Duration: 10/01/2017 to 09/30/2022

Administrative Advisor(s):

NIFA Reps:

Non-Technical Summary

Statement of 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 reduced molecules (e.g. sugars, lipids, amino acids and other cell building blocks), and oxygen is produced through photosynthetic water oxidation. Thus, plant and algal photosynthesis affects global geochemical processes, in particular carbon cycling (36), 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 (128, 143). Some of the greatest challenges humankind is currently facing – 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 (i.e. agricultural crops) (35). Therefore, the need for state-of-the-art photosynthesis research to improve the efficiency 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 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 the plant response. 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 (47).  For well-fertilized crops, HI has approached the maximum achievable for many crops, and future yield gains will depend on increasing total production (123).  Greater productivity, which is necessary to meet the food, feed, fuel and fiber needs of a growing world population (104), requires improving the basic efficiency of the photosynthetic process for light energy capture and the utilization of this energy for the synthesis and utilization of organic molecules (123).  For maximum benefit, the efficiency of the process must be increased under both optimal and sub-optimal conditions, consonant with the changes in temperature, CO₂, and precipitation anticipated from climate change. 

Importance and Consequences

Global climatic trends are negatively impacting the yields of our major crops used for the production of food, feed, fiber, and biofuels (80). 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 production in a way that is sustainable, that is not in competition with food, and that reduces greenhouse gas production requires novel approaches and non-traditional crops including possibly algae (116, 134). Only a concerted and vertically integrated effort encompassing all aspects of photosynthesis will ensure that appropriate solutions will 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 crops 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 technologies have advanced to the point that a wealth of information can be generated for any species of plant or alga (e.g., date and oil palm) (1, 20), in a relative short time span and at reasonable costs. Thus, the raw material for genetic engineering of novel crops or algal strains is readily available. Large-scale phenomics focused on chloroplast proteins in model plants such as Arabidopsis has become possible (81). Moreover, using comparative genomics, reconstruction of metabolism from gene expression and genomic data has become readily feasible for any organism. One example is the recent metabolic reconstruction of the alga Chlamydomonas reinhardtii (23). In addition, our knowledge and analysis of primary metabolism of plants and metabolic networks has advanced to levels (76) that can enable the rational design of novel crops, which will meet our needs. Despite this progress, the task of genetically transforming crop plants and analyzing the consequences (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 (98). 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 the basic discovery process and to devise strategies for the improvement of photosynthesis in crops, facile model organisms such as Arabidopsis (119) or model microalgae and cyanobacteria such as Chlamydomonas (99) and Synechocystis (25) will have to be employed to quickly identify the most promising directions for photosynthetic pathway and crop improvement. Following this guidance, collaboration with geneticists and breeders, associating traits with genomic regions of mapping populations can support marker-assisted selection (50). Such an integrated approach requires multifaceted expertise and, thus, will benefit from synergy derived from a multi-investigator effort. 

Multi State Effort

Providing a conceptual framework through the current project, this North Central Regional Group of scientists and others located throughout the US has already successfully made progress on understanding diverse aspects of photosynthesis bringing together a complementary set of expertise. 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 North Central Region and other participating states. Participating partners 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. Identify 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 biochemical production factory. As semi-autonomous organelles, chloroplasts do not function by themselves, but rely on extensive communication with other organelles 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 (16, 73). As primary photosynthate and many other metabolites (e.g., fatty acids), 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 photosynthesis occurs on a specialized structure called the thylakoid membrane, which is itself dynamically remodeled in response to development and stress cues. The architectural dynamics of the 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 transformation, the development of the thylakoid membranes under developmental and stress regimens, and the development of tools that can be used to assess thylakoid and inner envelope connectivity (MI-ABR, NE-AES, WA-AES). 

2. Photosynthetic Capture and Photorespiratory Release of CO₂. Photosynthetic carbon fixation and photorespiratory release of CO₂ have long been recognized as limitations to crop productivity (46, 94, 100, 121). Considerable focus will be placed on engineering improvements in the organization of the ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) and the heat-sensitive Rubisco activase (RCA) that regulates its function. Microbial CO₂-concentrating mechanisms will be investigated for potential improvements in the photosynthesis of algae and crop plants. Studies will examine the mechanisms of RCA regulation in soybean including redox regulation, protein phosphorylation and possible ‘cross-talk’ between the two regulatory processes. Emphasis will also be placed on the mechanisms that control carbon flux through primary and secondary metabolic pathways. Likely impacts will be insights into the regulation of carbon fixation and the generation of modified enzymes that improve primary and secondary carbon metabolism in plants. (IL-ARS, MI-ABR, MO-AES, NV-AES, WA-AES). 

3. Mechanisms Regulating Photosynthate Partitioning. Manipulation of carbon partitioning and understanding its regulation is central to advances in yield formation in cereal and oilseed crops, to the design of new crops for biofuel or commodity chemicals production that, e.g., divert carbon from carbohydrate synthesis into useful triacylglycerols (34), or terpenoid products (85). This work will lead to the development and commercial application of novel organisms that are endowed with the heterologous expression of terpene synthases (from higher plants) in either production-type plants or microalgae. The work will further seek to divert carbon flux and to limit catabolic reactions resulting in respiration that reduces the fraction of photosynthate available for yield formation. Transport phenomena at the cell, tissue and whole plant level will be considered, sugar-sensing mechanisms will be explored, as well as partitioning between storage carbohydrates, lipids, terpenoids and stress-protective compounds. Likely impacts will be defining the mechanisms that regulate photosynthate partitioning into the biosynthetic pathways for sucrose, starch, sugar alcohols, terpenoids, and lipids. Strategies will be developed to alter carbon partitioning by engineering bypass pathways to overcome rate and yield barriers to commercial application. (CA-AES, FL-AES, IL-ARS, KS-AES, MI-ABR, MO-ARS, MT-AES, OH-AES, VA-AES, WA-AES). 

4. Developmental and Environmental Limitations to Photosynthetic Productivity. Factors such as leaf anatomy (122) or environmental stress conditions such as high light (90), excess salinity, nitrogen deficiency, drought or heat stress (111) greatly affect photosynthesis. Leaf stomata, regulating photosynthetic productivity as a ‘gateway’ for CO₂ influx, are subject to complex active regulation. In the case of drought, the plant hormone abscisic acid (ABA) inhibits stomatal opening and promotes stomatal closure, supporting water conservation. A dynamic model of ABA-induced stomatal closure includes more than 40 identified network components (75); additional plant water status regulation is responsive to factors including light intensity, temperature, vapor pressure, and leaf CO₂ partial pressure. Feedback regulation of photosynthesis by sugar sensing and signal transduction mechanisms are known, but incompletely understood. Frequently, these regulatory pathways result from a cascading sequence of processes. Particular emphasis will be placed on identifying processes driving responses to abiotic stresses (e.g., temperature, water, and salinity), nitrogen use, and global atmospheric change. Likely impacts will be identifying genomic regions associated with developmental and environmental factors that influence photosynthetic productivity at the whole plant and canopy levels. (IL-AES, IL-ARS, KS-AES, MI-ABR, MO-AES, MS-AES, NV-AES, OH-AES).

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 (812 hits alone) and in combination with other terms such as lipids (21 hits), Rubisco (60 hits), carbon fixation (40 hits) etc. were 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. Knowledge of the apparent self-assembly and self-repair of the photosynthetic membrane, and the essential roles that membrane architecture through lipid and protein dynamics play in this process is expected to guide efforts in developing artificial photosynthetic membranes and enhance engineering of plant feedstocks.

Collaborators focus on two major sub-objectives: 1.1 Chloroplast membrane lipid and protein dynamics and 1.2 Dynamics of chloroplast membrane architecture. The first sub-objective enhances knowledge of production, delivery, and photosynthetic relevance of individual molecular components of chloroplast membranes, while the second addresses the importance of the three-dimensional arrangement of multiple components simultaneously.

1.1 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) (30, 37). 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 (33, 41, 42, 61, 69, 87, 137, 138, 139). Current investigations include biosynthesis and photosynthetic relevance of two hypothesized chloroplast PG pools. 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 (14, 129). Current investigations include collaborations with the Benning lab (MI-ABR) to determine suppressors of the response and independent investigations into other lipid responses to the same subcellular signal. The Stone lab (NE-AES) has expanded knowledge of plant programmed cell death responses to include the critical chloroplast protein AtDJ1C (77). Current investigations include determining the precise role of DJ1C in the chloroplast, and in collaboration with the Roston lab, its sub-organellar location.

1.2 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. 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 (52, 65, 66, 97), and facilitate diffusion of small electron carriers leading to efficient photosynthetic electron transport (52, 67, 68). Alternate membrane structures include, large invaginations of the inner envelope membrane (24, 89), small vesicles in the stroma (79, 133), and membrane contact sites between the inner and thylakoid membranes (22). 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. They have recently defined the role of semi-crystalline protein array formation in thylakoid membranes (124) and of architectural switches in thylakoid membranes for protein repair processes (97). The Roston lab (NE-AES), in addition to work described above is beginning to target understanding of formation and maintenance of the thylakoid lipid bilayer by investigating alternate chloroplast membrane structures.

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 three sub-objectives: 2.1 Rubisco activase, 2.2 Electron transport and regulation of the Calvin-Benson cycle, and 2.3 C₄ and CAM CO₂ concentrating mechanisms. The first sub-objective addresses an initial step of carbon fixation in most plants, the second sub-objective follows carbon after carboxylation, and the third sub-objective traces CO₂ accumulation mechanisms that optimize the operation of Rubisco. 

2.1 Rubisco activase. An initial step of carbon fixation, in most plants, is the carboxylation of ribulose 1,5-bisphosphate by Rubisco. Rubisco is maintained in an active state by another chloroplast protein, Rubisco activase (RCA). Rubisco is typically inactive at low light, high CO₂, and high temperature. The inactivation at low light and high CO₂ could be adaptive, but deactivation at high temperature is less well understood. In all cases the mechanisms regulating RCA are unknown. In this sub-objective, we aim to understand the role of Rubisco activase in regulating light utilization by plants. The Huber lab (IL-ARS) investigates regulation of RCA. RCA catalyzes the release of tightly-bound inhibitors from the active site of Rubisco. Rubisco activity may be increased by altering the interaction of Rubisco with RCA, improving the thermal stability of RCA (93, 117), or removing the post-translational modifications that reduce RCA activity. Past work (AZ-ARS, NE-AES) identified a substrate recognition domain in activase that directly interacts with a region in Rubisco (72, 92); (51) reported the X-ray crystal structure of this “lever-arm” domain. Arabidopsis and rice RCAs are composed of two subunits: a large 47-kDa α-subunit and a smaller 43-kDa β-subunit. The activity of RCA is regulated by ADP inhibition and redox control such that the activation state of Rubisco in planta parallels light intensity. Redox control of soybean RCA involves two Cys residues located in the large α-subunit of the protein that when oxidized form a disulfide that dramatically enhances ADP inhibition.  Recent results with Arabidopsis indicate that removal of the redox-regulated α-subunit (to leave only the β-subunit) prevents the deactivation of Rubisco at low light intensity such that growing plants in an alternating low light/high light regime results in more effective use of light energy and greater biomass accumulation. We will attempt to translate these results from Arabidopsis to rice and other crop plants through genome engineering. 

2.2 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 (MI-ABR) and Cousins (WA-AES) labs are exploring how CO₂ capture in the Calvin-Benson cycle is regulated and connected with electron transport regulation as well as CO₂ supply. 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. This project will lead to studies of the interaction of carbon metabolism and cyclic electron flow.

The Fritschi group (MO-AES) is exploring the genetics underlying leaf and canopy level pigment composition and leaf chlorophyll fluorescence characteristics in soybean.  Based on genome-wide association analyses, this research has identified numerous loci putatively associated with photosynthetic light reactions (30A, 30B, 50A).  These efforts have also led to the identification of genotypes with phenotypic extremes that will be leveraged for physiological and genetic studies.  Effects of high temperatures on photosynthetic light reactions will be characterized in phenotypic extremes. 

2.3 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) is studying carbon uptake in relation to water use in C₄ plants. Populations of two Setaria species and RIL have been grown in the field. Contents of ¹³C and ¹⁸O have been measured and mapped to find QTLs. 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) group has investigated the molecular basis of CAM evolution and the roles of storage carbohydrates and circadian-clock control of CAM. CAM evolution progresses from C₃ photosynthesis to weak and then to strong CAM. Strong CAM is correlated with reciprocal fluxes in acidity and storage carbohydrates, tissue succulence, isogene recruitment, and leaf-specific and circadian clock controlled mRNA expression (113-115). A colorimetric assay was developed to measure leaf pH and screen a mutagenized population of common ice plant (Mesembryanthemum crystallinum) to isolate CAM-deficient mutants (28). One mutant showed greater H₂O₂ content and Cu/Zn-superoxide dismutase mRNA expression, indicating that CAM might contribute to alleviation of oxidative stress (121). Feeding of sugars restored nocturnal accumulation of organic acids, indicating that CAM possesses some flexibility for utilizing different carbohydrate sources (28). Gene expression profiling showed changes associated with induction of CAM by salinity stress (29). Many genes undergo shifts in their circadian-clock output phase during the stressed-induced shift from C₃ photosynthesis to CAM providing new CAM-specific markers. 

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 Metabolic control, 3.2 Sugar-mediated signaling mechanisms, and 3.3 Transformation of assimilates into high-energy or high-value compounds. The first sub-objective addresses starch biosynthesis, degradation, and assimilate transport, the second sub-objective examines signaling mechanisms, and the third sub-objective targets barriers to commercial development of novel organisms and useful bio-products.

3.1 Metabolic Control. Starch synthesis is an important target for manipulating source-sink relationships as a means to increase the genetic yield potential of crop plants (53, 127) and plant productivity. 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 (106). 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. WA-AES research yielded insights into the roles of the large and small subunits that comprise the heterotetrameric subunit enzyme structure using a biochemical-genetic approach (13, 48, 54-58, 64). Unpublished results from the Okita lab (WA-AES) indicate that the construct Pho1ΔL80 interacts with the PsaC subunit of photosystem II and possibly controls electron flow to ferredoxin and NADPH production.

The Allen lab (MO-ARS) investigates the spatial and temporal dynamics of central carbon metabolism in plant cells (3) including the development of comprehensive metabolic flux maps to assess resource allocation and carbon partitioning in oilseeds (4, 5) and in leaves (82). The methods have been built to describe subcellular aspects of metabolism using isotopes that label spatially distinct compounds according to the fluxes in different subcellular or cellular locations (6-8, 83).

The Jagadish lab (KS-AES) investigates effects of high night temperature (HNT) on dark respiration (Rn) in relation to grain-filling processes in cereal crops. Dark respiration response to HNT, for susceptible rice cultivars, was greatest during the post flowering phase (9). 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). 

Work by MI-ABR on leaf starch degradation pathways (27, 110, 118, 130-132) showed that β-maltose is the primary molecule for export of carbon from chloroplasts at night. Export of maltose in preference to triose phosphates or glucose lowers the ATP required for starch to sucrose conversion at night from four to three (130). 

3.2. Sugar-mediated signaling mechanisms. The starch degradation pathway can provide important insights into sugar-mediated signaling mechanisms because leaf starch metabolism is a major source of glucose for the sugar-sensing hexokinase reaction (110).  In this sub-objective, we aim to understand the mechanisms of sugar-mediated signaling. OH-AES and VA-AES focus on components of sugar-mediated signal transduction pathways as targets for increasing plant yield (59, 60, 112). OH-AES addressed the dissection of hexokinase-dependent and independent sugar signaling mechanisms (60, 135), signal crosstalk between sugar and hormones (96, 142), and more recently the transcription factor networks involved in sugar signal transduction (62, 95). The sucrose non-fermenting related kinase-1 (SnRK1) (120) gene encodes a highly conserved energy and stress sensor that VA-AES and other groups have identified as a key regulator of sugar-mediated signaling in plants (10-12). Research from FL-AES focused on differential sugar-responsiveness of genes for sucrose metabolism, and C/N balance, to produce a global framework for selective advantages of feast-and-famine genes in multicellular organisms (70, 71, 136). FL-AES also helped establish a national resource for corn (Zea mays L.) knock-out mutants that compares with that for Arabidopsis (MaizeGDB.org (84, 109)). 

3.3 Transformation of assimilates into high energy or high value compounds. Novel organisms are capable of generating industrial feedstocks, although barriers limit commercial development. In this sub-objective, we aim to understand and overcome constraints to commercial production of high-energy or high-value compounds, derived from photosynthetic assimilates. The Benning lab (MI-ABR) studies the conversion of photosynthate into high energy compounds such as triacylglycerols (TAGs) in plants and algae. TAG production can be developmentally regulated (e.g., in developing oil seeds) (34) or stress induced (e.g., in nutrient-starved algae (78) or freeze-exposed plant leaves (88)). In algae, nutrient stress induces a state of quiescence (125), which leads to cessation of cell growth and cell division, while at the same time large-scale changes in metabolism occur including the accumulation of TAGs (86). Overcoming this apparent conundrum, which hampers the efficient production of TAG from algal aquaculture, is one of the goals of the Benning lab.  

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 have investigated developmental and environmental limitations to photosynthesis. These studies include nitrogen- and water-use efficiency and stress physiology (heat, salt, drought, cold), as well as the underlying mechanisms that signal plants to respond to stress. Nitrogen acquisition and its use are key to increasing the efficiency of light interception and its utilization. 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 (17,101). The response of plants to abiotic stress is 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 two major sub-objectives: 4.1 Abiotic stress responses and 4.2 Stress signal transduction. The first sub-objective investigates responses to abiotic stresses including nutrient and water deficits, chilling, heat, and salt; the second sub-objective addresses signal transduction pathways related to stress responses. 

4.1 Abiotic stress responses. Nutrient deficiencies, water deficits, cold and hot temperatures, and excessive salts impair photosynthetic processes in diverse ways. Breeding and selection for improved plant performance under different stresses can be accelerated by the development of high-throughput methods for phenotyping plant responses (101). In this sub-objective, we aim to understand the relation of abiotic stress on components of primary productivity including the photosynthetic membrane, CO₂ capture, and regulation of assimilate flow. Using advanced maize and soybean genetics, the Below lab (IL-AES) is focused on intensifying agriculture production by lowering N required per unit of carbon fixed and partitioned to grain (108). Their work shows (i) soil N depletion is likely to occur in years with above-average precipitation (44) and (ii) continuous corn yield penalty worsens over the years, with a corresponding decrease in nitrogen-use efficiency (45). Investigations including genetics, and a range of management practices including population, fertility, etc. had weather determining the greatest impact on corn yield (15, 19, 105). The Cushman group (NV-AES) is investigating Dunaliella salina Teodoresco, a unicellular, halophytic green alga, which produces massive amounts of b-carotene, and is a potential feedstock for biofuel production (102). The Cushman lab has obtained ESTs for 2,831 clones representing 1,401 unique transcripts from a complementary DNA (cDNA) library (2) to capture mRNA expression under high and low light and dark conditions, NaCl, anaerobic, and nutrient deprivation stress. 

Research on high temperature stress is a major focus with the Sharkey (MI-ABR), Cushman (NV-AES), Jagadish, Aiken and Prasad (KS-AES), Fritschi (MO-AES), and Harper (NV-AES) groups all participating in the effort, while Cushman’s group (NV-AES) also works with cold tolerance in Camelina sativa. Sharkey’s group has shown that 1) photosystem I becomes more reduced and the stroma becomes more oxidized (107), 2) the proton motive force (pmf) decreases and 3) the proportion of the pmf that is accounted for by the pH component declines (140, 141). Prasad’s group have examined the inhibitory effects of heat stress mediated through ethylene response, production of reactive oxygen species, and subsequent membranes damage (31, 32). The Jagadish and Prasad labs have 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 (91). 

4.2 Stress signal transduction. Abiotic stress responses involves cellular and molecular signaling pathways, often involving protein kinases. Calcium-dependent Protein Kinases (CPKs) have been implicated in regulating responses to biotic and abiotic stress, as well as multiple aspects of metabolism, vegetative development, and sexual reproduction. Numerous genetic and phenotypic studies have shown that sugar responsive Tandem Zinc Finger (TZF) proteins are potent regulators of hormone-mediated growth and stress responses (18). In this sub-objective, we aim to understand the role of protein kinases and sugar responsive genes in plant growth, reproduction, and responses to the environment. The Harper group (NV-AES) in collaboration with Cushman group (NV-AES) has obtained evidence for in vivo interactions regulating subcellular organization, signaling and metabolic regulation with most requiring phosphorylation of a binding site on the client. Many of these phospho-interactions are thought to be mediated by CPKs (21). Hence, a specific collaboration with Huber group (IL-ARS) to understand the multiple ways in which CPKs are regulated in biotic and abiotic stresses. Recently, the Huber lab has obtained evidence from in vitro kinase assays that some CPKs can autophosphorylate a tyrosine residue at the start of the kinase domain, resulting in a down-regulation of kinase activity (i.e., an auto-inactivation mechanism), which requires further understanding. The Li group (MS-AES) has identified an abscisic acid-activated protein kinase in Vicia faba and shown it to be a positive regulator of abscisic acid (ABA)-induced stomatal closure (74), leading to identifying several phosphoproteins regulated by ABA and drought (49, 63). The Harper lab has multiple research projects on lipid flippases, cyclic nucleotide-gated channels, lipid-stimulated kinases, calcium pumps, and sodium efflux mechanism.  A common theme is the role of these enzymes in plant growth, reproduction, and responses to the environment.   

The Jang lab (OH-AES) investigates sugar sensing and signal transduction mechanisms in higher plants, focusing on the dissection of hexokinase-dependent and independent sugar signaling mechanisms (59, 135), crosstalk between sugar and hormone signal transduction pathways (142, 97), and more recently the regulatory networks involved in sugar signal transduction (95, 62). Through multiple transcriptome analyses, basic leucine zipper (bZIP) and tandem zinc finger (TZF) proteins were identified as potential key players in plant sugar signal transduction. Genetic analysis indicated that bZIP transcription factors act as positive regulators in sugar starvation response. Their current work focuses on the characterization of a family of sugar responsive TZF genes.

Objectives

1. Identify strategies to optimize the assembly and function of the photosynthetic membrane.
   
2. Identify strategies to modify biochemical and regulatory factors that affect 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

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 that affect 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-AES) in part by collaboration, DNA and protein manipulation (e.g., electrophoresis, immunoblotting, cloning, native and heterological expression, enzyme assays), and biological fractionations to the sub-organellar level. 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 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 biosynthesis and function of the three glycolipids of the photosynthetic membrane are reasonably well known, but the biosynthesis and specific function of 16:1trans in phosphatidylglycerol remains to be clarified. In addition to the techniques describe above, the Benning lab (MI-ABR) couples their extensive collection of chloroplast lipid mutants in Arabidopsis with detailed lipid profiling using gas chromatography and liquid chromatography coupled with mass-spectrometry to determine the relevance of the 16:1trans pool. They also use advanced, non-invasive phenotyping based on chlorophyll fluorescence with combinations of mutations in strategic lipid genes to characterize the hypothesized PG pools. This will be done in collaboration with T. Sharkey and D. Kramer (MI-ABR) A specific desaturase is implicated in production of the 16:1 fatty acid, and detailed cell biological and biochemical analyses will be employed to determine its sub-chloroplast location, its substrate and its cofactors. 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. The Stone lab (NE-AES) will use common protein production platforms in bacteria and yeast, to investigate hypothetical functions of AtDJ1C. Further, partial loss-of-function (RNAi) lines with varying levels of DJ1C expression (and varying levels of chloroplast dysfunction) have been generated, and are now under investigation. In collaboration with the Roston lab they have initiated experiments aimed at understanding the exact location of AtDJ1C in chloroplasts using chloroplast import studies. 

1.2 Dynamics of chloroplast membrane architecture. The focus is on critical arrangements of chloroplast membranes enhancing photosynthesis. Identifying functional consequences of structural alterations is being explored by the Kirchhoff lab (WA-AES) by coupling the common methods described above with biophysical, microscopic, and computer analysis/modelling techniques. Biophysical approaches include polarographic oxygen measurements, difference absorption spectroscopy, time-resolved and steady-state fluorescence, and absorption spectroscopy at room- and cryogenic temperatures. Computational analysis allows quantification of protein distributions and generation of scale membrane models. These approaches allow in-depth functional characterization of the photosynthetic apparatus, including changes on the mesocopic level (the organization of many protein complexes within the membrane). The Roston lab (NE-AES) is studying generation of alternate chloroplast membrane structures (vesicles, invaginations, contact sites) by initiating production of multiple fluorescent visualization systems that determine thylakoid/inner envelope membrane continuity in Arabidopsis thaliana. Each system is designed to target a different alternate membrane structure in real time taking advantage of modern fluorescent protein varieties. Both labs take advantage of state-of-the-art confocal laser scanning microscopy (including diffusion measurements by FRAP) and electron microscopy combined with image analysis tools to image membrane ultrastructure. 

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 maintenance of Rubisco, the key carbon-fixing enzyme, in an active state; 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 measure changes in rates of Rubisco activity, photosynthetic electron transport and rates of carbon assimilation. 

2.1 Rubisco activase. The focus is on effects of Rubisco Activase (RCA) regulation on photosynthetic induction following transfer from low light to high light. The Huber lab (IL-ARS) will use the CRISPR-Cas9 system in the case of rice, to engineer a stop codon in the single RCA gene to truncate the protein and remove the C-terminal extension required for redox regulation. We will determine whether removal of RCA redox regulation increases photosynthetic induction following transfer from low light to high light; if so, we will examine how plant growth responds in the field where variable light intensity occurs naturally. We will also investigate the possible diel regulation of RCA activity involving protein phosphorylation, which is largely unexplored at present, but could interact with redox regulation of the enzyme as has been shown for certain other redox regulated enzymes. Because transformation of Arabidopsis is more rapid than soybean we will perform site-directed mutagenesis of the redox-sensitive Cys residues in Arabidopsis RCA to determine whether removal of redox regulation or the α-subunit itself is essential for the enhancement of plant growth in a changing light environment.  Phosphorylation of RCA will focus on the Thr-78 phosphosite; site-specific antibodies will be developed to monitor this site in relation to changes in light intensity and directed mutagenesis will be used to probe function in vivo. 

2.2 Electron transport and regulation of the Calvin-Benson cycle. The focus is on the interaction of carbon metabolism and cyclic electron flow. When carbon follows the G6P shunt extra ATP is likely required, resulting in cyclic electron flow around photosystem I. The Sharkey lab (MI-ABR) will test this pathway by feeding ¹⁴C-labeled glucose labeled in either the C1 or C2 position. Label in the C1 position is lost if the glucose enters the Calvin-Benson cycle through the oxidative branch of the pentose phosphate pathway, but label in the C2 position should be retained. The Cousins lab (WA-AES) will measure gas exchange and chlorophyll fluorescence, using a multiphase saturation flash, on fully expanded leaves with a 6400-40 Leaf Chamber Fluorometer.  Additionally, rates of cyclic electron flux will be estimated from the dark interval relaxation of the electrochromic shift at 520 nm in a custom-built spectroscope. Furthermore, measurements of O₂ and CO₂ exchange in response to O₂ concentrations in a closed leaf chamber attached to the mass spectrometer will be used to measure rates of Rubisco oxygenation and the rates of photorespiratory CO₂ release in the light. The Fritschi lab (MO-AES) will using diversity panels and bi-parental mapping populations to investigate the genetics underlying chlorophyll fluorescence characteristics in soybean.  These investigations will be complemented with manipulated field and controlled environment studies on extreme phenotypes. 

2.3 C₄ and CAM CO₂ concentrating mechanisms. The focus is on the full spectrum of gene expression changes that are associated with the C₃ to CAM evolutionary progression, to be characterized by the Cushman group (NV-AES). Increased, leaf-specific, and circadian-clock controlled mRNA expression patterns will be assessed using real-time, qRT-PCR for a set of well-studied CAM marker genes (e.g., PEPC) and correlated with traditional diagnostic indicators of CAM. High-throughput Illumina sequencing (RNA-Seq) will also be used. Cis-acting elements acquired (or lost) during CAM evolution will be identified by bioinformatic pattern matching tools. Integrated transcriptomic, proteomic, and metabolomics expression pattern data sets will also be collected from wild-type and CAM-deficient mutant plants performing C₃ photosynthesis and CAM over both diel and circadian (constant temperature and light) conditions in order to assess gene family members that are recruited specifically for the CAM pathway and to evaluate the effects of storage carbohydrates on circadian clock output phasing. Selected cis-elements will be identified bioinformatically and cognate transcription factors will be tested for functional roles in controlling circadian-clock controlled mRNA expression patterns using gain-of-function and loss-of-function promoter::luciferase transient reporter assays in attached leaves. Measurements of bulk leaf carbon isotopic signature (δ¹³C) and bulk leaf oxygen enrichment above source water (Δ¹⁸O) will be used to screen for variation in water usage and carbon uptake across Setaria RILs.  Additionally, we will use direct measurements of Δ¹³C and Δ¹⁸O with a Tunable Diode Laser (TDL) coupled to a leaf gas exchange system to measure the internal conductance of CO₂ in RILs with variation in water usage and carbon uptake. 

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, lipids, and protein kinase signaling cascades. Mass spectrometry, isotopic labeling studies, and enzyme assays will be used to discern regulatory properties of photosynthate portioning. 

3.1 Metabolic control. The focus is on processes regulating starch biosynthesis and degradation in relation to assimilate transport. Studies by the Giroux lab (MT-AES) address the degree to which leaf and seed starch biosynthesis control whole plant growth and productivity. They will test transgenic cereals (rice and wheat) that over express leaf, seed, or both leaf and seed starch biosynthesis, as well as wild-type and leaf starch knockout rice for alterations in transcripts and metabolites. Metabolites and transcripts associated with differences in physiological measurements will be identified throughout the photoperiod as well as throughout development. The Allen lab (MO-ARS) will assess central carbon metabolism and flux in oilseeds, primarily soybean and Camelina, using metabolite measurements with mass spectrometry and isotopic labeling studies. The Jagadish lab (KS-AES) will investigate genetic factors regulating dark respiration responses to increased night temperature in wheat. The Huber group (IL-ARS) will determine whether phosphorylation of key enzymes, including nitrate reductase and sucrose phosphate synthase is regulated in vivo by oxidation of methionine residues near regulatory phosphorylation sites. The Okita group (WA-AES) will examine the roles of AGPase subunits in enzyme catalysis and allosteric regulation. Substrate binding properties will be determined using Isothermal Titration Calorimetry. The catalytic and potential regulatory properties of Pho1 will be discerned by determining the crystal structure of rice Pho1 complex with a malto-oligosaccharide and the effector ADP-glucose. 

3.2 Sugar-mediated signaling mechanisms. The focus is on roles sugars play in signaling in plants. To determine the roles of specific factors in sugar signaling, the Jang group (OH-AES) will dissect the relationship between Arabadopsis thaliana Tandem Zinc Finger1 and the Mitogen Activated Protein Kinase cascade, an evolutionarily conserved stress signal transduction pathway. The Gillaspy lab (VA-AES) studies two sugar/energy sensing systems, the SnRK1 protein kinase, which functions as an energy and stress sensor, and inositol polyphosphates that are hypothesized to act in signaling the sugar/energy status of the plant cell. During the next five years work in Arabidopsis will focus on further defining components of these signaling systems. 

3.3 Transformation of assimilates into high energy or high value compounds. The focus is on conversion of carbon fixed from photosynthesis into high-value products. The Benning lab (MI-ABR) will use, and has long standing experience applying, a combination of genetic, molecular and biochemical approaches to functionally dissect the relevant aspects of lipid metabolism in plants and algae. The focus will initially be on the algal model Chlamydomonas and the plant Arabidopsis. The Melis lab (CA-AES) will work to enhance photosynthate partitioning toward specific terpene hydrocarbons. This will be implemented with model cyanobacterial (Synechocystis) and microalgal (Chlamydomonas) strains. Recombinant DNA constructs, and culturing conditions were described (25, 26, 38-40). DNA constructs and transformation strategies will be designed to enhance carbon flux through the MEP and alternative heterologous pathways.  The effect of a heterologous terpene carbon sink on the light reactions of photosynthesis will be assessed. 

4. Develop strategies to overcome limitations to photosynthetic productivity caused by developmental and environmental factors. The scope of Objective 4 includes plant productivity responses to abiotic stresses (nutrient and water deficits, chilling, heat and salinity stresses) and stress-related signal transduction. In this objective, molecular and field approaches will be utilized. 

4.1 Abiotic stress responses. The focus is on responses to nutrient and water deficits as well as cold, heat and salt stresses, which affect multiple components of primary productivity. Investigations at scales ranging from field to molecular scales will address a range of responses to abiotic stresses.

Nitrogen deficits: IL-AES will evaluate multiple commercial hybrids with different genetic backgrounds, grown in three locations across Illinois, with three nitrogen fertilizer levels (0, 60, and 280 lb N/acre), at three population intensities (32,000, 38,000, and 44,000 plants/ acre), at two different row spacing (30” versus 20”) and four replications; results will be characterized with ‘workhorse’ and ‘racehorse’ indices.  

Temperature stress: MI-ABR will investigate the effects of temperature on proton conductance (pmf), stromal redox status, and PSII and PSI function, using mutants that vary in membrane properties. KS-AES will utilize physiological techniques such as canopy temperature measurements, gas exchange instrumentation, leaf fluorescence, pollen viability, seed-set percentage, and harvest index to quantify effects of heat stress using growth chambers and unique field-based tents, and develop remote sensing methods for assessing stress responses. Samples collected across these spatial scales will be used for a range of biochemical estimations (e.g., lipidomics, reactive oxygen species, sugar, and carbohydrate content, etc.). MO-AES will investigate photosynthetic light reactions in soybean in responses to high temperature under modulated field experiments and in controlled environments.

Salinity stress: NV-AES will create a transcriptome sequence database for all tissues from Camelina sativa plants growing under a combined stress regime including salinity.  By transforming C. sativa with potential stress protection genes, the limits of abiotic stress tolerance of Camelina (e.g., drought, heat, cold, salinity) will be established.

Drought and water-use efficiency:  KS-AES will conduct field studies using rain out shelters to evaluate genotypes of sorghum, wheat, and soybean for tolerance to drought stress. To assess drought stress, canopy conductance will be inferred from digital images of vegetative indices and thermal irradiance. NV-AES will use transcriptomics and transformation to determine and manipulate drought tolerance in C. sativa.  The Harper lab will use multiple strategies to improve biomass, water-use efficiency, and salt tolerance in crop plants such as C. sativa and soybean. This project will involve measurements of photosynthesis and whole plant physiology, and will be done in collaboration with the Cushman (NV-AES) and Huber (IL-ARS) labs.

4.2 Stress signal transduction. The focus is on the role of signal transduction in stress sensing and tolerance responses, to be studied by NV-AES, OH-AES and MS-AES. The MS-AES will identify the phosphoproteins regulated by ABA-activated protein kinases using T-DNA insertion knockout mutants and RNAi knockdown lines and then verify them as substrates of the ABA-activated protein kinases. The NV-AES will identify phosphorylation events that are associated with abiotic stress responses by investigating the role of CPKs in the phospho-regulation of plant development and metabolism, particularly the involvement of phospho-regulation of protein-protein interactions.  The long-term goal is to engineer plants with improved stress tolerance by altering specific phosphorylation sites, or modifying an entire CPK signaling pathway.  This project will involve collaborations with the Cushman and Harper labs (NV-AES), the Huber group (IL-ARS), and complement lipid signaling work by the Gillaspy lab (VA-AES). The central hypothesis for the Jang lab (OH-AES) is that Tandem Zinc Finger (TZF) proteins play key roles in integrating sugar/hormone signal transduction and RNA regulation in gene expression. Specific objectives of this research are to determine: 1) The roles of TZF proteins in sugar, hormone, and stress signal transduction; 2) Molecular mechanisms underlying the effects of TZF proteins in gene expression; and 3) Use TZF proteins as novel tools in enhancing growth and stress tolerance in crop plants. Molecular, cellular, and biochemical analyses will be conducted to determine if TZF proteins are phosphorylated by MAPKs. Global identification of soybean TZF protein target genes by expression profiling will also be conducted.

Measurement of Progress and Results

Outputs

• Collectively, the research will produce data related to thylakoid membrane architecture; Rubisco activase; organic acid and lipid metabolism; photosynthate partitioning; stress-dependent phosphorylation sites; and photosynthetic responses to heat, chilling, and drought stress.
• Information developed by the project will pertain to thylakoid-inner envelope membrane connectivity, regulation of Rubisco activase, algal quiescence and triacylglycerol or terpene accumulation, the interface of RNA regulation and signal transduction, and maize hybrid response to fertility and plant spacing factors.
• Biological materials produced by the project will include a variety of DNA plasmids expressing photosynthetically relevant genes and gene fusion constructs; gene fusion constructs and synthetic operon configuration permitting high levels of heterologous transgenes; transgenic rice and Arabidopsis and Camelina cultivars with enhanced drought, heat and saline stress tolerance.

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 Rubisco activase could increase CO2 assimilation over the growing season, with corresponding yield increase.
• Insights into algal cell growth and photosynthesis, quiescence, 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 (e.g., tandem zinc finger-dependent pathways) function can support strategies to engineer plants with improved stress tolerance, sustaining agricultural productivity.

Milestones

(2018):DNA plasmids expressing photosynthetically relevant genes and gene fusion constructs will be developed (1, 2, 3, 4; CA-AES, IL-ARS, MI-ABR, MS-AES, NE-AES, NV-AES, OH-AES, VA-AES, WA-AES). The single RCA gene in rice will be engineered as required for redox regulation, using the CRISPR-Cas9 system (2.1, IL-ARS). The G6P shunt pathway be tested by feeding 14C-labeled glucose labeled in either the C1 or C2 position (2.2, MI-ABR, WA-AES). Integrated transcriptomic, proteomic, and metabolomics expression pattern data sets will be collected from wild-type and CAM-deficient mutant plants (2.3, NV-AES).

(2019):The biosynthesis and specific function of 16:1trans in phosphatidylglycerol and the role of oligogalactolipids in chloroplast membrane will be assessed (1.1, NE-AES, MI-ABR). Experiments will establish whether removal of RCA redox regulation increases photosynthetic induction following transfer from low light to high light (2.1, IL-ARS). Drought stress will be quantified by inferring canopy conductance from digital images of vegetative indices and thermal irradiance (4.1, KS-AES). Temperature effects on proton conductance (pmf), stromal redox status, and PSII and PSI function, will be assessed using mutants that vary in membrane properties (4.1, MI-ABR). Heat stress will be quantified by canopy temperature, gas exchange, leaf fluorescence, pollen viability, seed-set percentage, and harvest index (4.1, KS-AES). The role of phosphorylation events in abiotic stress responses will be assessed, including the involvement of phospho-regulation of protein-protein interactions (4.2, IL-ARS, NV-AES, VA-AES).

(2020):A functional characterization of the photosynthetic apparatus will include changes on the mesocopic level (i.e., the organization of many protein complexes within the membrane (1.2, WA-AES). Diel regulation of RCA activity involving protein phosphorylation will be investigated (2.1, IL-ARS). Functional roles of transcription factors in circadian-clock controlled mRNA expression patterns will be tested using gain-of-function and loss-of-function promoter::luciferase transient reporter assays in attached leaves of wild-type and CAM-deficient mutant plants (2.3, NV-AES). Several studies will assess starch biosynthesis and degradation in relation to assimilate transport (IL-ARS, KS-AES, MT-AES, WA-AES). Work in Arabidopsis will focus on further defining components of sugar-mediated signaling systems, including TZF and SnRK1 proteins (3.2, OH-AES, VA-AES). Gene fusion constructs and synthetic operon configuration permitting high levels of heterologous transgenes will be developed (3.3, CA-AES).

(2021):Studies will assess how CO2 capture in the Calvin-Benson cycle is regulated and connected with electron transport regulation as well as CO2 supply (2.2, MI-ABR, WA-AES). The potential to enhance photosynthate partitioning toward triacylglycerol or specific terpene hydrocarbons and alternative heterologous pathways will be assessed (3.3, MI-ABR, CA-AES). Commercial maize hybrids, differing in genetic backgrounds and growing conditions (locations, nitrogen fertilizer levels, population intensities, and row spacing), will be characterized with ‘workhorse’ and ‘racehorse’ indices (4.1, IL-AES).

(2022):Alternate chloroplast membrane structures in Arabidopsis thaliana will be assessed using multiple fluorescent visualization systems (1.2, NE-AES). Increased, leaf-specific, and circadian-clock controlled mRNA expression patterns will be assessed using real-time, qRT-PCR for a set of well-studied CAM marker genes (e.g., PEPC; 2.3, NV-AES). The limits of abiotic stress tolerance of Camelina (e.g., drought, heat, cold, salinity) will be assessed by transforming C. sativa with potential stress protection genes (4.1, NV-AES). The roles, which CPK and TZF proteins play in integrating sugar/hormone signal transduction and RNA regulation in gene expression will be assessed (4.2, NV-AES, OH-AES, VA-AES). Transgenic rice and Arabidopsis and Camelina cultivars with enhanced drought, heat and saline stress tolerance will be developed (IL-ARS, NV-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-9 (Conservation and Utilization of Plant Genetic Resources), NC-1203 (Lipids in Plants) and S-1069 (Research and Extension for Unmanned Aerial Systems Applications in Agriculture and Natural Resources). 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 NIFA Representative.

Literature Cited

1. Al-Dous, E. K., B. George, M. E. Al-Mahmoud, M. Y. Al-Jaber and others. 2011. De novo genome sequencing and comparative genomics of date palm (Phoenix dactylifera). Nat.Biotechnol. 29:521-527


2. Alkayal, F., R. L. Albion, R. L. Tillett, L. T. Hathwaik and others. 2010. Expressed sequence tag (EST) profiling in hyper saline shocked Dunaliella salina reveals high expression of protein synthetic apparatus components. Plant Sci. 179:437-449


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


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


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


6. 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


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


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


9. 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). Physiol. Plant. doi:10.1111/ppl.12485


10. Ananieva, E. A. and G. E. Gillaspy. 2009. Switches in nutrient and inositol signaling. Plant Signal Behav 2009/10/02:304-306


11. Ananieva, E. A., G. E. Gillaspy, A. Ely, R. N. Burnette and others. 2008. Interaction of the WD40 domain of a myoinositol polyphosphate 5-phosphatase with SnRK1 links inositol, sugar, and stress signaling. Plant Physiology 148:1868-1882


12. Baena-Gonzalez, E., F. Rolland, J. M. Thevelein, and J. Sheen. 2007. A central integrator of transcription networks in plant stress and energy signalling. Nature 2007/08/03:938-942


13. 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


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. Bender, RR, JW Haegele, ML Ruffo, FE Below. Nutrient uptake, partitioning, and remobilization in modern, transgenic insect-protected maize hybrids. Agron J 105:161-170.


16. Benning, C. 2009. Mechanisms of lipid transport involved in organelle biogenesis in plant cells. Annu.Rev.Cell Dev.Biol. 25:71-91


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


18. Bogamuwa, S.P., J.C. Jang. 2014. Tandem CCCH zinc finger proteins in plant growth, development and stress response, Plant Cell Physiol, 55: 1367-1375.


19. Boomsma CR, JB Santini, M Tollenaar, TJ Vyn. 2009. Maize morphological responses to intense crowding and low nitrogen availability: An analysis and review.  Ag. J. 101: 1426-1452.


20. Bourgis, F., A. Kilaru, X. Cao, G. F. Ngando-Ebongue and others. 2011. Comparative transcriptome and metabolite analysis of oil palm and date palm mesocarp that differ dramatically in carbon partitioning. Proc. Natl. Acad.Sci. U.S.A 108:12527-12532


21. 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


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


23. Chang, R. L., L. Ghamsari, A. Manichaikul, E. F. Hom and others. 2011. Metabolic network reconstruction of Chlamydomonas offers insight into light-driven algal metabolism. Mol.Syst.Biol. 7:518


24. Charuvi D, 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.


25. Chaves JE, Kirst H, Melis A. 2015. Isoprene production in Synechocystis under alkaline and saline growth conditions. J Appl Phycol 27:1089–1097 DOI: 10.1007/s10811-014-0395-2


26. 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


27. Cheng, L., R. Zhou, E. J. Reidel, T. D. Sharkey and others. 2005. Antisense inhibition of sorbitol synthesis leads to up-regulation of starch synthesis without altering CO₂ assimilation in apple leaves. Planta 220:767-776


28. Cushman, J. C., S. Agarie, R. L. Albion, S. M. Elliot and others. 2008. Isolation and characterization of mutants of common ice plant deficient in crassulacean acid metabolism. Plant Physiol 147:228-238


29. Cushman, J. C., R. L. Tillett, J. A. Wood, J. M. Branco and others. 2008. Large-scale mRNA expression profiling in the common ice plant, Mesembryanthemum crystallinum, performing C3 photosynthesis and Crassulacean acid metabolism (CAM). J.Exp.Bot 59:1875-1894


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

30A. Dhanapal, A.P., J.D. Ray, S.K. Singh, V. Hoyos-Villegas, J.R. Smith, L.C. Purcell, C.A. King, F.B. Fritschi. 2015. Association mapping of total carotenoids in diverse soybean genotypes based on leaf extracts and high-throughput canopy spectral reflectance measurements. PLOS ONE, DOI:10.1371/jounal.pone.0137213

30B. Dhanapal, A.P., J.D. Ray, S.K. Singh, V. Hoyos-Villegas, J.R. Smith, L.C. Purcell, and F.B. Fritschi. 2016. Genome-wide association mapping of soybean chlorophyll traits based on canopy spectral reflectance and leaf extracts. BMC Plant Biology 16:174.  DOI 10.1186/s12870-016-0861-x

31. Djanaguiraman, M., P. V. Prasad, and M. Seppanen. 2010. Selenium protects sorghum leaves from oxidative damage under high temperature stress by enhancing antioxidant defense system. Plant Physiol Biochem. 48:999-1007


32. Djanaguiraman, M., J. A. Sheeba, D. D. Devi, U. Bangarusamy and others. 2010. Nitrophenolates spray can alter boll abscission rate in cotton through enhanced peroxidase activity and increased ascorbate and phenolics levels. J Plant Physiol 167:1-9


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


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


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


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


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


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


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


40. Formighieri C, Melis A. 2015. A phycocyaninphellandrene 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


41. 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. Biol. Chem. 276(34):31806.


42. Gao J, 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.


43. Genkov, T., M. Meyer, H. Griffiths, and R. J. Spreitzer. 2010. Functional hybrid rubisco enzymes with plant small subunits and algal large subunits: engineered rbcS cDNA for expression in Chlamydomonas. J Biol Chem 285:19833-19841


44. Gentry, L. E., M. B. David, F. E. Below, T. V. Royer and others. 2009. Nitrogen mass balance of a tile-drained agricultural watershed in East-Central Illinois. J Environ.Qual. 38:1841-1847


45. Gentry, LF, ML Ruffo, FE Below. 2013. Identifying factors controlling the continuous corn yield penalty. Agron J. 105:295-303. doi:10.2134/agronj2012.0246


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


47. Godfray, H. C., J. R. Beddington, I. R. Crute, L. Haddad and others. 2010. Food security: the challenge of feeding 9 billion people. Science 327:812-818


48. 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


49. He, H. and J. Li. 2008. Proteomic analysis of phosphoproteins regulated by abscisic acid in rice leaves. Biochem.Biophys.Res.Commun. 371:883-888


50. Heffner, E. L., M.R. Sorrels, and J-L. Jannink. 2009. Genomic selection for crop improvement. Crop Science 49:1-12.
    
    50A. Herritt, M., A.P. Dhanapal, and F.B. Fritschi. 2016. Identification of genomic loci associated with the photochemical reflectance index by genome-wide association study in soybean. The Plant Genome 9, DOI: 10.3835/plangenome2015.08.0072.

51. Henderson, J. N., A. M. Kuriata, R. Fromme, M. E. Salvucci and others. 2011. Atomic resolution X-ray structure of the substrate recognition domain of higher plant Rubisco activase. J.Biol.Chem.


52. 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.


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


54. 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


55. 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


56. 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


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


58. 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


59. Jang, J. C., P. Leon, L. Zhou, and J. Sheen. 1997. Hexokinase as a sugar sensor in higher plants. Plant Cell. 1997/01/01:5-19


60. Jang, J. C. and J. Sheen. 1997. Sugar sensing in higher plants. Trends Plant Sci. 2:208-214


61. Jarvis 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.


62. Kang, S.G., J. Price, P.C. Lin, J.C. Hong, J.C. Jang. 2010. The Arabidopsis bZIP1 transcription factor is involved in sugar signaling, protein networking, and DNA binding, Mol Plant, 3:361-373.


63. Ke, Y., G. Han, H. He, and J. Li. 2009. Differential regulation of proteins and phosphoproteins in rice under drought stress. Biochem.Biophys.Res.Commun. 379:133-138


64. 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


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


66. Kirchhoff H. 2014. 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.


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


68. Kirchhoff H, 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.


69. Klaus D, 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.


70. Koch, K. E. 1996. Carbohydrate-modulated gene expresion in plants. Annu.Rev.Plant Physiol Plant Mol.Biol. 47:509-540


71. Koch, K. E., K. D. Nolte, E. R. Duke, D. R. McCarty and others. 1992. Sugar levels modulate differential expression of Maize sucrose synthase genes. Plant Cell 4:59-69


72. Li, C., M. E. Salvucci, and A. R. Portis, Jr. 2005. Two residues of rubisco activase involved in recognition of the Rubisco substrate. J.Biol.Chem. 280:24864-24869


73. Li, H. M. and C. C. Chiu. 2010. Protein transport into chloroplasts. Annu.Rev.Plant Biol. 61:157-180


74. Li, J., X. Q. Wang, M. B. Watson, and S. M. Assmann. 2000. Regulation of abscisic acid-induced stomatal closure and anion channels by guard cell AAPK kinase. Science 287:300-303


75. Li, S., S.M. Assmann and R. Albert. 2006. Predicting essential components of signal transduction networks: A dynamic model of guard cell abscisic acid signaling. PLoS Biol. 4(10): e312. DOI: 1371/journal.pbio.0040312


76. 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


77. Lin J.S.,J. Nazarenus, J.L. Frey, X. Liang, M.A. Wilson and J.M Stone. 2011. A Plant DJ-1 Homolog Is Essential for Arabidopsis thaliana Chloroplast Development. PLoS One 6(8) e23731.


78. Liu, B., and Benning, C. 2013. Lipid metabolism in microalgae distinguishes itself. Curr Opin Biotechnol 24, 300-309.


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


80. Lobell, D. B., W. Schlenker, and J. Costa-Roberts. 2011. Climate trends and global crop production since 1980. Science 333:616-620


81. Lu, Y., L. J. Savage, I. Ajjawi, K. M. Imre and others. 2008. New connections across pathways and cellular processes: industrialized mutant screening reveals novel associations between diverse phenotypes in Arabidopsis. Plant Physiol 146:1482-1500


82. 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


83. 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


84. McCarty, D. R., A. M. Settles, M. Suzuki, B. C. Tan and others. 2005. Steady-state transposon mutagenesis in inbred maize. Plant J. 44:52-61


85. Melis A. 2013. Carbon partitioning in photosynthesis. Curr Opin Chem Biol. 17:453–456; http://dx.doi.org/10.1016/j.cbpa.2013.03.010


86. Miller, R., Wu, G., Deshpande, R.R., Vieler, A., Gartner, K., Li, X., Moellering, E.R., Zauner, S., Cornish, A.J., Liu, B., Bullard, B., Sears, B.B., Kuo, M.H., Hegg, E.L., Shachar-Hill, Y., Shiu, S.H., and Benning, C. 2010. Changes in transcript abundance in Chlamydomonas reinhardtii following nitrogen deprivation predict diversion of metabolism. Plant Physiol 154, 1737-1752.


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


88. Moellering, E.R., and Benning, C. 2011. Galactoglycerolipid metabolism under stress: a time for remodeling. Trends Plant Sci 16, 98-107.


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


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


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


92. Ott, C. M., B. D. Smith, A. R. Portis, Jr., and R. J. Spreitzer. 2000. Activase region on chloroplast ribulose-1,5-bisphosphate carboxylase/oxygenase. Nonconservative substitution in the large subunit alters species specificity of protein interaction. J.Biol.Chem. 275:26241-26244


93. Parry, M. A., M. Reynolds, M. E. Salvucci, C. Raines and others. 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. Price, J., A. Laxmi, S.K. St Martin, J.C. Jang. 2004 Global transcription profiling reveals multiple sugar signal transduction mechanisms in Arabidopsis, Plant Cell, 16: 2128-2150.


96. Price, J., T. Li, S. Kang, J. Na, J.C. Jang. 2003. Mechanisms of glucose signaling during germination of Arabidopsis thaliana, Plant Physiol, 132: 1424-1438.


97. Puthiyaveetil S, 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.


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


99. 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


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


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. Ramos, A. A., J. Polle, D. Tran, J. C. Cushman and others. 2011. The unicellular green alga Dunaliella salina as a model for abiotic stress tolerance: Genetic advances and future prospects. Algae 26:3-20.


103. Reynolds, M., D. Bonnett, S. C. Chapman, R. T. Furbank and others. 2011. Raising yield potential of wheat. I. Overview of a consortium approach and breeding strategies. J.Exp.Bot 62:439-452


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


105. Ruffo, M., LF Gentry, AS Henninger, JR Seebauer, FE Below. 2015. Evaluating management factor contributions to reduce corn yield gaps. Agron J. 107:495-505.


106. 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.


107. Schrader, S. M., K. R. Kleinbeck, and T. D. Sharkey. 2007. Rapid heating of intact leaves reveals initial effects of stromal oxidation on photosynthesis. Plant Cell Environ. 30:671-678


108. Seebauer, J. R., G. W. Singletary, P. M. Krumpelman, M. L. Ruffo and others. 2010. Relationship of source and sink in determining kernel composition of maize. Journal of Experimental Botany 61:511-519


109. Settles, A. M., D. R. Holding, B. C. Tan, S. P. Latshaw and others. 2007. Sequence-indexed mutations in maize using the UniformMu transposon-tagging population. BMC Genomics 8:116


110. Sharkey, T. D., M. Laporte, Y. Lu, S. Weise and others. 2004. Engineering plants for elevated CO(2): a relationship between starch degradation and sugar sensing. Plant Biol.(Stuttg) 6:280-288


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


112. Sheen, J., L. Zhou, and J. C. Jang. 1999. Sugars as signaling molecules. 2:410-418


113. Silvera, K., K. M. Neubig, W. M. Whitten, N. H. Williams and others. 2010. Evolution along the Crassulaceaen continuum. Funct.Plant Biol. 37:995-1010


114. Silvera, K., L. S. Santiago, J. C. Cushman, and K. Winter. 2010. Incidence of Crassulaceaen acid metabolism in Orchidaceae derived from carbon isotope ratios: A checklist of the flora of Panamaand Costa Rica. Bot.J.Linnean Soc. 163:194-222


115. Silvera, K., L. S. Santiago, J. C. Cushman, and K. Winter. 2009. Crassulacean acid metabolism and epiphytism linked to adaptive radiations in the Orchidaceae. Plant Physiol 149:1838-1847


116. Somerville, C., H. Youngs, C. Taylor, S. C. Davis and others. 2010. Feedstocks for lignocellulosic biofuels. Science 329:790-792


117. Spreitzer, R. J. and M. E. Salvucci. 2002. Rubisco: Structure, regulatory interactions, and possibilities for a better enzyme. Annu.Rev.Plant Biol. 53:449-475


118. Steichen, J. M., R. V. Petty, and T. D. Sharkey. 2008. Domain characterization of a 4-alpha-glucanotransferase essential for maltose metabolism in photosynthetic leaves. J.Biol.Chem. 283:20797-20804


119. 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


120. Sugden, C., P. G. Donaghy, N. G. Halford, and D. G. Hardie. 1999. Two SNF1-related protein kinases from spinach leaf phosphorylate and inactivate 3-hydroxy-3-methylglutaryl-coenzyme A reductase, nitrate reductase, and sucrose phosphate synthase in vitro. Plant Physiology 120:257-274


121. 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


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


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


124. Tietz S, S. Puthiyaveetil, H.M. Enlow, R. Yarbrough, M. Wood and others. 2015. Functional Implications of Photosystem II Crystal Formation in Photosynthetic Membranes. Biol. Chem. 290(22):14091.


125. Tsai, C.H., Warakanont, J., Takeuchi, T., Sears, B.B., Moellering, E.R., and Benning, C. 2014. The protein Compromised Hydrolysis of Triacylglycerols 7 (CHT7) acts as a repressor of cellular quiescence in Chlamydomonas. Proc Natl Acad Sci USA 111: 15833-15838.


126. Tsai, C.H., Zienkiewicz, K., Amstutz, C.L., Brink, B.G., Warakanont, J., Roston, R., and Benning, C. 2015. Dynamics of protein and polar lipid recruitment during lipid droplet assembly in Chlamydomonas reinhardtii. Plant J 83, 650-660.


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


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


129. Vu HS, 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.


130. Weise, S. E., K. S. Kim, R. P. Stewart, and T. D. Sharkey. 2005. beta-Maltose is the metabolically active anomer of maltose during transitory starch degradation. Plant Physiol 137:756-761


131. Weise, S. E., S. M. Schrader, K. R. Kleinbeck, and T. D. Sharkey. 2006. Carbon balance and circadian regulation of hydrolytic and phosphorolytic breakdown of transitory starch. Plant Physiol 141:879-886


132. Weise, S. E., K. J. van Wijk, and T. D. Sharkey. 2011. The role of transitory starch in C(3), CAM, and C(4) metabolism and opportunities for engineering leaf starch accumulation. J.Exp.Bot 62:3109-3118


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


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


135. Xiao, W., J. Sheen, J.C. Jang. 2000. The role of hexokinase in plant sugar signal transduction and growth and development. Plant Mol Biol, 44: 451-461.


136. Xu, J., W. T. Avigne, D. R. McCarty, and K. E. Koch. 1996. A similar dichotomy of sugar modulation and developmental expression affects both paths of sucrose metabolism: Evidence from a Maize invertase gene family. Plant Cell 8:1209-1220


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


138. 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.


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


140. Zhang, R., J. A. Cruz, D. M. Kramer, M. E. Magallanes-Lundback and others. 2009. Moderate heat stress reduces the pH component of the transthylakoid proton motive force in light-adapted, intact tobacco leaves. Plant Cell and Environment 32:1538-1547


141. Zhang, R. and T. D. Sharkey. 2009. Photosynthetic electron transport and proton flux under moderate heat stress. Photosynthesis Research 100:29-43


142. Zhou, L., J.C. Jang, T.L. Jones, J. Sheen. 1998. Glucose and ethylene signal transduction crosstalk revealed by an Arabidopsis glucose-insensitive mutant, Proc Natl Acad Sci U S A, 95: 1998 10294-10299.


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

Attachments

Land Grant Participating States/Institutions

CA, IL, IN, KS, MD, MI, MO, MS, MT, NE, NV, RI, TX, WA, WI

Non Land Grant Participating States/Institutions

Danforth Plant Science Center, USDA-ARS/Missouri