Plant Science

In response to environmental changes, chloroplasts, the cellular organelles responsible for photosynthesis, undergo intracellular repositioning, a phenomenon known as chloroplast movement. Observing chloroplast movement within leaf tissues remains technically challenging in leaves consisting of multiple cell layers, where light scattering and absorption hinder deep tissue visualization. This limitation has been particularly problematic when analyzing chloroplast movement in the mesophyll cells of C₄ plants, which possess two distinct types of concentrically arranged photosynthetic cells. In response to stress stimuli, mesophyll chloroplasts aggregate toward the inner bundle sheath cells. However, conventional methods have not been able to observe these chloroplast dynamics over time in living cells, making it difficult to assess the influence of adjacent bundle sheath cells on this movement. Here, we present a protocol for live leaf section imaging that enables long-term and detailed observation of chloroplast movement in internal leaf tissues without chemical fixation. In this method, a leaf blade section prepared either using a vibratome or by hand was placed in a groove made of a silicone rubber sheet attached to a glass slide for microscopic observation. This technique allows for the quantitative tracking of chloroplast movement relative to the surrounding cells. In addition, by adjusting the sectioning angle and thickness of the unfixed leaf sections, it is possible to selectively inactivate specific cell types based on their size and shape differences. This protocol enables the investigation of the intercellular interactions involved in chloroplast dynamics in leaf tissues.

Amyloplasts, non-photosynthetic plastids specialized for starch synthesis and storage, proliferate in storage tissue cells of plants. To date, studies of amyloplast replication in roots and the ovule nucelli from various plant species have been performed using electron and fluorescence microscopy. However, a complete understanding of amyloplast replication remains unclear due to the absence of experimental systems capable of tracking their morphology and behavior in living cells. Recently, we demonstrated that Arabidopsis ovule integument could provide a platform for live-cell imaging of amyloplast replication. This system enables precise analysis of amyloplast number and shape, including the behavior of stroma-filled tubules (stromules), during proplastid-to-amyloplast development in post-mitotic cells. Here, we provide technical guidelines for observing and quantifying amyloplasts using conventional fluorescence microscopy in wild-type and several plastid-division mutants of Arabidopsis.

Cell subfractionation is a common technique employed in many research laboratories to isolate organelles or intracellular compartments for the study of metabolism or biomolecule purification. While numerous protocols exist for isolating organelles, few are specifically designed for starting materials in the milligram range. Here, we present a detailed milligram-scale miniprep protocol for purifying intact chloroplasts from Arabidopsis thaliana leaves. This chloroplast miniprep procedure is suitable for applications such as confocal microscopy, western blotting, enzymatic assays, and other downstream analyses.

In live-cell imaging, autofluorescence is often regarded as a negative factor that interferes with the accurate visualization of target fluorescence due to a phenomenon known as crosstalk. However, autofluorescence has also been effectively utilized as an organellar marker. For instance, the intense autofluorescence of chlorophyll in the red wavelength is widely used to visualize chloroplasts, the photosynthetic organelle in plants. Recently, we demonstrated that nuclei in plant cells emit phytochrome-derived autofluorescence in the red to infrared wavelength range, which can be visualized by a conventional confocal microscope equipped with a 640 nm laser. Here, we present protocols for growing plants and conducting confocal imaging of the near-infrared autofluorescence of nuclei in Arabidopsis thaliana.

Recordings of electric potential changes on plant surfaces have been utilized to identify the components and mechanisms involved in the formation and transmission of systemic signals elicited by stimuli such as herbivory, wounding, or burning. The recorded responses, commonly referred to as slow wave or variation potentials, exhibit striking variability in their waveform. The extent to which this variability is due to differences in experimental procedures or plant biological variability remains unclear. Here, we provide a detailed and robust protocol refined from years of experience in conducting leaf surface potential recordings of Arabidopsis thaliana in response to mechanical wounding. This protocol serves as a comprehensive tutorial covering plant growth, procedures for reproducible mechanical wounding, critical aspects of electrophysiological recordings, and statistical analysis of surface potential recordings. It particularly emphasizes the construction and maintenance of electrodes, placement of the reference or ground electrode, mechanisms for wounding, and data analysis. This protocol aims to promote and facilitate the adoption, standardization, and interoperability of plant surface potential recordings among research groups, thereby increasing the reproducibility and comparability of data within the field.

Extracellular vesicles (EVs) are membrane-bound, non-replicating particles released by virtually all types of cells. EVs concentrate and deliver a plethora of biomolecules driving very important biological functions, including intercellular communication not only between cells of the same organism but also across different kingdoms. Plant extracellular vesicles (PEVs) are a promising alternative to mammalian EVs in biomedical applications. Here, we present an optimized and reproducible protocol for isolating PEVs from the hairy root (HR) cultures of medicinal plants Salvia dominica and S. sclarea. Our methodological approach introduces a significant advancement in the standardization of HR-EVs purification processes from plant biotechnological platforms, paving the way for their broader application across various sectors, including agriculture, pharmaceuticals, and nutraceuticals.

In nature, filamentous fungi interact with plants. These fungi are characterized by rapid growth in numerous substrates and under minimal nutrient requirements. Investigating the interaction of these fungi with their plant hosts under controlled conditions is of importance for many researchers aiming to proceed with molecular or microscopical investigations of their favorite plant–fungus interaction system. The speed of growth of these fungi complicates transferring plant–fungal interaction systems in laboratory conditions. The issue is more complicated when monoxenic conditions are desired, to ensure that only two members (a fungus and a plant) are present in the system under study. Here, two simple closed systems for investigating plant–filamentous fungi associations under laboratory, monoxenic conditions are described, along with their limitations. The plant and fungal growth conditions, methods for sampling, staining, sectioning, and subsequent microscopical imaging of colonized plant tissues with affordable, common laboratory tools are described.

CRISPR/Cas9 genome editing technology has revolutionized plant breeding by offering precise and rapid modifications. Traditional breeding methods are often slow and imprecise, whereas CRISPR/Cas9 allows for targeted genetic improvements. Previously, direct delivery of Cas9-single guide RNA (sgRNA) ribonucleoprotein (RNP) complexes to grapevine (Vitis vinifera) protoplasts has been demonstrated, but successful regeneration of edited protoplasts into whole plants has not been achieved. Here, we describe an efficient protocol for obtaining transgene/DNA-free edited grapevine plants by transfecting protoplasts isolated from embryogenic callus and subsequently regenerating them. The regenerated edited plants were comparable in morphology and growth habit to wild-type controls. This protocol provides a highly efficient method for DNA-free genome editing in grapevine, addressing regulatory concerns and potentially facilitating the genetic improvement of grapevine and other woody crop plants.

Understanding how multicellular organisms are shaped requires high-resolution, quantitative data to unravel how biological structures grow and develop over time. In recent years, confocal live imaging has become an essential tool providing insights into developmental dynamics at cellular resolution in plant organs such as leaves or meristems. In the context of flowers, growth tracking has primarily been limited to sepals, the outermost floral organs, or the post-fertilization gynoecium, which are easily accessible for microscopy. Here, we describe a detailed pipeline for the preparation, dissection, and confocal imaging of the development of internal reproductive floral organs of Arabidopsis thaliana including both the stamen and gynoecium. We also discuss how to acquire high-quality images suitable for efficient 2D and 3D segmentation that allow the quantification of cellular dynamics underlying their development.

In plants, the first interaction between the pollen grain and the epidermal cells of the stigma is crucial for successful reproduction. When the pollen is accepted, it germinates, producing a tube that transports the two sperm cells to the ovules for fertilization. Confocal microscopy has been used to characterize the behavior of stigmatic cells post-pollination [1], but it is time-consuming since it requires the development of a range of fluorescent marker lines. Here, we propose a quick, high-resolution imaging protocol using tabletop scanning electron microscopy. This technique does not require prior sample fixation or fluorescent marker lines. It effectively captures pollen grain behavior from early hydration (a few minutes after pollination) to pollen tube growth within the stigma (1 h after pollination) and is particularly efficient for tracking pollen tube paths.