Duration: 10/01/2021 to 09/30/2026

Administrative Advisor(s):

NIFA Reps:

Non-Technical Summary

Statement of Issues and Justification

The need

The North Central (NC) committee, Lipids In Plants: Improving and Developing Sustainability of Crops (or “LIPIDS of Crops”) will work together to characterize lipid-related metabolism and traits relevant for crop improvement and to develop crops with improved yield, biotic and abiotic stress tolerance, and/or nutritional and industrial qualities. In doing so, the group will directly address different Program Themes of the USDA 2020 - 2025 Science Blueprint, including Ag Climate Adaptation, Food and Nutrition Translation and Value-Added Innovations. 

Lipids play critical structural, metabolic, and regulatory roles in all aspects of plant growth and development and in responses to environmental challenges. In addition, plant oils are an important source of nutrition for humans and livestock, and their trade generates billions of dollars annually. Dependence on plant oils for renewable chemical feedstocks and biofuel uses has also increased. Despite the importance of plant lipids, many fundamental questions remain unanswered about lipid compositions, metabolism, and function, impeding improvement of crop productivity and oilseed yields and fatty acid quality for human and livestock nutrition and industrial uses.

 

The importance of the work

Each plant is a green factory that uses solar energy and carbon dioxide to produce tens of thousands of compounds. Many thousands are non-water soluble compounds called lipids; these include photosynthetic pigments that harvest sunlight, building blocks of cellular membranes, protective and structural materials, seed oils, vitamins, plant (phyto-) hormones, signaling molecules, and essential oils that attract pollinators or repel insect and microbial predators. Despite their biological and economic importance, lipids are the least explored central metabolites. Limited understanding of plant lipid metabolism and function limits our ability to increase production of food and renewable resources for the growing population. In fact, the physiological functions of many plant genes putatively encoding lipid-metabolizing enzymes are unknown and their substrates are not defined. Although many plant lipids function as hormones and signaling messengers, the regulatory roles and networks through which lipids mediate plant growth and development are woefully unclear. Answers to these questions are essential for informed crop improvement through breeding and biotechnology.

 

The technical feasibility of the research

LIPIDS of Crops will improve and extend methods for lipid characterization and measurement, identify and characterize lipid related metabolism and traits relevant for crop improvement, and develop crops with improved yield and/or functionality. Approaches for lipid analysis will include mass spectrometry, imaging approaches, and flux analysis. One hindrance to significant expansion of current knowledge of plant lipids is less-than-adequate methodology to address central questions about lipid metabolism and lipid function. LIPIDS of Crops members will combine forces and share information to improve and implement lipid analytical strategies that will advance understanding of plant metabolism, regulation, development, and stress response, and enable plant improvements to increase food, feed, and seed oil production. Approaches for identifying and characterizing lipid-related traits will include analysis of genetic variants in lipid-related genes, biochemical approaches, and physiological analyses. Development of crops with improved yield and/or functionality will be via marker-assisted breeding or transformation with the relevant genes. GInitial group members have the relevant technical capabilities and access to instrumentation to pursue all aspects of the research.

 

The advantages of doing the work as a multistate effort

The multistate effort will provide a framework to enhance collaboration and sharing of resources, expertise, and instrumentation to accelerate progress among the high concentration of plant lipid researchers in the North Central Region and colleagues across the country.

 

Likely impacts of the work

The impacts will be

• a collaborative plant lipid research group, poised to participate in new opportunities
• more efficient analytical pipeline to obtain information on lipid metabolism and traits relevant for crop improvement,
• better understanding and improvement of plant growth during development, and under biotic and abiotic stress conditions
• knowledge of seed metabolism and biology for improvement of seed oils
• improvement of crop productivity and quality for human and livestock nutrition and the bio-based economy.

Related, Current and Previous Work

Information in CRIS and in the literature has been reviewed and incorporated into this section. While individual and small group projects described in CRIS deal with lipid analysis, lipid metabolism, and seed oil improvement, there is no other project that incorporates all these themes. The multistate group will continue to work together to improve tools and understand lipid metabolism, and to bring the tools and knowledge to guide crop improvement.

 

1. Methods for lipid characterization and measurement

Lipidomic analyses

The term “lipidomics” describes comprehensive analysis of the lipids in a tissue or other sample, typically by mass spectrometry. “Lipidome” refers to the entire array of lipid molecular species present in a cell, tissue, organism, or other biological unit. In the past twenty years, mass spectrometry-based lipidomics has given rise to extensive data on the range of lipids present in the model plant Arabidopsis thaliana, including glycerolipids, sphingolipids, sterol derivatives, storage lipids, cuticular lipids, and others (Markham et al., 2007; Perera et al., 2010; Ibrahim et al., 2011; Samarakoon et al., 2012; Schrick et al., 2012; Okazaki et al., 2013; Koo et al., 2014; Li et al., 2014; Nilsson et al., 2014; Shiva et al., 2020; Gonzalez-Solis et al., 2020). Many of these data were obtained by members of this multistate group. Among LIPIDS of Crops members, the Cahoon, Durrett, Koo, Kosma, Lee, Markham, Nikolau, Wang, Welti and Yandeau-Nelson groups use mass spectrometry directly to obtain lipid compositional data, while nearly all groups use such data. Individual groups use specific technologies (types of mass spectrometers) and approaches (e.g., direct infusion or liquid chromatography) and have unique abilities to characterize specific lipids. For example, the Nikolau, Kosma, and Yandeau-Nelson groups specialize in characterization of lipids found in plant cuticles and suberin while Markham’s group specializes in sphingolipids.

While considerable information on Arabidopsis is available, comprehensive data on crop plant lipids have just begun to be assembled (Riedelsheimer et al., 2013; Narayanan et al., 2016a; 2016b; 2018; 2020) and data on most crops, as well as comprehensive data, are still lacking.  Similarly, while analytical methods have been developed for many abundant, as well as some minor, classes of lipids (e.g., Song et al., 2020), analyses of many lipid intermediates and other minor lipid species have not been developed. Additionally, available analyses have not been integrated to provide an all-inclusive view of lipid metabolism.

 

Visualizing and localizing lipids

Mass spectrometry imaging technology can be used to decipher lipid localization at high-spatial resolution directly on plant tissues (typically off the plant). The Lee group is a key developer of this technology, which can now map lipids at ~10 µm resolution (Horn et al., 2012; Lee et al., 2012; Hansen et al., 2019). The Nikolau and Yandeau-Nelson groups collaborate with Lee, using MALDI-MS to evaluate the spatial distribution of lipids (Cha et al., 2008; Korte et al., 2015; Duenas et al., 2017, Feenstra et al., 2017, Bhunia et al., 2018). In addition, the Lee and Durrett groups collaborate to quantitatively visualize unusual acetyl-TAGs synthesized in transgenic camelina seeds. The Schrick group recently developed a click chemistry method to visualize phospholipids in plant tissues, discovering an abundance of these lipids in guard cells (Paper et al., 2018).

 

Data on plant lipids

The vast majority of lipid characterization data are not currently deposited in a database.  Plant/Eukaryotic and Microbial Systems Resource (PMR: http://metnetdb.org/PMR/) is a database designed for archiving, analyzing, and disseminating metabolomic and transcriptomic data and associated metadata by the Wurtele lab (Hur et al., 2013). PMR provides for analysis and comparison within and across experiments, and transcriptomics data can be co-analyzed with metabolomics data. Currently PMR contains data from Nikolau, Welti, Yandeau-Nelson and Wurtele labs (e.g., Vu et al., 2015; Shiva et al., 2018), as well as from over 20 other US and international research groups.

 

2. Identify lipid-related mechanisms to increase agricultural resilience

Roles of lipids and lipid-related gene products

Lipids in plants form membrane bilayers and plant cuticles, store energy, particularly in seeds, and serve as signals involved in plant development, growth, and stress responses. Despite the importance of lipids in plants, large gaps remain in our knowledge of their metabolism. In the following paragraph, the state of knowledge for lipid metabolism in the model plant Arabidopsis thaliana will be described. Though there are many gaps in our knowledge of Arabidopsis lipid metabolism and the gene products involved, even less specific information about lipid metabolism is available for crop plants.  

In Arabidopsis, over 900 genes have been identified as putatively involved in lipid metabolism (http://aralip.plantbiology.msu.edu/: Li-Beisson et al., 2013, Troncoso-Ponce et al., 2013). Lipid metabolism is very dynamic with compositions changing diurnally,  in response to stress, and during development (Kilaru et al., 2012; Nakamura et al., 2014; Vu et al., 2014; Shiva et al., 2020). However, many enzymes acting at specific developmental time points are not defined, nor, in most cases, are the relative fluxes through co-existing enzymes and pathways. In fact, less than 300 Arabidopsis lipid-metabolizing gene products have been characterized biochemically or by mutant analysis, and knowledge of the rest depends primarily on DNA sequence similarities to characterized genes from other species. Beyond this, there are about 30 lipid-related biochemical activities with no proposed relationships to genes (http://aralip.plantbiology.msu.edu/; Li-Beisson et al., 2013), suggesting that additional lipid-metabolizing genes have yet to be identified. Additionally, many examples exist of genes that encode proteins with sequence similarity, but different lipid-related functionality, indicating that detailed genetic, biochemical, and physiological analysis will be required to decipher their functions. For example, 300 genes encoding lipases have been identified (i.e., lipid-hydrolyzing enzymes) (Troncoso-Ponce et al., 2013).  Current knowledge suggests that these enzymes vary in the range of substrates that they hydrolyze in planta as well as in the timing and location of their expression. For example, Wang’s group has characterized 12 phospholipase Ds (PLDs) that hydrolyze the alcohols from phospholipid head groups. Individual PLD family members play distinctive roles in germination and seed viability, seed oil composition, stress tolerance, root hair patterning, pollen tube growth, and senescence (Li et al., 2009; Kim and Wang, 2020). One particularly well-characterized PLD, phospholipase Dα1, acts in an abscisic acid-initiated signaling cascade via a protein phosphatase to trigger stomatal closure (Li et al., 2009; Li et al., 2019). Some lipases directly hydrolyse TAGs in seeds. To increase retention of TAGs in seeds, the Allen, Durrett and Dhankher groups are using RNAi and CRISPR-Cas genome editing to inactivate the SDP1 lipase in camelina and soybean seeds. To add to the complexity, interactions among various lipid metabolizing enzymes can regulate functionality. For instance, Wang and Markham demonstrated interplay between sphingolipid-metabolizing and phospholipid-metabolizing enzymes in regulation of stomatal closure in response to abscisic acid (Guo et al., 2012). Thus, lipid-metabolizing enzymes perform functions critical to proper plant growth and development, seed oil formation and its regulation, and tolerance to biotic and abiotic stressors, but much remains to be understood even in the best-studied model plant species. Members of the group will extend this knowledge to crop plants, so that lipid-related information can be used to optimize crop performance, yield, and functionality.

 

Characterization of mechanisms and functions of lipid-related traits

Virtually all members of the “LIPIDS of Crops” group are involved in characterization of lipid-related traits in plants. The Wang and Welti groups characterized the function of a number of lipases that are involved in stress and seed metabolism (Zien et al., 2001; Welti et al., 2002; Li et al., 2004; Li et al. 2006; 2011; 2013; Hong et al., 2008; 2009; Peters et al., 2010; Yang et al., 2012; Li and Wang, 2019; Li et al. 2019; Kim and Wang, 2020). Roston has characterized a galactolipid-remodeling enzyme that mitigates freezing damage and a lipid transporter (Roston et al. 2011; 2012; 2014; Vu et al., 2015). The Cahoon, Markham, and Stone groups have characterized genes involved in sphingolipid metabolism; these regulate cell growth, stomatal opening, and the induction of cell death in the plant immune response (Kimberlin et al., 2013).  Schrick’s lab is analyzing putatively lipid-binding transcription factors that regulate cell-type differentiation in plants (Schrick et al., 2004; Schrick et al., 2014) and, Schrick, Welti, and Narayanan have investigated the role of enzymes involved in production of sterol derivatives that vary in level during plant stress responses (Schrick et al., 2012; Narayanan et al., 2016; 2016a; 2018; 2020; Shiva et al, 2020; Zoong-Lwe et al., 2020), and in development of the root epidermis (Pook et al., 2017). The Nikolau, Yandeau-Nelson, and Kosma groups are working to understand the roles of cuticular and suberic lipids specifically in the crop plants, maize, and potato (Alexander et al., 2020; Bhunia et al., 2018; Campbell et al., 2019; Dennison et al., 2019; Loneman et al., 2017). Nikolau and Yandeau-Nelson are characterizing the structure, function and underlying genetic network for cuticular lipids on maize silks, which are stigmatic floral tissues that are conduits for pollen fertilization and are therefore integral to yield. Louis, Koo, Hoffmann-Benning, and their colleagues have identified and characterized proteins involved in lipid signaling in response to abiotic and biotic stressors, (Louis et al., 2010; Guellette et al., 2012; Nakata et al., 2013; Koo et al., 2014; Smith et al., 2014; Barbaglia et al., 2016; Zhang et al., 2016; Poudel et al., 2016; 2019; Yurchenko et al., 2018). The Nikolau group identified all enzymatic components that constitute the plant mitochondrial Type II fatty acid synthase system, and found that some of these components are dual localized between mitochondria and plastids (Guan et al., 2015; Guan and Nikolau, 2016; Guan et al., 2017; Fu et al., 2020a; Guan et al., 2020). Allen, Bates, Cahoon, Durrett, Nikolau, Dhankher, Thelen, and Wang are working to understand steps in lipid synthesis that are critical to filling of oil seeds (e.g., Kambhampati et al., 2020; Nam et al., 2020; Hajduch et al., 2010; Lu et al., 2011; Ngaki et al., 2012; Manandhar-Shrestha et al., 2013; Bates et al., 2014; Guo et al., 2014; Liu et al., 2015; Li et al., 2015a, Abdullah et al., 2016; 2018; Aulakh and Durrett, 2019; Ye et al., 2020a, 2020b). Additionally, Bates, Cahoon, Kosma and Durrett are characterizing activities that encode enzymes catalyzing the formation of unusual, chemically active, and valuable fatty acids for potential incorporation into crop plants (e.g., Durrett et al., 2010; Bates et al., 2014; Kim et al., 2015a; 2015b; Malik et al., 2015; Nguyen et al., 2015; Tran et al., 2017; Karki and Bates, 2018; Busta et al., 2019; Karki et al., 2019; Shockey et al., 2019). The Nikolau and Yandeau-Nelson groups have characterized a wide range of plant and bacterial fatty acyl-ACP thioesterases, which terminate the Type II fatty acid synthase catalyzed reaction, leading to the generation of fatty acids of different chain lengths (Jing et al., 2018a, 2018b).

Whereas many genes that function in central lipid metabolism have been known for some time, perhaps surprisingly, some have been discovered only recently, and the roles of many genes in major lipid synthetic pathways still are not fully understood (e.g., Lu et al., 2009; Zhang et al., 2009; Ngaki et al., 2012; Fan et al., 2014; Lager et al., 2015; Glab et al., 2016; Shockey et al., 2016; Aulakh and Durrett, 2019). In comparison, genes involved in lipid signaling have only begun to be revealed, and it is likely that a significant proportion of unstudied, putatively lipid-related, genes play signaling or membrane remodeling roles.

The plasma membrane of plant cells provides the primary barrier to bacterial, fungal, and insect pathogens and to osmotic adjustments associated with drought and freezing. In addition, membranes provide a repository of lipids that can generate signal molecules for localized or distal responses to environmental perturbations. Recognizing membranes as key targets for crop improvement, the Markham and Clemente labs have modified sphingolipid composition in sorghum and soybean to enhance cold tolerance and potentially broaden the temporal and geographic ranges for production of these crops. Rational modifications of sphingolipid metabolism are also being pursued by the Markham and Cahoon labs to improve resistance to bacterial and fungal pathogens. In addition, the Wang, Koo, and Louis labs have developed strategies for altering production of signaling lipids, such as phosphatidic acid (PA), for improved drought and insect tolerance (Li et al., 2009; Peters et al., 2010; Yang et al., 2012; Lu et al., 2013; Zhao et al., 2013; Toyota et al., 2018; Poudel et al., 2019; Kim and Wang, 2020). The Nikolau group has used tools of synthetic biology to identify novel enzymes that can be used to bioengineer organisms to produce unusual lipid molecules that do not occur naturally. These lipids include bi-functional fatty acids (Garg et al., 2016; Sturms et al., 2017) or amide-linked fatty acids (Rizhsky et al., 2016), and these lipids offer new green chemistries for industrial applications. 

Methods to characterize lipid-related traits

The members of the “LIPIDS of Crops” group use complementary biochemical, cell biology, and genetic approaches to identify the role of lipid-related genes. Analysis of insertional and chemically induced knockout and knockdown mutations and ectopic or overexpression of genes putatively associated with lipid metabolism are frequently used strategies. Mutations or changes in expression can be analyzed in terms of effects on plant physiology, development, lipid composition, and/or lipid localization. These approaches are used routinely by Bates, Cahoon, Durrett, Hoffmann-Benning, Koo, Louis, Markham, Nikolau, Roston, Schrick, Stone, Thelen, Wang, and Yandeau-Nelson groups (e.g., Li et al., 2004; Li et al., 2011; Koo et al., 2014; Li et al., 2015b; Shockey et al., 2016, 2019; Zhang et al., 2016; Aulakh and Durrett, 2019; Karki et al., 2019; Poudel et al., 2019; Regmi et al., 2020, McNinch et al., 2020; Ye et al., 2020; Hoffmann-Benning, 2021). These methods may be complemented with expression of a protein of interest in bacteria or yeast to analyze enzyme activity in vitro, as demonstrated by work in the Cahoon, Koo, Schrick, and Wang groups (e.g., Yang et al., 2011; 2012; Koo et al., 2014; Stucky et al., 2015; Zhang et al., 2016). Genome wide association of lipid-related traits can be used to identify genes potentially regulating levels of particular lipids (Riedelsheimer et al., 2013). A tool that has been very useful in discovering new pathways in central lipid metabolism,, is flux analysis, which is performed by the Allen and Bates groups (e.g., Ma et al., 2014; Allen et al., 2015). Flux analysis in combination with gene mutation or overexpression can aid in defining the role of specific gene products in lipid-metabolizing pathways and identify bottlenecks within lipid engineering (e.g., Xu et al., 2003; Bates et al., 2014; Yang et al., 2017; Karki et al., 2019; Zhou et al., 2020; Regmi et al., 2020).

 

3. Crops with improved yield and/or functionality

Traits that increase quantity of plant products

Vegetable oils, consisting primarily of triacylglycerols (TAGs), are major caloric sources in human and livestock diets and combustible fuel sources for biodiesel and jet fuel production.  Vegetable oils are also widely used as a medium for fried food preparation, and the fatty acid components of vegetable oils can serve as sustainable hydrocarbon feedstocks for industrial applications, including lubricants and nylon precursors that are currently obtained from petrochemicals (Lu et al., 2011). An increasing global demand for vegetable oils to supply these expanding markets has resulted in a critical need to increase production per acre and to develop new vegetable oil sources to supplement those currently obtained largely from seeds and fruit mesocarp (Lu et al., 2011). Research in several labs of this multi-state group has been directed at meeting the global vegetable oil demand. The Dhankher, Wang, and Clemente labs, for example, have explored approaches to enhance carbon and fatty acid flux into the synthesis of TAGs by seed-specific expression of key metabolic genes and transcription factors such as DIACYLGLYCEROL ACYLTRANSFERASE1 (DGAT1), GLYCEROL-3-PHOSPHATE DEHYDROGENASE1 (GPD1) and WRINKLED1 (WRI1) in camelina and soybean (Clemente and Cahoon, 2009; Li et al., 2013; Lu et al., 2015; Chhikara et al., 2018). Dhankher is also characterizing dual function wax synthase/diacylglycerol acyltransferase (WSD/DGAT) genes to increase cuticular wax deposition on leaf surfaces for increasing abiotic stress tolerance in plants. The Thelen lab has examined the regulation of acetyl-CoA carboxylase (ACCase), a rate-limiting enzyme that provides the malonyl-CoA building block for fatty acid synthesis, and identified novel BADC and CTI subunits to ACCase that regulate fatty acid synthesis (Salie and Thelen 2016; Salie et al. 2016; Ye et al., 2020a, 2020b). The Nikolau group has characterized enzyme systems that generate the precursors of plastidic fatty acid biosynthesis, namely acetyl-CoA (Sofeo et al., 2019; Fu et al., 2020a, 2020b) and malonyl-CoA (Shivaiah et al., 2020); these characterizations exemplify the complex metabolic network that regulates the supply of the fatty acid building blocks required for the assembly of complex lipids.  The Clemente and Cahoon labs have expressed genes for TAG biosynthesis and storage in sorghum leaves and stalks to generate vegetable oils as co-products of lignocellulosic biomass, while Dhankher group is engineering oil seed crops with novel stress-associated proteins for increasing productivity and yield under extreme environmental conditions (Dixit et al., 2018).  Moreover, the Allen and Bates group’s protocols for carbon flux analysis and measurement of biosynthetic intermediates are expected to increase the predictability of biotechnological and breeding approaches for enhancement of TAG production in seeds and other plant organs (Allen et al., 2014; Allen et al., 2015; Regmi et al., 2020; Zhou et al., 2020). Modification of fatty acid compositions and fatty acid storage forms offer opportunities for enhancing the functionality and value of vegetable oils. The Cahoon and Clemente labs, for example, have engineered soybeans to produce long-chain omega-3 polyunsaturated fatty acids (PUFAs) and carotenoids for use of soybean as a sustainable, land-based source of aquaculture feed (Clemente and Cahoon, 2009; Park et al., 2017; Konda et al., 2020). The Clemente lab has also modified levels of fatty acid unsaturation in soybean seeds to produce a vegetable oil formulation for soft-spread margarine without the need for hydrogenation (Graef et al., 2009; Park et al., 2014). Similarly, the Cahoon and Durrett labs have conducted extensive metabolic engineering in camelina to generate vegetable oils with novel fatty acid compositions or oil storage forms for use as sustainable substitutes to petroleum as fuel and industrial feedstocks (Kim et al., 2015a; 2015b; Liu et al., 2015; Nguyen et al., 2015; Bansal et al., 2018).

To complement biotechnological approaches, the Cahoon and Durrett labs are actively involved in screening mutant camelina germplasm for altered fatty acid profiles to identify breeding lines that can be used to produce oils with improved oxidative stability for food processing and biodiesel applications (Neumann et al., 2021).The Allen and Durrett labs are studying the inverse correlation between protein and lipid in seed (Kambhampati et al., 2020). The Allen and Bates labs utilize flux analysis to understand how lipid metabolism adapts to bioengineering and identify new gene targets to improve plant oil production (Bates et al., 2011, 2014; Yang et al., 2017; Regmi et al., 2020; Zhou et al., 2020).  

 

Successes enabled by LIPIDS of Crops

Joint publications: 16 (highlighted in “Literature Cited” section)

Funded collaborative grants:

    National Science Foundation: 3

    United Soybean Board: 4

    US Department of Agriculture: 3

 

Objectives

1. Improve and extend methods for lipid characterization and measurement.
   
2. Identify lipid-related mechanisms to increase agricultural resilience.
   
3. Develop crops with improved yield and/or functionality.
   

Methods

Objective 1: Improve and extend methods for lipid characterization and measurement

(Lee, Markham, Welti, Wurtele; KS AES, IA AES, NE AES, Iowa State University).

Extend and improve lipidomic analyses

We will increase knowledge of the lipidomes of crop systems, increase capabilities for quantifying crop lipids, and increase analytical throughput to support projects that require analysis of large numbers of samples. Quantitative analyses of lipid molecular species of crop plants will be improved by adding lipid molecular species of biochemical and functional importance. Direct infusion and/or lipid chromatographic approaches will continue to be employed. In general, direct infusion (e.g., Welti et al. 2002; Song et al., 2020) is simple and appropriate for analysis of lipids present at relatively high levels, whereas liquid chromatography (e.g., Markham et al., 2007; Guelette et al., 2012) adds specificity and sensitivity for less abundant lipids.  Previously our group developed internal standard mixtures that are available to the scientific community. Now, the multistate group will develop plant reference materials to provide a method for normalization of analyses. The Markham and Welti groups will measure the reference materials, including leaf and seed extracts from camelina, soybean, and sorghum, to obtain consensus compositions. Having reference standards allows amounts of every lipid molecular species in samples to be directly compared among labs and among analyses performed at different times. Quantification protocols will be shared and published. The Welti lab will continue to maintain and update a web-based tool (LipidomeDB Data Calculation Environment; Fruehan et al., 2018; Song et al., 2020) for lipidomic data processing. 

 

Lipid atlas

Welti, Markham, and Lee groups, in collaboration with Durrett, will work toward developing a semi-quantitative and semi-targeted catalog of the occurrence and amounts of lipids from Arabidopsis and crop plants. They plan to develop tools to visualize and compare lipid levels across tissues and stress conditions, with the long-range goal of integrating lipidomic data with genomic and transcriptomic data.

 

Visualize and localize lipids within tissues

Lipids will be visualized and localized in plant tissues using mass spectrometry-based visualization. Mass spectrometry imaging technology to decipher lipid localization at high-spatial resolution directly in plant tissues will continue to be developed in Lee lab. The technology will be applied to lipids in crops via collaborations with team members (Objective 2) to better understand the intra-tissue organization of lipid biosynthesis.

One major challenge in MALDI-MS visualization and localization of lipids is maintaining high sensitivity while analyzing a small sampling area. Lee lab will approach this challenge by developing on-tissue derivatization techniques. Depending on chemical functional groups, appropriate chemical reactions will be developed to enhance ionization efficiencies. Additionally, MALDI-2 (post-ionization of laser plume) will be adopted. Use of this technology can increase certain lipid ion signals up to one hundred times. In order to make the analyses more quantitative, direct ESI infusion of standard will be developed while MALDI is scanning the tissue samples. An additional challenge in MALDI-MS imaging is to identify lipids without chromatographic separation. Lee lab will develop hydrogen-deuterium exchange to improve identification in addition to current accurate mass and MS/MS measurement. Furthermore, to facilitate the localization of metabolically incorporated labels, MS imaging will be developed for 13C and/or 15N isotopomers.

 

Archiving, analysis, and dissemination of data  

The “LIPIDS of Crops” multistate group plans to utilize the Plant/Eukaryotic and Microbial Systems Resource (PMR; http://metnetweb.gdcb.iastate.edu/PMR/) at Iowa State University for archiving, analyzing, and disseminating data resulting from group activities. Welti lab will continue to maintain LipidomeDB Data Calculation Environment (http://lipidome.bcf.ku.edu:8080/Lipidomics/), a tool to process mass spectrometry data. Mass spectrometry imaging data will become available to the public once published mostly via MetaSpace (https://metaspace2020.edu/).

 

Objective 2: Identify lipid-related mechanisms to increase agricultural resilience

(Allen, Bates, Cahoon, Hoffmann-Benning, Koo, Louis, Markham, Narayanan, Nikolau, Roston, Schrick, Stone, Wang, Welti, Yandeau-Nelson; IA AES, KS AES, MI AES, MO AES, NE AES, WA AES, Donald Danforth Plant Science Center, USDA-ARS/Missouri)

In tackling Objective 2, the LIPIDS of Crops group will use the lipid analysis, lipid localization, data storage, and data analysis tools generated in Objective 1 as components of their approaches to identify and characterize lipid-related signaling molecules, metabolism, and traits in plants. Thus, improvement of the tools in Objective 1 and efficient communication to the rest of the group will speed progress of Objective 2. Further, knowledge about the function of specific genes and their products will be expanded and ultimately tested toward the end of improving crop resilience and seed oil described in Objective 3. The group will use multiple approaches to identify gene products that affect lipid changes relevant to crop resilience and oil production. Some researchers will analyze the function of genes that have been tentatively identified as lipid-related and contributing to plant resilience. The research can be grouped into three categories: (a) understanding relevant signaling pathways/interactions; (b) determining changes in lipid metabolism underlying environmental and developmental responses; and (c) investigating how alterations in lipid metabolism can be employed for crop improvement. 

 

Understanding relevant signaling pathways/interactions. 

The Hoffmann-Benning lab has identified several predicted phloem-localized lipid-interacting proteins that affect plant development in response to abiotic stress, particularly drought (Guelette et al., 2012; Benning et al., 2012). They are investigating the mechanics of  protein-lipid interactions and how these affect protein-protein interactions and stress response. One group of proteins, annexins, will be studied in collaboration with the Stone and Schrick labs. A second, phosphatidic acid binding protein that appears to confer drought tolerance will be studied for its lipid-binding properties, essential amino acids required for protein-lipid interaction, and their role in protein localization and function (Barbaglia et al., 2016). The latter can be, in part, accomplished through MALDI-imaging with Lee.

The Welti and Schrick labs will collaborate on characterization of lipid-related genes with diverse functions in development and stress responses. Schrick will analyze transcription factors with Steroidogenic Acute Regulatory protein (StAR)-related lipid transfer (START) domains (Schrick et al., 2004; Schrick et al. 2014), involved in regulation of plant development and post-translational responses to environmental cues. Wang and Schrick labs are working to define potential moonlighting functions of metabolic enzymes that control the expression of transcriptional regulators (Cai et al., 2020).

 

Determining the changes in lipid metabolism underlying environmental and developmental responses.

Markham, Cahoon, and Stone labs collaborate to analyze enzymes involved in sphingolipid metabolism, in particular, thoses of proven importance in the responses of plants to pathogens and temperature (Stone et al. 2000, Asai et al. 2000, Kimberlin et al. 2013, Kimberlin et al. 2016). Similarly, Nikolau, Bates, and Thelen will study the regulation of multienzyme complexes that produce the lipid building blocks, acetyl-CoA and malonyl-CoA and thus represent a critical response to multiple environmental factors (Guan and Nikolau, 2016; Salie et al., 2016; Ye et al., 2020a, 2020b). The Nikolau group will focus on the regulation of acetyl-CoA and malonyl-CoA precursor production in three distinct subcellular compartments (plastids, mitochondria and cytosol), each of which generates distinct pools of fatty acids (Fu et al., 2020a,b; Guan et al., 2017, 2020; Sofeo et al., 2019; Shiavaiah et al., 2020). Louis, Koo, and Hoffmann-Benning groups are continuing to identify and characterize proteins involved in lipid signaling in response to abiotic and biotic stressors (Louis et al., 2010; Guellette et al., 2012; Nakata et al., 2013; Koo et al., 2014; Smith et al., 2014; Barbaglia et al., 2016; Zhang et al., 2016; Poudel et al., 2016; 2019; Yurchenko et al., 2018; Varsani et al., 2019; Poudel et al., 2019; Grover et al., 2020; Fernández-Milmanda et al., 2020). Narayanan’s group, with Welti, will investigate lipid-related mechanisms related to heat tolerance in crops, aiming to identify lipid-related molecular markers for high throughput screening for heat tolerance (Narayanan et al., 2016a, 2016b, 2018, 2020). On the opposite end of the temperature scale, Schrick and Roston labs are collaborating to test the role of sterols in low temperature responses. Genes regulating membrane lipids moderating low-temperature tolerance will be investigated by the Roston and Markham labs (Barnes et al., 2016; Michaelson et al., 2016). The Wang and Roston labs are both interested in the changes in lipids due to diurnal cycles (Maatta et al., 2012; Cai et al., 2020; Kenchanmane Raju et al., 2018); Wang lab is studying how lipid signaling is integrated into the clock mechanism, while Roston lab is studying how lipid abundance changes on a 24-hour cycle affect thermotolerance. Yandeau-Nelson and Nikolau will identify environmental factors impacting cuticular lipids, and the protective capacities of specific cuticular lipid compositions against environmental stressors.

 

Investigating how alterations in lipid metabolism can be employed for crop improvement.

Here the emphasis is on investigating near-production features that could immediately add agricultural value. Welti and Koo will analyze a genome-wide association study conducted in Arabidopsis to identify genes that affect lipid composition. Koo is collaborating with Thelen to test genes that can increase oil in camelina and Arabidopsis and study their impact on plant resistance to insects (Yurchenko et al., 2018). Koo is collaborating with Welti to complete lipidomic analysis on high oil transgenic lines. Thelen, Koo, Allen, and Bates are working to understand metabolic constraints in plants engineered to produce higher levels of seed oil. Nikolau, and Cahoon will study enzymes involved in production of novel lipid molecules with physical and chemical properties pertinent to nutrition or industrial applications. 

To analyze the function of lipid-related genes, knock-down (e.g., RNAi), gene-edited or knock-out mutants and lines exhibiting over-expression will be analyzed for physiological function, biochemical function, and lipid compositional differences as compared to wild-type lines. Proteins will be assessed by expression in yeast or bacterial systems, as appropriate. Enzymatic activities associated with in vitro-expressed proteins will be assessed, and hypotheses about substrates and products will be developed, using a comprehensive lipidomics approach to measure lipid composition before and after the proteins are incubated with plant lipids. Substrate requirements will be ascertained more completely by incubating the enzymes with various purified lipids and combinations of purified lipids.

 

Objective 3. Develop crops with improved yield and/or functionality 

(Allen, Bates, Cahoon, Clemente, Durrett, Kosma, Markham, Nikolau, Dhankher, Thelen; KS AES, MA AES, MO AES, NE AES, NV AES, WA AES, Donald Danforth Plant Science Center, USDA-ARS/Missouri)

The multistate project is anticipated to more rapidly advance efforts to improve the quantity and quality of plant products by bringing together complementary expertise and resources not available in any one location. For example, synthetic biology approaches to increase oil or specialized-product synthesis in seeds, as well as in leaves, will be extended by systems level analyses to uncover metabolic bottlenecks. Findings from Objective 2 will provide a basic understanding of plant lipid metabolism to enable more precise and rational strategies for crop improvement. Methods developed in Objective 1 will allow a more global assessment of intended and unintended impacts of specific biochemical alterations on lipid metabolic networks to further guide biotechnological and breeding approaches for plant product enhancement.

 

Increasing the quantity and quality of plant products in camelina and soybean seeds

The expression of different lipid biosynthetic enzymes and regulatory proteins will be manipulated to increase oil production in seed. Thelen lab is studying the regulation of the committed step for fatty acid synthesis, catalyzed by acetyl-CoA carboxylase (ACCase). Novel forms of regulation have been discovered, most notably the discovery of BADC and CTI effectors; these are being leveraged through transgenics to produce plants with enhanced ACCase activity.  Such efforts have resulted in higher seed and leaf oil in both Arabidopsis and camelina. The Clemente group will transform soybean with similar constructs to determine the impact on the seed oil.

Allen and Durrett labs are working to enhance protein quality in soybeans by augmenting composition with genes aimed at increased lipid levels, maintaining protein levels, and reducing carbohydrates. Genes are also being altered to improve protein amino acid composition.

The Dhankher group will characterize candidate genes controlling bottlenecks in oil production in camelina using both overexpression and RNAi approaches. They will overexpress camelina MGAT1, PDCT1 and WRI1 genes and suppress the SDP1 lipase, in collaboration with Cahoon, Durrett, and others. Further, Dhankher group has identified novel stress-related genes that provide strong tolerance to drought and heavy metals stress in Arabidopsis. His group will transfer these genes to camelina and Brassica juncea along with TAG pathway genes to increase productivity of oilseed crops on marginal nutrient-poor soils under environmental stress conditions.

In addition to increasing the amount of oil produced in seeds, groups will collaborate to increase the synthesis of specific lipid molecules valuable for human health, animal feeds, and industrial applications. Cahoon and Clemente labs are working to develop soybean germplasm with seed oil and carotenoid compositions tailored for aquaculture feedstocks. They will introduce seed-specific transgenes to extend the long-chain omega-3 polyunsaturated fatty acid (PUFA) pathway to produce oils with the fish oil-type fatty acid docosahexaenoic acid (DHA) in soybean seed. Additional transgene stacks will target production of enhanced vitamin E antioxidants for stabilization of PUFA-rich oils and WRI1 and DGAT1 genes to address observed reductions in seed oil content that accompany omega-3 PUFA biosynthesis. Cahoon and Clemente labs will also target production of the carotenoid pigment astaxanthin for improvement of soybean as an aquaculture feedstock. They will increase current production levels of astaxanthin, which is supplemented at high costs in aquaculture feeds to obtain the consumer-desired red pigmentation. This research will also use camelina as an oilseed model to test candidate genes for higher astaxanthin purity and total production as well as to address negative seed development and germination phenotypes associated with astaxanthin production.

To develop camelina lines with increased levels of acetyl-TAGs, Durrett group will target candidate lipase genes implicated in breakdown of storage lipids. In addition, Durrett will optimize timing of acetyl-TAG production through the use of seed specific promoters,using Goldenbraid cloning technology, including genetic elements developed in the Cahoon lab, to enable easy use of promoters by other group members for their own synthetic biology projects.

Interestingly, synthesis of either astaxanthin or acetyl-TAG in camelina seeds results in negative germination phenotypes with germination delayed and emerging cotyledons pale. Cahoon and Durrett will coordinate investigation of these phenotypes to see if a common mechanism explains the similar phenotypes resulting from overexpression of very different biosynthetic enzymes. 

The multistate project will take advantage of complementary expertise available among the participants to extensively phenotype at the molecular level the impact of the biochemical manipulations described above. This synergistic approach will identify metabolic bottlenecks or other restrictions (e.g. synthesis of toxic intermediates) that limit accumulation of seed oil and/or specialized molecules. For example, using improved lipid-imaging methods developed in Objective 1, Lee group will image the localization of target molecules (e.g. acetyl-TAG) in transgenic lines to identify areas in seed tissues with low production. Bates lab will analyze metabolic flux in transgenic seeds to reveal metabolic bottlenecks limiting target molecule production. Data from such experiments will be used to design new transgenic lines engineered to overcome these bottlenecks.

Crop yields can also be increased by improved resistance to biotic and abiotic stress. With this in mind, Kosma lab is focused on comprehending molecular genetic and biochemical regulation of wound suberin deposition. Kosma lab is working with potato and camelina to improve tuber and leaf wound-healing capacity. Kosma’s group has identified 9 transcription factors (4 in Arabidopsis and 5 in potato) regulating wound suberin deposition at the level of transcription. These are being leveraged to identify further components of the networks that regulate wound suberin and generate potato and camelina lines with enhanced wound suberin deposition and resistance to opportunistic pathogens. Preliminary evidence suggests that these transcription factors may be involved in regulating membrane lipid remodeling during wounding stress. Kosma will collaborate with Welti to clarify the role of the transcription factors in wound-induced membrane lipid remodeling.

 

Increasing the quantity and quality of plant products in vegetative tissue

Clemente, Cahoon, and Thelen labs will continue collaboration to produce vegetative oils and oils with specialty fatty acids in sorghum stems. The goal of this research is to provide co-products to increase the economic viability of grass feedstocks for lignocellulosic-derived biofuels. This research will include modular assembly of transgenes to enhance carbon flux into fatty acid and TAG production, improve TAG storage, and protect vegetative TAG from lipolytic degradation. For example, the Thelen group has demonstrated that silencing the CTI gene family by CRISPR in Arabidopsis produced higher leaf TAG but had no effect on seed TAG levels. Additional studies will explore the introduction of transgenes to produce oils, e.g., acetyl-TAGs (collaborating with Durrett) to add value to vegetative TAGs. Transgenic sorghum lines will be subjected to extensive phenotypic and genotypic characterizations. Production bottlenecks observed in these analyses will provide the basis for improved genetic designs in subsequent engineering iterations in the design-build-test-learn cycle. The Clemente lab will extend vegetative oil research for improvement of feedstock value of hemp as an emerging fuel and fiber crop for the Midwest. This research will include development of a reliable Agrobacterium-based transformation protocol for hemp and use of this protocol to introduce transgene stacks for high-value vegetative oil production as a co-product for hemp fiber markets.

Measurement of Progress and Results

Outputs

• Comprehensive and improved rapid methods of lipid analysis by mass spectrometry.
• Improved methods of lipid visualization and localization in plant tissues of lipids not previously visualized.
• Lipidomic data of the entire “LIPIDS of Crops” group available to the scientific public.
• Identification and characterization of previously uncharacterized genes involved in lipid metabolism in model and crop plants, including establishment of linkages between lipid-related genes and specific aspects of oil accumulation and/or stress response and/or resilience.
• Camelina and soybean plants with seed composition optimized for one or more food, feed, fuel, or industrial applications.
• One or more transgenic crop plants with improved resilience to one or more stresses.

Outcomes or Projected Impacts

• A collaborative plant lipid research group, poised to participate in new opportunities.
• An efficient analytical pipeline for plant biologists to obtain information on lipid metabolism and lipid-related traits relevant for crop improvement.
• A better understanding and improvement of plant growth under biotic and abiotic stress conditions, which scientists can use to improve crop plants by breeding and transgenic approaches.
• Knowledge of seed metabolism and biology for improvement of quantity and quality of seed oil.
• Improvement of crop productivity and quality for human and livestock nutrition, benefiting producers, consumers, and the bio-based economy.

Milestones

(2022):Development of on-tissue chemical derivatization for mass spectrometry imaging. Identification of genes with altered expression in transgenic plants with modified lipid metabolism. Quantitative analysis of the role of surface lipids in crop resilience against biotic and/or abiotic stress. Quantitative analysis of the stress response of phloem lipid-binding proteins Characterization of the mechanism underlying subcellular localization of transcription factors in response to lipid changes. Quantification of seed composition in soybean lines engineered for reduced lipid turnover. Camelina engineered for reduced lipase activity and overexpression of acyltransferase genes. Validation of ≥3 candidate genes for enhanced astaxanthin production in model systems. Phenotypic and genotypic data for sorghum lines engineered with first iteration of a vegetative oil construct completed.

(2023):Identification of gene transcripts and/or lipids with altered levels due to changing environmental factors in wildtype, mutant, and transgenic plants. Mutagenesis of amino acids essential for protein-lipid interaction followed by lipid-binding studies and protein localization studies. Characterization of genes involved in synthesizing surface lipids protective against environmental stresses. Lipid metabolic fluxes determined for oilseeds engineered for enhanced output traits. Engineered potato and camelina generated for enhanced suberin and other wound healing targets. Oil content and composition data for camelina lines with reduced lipase activity and overexpression of acyltransferase genes. Effects of enhanced ACCase on soybean seed oil, protein, and carbohydrates determined Oil content and compositional analyses of DHA soybeans measured. Proof-of-principle for a hemp transformation protocol completed.

(2024):Development of on-tissue hydrogen-deuterium exchange to improve lipid identification. Analysis of post-translational regulation of lipid-binding transcription factors during growth and development. Characterization of gene transcripts and/or lipids with altered levels due to changing environmental factors in wildtype, mutant, and transgenic plants. Second-generation camelina lines engineered to overcome existing bottlenecks. Quantification of protein, lipid, and carbohydrate contents of soybean seed with altered protein amino acid quality. Wound-healing and lipidomic measurements of engineered potato and camelina lines completed. Field trials of DHA-producing soybean lines conducted. Metabolomics of first-iteration of improved camelina astaxanthin lines completed. Phenotypic and genotypic measurements of second-generation of sorghum vegetative oil lines completed.

(2025):Identification of lipid-related interacting proteins and underlying molecular mechanisms. Quantification of environmental (and/or genetic) perturbations on the partitioning of acyl substrates into storage and membrane lipids. Stacking of oil production and stress-related genes in camelina completed. DHA-vitamin E-stacked soybean lines tested in the field. Initial hemp transformation events for vegetative oil production completed.

(2026):Field-testing of soybeans engineered for modified carbohydrate and lipid metabolism conducted. Phenotypic analysis of second-generation potato and camelina lines for improved wound healing completed. Soybean lines with optimized astaxanthin-production genes evaluated. Third-generation of vegetative oil-enhanced sorghum characterized. Field-testing of vegetative oil hemp conducted.

Projected Participation

View Appendix E: Participation

Outreach Plan

The “LIPIDS of Crops” group will publish its results in scientific journals.  Lipidomic and transcriptomic data stemming from the work of the “LIPIDS of Crops” group will be housed in Plant/Eukaryotic and Microbial Systems Resource (metnetdb.org/PMR/). There, lipidomics and transcriptomic data can be viewed by its owners, who can use PMR’s statistical tools for data analysis; the data become available to the community upon publication of the results. “LIPIDS of Crops” members also will present their results at scientific meetings.

Method advances also will be published. Kansas Lipidomics Research Center (KLRC) has a website that includes extraction and other protocols and information. To enable the adoption of methods in other labs beyond the multistate group, KLRC personnel also share methods files that drive acquisition of data by a mass spectrometer with labs who own similar instruments.  Internal standard mixes are available for a fee that reflects the cost of the components, their quantification, and assembly.  As feasible, lipidomics methods developed during the work of the multistate group will be adapted at KLRC and the analyses will be made available to the scientific community on a fee-for-service basis.  KLRC’s fees are designed to cover instrument maintenance, technical support, materials used in analysis, and data processing.  Quantitative data are provided in spreadsheets.

Members of the multi-state group are active faculty members with postdoctoral trainees, graduate students, and undergraduate researchers in their laboratories.  As an integral part of our collaborations, we exchange postdocs and/or students among laboratories, with planned visits ranging from a few days to several weeks, to aid joint projects, to improve student training, and to exchange information and expertise among labs.

In addition, members of our group have recently served as American Society of Plant Biologists (ASPB) Midwest Section Representative (Cahoon, 2015-18), Midwest Section Chair (Schrick, 2018-19), and Northeast Section Chair (Dhankher, 2017-2018).  ASPB Midwest Section meetings are held annually and thus serve as a second meeting opportunity for many group members.  Students and postdocs from our groups are encouraged to present their work at this regional meeting.  “LIPIDS of Crops” get-togethers at the Midwest ASPB meetings foster interactions among student and postdoc members of the “LIPIDS of Crops” groups.  In addition, Yandeau-Nelson is the Chair of the 2021 Maize Genetics Meeting at which a growing number of researchers are focused on lipid research, so similar LIPIDS networking sessions can be held at that meeting.  

Several members of the “LIPIDS of Crops” group interact with farmers and students on the subject of GMO crops and other topics related to industrial applications in agriculture.  Hoffmann-Benning has provided lectures and will continue to speak  to undergraduates majoring in non-science areas at Michigan State University about the generation and use of GMOs.  Clemente regularly addresses farmers and public about the advantages of GMOs (e.g., Crossley, 2015). Schrick published a laboratory exercise that introduces the utility of plant products to students interested in STEM careers (Mukherjee et al., 2019). Kosma will continue to host workshops on plant biotechnology for Future Farmers of America highschool students and Nevada Agriculture Teachers’ Association members.

The Yandeau-Nelson team focuses on increasing participation of STEM-underrepresented groups in research and in pursuing STEM careers.  The team collaborates with Iowa State University’s Science Bound program, which is designed to increase the number of ethnically and socio-economically diverse students who earn STEM degrees.  Science Bound partners with mid-Iowa school districts to engage interested 8th-12th students from STEM under-represented backgrounds in a science-intensive program that if completed successfully, results in a tuition scholarship to attend ISU in a STEM major. Yandeau-Nelson’s team hosts the interactive “Plant’s Molecular Raincoat” module, at which students participating in Science Bound Saturdays learn about plant biology, genetics and particularly, the protective capacity of the plant cuticle. Recently, this module has been modified to be offered in a hybrid format to increase accessibility during the COVID-19 pandemic and beyond.  Moreover, the team recruits Science Bound high school students to conduct research with the team during the summers and train Science Bound undergrads as research assistants who are integral to the team’s research efforts.

Organization/Governance

The “LIPIDS of Crops” group will use the Standard Governance for multistate research activities.  These include the election of a Chair, a Chair-elect, and a Secretary. All officers will continue to be elected for three-year terms which are offset by one year to provide continuity.  Current officers are  Doug Allen, Chair; Susanne Hoffmann-Benning, Vice Chair; and Tom Clemente, Secretary.  At each annual meeting, a new Secretary is elected, the previous Vice-Chair becomes Chair, and the previous Secretary becomes Vice-Chair.  The Chair is responsible for organizing the next year’s meeting.  Administrative guidance is provided by an assigned Administrative Advisor and a CSREES Representative.

Literature Cited

(Joint publications since 2017 between LIPIDS of Crops members are marked with *)

Abdullah HM, Chhikara S, Akbari P, Schnell DJ, Pareek A, Dhankher OP (2018) Comparative transcriptome and metabolome analysis revealed the bottlenecks for increasing the seed and oil yields in transgenic Camelina sativa expressing diacylglycerol acyltransferase 1 and glycerol-3-phosphate dehydrogenase. Biotechnology for Biofuels, 11: 335

Abdullah HM, Paulose B, Schnell DJ, Pareek Ashwani, Dhankher OP (2016) Transcriptome profiling of Camelina sativa to identify genes involved in triacylglycerol biosynthesis and accumulation in developing seeds. Biotechnology for Biofuels 9:136

Alexander LE, Okazaki Y, Schelling MA, Davis A, Zheng X, Rizhsky L, Yandeau-Nelson MD, Saito K, Nikolau BJ (2020) Evaluation of the functional role of the maize Glossy2 and Glossy2-like genes in cuticular lipid deposition. bioRxiv: 2020.2002.2027.968321

Allen DK, Bates PD, Tjellstrom H (2015) Tracking the metabolic pulse of plant lipid production with isotopic labeling and flux analyses: Past, present and future. Prog Lipid Res 58: 97-120

Allen DK, Evans BS, Libourel IG (2014) Analysis of isotopic labeling in peptide fragments by tandem mass spectrometry. PLoS One 9: e91537

Asai T, Stone JM, Heard JE, Kovtun Y, Yorgey P, Sheen J, Ausubel FM (2000) Fumonisin B1-induced cell death in Arabidopsis protoplasts requires salicylate-, jasmonate- and ethylene-signaling pathways. Plant Cell 12: 1823-1835 

Aulakh K and Durrett TP (2019) The plastid lipase PLIP1 is critical for seed viability in diacylglycerol acyltransferase 1 mutant seed. Plant Physiol 180: 1962-1974

* Bansal S, Kim HJ, Na G, Hamilton ME, Cahoon EB, Lu C, Durrett TP (2018) Towards the synthetic design of camelina oil enriched in tailored acetyl-triacylglycerols with medium-chain fatty acids. J Exp Bot 69: 4395-4402

Barbaglia AM, Tamot B, Greeve V, Hoffmann-Benning S (2016) Phloem proteomics reveals new lipid-binding proteins with a putative role in lipid-mediated signaling. Frontiers in Plant Science/Plant Physiology; Research topic: How plants deal with stress: exploration through proteome investigation. Front Plant Sci 7:563

Barnes AC, Benning C, Roston RL (2016) Chloroplast membrane remodeling during freezing stress is accompanied by cytoplasmic acidification activating SENSITIVE TO FREEZING2. Plant Physiol 171: 2140-2149

Bates PD, Browse J (2011) The pathway of triacylglycerol synthesis through phosphatidylcholine in Arabidopsis produces a bottleneck for the accumulation of unusual fatty acids in transgenic seeds. Plant Journal 68: 387-399

Bates PD, Johnson SR, Cao X, Li J, Nam J-W, Jaworski JG, Ohlrogge JB, Browse J (2014) Fatty acid synthesis is inhibited by inefficient utilization of unusual fatty acids for glycerolipid assembly. Proc Natl Acad Sci USA 111: 1204-1209

Benning UF, Tamot B, Guelette BS, Hoffmann-Benning S (2012) New aspects of phloem-mediated long-distance lipid signaling in plants. Front Plant Sci 3: 53

Bhunia RK, Showman LJ, Jose A, Nikolau BJ (2018) Combined use of cutinase and high-resolution mass-spectrometry to query the molecular architecture of cutin. Plant Methods 14: 117

* Busta L, Yim WC, LaBrant EW, Wang P, Grimes L, Malyszka K, Cushman JC, Santos P, Kosma DK, Cahoon EB (2018) Identification of genes encoding enzymes catalyzing the early steps of carrot polyacetylene biosynthesis. Plant Physiol 178: 1507-1521

Cai G, Kim SC, Li J, Zhou Y, Wang X (2020). Transcriptional regulation of lipid catabolism during seedling establishment. Mol Plant 13: 984-1000

Campbell AA, Stenback KE, Flyckt K, Hoang T, Perera MAD, Nikolau BJ (2019) A single-cell platform for reconstituting and characterizing fatty acid elongase component enzymes. PLoS One 14: e0213620

Cha S, Zhang H, Ilarslan HI, Wurtele ES, Brachova L, Nikolau BJ, Yeung ES (2008) Direct profiling and imaging of plant metabolites in intact tissues by using colloidal graphite-assisted laser desorption ionization mass spectrometry. Plant J 55: 348-360

Chhikara S, Abdullah HM, Akbari P, Schnell D, Dhankher OP (2018) Engineering Camelina sativa (L.) Crantz for enhanced oil and seed yields by combining diacylglycerol acyltransferase1 and glycerol-3-phosphate dehydrogenase expression. Plant Biotechnol J 16: 1034-1045

Clemente TE, Cahoon EB (2009) Soybean oil: genetic approaches for modification of functionality and total content. Plant Physiol 151: 1030-1040

Crossley T (2015) Biotechnology: Genetic diversity is the key to crop improvement. Story on a talk by Tom Clemente. Wheat Life. http://wheatlife.org/Issues/01_WLJan15web.pdf

* Dennison T, Qin W, Loneman DM, Condon SGF, Lauter N, Nikolau BJ, Yandeau-Nelson MD (2019) Genetic and environmental variation impact the cuticular hydrocarbon metabolome on the stigmatic surfaces of maize. BMC Plant Biol 19: 430

Dixit A, Tomar P, Vaine E, Abdullah HM, Hazen S, Dhankher OP (2018). A stress-associated protein, AtSAP13, from Arabidopsis thaliana provides tolerance to multiple abiotic stresses. Plant Cell Environ 41: 1171-1185

* Duenas M, Klein A, Alexander L, Yandeau-Nelson MD, Nikolau BJ, Lee Y-J (2017) High-spatial resolution mass spectrometry imaging reveals the genetically programmed, developmental modification of the distribution of thylakoid membrane lipids among individual cells of the maize leaf.  Plant J, 89: 825-838

Durrett TP, McClosky DD, Tumaney AW, Elzinga DA, Ohlrogge J, Pollard M (2010) A distinct DGAT with sn-3 acetyltransferase activity that synthesizes unusual, reduced-viscosity oils in Euonymus and transgenic seeds. Proc Natl Acad Sci U S A 107: 9464-9469

Fan J, Yan C, Roston R, Shanklin J, Xu C (2014) Arabidopsis lipins, PDAT1 acyltransferase, and SDP1 triacylglycerol lipase synergistically direct fatty acids toward β-oxidation, thereby maintaining membrane lipid homeostasis. Plant Cell 26: 4119-4134

Fernández-Milmanda GL, Crocco CD, Reichelt M, Mazza CA, Köllner TG, Zhang T, Cargnel MD, Lichy MZ, Koo AJ, Austin AT, Gershenzon J, Ballaré CL (2020) A light-dependent molecular link between competition cues and defense responses in plants. Nature Plant 6: 223-230

* Feenstra AD, Alexander LE, Song Z, Korte AR, Yandeau-Nelson MD, Nikolau BJ, Lee YJ (2017) Spatial mapping and profiling of metabolite distributions during germination. Plant Physiol. 174, 2532-2548

Fruehan C, Johnson D, Welti R(2018) LipidomeDB Data Calculation Environment has been updated to process direct-infusion multiple reaction monitoring data. Lipids. 53:1019-1020

Fu X, Guan X, Garlock R, Nikolau BJ (2020a) Mitochondrial fatty acid synthase utilizes multiple acyl carrier protein isoforms. Plant Physiol 183: 547-557

Fu X, Yang H, Pangestu F, Nikolau BJ (2020b) Failure to maintain acetate homeostasis by acetate-activating enzymes impacts plant development. Plant Physiol 182: 1256-1271

Garg S, Rizhsky L, Jin H, Yu X, Jing F, Yandeau-Nelson MD, Nikolau BJ (2016) Microbial production of bi-functional molecules by diversification of the fatty acid pathway. Metab Eng 35: 9-20

Gląb B, Beganovic M, Anaokar S, Hao M-S, Rasmusson A, Patton-Vogt J, Banaś A, Stymne S, Lager I (2016) Cloning of glycerophosphocholine acyltransferase (GPCAT) from fungi and plants; a novel enzyme in phosphatidylcholine synthesis. J Biol Chem 291: 25066-25076

* Gonzalez-Solis A, Han G, Gan L, Li Y, Markham JE, Cahoon RE, Dunn TM, Cahoon EB (2020) Unregulated sphingolipid biosynthesis in gene-edited Arabidopsis ORM mutants results in nonviable seeds with strongly reduced oil content. Plant Cell 32, 2474-2490.

Graef G, LaVallee BJ, Tenopir P, Tat M, Schweiger B, Kinney AJ, Van Gerpen JH, Clemente TE (2009) A high-oleic-acid and low-palmitic-acid soybean: agronomic performance and evaluation as a feedstock for biodiesel. Plant Biotechnol J 7: 411-421

Grover S, Varsani S, Kolomiets MV, Louis J (2020) Maize defense elicitor, 12-oxo-phytodienoic acid, prolongs aphid salivation. Commun Integr Biol 1: 63-66

Guan X, Chen H, Abramson A, Man H, Wu J, Yu O, Nikolau BJ (2015) A phosphopantetheinyl transferase that is essential for mitochondrial fatty acid biosynthesis. Plant J 84: 718-732

Guan X, Nikolau BJ (2016) AAE13 encodes a dual-localized malonyl-CoA synthetase that is crucial for mitochondrial fatty acid biosynthesis. Plant J 85: 581-593

Guan X, Okazaki Y, Lithio A, Li L, Zhao X, Jin H, Nettleton D, Saito K, Nikolau BJ (2017) Discovery and characterization of the 3-hydroxyacyl-ACP dehydratase component of the plant mitochondrial fatty acid synthase system. Plant Physiol 173: 2010-2028

Guan X, Okazaki Y, Zhang R, Saito K, Nikolau BJ (2020) Dual-localized enzymatic components constitute the fatty acid synthase systems in mitochondria and plastids. Plant Physiol 183: 517-529

Guelette BS, Benning UF, Hoffmann-Benning S (2012) Identification of lipids and lipid-binding proteins in phloem exudates from Arabidopsis thaliana. J Exp Bot 63: 3603-3616

Guo L, Ma F, Wei F, Fanella B, Allen DK, Wang X (2014) Cytosolic phosphorylating glyceraldehyde-3-phosphate dehydrogenases affect Arabidopsis cellular metabolism and promote seed oil accumulation. Plant Cell 26: 3023-3035

Guo L, Mishra G, Markham JE, Li M, Tawfall A, Welti R, Wang X (2012) Connections between sphingosine kinase and phospholipase D in the abscisic acid signaling pathway in Arabidopsis. J Biol Chem 287: 8286-8296

Hajduch M, Hearne LB, Miernyk JA, Casteel JE, Joshi T, Agrawal GK, Song Z, Zhou M, Xu D, Thelen JJ (2010) Systems analysis of seed filling in Arabidopsis: using general linear modeling to assess concordance of transcript and protein expression. Plant Physiol 152: 2078-2087

Hansen RL, Guo Hm, Yin Y, Lee YJ (2019) FERONIA mutation induces high levels of chloroplast-localized Arabidopsides which are involved in root growth. Plant J 97, 341-351

Hoffmann-Benning S (2021) Collection and analysis of phloem lipids. In: Methods in Molecular Biology; Dorothea Bartels, Peter Dörmann eds; Springer Nature. In press.

Hong Y, Devaiah SP, Bahn SC, Thamasandra BN, Li M, Welti R, Wang X (2009) Phospholipase Dε and phosphatidic acid enhance Arabidopsis nitrogen signaling and growth.  Plant J 58: 376-387

Hong Y, Pan X, Welti R, Wang X (2008) Phospholipase Dα3 regulates Arabidopsis response to salinity and water deficits. Plant Cell 20: 803-816

Horn PJ, Korte AR, Neogi PB, Love E, Fuchs J, Strupat K, Borisjuk L, Shulaev V, Lee YJ, Chapman KD (2012) Spatial mapping of lipids at cellular resolution in embryos of cotton. Plant Cell 24: 622-636

Hur M, Campbell AA, Almeida-de-Macedo M, Li L, Jose A, Crispin M, Nikolau BJ, Wurtele ES (2013) A global approach to analysis and interpretation of metabolic data for plant natural product discovery. Nat Prod Rep 30: 565-583

Ibrahim A, Schütz AL, Galano JM, Herrfurth C, Feussner K, Durand T, Brodhun F, Feussner I (2011) The alphabet of galactolipids in Arabidopsis thaliana. Front. Plant Sci 2: 95.

Jing F, Yandeau-Nelson MD, Nikolau BJ (2018a) Identification of active site residues implies a two-step catalytic mechanism for acyl-ACP thioesterase. Biochem J 475: 3861-3873

* Jing F, Zhao L, Yandeau-Nelson MD, Nikolau BJ (2018b) Two distinct domains contribute to the substrate acyl chain length selectivity of plant acyl-ACP thioesterase. Nat Commun 9: 860

Karki N, Bates PD (2018) The effect of light conditions on interpreting oil composition engineering in Arabidopsis seeds. Plant Direct 2: e00067

Karki N, Johnson BS, Bates PD (2019) Metabolically distinct pools of phosphatidylcholine are involved in trafficking of fatty acids out of and into the chloroplast for membrane production. Plant Cell 31: 2768-2788

* Kambhampati S, Aznar-Moreno JA, Hostetler C, Caso T, Bailey SR, Hubbard AH, Durrett TP, Allen DK (2020) On the inverse correlation of protein and oil: Examining the effects of altered central carbon metabolism on seed composition using soybean fast neutron mutants. Metabolites 10: 18

Kenchanmane Raju SK, Barnes AC, Schnable JC, Roston RL (2018) Low-temperature tolerance in land plants: Are transcript and membrane responses conserved? Plant Sci 276: 73-86

Kilaru A, Tamura P, Isaac G, Welti R, Venables BJ, Seier E, Chapman KD (2012) Lipidomic analysis of N-acylphosphatidylethanolamine molecular species in Arabidopsis suggests feedback regulation by N-acylethanolamines. Planta 236: 809-824 

Kim HJ, Silva JE, Iskandarov U, Andersson M, Cahoon RE, Mockaitis K, Cahoon EB (2015a) Structurally divergent lysophosphatidic acid acyltransferases with high selectivity for saturated medium chain fatty acids from Cuphea seeds. Plant J 84: 1021-1033

Kim HJ, Silva JE, Vu HS, Mockaitis K, Nam JW, Cahoon EB (2015b) Toward production of jet fuel functionality in oilseeds: identification of FatB acyl-acyl carrier protein thioesterases and evaluation of combinatorial expression strategies in Camelina seeds. J Exp Bot 66: 4251-4265

Kim SC, Wang X (2020) Phosphatidic acid: an emerging versatile class of cellular mediators. Essays Biochem. 64: 533-546

Kimberlin AN, Majumder S, Han G, Chen M, Cahoon RE, Stone JM, Dunn TM, Cahoon EB (2013) Arabidopsis 56-amino acid serine palmitoyltransferase-interacting proteins stimulate sphingolipid synthesis, are essential, and affect mycotoxin sensitivity. Plant Cell 25: 4627-4639

* Kimberlin AN, Han G, Luttgeharm KD, Chen M, Cahoon RE, Stone JM, Markham JE, Dunn TM, Cahoon EB (2016) ORM expression alters sphingolipid homeostasis and differentially affects ceramide synthase activity. Plant Physiology 172: 889-900

* Konda AR, Nazarenus TJ, Nguyen H, Yang J, Gelli M, Swenson S, Shipp JM, Schmidt MA, Cahoon RE, Ciftci ON, Zhang C, Clemente TE, Cahoon EB (2020) Metabolic engineering of soybean seeds for enhanced vitamin E tocochromanol content and effects on oil antioxidant properties in polyunsaturated fatty acid-rich germplasm. Metab Eng 57 : 63-73

Koo AJ, Thireault C, Zemelis S, Poudel AN, Zhang T, Kitaoka N, Brandizzi F, Matsuura H, Howe GA (2014) Endoplasmic reticulum-associated inactivation of the hormone jasmonoyl-L-isoleucine by multiple members of the cytochrome P450 94 family in Arabidopsis. J Biol Chem 289: 29728-29738

Korte AR, Yandeau-Nelson MD, Nikolau BJ, Lee YJ (2015) Subcellular-level resolution MALDI-MS imaging of maize leaf metabolites my MALDI-linear ion trap-Orbitrap mass spectrometer. Anal Bioanal Chem 407: 2301-2309

Lager I, Glab B, Eriksson L, Chen G, Banas A, Stymne S (2015) Novel reactions in acyl editing of phosphatidylcholine by lysophosphatidylcholine transacylase (LPCT) and acyl-CoA:glycerophosphocholine acyltransferase (GPCAT) activities in microsomal preparations of plant tissues. Planta 241: 347-358

Lee Y-J, Perdian DC, Song Z, Yeung E, Nikolau N (2012) Use of mass-spectrometry for imaging metabolites in plants. Plant J 70: 81-95

Li J, Wang X. 2019. Phospholipase D and phosphatidic acid in plant immunity. Plant Sci. 279, 45-50.

Li L, Hur M, Lee JY, Zhou W, Song Z, Ransom N, Demirkale CY, Nettleton D, Westgate M, Arendsee Z, Iyer V, Shanks J, Nikolau B, Wurtele ES (2015a) A systems biology approach toward understanding seed composition in soybean. BMC Genomics. 16 Suppl 3: S9

Li L, Zheng W, Zhu Y, Ye H, Tang B, Arendsee ZW, Jones D, Li R, Ortiz D, Zhao X, Du C, Nettleton D, Scott MP, Salas-Fernandez MG, Yin Y, Wurtele ES (2015b) QQS orphan gene regulates carbon and nitrogen partitioning across species via NF-YC interactions. Proc Natl Acad Sci USA 112: 14734-14739

Li M, Bahn SC, Fan C, Li J, Phan T, Ortiz M, Roth MR, Welti R, Jaworski J, Wang X (2013) Patatin-related phospholipase pPLAIIIdelta increases seed oil content with long-chain fatty acids in Arabidopsis. Plant Physiol 162: 39-51

Li M, Bahn SC, Guo L, Musgrave W, Berg H, Welti R, Wang X (2011) Patatin-related phospholipase pPLAIIIβ-induced changes in lipid metabolism alter cellulose content and cell elongation in Arabidopsis.  Plant Cell 23: 1107-1123

Li M, Baughman E, Roth MR, Han X, Welti R, Wang X (2014) Quantitative profiling and pattern analysis of triacylglycerol species in Arabidopsis seeds by electrospray ionization mass spectrometry. Plant J 77: 160-17

Li M, Hong Y, Wang X (2009) Phospholipase D- and phosphatidic acid-mediated signaling in plants. Biochim Biophys Acta 1791: 927-935

Li M, Welti R, Wang X (2006) Arabidopsis phospholipase Dβ1 modulates defense responses to bacterial and fungal pathogens. Plant Physiol 142: 750-761

Li W, Li M, Zhang W, Welti R, Wang X (2004) The plasma membrane-bound phospholipase Dδ enhances freezing tolerance in Arabidopsis thaliana. Nature Biotechnol 22: 427-433

Li W, Song T, Wallrad L, Kudla J, Wang X, Zhang W. 2019. Tissue-specific accumulation of pH-sensing phosphatidic acid determines plant stress tolerance. Nat Plants. 5, 1012-1021.

Li-Beisson Y, Shorrosh B, Beisson F, Andersson MX, Arondel V, Bates PD, Baud S, Bird D, DeBono A, Durrett TP, Franke RB, Graham IA, Katayama K, Kelly AA, Larson T, Markham JE, Miquel M, Molina I, Nishida I, Rowland O, Samuels L, Schmid KM, Wada H, Welti R, Xu C, Zallot R, Ohlrogge J (2013) Acyl-lipid metabolism. Arab. Book: 11: e0161. 

Liu J, Rice A, McGlew K, Shaw V, Park H, Clemente T, Pollard M, Ohlrogge J, Durrett TP (2015) Metabolic engineering of oilseed crops to produce high levels of novel acetyl glyceride oils with reduced viscosity, freezing point and calorific value. Plant Biotechnol J 13: 858-865

* Loneman DM, Peddicord L, Al-Rashid A, Nikolau BJ, Lauter N, Yandeau-Nelson MD (2017) A robust and efficient method for the extraction of plant extracellular surface lipids as applied to the analysis of silks and seedling leaves of maize. PLoS One 12: e0180850

Louis J, Kukula K-L, Singh V, Reese JC, Jander G, Shah, J (2010) Antibiosis against the green peach aphid requires the Arabidopsis thaliana MYZUS PERSICAE INDUCED LIPASE1 gene. Plant J 64: 800-811

Lu C, Napier JA, Clemente TE, Cahoon EB (2011) New frontiers in oilseed biotechnology: meeting the global demand for vegetable oils for food, feed, biofuel, and industrial applications. Curr Opin Biotechnol 22: 252-259

Lu C, Xin Z, Ren Z, Miquel M, Browse J (2009) An enzyme regulating triacylglycerol composition is encoded by the ROD1 gene of Arabidopsis. Proc Natl Acad Sci USA 106: 18837-18842

Lu S, Bahn SC, Qu G, Qin H, Hong Y, Xu Q, Zhou Y, Wang X (2013) Increased expression of phospholipase Dalpha1 in guard cells decreases water loss with improved seed production under drought in Brassica napus. Plant Biotechnol J 11: 380-389

Lu S, Yao S, Wang G, Guo L, Zhou Y, Hong Y, Wang X (2016) Phospholipase De enhances Brassica napus growth and seed production in response to nitrogen availability. Plant Biotechnol J  14: 926-937

Ma F, Jazmin LJ, Young JD, Allen DK (2014) Isotopically nonstationary 13C flux Arabidopsis thaliana leaf metabolism due to high light acclimation. Proc Natl Acad Sci USA 111: 16967–16972

Malik MR, Yang W, Patterson N, Tang J, Wellinghoff RL, Preuss ML, Burkitt C, Sharma N, Ji Y, Jez JM, Peoples OP, Jaworski JG, Cahoon EB, Snell KD (2015) Production of high levels of poly-3-hydroxybutyrate in plastids of Camelina sativa seeds. Plant Biotechnol J 13: 675-688

Manandhar-Shrestha K, Tamot B, Pratt EP, Saitie S, Bräutigam A, Weber AP, Hoffmann-Benning S (2013) Comparative proteomics of chloroplasts envelopes from bundle sheath and mesophyll chloroplasts reveals novel membrane proteins with a possible role in C4-related metabolite fluxes and development. Front Plant Sci 4: 65

Markham JE, Jaworski JG (2007) Rapid measurement of sphingolipids from Arabidopsis thaliana by reversed-phase high-performance liquid chromatography coupled to electrospray ionization tandem mass spectrometry. Rapid Commun Mass Spectrom 21: 1304-1314

McNinch C, Chen K, Dennison T, Lopez M, Yandeau-Nelson MD, Lauter N. (2020)  A multi-genotype maize silk expression atlas reveals how exposure-related stresses are mitigated following emergence from husk leaves.  Plant Genome Oct 14:e20040.

Michaelson LV, Napier JA, Molino D, Faure J-D (2016) Plant sphingolipids: Their importance in cellular organization and adaption. Biochim Biophys Acta 1861: 1329–1335

Mukherjee, T, Lerma-Reyes, R, Thompson, K, Schrick, K (2019). Making glue from seeds and gums: Working with plant-based polymers to introduce students to plant biochemistry. Biochem Mol Biol Educ 47:468-475  

Nakamura Y, Teo NZX, Shui G, Chua CHL, Cheong W-F, Parameswaran S, Koizumi R, Ohta H, Wenk MR, Ito T (2014) Transcriptomic and lipidomic profiles of glycerolipids during Arabidopsis flower development. New Phytol 203: 310-322

Nakata M, Mitsuda N, Herde M, Koo AJ, Moreno JE, Suzuki K, Howe GA, Ohme-Takagi M (2013) A bHLH-type transcription factor, ABA-INDUCIBLE BHLH-TYPE TRANSCRIPTION FACTOR/JA-ASSOCIATED MYC2-LIKE1, acts as a repressor to negatively regulate jasmonate signaling in Arabidopsis. Plant Cell. 25: 1641-1656

Nam JW, Jenkins LM, Li J, Evans BS, Jaworski JG, Allen DK (2020) A general method for quantification and discovery of acyl groups attached to acyl carrier proteins in fatty acid metabolism using LC-MS/MS. Plant Cell 32: 820-832

Narayanan S, Tamura PJ, Roth MR, Prasad PV, Welti R (2016a) Wheat leaf lipids during heat stress: I. High day and night temperatures result in major lipid alterations. Plant Cell Environ 39: 787-803

Narayanan S, Prasad PV, Welti R (2016b) Wheat leaf lipids during heat stress: II. Lipids experiencing coordinated metabolism are detected by analysis of lipid co-occurrence. Plant Cell Environ 39: 608-617

* Narayanan S, Prasad PVV, Welti R(2018) Alterations in wheat pollen lipidome during high day and night temperature stress. Plant Cell Environ 41: 1749-1761

* Narayanan S, Zoong-Lwe ZS, Gandhi N, Welti R, Fallen B, Smith JR, Rustgi S (2020) Comparative lipidomic analysis reveals feat stress responses of two soybean genotypes differing in temperature sensitivity. Plants (Basel) 9: 457

* Neumann NG, Nazarenus TJ, Aznar-Moreno JA, Rodriguez-Aponte SA, Mejias Veintidos VA, Comai L, Durrett TP, Cahoon EB (2021) Generation of camelina mid-oleic acid seed oil by identification and stacking of fatty acid biosynthetic mutants. Ind Crop Prod 159: 113074

Ngaki MN, Louie GV, Philippe RN, Manning G, Pojer F, Bowman ME, Li L, Larsen E, Wurtele ES, Noel JP (2012) Evolution of the chalcone-isomerase fold from fatty-acid binding to stereospecific catalysis. Nature 485: 530-533

Nguyen HT, Park H, Koster KL, Cahoon RE, Shanklin J, Clemente TE, Cahoon EB (2015) Redirection of metabolic flux for high levels of omega-7 monounsaturated fatty acid accumulation in camelina seeds. Plant Biotechnol J 13: 38-50

Nilsson AK, Johansson ON, Fahlberg P, Steinhart F, Gustavsson MB, Ellerström M, Andersson MX (2014) Formation of oxidized phosphatidylinositol and 12-oxo-phytodienoic acid containing acylated phosphatidylglycerol during the hypersensitive response in Arabidopsis. Phytochemistry 101: 65-75

Okazaki Y, Otsuki H, Narisawa T, Kobayashi M, Sawai S, Kamide Y, Kusano M, Aoki T, Hirai MY, Saito K (2013) A new class of plant lipid is essential for protection against phosphorus depletion. Nat Commun 4: 1510

Paper, JM, Mukherjee, T, Schrick, K (2018) Bioorthogonal click chemistry for fluorescence imaging of choline phospholipids in plants. Plant Methods 14: 31

Park H, Graef G, Xu Y, Tenopir P, Clemente TE (2014) Stacking of a stearoyl-ACP thioesterase with a dual-silenced palmitoyl-ACP thioesterase and 12 fatty acid desaturase in transgenic soybean. Plant Biotechnol J 12: 1035-1043

* Park H, Weier S, Razvi F, Pena PA, Sims NA, Lowell J, Hungate C, Kissinger K, Key G, Fraser P, Napier JA, Cahoon EB, Clemente TE (2017) Towards the development of a sustainable soya bean-based feedstock for aquaculture. Plant Biotechnol J 15: 227-236

Perera MA, Qin W, Yandeau-Nelson M, Fan L, Dixon P, Nikolau BJ (2010) Biological origins of normal-chain hydrocarbons: a pathway model based on cuticular wax analyses of maize silks. Plant J 64: 618-632

Peters C, Li M, Narasimhan R, Roth M, Welti R, Wang X (2010) Nonspecific phospholipase C NPC4 promotes responses to abscisic acid and tolerance to hyperosmotic stress in Arabidopsis. Plant Cell 22: 2642-2659

Pook, VG, Nair, M, Ryu, KH, Arpin, JC, Schiefelbein, J, Schrick, K, DeBolt, S (2017) SCRAMBLED receptor requires UDP-Glc:sterol glucosyltransferase 80B1 in Arabidopsis roots. Sci Rep 7: 5714

Poudel AN, Zhang T, Kwasniewski M, Nakabayashi R, Saito K, Koo AJ (2016) Mutations in jasmonoyl-L-isoleucine-12-hydroxylases suppress multiple JA-dependent wound responses in Arabidopsis thaliana. Biochim Biophys Acta 1861: 1396-1408

Poudel AN, Holtsclaw RE, Kimberlin A, Sen S, Zeng S, Joshi T, Lei Z, Sumner LW, Singh K, Matsuura H, Koo AJ (2019) 12-Hydroxy-jasmonoyl-l-isoleucine is an active jasmonate that signals through CORONATINE INSENSITIVE 1 and contributes to the wound response in Arabidopsis. Plant Cell Physiol 60: 2152-2166

Regmi A, Shockey J, Kotapati HK, Bates PD (2020) Oil-producing metabolons containing DGAT1 use separate substrate pools from those containing DGAT2 or PDAT. Plant Physiol 184: 720-737

Riedelsheimer C, Brotman Y, Méret M, Melchinger AE, Willmitzer L (2013) The maize leaf lipidome shows multilevel genetic control and high predictive value for agronomic traits. Sci Rep. 3: 2479

Rizhsky L, Jin H, Shepard MR, Scott HW, Teitgen AM, Perera MA, Mhaske V, Jose A, Zheng X, Crispin M, Wurtele ES, Jones D, Hur M, Gongora-Castillo E, Buell CR, Minto RE, Nikolau BJ (2016) Integrating metabolomics and transcriptomics data to discover a biocatalyst that can generate the amine precursors for alkamide biosynthesis. Plant J 88: 775-793

Roston RL, Gao J, Murcha MW, Whelan J, Benning C (2012) TGD1, -2, and -3 proteins involved in lipid trafficking form ATP-binding cassette (ABC) transporter with multiple substrate-binding proteins. J Biol Chem. 287: 21406-21415

Roston RL, Gao J, Xu C, Benning C (2011) Arabidopsis chloroplast lipid transport protein TGD2 disrupts membranes and is part of a large complex. Plant J 66: 759-769

Roston RL, Wang K, Kuhn LA, Benning C (2014) Structural determinants allowing transferase activity in SENSITIVE TO FREEZING 2, classified as a family I glycosyl hydrolase. J Biol Chem 289: 26089-26106

Salie MJ, Zhang N, Lancikova V, Xu D, Thelen JJ (2016). A family of negative regulators targets the committed step of de novo fatty acid biosynthesis. Plant Cell 28: 2312-2325.

Salie MJ, Thelen JJ (2016) Regulation and structure of the heteromeric acetyl-CoA carboxylase. Biochim Biophys Acta. 186: 1207-1213

Samarakoon T, Shiva  S, Lowe K, Tamura P, Roth MR, Welti R (2012) Arabidopsis thaliana membrane lipid molecular species and their mass spectral analysis. Meth Mol Biol 918: 179-268

Schrick K, Bruno M, Khosla A, Cox PN, Marlatt SA, Roque RA, Nguyen HC, He C, Snyder MP, Singh D, Yadav G (2014) Shared functions of plant and mammalian StAR-related lipid transfer (START) domains in modulating transcription factor activity. BMC Biol 12: 70

Schrick K, Nguyen D, Karlowski WM, Mayer KF (2004) START lipid/sterol-binding domains are amplified in plants and are predominantly associated with homeodomain transcription factors. Genome Biol 5: R41

Schrick K, Shiva S, Arpin J, Delimont N, Isaac G, Tamura P, Welti R (2012) Steryl glucoside and acyl steryl glucoside analysis of Arabidopsis seeds by electrospray ionization tandem mass spectrometry.  Lipids 47: 185-193

Shiva S, Enninful R, Roth MR, Tamura P, Jagadish K, Welti R (2018) An efficient modified method for plant leaf lipid extraction results in improved recovery of phosphatidic acid. Plant Methods 14: 14. 

* Shiva S, Samarakoon T, Lowe KA, Roach C, Vu HS, Colter M, Porras H, Hwang C, Roth MR, Tamura P, Li M, Schrick K, Shah J, Wang X, Wang H, Welti R (2020) Leaf lipid alterations in response to heat stress of Arabidopsis thaliana. Plants (Basel) 9, 845.

Shivaiah KK, Ding G, Upton B, Nikolau BJ (2020) Non-catalytic subunits facilitate quaternary organization of plastidic acetyl-CoA carboxylase. Plant Physiol 182: 756-775

Shockey J, Regmi A, Cotton K, Adhikari N, Browse J, Bates PD (2016) Identification of Arabidopsis GPAT9 (At5g60620) as an essential gene involved in triacylglycerol biosynthesis. Plant Physiol 170: 163-179

Shockey J, Lager I, Stymne S, Kotapati HK, Sheffield J, Mason C, Bates PD (2019) Specialized lysophosphatidic acid acyltransferases contribute to unusual fatty acid accumulation in exotic Euphorbiaceae seed oils. Planta 249: 1285-1299

Smith JM, Leslie ME, Robinson SJ, Korasick DA, Zhang T, Backues SK, Cornish PV, Koo AJ, Bednarek SY, Heese A (2014) Loss of Arabidopsis thaliana dynamin-related protein 2B reveals separation of innate immune signaling pathways. PLoS Pathog 10: e1004578

Smith-Hammond CL, Swatek KN, Johnston ML, Thelen JJ, Miernyk JA (2014) Initial description of the developing soybean seed protein Lys-N(ε)-acetylome. J Proteomics 96: 56-66

* Sofeo N, Hart JH, Butler B, Oliver DJ, Yandeau-Nelson MD, Nikolau BJ (2019) Altering the substrate specificity of acetyl-CoA synthetase by rational mutagenesis of the carboxylate binding pocket. ACS Synth Biol 8: 1325-1336

Song Y, Vu HS, Shiva S, Fruehan C, Roth MR, Tamura P, Welti R (2020) A lipidomic approach to identify cold-induced changes in Arabidopsis membrane lipid composition. Methods Mol Biol. 2156:187-202. 

Stucky DF, Arpin JC, Schrick K (2015) Functional diversification of two UGT80 enzymes required for steryl glucoside synthesis in Arabidopsis. J Exp Bot 66: 189-201

Stone JM, Heard JE, Asai T, Ausubel FM (2000) Simulation of fungal-mediated cell death and selection of Arabidopsis fumonisin B1-resistant (fbr) mutants. Plant Cell 12: 1811-1822.

Sturms R, White D, Vickerman KL, Hattery T, Sundararajan S, Nikolau BJ, Garg S (2017) Lubricant properties of ω − 1 hydroxy branched fatty acid-containing natural and synthetic lipids. Tribology Letters 65: 99

Toyota M, Spencer D, Sawai-Toyota S, Jiaqi W, Zhang T, Koo AJ, Howe GA, Gilroy S (2018) Glutamate triggers long-distance, calcium-based plant defense signaling. Science 361: 1112-1115

Troncoso-Ponce MA, Cao X, Yang Z, Ohlrogge JB (2013) Lipid turnover during senescence. Plant Sci 205-206: 13-19

Tran TNT, Shelton J, Brown S, Durrett TP (2017) Membrane topology and identification of key residues of EaDAcT, a plant MBOAT with unusual substrate specificity. Plant J 92:82-94.

Varsani S, Grover S, Zhou S, Koch KG, Huang PC, Kolomiets MV, Williams WP, Heng-Moss T, Sarath G, Luthe DS, Jander G (2019) 12-Oxo-phytodienoic acid acts as a regulator of maize defense against corn leaf aphid. Plant Physiol, 179: 1402-1415

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

Vu HS, Shiva S, Roth MR, Tamura P, Zheng L, Li M, Sarowar S, Honey S, McEllhiney D, Hinkes P, Seib L, Williams TD, Gadbury G, Wang X, Shah J, Welti, R (2014) Lipid changes after leaf wounding in Arabidopsis thaliana: expanded lipidomic data form the basis for lipid co-occurrence analysis. Plant J 80: 728-743

Welti R, Li W, Li M, Sang Y, Biesiada H, Zhou H-E, Rajashekar CB, Williams TD, Wang X (2002) Profiling membrane lipids in plant stress responses. Role of phospholipase D in freezing-induced lipid changes in Arabidopsis. J Biol Chem 277: 31994-32002

Xu C, Fan J, Riekhof W, Froehlich JE, Benning C (2003) A permease-like protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J 22: 2370–2379

Yang W, Cahoon RE, Hunter SC, Zhang C, Han J, Borgschulte T, Cahoon EB (2011) Vitamin E biosynthesis: functional characterization of the monocot homogentisate geranylgeranyl transferase. Plant J 65: 206-217.

Yang WY, Zheng Y, Bahn SC, Pan XQ, Li MY, Vu HS, Roth MR, Scheu B, Welti R, Hong YY, Wang XM (2012) The patatin-containing phospholipase A pPLAIIalpha modulates oxylipin formation and water loss in Arabidopsis thaliana. Mol Plant 5: 452-460

* Yang W, Wang G, Li J, Bates PD, Wang X, Allen DK (2017) Phospholipase Dzeta enhances diacylglycerol flux into triacylglycerol. Plant Physiol 174: 110-123

* Ye Y, Fulcher YG, Sliman DJ, Day MT, Schroeder MJ, Koppisetti RK, Bates PD, Thelen JJ, Van Doren SR (2020) The BADC and BCCP subunits of chloroplast acetyl-CoA carboxylase sense the pH changes of the light–dark cycle. J Biol Chem 295: 9901-9916

Ye Y, Nikovics K, To A, Lepiniec L, Fedosejevs ET, Van Doren SR, Baud S, Thelen JJ. (2020) Docking of acetyl-CoA carboxylase to the plastid envelope membrane attenuates fatty acid production in plants.  Nature Comm.  In press

Yurchenko O, Kimberlin A, Mehling M, Koo AJ, Chapman KD, Mullen RT, Dyer JM (2018) Response of high leaf-oil Arabidopsis thaliana plant lines to biotic or abiotic stress. Plant Signal Behav 13: e1464361

Zhang M, Fan J, Taylor DC, Ohlrogge JB (2009) DGAT1 and PDAT1 acyltransferases have overlapping functions in Arabidopsis triacylglycerol biosynthesis and are essential for normal pollen and seed development. Plant Cell 21: 3885-3901

Zhang T, Poudel AN, Jewell JB, Kitaoka N, Staswick P, Matsuura H, Koo AJ (2016) Hormone crosstalk in wound stress response: wound-inducible amidohydrolases can simultaneously regulate jasmonate and auxin homeostasis in Arabidopsis thaliana. J Exp Bot 67: 2107-2120

Zhao J, Devaiah SP, Wang C, Li M, Welti R, Wang X (2013) Arabidopsis phospholipase Db1 modulates defense responses to bacterial and fungal pathogens. New Phytol 199: 228-240

* Zhou XR, Bhandari S, Johnson BS, Kotapati HK, Allen DK, Vanhercke T, Bates PD (2020) Reorganization of acyl flux through the lipid metabolic network in oil-accumulating tobacco leaves. Plant Physiol 182: 739-755

Zien CA, Wang C, Wang X, Welti R (2001) In-vivo substrates and the contribution of the common phospholipase D, PLD alpha, to wound-induced metabolism of lipids in Arabidopsis. Biochim Biophys Acta 1530: 236-248

* Zoong Lwe ZS, Welti R, Naveed S, Rustgi S, Anc D, Narayanan S (2020) Heat stress elicits remodeling in the anther lipidome of peanut. Sci Rep In press.

Attachments

Land Grant Participating States/Institutions

DE, IA, KS, LA, MA, MI, MO, NE, NV, SC, TX, WA

Non Land Grant Participating States/Institutions

Danforth Plant Science Center, Iowa State University - College of Vet Med, USDA-ARS/Missouri