植物科学

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.

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.

In the model alga Chlamydomonas reinhardtii, CRISPR (clustered regularly interspaced short palindromic repeat)-based gene editing using Cas (CRISPR-associated) enzymes enables both (a) insertion of large gene cassettes and (b) the creation of knockouts based on the introduction of indels, and specific mutations via mutation-directing oligonucleotides. Owing to the relatively low efficiency of this process, selection markers are frequently used to enrich the candidate pool prior to screening, which typically employs PCR. Unfortunately, few selection markers are available for Chlamydomonas. Furthermore, each marker requires different selection media, and deletion of the selectable marker can be difficult. When multiple successive gene editing steps are required, the use of these markers becomes onerous. The SCREAM (sequential CRISPR via recycling endogenous auxotrophic markers) technique employs an endogenous gene as a marker, the mutation of which can be selected both in the forward (loss of function) and reverse (gain of function) directions. During the first gene editing step, crRNA and mutation-directing oligonucleotides are provided for both the marker and the first target gene (Target 1). Candidates with edited marker genes are selected by loss of marker function, prior to screening for the desired modification of the first target gene. Using a successful candidate, a subsequent gene editing step directs reversion of the mutant marker gene to wild-type status, with candidates being selected on auxotrophic media to detect the regain of function of the auxotrophic marker to wild type (i.e., reversion). Simultaneously, a second target gene modification is produced using Target 2–specific crRNA and oligonucleotides. Revertants, now with a wild-type auxotrophic marker, are then screened for the specific mutation of Target 2. This reversion strategy enables a single selectable marker to be reused indefinitely, facilitating the creation of many successive mutations in a single cell line. As the marker can be completely reconstituted, strains can be created in which only the target gene is altered. Employment of homology-directed repair, using single-stranded oligonucleotides for mutation creation, enables the creation of site-directed mutants, tag insertion, and gene knockouts or reversion, rather than the insertion of large gene cassettes. In this implementation, nitrate reductase is used as the endogenous auxotrophic marker, and the adenine phosphoribosyltransferase gene is used as an example of a target gene.

Chromera velia is an apicomplexan alga uniquely positioned as the closest photosynthetic relative to apicomplexan parasites (Sporozoa), which include the human pathogens that cause malaria (Plasmodium) and toxoplasmosis (Toxoplasma). Under favorable conditions, C. velia forms motile zoospores that contribute to dispersal and possibly host interaction. However, zoospores coexist with other developmental stages in culture, making their isolation technically challenging. Previous studies characterized the phototactic behavior of zoospores in several taxa, yet this response has not been used to separate motile zoospores from mixed cultures. Other reported methods for zoospore recovery relied instead on physical or chemical principles such as passive filtration, differential centrifugation, or column-based purification, all of which can compromise zoospore motility and viability through mechanical shear or osmotic changes. To address this limitation, we developed a non-invasive, simple, and effective method for rapid zoospore isolation depending entirely on their negative phototaxis response. Using a directional light gradient, the method enables reliable collection of active, motile zoospores without specialized equipment or chemical treatments. Our protocol is straightforward to reproduce, relies on standard laboratory equipment, can be completed in under two hours, and yields a zoospore fraction of sufficient quality for live-imaging, motility assays, and downstream molecular and -omics applications. It may also be adapted to other flagellated protists with light-responsive motile stages.

Chlamydomonas reinhardtii is a model green alga extensively used to study photosynthesis and cilia using molecular biology and genetics. Electroporation is a very common technique to integrate DNA into the nuclear genome, which is essential to generate mutant collections and express transgenes. Here, we describe a simple, fast, and efficient protocol to transform strains with an intact cell wall. The technique achieves good transformation efficiency without cell wall digestion or the use of commercial kits and is compatible with the widely available Gene Pulser electroporation system.