植物科学

Identifying substrates of protein phosphatases has been technically challenging and has hampered progress in the field of plant sciences. Small molecule inhibitors of protein phosphatases have aided in uncovering classes of phosphatases that target substrates, but that too has severe limitations. Here, we describe a method that enriches phosphorylated substrates using TiO₂ and phospho-tyrosine antibodies in phosphatase knockout lines of Arabidopsis thaliana. When compared to wild-type plants, this approach permits identification of putative substrates and specific phosphorylation sites by mass spectrometry, allowing for further in vitro or functional validation. The key to the approach described here is the use of phosphatase knockout lines to maintain substrates in a phosphorylated state and using phospho-tyrosine antibodies to enrich for tyrosine phosphorylated peptides.

Efficient protein synthesis in eukaryotic cells typically requires a 5′ cap structure on messenger RNAs (mRNAs). However, under stress conditions or in viral infection, translation can also occur independently of the cap via internal ribosomal entry sites (IRES). IRES elements are therefore key regulators of protein expression in both viral and cellular contexts. Here, we describe a cell-free protocol to quantitatively assess cap-independent translation using wheat germ extract (WGE) and a firefly luciferase (FLuc) reporter. The protocol includes template preparation, RNA synthesis, and luminescence measurement following in vitro translation in WGE. This method enables rapid and robust comparison of translation activity under controlled conditions and can additionally be applied to evaluate mRNA modifications designed to enhance translation efficiency.

Small ubiquitin-like modification (SUMOylation) is a crucial post-translational modification that modulates protein stability, localization, and interaction dynamics. Despite the identification of thousands of putative small ubiquitin-like modifier (SUMO) substrates, functional validation remains challenging due to the low abundance and highly dynamic nature of SUMOylated proteins. Here, we present a protocol for detecting protein SUMOylation, integrating bioinformatic site prediction, and rapid substrate screening via in vivo tobacco transient expression and in vitro E. coli assay, followed by precise validation using transgenic Arabidopsis lines. However, detection of low-abundance SUMOylated proteins may require coupling with mass spectrometry, and the in vitro system does not fully recapitulate the complex regulatory network in vivo. This workflow provides a useful tool for studying SUMOylation in plants.

Peroxisomal β-oxidation is a key step in jasmonic acid biosynthesis. Quantitative biochemical characterization of enzymes involved in the β-oxidation pathway is essential for validating their catalytic functions and comparing differences among genetic variants. Existing enzyme activity assays largely rely on chromatographic techniques to quantify substrate consumption or product formation, but these approaches are not well-suited for high-throughput or continuous kinetic measurements. Here, we describe a spectrophotometric assay based on a plate reader determining OsAIM1 enzymatic activity by monitoring the decrease in NADH absorbance at 340 nm. The method employs a 96-well plate reaction system, enabling real-time kinetic measurements and providing a standardized workflow for calculating reaction rates. Reaction components, protein concentration ranges, and data processing parameters were systematically optimized to ensure linearity, reproducibility, and quantitative accuracy. This assay is simple to perform, requires small reaction volumes, and offers relatively high throughput, making it suitable for functional characterization and kinetic analysis of NADH-dependent enzymes.

The activity of chloroplast ATP synthase (CF_oCF₁) is precisely regulated through a thioredoxin (Trx)-mediated dithiol/disulfide reaction in response to varying light conditions. This regulatory mechanism is further controlled by ΔpH formation across the thylakoid membrane. To better understand this complicating regulatory function of CF_oCF₁, a method is required to evaluate the extent of CF_oCF₁ reduction by Trx under controlled ΔpH conditions and to directly evaluate the redox state of CF_oCF₁. In this study, we present a simple in vitro procedure to assess the CF_oCF₁ reduction system using spinach thylakoids. The method consists of three key steps: (A) simple preparation of intact thylakoids from spinach leaves; (B) reduction of CF_oCF₁ on the thylakoid membrane using recombinant Trx under light irradiation; and (C) in situ determination of the redox state of CF_oCF₁ by labeling thiol groups with a maleimide reagent followed by protein detection using western blotting. The redox state of CF_oCF₁ was determined by mobility shifts on non-reducing SDS-PAGE. This protocol provides a refined strategy for elucidating the regulatory mechanism controlling energy conversion by CF_oCF₁ under fluctuating photosynthetic conditions.

Anthocyanins are specialized flavonoid pigments that play critical roles in plant coloration, photoprotection, and responses to environmental stress. Arabidopsis thaliana serves as a valuable genetic model for dissecting anthocyanin biosynthesis and regulatory networks. Conventional methods for anthocyanin quantification, such as crude spectrophotometric assays, often compromise pigment integrity, yield inconsistent results, and provide limited information on compound composition. Here, we describe a simple, reproducible, and high-fidelity protocol for the induction, extraction, quantification, and chromatographic profiling of anthocyanins in Arabidopsis thaliana seedlings. The workflow employs well-defined anthocyanin-inductive conditions (AIC), methanol/formic acid extraction, lyophilization for dry-weight normalization, and dual quantification via spectrophotometry and High-performance liquid chromatography with diode-array detection (HPLC-DAD) analysis. This protocol enables accurate comparison between wild-type and mutant genotypes, facilitating both mutant screening and metabolic pathway analysis. The approach minimizes pigment degradation, enhances reproducibility across replicates, and offers a robust tool for research in plant metabolism, stress physiology, and flavonoid biochemistry.

The plant cell wall is a dynamic and complex extracellular matrix that not only provides structural integrity and determines cell shape but also mediates intercellular communication. Among its major components, pectins play essential roles in cell adhesion, wall porosity, hydration, and flexibility. Rhamnogalacturonan-I (RG-I), a structurally diverse pectic polysaccharide, remains one of the least understood components of the plant cell wall. Its backbone is substituted with arabinan, galactan, and arabinogalactan side chains that vary in length, branching, and composition across tissues, species, and developmental stages. In addition, RG-I can undergo modifications such as backbone acetylation, further contributing to its structural complexity and functional diversity. To advance understanding of RG-I, we present a detailed method for isolating RG-I from the model plant Arabidopsis thaliana. Leveraging Arabidopsis as a model system provides major advantages owing to its well-characterized genome and powerful molecular toolkit, enabling deeper investigation into the roles of RG-I in plant development and responses to environmental stress. Our method consists of two major steps: an initial chemical extraction using oxalate, followed by endo-polygalacturonase (EPG) digestion to fragment the pectic domains. An advantage of this approach is that it produces a dry material that can be stored at room temperature without special handling and does not introduce chemicals that may interfere with downstream analyses. The purified RG-I can be used for detailed compositional and structural analyses, as well as for functional studies of enzymes involved in pectin biosynthesis, modification, and degradation. Although this protocol was developed for isolating RG-I from Arabidopsis rosette leaves, it is also applicable to other Arabidopsis organs and other plant species.

The protochlorophyllide (Pchlide) level is a crucial indicator of plant fitness. Precise quantification of Pchlide content is necessary not only in studies of flu-related mutants that over-accumulate Pchlide in the dark but also for research on plants suffering from environmental stresses. Due to its low content and interference of chlorophylls, quantitative determination of Pchlide content is a challenge. Here, we describe an optimized protocol for Pchlide extraction from Arabidopsis thaliana seedlings and subsequent analysis using high-performance liquid chromatography (HPLC) coupled with fluorescence detection. Divinyl-Protochlorophyllide (DV-Pchlide, the major form of Pchlide in plants) quantification is achieved by interpolating fluorescence peak areas against an experimentally derived standard curve. This protocol provides a reliable workflow for Pchlide quantification, facilitating the deciphering of the underlying mechanism of plant environmental resilience.

In plants, the apoplast contains a diverse set of proteins that underpin mechanisms for maintaining cell homeostasis, cell wall remodeling, cell signaling, and pathogen defense. Apoplast protein composition is highly regulated, primarily through the control of secretory traffic in response to endogenous and environmental factors. Dynamic changes in apoplast proteome facilitate plant survival in a changing climate. Even so, the apoplast proteome profiles in plants remain poorly characterized due to technological limitations. Recent progress in quantitative proteomics has significantly advanced the resolution of proteomic profiling in mammalian systems and has the potential for application in plant systems. In this protocol, we provide a detailed and efficient protocol for tandem mass tag (TMT)-based quantitative analysis of Arabidopsis thaliana secretory proteome to resolve dynamic changes in leaf apoplast proteome profiles. The protocol employs apoplast flush collection followed by protein cleaning using filter-aided sample preparation (FASP), protein digestion, TMT-labeling of peptides, and mass spectrometry (MS) analysis. Subsequent data analysis for peptide detection and quantification uses Proteome Discoverer software (PD) 3.0. Additionally, we have incorporated in silico–generated spectral libraries using PD 3.0, which enables rapid and efficient analysis of proteomic data. Our optimized protocol offers a robust framework for quantitative secretory proteomic analysis in plants, with potential applications in functional proteomics and the study of trafficking systems that impact plant growth, survival, and health.

Protein isolation combined with two-dimensional electrophoresis (2-DE) is a powerful technique for analyzing complex protein mixtures, enabling the simultaneous separation of thousands of proteins. This method involves two distinct steps: isoelectric focusing (IEF), which separates proteins based on their isoelectric points (pI), and sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), which separates proteins by their relative molecular weights. However, the success of 2-DE is highly dependent on the quality of the starting material. Isolating proteins from plant mature roots is challenging due to interfering compounds and a thick, lignin-rich cell wall. Bacterial proteins and metabolites further complicate extraction in legumes, which form symbiotic relationships with bacteria. Endogenous proteases can degrade proteins, and microbial contaminants may co-purify with plant proteins. Therefore, comparing extraction methods is essential to minimize contaminants, maximize yield, and preserve protein integrity. In this study, we compare two protein isolation techniques for lupine roots and optimize a protein precipitation protocol to enhance the yield for downstream proteomic analyses. The effectiveness of each method was evaluated based on the quality and resolution of 2-DE gel images. The optimized protocol provides a reliable platform for comparative proteomics and functional studies of lupine root responses to stress, e.g., drought or salinity, and symbiotic interactions with bacteria.