Plant Science

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.

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.

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.

Plant-derived extracellular vesicles (PDEVs) have emerged as important mediators of intercellular communication and hold growing potential in therapeutic applications. However, standardized methods for their isolation, particularly from Piper betle leaves (PBL), remain unexplored. Existing apoplastic fluid washing (AFW) extraction techniques typically rely on manual syringe infiltration, which often leads to inconsistent pressure control, variable yields, and increased risk of tissue damage. This protocol describes a vacuum-assisted AFW extraction method optimized for the recovery of intact extracellular vesicles (EVs) from PBL. The workflow features controlled negative pressure using a vacuum pump and chamber to achieve more efficient leaf infiltration compared to infiltration using the syringe method and reproducible apoplastic fluid (AF) collection with subsequent low-speed centrifugation steps, to ensure minimal contamination and preservation of vesicle integrity. Piper betle–derived extracellular vesicle (PBdEV) isolation and purification steps are performed using size exclusion chromatography (SEC). The size and concentration of PBdEVs were confirmed using nanoparticle tracking analysis (NTA), whereas the cup-shaped and lipid bilayer morphology of the EVs were confirmed using transmission electron microscopy (TEM). The method is scalable and adaptable to various leaf morphologies and physiological states, making it suitable for both exploratory and high-throughput studies. Overall, this protocol provides a more consistent, efficient, and tissue-preserving alternative to traditional syringe-based AF extraction methods, offering higher-quality EV preparations for plant EV research.

Calcium ions serve as a universal secondary messenger, integrating diverse external signals, such as light, herbivory, and mechanical stimuli, within plant cells. However, the visualization and mechanistic dissection of calcium signaling specifically in response to mechanical stimulation remain technically challenging and underexplored in most plants. Previous studies have been largely confined to a few model systems, including Arabidopsis; here, we introduce a live-cell imaging approach using the stigmas of Torenia fournieri. This in vitro system enables multiscale observation of calcium signal patterns following controlled mechanical stimulation. This versatile platform not only simplifies the design of calcium imaging assays but also provides a tractable system for functionally validating other key molecular components in this signaling pathway.

While traditional kinetic studies of the cytochrome b₆f complex have frequently relied on measurements within the complex environment of intact leaves or whole-organism systems, such approaches can be limited by overlapping signals and physiological variables. This protocol advances existing frameworks by introducing a streamlined, multi-wavelength spectroscopic approach utilizing a reconstituted in vitro system to elucidate the inter-complex electron transfer kinetics between photosystem I and cytochrome b₆f. Utilizing the JTS-150 pulsed spectrometer, supplied with a Smart Lamp, we monitored the redox transitions of P700⁺ and Cytf by simultaneously measuring the absorbance changes of our isolated complexes system in six different wavelengths (546, 554, 563, 574, 705, and 740 nm). Kinetic analysis was divided into two phases: laser-induced flash kinetics and steady-state actinic induction. We resolved the second-order re-reduction of P700⁺ by plastocyanin, accounting for detector saturation constraints with a 2 ms post-flash delay. Steady-state measurements under actinic light revealed complex Cytf turnover, characterized by a double-exponential decay. Furthermore, dark relaxation kinetics were used to quantify ferredoxin-mediated re-reduction of the cytochrome pool. By allowing the incorporation of specific regulatory and inhibitory factors, this methodology sets the ground for the deconvolution of competing electron pathways. It can therefore be used as a robust framework for assessing the mechanism of regulatory processes on photosynthetic flux.

Plant genome editing is a powerful approach for modifying plant DNA to investigate gene function and to engineer desirable traits. Several genome-editing technologies have been developed, among which CRISPR/Cas systems and transcription activator-like effector nucleases (TALENs) are widely used to introduce targeted double-stranded DNA breaks. While CRISPR/Cas systems are highly efficient for nuclear genome editing, their application to plant organellar genomes remains limited, largely due to difficulties in guide RNA delivery into mitochondria and chloroplasts. Here, we present a detailed and reproducible protocol for constructing TALEN-based binary vectors for targeted genome editing in Arabidopsis thaliana. This protocol describes the assembly of TALE repeat arrays, the generation of nuclear-, mitochondrial-, and plastid-targeted TALEN expression vectors using MultiSite Gateway cloning, and subsequent Agrobacterium-mediated plant transformation and genotyping. The workflow enables the production of nTALENs, mitoTALENs, and ptpTALENs using a unified vector design strategy. In addition, the protocol briefly outlines the construction principles of TALE-based cytidine deaminases (TALECDs) for targeted C-to-T base editing in plant organellar genomes. The protocol provides a flexible and robust framework for plant nuclear and organellar genome editing and can be readily adapted to different target genes and experimental purposes. Its modular design and compatibility with standard molecular cloning techniques make it accessible to laboratories aiming to perform precise genome manipulation in plants.

We present a protocol to allow continuous assessment of cell death in Arabidopsis thaliana (L.) seedlings by measuring the release of electrolytes from dying cells upon heat shock. The electrolyte leakage assay is a well-established method to quantify the extent of cell death of plant tissues exposed to pathogen infection, since the activation of the immune response leads to compromised membrane integrity and to the release of ions from the dying cell. This prolonged release of electrolytes is considered a hallmark of regulated cell death in plants. Heat shock in plants induces ferroptosis-like cell death, which can be suppressed either pharmacologically, using inhibitors such as ferrostatin, or genetically through knockout of ferroptosis-related genes. Here, we have adapted the electrolyte leakage assay to quantify cell death in young Arabidopsis seedlings exposed to a heat shock previously shown to induce ferroptosis-like cell death. We also illustrate how this method can be used to assess activation of ferroptosis-like cell death in whole Arabidopsis seedlings using ferrostatin or knockout mutants of potential gene candidates involved in ferroptosis-like cell death.

Aloe vera has long been used for its diverse pharmacological properties, motivating continued interest in isolating and preserving the bioactive molecules responsible for its therapeutic potential. More recently, Aloe vera–derived extracellular vesicles (Av-EVs) have emerged as nanoscale, cell-free carriers capable of retaining and delivering these properties, making them attractive for various biomaterials, nanomedicine, and regenerative medicine applications. Multiple techniques are available for extracellular vesicle isolation. These include ultracentrifugation, polymer-based precipitation, size-exclusion chromatography, immunoaffinity capture, ultrafiltration, density gradient separation, and emerging microfluidic platforms. Each method presents distinct trade-offs in purity, yield, scalability, and downstream compatibility. Despite this diversity, standardized workflows tailored to Av-EV isolation remain limited, and the influence of homogenization-induced shear forces and plant maturity on vesicle recovery and characterization has not been systematically addressed. Here, we present a reproducible protocol for isolating Av-EVs from Aloe vera gel employing two distinct homogenization strategies: manual, no-shear force (NB EVs), and blender-based shear-force homogenization (B EVs). The workflow covers gel preparation, serial centrifugation for debris removal, ultracentrifugation as the gold standard for vesicle enrichment, and final sterile filtration. This protocol enables consistent recovery of Av-EVs suitable for physicochemical characterization and functional analyses. It is simple and relies on commonly available laboratory equipment, facilitating broad adoption by ultracentrifugation users and offering adaptability to diverse research projects involving purified Aloe vera gel and Av-EVs, including studies focused on wound healing, fibrotic scarring, and regenerative processes, where coordinated antioxidant, anti-inflammatory, antimicrobial, immunomodulatory, and moisturizing responses are of interest.

Reactive oxygen species (ROS) are central regulators of plant development and stress responses, with hydrogen peroxide (H₂O₂) acting as a key signaling molecule whose spatial distribution determines adaptive versus damaging outcomes. Accurate detection of H₂O₂ at tissue and cellular resolution is therefore essential for understanding redox-dependent regulation of plant growth. A variety of techniques have been used to monitor H₂O₂, including bulk spectrophotometric and fluorometric assays, genetically encoded sensors for real-time measurements, and chemical probes for in situ detection. While these approaches differ in sensitivity, specificity, and temporal resolution, many are limited by a lack of spatial information, technical complexity, or dependence on transgenic material. Here, we present a detailed protocol for 3,3′-diaminobenzidine (DAB)-based histochemical detection of H₂O₂ in seedling roots, covering staining, imaging, and semi-quantitative image analysis using open-source software (FIJI/ImageJ). The method relies on peroxidase-mediated oxidation of DAB, resulting in a stable, light-resistant, and insoluble precipitate that enables visualization of H₂O₂ accumulation with high spatial resolution. This protocol provides a robust, accessible, and genetically independent approach for spatial analysis of H₂O₂ in plant tissues. Its simplicity, compatibility with diverse genotypes and treatments, and suitability for semi-quantitative analysis make it a valuable tool for examining the spatial distribution of H₂O₂, thereby providing spatial insight into redox-related regulatory processes during plant development and stress responses.