Duration: 10/01/2026 to 09/30/2031

Administrative Advisor(s):

NIFA Reps:

Non-Technical Summary

Lipids constitute a large group of critical plant components necessary for crop development and growth and for plant responses to environmental challenges. Plant seed oils are lipids that are an important source of nutrition for humans and livestock; seed oil trade generates billions of dollars annually. Dependence on plant oils for chemical feedstocks and biofuels has increased. Despite the importance of plant lipids in crop productivity and to the bioeconomy, many fundamental questions about lipid compositions, metabolism, and function remain unanswered, impeding improvement of crop productivity, oilseed yields, and oil quality for human and livestock nutrition and industrial uses.

The LIPIDS of Crops team will work on behalf of our target audiences, which include both other plant scientists seeking to improve plant productivity, farmers, and others building the agricultural economy. The team will (1) develop a more efficient analytical pipeline to obtain information on lipid metabolism and traits relevant for crop improvement, (2) facilitate a better understanding and improvement of plant growth during development and under biotic and abiotic stress conditions to improve crop survival and yield, (3) increase our knowledge of seed metabolism and biology for improvement of seed oils as the group develops crops with improved productivity and quality for human and livestock nutrition and the bio-based economy.

 

Statement of Issues and Justification

The Need 

The North Central (NC) committee, Lipids In Plants: Improving and Developing Sustainability of Crops (“LIPIDS of Crops”), will continue to collaborate to elucidate lipid-related metabolism and traits relevant for crop improvement and to develop crops with enhanced yield, resilience to biotic and abiotic stress, and improved nutritional and industrial quality. This multistate project directly supports the USDA Science and Research Strategy 2023–2026, aligning with the priority areas of Accelerating Innovative Technologies and Practices, and Bolstering Nutrition Security and Health. 

Lipids are central to plant structure, metabolism, and signaling and are crucial determinants of how plants grow, develop, and respond to environmental challenges. Plant-derived oils and lipid-based compounds are vital not only for human and animal nutrition but also for renewable industrial applications that advance the U.S. bio-based economy. Despite their importance, many aspects of lipid metabolism, regulation, and function remain poorly understood, limiting opportunities to enhance crop productivity and quality, advances that are essential for ensuring an adaptive, nutritious, and economically strong agricultural system that benefits all Americans. 

The Importance of the Work 

Each plant is a biochemical factory that converts sunlight and carbon dioxide into thousands of metabolites, many of which are lipids. These include membrane-forming phospholipids, glycolipids and sterols, protective cuticular waxes and cutin monomers, seed oils, signaling molecules, plant (phyto-) hormones, and pigments essential for photosynthesis and stress tolerance. Despite their biological and economic importance, lipids are the least explored central metabolites. 

Greater insight into lipid metabolism will unlock opportunities to develop stress-resilient, nutrient-dense, and value-added crops, supporting USDA’s goals of improving nutrition security, reducing environmental impact, and strengthening rural economies. Many genes predicted to encode lipid-metabolizing enzymes have unknown functions, and the regulatory networks through which lipid signals control plant development and stress responses are still being defined. Addressing these knowledge gaps is essential for both resilient agriculture and innovative crop biotechnology, ensuring that U.S. agriculture remains globally competitive while contributing to a sustainable bioeconomy. 

The Technical Feasibility of the Research 

The LIPIDS of Crops team integrate complementary expertise in lipidomics, biochemistry, plant physiology, genomics, and biotechnology. Members will employ and advance state-of-the-art analytical methods, including high-resolution mass spectrometry, imaging-based lipid mapping, and metabolic flux analysis, to characterize lipid pathways and their roles in plant performance. By refining these technologies and standardizing lipid analytical pipelines across institutions, the project will help meet USDA’s objective to accelerate innovation and translate research into action. 

Research approaches will include genetic, biochemical, and physiological analyses of lipid-related traits and their impact on yield and stress tolerance. Improved analytical capacity and collaborative data sharing will enable discovery-driven advances in lipid metabolism and the development of crops optimized through marker-assisted breeding and biotechnological transformation. 

The Advantages of Doing the Work as a Multistate Effort 

Conducting this project as a multistate effort maximizes efficiency, collaboration, and innovation, key tenets of the USDA Science and Research Strategy. The North Central Region has a high concentration of leading plant lipid researchers and shared access to advanced analytical facilities. By coordinating expertise, data, and instrumentation, the group will catalyze progress that no single institution could achieve independently. The multistate framework also facilitates training and mentorship, broadening participation in cutting-edge agricultural research across different institutions. 

Likely Impacts of the Work 

The outcomes of this renewal will include: 

• A strengthened, collaborative research network focused on plant lipid metabolism and its role in sustainable agriculture. 

• Accelerated innovation in lipidomics and other analytical technologies for crop improvement. 

• Deeper mechanistic understanding of lipid-mediated regulation of plant growth, development, and stress responses. 

• Development of crops with improved yield and quality to enhance food, feed, and bio-based product markets. 

• Tangible contributions to USDA’s goals of adaptive agriculture, nutrition security, and enhancing the bio-economy. 

Related, Current and Previous Work

This multistate project integrates advanced analytical tools with fundamental studies of lipid biochemistry and metabolism to uncover the mechanisms that govern lipid synthesis, modification, storage, and signaling in plants. These efforts also address how lipids contribute to cellular homeostasis and plant resilience to environmental stresses. Insights from this work provide the foundation for engineering crops with improved oil content and composition, enhanced stress tolerance, and expanded functionality for agricultural and industrial applications. A review of the USDA-NIFA Data Gateway indicates that, while many projects address individual aspects of lipid biology or crop improvement, none matches the breadth and integration of this consortium, which connects basic discovery to translational application. The following literature review highlights major accomplishments achieved during the current funding period and emphasizes the collaborative work among project members that continue to drive progress toward shared goals. 

Objective 1. Develop systems-based analytical methods to characterize lipid metabolism. 

Mass spectrometry has revolutionized our ability to measure metabolites (including lipids) and proteins, providing key insights into the function of genes, proteins, and metabolites in plant physiology. Nearly 25 years after electrospray ionization mass spectrometry enabled direct analysis of intact lipids and metabolites, advances in instrumentation have greatly expanded sensitivity and coverage. However, standardization and development of new methods are still needed, particularly for plant metabolites, which are not comprehensively included in commercial analytical platforms. In addition, lipidomics is being integrated with other systems biology approaches, including transcriptomics, proteomics, and flux analysis, to better connect molecular profiles with specific biological mechanisms associated with lipid signaling, metabolic regulation, and stress resilience. 

NC1203 researchers have played and continue to play leading roles in the development of metabolomics, proteomics, and isotope-labeling approaches for plant biology. They (Allen (Danforth Plant Science Center: DPSC), Cahoon (Univ of Nebraska-Lincoln; UNL), Durrett (Kansas State; KSU), Koo (Univ of Missouri; MU), Kosma (Univ of Nevada-Reno; UNR), Lee (Iowa State; ISU), Markham (UNL), Wang (DPSC), Welti (KSU)) have developed mass spectrometry-based analytical methods for a wide range of lipids and metabolites, including phospholipids, glycolipids, sphingolipids, neutral glycerolipids, acyl-CoAs, acyl-ACPs, stress-induced lipids, amino acids, sugar/central metabolites, phytohormones, plant cuticle components, and additional more specialized compounds [e.g., Welti et al 2002; Koo et al 2014; Li et al 2014; Markham and Jaworski 2007; Markham et al 2006; Song et al 2020; Vu et al 2014; Jenkins et al 2021; Nam et al 2020; Koley et al 2022; Koley et al 2025]. Thelen’s group (MU) has developed methods to quantify proteins and their modifications, including a multiplexed AQUA-MRM assay for absolute quantification of specific proteins [Wilson and Thelen 2018; Fan et al 2024]. Allen and Bates (Washington State) have developed approaches to calculate metabolic flux based on measurements of both stable and radioisotope-labeled compounds over time [Allen et al 2015; Kotapati and Bates 2021; Kambhampati et al 2024]. Lee has focused on spatial analysis of lipids and other small molecules and has combined isotope labeling with spatial analysis to provide insights into membrane lipid restructuring and biosynthesis [Tat and Lee 2024; Na and Lee 2024]. Almost all the researchers in NC1203 are utilizing transcriptomics in conjunction with other omics approaches [e.g., Wahrenburg et al 2021, Muthan et al 2024, Chen et al 2024a, Kenchanmane Raju et al 2024, Kataya et al 2025]. 

Some highlights of LIPIDS of Crops work in the current project (2021-2026) include the Lee group’s optimization, in collaboration with other NC1203 researchers, of MALDI-MS imaging conditions for camelina and pennycress seed lipids and development of mass spectrometry imaging with isotope labeling of duckweed, Arabidopsis, and maize [Rensner et al 2022; Alkotami et al 2024; Lee et al 2024]. The Lee group also developed a new mass spectrometry imaging (MSI) technique to determine carbon-carbon double bond positions of PC lipids, called OzMALDI, by introducing ozone gas into the MALDI source [Rensner et al 2025]. Using this method, they successfully visualized PC double-bond isomers in camelina and soybean seeds engineered by Cahoon’s lab. The Lee group also demonstrated the use of unsupervised machine learning for mass spectrometry imaging data analysis after in vivo isotope labeling [Johnson et al 2025]. The Bates, Durrett and Welti groups developed and optimized three approaches for analysis of molecular species of diacylglycerol, a key metabolic intermediate in lipid biosynthesis, catabolism, and signaling [Parchuri et al 2023]. The Bates lab developed GC-FID-based derivatization methods for rapid characterization of whole seed fatty acid composition and quantity for a variety of current and emerging oilseed crops that allows quicker screening of seed lipid content [Garneau et al 2025]. The Thelen lab’s multiplexed AQUA-MRM assay was applied to absolute quantitation of Arabidopsis acetyl-CoA carboxylase catalytic and effector proteins, demonstrating the utility of this approach for study of the spatiotemporal regulation of this multienzyme complex that catalyzes the committed step of de novo fatty acid synthesis [Wang et al 2022]. 

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

Lipids play diverse and dynamic roles in plant biology, functioning not only as structural and storage molecules but also as signals, signal regulators, and essential elements of plant stress responses. This objective focuses on fundamental mechanisms through which lipid composition, organization, and metabolism contribute to plant resilience and adaptation. By uncovering these processes, we are identifying strategies and targets for improving stress tolerance and productivity in agricultural crops. Notably, much basic research in this objective involves newer areas of lipid biology including signaling and stress tolerance, while long-studied areas such as oil accumulation are progressing towards translational advances highlighted in Objective 3. 

Lipids as barriers that protect plants from biotic and abiotic stresses: 

The Yandeau-Nelson (ISU) and Kosma groups are characterizing the transcriptional networks underlying cuticle and suberin biosynthesis, which is important in protecting plants from biotic and abiotic stresses [Kosma et al., 2025]. Using integrated transcriptomic and metabolomic approaches, Yandeau-Nelson identified genetic networks underlying cuticle biosynthesis in maize seedlings [Chen et al 2024a] and maize silks and functionally characterized transcriptional regulators [Castorina et al 2023], and enzymes [Alexander et al 2020, Alexander et al 2024] involved in cuticle synthesis pathways. Relationships in cuticular wax biosynthesis have been inferred from the cuticular wax products that accumulated on the silk surface, establishing the complexity of product–precursor relationships within the silk cuticular wax-producing network [Chen et al 2024b]. Using similar approaches, Kosma identified five transcription factors that regulate suberin deposition in potatoes, four of which play roles in regulating tuber wound suberin biosynthesis [Wahrenburg et al 2021]. Other work investigated the functional role of transcriptional regulators of tomato leaf and fruit cutin biosynthesis in plants growing under saline or drought conditions [Bonarota et al 2024]. 

Lipids as signals: 

The Koo and Wang labs study the role of lipid signals in plant development and stress responses. Signaling roles of phosphatidic acid and diacylglycerol have been discovered [reviewed recently in Gong et al 2024; Yao et al 2024; Yang et al 2025; Yao et al 2025]. Additionally, the initial steps of jasmonate (JA) synthesis were investigated, revealing that a phospholipase A1 enzyme provides the JA precursor (by hydrolyzing linolenic acid from plastidial membranes) through a post-transcriptional regulatory mechanism conserved in Arabidopsis and N. benthamiana [Kimberlin et al 2022; Holtsclaw et al., 2024]. 

The Schrick lab is studying homeodomain leucine-zipper IV (HD-Zip IV) transcription factors important for differentiation of the epidermis that possess START lipid-binding domains [Schrick et al 2023]. The START domain is critical for protein stability as well as for transcription factor dimerization [Mukherjee et al 2022]. Recent work demonstrated that lysophospholipids are binding partners of the START domain, suggesting a mechanistic link between elongation growth control and phospholipid metabolism [Wojciechowska et al 2024]. Lysophospholipids are underrepresented in plant membranes and may serve as signaling molecules to modulate transcription factor function.  

Lipids as mediators of plant–microbe interactions: 

The Huang (Louisiana State) and Santos (UNR) groups are uncovering how diverse classes of lipids shape plant–microbe interactions, both by mediating microbial behavior and by contributing to plant defense. These studies span pathogen-derived lipids that influence infection outcomes and plant-derived defensive lipids whose biosynthesis and modes of action are only beginning to be understood. Huang investigates the role of lipid droplets during interactions between crops and fungal pathogens of soybeans, tomatoes, and strawberries, including Cercospora, Botrytis, and Agroathelia species. They demonstrated that fungal lipid droplets can serve as energy sources for reproduction or to sequester the fungal toxins, such as Cercosporin, produced by Cercospora fungus [de Novaes et al., 2024]. Santos studies falcarins, naturally occurring C17 polyacetylenic lipids [Busta et al 2018; Santos et al 2022]. Although falcarins were described more than 50 years ago and prior work suggests they contribute to resistance against several agriculturally important fungal pathogens, their modes of action and the regulatory mechanisms of their biosynthesis are still largely unknown. Santos, with Kosma and Cahoon, is investigating falcarin biosynthesis in carrot, which naturally synthesizes these compounds, with a focus on improving carrot plant resistance to devastating fungal pathogens [Santos et al 2022]. Both the Huang and Santos groups are also studying complex lipid metabolic pathways to develop novel antifungal compounds. 

Lipids as modulators of temperature tolerance: 

An interest of several groups includes lipid fluctuations in response to temperatures. The Roston laboratory (UNL) is establishing mechanistic and evolutionary frameworks for how plant membrane lipid remodeling supports low temperature resilience [Shomo et al 2024a]. The group demonstrated that triacylglycerol (TAG) metabolism buffers membrane stress across temperature extremes by showing all Arabidopsis DGAT and PDAT acyltransferases are active during both cold and heat, overturning the previous model of isoform specialization [Shomo et al., 2024b]. The Welti lab correlated responses to freezing with lipid compositional changes to propose roles of specific lipid in Arabidopsis freezing tolerance [Vu et al 2022]. Through lipidomics with the Welti lab and transcriptomics in panicoid grasses, the Roston lab uncovered rhythmic lipid and gene expression patterns during chilling stress, revealing that diurnal timing strongly influences cold tolerance [Kenchanmane Raju et al 2024]. Related evolutionary studies across bryophytes and angiosperms showed that oligogalactolipid accumulation, a response to severely low temperatures, arose independently in multiple lineages [Barnes et al 2023]. The Narayanan (Clemson U) and Welti labs collaborated to define lipid metabolic changes underlying heat tolerance in soybean and peanut and to develop molecular markers for screening germplasm. They found that heat stress drives removal of 18:3 and 18:2 fatty acids from membranes and their sequestration into TAGs and sterol esters, lowering membrane unsaturation. Reduced FAD3 expression was identified as a key factor decreasing 18:3 levels. Based on heat-adaptive lipid remodeling, these labs were able to identify potential QTLs for heat tolerance, which they will seek to verify in future studies [Spivey et al 2023]. 

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

Analytical tools developed in Objective 1 and the fundamental insights into lipid function uncovered in Objective 2 have expanded our understanding of how plants synthesize, regulate, and adapt their lipid metabolism under diverse conditions. These advances now provide a foundation for identifying metabolic control points that influence carbon flux into storage lipids. Building on this knowledge, Objective 3 integrates biochemical, genetic, and engineering approaches to develop crops with improved yield or enhanced functional properties.  

Traits that increase the quantity of plant products; 

Malonyl-CoA production by acetyl-CoA carboxylase (ACCase) is the committed step in de novo fatty acid synthesis, which if increased, would lead to higher oil accumulation. In plants, this pathway occurs in the plastids, where ACCase typically functions as a heteromeric multi-subunit complex. Although the catalytic subunits are better characterized, recent work has identified novel effector subunits in plants, including the BADC, RFS, and CTI gene families [Conrado et al 2024]. Additionally, the PII effector protein is present in autotrophs, as well as bacteria. The Thelen, Bates, Durrett, Van Doren (MU), and Yokom (MU) labs are actively defining the roles of these effector proteins through molecular genetic and structural studies. A recently published review of these novel effector proteins suggests post-translational modifications and alternative gene splicing may be yet another layer of regulation for ACCase [Balbuena et al 2025]. Gene knockouts for most of these effector proteins produce complex phenotypes, but generally increased TAG accumulation in the principal organ where the effector protein is expressed, e.g., leaves for CTI [Ye et al 2020a] and seed for BADC. This has led to the overall conclusion that these small effector proteins are negative regulators for ACCase, although this may be an oversimplification given the pleiotropy observed in PII / BADC stacked mutants [Garneau et al 2025]. BADC appears to respond to pH changes that link its impact to the diurnal cycle [Ye et al 2020b]. Such results confirm that a better understanding of these plant-unique effector proteins will require the collaborative research efforts of the NC1203 team, including bioanalytical (AQUA-MRM), biophysical (CryoEM, NMR, micro-scale thermophoresis), and molecular genetic approaches. 

In addition to understanding oil production in seeds, NC1203 researchers have collaborated to engineer increased TAG production in vegetative tissue as feedstocks for renewable diesel and sustainable aviation fuel (SAF) production [Park et al 2021; Park et al 2025]. In one collaboration, the Koo, Welti, and Allen labs together demonstrated that the inducible expression of a plastid-localized phospholipase A1 (PLA1) leads to the accumulation of TAGs in leaves [Kimberlin et al 2025]. Lipidomics and FAME analyses revealed that the accumulated TAGs were primarily composed of 18:3 fatty acids derived from MGDG and DGDG. Similar results obtained in N. benthamiana and soybean showcase the potential of this technology for crop engineering. Work in the Cahoon and Clemente labs led to the successful development and field testing of sorghum prototypes that produce elevated vegetative oil concentrations. In this study, constraints in carbon partitioning into fatty acid production and induction in fatty acid catabolism were identified as processes that limit maximal oil accumulation [Park et al 2025]. These bottlenecks will be examined in the proposed research in new design-build-test-learn iterations that will test hypotheses to enhance fatty acid production, while limiting their breakdown, toward sorghum biodesigns with commercially relevant vegetative oil levels. 

Through CRISPR-Cas9 gene editing of suberin genes, Kosma and Santos have developed a new potato cultivar with reduced sprouting and enhanced storage life that stands to reduce potato tuber post-harvest storage losses due to premature sprouting and associated tuber shrinkage [Vulavala et al 2024].  

Traits that increase the value of plant products 

Plant oils are important feedstocks for fuels, industrial chemicals, and bioproducts, yet most oilseed crops synthesize only a limited set of fatty acids. In contrast, plants collectively produce a wide range of lipid structures with valuable chemical properties. Recent progress in enzyme discovery and seed lipid metabolism now enables engineering crops to accumulate these unusual fatty acids, but such efforts often reveal unanticipated metabolic bottlenecks. Identifying the adaptations that constrain engineered lipid accumulation is essential for rational metabolic design and resilient crop improvement. The Cahoon, Allen, Bates, Durrett, Thelen, and Welti labs are jointly addressing these challenges by defining bottlenecks in medium-chain fatty acid engineering for sustainable aviation fuels in camelina and pennycress [Kim et al., 2015; Bansal et al., 2018]. 

Hydroxylated fatty acids (HFA) are valued by the chemical industry because the hydroxyl group enables chemical transformations that produce high-value polymers, lubricants, surfactants, and specialty chemicals. However, natural sources are not viable crops in the US due to the required growing climate, low yields, or toxicity. Attempts to bioengineer HFA or other unusual fatty acids into high yielding oilseed crops have had limited success. Bates and Parchuri (KSU) collaborated to discover how the native species Physaria fendleri accumulates HFA. They discovered that triacylglycerol (TAG) remodeling changes the oil fatty acid composition after initial synthesis and demonstrated in model species that engineering TAG remodeling can be a new tool to help control seed oil composition [Bhandari and Bates 2021; Parchuri et al 2024]. 

Acetyl-TAGs are an unusual class of triacylglycerols in which an acetyl group occupies the sn-3 position, resulting in oils with reduced viscosity and improved cold-temperature properties. These features make acetyl-TAGs attractive for use in drop-in biofuels and other bio-based industrial products. The Durrett lab, together with the Lee and Allen groups, generated camelina and pennycress lines that accumulate nearly pure acetyl-TAG in seeds (95 mol% in camelina and 98 mol% in pennycress) [Alkotami et al 2024]. MALDI-MS imaging showed that the small amount of remaining endogenous TAG is confined to the embryonic axis, highlighting potential strategies to engineer seeds that synthesize only acetyl-TAG. 

Other examples are the long-chain omega-3 oil and astaxanthin traits developed in soybeans by the Cahoon lab, in collaboration with the UNL Plant Transformation Core Research Facility (PTCRF) [Kim et al 2025]. These traits are designed to enhance the value of soybean oil as an aquaculture feedstock to expand market opportunities for US soybean farmers. To continue progress toward commercialization, project teams will carry out regulated field assessments and additional studies needed for advancing these traits. 

 

Successes enabled by LIPIDS of Crops (2021 to present) 

Publications: 105. Joint publications: 40 (highlighted in “Literature Cited” section). 

Patents: 4 

 

Funded collaborative grants: 

National Science Foundation: 2 

United Soybean Board: 5 

US Department of Agriculture: 3 

US Department of Energy: 1 

Objectives

1. Develop systems-based analytical methods to characterize lipid metabolism.
   
2. Identify lipid-related mechanisms to increase agricultural resilience
   
3. Develop crops with improved yield and/or functionality
   

Methods

Objective 1: Develop systems-based analytical methods to characterize lipid metabolism 

(Allen, Bates, Durrett, Kosma, Lee, Thelen, Welti; KS AES, MO AES, NV AES, WA AES, Donald Danforth Plant Science Center, Iowa State University) 

This objective will develop and refine analytical tools that underpin and enhance the research in the other two objectives, with particular emphasis on integrating lipid analysis with complementary systems biology approaches. By expanding capabilities in lipidomics, metabolite flux analysis, spatial metabolite imaging, and proteomics, we will generate high-resolution, multi-layer datasets and platforms that can be deployed across NC1203 to accelerate discovery and trait development in diverse plant systems. 

Lipid analysis:  

We will extend lipidomics capability by publishing a semi-quantitative and semi-targeted catalog of the occurrence and amounts of lipids from selected plant species across the plant kingdom from seedless vascular plants to angiosperms. These data are derived from comprehensive fatty acid-based negative precursor and positive neutral loss scanning by the Welti and Durrett groups, using a triple quadrupole mass spectrometer. We also plan to enhance our analysis of plant triacylglycerols by defining each of the three fatty acids in nearly all measured triacylglycerol molecular species, based on a method developed for animal lipids [Cabruja et al 2021].  

Other metabolite analysis: 

The Allen and Bates labs will collaborate to develop software for the quantitation of lipid fluxes through different branches of the lipid metabolic network utilizing radio isotopic labeling data. This software will aid in understanding how lipid metabolism changes due to genetic changes from breeding or bioengineering. Analysis of biopolymer suberin macro- and intermolecular structure has long been hindered by its recalcitrance to dissolution in traditionally employed organic solvents. Kosma has developed a novel chemical method for dissolving suberin into oligomeric structures amenable to analysis via GC-MS and other liquid-based chromatographic and/or mass spectroscopic measurements. To date, Kosma has been able to dissolve 80% of total suberin using this new method. Kosma will work with Welti to provide a quantitative analysis of the inter-molecular linkages of suberin using GC-MS- and LC/ESI-MS/MS-based methods. 

Spatial metabolic analysis:  

Recently, Kansas State received a new quadrupole time-of-flight mass spectrometer, funded by an NSF Major Research Instrumentation grant (PI: Durrett; Co-PIs include Schrick and Welti). This instrument is being installed in the Kansas Lipidomics Research Center, and its capabilities include desorption electrospray imaging (DESI). DESI can image fresh or even live tissues, and we will bring this technology to NC1203 and the larger plant biology community. This will be a complement to the image analysis provided by the Lee lab using MALDI imaging techniques. The Lee group will continue to develop MALDI-based MSI techniques and apply them to plant biology systems. They will employ on-tissue chemical derivatizations (OTCD) to enhance ion signals and improve metabolite annotations, as annotation options of metabolites can be differentiated by their ability or lack of ability to react with the derivatizing reagent. 

Proteomics: 

The Thelen lab is capable of label-free, global quantitation of 8-10k proteins and ~15k phosphopeptides, and is advancing targeted, absolute quantitation using the AQUA-MRM approach. This assay has been successfully used to quantity subunits of ACCase in Arabidopsis, ultimately leading to increased oil content in seeds [Wilson and Thelen 2018; Wang et al 2022] and to study the effect of acyl-lipid metabolism mutants on fatty acid synthesis [Garneau et al 2025; Fan et al 2024], making it a powerful tool for dissecting fatty acid synthesis in Arabidopsis by NC1203 collaborators. The Thelen lab will develop an expanded AQUA-MRM assay that quantifies each of the enzymatic steps from acetate to 18:1 production, a total of 34 proteins, in the oil seed crop pennycress. This pennycress assay will be fully validated and made available to the NC1203 community.  

 

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

(Huang, Koo, Louis, Narayanan, Peck, Roston, Santos, Schrick, Welti, Yandeau-Nelson; IA AES, KS AES, LA AES, MO AES, NE AES, NV AES) 

Objective 2 will use the analytical and systems-level tools developed in Objective 1 to continue developing a mechanistic-level understanding of the relationship of lipids as protective barriers, regulators of stress and defense pathways, and mediators of plant interactions with their environment. This objective seeks to reveal fundamental mechanisms that enable crops to maintain productivity under biotic and abiotic stress, toward future engineering goals. 

Assessing lipids as barriers protecting plants from biotic and abiotic stresses:  

The Yandeau-Nelson group will use synthetic biology approaches to 1) express and analyze maize cuticle biosynthesis pathways in yeast, to unravel the role of genetic redundancy in the fatty acid elongation pathway [Alexander et al 2024] and the hydrocarbon biosynthesis pathway, and 2) explore the transcriptional regulation of cuticle biosynthesis by expressing predicted cuticle-related transcription factors [Chen et al 2024a] in plant roots, which are mostly devoid of a cuticle. The group has recently identified mutants of cuticle-related genes that alter cuticular wax composition on maize silks. The Yandeau-Nelson and Louis (UNL) groups will test whether aphid feeding on maize silks is impacted in the cuticle-impaired mutants. This work will deepen our understanding of genetic architecture underlying cuticle synthesis and deposition and clarify structure-function relationships that contribute to resilience to drought and insect herbivory, with the goal of sustaining crop productivity under stress.  

Defining the roles of lipids as signals: 

The Koo lab will determine the biochemical regulation of PLA1 lipases for wound-elicited JA biosynthesis. They will collaborate with the Thelen lab to identify PLA1 protein interactors using co-immunoprecipitation proteomics followed by a reverse genomics approach. Koo will also characterize a novel inhibitor of the JA-dependent wound response. The Koo and Peck (MU) labs will collaborate to carry out a structure-function analysis of GLR3.6 ion channels, which are involved in systemic JA biosynthesis [Toyota et al 2018]. Welti and Koo will analyze data on variation in levels of JA and related compounds across Arabidopsis accessions to determine other genes/proteins affecting plant response to wounding. 

The Schrick lab will determine lipid- and adaptor protein binding mechanisms that control gene expression by using molecular dynamics simulations to characterize the lipid binding domain of HD-Zip IV transcription factors. Genetic and biochemical experiments will identify the downstream targets induced by lysophospholipid binding to these regulatory proteins. Loss-of-function mutants in associated adaptor proteins display increases in glucosinolate accumulation in sepals, the outer organ that protects flower buds from pathogen attack. The Schrick lab will collaborate with the Durrett, Welti, Koo, and Lee labs to engineer the identified glucosinolate-related traits from the Arabidopsis model into the oilseed crops, pennycress and camelina.   

Characterizing lipids as mediators of plant–microbe interactions:  

The Huang group will identify the role of lipids in Cercospora infection of soybean by infecting plants that are susceptible or resistant to the fungi and collaborating with the Welti group to determine the lipid compositional differences and their roles in disease-resistant response.  The Louis lab will identify the role of lipids in monocrop crop defense against insect pests (e.g., aphids, caterpillars) using multi-omic approaches. The Santos lab will identify yeast mutants susceptible or resistant to falcarin treatment and investigate the mutants further to shed light on the mode of action of the falcarins. To understand the role and regulation of the falcarin biosynthetic pathway in plant defense against necrotrophic fungi, stably transformed, falcarin-deficient carrot plants, provided by collaborator Cahoon, will be characterized for enhanced susceptibility or resistance to the fungus S. sclerotiorum. By comparing the transgenic and wild-type carrots using transcriptomics, targeted and untargeted metabolomics, molecular biology, and microbiology approaches, the regulation of carrot falcarin biosynthesis by specific necrotrophic pathogens, phytohormones, and other signaling molecules will be unveiled. Santos will collaborate with Kosma for targeted metabolomics. 

Investigating lipids as modulators of temperature tolerance: 

The Roston lab will collaborate with Welti lab to assess the effect of cold stress on lipid composition of crop plants across a series of timepoints to understand the effect of time on lipid changes post stress. To impose cold stress, they will employ a series of custom cold chambers and recovery chambers and will characterize the extent of death by leaf senescence/biomass and with electrolyte leakage. Photosynthetic fluorescence parameters will serve as a readout for photosystem impacts. The Narayanan and Welti labs will evaluate soy breeding populations such as a recombinant inbred line population (derived from a genetic cross between a heat-tolerant and a susceptible genotype) for developing molecular markers for heat tolerance. They will (1) screen the populations for physiological responses to heat stress, (2) elucidate lipid metabolic changes in tolerant and susceptible genotypes under heat stress, and identify changes associated with tolerance, and (3) determine genomic regions and molecular markers associated with heat-induced lipid metabolic changes and other heat tolerance-related traits. 

 

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

(Allen, Bates, Butler, Cahoon, Dhankher, Durrett, Gates, Kosma, Koo, Louis, Mukherjee, Obata, Parchuri, Sanjaya, Thelen, Van Doren, Yokom; KS AES, MA AES, MO AES, NE AES, NV AES, WA AES, Donald Danforth Plant Sciences Center, Texas A&M University – San Antonio, West Virgina State University) 

Global demand for plant-derived products is expected to rise sharply in coming decades, requiring crops that produce higher yields and more valuable lipid profiles on limited land [Gautum et al 2025; Park et al 2021]. Using tools developed in Objective 1 and building on basic research into the mechanisms of lipid accumulation, Objective 3 will develop strategies to boost oil accumulation and tailor lipid compositions, while advancing field evaluation and functionality testing to support translation of engineered traits. Together, these efforts will enable the development of high-value lipid products that strengthen U.S. agriculture and the bioeconomy. 

Increasing the quantity of products from crop seeds  

The Allen, Bates, Durrett, Dhankher (Univ of Massachusetts), Mukherjee (Texas A&M-San Antonio), and Parchuri labs will investigate mechanisms of triacylglycerol turnover (and remodeling) that limit total seed oil and select fatty acid accumulation. The team is elucidating the cellular and biochemical mechanisms underlying TAG turnover (predominantly by lipases, e.g. SDP1, TAGL1) and developing CRISPR-based genome editing and/or RNAi approaches to produce high oil yielding soybean, camelina, canola, and pennycress.  The ultimate goal is to overcome the mechanisms that limit seed oil or specific fatty acid accumulation and integrate this with engineering approaches that enhance oil synthesis or create designer fatty acid profiles to greatly increase total plant oil yields. 

Collaborations between project members will characterize key enzymes and regulatory factors involved in lipid biosynthesis and modify their expression to increase oil accumulation in seeds. The Sanjaya, Cahoon, and Durrett labs will investigate strategies to modify seed oil composition in crop plants by leveraging key genes and transcription factors that regulate lipid biosynthesis in both higher plants and algae [Muthan et al 2024]. The Thelen and Bates labs study the regulation of ACCase by effector proteins (BADC, CTI, PII), to understand total fatty acid synthesis. The Gates (MU), Yokom and Van Doren groups are pursuing biophysical and structural characterization of enzyme complexes via NMR, cryo-electron microscopy, and biochemical assays. The Durrett and Thelen groups will generate and characterize mutations in BADC and CTI genes in pennycress to develop seeds with increased oil content. It is anticipated that compensatory responses, ranging from acetyl-CoA supply through de novo fatty acid synthesis and desaturation, will occur in response to altered homeostatic control of ACCase in pennycress that will be studied by proteomics and AQUA-MRM assays (Objective 1) to understand how modulation of fatty acid synthesis effects metabolism. 

The Butler lab (UNL) is studying the impact of a major seed protein quantitative trait locus (QTL), cqSeed protein-003 on seed quality traits (oil and protein) in soybean. The QTL was recently mapped to gene model Glyma.20G085100 (Gm20P), with its paralog present on chromosome 10, Glyma.10G134400 (Gm10P) and both were knocked out in gene edited lines of soybean (gm10p gm20p). The gm10p gm20p double mutants were significantly lower in protein than the gm20p single mutant and wildtype and exhibited an increase in oil and starch content [Quach et al 2025]. Future research will determine the mechanisms of Gm20P and Gm10P and new targets for seed quality improvement.  

Increasing the quality and functionality of products in crop seeds  

The Cahoon, Durrett, Bates, Welti, Thelen, and Allen labs are integrating multi-omics analyses across multiple oilseed species to understand why naturally occurring medium-chain fatty acid (MCFA) traits fail to translate efficiently into camelina and pennycress engineered for C10 fatty acid production. Candidate genes implicated in MCFA synthesis or flux will be evaluated by targeted up- or down-regulation in these species. In parallel, seeds from MCFA-rich species such as Cuphea and elm will be mined to identify additional engineering targets. In addition, the Durrett lab will enhance the accumulation of unusual fatty acids by altering endogenous lipid-metabolic pathways in camelina and pennycress, including genome editing of competing LPAT and DGAT activities to generate platform lines optimized for heterologous enzyme expression. The Cahoon lab will assemble and deliver specialized enzyme constructs for MCFA production into these improved chassis lines. The Obata lab (UNL) will support the improvement of engineered plants by performing GC-MS-based metabolomics to characterize primary metabolism in engineered plants, helping to identify metabolic bottlenecks and perturbations affecting lipid production and plant phenotypes. 

The Parchuri lab, in collaboration with the Welti and Bates labs, will investigate lipid remodeling networks regulating the high-level accumulation of 20-carbon fatty acids in various Brassicaceae crop species, including production of the unusual 20-carbon monounsaturated fatty acid, Δ⁵-eicosenoic acid, in meadowfoam (Limnanthes alba) seeds. Using metabolic flux studies, lipidomics, and transcriptomics, we will elucidate the Δ⁵-eicosenoic acid-rich TAG biosynthetic pathway and characterize genes involved in lipid remodeling and TAG assembly. In collaboration with the Cahoon and Durrett labs, the Parchuri lab will also engineer Camelina sativa to accumulate Δ⁵-eicosenoic acid in seed oil through synthetic biology approaches. 

The Dhankher lab is engineering camelina for hyperaccumulation of nickel by overexpressing and mutating genes involved in uptake, transport and sequestration of metals in aboveground leafy biomass. Biomass will be used to recover nickel for use in EV batteries and industrial uses, and seed oil can be used for biofuel production. Further, Dhankher lab is also increasing the tolerance to toxic metals in oilseed crops including camelina, Brassica juncea and Crambe abtyssinica for enabling their cultivation on marginal lands.   

Enhancing production and functionality in non-seed tissues of crops  

The Butler and Louis labs are engineering lines of sorghum for enhanced flavonoid content in vegetative tissues by over-expressing the sorghum Bmr30 (Brown midrib 30) and maize bHLH (R-S) (basic helix-loop-helix R-S locus) genes via constitutive promoters. Bmr30 encodes a chalcone isomerase (CHI) enzyme and loss-of-function results in a reduction in the flavonoid, tricin, and reduced lignin deposition [Tetreault et al 2021]. bHLH (R-S) encodes a helix-loop-helix transcription factor and has been associated with increased anthocyanin production [Consonni et al 1993]. Seed tissues of resulting events will also be evaluated for yield, seed quality, and flavonoid content.  

The Koo, Welti, and Allen labs will collaborate to enhance oil production in vegetative tissues by combining the inducible PLA1 technology with other pathway engineering strategies. The Koo and Bates labs will investigate the mechanism of plastidial PLA1-dependent TAG accumulation in leaves using genetic and isotopic tracing. The Cahoon and Butler labs will build on their success in development of sorghum enriched in vegetative oils by exploring additional DBTL cycles that test strategies, including evaluation of high-activity DGATs to enhance fatty acid pull into TAG storage and downregulation of native lipases to limit TAG turnover [Park et al 2025].  

Santos and Kosma will investigate six suberin-regulating transcription factors that they have identified [Wahrenburg et al 2022; Kosma et al 2025] for their potential in mitigating potato tuber post-harvest storage losses by determining the transcriptional networks controlled by these transcription factors and analysis of suberin and post-harvest storage-related phenotypes of CRISPR-Cas9 edited potato accessions targeting these six transcription factors.

Toward translation of biodesigned crops  

The Cahoon, Butler, Durrett, Santos, and Kosma labs will take advantage of transgenic field-testing capacity at the Eastern Nebraska Research, Extension and Education Center (ENREEC) to evaluate a suite of engineered traits across multiple crops. The Cahoon and Butler labs are advancing soybean lines with seed oils enriched in the omega-3 fatty acid EPA and soybean and camelina lines that produce astaxanthin-rich oils, both optimized to minimize yield drag and maintain normal oil content. Field testing of these lines will occur at the ENREEC, with collaborations planned for oil evaluation in aquaculture feeding studies. Field trials with the new potato cultivar developed by Santos and Kosma will be conducted in Nebraska with Cahoon to determine yields and to evaluate post-harvest storage losses  In addition, the Cahoon and Durrett labs will assess high–acetyl-TAG camelina lines (~95% acetyl-TAG) and will expand field-testing capability to Kansas State University to support multi-location evaluation of other lipid traits developed by NC1203 researchers. 

 

Measurement of Progress and Results

Outputs

• Shared analytical and engineering platforms (Objectives 1, 3): Deployment of standardized lipid analysis and engineering pipelines, from AQUA-MRM and flux models to genome-edited “platform” lines, and design-build-test-learn (DBTL) workflows shared among participants.
• Lipidomics and multi-omics data resources (Objectives 1, 2): Generation and public dissemination of integrated lipidomic, proteomic, and fluxomic datasets from developing seeds and stress-treated vegetative tissues, supporting community-scale modeling of lipid metabolism and regulation.
• Validated lipid pathway targets and regulatory components (Objectives 2, 3): Identification and biochemical or structural validation of key lipid metabolic enzymes and regulatory proteins, lipid remodeling enzymes, and kinases regulating lipid metabolism.
• Developed germplasm and research lines (Objectives 2, 3): Development of crop and model species lines with enhanced seed and vegetative oil content, customized oil composition, or demonstrated tolerance to abiotic or biotic stresses.
• Field evaluation and pre-commercial material distribution (Objective 3): Multi-site field evaluation of engineered crops with accompanying datasets on agronomic performance, oil yield and composition. Provision of scaled seed/biomass and oil samples for further testing in industry application evaluation (e.g., aquaculture feed trials, fuel performance testing).

Outcomes or Projected Impacts

• Advancement of lipid metabolic engineering knowledge: Foundational insights into lipid biosynthesis, regulation, storage, and their roles mediating plant stress responses. These discoveries will enable more rationale design of traits that increase oil accumulation, improve lipid composition, and enhance resilience to abiotic and biotic stresses.
• Higher-value and higher-yielding crop products: Development of oil-rich, designer-lipid, and stress-tolerant crops (e.g. soybean, camelina, pennycress, sorghum, maize, carrot, potato) will expand the availability of plant products for food, feed, and industrial uses, supporting profitability and U.S. agricultural competitiveness.
• Enhanced nutritional and industrial oil quality: Production of seed oils containing health-promoting long-chain fatty acids (EPA, DHA) and specialized industrial lipids (acetyl-TAGs, MCFAs, HFAs) will broaden options for renewable fuels, aquaculture, nutraceuticals, and biomanufacturing sectors.
• Workforce development and shared research infrastructure: Availability of standardized datasets, characterized lipid pathway components, methods for translation of lab results, and shared analytical platforms will facilitate coordinated research and data integration, improving the community of researchers. Training of early career interdisciplinary scientists in lipid biochemistry, plant biotechnology, and plant product development will further support a strong U.S. future in plant lipids.

Milestones

(2027):Identify and prioritize lipid pathway targets and regulatory nodes for modification or mutant characterization. Initiate genome editing and transgenic construct development in soybean, camelina, pennycress, tobacco, carrot, potato, Arabidopsis, maize and sorghum. Establish shared analytical workflows (AQUA-MRM, lipidomics, fluxomics, and genomics) Develop multi-site coordination frameworks for confined field testing. Characterize phosphorylation events associated with low-temperature responses in crops to identify regulatory steps influencing lipid metabolism. Determine key molecular features of lipid binding from molecular dynamics simulations. Assess structural features of the biotin carboxylase component of ACCase. Explore the lipid biosynthesis and metabolism inhibitors that affect plant-microbe and plant-insect interactions. Build heterologous systems to test functions of lipid transporters. Optimize chemical techniques for characterizing lipid polymer (suberin) structure.

(2028):Create first-generation engineered lines (soybean, camelina, pennycress, carrot, potato, sorghum, maize, etc.) and platform backgrounds. Begin metabolic, biochemical, and structural characterization of ACCase regulatory components and TAG synthesis or remodeling enzymes. Conduct preliminary confined field trials of existing EPA, astaxanthin, acetyl-TAG, and high-vegetative-oil lines. Experimentally test molecular dynamics simulation-inspired lipid-binding mechanisms. Complete pilot DESI imaging experiments on at least one crop. Identify the mode of action of lipid biosynthesis and metabolism inhibitors in suppressing crop disease development. Characterize plant transporter function and substrate specificity in heterologous systems. Test plant-insect and plant-microbe interactions in plant mutants of lipid pathways. Optimization of MS/MS techniques for characterizing lipid polymer (suberin) structure.

(2029):Characterization of first-generation engineered lines and platform backgrounds. Integrate multiple systems biology analyses to refine targets. Expand field evaluations across ENREEC and Kansas State capacity; collect yield and oil quality data. Begin scaled material production for downstream oil functionality testing. Experimentally test molecular dynamics simulation-inspired lipid-binding mechanisms. Determine insect resistance of leaf-oil engineered plant lines.

(2030):Complete multi-site field testing and trait validation.  Conduct aquaculture feed studies and biofuel/lubricant performance analyses with partners.  Quantify and analyze lipid compositional changes triggered by temperature changes in crops to assess their functional relevance. Identify and functionally test kinases involved in temperature activation of lipid metabolic enzymes to determine additional regulatory targets for crop improvement. Create second-generation engineered lines based on phenotypes and system analyses of first-generation lines Identify the effect of inhibiting lipid biosynthesis on the fungal toxin delivery to the plant host.

(2031):Characterization of second-generation engineered lines Develop and assemble the project renewal Finalize trait performance datasets and publish integrative findings.  Transition high-potential traits to industry. Develop crop disease management tools based on lipid biosynthesis and metabolism targets in plant-microbe and plant-insect interactions

Projected Participation

View Appendix E: Participation

Outreach Plan

The “LIPIDS of Crops” group will publish its results in scientific journals and make data available via appropriate databases. “LIPIDS of Crops” members also will present their results at scientific meetings. Method advances also will be published. For example, the Kansas Lipidomics Research Center has a website that includes extraction and other protocols and information. 

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 are active in organizing scientific meetings that promote the dissemination of new knowledge and foster collaboration and professional networking, with a strong emphasis on supporting junior scientists.  For example, all participants frequently attend the Gordon Research Conference on Plant Lipids: Structure, Metabolism and Function held most recently in 2025 in Pomona, California. Members of the LIPIDS of Crops group have served and continue to serve as Chairs and Vice-Chairs, spearheading the organization of the meeting to benefit the community. This will continue in 2027 with Chairs Yandeau-Nelson and Allen, and Vice-Chairs Bates and Narayanan. In 2024, Cahoon and Welti along with Kent Chapman chaired the International Symposium on Plant Lipids held in Lincoln, NE which brought together approximately 250 plant lipid scientists from around the world and was followed by a joint conference on camelina co-organized by Cahoon. 

NC1203 participants maintain a strong commitment to extensive outreach activities that promote STEM learning, disseminate research findings to stakeholders, and enhance public engagement with plant and agricultural sciences. 

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. At each annual meeting, a new secretary is elected, the chair-elect becomes chair, and the previous Secretary becomes chair-elect. The chair is responsible for organizing the next year’s meeting. The secretary is responsible for submitting the annual report. Administrative guidance is provided by an assigned Administrative Advisor and a NIFA Representative 

Literature Cited

* Publications since 2021 from LIPIDS of Crops members.  

** Joint publications since 2021 

* Abdullah HM, Rodriguez J, et al. (2021) Increased cuticle waxes by overexpression of WSD1 improves osmotic stress tolerance in Arabidopsis thaliana and Camelina sativa. Int. J. Mol. Sci. 22:5173. https://doi.org/10.3390/ijms22105173.  

* Abdullah HM, Pang N, et al. (2024). Overexpression of the phosphatidylcholine:diacylglycerol cholinephosphotransferase (PDCT) gene increases carbon flux toward triacylglycerol synthesis in Camelina sativa seeds. Plant Physiol Biochem 208:108470. doi:10.1016/j.plaphy.2024.108470 

** Ahmad B, Lerma-Reyes R, et al. (2024). Nuclear localization of HD-Zip transcription factor GLABRA2 is driven by Importin α. J Exp Bot 75:6441–6461. doi:10.1093/jxb/erae326 

** Alexander LA, Winkelman D, et al. (2024). The impact of the GL2 and GL2-LIKE BAHD-proteins in affecting the product profile of the maize fatty acid elongase. Frontiers in Plant Science 15:1403779. doi: 10.3389/fpls.2024.1403779 

Alexander LE, Okazaki Y, et al. (2020). Maize Glossy2 and Glossy2-like genes have overlapping and distinct functions in cuticular lipid deposition. Plant Physiology 183:840-853. doi: 10.1104/pp.20.00241 

** Alkotami L, White DJ, et al. (2024). Targeted engineering of camelina and pennycress seeds for ultrahigh accumulation of acetyl-TAG. Proc Natl Acad Sci U S A. 121:e2412542121. doi: 10.1073/pnas.2412542121 

Allen DK, Bates PD, Tjellström H. (2015). Tracking the metabolic pulse of plant lipid production with isotopic labeling and flux analyses: Past, present and future. Progress in Lipid Research 58:97-120. doi: 10.1016/j.plipres.2015.02.002 

* Arias CL, Quach T, et al. (2022). Expression of AtWRI1 and AtDGAT1 during soybean embryo development influences oil and carbohydrate metabolism. Plant Biotechnol J 20:1327–1345. 

* Azeez A, Bates PD. (2024). Self-incompatibility–based functional genomics for rapid phenotypic characterization of seed metabolism genes. Plant Biotechnol J 22:2688–2690. doi:10.1111/pbi.14383 

** Azeez A, Parchuri P, Bates PD. (2022). Suppression of Physaria fendleri SDP1 increased seed oil and hydroxy fatty acid content while maintaining oil biosynthesis through triacylglycerol remodeling. Front Plant Sci 13:931310. 

** Aznar-Moreno JA, Mukherjee T, et al. (2022). Suppression of SDP1 improves soybean seed composition by increasing oil and reducing undigestible oligosaccharides. Front Plant Sci 13:863254. 

* Balbuena TS, Lemes Jorge G, et al. (2025). Novel effector proteins of the plant heteromeric acetyl-CoA carboxylase. J Exp Bot. In press. 

Bansal S, Kim HJ, et al. (2018). Towards the synthetic design of camelina oil enriched in tailored acetyl-triacylglycerols with medium-chain fatty acids. J Exp Bot 69:4395-4402. doi: 10.1093/jxb/ery225 

* Barnes AC, Myers JL, et al. (2023). Oligogalactolipid production during cold challenge is conserved in early diverging lineages. Journal of Experimental Botany 74:5405–5417. doi: 10.1093/jxb/erad241 

* Bhandari S, Bates PD. (2021). Triacylglycerol remodeling in Physaria fendleri indicates oil accumulation is dynamic and not a metabolic endpoint. Plant Physiol 187:799-815. doi: 10.1093/plphys/kiab294 

* Blume RY, Kalendar R, et al. (2023). Overcoming genetic paucity of Camelina sativa: Possibilities for interspecific hybridization conditioned by the genus evolution pathway. Front Plant Sci 14:1259431. 

* Blume RY, Hotsuliak VY, et al. (2024). Genome-wide identification and diversity of FAD2, FAD3, and FAE1 genes in camelina species. BMC Biotechnol 24:107. doi:10.1186/s12896-024-00936-4 

* Bonarota MS, Kosma DK, Barrios-Masias FH. (2022). Salt tolerance mechanisms in the Lycopersicon clade and their trade-offs. AoB Plants 14:plab072. https://doi.org/10.1093/aobpla/plab072 

* Bonarota MS, Kosma D, Barrios-Masias FH. (2024). Physiological characterization of the tomato cutin mutant cd1 under salinity and nitrogen stress. Planta 260:64. 

* Burns MJ, Renk JS, et al. (2021). Predicting moisture content during maize nixtamalization using machine learning with NIR spectroscopy. Theor Appl Genet 134:3743–3757. 

Busta L, Yim WC, et al. (2018). Identification of genes encoding enzymes catalyzing the early steps of carrot polyacetylene biosynthesis. Plant Physiology 178:1507–1521. doi: 10.1104/pp.18.01195 

* Busta L, Chapman KD, Cahoon EB. (2022). Better together: Protein partnerships for lineage-specific oil accumulation. Curr Opin Plant Biol 66:102191. 

** Busta L, Dweikat I, et al. (2022). Chemical and genetic variation in feral Cannabis sativa populations across the Nebraska climate gradient. Phytochemistry 200:113206. 

Cabruja M, Priotti J, et al. (2021). In-depth triacylglycerol profiling using MS3 Q-Trap mass spectrometry. Anal Chim Acta 1184:339023. doi: 10.1016/j.aca.2021.339023 

* Cahoon EB, Kim P, et al. (2024). Sphingolipid homeostasis: How do cells know when enough is enough? Implications for plant pathogen responses. Plant Physiol 197:kiae460. doi:10.1093/plphys/kiae460 

* Cardona JB, Grover S, et al. (2023a). Sugars and cuticular waxes impact sugarcane aphid (Melanaphis sacchari) colonization on different developmental stages of sorghum. Plant Sci 330:111646. 

* Cardona JB, Grover S, et al. (2023b). Sorghum cuticular waxes influence host plant selection by aphids. Planta 257:22. 

* Castorina G, Bigelow M, et al. (2023). Roles of the MYB94/FUSED LEAVES1 (ZmFDL1) and GLOSSY2 (ZmGL2) genes in cuticle biosynthesis and potential impacts on Fusarium verticillioides growth on maize silks. Frontiers in Plant Science 14:1228394. doi: 10.3389/fpls.2023.1228394 

* Chen M, Wang S, et al. (2023). Plastid phosphatidylglycerol homeostasis influences polar lipid synthesis in Arabidopsis. Metabolites 13:318. 

** Chen K, Alexander L, et al. (2024). Dynamic relationships among pathways that produce the hydrocarbons and VLCFAs of maize silk cuticular waxes. Plant Physiology 195:2234-2255. doi: 10.1093/plphys/kiae150 

** Chen K, Bhunia R, et al. (2024). Cuticle development and underlying transcriptome-metabolome associations during early seedling establishment in maize. Journal of Experimental Botany 20:6500-6522. doi: 10.1093/jxb/erae311 

* Cheng D, Li L, et al. (2022). Heterologous expression and characterization of plant wax ester–producing enzymes. Metabolites 12:577. 

* Chhikara S, Singh Y, et al. (2024). Overexpression of bacterial γ-glutamylcysteine synthetase increases toxic metal/loid tolerance and accumulation in Crambe abyssinica. Plant Cell Rep 43:270. doi:10.1007/s00299-024-03351-3 

** Chu KL, Koley S, et al. (2022). Metabolic flux analysis of the non-transitory starch tradeoff for lipid production in mature tobacco leaves. Metab Eng 69:231–248. 

* Colak N, Kurt-Celebi A, et al. (2025). Salicylic acid priming before cadmium exposure increases wheat growth but does not uniformly reverse cadmium effects on membrane glycerolipids. Plant Biol 27:79–91. doi:10.1111/plb.13736 

* Conrado AC, Lemes Jorge G, et al. (2024). Evolution of the regulatory subunits for the heteromeric acetyl-CoA carboxylase. Philos Trans R Soc Lond B Biol Sci 379:20230353. doi: 10.1098/rstb.2023.0353 

Consonni G, Geuna F, et al. (1993). Molecular homology among members of the R gene family in maize. Plant J 3:335-46. doi: 10.1111/j.1365-313x.1993.tb00185.x  

* Deng M, Chen H, et al. (2023). Genetic improvement of tocotrienol content enhances the oxidative stability of canola oil. Front Plant Sci 14:1247781. 

de Novaes MIC, Robertson C, et al. (2024). Distribution and Sequestration of Cercosporin by Cercospora cf. flagellaris. Phytopathology 114:1822-1831. 

* Dong J, Croslow SW, et al. (2025). Enhancing lipid production in plant cells through automated high-throughput genome engineering and phenotyping. Plant Cell 37:koaf026. doi:10.1093/plcell/koaf026 

* Du ZY, Hoffmann-Benning S, et al. (2021). Editorial: Lipid metabolism in development and environmental stress tolerance for engineering agronomic traits. Front Plant Sci 12:739786. 

** Durrett TP, Welti R. (2021). The tail of chlorophyll: Fates for phytol. J Biol Chem 296:100802. 

** Esterhuizen L, Ampimah N, et al. (2025). AraRoot – A comprehensive genome-scale metabolic model for the Arabidopsis root system. In Silico Plants. doi:10.1093/insilicoplants/diaf003 

* Fan J, Sah SK, et al. (2024). Arabidopsis trigalactosyldiacylglycerol1 mutants reveal a critical role for phosphtidylcholine remodeling in lipid homeostasis. Plant J 120:788-798. doi: 10.1111/tpj.17020 

* Fliege CE, Ward RA, et al. (2022). Fine mapping and cloning of the major seed protein QTL on soybean chromosome 20. Plant J 110:114–128. 

* Gan L, Park K, et al. (2022). Divergent evolution of extreme production of variant plant monounsaturated fatty acids. Proc Natl Acad Sci USA 119:e2201160119. 

** Garneau MG, Lemes Jorge G, et al. (2025). PII interactions with the acetyl-CoA carboxylase subunits BADC and BCCP co-regulate lipid and nitrogen metabolism in Arabidopsis. Plant Physiol. In press. 

** Garneau MG, Parchuri P, et al. (2025). Rapid quantification of whole seed fatty acid amount, composition, and shape phenotypes from diverse oilseed species with large differences in seed size. Plant Methods 21:67. doi: 10.1186/s13007-025-01388-3 

* Gautam B, Kim H, et al. (2025). Meeting liquid biofuel and bioproduct goals: Biotechnological design of the intermediate oilseeds pennycress and camelina, and beyond. J Experimental Botany. doi: 10.1093/jxb/eraf415 

* Gong Q, Yao S, et al. (2024) Fine-tuning phosphatidic acid production for optimal plant stress responses. Trends Biochem Sci. 8:663-666. doi: 10.1016/j.tibs.2024.05.008. 

* Grover S, Puri H, et al. (2022). Dichotomous role of jasmonic acid in sorghum defense against aphids. Mol Plant Microbe Interact. 35:755-767 doi:10.1094/MPMI-01-22-0005-RHoffmann-Benning S. (2021). Collection and analysis of phloem lipids. In: Bartels D, Dörmann P (eds). Plant Lipids: Methods and Protocols. Methods in Molecular Biology 2295:351–361. 

* Hoffmann-Benning S, Simon-Plas F. (2024). Editorial: Lipid signaling in plant physiology. Plant Sci 334:112088. doi:10.1016/j.plantsci.2024.112088 

* Holtsclaw RE, Mahmud S, Koo AJ. (2024). Identification and characterization of GLYCEROLIPASE A1 for wound-triggered JA biosynthesis in Nicotiana benthamiana leaves. Plant Mol Biol 114:4. doi: 10.1007/s11103-023-01408-7 

* Huang F, Chen P, et al. (2022). Genome assembly of the Brassicaceae diploid Orychophragmus violaceus reveals complex whole-genome duplication and evolution of dihydroxy fatty acid metabolism. Plant Commun 100432. doi:10.1016/j.xplc.2022.100432. 

* Hurst JP, Yobi A, et al. (2023). Large and stable genome edits at the sorghum alpha kafirin locus result in changes in chromatin accessibility and globally increased expression of genes encoding lysine enrichment. Front Plant Sci 14:1116886. 

* Jenkins LM, Nam JW, et al. (2021). Quantification of acyl-acyl carrier proteins for fatty acid synthesis using LC-MS/MS. Plant Lipids: Methods and Protocols, Methods in Molecular Biology 2295. doi: 10.1007/978-1-0716-1362-7_13 

** Johnson BS, Allen DK, Bates PD. (2025). Triacylglycerol stability limits futile cycles and inhibition of carbon capture in oil-accumulating leaves. Plant Physiol 197:kiae121. doi:10.1093/plphys/kiae121 

* Johnson RLB, Tat VT, Lee YJ. (2025). Unsupervised machine learning for mass spectrometry imaging data analysis with in vivo isotope labeling. Analyst 150:4404-4413. doi: 10.1039/d5an00649j 

* Kakati JP, Zoong Lwe Z, Narayanan S. (2022). Heat stress during the early flowering stage did not affect seed fatty acid contents in conventional oleic peanut varieties. Peanut Sci 49:1. 

* Kambhampati S, Hoffman A, et al. (2021). Dynamic flux balance analysis of leaf metabolism in response to drought stress. Metab Eng 63:125–138. 

* Kambhampati S, Hubbard AH, et al. (2024). Stable Isotope Labeled Pathway Elucidation (SIMPEL): Using stable isotopes to elucidate dynamics of context-specific metabolism. Communications Biology 7:172-183. 

** Kataya A, Nascimento JRS, et al. (2025). Comparative omics reveals unanticipated metabolic rearrangements in a high-oil mutant of plastid acetyl-CoA carboxylase. J Proteome Res. doi: 10.1021/acs.jproteome.4c00947 

* Kenchanmane Raju SK, Zhang Y, et al. (2024). Rhythmic lipid and gene expression responses to chilling in panicoid grasses. Journal of Experimental Botany 75:5790–5804. doi: 10.1093/jxb/erae247

* Kim H, Liu L, et al. (2025). Oilseed-based metabolic engineering of astaxanthin and related ketocarotenoids using a plant-derived pathway: Lab-to-field-to-application. Plant Biotechnol J 23:3451-3464. doi: 10.1111/pbi.70148 

Kim HJ, Silva JE, et al. (2015). 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. Journal of Experimental Botany 66:4251-4265 

** Kim P, Mahboob S, et al. (2024). Characterization of soybean events with enhanced expression of microtubule-associated protein 65-1 (MAP65-1). Mol Plant Microbe Interact 37:62–71. doi:10.1094/MPMI-09-23-0134-R 

* Kim SC, Yao S, et al. (2022). Phospholipase Dδ and phosphatidic acid mediate heat-induced nuclear localization of glyceraldehyde-3-phosphate dehydrogenase in Arabidopsis. Plant J 112:786–799. doi:10.1111/tpj.15981. 

* Kimberlin A, Holtsclaw RE, et al. (2022). On the initiation of jasmonate biosynthesis in wounded leaves. Plant Physiol 189:1925-1942. 

** Kimberlin AN, Mahmud S, et al. (2025). Increasing oil production in leaves by engineering plastidial phospholipase A1. Plant J 121:e70088. doi: 10.1111/tpj.70088 

* Koley S, Chu KL, et al. (2022). An efficient LC-MS method for isomer separation and detection of sugars, phosphorylated sugars, and organic acids. Journal of Experimental Botany 73:2937-295. 

* Koley S, Chu KL, et al. (2022). Metabolic synergy in Camelina reproductive tissues for seed development. Sci Adv 8:abo7683. doi:10.1126/sciadv.abo7683. 

** Koley S, Jyoti P, et al. (2025). Persistent fatty acid catabolism during plant oil synthesis. Cell Reports 44:115492. 

** Konda AR, Gelli M, et al. (2023). Vitamin E biofortification: Maximizing oilseed tocotrienol and total vitamin E tocochromanol production by use of metabolic bypass combinations. Metab Eng 79:66–77.

* Koo AJ, Thireault C, et al. (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. doi: 10.1074/jbc.M114.603084

* Koo AJ, Arimura GI. (2022). Molecular biology of chemical defenses. Plant Mol Biol 109:351–353. 

* Koppisetti RK, Fulcher YG, Van Doren SR. (2021). The fusion peptide of SARS-CoV-2 spike rearranges into a wedge inserted in bilayered micelles. J Am Chem Soc 143:13205–13211.

* Kosma DK, Graça J, Molina I. (2025). Update on the structure and regulated biosynthesis of the apoplastic polymers cutin and suberin. Plant Physiology 197:kiae653. 

* Kotapati HK, Bates PD. (2021). Plant Lipids: Methods and Protocols 2295:59-80. Doi: 10.1007/978-1-0716-1362-7_5

* Kulke M, Kurtz E, et al. (2024). PLAT1/PLAFP binds the Arabidopsis plasma membrane and inserts a lipid. Plant Sci 338:111900. doi:10.1016/j.plantsci.2023.111900 

* Lee YJ, Hapuarachchige P, et al. (2024). Visualizing 13C-labeled metabolites in maize root tips with mass spectrometry imaging. J Am Soc Mass Spectrom 35:7. doi: 10.1021/jasms.4c00042 

* Levy JG, Mendoza-Herrera A, et al. (2023). Evaluation of the effect of ‘Candidatus Liberibacter solanacearum’ haplotypes in tobacco infection. Agronomy 13:569. 

Li M, Baughman E, et al. (2014). Quantitative profiling and pattern analysis of triacylglycerol species in Arabidopsis seeds by electrospray ionization mass spectrometry. Plant J 77:160-172. doi: 10.1111/tpj.12365 

* Li J, Su Y, et al. (2023). Phosphate deficiency modifies lipid composition and seed oil production in Camelina. Plant Sci 111636. doi:10.1016/j.plantsci.2023.111636. 

* Li J, Yao S, et al. (2025). Non-specific phospholipase C4 improves phosphorus remobilization from old to young leaves in Camelina. Plant Cell Environ. doi:10.1111/pce.15122 

* Li S, Zhang X, et al. (2025). Deciphering the core shunt mechanism in Arabidopsis cuticular wax biosynthesis and its role in environmental adaptation. Nat Plants 11:165–175. doi:10.1038/s41477-024-01892-9

* Li Z, Kim M, et al. (2025). Knocking out the carboxyltransferase interactor 1 (CTI1) in Chlamydomonas boosted oil content by fivefold without affecting cell growth. Plant Biotechnol J 23:1230-1242. doi: 10.1111/pbi.14581 

* Li-Beisson Y, Roston RL. (2024). Plant and algal lipids: In all their states and on all scales. Plant Cell Physiol 65:823–825 doi:10.1093/pcp/pcae061 

* Liu P, Xie T, et al. (2023). Mechanism of sphingolipid homeostasis revealed by structural analysis of Arabidopsis SPT–ORM1 complex. Sci Adv 9:eadg0728.

** Lusk HJ, Neumann N, et al. (2022). Lipidomic analysis of Arabidopsis T-DNA insertion lines identifies C-terminal alterations in FATTY ACID DESATURASE 6. Plant Cell Physiol 63:1193–1204. 

* Maitra S, Viswanathan MB, et al. (2022). Bioprocessing, recovery, and mass balance of vegetative lipids from metabolically engineered “oilcane.” ACS Sustain Chem Eng 10:16833–16844.  

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. doi: 10.1002/rcm.2962 

Markham JE, Li J, Cahoon EB, Jaworski JG. (2006). Separation and identification of major plant sphingolipid classes from leaves. J Biol Chem 281:22684-22694. doi: 10.1074/jbc.M604050200

* McGuire ST, Shockey J, Bates PD. (2025). The first intron and promoter of Arabidopsis DIACYLGLYCEROL ACYLTRANSFERASE1 exert synergistic effects on pollen and embryo lipid accumulation. New Phytol 245:263–281.

* Meher PK, Begam S, et al. (2022). ASRmiRNA: Abiotic stress-responsive miRNA prediction in plants using machine learning algorithms. Int J Mol Sci 23:1612.

** Morley SA, Ma F, et al. (2023). Expression of malic enzyme reveals subcellular carbon partitioning of storage reserve production in soybean. New Phytol 239:1834–1851. 

* Mugume Y, Ding G, et al. (2022). Complex changes in membrane lipids associated with modification of autophagy in Arabidopsis. Metabolites 12:190. 

** Mukherjee T, Subedi B, et al. (2022). The START domain mediates Arabidopsis GLABRA2 dimerization and turnover independently of homeodomain DNA binding. Plant Physiol 190:2315-2334. doi: 10.1093/plphys/kiac383 

* Mulaudzi T, Sias G, et al. (2023). Seed priming with MeJA prevents salt-induced growth inhibition and oxidative damage in Sorghum bicolor. Int J Mol Sci 24:10368. 

** Murphy KM, Johnson BS, et al. (2025). Excessive leaf oil modulates the plant abiotic stress response via reduced stomatal aperture in Nicotiana tabacum. Plant J 121:e70067.  doi: 10.1111/tpj.70067 

** Muthan B, Wang J, et al. (2024). Mechanisms of Spirodela polyrhiza tolerance to FGD wastewater-induced heavy-metal stress: Lipidomics, transcriptomics, and functional validation. J Hazard Mater 469:133951. doi: 10.1016/j.jhazmat.2024.133951 

* Na S, Lee YJ. (2024). Mass spectrometry imaging of Arabidopsis thaliana with in vivo D2O labeling. Frontiers in Plant Science 15. Doi.org/10.3389/fpls.2024.1379299 

Nam JW, Jenkins LM, et al. (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. 

* Neumann N, Fei T, et al. (2023). Defining the physical properties of blends of acetyl-triacylglycerols derived from transgenic oilseeds. J Am Oil Chem Soc 101:197–204. doi:10.1002/aocs.12746 

* Neumann N, Harman M, et al. (2024). Arabidopsis diacylglycerol acyltransferase1 mutants require fatty acid desaturation for normal seed development. Plant J. 119: 916-926.  doi: 10.1111/tpj.16805 

* Nguyen D, Groth N, et al. (2024). Project ChemicalBlooms: Collaborating with citizen scientists to survey chemical diversity of epicuticular wax blooms. Plant Direct 8:e588. doi:10.1002/pld3.588 

* Nwafor CC, Li D, et al.  (2022). Genetic and biochemical investigation of seed fatty acid accumulation in Arabidopsis. Front Plant Sci 13:942054. 

* Ojeda-Rivera JO, Barnes AC, et al. (2025). Designing a nitrogen-efficient cold-tolerant maize for modern agricultural systems. Plant Cell 37:koaf139. doi:10.1093/plcell/koaf139. 

* Osinuga A, Solís AG, et al. (2024). Deciphering sphingolipid biosynthesis dynamics in Arabidopsis cell cultures. iScience 27:110675. doi:10.1016/j.isci.2024.110675 

** Parchuri P, Pappanoor A, et al. (2022). Lipidome analysis and characterization of Buglossoides arvensis acyltransferases. Plant Sci 324:111445. 

** Parchuri P, Bhandari S, et al. (2024). Identification of triacylglycerol remodeling mechanism to synthesize unusual fatty acid containing oils. Nature Communications 15:3547. doi: 10.1038/s41467-024-47995-x 

** Parchuri P, Garneau MG, et al. (2023). Comparison of TLC, HPLC, and direct-infusion ESI-MS methods for the identification and quantification of diacylglycerol molecular species. Methods Enzymol 683:191-224. doi: 10.1016/bs.mie.2022.09.011 

** Park K, Sanjaya S, et al. (2021). Toward sustainable production of value‐added bioenergy and industrial oils in oilseed and biomass feedstocks. Gcb Bioenergy 13: 1610-1623. 

** Park K, Quach T, et al. (2025). Development of vegetative oil sorghum: From lab-to-field. Plant Biotechnol J 23:660-673. doi: 10.1111/pbi.14527 

* Qin P, Chen P, et al. (2024). Vitamin E biofortification: Enhancement of seed tocopherol concentrations by altered chlorophyll metabolism. Front Plant Sci 15:1344095. doi:10.3389/fpls.2024.1344095

* Quach TN, Nguyen H, et al. (2023). Introduction of genome editing reagents and genotyping of edited alleles in soybean. In Plant Genome Engineering: Methods and Protocols (pp. 273-285). New York, NY: Springer US.  doi:10.1007/978-1-0716-3131-7_17

* Quach TN, Sato SJ, et al. (2023). A facile Agrobacterium-mediated transformation method for Chlamydomonas reinhardtii. In Vitro Cell Dev Biol Plant 59:671–683. 

* Quach T, Nguyen H, et al. (2025). Editing of the gene model underlying the major protein quantitative trait loci (QTL), cqSeed protein-003, and its paralog in soybean. J Experimental Botany. Accepted. 

** Rensner JJ, Kim H, et al. (2025). OzMALDI: A gas-phase, in-source ozonolysis reaction for efficient double-bond assignment in mass spectrometry imaging with matrix-assisted laser desorption/ionization. Anal Chem 97:7447–7455

* Rensner JJ, Lee YJ. (2022). Efficient hydrogen-deuterium exchange in matrix-assisted laser desorption/ionization mass spectrometry imaging for confident metabolite identification. Anal Chem 94:11129-11133. doi: 10.1021/acs.analchem.2c00978

* Rhee SY, Anstett DN, et al. (2025). Resilient plants, sustainable future. Trends Plant Sci 30:382–388. doi:10.1016/j.tplants.2024.11.001 

* Richter M, Segal LM, et al. (2024). Membrane fluidity control by Magnaporthe oryzae acyl-CoA binding protein sets thermal limits for rice cell colonization. PLoS Pathog 20:e1012738. doi:10.1371/journal.ppat.1012738 

** Santos P, Busta L, et al. (2022). Structural diversity, biosynthesis, and function of plant falcarin-type polyacetylenic lipids. Journal of Experimental Botany 73:2889–2904. doi: 10.1093/jxb/erac006 

* Saucedo-García M, González-Solís A, et al. (2023). Sphingolipid long-chain base signaling in compatible and non-compatible Arabidopsis–pathogen interactions. Int J Mol Sci 24:4384. 

* Schrick K, Ahmad B, Nguyen HV. (2023). HD-Zip IV transcription factors: Drivers of epidermal cell fate integrate metabolic signals. Curr Opin Plant Biol 75:e102407. doi: 10.1016/j.pbi.2023.102417

* Scott S, Cahoon EB, Busta L. (2022). Variation on a theme: Structures and biosynthesis of specialized fatty acid natural products in plants. Plant J 111:954–965. 

** Shockey J, Parchuri P, et al. (2023). Assessing the biotechnological potential of cotton type-1 and type-2 diacylglycerol acyltransferases. Plant Physiol Biochem 196:940–951.

* Shomo ZD, Li F, Smith CN, et al. (2024). From sensing to acclimation: The role of membrane lipid remodeling in plant responses to low temperatures. Plant Physiol 196:1737–1757. doi: 10.1093/plphys/kiae382

* Shomo ZD, Mahboub S, et al. (2024). All members of the Arabidopsis DGAT and PDAT acyltransferase families operate during high and low temperatures. Plant Physiology 195:685–697. doi: 10.1093/plphys/kiae074

* Singh G, Le H, et al. (2024). Overexpression of CsGGCT2;1 reduces arsenic toxicity and accumulation in Camelina sativa. Plant Cell Rep 43:14. doi:10.1007/s00299-023-03091-w

* Singh G, Aftab SO, Dhankher OP. (2025). Arabidopsis oxoprolinase1 maintains glutamate homeostasis and promotes arsenite and mercury tolerance. Plant J 122:e70154. doi:10.1111/tpj.70154

Song Y, Vu HS, et al. (2020). A lipidomic approach to identify cold-induced changes in Arabidopsis membrane lipid composition. Methods Mol Biol 2156:187-202. doi: 10.1007/978-1-0716-0660-5_14

** Spivey WW, Rustgi R, et al. (2023). Lipid Modulation contributes to heat stress adaptation in peanut. Frontiers in Plant Science 14:1299371.

* Stenback KE, Flyckt KS, et al. (2022). Modifying yeast very-long-chain fatty acid biosynthesis by expression of plant 3-ketoacyl-CoA synthases. Sci Rep 12:13235.

* Subedi B, Schrick K. (2022). EYFP fusions to HD-Zip IV transcription factors enhance stability and cause phenotypic changes in Arabidopsis. Plant Signal Behav 17:2119013.

* Surber SM, Thien Thao NP, et al. (2024). Exploring cotton SFR2’s role in cold stress response. Plant Signal Behav 19:2362518.

* Tat VT, Lee YJ. (2024). Spatio-temporal study of galactolipid biosynthesis in duckweed using mass spectrometry imaging and in vivo isotope labeling. Plant Cell Physiol:pcae032

Tetreault HM, Gries T, et al. (2021). The sorghum (Sorghum bicolor) brown midrib 30 gene encodes a chalcone isomerase required for cell wall lignification. Frontiers in Plant Science 12:732307. doi: 10.3389/FPLS.2021.732307

* Toyinbo JO, Saripalli G, et al. (2025). Impact of mutations in soybean oleate and linoleate desaturases on seed germination under heat stress. Crops 5:2.

* Vadde BVL, Russell NJ, et al. (2024). The transcription factor ATML1 maintains giant cell identity by inducing synthesis of its own long-chain fatty acid-containing ligands. bioRxiv. doi:10.1101/2024.03.14.584694. 

* Villalobos JA, Cahoon RE, v. (2024). Glucosylceramides impact cellulose deposition and cellulose synthase complex motility in Arabidopsis. Glycobiology 34:cwae035. doi:10.1093/glycob/cwae035

Vu HS, Shiva S, et al. (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. doi: 10.1111/tpj.12659 

** Vu HS, Shiva S, et al. (2022). Specific changes in Arabidopsis thaliana rosette lipids during freezing can be associated with freezing tolerance. Metabolites 12:385. doi: 10.3390/metabo12050385

* Wang H-L, Ding B-J, et al. (2022). Insect pest management with sex pheromone precursors from engineered oilseed plants. Nat Sustain. doi:10.1038/s41893-022-00949-x.

** Wahrenburg Z, Benesch E, et al. (2021). Transcriptional regulation of wound suberin deposition in potato cultivars with differential wound healing capacity. Plant J 107:77-99. doi: 10.1111/tpj.15275

** Wang M, Garneau MG, et al. (2022). Overexpression of pea α-carboxyltransferase in Arabidopsis and Camelina increases fatty acid synthesis leading to improved seed oil content. Plant J 110:1035-1046.

* Wang S, Blume RY, et al. (2024). Chromosome-level assembly of Camelina neglecta, a diploid model for camelina biotechnology. Biotechnol Biofuels Bioprod 17:17. doi:10.1186/s13068-024-02466-9

* Wilson RS, Thelen JJ. (2018). In vivo quantitative monitoring of subunit stoichiometry for metabolic complexes. J Proteome Res 17:1773-1783.

* Winkelman DC, Nikolau BJ. (2022). The effects of carbon source and growth temperature on fatty acid profiles of Thermobifida fusca. Front Mol Biosci 9:8962261.

** Wojciechowska I, Mukherjee T, et al. (2024). Arabidopsis PROTODERMAL FACTOR2 binds lysophosphatidylcholines and transcriptionally regulates phospholipid metabolism. New Phytol 244:1498-1518. doi: 10.1111/nph.19917

* Yang B, Li J, et al. (2023). Non-specific phospholipase C4 hydrolyzes phosphosphingolipids and phosphoglycerolipids and promotes rapeseed growth and yield. J Integr Plant Biol. doi:10.1111/jipb.13560.

* Yang B, Fan R, et al. (2025) Non-specific phospholipase Cs and their potential for crop improvement. J Exp Bot. doi: 10.1093/jxb/eraf334. 

* Yao S, Kim SC, et al. (2024) Phosphatidic acid signaling and function in nuclei. Prog Lipid Res. 93:101267. doi: 10.1016/j.plipres.2023.101267.

* Yao S, Yang B, et al. (2025) Phosphatidic acid signaling in modulating plant reproduction and architecture. Plant Commun. 6:101234. doi: 10.1016/j.xplc.2024.101234.

Ye Y, Fulcher YG, et al. (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. doi: 10.1074/jbc.RA120.012877

Ye Y, Nikovics K, et al. (2020). Docking of acetyl-CoA carboxylase to the plastid envelope membrane attenuates fatty acid production in plants. Nat Commun 11:6191. doi: 10.1038/s41467-020-20014-5 

PATENTS

* Berman D, Chapman KD, Romsdahl DB, Minto RE, Zhang C, Cahoon EB. (2021). Liquid and semisolid lubricant compositions, methods of making, and uses thereof. US Patent 11,136,525. 

* Koo AJ, Kimberlin A. (2022). Genetic means to increase neutral oil in vegetative tissues by conditional induction of membrane lipid hydrolysis. US Patent P13628US01. 

** Neumann N, Nazarenus TJ, Aznar Moreno JA, Durrett TP, Cahoon EB. (2023). Improved camelina plants and plant oil, and uses thereof. US Patent Application 17/770,466.

** Vulavala VKR, Kosma DK, Santos P. (2024). Potato variety named ‘UNR-01’. US Patent App. 18/739,142. 

 

Attachments

Land Grant Participating States/Institutions

IA, KS, LA, MA, MO, NE, NV, SC, WA, WV

Non Land Grant Participating States/Institutions

Danforth Plant Science Center, Iowa State University - College of Vet Med, Texas A&M University