植物科学

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.

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.

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.

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.

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.

Rice lodicules are specialized floral organs located at the base of the ovary that undergo dynamic morphological changes during the flowering period. Water uptake–driven swelling and subsequent dehydration-induced shrinkage of the lodicules trigger floret opening and closure, respectively. Although lodicules play a central role in floret movement, standardized methods for quantitatively monitoring their temporal morphological changes remain limited. Here, we describe a detailed and reproducible workflow for lodicule sampling, dissection, imaging, and quantitative morphometric analysis. Florets are collected at predefined clock time points during the flowering period, and lodicules are carefully isolated under a stereomicroscope. High-resolution imaging is performed under consistent acquisition settings, followed by precise measurement of lodicule length, width, and thickness using image analysis software. This protocol emphasizes positional consistency in sampling, uniform imaging parameters, and standardized data analysis to enhance reproducibility. This method is suitable for evaluating the effects of genetic background or environmental conditions on lodicule morphology. By providing a standardized analytical framework, this protocol enables accurate and quantitative morphometric analysis of rice lodicules during floret opening.

Conventional light microscopy is limited in resolution by the diffraction limit of light, restricting the visualization of the nanoscale organization of biomolecules. Expansion microscopy (ExM) has emerged as a powerful technique to overcome this barrier by physically expanding the specimen embedded in a swellable hydrogel without requiring specialized or high-cost imaging hardware. ExM is widely used in animal models, whereas its application to plant tissues has been challenging due to their multicellularity, in which each cell is encompassed by the rigid cell wall, which resists the expansion forces and prevents isotropic swelling. Here, we describe a robust and optimized ExM protocol specifically designed for Arabidopsis thaliana root tissues. This protocol details critical steps, including immunostaining, anchoring, gelation, denaturation, cell wall digestion, and expansion. Our method achieves an expansion factor of approximately 4.3×, enabling an effective lateral resolution of ~60 nm using a standard confocal microscope. We demonstrate the visualization of microtubules with preserved ultrastructure. This accessible protocol allows plant researchers to perform super-resolution imaging without specialized optical equipment, facilitating detailed structural analysis of plant cells.

Cellular function relies on a network of precisely regulated interactions among macromolecules such as proteins, peptides, carbohydrates, and nucleic acids. These molecular interactions regulate vital processes, including signaling, structural organization, and developmental patterning. Biolayer interferometry (BLI) is a label-free optical biosensing technique that enables real-time quantification of such interactions. This protocol describes how to use BLI to assess the binding affinity between a biotinylated plant peptide hormone (RALF1) and cell wall–derived oligogalacturonides (OG25–50) on the Octet RED96 platform. Streptavidin-coated biosensors are employed to immobilize the ligand, while analyte binding is monitored through wavelength shifts in the reflected light. The protocol includes detailed steps for sensor preparation, assay setup, software configuration, and kinetic data analysis. While optimized for plant peptide–matrix interactions, the method is broadly adaptable to other macromolecular systems across biological disciplines.

Understanding cellular growth dynamics in plants requires precise, long-term imaging of developing tissues. Cauline leaves are produced during the transition from vegetative to reproductive development and provide a useful system for studying how laminar organs diversify in form and function. While other laminar organs, such as rosette leaves and sepals, have been extensively studied, early cauline leaf development remains technically challenging to capture due to their concealed position, curved morphology, and the presence of dense trichomes. Here, we provide a complete pipeline for the dissection, confocal imaging, 2.5D segmentation, and image analysis of initiating cauline leaves in Arabidopsis thaliana. This method enables reproducible, high-resolution imaging of cauline leaves, supporting robust quantitative analysis of growth across developmental stages at cellular scale resolution.

Volume electron microscopy based on serial sectioning allows for three-dimensional (3D) visualization and analysis of the internal structures of tissues, cells, and organelles. One such technique, focused ion beam (FIB) scanning electron microscopy (SEM), has the advantages of nanoscale sectioning and high z-resolution, but the disadvantage of limited volume processing. Because of this limitation, targeting localized objects by FIB-SEM is difficult. Here, we developed a FIB-SEM observation workflow that enables the analysis of the filiform apparatus of synergid cells enclosed in the Arabidopsis ovule. In this protocol, plant samples are stained, embedded, trimmed, and carbon-coated while maintaining their orientation within the tissue. Then, sequential observations are performed using Cut & See function of FIB-SEM, followed by image processing for 3D reconstruction. Utilization of multi-scanning and image cropping from high-resolution data helps to identify localized targets within plant tissue. The filiform apparatus, which is an invaginated cell wall structure of the synergid cells, shows distinct contrast in each image, allowing for segmentation using brightness-based binarization. Such segmentation avoids the need to manually trace complex structures and facilitates 3D reconstruction by volume electron microscopy.