NC1200: Regulation of Photosynthetic Processes

(Multistate Research Project)

Status: Active

NC1200: Regulation of Photosynthetic Processes

Duration: 10/01/2022 to 09/30/2027

Administrative Advisor(s):


NIFA Reps:


Statement of Issues and Justification

STATEMENT OF THE ISSUES AND JUSTIFICATION

Necessity of Photosynthesis Research

Photosynthesis is essential to life on earth as it converts sunlight into biochemical energy used by nearly all life forms. It is the primary process for generating plant biomass. As a result of photosynthesis, carbon dioxide, as well as inorganic nitrogen and sulfur, are converted into organic molecules (e.g., sugars, lipids, amino acids and other cell building blocks). Oxygen is generated, as a byproduct, through photosynthetic water oxidation. Thus, plant and algal photosynthesis affects global geochemical processes, in particular carbon cycling (29), and is an important factor to be considered in global climate change models. Aside from these fundamental aspects of photosynthesis, agricultural production of food, feed, fiber, natural chemicals, and biofuel feedstocks is directly affected by limitations in the rate and yield of photosynthesis and the capacity to utilize fixed carbon (118, 132). Some of the greatest challenges that confront humankind – feeding an exponentially growing global population, supplying sufficient energy to sustain this global population, and averting negative environmental impacts due to human activities – can or need to be addressed using photosynthetic organisms (28). Therefore, the need for state-of-the-art photosynthesis research to improve the efficiency and productivity of this process in traditional crops or for the development of novel crops and products has never been more urgent.

Collaborators participating in this regional project will place considerable focus on understanding and improving the response of photosynthesis to genetic, developmental and environmental factors that limit productivity. The research spans all levels of organization from the genetic, molecular and cellular through the leaf, whole plant, and canopy.  Particular emphasis will be placed on abiotic stresses (i.e., heat, cold, drought and salinity), nitrogen- and water-use efficiency, carbon flux pathways, and the signal transduction mechanisms that initiate plant responses, because these are critical to developing a climate-smart agriculture (100). Factors that enhance or limit agricultural productivity generally do so by impacting photosynthesis.  The gains in yield of the major crops over the past half-century have come about primarily from breeding for greater harvest index (HI) and through management, particularly the application of fertilizer (39).  For well-fertilized crops, HI has approached the maximum achievable for many crops, and future yield gains will depend on increasing total production (113).  Greater productivity, which is necessary to meet the food, feed, fuel and fiber needs of a growing world population (102), requires improving the basic efficiency of the photosynthetic process for light energy capture, and the conversion of this energy to chemical form for the synthesis and utilization of organic molecules (113).  For maximum benefit, the efficiency of photosynthesis must also be resilient to changes in temperature, CO2, and precipitation / drought anticipated from climate change.

Importance and Consequences

Global climatic trends are negatively impacting the yields of major crops used for the production of food, feed, fiber, and biofuels (92). Sustaining the productivity of traditional crops will require improvements in the efficiency of photosynthesis that go beyond traditional breeding and selection (93). Similarly, the development of novel feedstocks for biofuel/chemicals generation in a way that is sustainable, commercially-viable, not in competition with food production, and that mitigates greenhouse gas emissions requires novel approaches and non-traditional crops including algae (109, 123) and cyanobacteria (30). Only a concerted and vertically-integrated effort encompassing all aspects of photosynthesis will ensure that appropriate solutions can be found for some of these most pressing problems currently facing society. Innovative thinking will be required that does not stop at traditional agricultural systems and crops, but may enable transitioning to new systems dedicated to the production of biofuels and chemicals instead of food. As photosynthesis is at the basis of biomass production, we need to find innovative ways to overcome its limitations. Failure to do so now will limit our future ability to produce sufficient food, feed, fiber, and fuel in a rapidly changing climate.

Technical Feasibility

Genomic resources and next generation sequencing and transformation technologies have advanced to the point that a wealth of information can be generated for any species of plant or alga (e.g., 10,000 plant genomes project) (116), in a relative short time span and at reasonable costs. Thus, the raw material for the genetic manipulation of novel crops or algal strains is readily available. Large-scale phenomics focused on chloroplast proteins in multiple plant species including genetic models such as Arabidopsis (24) and crops (127). Moreover, using comparative genomics, reconstruction of metabolism from gene expression and genomic data has become readily feasible for any organism. One example is the metabolic reconstruction of the alga Chlamydomonas reinhardtii (53) and the cyanobacterium Synechocystis sp PCC 6803 (64). In addition, our knowledge and analysis of primary metabolism of photosynthesis in plants and associated metabolic networks has advanced to levels (71) that can enable the rational design of novel crops, for example, establishing crassulacean acid metabolism (CAM) in plants with C3 metabolism (129). Despite this progress, the task of genetically transforming crop plants and analyzing the resulting phenotypes is still tedious and time consuming. Synthetic biology efforts with plants, involving stacking or replacing multiple genes, are lagging behind those for bacterial systems (97). Moreover, photosynthesis is one of the most complex processes found in nature, requiring hundreds of genes and proteins, and multiple and overlapping levels of regulation. Thus, to enable rapid progress in basic discovery and to devise strategies for the improvement of photosynthesis in crops, facile model organisms such as Arabidopsis (110) or model microalgae and cyanobacteria such as Chlamydomonas (98) and Synechocystis (20, 117) will have to be employed to quickly identify the most promising directions for photosynthetic pathway and crop productivity improvement. Following this guidance, collaboration with geneticists and breeders, associating traits with genomic regions of mapping populations or diversity panels can support marker-assisted selection (22, 41). Such an integrated approach requires multifaceted expertise and, thus, will benefit from synergy derived from a multi-state investigator effort.

Multi State Effort

Providing a conceptual framework through the current project, this group of scientists, bringing together a complementary set of expertise, and others located throughout the US have already successfully made progress on understanding diverse aspects of photosynthesis. While global issues as laid out above are addressed, practical solutions to these problems often have local solutions (e.g., by taking into account climatic zones to which specific crop species or algae are adapted). Continued effort by the current group will contribute towards these main goals while also enabling local solutions of particular value to the participating states. Partners in this endeavor are listed for each focus area below.

Likely Impacts

Efforts by the group are organized into four themes (Objectives). While the details and outcomes will be discussed below in the main body of the proposal, likely impacts falling under these themes can be briefly summarized as follows:

  1. Photosynthetic Capture and Photorespiratory Release of CO2. Photosynthetic carbon fixation and photorespiratory release of CO2 have long been recognized as limitations to crop productivity (37, 94, 99, 111). Carbon metabolism in photosynthesizing leaves involves the Calvin-Benson cycle but also other carbon fluxes. A more complete understanding of these carbon-reaction processes will identify mechanisms leading to improved photosynthetic efficiency. The carboxylating and oxygenating enzyme, Rubisco, is central to these efforts. As temperature increases, photorespiratory pressure in C3 plants increases due to decreased Rubisco specificity for CO2 relative to O2. However, the carboxylating activity of Rubisco is greater in C4 and CAM photosynthetic pathways, relative to that in C3 photosynthesis, in spite of the warmer ecozones these occupy. Understanding the gene expression regulation and regulatory network of C4 and CAM metabolic activities is the key to fully elucidate the evolutionary history of C4 and CAM photosynthesis, and to take advantage of these for improved crop productivity. The rates of photosynthesis in C3, C4 and CAM plants can be limited by the availability of CO2 at the initial site of carboxylation.  Understanding how leaf biochemical and anatomical traits influence the concentration of CO2 at the site of carboxylation may guide strategies to increase photosynthetic rates. Studies will utilize genomic analysis, loss of function mutants, 13CO2 isotope studies and metabolite analysis to understand carbon fluxes involved in photorespiration, transcriptional regulation of C4 and CAM metabolism and CO2 availability at the initial site of carboxylation. Likely impacts include an instrumental knowledge of factors regulating carbon fluxes in light-independent processes including effective CO2 concentrations at carboxylation sites, the costs and benefits of ancillary carbon paths in photosynthesizing cells, and circadian regulation in C3, C4 and CAM photosynthesis, with application to crop improvement (MI-ABR, NV-AES, TX-AES, WA-AES).
  2. Strategies to optimize the assembly and function of the photosynthetic membrane. Chloroplasts are the organelles that perform photosynthesis in both plants and algae. Chloroplasts also contain a large number of enzymes, highlighting the role of this organelle as a primary biochemical production factory. As semi-autonomous organelles, chloroplasts do not function by themselves, but rely on extensive communication with other organelles and compartments within the cell, and with the whole plant. The import of nuclear-encoded proteins or membrane lipids assembled at the endoplasmic reticulum provide two examples of such interactions (66, 114). As primary photosynthate and many other metabolites (e.g., fatty acids and most isoprenoids), are only synthesized in chloroplasts, they have to be exported to be available to other cellular compartments. The integration of chloroplast biogenesis into overall cell development requires intricate signaling processes, as does the adjustment of the photosynthetic electron transport chain to changing conditions. Within the chloroplast, the capture and conversion of light energy by photosynthesis occurs at a specialized structure called the thylakoid membrane, which is itself dynamically remodeled in response to developmental and stress cues. The architectural dynamics of the stacked grana thylakoids are involved in regulating and maintaining photosynthetic Studies will focus on how thylakoid membranes change their shape, the functional consequences of structural alterations, and effects of whole-plant stresses and developmental cues on the thylakoid membrane and chloroplast lipid changes. Likely impacts are a better understanding of photosynthetic energy transduction and transformation, the development of the thylakoid membranes under developmental and stress regimens, and the development of tools that can be used to assess chloroplast membrane connectivity (MI-ABR, NE-ARD, WA-AES).
  3. Mechanisms Regulating Photosynthate Partitioning. Manipulation of carbon partitioning and understanding its regulation is central to advances in yield and heterologous product formation in cereal, oilseed crops, and algae. This effort will contribute with the design of new photosynthetic systems for specialty and commodity chemicals production that possess enhanced carbon flux to innate or new sinks, e.g., starch, useful triacylglycerols (27), and antioxidant or isoprenoid products (81). Several related efforts are expected to have positive impacts on plant growth rates and yield. The “starch partitioning” project, comprising the mapping of genes impacting leaf starch levels will use field and greenhouse studies to determine the impact of sink capacity on photosynthetic rates and plant productivity. This will lead to information on mechanisms regulating photosynthate partitioning toward starch biosynthesis / accumulation. Another approach seeks to evaluate the function of PHO1, a higher plant plastid-localized phosphorylase in starch biosynthesis and its newly discovered interaction with photosystem I. The role of PHO1 in starch partitioning and photosynthesis will be investigated in relation to enhanced plant growth rates and grain yields. The “lipid partitioning” project will investigate plants with altered metabolism to accumulate significant storage reserves, such as lipids, in leaves. Metabolic studies will assess the functional and productivity consequences of photosynthetic carbon partitioning to lipid and storage in leaves, converting plant leaves into lipid storage tissues. The “antioxidants enhancement project” investigates evolutionary aspects of leaf antioxidant content in relation to biomass accumulation. Small antioxidant amounts, applied exogenously to plants, improve photosynthetic efficiency by altering the partitioning of light to photochemistry or non-photochemical quenching. This project will investigate the underlying reasons and explore the possibility of improving antioxidant pools in vivo. The “isoprenoids partitioning” project aims to enhance flux toward the synthesis of plant essential oils and related compounds with applications in flavor, fragrance, synthetic chemistry, and biopharmaceuticals. All of the above projects would benefit from the “fusion constructs” project, which will deliver a platform for substantially enhanced carbon partitioning toward a target biosynthetic pathway, via pathway enzyme over-expression (117). This would alleviate rate- and yield-limiting catalytic steps in the generation of endogenous and heterologous compounds.  Likely impacts include definition of the mechanisms that regulate photosynthate partitioning into the biosynthetic pathways for sucrose, starch, lipids, antioxidants, and isoprenoids. The effect of innate or heterologous sink strength on rates and capacity of photosynthesis will be assessed in each case. The above comprise an integrated approach to understanding carbon partitioning and its effect on product synthesis and accumulation.  (CA-AES, MO-ARS, MT-AES, NE-ARD, WA-AES).
  4. Developmental and Environmental Limitations to Photosynthetic Productivity. Factors such as leaf anatomy (112) or environmental stress conditions, such as high light (86), excess salinity, phosphorous deficiency, drought or heat stress (105) greatly affect photosynthesis and plant productivity. Moreover, leaf stomata, affecting photosynthetic productivity as a ‘gateway’ for CO2 influx, are subject to complex active regulation, responsive to factors including light intensity, temperature, vapor pressure, and leaf CO2 partial pressure. Heat stress during flowering and grain filling alters the flux of assimilates in wheat, corn, and sorghum (54, 95). Included are pollen infertility, which reduces grain number, while diminished assimilate flow reduces grain weight. Genomic analysis in this objective seeks to identify critical loci encoding heat and water stress tolerance traits. An additional approach combines adaptive traits (CAM, tissue succulence, thick cuticles and epicuticular wax, low stomatal density with high responsiveness and rectifier-like roots) to develop crops that are resilient to severe heat and water-deficit stress. Abiotic stress signaling systems involve inositol pyrophosphates (signal for phosphorus uptake and utilization) and calcium-dependent protein kinases (regulating responses to biotic and abiotic stress, metabolism, vegetative development and sexual reproduction). Small signaling peptides are known to regulate plant development. However, the functions of dehydration-stimulated peptides on water-limited photosynthesis have not been investigated. Studies will utilize genomic analysis, loss of function mutants, and metabolite analysis to identify factors regulating photosynthetic and photosynthate partitioning responses to abiotic stresses such as heat, water deficits, excess salinity and phosphorus deficiencies/toxicity. Activities in Objective 4 will thus be integrated with those of Objective 3 to provide a more holistic perspective on photosynthate partitioning, as this is defined by genetic and environmental stress conditions. Likely impacts include better understanding of the effect of abiotic stress on carbon fluxes, development of genomic tools for crop improvement and novel genetic resources for xeric conditions. (IN-AES, KS-AES, MO-AES, MS-AES, NE-ARD, NV-AES, VA-AES, TX-ARS).

Related, Current and Previous Work

Related current and previous work

Comprehensive CRIS database searches were performed by individual participants using specific and more general search terms such as photosynthesis alone (500 hits alone) and in combination with other terms such as lipids (26 hits), Rubisco (25 hits), carbon fixation (20 hits) etc.; conducted as relevant to the topic. Aside from projects associated with NC1200 members, no other NC group, investigating mechanistic and regulatory processes related to photosynthesis, was detected. It was apparent that no other regional project examines multiple scales of photosynthesis—from molecules to whole plant/field responses.

1. Identify strategies to optimize the assembly and function of the photosynthetic membrane. The conversion of sunlight into chemical energy by plant photosynthesis requires a specialized photosynthetic membrane forming the thylakoids inside chloroplasts. Chloroplast membranes are highly dynamic, changing their shape and composition rapidly in response to environmental cues. These dynamics are the basis for the functionality, regulation, and maintenance of photosynthetic performance under field conditions. Knowledge of the apparent self-assembly and self-repair of the photosynthetic membrane is expected to guide efforts in developing artificial photosynthetic membranes and enhance engineering of plant feedstocks. Compositional lipid and protein dynamics play an essential role in shaping the membrane architecture. Collaborators focus on two major sub-objectives: 1.1 Dynamics of chloroplast membrane architecture and 1.2 Chloroplast membrane lipid and protein dynamics. The first sub-objective addresses the importance of the three-dimensional arrangement of multiple components simultaneously, while the second enhances knowledge of production, delivery, and photosynthetic relevance of individual molecular components of chloroplast membranes.

1.1 Dynamics of chloroplast membrane architecture. The architecture of membranes inside the chloroplast is complex and dynamic. At its simplest iteration, the chloroplast consists of three independent membrane systems: the outer envelope membrane encapsulating the inner envelope membrane encapsulating the stroma and the photosynthetic thylakoid membrane with thylakoid lumen. However, variations in the thylakoid membrane structure and appearance of alternate membrane structures have long been observed in response to environmental cues and affect photosynthetic efficiency. Examples of thylakoid dynamics include light-responsive decreased grana thylakoids in the lateral dimension and increased vertical swelling of the thylakoid lumen, which respectively optimize molecular repair processes (42, 60,62, 96, 69), and facilitate diffusion of small electron carriers leading to efficient photosynthetic electron transport (42, 62, 59, 46). Alternate membrane structures include large invaginations of the inner envelope membrane (19, 84), small vesicles in the stroma (75, 122), and membrane contact sites between the inner and thylakoid membranes (18). Thus, knowledge of how chloroplast membranes change their shape is required for an in-depth understanding of photosynthetic energy transformation. In this sub-objective, we aim to understand architectural dynamics of chloroplast membranes and relate them to the functionality, regulation, and maintenance of energy conversion. The Kirchhoff lab (WA-AES) targets understanding of thylakoid structural dynamics on the whole membrane and supramolecular membrane-protein organization levels. These topics includes the light-induced vertical swelling and subsequent shrinkage of the thylakoid membrane system in the dark; the role of light-dependent swelling/shrinkage in control of diffusion dependent electron transport; lateral changes of the stacked grana thylakoid area accompanied by lateral redistribution of protein complexes; the role of lipids and small regulatory and structural protein in controlling the supramolecular protein organization and the functionality of membrane-embedded photosynthetic protein complexes; and influence of thylakoid membrane composition on the balance of physicochemical forces involved in membrane stacking. These different levels support a holistic understanding of structure/function dynamics in plant thylakoid membranes. The Roston lab (NE-ARD) studies alternate chloroplast membrane structures (vesicles, invaginations, contact sites) (65) by initiating production of multiple fluorescent visualization systems, developed in the lab, that determine thylakoid/inner envelope membrane continuity in Arabidopsis thaliana. Each system is designed to expand our understanding of the dynamic relationships between the membranes that contribute to photosynthetic stability.

1.2 Chloroplast membrane lipid and protein dynamics. Chloroplasts have unique membrane lipid and protein compositions. First examined over 50 years ago, chloroplasts are composed of specific lipids required for photosynthetic efficiency including galactolipids (MGDG, DGDG), sulfolipids (SQDG), and specific variants of phosphatidylglycerol (PG) (25, 34). Despite their long history, critical features of these lipids remain unknown. In this sub-objective, we aim to understand the synthesis, maintenance, and turnover of critical chloroplast lipids and proteins. The Benning lab (MI-ABR) has been instrumental in defining biosynthesis and turnover of thylakoid lipids, and regulation of those processes (26, 35, 36, 55, 63, 83, 124, 125, 128). Current investigations include biosynthesis, turnover, and transport of a key intermediate of thylakoid lipid biosynthesis (66, 67), phosphatidic acid, at the inner envelope membrane and determining the biosynthesis and physiological role of specific PG molecular species (47). The Roston lab (NE-AES) has recently defined cellular signals required to alter chloroplast galactolipid levels in response to abiotic stress that are critical for plant survival (13, 14, 119).

2. Identify strategies to modify biochemical and regulatory factors that impact the photosynthetic capture and photorespiratory release of CO2. The use of chemical energy to capture and convert CO2 into sugars is fundamental to plant growth and function. Almost all energy from sunlight captured by photosynthetic membranes is stored as carbon, and these carbon products supply the plant’s needs for energy and growth as well as food, feed, fiber, and fuel. Acquisition and metabolism of carbon can determine how fast the energy of sunlight can be turned into products. Knowledge of processes regulating photosynthetic capture and release of CO2 is expected to guide discovery and development of more productive and resilient plant growth. Collaborators focus on two sub-objectives: 2.1 Electron transport and regulation of the Calvin-Benson cycle, and 2.2 C4 and CAM CO2 concentrating mechanisms. The first sub-objective follows carbon after carboxylation and the second sub-objective traces CO2 accumulation mechanisms that optimize the operation of Rubisco.

2.1 Electron transport and regulation of the Calvin-Benson cycle. After carboxylation, carbon flows into the Calvin-Benson cycle until it is exported from the cycle for sucrose and starch synthesis. Recent work indicates that about 15% of the carbon follows the oxidative pentose phosphate pathway, called the glucose-6-phosphate (G6P) shunt, instead of the canonical Calvin-Benson cycle. In this sub-objective, we aim to understand the interaction of carbon metabolism and cyclic electron flow. The Sharkey and Walker labs (MI-ABR) are exploring assumptions about how flux proceeds through photosynthetic metabolism. A major advance was discovery that a glucose-6-phosphate shunt involving the oxidative pentose phosphate pathway likely occurs simultaneously with the Calvin Benson cycle (107). This metabolism releases CO2 at the same time CO2 is being assimilated in the Calvin-Benson cycle (126). This carbon pathway also allows carbon from a large pool of unlabeled glucose and sucrose to enter the Calvin-Benson cycle as ribulose 5-phosphate, limiting the degree of label (13CO2 or 12CO2) that can accumulate in Calvin-Benson cycle intermediates (106). This explains the mysterious lack of complete labeling that has been observed for many years. It was also shown that the lack of label varies with stress. We also investigated the conditions under which a glucose 6-phosphate transporter is expressed in photosynthesizing leaves (120). This collaboration helped determine that the biochemical source of the CO2-releasing process known as respiration in the light. Respiration in the light is a CO2-releasing process that is distinct from photorespiration and with unclear metabolic origins. We have determined that it does not come from the TCA cycle and instead likely originates via the G6P shunt (126). The G6P shunt may be critical for filling the Calvin-Benson cycle with intermediates in the morning and during light flecks. When carbon follows the G6P shunt, extra ATP is likely required resulting in cyclic electron flow around photosystem I. Currently, the Walker lab is characterizing how a C3 extremophile, Rhyza stricta, has adapted to process high photorespiratory rates at temperatures as high as 50°C. Current work indicates key photorespiratory enzymes appear important to maintain high fluxes under elevated temperatures. This project will lead to studies of the interaction of carbon metabolism and cyclic electron flow.

2.2 C4 and CAM CO2-concentrating mechanisms. CO2-concentrating mechanisms increase CO2 fixation rates and reduce water losses. In C3 plants, CO2 uptake relies on diffusion. This is costly in terms of water loss. Two mechanisms have evolved in terrestrial plants that significantly reduce the water cost of CO2 uptake. In C4 plants a spatially segregated series of reactions allows active accumulation of CO2 near Rubisco. This reduces the rate of photorespiration and also reduces the water cost per CO2. CAM plants have similar reactions, but separated in time, not space. This reduces water cost even more although often at the expense of overall rate. In this sub-objective, we aim to understand gene expression which underlies C4 and CAM CO2 accumulation mechanisms. The Cousins group (WA-AES) studies carbon uptake in relation to water use in C4 plants. Collaboration between Cousins and Allen (MO-ARS) labs investigated the biochemical flexibility of the CO2 concentrating mechanisms in the C4 plant maize. The Cousins lab measured and mapped contents of 13C and 18O in populations of three C4 species (recombinant inbred lines derived from two Setaria species; sorghum and maize bioenergy genomic panels) to identify genomic regions affecting the carbon water flux ratio as well as key leaf-level traits that influence whole plant resource use efficiency (e.g. water and nitrogen use efficiency). Effects of water status and planting density will be studied to find genes that are important for regulating the tradeoff between water usage and carbon uptake. The Cushman (NV-AES) lab investigated the introduction of CAM into model plants, such as Arabidopsis (73) with current efforts aimed at moving CAM into C3 photosynthesis crops, such as soybean. Current work is focused on engineering optimized versions of CAM into model and crop species alone and in combination with engineered tissue succulence, which is expected to optimize the functional performance of engineered CAM. The Yu lab (TX-AES) studies circadian regulation and regulatory network of CAM metabolic activities in pineapple (Ananas comosus (L.) Merr.), the most economically important tropical fruit crop utilizing CAM. This lab collaborated in sequencing the pineapple genome (82), identified and classified transcription factors and transcription coregulators and analyzed their tissue-specific and diurnal expression patterns (108).

3. Identify strategies to manipulate photosynthate partitioning. Assimilate flow constitutes the means and mechanisms of plant primary productivity. Regulation of hexose conversion to starch and sucrose can maintain optimum photosynthetic CO2 fixation activity. Knowledge of regulatory pathways of assimilate flow can guide crop improvement to enhance utilization of photosynthate for food, fuel, fiber and industrial feedstocks.

Collaborators focus on 3.1 Starch partitioning, 3.2 Lipid partitioning, 3.3 Antioxidant enhancement and 3.4 Fusion constructs for enhanced carbon partitioning. The first sub-objective addresses starch biosynthesis, degradation, and assimilate transport, the second sub-objective examines conversion of leaves into lipid storage units, the third sub-objective investigates a novel source of antioxidants, and the fourth sub-objective targets barriers to commercial development of novel organisms and useful bio-products.

3.1 Starch partitioning. Starch synthesis is an important target for manipulating source-sink relationships leading to increased genetic yield potential of crop plants (Ho, 1988, Turgeon 1989) and plant productivity. Comparisons of historically-important versus current wheat varieties revealed that flag leaf starch has increased as has wheat yield in cultivars released over the past 100 years. In this sub-objective, we aim to understand processes regulating starch biosynthesis and degradation in relation to assimilate transport. The Giroux lab (MT-AES) reported that leaf starch biosynthesis contributes to crop yield, as over expression of leaf starch biosynthesis in rice modified whole plant processes resulting in increased plant biomass (104). Two regulatory enzymes, ADP glucose pyrophosphorylase (AGPase) and phosphorylase I, control different phases of starch biosynthesis. AGPase catalyzes the first committed step in the starch biosynthetic (maturation) pathway and is subjected to allosteric regulation and redox control. The Okita lab (WA-AES) used a biochemical-genetic approach to provide insights into the roles of the large and small subunit structure of AGPase (12, 40, 48, 49, 50, 51, 52, 58). Okita’s group also evaluated the function of PHO1, a higher plant plastid-localized phosphorylase in starch biosynthesis. Transgenic rice lines expressing a pho1 variant form (Pho1ΔL80) lacking the L80 peptide grow faster and have higher productivity (biomass) and grain yields, the latter due to larger grain size. Recent results indicate that the construct Pho1ΔL80 interacts with the PsaC subunit of photosystem II and possibly controls electron flow to ferredoxin and NADPH production (56).

3.2. Lipid partitioning. Plants produce proteins, lipids, and carbohydrates. High concentrations of lipids are limited to seeds—particularly oilseeds. While seeds are produced during approximately one third of the plant life cycle, vegetative tissues such as leaves are available for lipid storage much of the time in plant development. Plants altered to produce significant storage reserves in leaves, such as lipids, have untapped potential for agricultural production. In this sub-objective we will assess the functional and productivity consequences of photosynthetic carbon partitioning to lipid and storage in leaves, converting plant leaves into lipid storage tissues.  The Allen lab (MO-ARS) investigates the spatial and temporal dynamics of central carbon metabolism in plant cells (7), developing comprehensive metabolic flux maps of resource allocation and carbon partitioning in oilseeds (9, 10) and in leaves (77). The methods provide subcellular description of metabolism using isotopes that label spatially distinct compounds according to the fluxes in different subcellular or cellular locations (2, 5, 6, 79). This is shown for lipid production in tobacco leaves, described using radioactivity and biomass measurement techniques (131). Preliminary investigations indicate that changes in metabolism are associated with chloroplast membranes in response to altered temperature. Since plants respond to temperature by altering acyl chain composition in the membranes, we hypothesize that acyl carrier proteins may be involved in acyl chain handling and storage in plastoglobuli.

3.3 Antioxidant enhancement. Reactive Oxygen Species (ROS), such as singlet oxygen, superoxide radical, hydroxyl radical, peroxides, and hydrogen peroxide, have toxic effects because they react with cell components including membranes, lipids, proteins, and nucleic acids. Their accumulation in plant cells can inhibit photosynthesis, resulting in loss of fitness and productivity. Improved ROS scavenging by antioxidant compounds protects photosynthesis and mitigates productivity losses. In this sub-objective we will install in crop plants a novel biosynthetic pathway leading to accumulation of a small antioxidant heterologous compound from anaerobic methane-producing archaea, which has been shown to enhance plant growth, when applied exogenously. Collaboration among the Roston, Glowacka, Buan and Stone labs (NE-ARD) examine effects of antioxidants on photosynthesis, with focus on an antioxidant from archaea. Preliminary data indicates that this compound may change non-photochemical quenching and carbon assimilation processes. Strategies for applications that confer growth benefits have been identified for Arabidopsis thaliana and three other species.

3.4 Fusion constructs for enhanced carbon partitioning. In synthetic biology, including the generation of agricultural bioproducts, yield of process often depends on the concentration of the pathway-catalyzing recombinant enzymes. However, heterologous proteins are almost always unwelcomed by the host cell and either pellet as inclusion bodies or are degraded by the cell—a barrier to the meaningful application of synthetic biology approaches in plants and algae. Another barrier has been the very low-level accumulation of recombinant enzymes, often compounding slow Kcat (Vmax) of individual steps, leading to low yield of products. This sub-objective will deliver a platform for substantially enhanced carbon partitioning toward a target biosynthetic pathway, via enzyme over-expression. This would alleviate rate- and yield-limiting catalytic steps in the generation of endogenous compounds, e.g., starch and lipid, as well as heterologous specialty and commodity products, including agricultural bioproducts, via the process of photosynthesis. The Melis lab (CA-AES) has examined production of high-value compounds and has applied transformation technologies in the model cyanobacterium Synechocystis for the heterologous production of monoterpene (β-phellandrene) hydrocarbons using genes and pathways from lavender and tomato. Novel fusion constructs generated an average of 10 mg product g-1 dry cell weight (dcw) compared with the 0.01 mg g-1 dcw measured with low-expressing constructs (i.e., a 1000-fold yield improvement). The terpene synthase fusion-protein approach is promising, as it enhanced rates and yield of β-phellandrene hydrocarbons production in these model photosynthetic microorganisms.

4. Develop strategies to overcome limitations to photosynthetic productivity caused by developmental and environmental factors. Collaborators investigate developmental and environmental limitations to photosynthesis. These studies include stress physiology (heat, salt, drought, cold), as well as the underlying mechanisms that signal plants to respond to stress. Cultivars differ in photosynthetic efficiency related to light interception and reflectance, leaf color, leaf rolling, leaf cuticle thickness, leaf wax content, and presence of awns in spikes (15, 101). The responses of plants to abiotic stresses are communicated at the cellular and molecular levels through a network of signaling pathways, often involving protein kinases and sugar responsive proteins. Knowledge of stress responses and stress signaling systems can integrate knowledge regarding photosynthetic membrane, CO2 capture and regulation of assimilate flow into commercial crop improvement, adaptive crop management, and utilization of novel organisms to generate industrial feedstocks. Collaborators focus on four major sub-objectives: 4.1 Heat stress responses, 4.2 Stomatal behavior, 4.3 Stress signal transduction and 4.4 Critical loci encoding stress responses. The first sub-objective addresses heat stress effects on photosynthetic and seed filling processes; the second examines stomatal behavior from evolutionary, novel trait and novel crop perspectives; stress signal transduction involving Calcium-dependent Protein Kinases (CPKs), inositol pyrophosphate and a novel peptide comprise the third sub-objective; the fourth seeks to identify genetic loci associated with regulation of stress responses and developmental processes.

4.1 Heat stress responses. Heat stress can impair enzyme function, alter membrane composition, limit pollination efficacy and diminish assimilate flow to developing seeds. In this sub-objective we aim to understand effects of high temperature on C3 and C4 photosynthetic physiology on assimilate flow to developing seeds. The Zhang lab (MO-AES) characterized C4 model plant Setaria viridis responses to high light or high temperatures at photosynthetic, ultrastructural, and transcriptomic levels and identified gene candidates for improving high light or high temperature tolerance in C4 crops (9). Genes and pathways with potential roles for heat stress tolerance in the model green alga Chlamydomonas reinhardtii were discovered following -omic analysis of cell physiologies, transcriptome, and proteome (130). Jagadish and Prasad investigated responses of photosynthesis, respiration and floret fertility in multiple field crops to heat stress conditions. The Aiken and Prasad groups have shown canopy temperature and chlorophyll fluorescence as potential drought screening tools and elucidated slow wilting trait in sorghum (87). The Jagadish lab found that dark respiration (Rn) response to high night temperature was greatest during the post flowering phase (11) with up to 5% yield reduction per 1oC increase in night-time temperatures. The increased Rn corresponded with reduced phloem unloading to sink tissue (reduced cell wall invertase activity), slower cell expansion (reduced vacuolar invertase activity), and limited substrate supply for starch synthesis (reduced starch synthase activity). Heat stress thresholds, established for wheat, are used to identify sources of heat stress tolerance during grain filling. 

4.2 Stomatal behavior. Regulation of stomatal opening, in leaves, is an essential determinant of light and water utilization. In this sub-objective, we aim to uncover evolutionary trails to high photosynthetic rates among Angiosperms, effects of novel traits on photosynthetic processes and primary productivity of a novel crop. The McAdam lab (IN-AES) reported macro-evolutionary changes in stomatal responses to environmental signals (80, 121).  The group evaluates the evolution of large stomatal apertures in relation to the high photosynthetic rates that evolved among Angiosperms. Complementary investigations address the evolution of hormonal signaling of drought stress in guard cells. Collaboration with the Zhang lab (MO-AES) focused on abscisic acid and stomata closure in response to drought and heat stress in Setaria (9). The Glowacka lab (NE-ARD) investigates a photosynthesis-related protein of photosystem II subunit S (PsbS) that suppresses stomatal opening while maintaining CO2 uptake; transpiration ratio decreased by 25% (38). Tobacco transformed with the PsbS trait showed increased photosynthetic capacity (increased maximum carboxylation) and up to 20% greater biomass under limiting field conditions. Since PsbS stimulates the thermal dissipation of excitation energy (observed as NPQ; 70, 85, 78), we hypothesize that increased expression of PsbS decreases the chloroplast-derived signal for stomatal opening in response to light and decreases water loss at the leaf level (17). The Cushman lab (NV-AES) develops novel strategies to improve drought and salinity tolerance. A key project focuses on enhancing tissue succulence through the overexpression of a member of helix-loop-helix transcription factor that improved biomass production (72) as well as salinity and water-deficit stress tolerance (74). Collaboration with the Harper lab (NV-AES) to increase biomass and improve water use efficiency of crop plants. The Cushman lab engineered tissue succulence and CAM metabolism into Arabidopsis thaliana.  The Harper lab (NV-AES) investigates use of stabilized oil bodies to improve photosynthate capture.  Collaborative goal is to determine if the combination of these strategies can yield a synergistic effect. Additional work develops use of highly productive CAM crop species, such cactus pear, as food, feed, and biofuel feedstocks (90).

4.3 Stress signal transduction. Regulatory processes are ‘triggered’ by developmental cues and sensory mechanisms that activate signal transduction systems. In this sub-objective we aim to improve understanding of roles of CPK, inositol pyrophosphates and dehydration-stimulated peptides in regulation of photosynthetic processes. The Harper group (NV-AES) collaborates with the Cushman group (NV-AES) to investigate the regulation of subcellular organization, signaling and metabolism with emphasis on those that require phosphorylation of a binding site on the client. Many of these phospho-interactions are thought to be mediated by CPKs (16). Phospho-proteomic analyses in the lab have identified multiple heat-stress dependent changes in phosphorylation, with candidate targets including proteins involved in the secretory pathway, transcription and translation, and central carbon metabolism (Harper, unpublished). The group evaluates the importance of an auto-inactivation mechanism in planta and how this impacts the ability of CPK34 encoding transgenes to rescue the near sterile phenotype of a cpk17/34 knockout. The Gillaspy lab (VA-AES) delineated key enzymes in the inositol pyrophosphate signaling pathway (23). The genes that encode these enzymes provide key insights into how plants produce these molecules (1). We have also transferred a trait for increased phosphorous acquisition to a cover crop species. The Li lab (MS-AES) investigated effects of drought and salinity on crop photosynthetic productivity (103, 76). Silicate application enhanced the protein abundance of NADPH-generating enzymes and detoxifying enzymes under water deficit stress; a result attributed to improved growth and photosynthesis of soybean plants grown under water limiting conditions. Recently, gene-encoded small peptides have emerged as regulatory signaling molecules involved in the control of plant growth and development (91). The group recently isolated two dehydration-stimulated peptides from rice plants subjected to water deficit and will investigate regulatory roles of the peptides for photosynthetic responses of rice to water limiting conditions.

4.4 Critical loci encoding stress responses. In this sub-objective we aim to identify critical loci in soybean and sorghum genome associated with control of photosynthetic processes. The Fritschi lab (MO-AES) identified genomic loci for photosynthetic gas exchange traits based on genome wide association studies (GWAS, 43), and QTL mapping (ongoing).  Photosynthetic responses to elevated temperature featured contrasting genotypes identified from the GWAS (44). Ongoing efforts explore photosynthetic characteristics in soybean genotypes differing in drought tolerance related traits (canopy wilting, canopy temperature, and carbon isotope discrimination). The Xin lab (TX-ARS) developed sorghum mutant populations with multiple distinctive traits. The sorghum mutant library will be expanded by deep-sequencing 1,000 additional sorghum mutant lines. The focus of current work is use of the mutant population to dissect control points of photosynthetic processes.

Objectives

  1. Identify strategies to optimize the assembly and function of the photosynthetic membrane.
  2. Identify strategies to modify biochemical and regulatory factors that impact the photosynthetic capture and photorespiratory release of CO2.
  3. Identify strategies to manipulate photosynthate partitioning.
  4. Develop strategies to overcome limitations to photosynthetic productivity caused by developmental and environmental factors.

Methods

Methods

1. Identify strategies to optimize the assembly and function of the photosynthetic membrane. The scope of Objective 1 includes investigation of the basic principles driving photosynthetic efficiency determined by compositional and architectural dynamics of the photosynthetic membrane. Thus, all investigators in this category use common molecular-biology resources, including Arabidopsis thaliana mutants, a subset of which have been generated in the Benning (MI-ABR), Roston and Stone labs (NE-ARD) in part by collaboration, DNA and protein manipulation (e.g., electrophoresis, immunoblotting, cloning, native and heterological expression, enzyme assays), biological fractionations to the sub-organellar level, as well as in vivo spectroscopy and electron microscopy. Created resources can thus be easily shared throughout the group. In addition, each laboratory adds specialty expertise required for their investigations, as follows:

1.1 Dynamics of chloroplast membrane architecture. The focus is on critical arrangements of chloroplast membranes enhancing photosynthetic energy conversion. Identifying functional consequences of structural alterations is being explored by the Kirchhoff lab (WA-AES) by coupling ultrastructural techniques with functional and compositional analyses. In detail, in vivo difference absorption spectroscopy and fluorescence spectroscopy, different sets of biochemical methods (e.g. membrane, sub-membrane, protein and lipid isolation, gel electrophoresis, BN-PAGE electrophoresis, thin-layer chromatography, gas chromatography, proteoliposomes technology), high-pressure freezing, electron- and fluorescence-microscopy, coarse grain computer simulations, image analysis, and mathematical modeling allows profound analysis of structure-function dynamics under different environmental conditions and in selected mutants. This work features the integration of experimental data into a dynamic coarse-grain computer model that will describe thylakoid dynamics and its functional consequences. The Roston lab (NE-ARD) will use multiple fluorescent markers of thylakoid/inner envelope membrane contact sites to investigate the condition resulting in changed inner envelope and thylakoid membrane connectivity. The fluorescent marker systems will be paired with traditional microscopy to identify stress conditions resulting in increased contact sites. Candidate thylakoid/inner envelope membrane contact site proteins have been identified using traditional fractionation techniques and predictions of chloroplast homologs of membrane contact site proteins. The role of these proteins in chloroplast contact sites will be investigated by co-localizing them with fluorescent reporters and confirmed, using traditional fractionation/electron microscopy techniques. These approaches allow in-depth functional characterization of the photosynthetic apparatus, including the organization of protein complexes within the membrane.

 1.2 Chloroplast membrane lipid and protein dynamics. The focus is on critical membrane lipid and protein changes required for efficient photosynthetic membranes, that are yet poorly understood. The Benning lab (MI-ABR) will investigate the role of the regulatory protein RBL10 in the regulation of lipid metabolism at the inner chloroplast envelope membrane. Characterizing its proteolytic activity and its potential substrates and identifying proteins interacting with RBL10, which is found in a large complex, will be key to a mechanistic understanding of the role of this rhomboid protease in chloroplast lipid metabolism. The Benning Lab will also investigate the identity, processing, topology and function of PA phosphatases associated with the chloroplast envelope membranes. The group will determine the function of the thylakoid-specific lipid 16:1Δ3t PG and its metabolism in response to the oxidative state of the chloroplast. Arabidopsis lipid mutants will be analyzed by photosynthetic phenotyping to determine effects of thylakoid metabolism on photosynthesis, in collaboration with T. Sharkey and D. Kramer (MI-ABR). The Roston lab (NE-AES) is investigating the role(s) of unusual chloroplast lipids, oligogalactolipids. Under freezing stress, oligogalactolipids are accumulated, are required for survival, and yet their role in the chloroplast remains unclear. The Roston lab will also couple a series of chloroplast lipid mutants developed in collaboration with the Benning lab, with detailed lipid profiling and precision environmental chambers to determine the function of the oligogalactolipids, and to identify alternate strategies to stabilize chloroplast membranes.

2. Identify strategies to modify biochemical and regulatory factors that affect the photosynthetic capture and photorespiratory release of CO2. The scope of Objective 2 includes flow of carbon through the Calvin-Benson cycle and gene-expression associated with C4 and CAM CO2 accumulating mechanisms. In this objective, common molecular biology resources will be used to modify the metabolic pathways involved in carbon capturing and metabolism. In addition, specialty resources will be used to develop and apply synthetic biology techniques and to measure and simulate changes in rates of Rubisco activity, photosynthetic electron transport and rates of carbon assimilation.

2.1 Electron transport and regulation of the Calvin-Benson cycle. The focus is on the interaction of carbon metabolism and cyclic electron flow. The Sharkey lab (MI-ABR) investigates the oxidative branch of the pentose phosphate pathway (glucose-6-phosphate/G6P shunt). Collaborative work by the Sharkey and Walker labs will use 13CO2 labeling, genetically modified plants (especially loss of function mutants) and metabolite measurements to determine the amount and degree of label of critical metabolites in specific plant cell compartments, using non-aqueous fractionation of metabolites. The group will also investigate a cytosolic bypass that allows G6P to reenter the chloroplast e.g., in plants lacking the photorespiratory enzyme hydroxypyruvate reductase (68). The Walker lab (MI-ABR) will continue physiological and biochemical interrogation of Rhyza stricta to determine the adaptive mechanisms used to process high rates of photorespiratory flux at elevated temperatures using gas exchange, online carbon isotope discrimination, and the temperature response of photorespiratory enzymes. Additional metabolic flux experiments will help determine how photorespiratory flux changes under different photorespiratory pressures in various species. These efforts will benefit from discussions with Tom Sharkey and Doug Allen (MO-ARS).

2.2 C4 and CAM CO2 concentrating mechanisms. The focus is on the full spectrum of gene expression changes that are associated with the C3 to C4 and CAM evolutionary progression. The Cousins lab (WA-AES) uses leaf carbon isotope composition (δ13C) to determine how water use efficiency differs in two C4 species of Setaria species and in Sorghum mapping populations. This research helps to better understand the relationship of leaf carbon isotopes and the influence photosynthetic water use efficiency to whole plant water use efficiency.  Collaborative with the Jagadish lab (KS-AES) will expand this work to test the impact of nighttime temperatures on maize. The Cushman lab (NV-AES) will continue to develop synthetic biology approaches to assemble large gene circuits to identify the most efficient versions of CAM through Gibson assembly, vector development, and stable plant transformation. Outcomes include generation of optimized synthetic carboxylation, decarboxylation, starch degradation, and complete CAM gene circuits. The resulting plants will be evaluated for improved growth, productivity, water-use efficiency, and tolerance of water-deficit and salinity. Empirical testing will be accompanied by detailed, genome-scale transcriptomic and metabolome profiling and diel flux balance analysis modeling to corroborate energetic efficiency predictions for each iteration of the synthetic CAM gene circuits. Plants expressing optimized CAM gene circuits will be evaluated with and without engineered tissue succulence in Arabidopsis and soybean in collaboration with the Harper lab (NV-AES). The Yu lab (TX-AES) will conduct comparative genomics and evolutionary genomics analysis investigate the molecular changes linked to C4 and CAM photosynthesis evolution. DNase-Seq will be used to identify a set of cis-elements that are involved in regulation of C4 and CAM photosynthesis. DAP-seq will be used for genome-wide identification and comparative analysis of binding sites of circadian oscillators in C3, C4, and CAM plants. These studies will also explore genome-wide identification of regulatory elements for candidate transcription factors that are involved in transcriptional regulation of C4 and CAM metabolic activities. Virus-induced gene silencing system, in combination with RNA-seq, will provide a functional validation of key regulatory elements of CAM photosynthesis.

3. Identify strategies to manipulate photosynthate partitioning. The scope of Objective 3 includes investigation into how photosynthate (fixed carbon from photosynthesis) partitioning impacts metabolism, growth, and synthesis of novel compounds. In this objective, common molecular-biology resources, including generated plasmids along with Arabidopsis thaliana mutants, Chlamydomonas and Synechocystis transformants, as well as rice and wheat transgenic plants, will be analyzed with regards to metabolite levels, including terpenes starch and lipids. Mass spectrometry, isotopic labeling studies, and enzyme assays will be used to discern regulatory properties of photosynthate portioning.

3.1 Starch partitioning. The focus is on processes regulating starch biosynthesis and degradation in relation to assimilate transport. The Giroux lab (MT-AES) will search for genes (and genetic markers) impacting leaf starch levels by applying QTL analysis to recombinant inbred line populations in wheat.  Near isogenic lines (NILs) of wheat that vary in leaf starch will be developed and used in field and greenhouse studies to determine the impact of leaf starch variation on photosynthetic rates and plant productivity. The Okita lab will investigate the role of PHO1 in starch partitioning via CRISPR-Cas12a approaches to identify specific segments of the L80 peptide, harboring cis-regulatory sequences, deletion of which enhances plant growth rates and grain yields. Candidate rice lines will be studied in more detail by gas exchange and photosynthetic fluorescence spectroscopy to assess PSI properties.  In vitro PSI assays using isolated thylakoid membranes will also be employed to see if Pho1 and Pho1ΔL80 affect the PSI redox properties.

3.2 Lipid partitioning. The focus is on consequences of converting plant leaves into lipid storage tissues. The Allen lab (MO-ARS) will assess central carbon metabolism and flux in oilseeds and leaves, primarily seeds of soybean and Camelina, and leaves of tobacco, soybean and other sources as applicable using metabolite measurements with mass spectrometry and isotopic labeling studies (89, 57, 10, 8). Such studies will provide insight specifically on how membrane composition mechanistically accounts for the exchange of acyl groups, and also describe the implications on central metabolism that result in tradeoffs between starch, sucrose, and lipid.

3.3 Antioxidant enhancement. The focus is on effects of archaeal antioxidants on photosynthesis. The Roston, Glowacka, Buan and Stone labs (UN-ARD) will use biochemical and genetic techniques to investigate whether archaeal thiol redox molecules and enzymes can be used to increase plant photosynthetic efficiency and abiotic stress responses. To determine effects on photosynthesis, plant measurements will include plant size, photosynthesis parameters Fv/Fm, photochemical quenching, non-photochemical quenching, carbon assimilation, transpiration, and chlorophyll content. We also use genetically encoded redox sensors and multiple –omics technology to identify non-photosynthetic effects. Future work will focus on how the antioxidant affects non-photosynthetic quenching (NPQ) and carbon assimilation using mutants, genetically encoded sensors and plants engineered to produce archaeal antioxidants.

3.4 Fusion constructs. The focus is on conversion of carbon fixed from photosynthesis into high-value products. The Melis lab (CA-AES) will work to enhance photosynthate partitioning toward specific terpene hydrocarbons and related plant essential oils. This will be implemented with model cyanobacterial (Synechocystis) and microalgal (Chlamydomonas) strains. Recombinant DNA constructs, and culturing conditions were described (20, 21, 31, 32, 33). Fusion constructs with varying orientation of the leader versus the trailer moieties will be applied and examined for stability and activity. Genomic DNA PCR analysis will be employed to test for plasmid insertion in the DNA of the recipient strains. RT-qPCR analysis will be employed to assay for transgene transcription and message abundance/stability. SDS-PAGE and quantitative Western blot analysis will test for level(s) of transgenic protein expression. Isolation of different “fusion construct” enzymes from their in vivo state via His-tagged elution will be employed to test for possible association with other non-fusion proteins that might confer stability to the heterologous protein. Measurements of photoautotrophic growth and product generation will test for transformant cell fitness and its ability to generate the desirable heterologous high-value product.

4. Develop strategies to overcome limitations to photosynthetic productivity caused by developmental and environmental factors. The scope of Objective 4 encompasses stress response and stress tolerance with particular attention to light utilization, water deficit and high temperature conditions. Investigations range from signal transduction in model organisms, mapping populations of diversity panels and bi-parental genetic combinations, effects of novel traits to the evolution of adaptive stomatal behavior. The linkage of gene-expression to adaptive responses underlies experimental approaches, under field and controlled conditions. Studies will utilize genomic analysis, loss of function mutants, and metabolite analysis to identify factors regulating photosynthetic and photosynthate partitioning responses to abiotic stresses such as heat, water deficits, excess salinity and phosphorus deficiencies/toxicity.  In addition, each lab adds specialty expertise required for their investigations.

4.1 Heat stress responses. The focus is on responses to heat stress, which affects multiple components of primary productivity. The Jagadish lab (KS-AES) will collaborate with the Prasad lab (KS-AES) to investigate genetic factors regulating dark respiration responses to increased night temperature in wheat and corn. A novel cyber-physical system maintains controlled night-time temperature differentials at field scale. Samples collected from a large-scale wheat association mapping panel will be processed and analyzed using genome-wide association study procedures. Protocols established for wheat will be applied to corn hybrids to identify genetic factors affecting responses to high night temperature during grain filling. The Zhang lab (MO-AES) will continue studies of photosynthetic responses to abiotic stresses, especially high temperatures, under controlled conditions. Experimental methods include photosynthetic measurements (gas exchange, spectroscopic measurements), -omics level analysis (transcriptomes and proteomes), and genetics (gene functional analysis) in both the C4 model plant Setaria viridis and the model green alga Chlamydomonas reinhardtii.

4.2 Stomatal behavior. The focus is on stomatal behavior in relation to primary productivity. The McAdam lab (IN-AES) will investigate the evolution of stomata in angiosperms and mechanical interactions with the epidermis. Comparative studies of stomatal mechanics and guard cell wall properties will focus on representative species that span the phylogeny of land plants. Corresponding studies will investigate the evolution of ABA responses to unfavorable photosynthetic conditions. The Glowacka lab (NE-ARD) will continue investigations with transformed tobacco to determine the effects of PsbS overexpression on seedling survival, growth and photosynthetic parameters such as stomata opening, NPQ, the light efficiency of photosynthesis and carboxylation capacity of RuBisCO. Tests will be conducted on homozygous T2 generations under different strengths of drought stress. The Cushman lab (NV-AES) will continue to perform field trials on highly productive CAM species as bioenergy feedstocks having significantly lower water requirements than current bioenergy feedstocks. These studies will identify highly productive cactus pear (Opuntia spp.) accessions as well as evaluate life cycle and life cycle costing analyses.

4.3 Stress signal transduction. The focus is on signal pathways involving CPKs, inositol pyrophosphate and novel peptides. The Harper lab (NV-AES) will continue to use molecular genetic and mass-spec proteomic approaches to investigate the role of CPKs in phospho-regulation of plant development and metabolism, with specific attention to stress response pathways that involve phospho-regulation of protein-protein interactions.  The purpose is to identify phosphorylation events that are associated with abiotic stress responses.  This project will involve collaborations with the Cushman lab (NV-AES) and is expected to lead to interactions on lipid signaling with the Gillaspy lab (VA-AES). The Gillaspy lab will develop applications for the inositol pyrophosphate signaling pathway. Genes encoding enzymes involved in inositol pyrophosphate synthesis or turnover will be transferred to Pennycress, a cover crop species, using an inducible promoter system. CRISPR will be used to remove a gene that likely increases phosphate use efficiency in a model plant species. Phosphate uptake, accumulation, and phosphate starvation response gene expression changes will be measured for both gain- and loss-of-function plants. Inositol pyrophosphates will be measured using radiolabeling followed by HPLC separation and quantification. We will examine the impact of elevating phosphate uptake on photosynthetic parameters. The Li lab (MS-AES) will investigate roles of two novel dehydration-stimulated peptides in regulating rice responses and affecting photosynthetic productivity under water limiting condition. They will determine the peptide amino acid sequences (mass spectrometry and Edman degradation), establishing the precursors of the peptides isolated. Overexpression and underexpression constructs in rice will support functional studies of peptide effects on growth and photosynthesis under drought condition. Agronomic responses to drought treatments will include plant height, leaf numbers, tiller numbers, tillers with panicles, and photosynthesis parameters.

4.4 Critical loci encoding stress responses. The focus is on associating gene expression with photosynthetic processes. The Fritschi lab (MO-AES) will use diversity panels and bi-parental mapping populations to investigate the genetics underlying photosynthetic light and carbon reactions of soybean genotypes. Genotypes selected as parents for these mapping populations will be grown under abiotic stress conditions (manipulated field as well as controlled environment); chlorophyll fluorescence and gas exchange techniques will be used to characterize their responses. The Xin lab (TX-ARS) will continue to develop a mutant library for the sorghum cultivar BTX623. Bioinformatic approaches, such as MutMap, will be used to identify causal mutations affecting photosynthesis, providing targets for genome editing and physiological analysis.

Measurement of Progress and Results

Outputs

  • Collectively, the research will produce data related to thylakoid membrane architecture; organic acid and lipid metabolism; C4 and CAM CO2-concentrating mechanisms; photosynthate partitioning; central carbon metabolism; photosynthetic responses to heat, chilling, and drought stress, and stress-dependent phosphorylation sites.
  • Information developed by the project will pertain to thylakoid-inner envelope membrane connectivity, genetic structure of CAM biomass feedstocks, regulatory networks of C3, C4 and CAM photosynthesis, novel strategies to enhance water use efficiency, evolution of CAM and ABA physiology, optimized synthetic versions of CAM and synergies with tissue succulence, yield impacts of modified starch biosynthesis, triacylglycerol or terpene accumulation, photo-protective effects of a novel archaeal antioxidant, stress responses in wheat and soybean, role of CPKs in phospho-regulation of stress responses, genomic regions affecting leaf starch levels in wheat, photosynthetic responses to high temperature, tolerance of high night temperatures, stomatal responses to PsbS protein overexpression, photosynthetic responses to two novel drought signaling peptides and to genes regulating phosphate uptake.
  • Biological materials produced by the project will include a variety of DNA plasmids expressing photosynthetically relevant genes, large gene circuits for synthetic versions of CAM, rice plasmids with modified starch biosynthesis, NILs for wheat varying in leaf starch, gene fusion constructs and synthetic operon configuration permitting high levels of heterologous protein expression, sources of high night temperature tolerance in wheat and heat tolerance in soybean, pennycress cover crops with enhanced phosphate uptake, a novel dehydration-induced peptide in rice, annotated genetic sequences of functional sorghum mutations that affect photosynthetic processes.

Outcomes or Projected Impacts

  • Knowledge of photosynthetic machinery responses to environmental dynamics and endogenous regulatory controls will guide improvements in crop plants and biofuel prospects for growth in specific regional climates.
  • Effective regulation of stomata and starch biosynthesis could increase CO2 assimilation over the growing season, with corresponding yield increase.
  • Insights into algal cell growth and photosynthesis, and expression of transgenes can support engineering and industrial production of novel crops generating biofuels and high-value products such as terpene hydrocarbons.
  • Understanding how signaling systems function can support strategies to engineer plants with improved stress tolerance, sustaining agricultural productivity.
  • Synthetic CAM gene circuits can be applied widely to other food, feed, fiber, and biofuel crops to improve their productivity, reduce photorespiration, improve water-use efficiency and drought/salinity stress tolerance under the hotter and drier environments of the future.
  • Developing warm night temperature tolerant cereals including wheat and corn will help sustain global food and nutritional security under predicted warmer climate.

Milestones

(2023):DNA plasmids expressing photosynthetically relevant genes and gene fusion constructs will be developed (1, 2, 3, 4; CA-AES, MI-ABR, MS-AES, NE-AES, NV-AES, VA-AES, TX-AES, WA-AES). Photosynthetic and stress responses to archaeal thiol redox molecules and enzymes will be tested (3.3, UN-ARD). A novel cyber-physical system controlling night-time temperature differentials will support genetic mapping of heat tolerance in wheat (4.1, KS-AES). Genes encoding enzymes involved in inositol pyrophosphate synthesis or turnover will be transferred to Pennycress, a cover crop species, using an inducible promoter system (4.3, VA-AES). Bioinformatic approaches will help identify causal mutations affecting photosynthesis in sorghum (4.4, TX-ARS).

(2024):Functional analysis of the thylakoid-specific lipid 16:13t PG and its metabolism will be determined in response to the oxidative state of the chloroplast (1.2, MI-ABR). The molecular changes linked to the evolution of C4 and CAM photosynthesis will use comparative genomics and evolutionary genomics analysis (2.4, TX-AES). Central carbon metabolism and flux in oilseeds and leaves (i.e., soybean, Camelina, and tobacco) will use metabolite measurements, mass spectrometry, and isotopic labeling (3.2, MO-ARS). Photosynthetic responses to high temperature stress will include photosynthetic measurements, -omics level analysis, and gene functional analysis in the C4 plant Setaria viridis and the C3 green alga Chlamydomonas reinhardtii (4.1, MO-Danforth).

(2025):The role(s) of unusual chloroplast lipids, oligogalactolipids, in membrane stability will be investigated in chloroplast lipid mutants using detailed lipid profiling and precision environmental chambers (1.2, NE-ARD, MI-ABR). Warm night temperature effects on respiration and grain fill in corn will be investigated (2.2, WA-AES, KS-AES). Genes (and genetic markers) impacting leaf starch levels will be investigated by applying QTL analysis to recombinant inbred line populations in wheat (3.1, MT-AES). Fusion constructs (in model cyanobacterial Synechocystis and microalgal Chlamydomonas strains) with varying orientation of the leader versus the trailer moieties will be applied and examined for stability and activity (3.4, CA-AES). Effects of the PSII subunit S protein on stomatal closure, photosynthetic parameters and plant development (tobacco) will be investigated (4.2, UN-ARD).

(2026):Functional consequences of thylakoid structural dynamics will integrate experimental data into a dynamic coarse-grain computer model, describing thylakoid dynamics and its functional consequences (1.1, WA-AES). Metabolic flux experiments will help determine how photorespiratory flux changes under different photorespiratory pressures (2.2, MI-ABR, MO-ARS). Modified starch biosynthesis, related to PHO1, will be investigated using gas exchange and photosynthetic fluorescence spectroscopy (3.1, WA-AES). The role of calcium-dependent protein kinases (CPKs) in stress-response pathways will be investigated using molecular genetic and mass-spec proteomic approaches (4.3, NV-AES, VA-AES). Overexpression and underexpression constructs of two novel dehydration-stimulated peptides in rice will support functional studies of peptide effects on growth and photosynthesis under drought conditions (4.3, MS-AES).

(2027):Synthetic biology approaches to assemble large gene circuits will investigate the most efficient versions of CAM through Gibson assembly, vector development, and stable plant transformation (2.2, NV-AES). The evolution of stomata in angiosperms will use comparative studies of stomatal mechanics and guard cell wall properties of representative species that span the phylogeny of land plants (4.2, IN-AES). Genetic studies of factors underlying photosynthetic light and carbon reactions of soybean genotypes will use gas exchange and chlorophyll fluorescence in diversity panels and bi-parental mapping populations (4.4, MO-AES).

Projected Participation

View Appendix E: Participation

Outreach Plan

The major advances and discoveries of the proposed research will be published in scientific journals and meeting proceedings. Peer-reviewed publication is the best practical method for evaluating the quality and impact of the research results. NC-1200 investigators have been successful in this endeavor, as illustrated by the large number of articles published in high impact, high quality journals (see Appendix). Research results will also be presented as platform lectures and poster presentations at local, regional, national, and international scientific meetings. Members will be encouraged to highlight NC-1200 contributions, utilizing professional social media sites such as professional web-pages and ‘projects’ listed in science networks such as ResearchGate. Annual reports will be shared with regional committees such as NE-9/S-009/W6/NC7 (Conservation and Utilization of Plant Genetic Resources) and NC-1203 (Lipids in Plants). In addition, NC-1200 members will seek opportunities to engage undergraduate students in research with programs such as the MSU annual summer intern program in plant genomics (www.plantgenomics.msu.edu).

Organization/Governance

The Standard Governance for multistate research activities will be implemented, which includes include the election of a Chair (organizes current annual meeting), a Chair-elect (organizes next annual meeting), and a Secretary (organizes the subsequent annual meeting). Rotating through these functions all officers are elected for at least two-year terms to provide continuity. Administrative guidance will be provided by an assigned Administrative Advisor and a CSREES Representative.

Literature Cited

1.          Adepoju, O., Williams, S. P., Craige, B., Cridland, C.A., Sharp, A., Brown, A., Land, E., Perera, I.Y., Mena, D., Sobrado, P. and Gillaspy, G. Inositol Trisphosphate Kinase and Diphosphoinositol Pentakisphosphate Kinase Enzymes Constitute the Inositol Pyrophosphate Synthesis Pathway in Plants. BioRxiv, doi: https://doi.org/10.1101/724914

2.          Allen, DK, RW LaClair, JB Ohlrogge, and Y Shachar-Hill. 2012. Isotope labeling of RuBisCO subunits provides in vivo information on subcellular biosynthesis and exchange of amino acids between compartments. Plant Cell and Environment 35(7):1232-1244

3.           Allen, DK, JB Ohlrogge, and Y Shachar-Hill. 2009. The role of light in soybean seed filling metabolism. Plant Journal 58, 220-234

4.           Allen, DK, and JD Young.  2013. Carbon and nitrogen provisions alter the metabolic flux in developing soybean embryos. Plant Physiology 161:1458-1475

5.           Allen, DK, J Goldford, J Gierse, D Mandy, C Diepenbrock, and IGL Libourel. 2014a. Quantification of peptide m/z distributions from 13C-labeled cultures with high resolution mass spectrometry. Analytical Chemistry 86:1894-1901

6.           Allen, DK, BS Evans, and IGL Libourel. 2014b. Quantification of isotopic labeling in fragmented peptides from tandem mass spectra. Plos One 9:e91537

7.           Allen, DK. 2016. Quantifying plant phenotypes with metabolic flux and isotopic labelling. Current Opinion in Biotechnology 37:45-52

8.           Allen, DK, JD Young 2020. Tracing metabolic flux through time and space with isotope labeling experiments. Current Opinion in Biotechnology 64:92-100.

9.           Anderson, C.M., Mattoon, E.M., Zhang, N. et al. 2021. High light and temperature reduce photosynthetic efficiency through different mechanisms in the C4 model Setaria viridis. Commun Biol 4, 1092. https://doi.org/10.1038/s42003-021-02576-2

10.         AuBuchon-Elder, T, V Coneva, DM. Goad, LM Jenkins, DK Allen, EA Kellogg. 2020. Sterile spikelets contribute to yield in sorghum and related grasses. Plant Cell 32:3500-3518

11.         Bahuguna, R.N., C.A. Solis, W. Shi and K.S.V. Jagadish. 2016. Post-flowering night respiration and altered sink activity account for high night temperature-induced grain yield and quality loss in rice (Oryza sativa L.). Physiol. Plant. doi:10.1111/ppl.12485

12.         Ballicora, M. A., M. J. Laughlin, Y. Fu, T. W. Okita and others. 1995. Adenosine 5'-diphosphate-glucose pyrophosphorylase from potato tuber: significance of the N-terminus of the small subunit for catalytic properties and heat stability. 109:245-251

13.         Barnes AC, Elowsky CG, Roston RL. 2019. An Arabidopsis protoplast isolation method reduces cytosolic acidification and activation of the chloroplast stress sensor SENSITIVE TO FREEZING 2. Plant signaling & behavior. 14:9, DOI: 10.1080/15592324.2019.1629270

14.         Barnes AC, Benning C, & Roston R. 2016. Chloroplast membrane remodeling during freezing stress is accompanied by cytoplasmic acidification activating SENSITIVE TO FREEZING 2. Plant Physiol. 171(3):2140-2149.

15.         Blum A. 1988. Plant breeding for stress environment. CRC Press Inc., Boca Raton, Florida, USA.

16.               Boursiac, Y., S. M. Lee, S. Romanowsky, R. Blank and others. 2010. Disruption of the vacuolar calcium-ATPases in Arabidopsis results in the activation of a salicylic acid-dependent programmed cell death pathway. Plant Physiol 154:1158-1171

17.         Busch, F. A. 2014. Opinion: The red-light response of stomatal movement is sensed by the redox state of the photosynthetic electron transport chain. Photosynthesis Research, 119(1–2), 131–140. https://doi.org/10.1007/s11120-013-9805-6

18.         Carde JP, Joyard J, & Douce R. 1982. Electron-microscopic studies of envelope membranes from spinach plastids. Biol. Cell 44(3):315.

19.         Charuvi D,V. Kiss, R. Nevo, E. Shimoni, Z. Adam, and Z. Reich. 2012. Gain and loss of photosynthetic membranes during plastid differentiation in the shoot apex of Arabidopsis. Plant Cell 24(3):1143.

20.         Chaves, J.E., H. Kirst, A. Melis. 2015. Isoprene production in Synechocystis under alkaline and saline growth conditions. J Appl Phycol 27:1089-1097.

21.         Chen H-C, Melis A. 2013. Marker-free genetic engineering of the chloroplast in the green microalga Chlamydomonas reinhardtii. Plant Biotech J. 11: 818–828; DOI: 10.1111/pbi.12073

22.         Cooper, M., C. Gho, R. Leafgren, T. et al..  2014. Breeding drought-tolerant maize hybrids for the US corn-belt: discovery to product. J. Exp. Bot. 65, 6191-6204

23.         Cridland, C. and Gillaspy, G. Inositol Pyrophosphate Pathways and Mechanisms: What Can We Learn from Plants? Molecules 25(12):2789. doi: 10.3390/molecules25122789.

24.         Cruz, J.A., L.J. Savage, R. Zegarac et al. 2016. Dynamic environmental photosynthetic imaging reveals emergent phenotypes. Cell Syst. 2(6):365-77.

25.         Debuch H. 1961. Über die Fettsäuren aus grünen Blättern und das Vorkommen der Δ3-trans Hexadecensäure Z Naturforsch B 16B:246.

26.         Dörmann P, Balbo I, & Benning C. 1999. Arabidopsis galactolipid biosynthesis and lipid trafficking mediated by DGD1. Science 284(5423):2181.

27.         Durrett, T. P., C. Benning, and J. Ohlrogge. 2008. Plant triacylglycerols as feedstocks for the production of biofuels. Plant J. 54:593-607

28.         Edgerton, M. D. 2009. Increasing crop productivity to meet global needs for feed, food, and fuel. Plant Physiol 149:7-13

29.         Falkowski, P., R. J. Scholes, E. Boyle et al. 2000. The global carbon cycle: a test of our knowledge of earth as a system. Science 290:291-296

30.         Farrokh, P., M. Sheikhpour, A.Kasaeian, H. et al. 2019. Cyanobacteria as an eco-friendly resource for biofuel production: A critical review. Biotech. Prog. 35(5); e2835.

31.         Formighieri C, Melis A. 2014. Regulation of β‑phellandrene synthase gene expression, recombinant protein accumulation, and monoterpene hydrocarbons production in Synechocystis transformants. Planta 240:309–324 DOI: 10.1007/s00425-014-2080-8

32.         Formighieri C, Melis A. 2014. Carbon partitioning to the terpenoid biosynthetic pathway enables heterologous β‑phellandrene production in Escherichia coli cultures. Arch Microbiol 196(12):853-861 DOI 10.1007/s00203-014-1024-9

33.         Formighieri C, Melis A. 2015. A phycocyanin·phellandrene synthase fusion enhances recombinant protein expression and β-phellandrene (monoterpene) hydrocarbons production in Synechocystis (cyanobacteria). Metab. Eng. 32:116–124 http://dx.doi.org/10.1016/j.ymben.2015.09.010

34.         Freeman Allen C, Good P, Davis HF, & Fowler SD. 1964. Plant and chloroplast lipids I. Separation and composition of major spinach lipids. Biochem. Biophys. Res. Commun. 15(5):424.

35.         Froehlich JE, Benning C, & Dörmann P. 2001. The digalactosyldiacylglycerol (DGDG) synthase DGD1 is inserted into the outer envelope membrane of chloroplasts in a manner independent of the general import pathway and does not depend on direct interaction with monogalactosyldiacylglycerol synthase for DGDG biosynthesis. J. Biol. Chem. 276(34):31806.

36.         Gao J, I. Ajjawi, A. Manoli, A. Sawin, C. Xu, J.E. Froehlich, R.L. Last and C. Benning. 2009. FATTY ACID DESATURASE4 of Arbaidopsis encodes a protein distinct from characterized fatty acid desaturases. Plant J. 60(5):832-9.

37.         Gibson, K., J.-K. Park, Y. Nagai, et al.. 2011. Exploiting leaf starch synthesis as transient sink to increase plant productivity and yields. Plant Sci. 181:275-281

38.         Głowacka, K., Kromdijk, J., Kucera, K., Xie, J., Cavanagh, A. P., Leonelli, L., Leakey, A. D. B., Ort, D. R., Niyogi, K. K., Long, S. P. 2018. Photosystem II Subunit S overexpression increases the efficiency of water use in a field-grown crop. Nature Communications, 9, 868. https://doi.org/10.1038/s41467-018-03231-x

39.         Godfray, H. C., J. R. Beddington, I. R. Crute, et al.. 2010. Food security: the challenge of feeding 9 billion people. Science 327:812-818

40.         Greene, T. W., S. E. Chantler, M. L. Kahn, G. F. Barry and others. 1996. Mutagenesis of the potato tuber ADPglucose pyrophosphorylase and characterization of an allosteric mutant defective in 3-phosphoglycerate activation. 93:1509-1513

41.         Heffner, E. L., M.R. Sorrels, and J-L. Jannink. 2009. Genomic selection for crop improvement. Crop Sci 49:1-12.

42.         Herbstova M, S. Tietz, C. Kinzel, M.V. Turkina, and H. Kirchhoff. 2012. Architectural switch in plant photosynthetic membranes induced by light stress. Proc Natl Acad Sci U S A 109(49):20130.

43.         Herritt, M., A.P. Dhanapal, L.C. Purcell, and F.B. Fritschi. 2018. Identification of genomic loci associated with 21 chlorophyll fluorescence phenotypes by genome-wide association study in soybean.  BMC Plant Biology 18:312. DOI: 10.1186/s12870-018-1517-9

44.         Herritt, M., and F.B. Fritschi. 2020. Characterization of photosynthetic phenotypes and chloroplast ultrastructural changes of soybean (Glycine max) in response to elevated air temperatures. Frontiers in Plant Science 11:153. DOI: 10.3389fpls.2020.00153.

45.         Ho, L. C. 1988. Metabolism and compartmentation of imported sugars in sink organs in relation to sink strength. 39:355-378

46.         Höhner R, Pribil M, Herbstová M, Lopez LS, Kunz HH, Li M, Wood M, Svoboda V, Puthiyaveetil S, Leister D, Kirchhoff H. 2020. Plastocyanin is the long-range electron carrier between photosystem II and photosystem I in plants. Proc. Natl. Acad. Sci. USA 117, 15354-15362.

47.         Horn, P.J.; Smith, M.D.; Clark, T.R.; Froehlich, J.E.; Benning, C. 2020. PEROXIREDOXIN Q stimulates the activity of the chloroplast 16:1(Delta3trans) FATTY ACID DESATURASE4. Plant J, 102, 718-729, doi:10.1111/tpj.14657.

48.         Hwang, S. K., S. Hamada, and T. W. Okita. 2006. ATP binding site in the plant ADP-glucose pyrophosphorylase large subunit. FEBS Letters 2006/12/02:6741-6748

49.         Hwang, S. K., S. Hamada, and T. W. Okita. 2007. Catalytic implications of the higher plant ADP-glucose pyrophosphorylase large subunit. Phytochemistry 2007/01/09:464-477

50.         Hwang, S. K., Y. Nagai, D. Kim, and T. W. Okita. 2008. Direct appraisal of the potato tuber ADP-glucose pyrophosphorylase large subunit in enzyme function by study of a novel mutant form. Journal of Biological Chemistry 2008/01/18:6640-6647

51.         Hwang, S. K., P. R. Salamone, H. Kavakli, C. J. Slattery et al., 2004. Rapid purification of the potato ADP-glucose pyrophosphorylase by poly-histidine mediated chromatography. Protein Expression and Purification 38:99-107

52.         Hwang, S. K., P. R. Salamone, and T. W. Okita. 2005. Allosteric regulation of the higher plant ADP-glucose pyrophosphorylase is a product of synergy between the two subunits. FEBS Letters 2005/02/16:983-990

53.         Imam, S., S. Schäuble, J. Valenzuela, and others. 2015. A refined genome-scale reconstruction of Chlamydomonas metabolism provides a platform for systems-level analyses. The Plant J 84, 1239–1256

54.         Jagadish SVK. 2020. Heat stress during flowering in cereals – effects and adaptation strategies. New Phytologist 226(6), 1567-1572.

55.         Jarvis P, P. Dormann, C.A. Peto, J. Lutes, C. Benning nd J. Chory. 2000. Galactolipid-deficiency and abnormal chloroplast development in the Arabidopsis MGD synthase 1 mutant. Proc. Natl. Acad. Sci. USA 97:8175.

56.         Kaan, K, S.-K. Hwang, M. Wood, S. Singh, A. Cousins, H. Kirchhoff, and T.W. Okita. 2021. The rice plastidial phosphorylase participates directly in both sink and source processes.  Plant Cell Physiol. 62: 125-142.  doi: 10.1093/pcp/pcaa146. PMID: 33237266.

57.         Kambhampati, S, J Aznar-Moreno, SR Bailey, JJ Arp, K Chu, TP Durrett, DK Allen. Temporal changes in soybean seed developmental metabolism leading to the accumulation of oil, protein and carbohydrates. Plant Physiology (in press)

58.         Kim, D., S. K. Hwang, and T. W. Okita. 2007. Subunit interactions specify the allosteric regulatory properties of the potato tuber ADP-glucose pyrophosphorylase. Biochemical and Biophysical Research Communications. 2007/08/21:301-306

59.         Kirchhoff H, C. Hall, M. Wood, M. Herbstova, O. Tsabari, R. Nevo, D. Charuvi, E. Shimoni and Z. Reich. 2011. Dynamic control of protein diffusion within the granal thylakoid lumen. Proc Natl Acad Sci USA 108(50):20248.

60.         Kirchhoff H. 2013. Structural constraints for protein repair in plant photosynthetic membranes. Plant Signal Behav 8(4):e23634.

61.         Kirchhoff H. 2014a. Structural changes of the thylakoid membrane network induced by high light stress in plant chloroplasts. Philosophical transactions of the Royal Society of London. Series B, Biological sciences 369(1640):20130225.

62.         Kirchhoff H. 2014b. Diffusion of molecules and macromolecules in thylakoid membranes. Biochim Biophys Acta 1837(4):495.

63.         Klaus D, H. Hartel, L.M. Fitzpatrick, J.E. Froehlich, C. Benning and P. Dormann. 2002. Digalactosyldiacylglycerol synthesis in chloroplasts of the  Arabidopsis thaliana dgd1  mutant. Plant Physiol. 128:885.

64.         Knoop, H., Y. Zilliges, W. Lockau and R. Steuer. 2010. The metabolic network of Synechosystis sp. PCC 6803: Systemic properties of autotrophic growth. Plant Physiol. 154:410-422.

65.         LaBrant E, Barnes AC, Roston RL. 2018. Lipid transport required to make photosynthetic membranes. Photosynthesis Res. 138, 345-360. doi: 10.1007/s11120-018-0545-5

66.         Lavell, A.A. and C. Benning. 2019. Cellular organization and regulation of plant glycerolipid metabolism. Plant Cell Physiol. 60(6):1176-1183.

67.         Lavell, A.; Froehlich, J.E.; Baylis, O.; Rotondo, A.; Benning, C. A. 2019. Predicted Plastid Rhomboid Protease Affects Phosphatidic Acid Metabolism in Arabidopsis thaliana. Plant J, 10.1111/tpj.14377, doi:10.1111/tpj.14377.

68.         Li J, Weraduwage SM, Preiser AL, Tietz S, Weise SE, Strand DD, Froehlich JE, Kramer DM, Hu J, Sharkey TD. 2019. A cytosolic bypass and G6P shunt in plants lacking peroxisomal hydroxypyruvate reductase. Plant Physiology 180: 783-792

69.         Li M, Mukhopadhyay R, Svoboda V, Oung HMO, Mullendore DL, Kirchhoff H. 2020. Measuring the dynamic response of the thylakoid architecture in plant leaves by electron microscopy. Plant Direct 4, e00280.

70.         Li, X.-P., BjoÈrkman, O., Shih, C., Grossman, A. R., Rosenquist, M., Jansson, S., Niyogi, K. K. 2000. A pigment-binding protein essential for regulation of photosynthetic light harvesting. Nature, 403, 391–395. https://doi.org/10.1038/35000131

71.         Libourel, I. G. and Y. Shachar-Hill. 2008. Metabolic flux analysis in plants: from intelligent design to rational engineering. Annu.Rev.Plant Biol. 59:625-650

72.         Lim SD, Yim, WC, Liu D, Hu R, Yang X, Cushman JC. 2018. A Vitis vinifera basic helix-loop-helix transcription factor enhances plant cell size, vegetative biomass and reproductive yield. Plant Biotechnology Journal. 16: 1595–1615. DOI: 10.1111/pbi.12898.

73.            Lim SD, Lee S, Choi W-G, Yim WC, Cushman JC. 2019. Laying the foundation for crassulacean acid metabolism (CAM) biodesign: Characterization of individual C4 metabolism genes from the common ice plant in Arabidopsis. Frontiers in Plant Science. 10: 101. DOI: 10.3389/fpls.2019.00101.

74.            Lim SD, Meyer JA, Yim WC, and Cushman JC. 2020. Plant tissue succulence engineering improves water-use efficiency, water-deficit stress attenuation, and salinity tolerance in Arabidopsis. The Plant Journal. 103: 1049–1072. DOI: 10.1111/tpj.14783

75.            Lindquist E, K. Solymosi, and H. Aronsson. 2016. Vesicles are persistent features of different Plastids. Traffic 17(10):1125.

76.            Liu Y, Wang B, Li J, Song Z, Lu B, Chi M, Yang B, Qin D, Lam YW, Li J, Xu D. 2017. Salt response analysis in two rice cultivars at seedling stage. Acta Physiologiae Plantarum 39:215.

77.      Ma, F, LJ Jazmin, JD Young, and DK Allen. 2014. Isotopically nonstationary 13C flux analysis of Arabidopsis thaliana leaf metabolism at varying light intensities. Proceedings of the National Academy of Sciences 111:16967-16972

78.      Malnoë, A. 2018. Photoinhibition or photoprotection of photosynthesis? Update on the (newly termed) sustained quenching component qH. Environmental and Experimental Botany, 154, 123–133. https://doi.org/10.1016/j.envexpbot.2018.05.005

79.      Mandy, D, J Goldford, H Yang, DK Allen, and IGL Libourel. 2014 Metabolic flux analysis using 13C peptide label measurements. Plant Journal 77:476-486

80.       McAdam SAM, Sussmilch FC. 2021. The evolving role of abscisic acid in cell function and plant development over geological time. Semin Cell Dev Biol 109: 39-45.

81.       Melis, A. 2013. Carbon partitioning in photosynthesis. Curr Opin Chem Biol. 17:453-456.

82.       Ming, R., R. VanBuren, C.M. Wai, H. Tang, M.C. Schatz, et al., 2015. The pineapple genome and the evolution of CAM photosynthesis. Nature Genetics 47, 1435–1442.

83.       Moellering ER, Muthan B, & Benning C. 2010. Freezing tolerance in plants requires lipid remodeling at the outer chloroplast membrane. Science 330(6001):226.

84.       Muhlethaler K & Freywyssling A. 1959 Entwicklung Und Struktur Der Proplastiden. Journal of Biophysical and Biochemical Cytology 6(3):507.

85.       Müller, P., Li, X. P., Niyogi, K. K. 2001. Non-photochemical quenching. A response to excess light energy. Plant Physiology, 125(4), 1558–1566. https://doi.org/10.1104/pp.125.4.1558

86.       Murchie, E. H. and K. K. Niyogi. 2011. Manipulation of photoprotection to improve plant photosynthesis. Plant Physiol 155:86-92

87.       Mutava, R. N., P. V. V. Prasad, M. R. Tuinstra, M. D. Kofoid et al., 2011. Characterization of sorghum genotypes for traits related to drought tolerance. Field Crops Research 123:10-18

88.       Nakamura, Y.; Tsuchiya, M.; Ohta, H. 2007. Plastidic phosphatidic acid phosphatases identified in a distinct subfamily of lipid phosphate phosphatases with prokaryotic origin. J Biol Chem, 282, 29013-29021, doi:10.1074/jbc.M704385200.

89.       Nam JW, Jenkins LM, Li J, Evans B, Jaworski JG, Allen DK (2020) A general method for quantification and discovery of acyl groups attached to acyl carrier proteins in fatty acid metabolism using LC-MS/MS. Plant Cell.  https://doi.org/10.1105/tpc.19.00954

90.       Neupane D, Mayer JA, Niechayev NA, Bishop CD, Cushman JC. 2021 Five-year field trial of the biomass productivity and irrigation response of  cactus pear (Opuntia spp.) as a bioenergy feedstock for arid lands. Global Change Biology: Bioenergy. 13(4): 719-741. DOI: 10.1111/gcbb.12805.

91.       Olsson V, Joos L, Zhu S, Gevaert K, Butenko MA, De Smet I. 2019. Look closely, the beautiful may be small: Precursor-derived peptides in plants. Annual Review of Plant Biology 70:153-186.

92.       Ortiz-Bobea, A., Ault, T.R., Carrillo, C.M. et al. 2021. Anthropogenic climate change has slowed global agricultural productivity growth. Nat. Clim. Chang. 11, 306–312

93.       Parry, M. A., M. Reynolds, M. E. Salvucci, C. et al. 2011. Raising yield potential of wheat. II. Increasing photosynthetic capacity and efficiency. J.Exp.Bot. 62:453-467

94.       Portis, A. R., Jr., C. Li, D. Wang, and M. E. Salvucci. 2008. Regulation of Rubisco activase and its interaction with Rubisco. J.Exp.Bot. 59:1597-1604

95.       Prasad PVV, R. Bhemanahalli, S.V.K. Jagadish. 2017. Field crops and the fear of heat stress - opportunities, challenges, and future directions. Field Crops Res. 200, 114-121.

96.       Puthiyaveetil S, O. Tsabari, T. Lowry, S. Lenhery, R. R. Lewis, Z. Reich and H. Kirchoff. 2014. Compartmentalization of the protein repair machinery in photosynthetic membranes. Proc Natl Acad Sci U S A 111(44):15839.

97.       Que, Q., M. D. Chilton, C. M. de Fontes, et al.. 2010. Trait stacking in transgenic crops: Challenges and opportunities. GM Crops 1:220-229

98.       Radakovits, R., R. E. Jinkerson, A. Darzins, and M. C. Posewitz. 2010. Genetic engineering of algae for enhanced biofuel production. Eukaryot. Cell 9:486-501

99.       Raines, C. A. 2011. Increasing photosynthetic carbon assimilation in C3 plants to improve crop yield: current and future strategies. Plant Physiol 155:36-42

100.     Raza A, Razzaq A, Mehmood SS, et al. 2019. Impact of Climate Change on Crops Adaptation and Strategies to Tackle Its Outcome: A Review. Plants (Basel). 2019;8(2):34.

101.     Ristic Z, Jenks MA. 2002. Leaf cuticle and water loss in maize lines differing in dehydration avoidance. Journal of Plant Physiology 159: 645-651.

102.     Roberts, L. 2011. 9 billion? Science 333:540-543

103.     Sah SK, Reddy KR, and Li J. 2016. Abscisic acid and abiotic stress tolerance in crop plants. Frontiers in Plant Science 7:571.

104.     Schlosser AJ, Martin JM, Beecher BS, Giroux MJ. 2014. Enhanced rice growth is conferred by increased leaf ADP-glucose pyrophosphorylase activity. Journal of Plant Physiology and Pathology 2:4.

105.     Sharkey, T. D. and R. Zhang. 2010. High temperature effects on electron and proton circuits of photosynthesis. J.Integr.Plant Biol. 52:712-722

106.     Sharkey TD, Preiser AL, Weraduwage SM, Gog L. 2020. Source of 12C in Calvin-Benson cycle intermediates and isoprene emitted from plant leaves fed with 13CO2. Biochemical Journal 477: 3237-3252

107.     Sharkey TD, Weise SE. 2016 The glucose 6-phosphate shunt around the Calvin-Benson cycle. Journal of Experimental Botany 67: 4067-4077

108.     Sharma, A., C.M. Wai, R. Ming, Q. Yu. 2017. Diurnal cycling transcription factors of pineapple revealed by genome-wide annotation and global transcriptomic analysis. Genome Biology and Evolution 9: 2170-2190

109.     Somerville, C., H. Youngs, C. Taylor, et al. 2010. Feedstocks for lignocellulosic biofuels. Science 329:790-792

110.     Stitt, M., J. Lunn, and B. Usadel. 2010. Arabidopsis and primary photosynthetic metabolism - more than the icing on the cake. Plant J. 61:1067-1091

111.     Sunagawa, H., J. C. Cushman, and S. Agarie. 2010. Crassulaceaen acid metabolism alleviates reactive oxygen species in the facultative CAM plant, the common ice plant, Mesembryanthemum crystallinum. Plant Prod.Sci. 13:246-260

112.     Terashima, I., Y. T. Hanba, D. Tholen, and U. Niinemets. 2011. Leaf functional anatomy in relation to photosynthesis. Plant Physiol 155:108-116

113.     Tester, M. and P. Langridge. 2010. Breeding technologies to increase crop production in a changing world. Science 327:818-822

114.     Thomson, C.M., P. Pulido, and R.P. Jarvis. 2020. Protein import into chloroplasts and its regulation by the ubiquitin-proteasome system. Biochem. Soc Trans. 48(1):71-82.

115.     Turgeon, R. 1989. The source-sink transition in leaves. 40:119-138

116.     Twyford, A.D. 2018. The road to 10,000 plant genomes. Nature Plants. 4:312-313.

117.     Valsami, E-A, M.E. Psychogyiou, A. Pateraki, E. et al. 2020. Fusion constructs enhance heterologous b-phellandrene production in Synechocystis sp. PCC 6803. J. Appl. Phycology. 32:2889-2902.

118.     Von Caemmerer, S. and J. R. Evans. 2010. Enhancing C3 photosynthesis. Plant Physiol 154:589-592

119.     Vu HS, R. Roston, S. Shiva, M. Hur, E.S. Wurtele, X. Wang, J. Shah and R. Welti. 2015. Modifications of membrane lipids in response to wounding of Arabidopsis thaliana leaves. Plant Signal Behav 10(9):e1056422.

120.     Weise SE, Liu T, Childs KL, Preiser AL, Katulski HM, Perrin-Porzondek C, Sharkey TD. 2019. Transcriptional regulation of the glucose-6-phosphate/phosphate translocator 2 is related to carbon exchange across the chloroplast envelope. Frontiers in Plant Science 10: 827

121.      Westbrook AS, McAdam SAM. 2021. Stomatal density and mechanics are critical for high productivity: insights from amphibious ferns. New Phytologist 229(2): 877-889.

122.      Westphal S, J. Soll, and U.C. Vothknecht. 2001. A vesicle transport system inside chloroplasts. FEBS Lett 506(3):257.

123.      Wijffels, R. H. and M. J. Barbosa. 2010. An outlook on microalgal biofuels. Science 329:796-799

124.      Xu C., J. Fan, W. Riekhof, J.E. Froehlich, and C. Benning.  2003. A permease-like protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J. 22(10):2370.

125.      Xu C., J. Fan, J. Froehlich, K. Awai, and C. Benning. 2005. Mutation of the TGD1 chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. Plant Cell 17:3094.

126.      Xu Y., Fu X., Sharkey T.D., Shachar-Hill Y. & Walker B. 2021. The metabolic origins of non-photorespiratory CO2 release during photosynthesis: A metabolic flux analysis. Plant Physiology 186, 297-314.

127.      Yang, W., H Feng, X Zhang, et al. 2020. Crop Phenomics and High-Throughput Phenotyping: Mol Plant 13, 187-214.

128.      Yu B. and C. Benning. 2003. Anionic lipids are required for chloroplast structure and function in Arabidopsis. Plant J. 36(6):762.

129.      Yuan, G., M. M. Hassan, D. Liu, et al. 2020. Biosystems design to accelerate C3-to-CAM progression. BioDesignResearch. 2020:3686791.

130.      Zhang et al., 2021. Systems-wide Analysis Revealed Shared and Unique Responses to Moderate and Acute High Temperatures in the Green Alga Chlamydomonas reinhardtii. bioRxiv 2021.08.17.456552 doi.org/10.1101/2021.08.17.456552
131.      Zhou, XR, S Bhandari, B Johnson, HK Kotapati, DK Allen, T Vanhercke, PD Bates. 2020. Reorganization of acyl flux through the lipid metabolic network in oil-accumulating tobacco leaves. Plant Physiology 182(2):739-755

132.      Zhu, X. G., S. P. Long, and D. R. Ort. 2010. Improving photosynthetic efficiency for greater yield. Annu.Rev.Plant Biol. 61:235-261

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CA, IL, IN, KS, MI, MS, MT, NE, NV, TX, VA, WA

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Danforth Plant Science Center, USDA-ARS/Missouri, USDA-ARS/TX
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