystis sp. strain PCC 6803. J. Bacteriol. 184, 3671–3681. doi: 10.1128/JB.184.13.3671-3681.2002 (62) Giovannoni, S. J., Turner, S., Olsen, G. J., Barns, S., Lane, D. J., and Pace, N. R. (1988) Evolutionary relationships among cyanobacteria and green chloroplasts. J. Bacteriol. 170, 3584– 3592. doi: 10.1128/jb.170.8.3584-3592.1988 (63) Gomez-Garcia, M. R., Fazeli, F., Grote, A., Grossman, A. R., and Bhaya, D. (2013) Role of polyphosphate in thermophilic Synechococcus sp. from microbial mats. J. Bacteriol. 195, 3309– 3319. doi: 10.1128/JB.00207-13 (64) Gonzalez-Esquer, C. R., Shubitowski, T. B., and Kerfeld, C. A. (2015) Streamlined construction of the cyanobacterial CO2-fixing organelle via protein domain fusions for use in plant synthetic biology. Plant Cell 27, 2637–2644. doi: 10.1105/tpc.15.00329 (65) Gorelova, O. A., Baulina, O. I., Rasmussen, U., and Koksharova, O. A. (2013) The pleiotropic effects of ftn2 and ftn6 mutations in cyanobacterium Synechococcus sp. PCC 7942. Protoplasma 250, 931–942. doi: 10.1007/s00709-012-0479-2 (66) Graan, T., and Ort, D. R. (1986) Detection of oxygen-evolving Photosystem II centers inactive in plastoquinone reduction. Biochim. Biophys. Acta Bioenerg. 852, 320–330. doi: 10.1016/0005-2728(86)90238-0 (67) Grabsztunowicz, M., Koskela, M. M., and Mulo, P. (2017) Post-translational modifications in regulation of chloroplast function: Recent advances. Front. Plant Sci. 8. doi: 10.3389/fpls.2017.00240 (68) Gray, J. J., Moughon, S., Wang, C., Schueler-Furman, O., Kuhlman, B., Rohl, C. A., and Baker, D. (2003) Protein–Protein Docking with Simultaneous Optimization of Rigid-body Displacement and Side-chain Conformations. J. Mol. Biol. 331, 281–299. doi: 10.1016/S0022- 2836(03)00670-3 (69) Guindon, S., Dufayard, J.-F., Lefort, V., Anisimova, M., Hordijk, W., and Gascuel, O. (2010) New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Syst. Biol. 59, 307–321. doi: 10.1093/sysbio/syq010 154 (70) Guitton, C., and Mache, R. (1987) Phosphorylation in vitro of the large subunit of the ribulose-1,5-bisphosphate carboxylase and of the glyceraldehyde-3-phosphate dehydrogenase. Eur. J. Biochem. 166, 249–254. doi: 10.1111/j.1432-1033.1987.tb13509.x (71) Gutu, A., and Kehoe, D. M. (2012) Emerging perspectives on the mechanisms, regulation, and distribution of light color acclimation in cyanobacteria. Mol. Plant 5, 1–13. doi: 10.1093/mp/ssr054 (72) Guzman, L. M., Belin, D., Carson, M. J., and Beckwith, J. (1995) Tight regulation, modulation, and high-level expression by vectors containing the arabinose PBAD promoter. J. Bacteriol. 177, 4121–4130. doi: 10.1128/jb.177.14.4121-4130.1995 (73) Hasse, D., Larsson, A. M., and Andersson, I. (2015) Structure of Arabidopsis thaliana rubisco activase. Acta Crystallogr. D Biol. Crystallogr. 71, 800–808. doi: 10.1107/S1399004715001182 (74) He, Y.-Y., and Häder, D.-P. (2002) Involvement of reactive oxygen species in the UV-B damage to the cyanobacterium Anabaena sp. J. Photochem. Photobiol. B 66, 73–80. doi: 10.1016/S1011-1344(01)00278-0 (75) Heber, U., Takahama, U., Neimanis, S., and Shimizu-Takahama, M. (1982) Transport as the basis of the kok effect. Levels of some photosynthetic intermediates and activation of light- regulated enzymes during photosynthesis of chloroplasts and green leaf protoplasts. Biochim. Biophys. Acta Bioenerg. 679, 287–299. doi: 10.1016/0005-2728(82)90299-7 (76) Hihara, Y., Kamei, A., Kanehisa, M., Kaplan, A., and Ikeuchi, M. (2001) DNA microarray analysis of cyanobacterial gene expression during acclimation to high light. Plant Cell 13, 793– 806. (77) Hosseinzadeh, P., Bhardwaj, G., Mulligan, V. K., Shortridge, M. D., Craven, T. W., Pardo- Avila, F., Rettie, S. A., Kim, D. E., Silva, D.-A., Ibrahim, Y. M., Webb, I. K., Cort, J. R., Adkins, J. N., Varani, G., and Baker, D. (2017) Comprehensive computational design of ordered peptide macrocycles. Science 358, 1461–1466. doi: 10.1126/science.aap7577 (78) Huang, J., Wang, J., and Xu, H. (2014) The circadian rhythms of photosynthesis, ATP content and cell division in Microcystisaeruginosa PCC7820. Acta Physiol. Plant. 36, 3315– 3323. doi: 10.1007/s11738-014-1699-1 (79) Huang, L., McCluskey, M. P., Ni, H., and LaRossa, R. A. (2002) Global gene expression profiles of the cyanobacterium Synechocystis sp. strain PCC 6803 in response to irradiation with UV-B and white light. J. Bacteriol. 184, 6845–6858. doi: 10.1128/JB.184.24.6845-6858.2002 (80) Iancu, C. V., Ding, H. J., Morris, D. M., Dias, D. P., Gonzales, A. D., Martino, A., and Jensen, G. J. (2007) The structure of isolated Synechococcus strain WH8102 carboxysomes as revealed by electron cryotomography. J. Mol. Biol. 372, 764–773. doi: 10.1016/j.jmb.2007.06.059 155 (81) Ito, H., Mutsuda, M., Murayama, Y., Tomita, J., Hosokawa, N., Terauchi, K., Sugita, C., Sugita, M., Kondo, T., and Iwasaki, H. (2009) Cyanobacterial daily life with Kai-based circadian and diurnal genome-wide transcriptional control in Synechococcus elongatus. Proc. Natl. Acad. Sci. 106, 14168–14173. doi: 10.1073/pnas.0902587106 (82) Jensen, T., and Bowen, C. (1961) Organization of the centroplasm in Nostoc pruniforme. Proc. Iowa Acad. Sci. 68, 86–89. (83) Jordan, D. B., and Chollet, R. (1983) Inhibition of ribulose bisphosphate carboxylase by substrate ribulose 1,5-bisphosphate. J. Biol. Chem. 258, 13752–13758. (84) Kaiser, E., Morales, A., and Harbinson, J. (2018) Fluctuating light takes crop photosynthesis on a rollercoaster ride. Plant Physiol. 176, 977–989. doi: 10.1104/pp.17.01250 (85) Kakani, V. G., Surabhi, G. K., and Reddy, K. R. (2008) Photosynthesis and fluorescence responses of C4 plant Andropogon gerardii acclimated to temperature and carbon dioxide. Photosynthetica 46, 420–430. doi: 10.1007/s11099-008-0074-0 (86) Kaplan, A., and Reinhold, L. (1999) CO2 concentrating mechanisms in photosynthetic microorganisms. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 539–570. doi: 10.1146/annurev.arplant.50.1.539 (87) Karpova, T., and McNally, J. G. (2006) Detecting protein–protein interactions with CFP- YFP FRET by acceptor photobleaching. Curr. Protoc. Cytom. 35, 12.7.1-12.7.11. doi: 10.1002/0471142956.cy1207s35 (88) Katoh, K., Kuma, K., Toh, H., and Miyata, T. (2005) MAFFT version 5: Improvement in accuracy of multiple sequence alignment. Nucleic Acids Res. 33, 511–518. doi: 10.1093/nar/gki198 (89) Kehoe, D. M., and Grossman, A. R. (1996) Similarity of a chromatic adaptation sensor to phytochrome and ethylene receptors. Science 273, 1409–1412. doi: 10.1126/science.273.5280.1409 (90) Kehoe, D. M., and Grossman, A. R. (1997) New classes of mutants in complementary chromatic adaptation provide evidence for a novel four-step phosphorelay system. J. Bacteriol. 179, 3914–3921. (91) Kerfeld, C. A., and Melnicki, M. R. (2016) Assembly, function and evolution of cyanobacterial carboxysomes. Curr. Opin. Plant Biol. 31, 66–75. doi: 10.1016/j.pbi.2016.03.009 (92) Kerfeld, C. A., Sawaya, M. R., Tanaka, S., Nguyen, C. V., Phillips, M., Beeby, M., and Yeates, T. O. (2005) Protein structures forming the shell of primitive bacterial organelles. Science 309, 936–938. doi: 10.1126/science.1113397 (93) Khan, S., Andralojc, P. J., Lea, P. J., and Parry, M. A. J. (1999) 2′‐Carboxy‐D‐arabitinol 1‐ phosphate protects ribulose 1,5‐bisphosphate carboxylase/oxygenase against proteolytic breakdown. Eur. J. Biochem. 266, 840–847. doi: 10.1046/j.1432-1327.1999.00913.x 156 (94) Kinney, J. N., Salmeen, A., Cai, F., and Kerfeld, C. A. (2012) Elucidating essential role of conserved carboxysomal protein CcmN reveals common feature of bacterial microcompartment assembly. J. Biol. Chem. 287, 17729–17736. doi: 10.1074/jbc.M112.355305 (95) Knight, S., Andersson, I., and Brändén, C.-I. (1990) Crystallographic analysis of ribulose 1,5-bisphosphate carboxylase from spinach at 2·4 Å resolution: Subunit interactions and active site. J. Mol. Biol. 215, 113–160. doi: 10.1016/S0022-2836(05)80100-7 (96) Kobayashi, Y., Inoue, Y., Shibata, K., and Heber, U. (1979) Control of electron flow in intact chloroplasts by the intrathylakoid pH, not by the phosphorylation potential. Planta 146, 481–486. doi: 10.1007/BF00380864 (97) Koksharova, O. A., and Wolk, C. P. (2002) A novel gene that bears a DnaJ motif influences cyanobacterial cell division. J. Bacteriol. 184, 5524–5528. doi: 10.1128/JB.184.19.5524- 5528.2002 (98) Krissinel, E., and Henrick, K. (2007) Inference of macromolecular assemblies from crystalline state. J. Mol. Biol. 774–797. doi: 10.1016/j.jmb.2007.05.022 (99) Kuriata, A., Chakraborty, M., Nathan Henderson, J., Hazra, S., J Serban, A., V T Pham, T., Levitus, M., and Maria Wachter, R. (2014) ATP and magnesium promote cotton short-form ribulose-1,5-bisphosphate carboxylase/oxygenase (rubisco) activase hexamer formation at low micromolar concentrations. Biochemistry 53. doi: 10.1021/bi500968h (100) Laskowski, R. A. (2017) The ProFunc function prediction server. Methods Mol. Biol. Clifton NJ 1611, 75–95. doi: 10.1007/978-1-4939-7015-5_7 (101) Lechno‐Yossef, S., Rohnke, B. A., Belza, A. C. O., Melnicki, M. R., Montgomery, B. L., and Kerfeld, C. A. (2019) Cyanobacterial carboxysomes contain an unique rubisco-activase-like protein. New Phytol. Early View. doi: 10.1111/nph.16195 (102) Li, L. A., and Tabita, F. R. (1997) Maximum activity of recombinant ribulose 1,5- bisphosphate carboxylase/oxygenase of Anabaena sp. strain CA requires the product of the rbcX gene. J. Bacteriol. 179, 3793–3796. (103) Li, L. A., Zianni, M. R., and Tabita, F. R. (1999) Inactivation of the monocistronic rca gene in Anabaena variabilis suggests a physiological ribulose bisphosphate carboxylase/oxygenase activase-like function in heterocystous cyanobacteria. Plant Mol. Biol. 40, 467–478. doi: 10.1023/a:1006251808625 (104) Li, L., Alvey, R. M., Bezy, R. P., and Kehoe, D. M. (2008) Inverse transcriptional activities during complementary chromatic adaptation are controlled by the response regulator RcaC binding to red and green light-responsive promoters. Mol. Microbiol. 68, 286–297. doi: 10.1111/j.1365-2958.2008.06151.x (105) Li, L., and Kehoe, D. M. (2005) In vivo analysis of the roles of conserved aspartate and histidine residues within a complex response regulator. Mol. Microbiol. 55, 1538–1552. doi: 10.1111/j.1365-2958.2005.04491.x 157 (106) Li, L., and Kehoe, D. M. (2008) Abundance changes of the response regulator RcaC require specific aspartate and histidine residues and are necessary for normal light color responsiveness. J. Bacteriol. 190, 7241–7250. doi: 10.1128/JB.00762-08 (107) Li, L.-A., Janet, L., Gibson, and Robert Tabita, F. (1993) The rubisco activase (rca) gene is located downstream from rbcS in Anabaena sp. strain CA and is detected in other Anabaena/Nostoc strains. Plant Mol. Biol. 21, 753–764. doi: 10.1007/BF00027109 (108) Liang, Y., Urano, D., Liao, K.-L., Hedrick, T. L., Gao, Y., and Jones, A. M. (2017) A nondestructive method to estimate the chlorophyll content of Arabidopsis seedlings. Plant Methods 13, 26. doi: 10.1186/s13007-017-0174-6 (109) Liberton, M., Austin, J. R., Berg, R. H., and Pakrasi, H. B. (2011) Unique thylakoid membrane architecture of a unicellular N2-fixing cyanobacterium revealed by electron tomography. Plant Physiol. 155, 1656–1666. doi: 10.1104/pp.110.165332 (110) Liu, H., Landry, M. R., Vaulot, D., and Campbell, L. (1999) Prochlorococcus growth rates in the central equatorial Pacific: An application of the ƒmax approach. J. Geophys. Res. Oceans 104, 3391–3399. doi: 10.1029/1998JC900011 (111) Lohrig, K., Müller, B., Davydova, J., Leister, D., and Wolters, D. A. (2009) Phosphorylation site mapping of soluble proteins: Bioinformatical filtering reveals potential plastidic phosphoproteins in Arabidopsis thaliana. Planta 229, 1123–1134. doi: 10.1007/s00425- 009-0901-y (112) Long, B. M., Badger, M. R., Whitney, S. M., and Price, G. D. (2007) Analysis of carboxysomes from Synechococcus PCC7942 reveals multiple rubisco complexes with carboxysomal proteins CcmM and CcaA. J. Biol. Chem. 282, 29323–29335. doi: 10.1074/jbc.M703896200 (113) Long, B. M., Rae, B. D., Badger, M. R., and Price, G. D. (2011) Over-expression of the β- carboxysomal CcmM protein in Synechococcus PCC7942 reveals a tight co-regulation of carboxysomal carbonic anhydrase (CcaA) and M58 content. Photosynth. Res. 109, 33–45. doi: 10.1007/s11120-011-9659-8 (114) Long, B. M., Tucker, L., Badger, M. R., and Price, G. D. (2010) Functional cyanobacterial β-carboxysomes have an absolute requirement for both long and short forms of the CcmM protein. Plant Physiol. 153, 285–293. doi: 10.1104/pp.110.154948 (115) Long, S. P., and Bernacchi, C. J. (2003) Gas exchange measurements, what can they tell us about the underlying limitations to photosynthesis? Procedures and sources of error. J. Exp. Bot. 54, 2393–2401. (116) Ludwig, M., Sultemeyer, D., and Price, G. D. (2000) Isolation of ccmKLMN genes from the marine cyanobacterium, Synechococcus sp PCC7002 (Cyanophyceae) and evidence that CcmM is essential for carboxysome assembly. J. Phycol. 36, 1109–1119. doi: 10.1046/j.1529- 8817.2000.00028.x 158 (117) van Lun, M., van der Spoel, D., and Andersson, I. (2011) Subunit interface dynamics in hexadecameric rubisco. J. Mol. Biol. 411, 1083–1098. doi: 10.1016/j.jmb.2011.06.052 (118) MacCready, J. S., Hakim, P., Young, E. J., Hu, L., Liu, J., Osteryoung, K. W., Vecchiarelli, A. G., and Ducat, D. C. (2018) Protein gradients on the nucleoid position the carbon-fixing organelles of cyanobacteria. eLife 7, e39723. doi: doi.org/10.7554/eLife.39723 (119) Mackinder, L. C. M., Chen, C., Leib, R. D., Patena, W., Blum, S. R., Rodman, M., Ramundo, S., Adams, C. M., and Jonikas, M. C. (2017) A spatial interactome reveals the protein organization of the algal CO2-concentrating mechanism. Cell 171, 133-147.e14. doi: 10.1016/j.cell.2017.08.044 (120) Mahinthichaichan, P., Morris, D. M., Wang, Y., Jensen, G. J., and Tajkhorshid, E. (2018) Selective permeability of carboxysome shell pores to anionic molecules. bioRxiv. doi: 10.1101/367714 (121) Mangan, N. M., and Brenner, M. P. (2014) Systems analysis of the CO2 concentrating mechanism in cyanobacteria. eLife (Milo, R., Ed.) 3, e02043. doi: 10.7554/eLife.02043 (122) de Marsac, N. T., and Houmard, J. (1988) [34] Complementary chromatic adaptation: Physiological conditions and action spectra, in Methods in Enzymology, pp 318–328. Academic Press. doi: 10.1016/0076-6879(88)67037-6 (123) Mcginn, P. J., Price, G. D., and Badger, M. R. (2004) High light enhances the expression of low-CO2-inducible transcripts involved in the CO2-concentrating mechanism in Synechocystis sp. PCC6803. Plant Cell Environ. 27, 615–626. doi: 10.1111/j.1365-3040.2004.01175.x (124) McGinn, P. J., Price, G. D., Maleszka, R., and Badger, M. R. (2003) Inorganic carbon limitation and light control the expression of transcripts related to the CO2-concentrating mechanism in the cyanobacterium Synechocystis sp. strain PCC6803. Plant Physiol. 132, 218– 229. doi: 10.1104/pp.019349 (125) McGurn, L. D., Moazami-Goudarzi, M., White, S. A., Suwal, T., Brar, B., Tang, J. Q., Espie, G. S., and Kimber, M. S. (2016) The structure, kinetics and interactions of the β- carboxysomal β-carbonic anhydrase, CcaA. Biochem. J. 473, 4559–4572. doi: 10.1042/BCJ20160773 (126) McKay, R. M. L., and Gibbs, S. P. (1991) Composition and function of pyrenoids: Cytochemical and immunocytochemical approaches. Can. J. Bot. 69, 1040–1052. doi: 10.1139/b91-134 (127) McKay, R. M. L., Gibbs, S. P., and Espie, G. S. (1993) Effect of dissolved inorganic carbon on the expression of carboxysomes, localization of Rubisco and the mode of inorganic carbon transport in cells of the cyanobacterium Synechococcus UTEX 625. Arch. Microbiol. 159, 21–29. doi: 10.1007/BF00244259 (128) Menon, B. B., Heinhorst, S., Shively, J. M., and Cannon, G. C. (2010) The carboxysome shell is permeable to protons. J. Bacteriol. 192, 5881–5886. doi: 10.1128/JB.00903-10 159 (129) Montgomery, B. L. (2016) Mechanisms and fitness implications of photomorphogenesis during chromatic acclimation in cyanobacteria. J. Exp. Bot. 67, 4079–4090. doi: 10.1093/jxb/erw206 (130) Montgomery, B. L., Lechno-Yossef, S., and Kerfeld, C. A. (2016) Interrelated modules in cyanobacterial photosynthesis: the carbon-concentrating mechanism, photorespiration, and light perception. J. Exp. Bot. 67, 2931–2940. doi: 10.1093/jxb/erw162 (131) Mueller-Cajar, O. (2017) The diverse AAA+ machines that repair inhibited rubisco active sites. Front. Mol. Biosci. 4. doi: 10.3389/fmolb.2017.00031 (132) Mueller-Cajar, O., Stotz, M., Wendler, P., Hartl, F. U., Bracher, A., and Hayer-Hartl, M. (2011) Structure and function of the AAA+ protein CbbX, a red-type rubisco activase. Nature 479, 194–199. doi: 10.1038/nature10568 (133) Mulo, P., Laakso, S., Mäenpää, P., and Aro, E.-M. (1998) Stepwise photoinhibition of Photosystem II. Plant Physiol. 117, 483–490. (134) Murata, K., Hagiwara, S., Kimori, Y., and Kaneko, Y. (2016) Ultrastructure of compacted DNA in cyanobacteria by high-voltage cryo-electron tomography. Sci. Rep. 6, 34934. doi: 10.1038/srep34934 (135) Nagarajan, R., and Gill, K. S. (2018) Evolution of rubisco activase gene in plants. Plant Mol. Biol. 96, 69–87. doi: 10.1007/s11103-017-0680-y (136) Neuwald, A. F., Aravind, L., Spouge, J. L., and Koonin, E. V. (1999) AAA+: A class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res. 9, 27–43. doi: 10.1101/gr.9.1.27 (137) Niederhuber, M. J., Lambert, T. J., Yapp, C., Silver, P. A., and Polka, J. K. (2017) Superresolution microscopy of the β-carboxysome reveals a homogeneous matrix. Mol. Biol. Cell 28, 2734–2745. doi: 10.1091/mbc.E17-01-0069 (138) Nierzwicki-Bauer, S. A., Balkwill, D. L., and Stevens, S. E., Jr. (1983) Three-dimensional ultrastructure of a unicellular cyanobacterium. J. Cell Biol. 97, 713–722. (139) Nishihara, K., Kanemori, M., Kitagawa, M., Yanagi, H., and Yura, T. (1998) Chaperone coexpression plasmids: Differential and synergistic roles of DnaK-DnaJ-GrpE and GroEL- GroES in assisting folding of an allergen of Japanese cedar pollen, Cryj2, in Escherichia coli. Appl. Environ. Microbiol. 64, 1694–1699. (140) Nitta, K., Nagayama, K., Danev, R., and Kaneko, Y. (2009) Visualization of BrdU-labelled DNA in cyanobacterial cells by Hilbert differential contrast transmission electron microscopy. J. Microsc. 234, 118–123. doi: 10.1111/j.1365-2818.2009.03162.x (141) Oakley, C. A., Hopkinson, B. M., and Schmidt, G. W. (2012) A modular system for the measurement of CO2 and O2 gas flux and photosynthetic electron transport in microalgae. Limnol. Oceanogr. Methods 10, 968–977. doi: 10.4319/lom.2012.10.968 160 (142) Ogawa, T., Amichay, D., and Gurevitz, M. (1994) Isolation and characterization of the ccmM gene required by the cyanobacterium Synechocystis PCC6803 for inorganic carbon utilization. Photosynth. Res. 39, 183–190. doi: 10.1007/BF00029385 (143) Omata, T., Gohta, S., Takahashi, Y., Harano, Y., and Maeda, S. (2001) Involvement of a CbbR homolog in low CO2-induced activation of the bicarbonate transporter operon in cyanobacteria. J. Bacteriol. 183, 1891–1898. doi: 10.1128/JB.183.6.1891-1898.2001 (144) Omata, T., Price, G. D., Badger, M. R., Okamura, M., Gohta, S., and Ogawa, T. (1999) Identification of an ATP-binding cassette transporter involved in bicarbonate uptake in the cyanobacterium Synechococcus sp. strain PCC 7942. Proc. Natl. Acad. Sci. 96, 13571–13576. doi: 10.1073/pnas.96.23.13571 (145) Parry, M. A. J., Keys, A. J., Madgwick, P. J., Carmo-Silva, A. E., and Andralojc, P. J. (2008) Rubisco regulation: A role for inhibitors. J. Exp. Bot. 59, 1569–1580. (146) Pattanaik, B., Busch, A. W. U., Hu, P., Chen, J., and Montgomery, B. L. (2014) Responses to iron limitation are impacted by light quality and regulated by RcaE in the chromatically acclimating cyanobacterium Fremyella diplosiphon. Microbiology, 160, 992–1005. doi: 10.1099/mic.0.075192-0 (147) Pearce, F. G. (2006) Catalytic by-product formation and ligand binding by ribulose bisphosphate carboxylases from different phylogenies. Biochem. J. 399, 525–534. doi: 10.1042/BJ20060430 (148) Peña, K. L., Castel, S. E., Araujo, C. de, Espie, G. S., and Kimber, M. S. (2010) Structural basis of the oxidative activation of the carboxysomal γ-carbonic anhydrase, CcmM. Proc. Natl. Acad. Sci. 107, 2455–2460. doi: 10.1073/pnas.0910866107 (149) Peterson-Forbrook, D. S., Hilton, M. T., Tichacek, L., Henderson, J. N., Bui, H. Q., and Wachter, R. M. (2017) Nucleotide dependence of subunit rearrangements in short-form rubisco activase from spinach. Biochemistry 56, 4906–4921. doi: 10.1021/acs.biochem.7b00574 (150) Pfeffer, M., and Peisker, M. (1998) CO2 gas exchange and phosphoenolpyruvate carboxylase activity in leaves of Zea mays L. Photosynth. Res. 58, 11. doi: 10.1023/A:1006188705423 (151) Pollock, S. V., Colombo, S. L., Prout, D. L., Godfrey, A. C., and Moroney, J. V. (2003) Rubisco activase is required for optimal photosynthesis in the green alga Chlamydomonas reinhardtii in a low-CO2 atmosphere. Plant Physiol. 133, 1854–1861. doi: 10.1104/pp.103.032078 (152) Portis, A. R., Salvucci, M. E., and Ogren, W. L. (1986) Activation of ribulosebisphosphate carboxylase/oxygenase at physiological CO2 and ribulosebisphosphate concentrations by rubisco activase. Plant Physiol. 82, 967–971. 161 (153) Potter, S. C., Luciani, A., Eddy, S. R., Park, Y., Lopez, R., and Finn, R. D. (2018) HMMER web server: 2018 update. Nucleic Acids Res. 46, W200–W204. doi: 10.1093/nar/gky448 (154) Price, G. D., and Badger, M. R. (1989) Isolation and characterization of high CO2- requiring-mutants of the cyanobacterium Synechococcus PCC7942. Plant Physiol. 91, 514–525. (155) Price, G. D., Badger, M. R., Woodger, F. J., and Long, B. M. (2008) Advances in understanding the cyanobacterial CO2-concentrating-mechanism (CCM): functional components, Ci transporters, diversity, genetic regulation and prospects for engineering into plants. J. Exp. Bot. 59, 1441–1461. doi: 10.1093/jxb/erm112 (156) Price, G. D., Howitt, S. M., Harrison, K., and Badger, M. R. (1993) Analysis of a genomic DNA region from the cyanobacterium Synechococcus sp. strain PCC7942 involved in carboxysome assembly and function. J. Bacteriol. 175, 2871–2879. doi: 10.1128/jb.175.10.2871- 2879.1993 (157) Price, G. D., Woodger, F. J., Badger, M. R., Howitt, S. M., and Tucker, L. (2004) Identification of a SulP-type bicarbonate transporter in marine cyanobacteria. Proc. Natl. Acad. Sci. 101, 18228–18233. doi: 10.1073/pnas.0405211101 (158) R Core Team. (2016) R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna, Austria. (159) Rae, B. D., Long, B. M., Badger, M. R., and Price, G. D. (2012) Structural determinants of the outer shell of β-Carboxysomes in Synechococcus elongatus PCC 7942: Roles for CcmK2, K3-K4, CcmO, and CcmL. PLoS ONE (Lei, B., Ed.) 7, e43871. doi: 10.1371/journal.pone.0043871 (160) Rae, B. D., Long, B. M., Badger, M. R., and Price, G. D. (2013) Functions, compositions, and evolution of the two types of carboxysomes: Polyhedral microcompartments that facilitate CO2 fixation in cyanobacteria and some proteobacteria. Microbiol. Mol. Biol. Rev. 77, 357–379. doi: 10.1128/MMBR.00061-12 (161) Rausenberger, J., Hussong, A., Kircher, S., Kirchenbauer, D., Timmer, J., Nagy, F., Schäfer, E., and Fleck, C. (2010) An integrative model for phytochrome B mediated photomorphogenesis: From protein dynamics to physiology. PLoS ONE 5. doi: 10.1371/journal.pone.0010721 (162) Read, B. A., and Tabita, F. R. (1992) A hybrid ribulose bisphosphate carboxylase/oxygenase enzyme exhibiting a substantial increase in substrate specificity factor. Biochemistry 31, 5553–5560. doi: 10.1021/bi00139a018 (163) Read, B. A., and Tabita, F. R. (1994) High substrate specificity factor ribulose bisphosphate carboxylase/oxygenase from eukaryotic marine algae and properties of recombinant cyanobacterial rubisco containing “algal” residue modifications. Arch. Biochem. Biophys. 312, 210–218. doi: 10.1006/abbi.1994.1301 162 (164) Reed, J. W., Nagatani, A., Elich, T. D., Fagan, M., and Chory, J. (1994) Phytochrome A and phytochrome B have overlapping but distinct functions in Arabidopsis development. Plant Physiol. 104, 1139–1149. (165) Rippka, R., Deruelles, J., Waterbury, J. B., Herdman, M., and Stanier, R. Y. (1979) Generic assignments, strain histories and properties of pure cultures of cyanobacteria. Microbiology, 111, 1–61. doi: 10.1099/00221287-111-1-1 (166) Roach, T., and Krieger-Liszkay, A. K. (2014) Regulation of photosynthetic electron transport and photoinhibition. Curr. Protein Pept. Sci. 15, 351–362. doi: 10.2174/1389203715666140327105143 (167) Robinson, S. P., and Portis, A. R. (1989) Adenosine triphosphate hydrolysis by purified rubisco activase. Arch. Biochem. Biophys. 268, 93–99. doi: 10.1016/0003-9861(89)90568-7 (168) Rockwell, N. C., and Lagarias, J. C. (2010) A brief history of phytochromes. Chemphyschem 11, 1172–1180. doi: 10.1002/cphc.200900894 (169) Rockwell, N. C., Martin, S. S., and Lagarias, J. C. (2012) Red/Green cyanobacteriochromes: Sensors of color and power. Biochemistry 51, 9667–9677. doi: 10.1021/bi3013565 (170) Rohnke, B. A., Singh, S. P., Pattanaik, B., and Montgomery, B. L. (2018) RcaE-dependent regulation of carboxysome structural proteins has a central role in environmental determination of carboxysome morphology and abundance in Fremyella diplosiphon. mSphere 3, e00617-17. doi: 10.1128/mSphere.00617-17 (171) Ryan, P., Forrester, T. J. B., Wroblewski, C., Kenney, T. M. G., Kitova, E. N., Klassen, J. S., and Kimber, M. S. (2018) The small RbcS-like domains of the β-carboxysome structural protein, CcmM, bind rubisco at a site distinct from that binding the RbcS subunit. J. Biol. Chem. 294, 2593–2603. doi: 10.1074/jbc.RA118.006330 (172) Salvucci, M. E., and Ogren, W. L. (1996) The mechanism of rubisco activase: Insights from studies of the properties and structure of the enzyme. Photosynth. Res. 47, 1–11. doi: 10.1007/BF00017748 (173) Santos, J. M., Freire, P., Vicente, M., and Arraiano, C. M. (1999) The stationary-phase morphogene bolA from Escherichia coli is induced by stress during early stages of growth. Mol. Microbiol. 32, 789–798. doi: 10.1046/j.1365-2958.1999.01397.x (174) Savage, D. F., Afonso, B., Chen, A. H., and Silver, P. A. (2010) Spatially ordered dynamics of the bacterial carbon fixation machinery. Science 327, 1258–1261. doi: 10.1126/science.1186090 (175) Schaefer, M. R., Chiang, G. G., Cobley, J. G., and Grossman, A. R. (1993) Plasmids from two morphologically distinct cyanobacterial strains share a novel replication origin. J. Bacteriol. 175, 5701–5705. 163 (176) Schneider, C. A., Rasband, W. S., and Eliceiri, K. W. (2012) NIH Image to ImageJ: 25 years of image analysis. Nat. Methods 9, 671–675. (177) Schopf, J. W. (2012) The fossil record of cyanobacteria, in Ecology of cyanobacteria II: Their diversity in space and time (Whitton, B. A., Ed.), pp 15–36. Springer, New York. (178) Schuster-Böckler, B., Schultz, J., and Rahmann, S. (2004) HMM Logos for visualization of protein families. BMC Bioinformatics 5, 7. doi: 10.1186/1471-2105-5-7 (179) Schwarz, D., Nodop, A., Hüge, J., Purfürst, S., Forchhammer, K., Michel, K.-P., Bauwe, H., Kopka, J., and Hagemann, M. (2011) Metabolic and transcriptomic phenotyping of inorganic carbon acclimation in the cyanobacterium Synechococcus elongatus PCC 7942. Plant Physiol. 155, 1640–55. (180) Seib, L. O., and Kehoe, D. M. (2002) A turquoise mutant genetically separates expression of genes encoding phycoerythrin and its associated linker peptides. J. Bacteriol. 184, 962–970. doi: 10.1128/jb.184.4.962-970.2002 (181) Seki, Y., Nitta, K., and Kaneko, Y. (2014) Observation of polyphosphate bodies and DNA during the cell division cycle of Synechococcus elongatus PCC 7942. Plant Biol. 16, 258–263. doi: 10.1111/plb.12008 (182) Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. (1996) Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68, 850–858. doi: 10.1021/ac950914h (183) Shibata, M., Katoh, H., Sonoda, M., Ohkawa, H., Shimoyama, M., Fukuzawa, H., Kaplan, A., and Ogawa, T. (2002) Genes essential to sodium-dependent bicarbonate transport in cyanobacteria FUNCTION AND PHYLOGENETIC ANALYSIS. J. Biol. Chem. 277, 18658– 18664. doi: 10.1074/jbc.M112468200 (184) Shibata, M., Ohkawa, H., Kaneko, T., Fukuzawa, H., Tabata, S., Kaplan, A., and Ogawa, T. (2001) Distinct constitutive and low-CO2-induced CO2 uptake systems in cyanobacteria: Genes involved and their phylogenetic relationship with homologous genes in other organisms. Proc. Natl. Acad. Sci. 98, 11789–11794. doi: 10.1073/pnas.191258298 (185) Shih, P. M., Wu, D., Latifi, A., Axen, S. D., Fewer, D. P., Talla, E., Calteau, A., Cai, F., Marsac, N. T. de, Rippka, R., Herdman, M., Sivonen, K., Coursin, T., Laurent, T., Goodwin, L., Nolan, M., Davenport, K. W., Han, C. S., Rubin, E. M., Eisen, J. A., Woyke, T., Gugger, M., and Kerfeld, C. A. (2013) Improving the coverage of the cyanobacterial phylum using diversity- driven genome sequencing. Proc. Natl. Acad. Sci. 110, 1053–1058. doi: 10.1073/pnas.1217107110 (186) Singh, shailendra pratap, and L Montgomery, B. (2012) Reactive oxygen species are involved in the morphology-determining mechanism of Fremyella diplosiphon cells during complementary chromatic adaptation. Microbiology 158, 2235–45. doi: 10.1099/mic.0.060475-0 164 (187) Singh, S. P., and Montgomery, B. L. (2011) Temporal responses of wild-type pigmentation and RcaE-deficient strains of Fremyella diplosiphon during light transitions. Commun. Integr. Biol. 4, 503–510. doi: 10.4161/cib.16788 (188) Singh, S. P., and Montgomery, B. L. (2013) Distinct salt-dependent effects impair Fremyella diplosiphon pigmentation and cellular shape. Plant Signal. Behav. 8, e24713. doi: 10.4161/psb.24713 (189) Singh, S. P., and Montgomery, B. L. (2014) Morphogenes bolA and mreB mediate the photoregulation of cellular morphology during complementary chromatic acclimation in Fremyella diplosiphon. Mol. Microbiol. 93, 167–182. doi: 10.1111/mmi.12649 (190) Singh, S. P., and Montgomery, B. L. (2015) Regulation of BolA abundance mediates morphogenesis in Fremyella diplosiphon. Front. Microbiol. 6. doi: 10.3389/fmicb.2015.01215 (191) Somerville, C. R., Portis, A. R., and Ogren, W. L. (1982) A mutant of Arabidopsis thaliana which lacks activation of RuBP carboxylase in vivo. Plant Physiol. 70, 381–387. (192) Sommer, M., Cai, F., Melnicki, M., and Kerfeld, C. A. (2017) β-Carboxysome bioinformatics: identification and evolution of new bacterial microcompartment protein gene classes and core locus constraints. J. Exp. Bot. 68, 3841–3855. doi: 10.1093/jxb/erx115 (193) Sommer, M., Sutter, M., Gupta, S., Kirst, H., Turmo, A., Lechno-Yossef, S., Burton, R. L., Saechao, C., Sloan, N. B., Cheng, X., Chan, L.-J. G., Petzold, C. J., Fuentes-Cabrera, M., Ralston, C. Y., and Kerfeld, C. A. (2019) Heterohexamers formed by CcmK3 and CcmK4 increase the complexity of beta carboxysome shells. Plant Physiol. 179, 156–167. doi: 10.1104/pp.18.01190 (194) Spreitzer, R. J., and Salvucci, M. E. (2002) Rubisco: Structure, regulatory interactions, and possibilities for a better enzyme. Annu. Rev. Plant Biol. 53, 449–475. doi: 10.1146/annurev.arplant.53.100301.135233 (195) Stamatakis, A. (2014) RAxML version 8: A tool for phylogenetic analysis and post- analysis of large phylogenies. Bioinformatics 30, 1312–1313. doi: 10.1093/bioinformatics/btu033 (196) Stotz, M., Mueller-cajar, O., Ciniawsky, S., Wendler, P., Hartl, F. U., Bracher, A., and Hayer-Hartl, M. (2011) Structure of green-type rubisco activase from tobacco. Nat. Struct. Mol. Biol. 18, 1366–70. doi: 10.1038/nsmb.2171 (197) Stowe-Evans, E. L., Ford, J., and Kehoe, D. M. (2004) Genomic DNA microarray analysis: Identification of new genes regulated by light color in the cyanobacterium Fremyella diplosiphon. J. Bacteriol. 186, 4338–4349. doi: 10.1128/JB.186.13.4338-4349.2004 (198) Sültemeyer, D., Klughammer, B., Ludwig, M., Badger, M. R., and Price, G. D. (1997) Random insertional mutagenesis used in the generation of mutants of the marine cyanobacterium Synechococcus sp. Strain PCC7002 with an impaired CO2 concentrating mechanism. Funct. Plant Biol. 24, 317–327. doi: 10.1071/pp96124 165 (199) Sun, Y., Casella, S., Fang, Y., Huang, F., Faulkner, M., Barrett, S., and Liu, L.-N. (2016) Light modulates the biosynthesis and organization of cyanobacterial carbon fixation machinery through photosynthetic electron flow. Plant Physiol. 171, 530–541. doi: 10.1104/pp.16.00107 (200) Sun, Y., Wollman, A. J. M., Huang, F., Leake, M. C., and Liu, L.-N. (2019) Single- organelle quantification reveals stoichiometric and structural variability of carboxysomes dependent on the environment. Plant Cell 31, 1648–1664. doi: 10.1105/tpc.18.00787 (201) Sutter, M., Greber, B., Aussignargues, C., and Kerfeld, C. A. (2017) Assembly principles and structure of a 6.5-MDa bacterial microcompartment shell. Science 356, 1293–1297. doi: 10.1126/science.aan3289 (202) Sutter, M., Roberts, E. W., Gonzalez, R. C., Bates, C., Dawoud, S., Landry, K., Cannon, G. C., Heinhorst, S., and Kerfeld, C. A. (2015) Structural characterization of a newly identified component of α-carboxysomes: The AAA+ domain protein CsoCbbQ. Sci. Rep. 5, 16243. doi: 10.1038/srep16243 (203) Sutter, M., Wilson, S. C., Deutsch, S., and Kerfeld, C. A. (2013) Two new high-resolution crystal structures of carboxysome pentamer proteins reveal high structural conservation of CcmL orthologs among distantly related cyanobacterial species. Photosynth. Res. 118, 9–16. doi: 10.1007/s11120-013-9909-z (204) Tabita, F. R. (1999) Microbial ribulose 1,5-bisphosphate carboxylase/oxygenase: A different perspective. Photosynth. Res. 60, 1–28. doi: 10.1023/A:1006211417981 (205) Tabita, F. R., and Colletti, C. (1979) Carbon dioxide assimilation in cyanobacteria: Regulation of ribulose, 1,5-bisphosphate carboxylase. J. Bacteriol. 140, 452–458. (206) Tanaka, S., Kerfeld, C. A., Sawaya, M. R., Cai, F., Heinhorst, S., Cannon, G. C., and Yeates, T. O. (2008) Atomic-level models of the bacterial carboxysome shell. Science 319, 1083–1086. doi: 10.1126/science.1151458 (207) Tcherkez, G. G. B., Farquhar, G. D., and Andrews, T. J. (2006) Despite slow catalysis and confused substrate specificity, all ribulose bisphosphate carboxylases may be nearly perfectly optimized. Proc. Natl. Acad. Sci. 103, 7246–7251. doi: 10.1073/pnas.0600605103 (208) Tchernov, D., Hassidim, M., Luz, B., Sukenik, A., Reinhold, L., and Kaplan, A. (1997) Sustained net CO2 evolution during photosynthesis by marine microorganism. Curr. Biol. 7, 723–728. doi: 10.1016/S0960-9822(06)00330-7 (209) Terauchi, K., Montgomery, B. L., Grossman, A. R., Lagarias, J. C., and Kehoe, D. M. (2004) RcaE is a complementary chromatic adaptation photoreceptor required for green and red light responsiveness. Mol. Microbiol. 51, 567–577. doi: 10.1046/j.1365-2958.2003.03853.x (210) Turmo, A., Gonzalez-Esquer, C. R., and Kerfeld, C. A. (2017) Carboxysomes: Metabolic modules for CO2 fixation. FEMS Microbiol. Lett. 364. doi: 10.1093/femsle/fnx176 166 (211) Ulijasz, A. T., and Vierstra, R. D. (2011) Phytochrome structure and photochemistry: Recent advances toward a complete molecular picture. Curr. Opin. Plant Biol. 14, 498–506. doi: 10.1016/j.pbi.2011.06.002 (212) Wang, C., Schueler-Furman, O., and Baker, D. (2005) Improved side-chain modeling for protein–protein docking. Protein Sci. Publ. Protein Soc. 14, 1328–1339. doi: 10.1110/ps.041222905 (213) Wang, H., Yan, X., Aigner, H., Bracher, A., Nguyen, N. D., Hee, W. Y., Long, B. M., Price, G. D., Hartl, F. U., and Hayer-Hartl, M. (2019) Rubisco condensate formation by CcmM in β-carboxysome biogenesis. Nature 566, 131. doi: 10.1038/s41586-019-0880-5 (214) Wang, H.-L., Postier, B. L., and Burnap, R. L. (2004) Alterations in global patterns of gene expression in Synechocystis sp. PCC 6803 in response to inorganic carbon limitation and the inactivation of ndhR, a LysR family regulator. J. Biol. Chem. 279, 5739–5751. doi: 10.1074/jbc.M311336200 (215) Wang, Q., Serban, A. J., Wachter, R. M., and Moerner, W. E. (2018) Single-molecule diffusometry reveals the nucleotide-dependent oligomerization pathways of Nicotiana tabacum rubisco activase. J. Chem. Phys. 148, 123319. doi: 10.1063/1.5005930 (216) Waterhouse, A., Bertoni, M., Bienert, S., Studer, G., Tauriello, G., Gumienny, R., Heer, F. T., de Beer, T. A. P., Rempfer, C., Bordoli, L., Lepore, R., and Schwede, T. (2018) SWISS- MODEL: Homology modelling of protein structures and complexes. Nucleic Acids Res. gky427. doi: 10.1093/nar/gkg520 (217) Watson, G. M., and Tabita, F. R. (1996) Regulation, unique gene organization, and unusual primary structure of carbon fixation genes from a marine phycoerythrin-containing cyanobacterium. Plant Mol. Biol. 32, 1103–1115. doi: 10.1007/bf00041394 (218) Wheeler, T. J., Clements, J., and Finn, R. D. (2014) Skylign: A tool for creating informative, interactive logos representing sequence alignments and profile hidden Markov models. BMC Bioinformatics 15, 7. doi: 10.1186/1471-2105-15-7 (219) Whitehead, L., Long, B. M., Price, G. D., and Badger, M. R. (2014) Comparing the in vivo function of α-carboxysomes and β-carboxysomes in two model cyanobacteria. Plant Physiol. 165, 398–411. doi: 10.1104/pp.114.237941 (220) Whitton, B. A., and Potts, M. (2012) Introduction to the cyanobacteria, in Ecology of cyanobacteria II: Their diversity in space and time (Whitton, B. A., Ed.), pp 1–13. Springer, New York. (221) Zacarias, L., and Reid, M. S. (1990) Role of growth regulators in the senescence of Arabidopsis thaliana leaves. Physiol. Plant. 80, 549–554. doi: 10.1111/j.1399- 3054.1990.tb05677.x 167 (222) Zarzycki, J., Axen, S. D., Kinney, J. N., and Kerfeld, C. A. (2013) Cyanobacterial-based approaches to improving photosynthesis in plants. J. Exp. Bot. 64, 787–798. doi: 10.1093/jxb/ers294 (223) Zhang, N., Kallis, R. P., Ewy, R. G., and Portis, A. R. (2002) Light modulation of rubisco in Arabidopsis requires a capacity for redox regulation of the larger rubisco activase isoform. Proc. Natl. Acad. Sci. 99, 3330–3334. doi: 10.1073/pnas.042529999 (224) Zhang, P., Battchikova, N., Jansen, T., Appel, J., Ogawa, T., and Aro, E.-M. (2004) Expression and functional roles of the two distinct NDH-1 complexes and the carbon acquisition complex NdhD3/NdhF3/CupA/Sll1735 in Synechocystis sp PCC 6803. Plant Cell 16, 3326– 3340. doi: 10.1105/tpc.104.026526 (225) Zimorski, V., Ku, C., Martin, W. F., and Gould, S. B. (2014) Endosymbiotic theory for organelle origins. Curr. Opin. Microbiol. 22, 38–48. doi: 10.1016/j.mib.2014.09.008 (226) Zmasek, C. M., and Godzik, A. (2013) Evolution of the animal apoptosis network. Cold Spring Harb. Perspect. Biol. 5. doi: 10.1101/cshperspect.a008649 168