植物科学

Symbiosomes are plant membrane–bound, organelle-like structures that enclose single or multiple bacteroids (differentiated rhizobia) within legume cells, serving specifically as the site for N₂ fixation. Although isolated symbiosomes have been used to study nitrogen fixation, ion transport, and metabolite translocation, conventional density gradient–based isolation methods for their isolation are labor-intensive. Here, we describe a simplified, density gradient–free protocol for rapid enrichment of symbiosome-containing fractions from mature legume nodules using the Medicago truncatula–Sinorhizobium meliloti symbiosis as an example. The method combines razor blade homogenization, filtration through a 40 μm cell strainer, and sequential low-speed centrifugation to remove debris and enrich symbiosome-associated fractions. The resulting material is compatible with downstream mass spectrometry–based proteomics and mutant phenotypic analyses. Compared with conventional approaches, this protocol is rapid, simple, and suitable for higher-throughput biochemical and proteomic studies of symbiosomes.

Plants growing in soil are exposed to several environmental stresses, such as soil compaction, flooding, or drought, and need to quickly acclimate to these conditions to survive. Such acclimations include suberization and lignification of the outer and inner parts of the root. Visualization of suberin and lignin depositions in roots can be done with several well-established staining techniques followed by microscopy, but these methods are mostly qualitative. Radial water loss assays provide a quick, robust, and quantitative method to evaluate the formation of suberized and/or lignified outer apoplastic barriers and how tightly they regulate water loss. Here, we provide a detailed step-by-step protocol, from harvesting the roots from soil conditions and cleaning them, to preparing them for water loss measurements, while using basic lab equipment. Since the data obtained is water loss as mass, quantitative and statistical comparisons can be performed. Finally, while in this protocol roots were grown in soil, the method can be applied to roots growing in hydroponics, agar plates, or any other growth conditions that allow non-destructive harvesting of roots. In summary, the protocol provides a fast and reliable method to assess differences in outer apoplastic barrier development of root tissues, allowing for fast root tissue screenings while obtaining quantitative data without expensive or highly specialized equipment.

Obtaining homozygous mutant and transgenic lines is a critical yet time-consuming step in Arabidopsis thaliana research. Conventional breeding procedures require seeds to undergo complete maturation and natural desiccation before harvest, followed by cold stratification to overcome seed dormancy. This process substantially prolongs generation turnover and delays genetic screening. Here, we describe a rapid germination strategy based on the use of partially dehydrated seeds collected approximately 15 days after pollination (DAP). At this developmental stage, embryos have reached physiological maturity, while the seeds have not yet entered deep desiccation-induced dormancy. After surface sterilization and short-term cold treatment (2–3 days at 4 °C), these seeds readily germinate on 1/2 MS medium and develop into normal seedlings. By bypassing the prolonged maturation and dormancy phases associated with naturally dried seeds, this protocol shortens each generation cycle by approximately 1–2 weeks. The method is particularly useful for accelerating the propagation of transgenic materials and the identification of homozygous mutant lines in Arabidopsis.

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.

Accurate quantification of plant disease severity is essential for evaluating host–pathogen interactions and assessing the effectiveness of disease management strategies. Traditional visual scoring methods and manual estimation of infected tissue are widely used but are often subjective and prone to observer bias. Digital image analysis offers an objective alternative by enabling automated identification and quantification of symptomatic plant tissues based on color and spatial characteristics.

Here, we present a MATLAB-based image processing protocol for differentiating diseased and healthy plant tissue from digital leaf images. The workflow involves acquisition of standardized leaf images, conversion of RGB images into hue-saturation-value (HSV) color space, segmentation of diseased tissue using defined HSV thresholds, refinement of the segmented mask through morphological operations, and extraction of the whole leaf area. The protocol then calculates the diseased area and total leaf area in pixels and computes the percentage of infected tissue. The method uses MATLAB together with the Image Processing Toolbox and can be implemented using simple scripts. This protocol enables rapid and reproducible quantification of disease severity in plant leaves exhibiting visually distinct symptoms such as necrotic lesions or blight patches. By minimizing observer bias and providing quantitative measurements of infected area, the protocol offers a practical and reproducible approach for plant disease phenotyping and evaluation of disease management strategies across diverse plant-pathogen systems where diseased tissues can be clearly distinguished from healthy tissues under reasonably controlled imaging conditions.

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.

Accurately quantifying the areas of necrotic lesions on plant leaves is essential for evaluating plant–pathogen interactions and disease resistance. Although digital image analysis methods using ImageJ are widely employed, they often require case-specific optimization and may not be readily applicable across different experimental conditions. Furthermore, many studies have used ImageJ for lesion measurement without providing methodological details, which limits reproducibility. Here, we present a simple, step-by-step ImageJ workflow for measuring irregular necrotic lesions using a standard personal computer and mouse. The procedure relies on manual lesion selection using the freehand selection tool, followed by Gaussian smoothing, binarization, and automated particle analysis to extract lesion area measurements. By balancing manual isolation with computational thresholding, this protocol eliminates the need for extensive parameter tuning. This approach provides an accessible, reliable alternative to time-consuming color thresholding methods, thereby improving transparency and reproducibility in lesion quantification. The workflow's reproducibility has been confirmed through both intra-user and inter-user analyses.

Plant roots dynamically respond to environmental changes and serve as an ideal system for studying cell development and gene regulation. Recent advances in imaging-based spatial transcriptomics have enabled high-resolution mapping of gene expression while preserving spatial context. However, existing sample preparation techniques remain inadequate for handling rigid plant tissues such as crop roots. Here, we present a detailed and practical protocol for preparing rigid plant tissue samples for imaging-based spatial transcriptomics. The workflow ensures effective tissue handling while maintaining RNA integrity and spatial organization. Within approximately eight days, samples can be processed and mounted onto commercial slides, making them ready for subsequent probe hybridization and imaging. This protocol also includes an integrated sample attachment test performed to assess slide quality. It has been optimized to produce consistent and reliable results across experiments. Overall, our method provides a robust solution for spatial transcriptomic analysis in rigid plant tissues, facilitating broader application of these technologies in plant research.

Super-resolution microscopy has transformed our ability to visualize subcellular structures, but its application in plant biology remains challenging due to the optical complexity of plant tissues. Here, we present a detailed protocol for tau-STED microscopy (Leica Microsystems), which combines stimulated emission depletion (STED) with fluorescence lifetime imaging (FLIM) to achieve nanoscale resolution while minimizing phototoxicity. This method leverages time-correlated single-photon counting (TCSPC) to separate fluorescence signals based on their lifetimes, enhancing signal specificity and enabling the visualization of elusive subcellular compartments in Arabidopsis thaliana root tips. The protocol covers sample preparation, fluorophore selection, microscope configuration, image acquisition, and data analysis, providing a step-by-step guide to optimize tau-STED imaging for plant cell biology. By addressing the unique challenges of plant tissue imaging, such as autofluorescence, refractive index mismatches, and light scattering, this approach facilitates super-resolution imaging of intracellular structures, including the plant endoplasmic reticulum–Golgi intermediate compartment (ERGIC). This protocol is designed to be accessible to researchers with basic microscopy experience and offers a robust framework for exploring subcellular dynamics in plants with unprecedented detail.

Ammonium (NH₄⁺) is the primary inorganic nitrogen source for rice (Oryza sativa L.). Substantial progress has been made in characterizing the functions of ammonium transporters (AMTs) in roots; however, the regulatory dynamics governing subcellular ammonium compartmentation after its entry into cells, particularly its vacuolar sequestration and efflux back to the external environment, remain poorly understood. This knowledge gap stems mainly from two factors: the difficulty of applying conventional detection methods at the organellar scale and interference caused by nonspecific ion adsorption to the cell wall of intact roots. To address these challenges, we present a detailed and reproducible protocol for real-time measurement of net NH₄⁺ fluxes in rice roots, root protoplasts, and isolated vacuoles using non-invasive micro-test technology (NMT). The protocol covers the preparation of protoplasts and vacuoles from rice roots, the configuration and calibration of the NMT system, and the step-by-step measurement of net NH₄⁺ fluxes at three distinct biological levels (intact roots, protoplasts, and vacuoles). By employing a unified sample preparation and measurement strategy, this protocol enables quantification of net uptake fluxes across the plasma membrane, characterization of net efflux dynamics under specific conditions, and indirect estimation of vacuolar sequestration capacity using the isolated vacuole system. Overall, this protocol provides a flexible and robust framework for studying NH₄⁺ homeostasis in plants and is readily adaptable to different crop species, treatment conditions, and experimental objectives. Owing to its modular design and compatibility with standard NMT equipment, it can be readily adopted by laboratories seeking to investigate nitrogen transport mechanisms in plants.