Plant Science

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.

Lysosome-related organelles (LROs) are a class of heterogeneous subcellular organelles conserved in eukaryotes, performing various functions. An important function of LROs is to mediate phosphorus and metal homeostasis. Chlamydomonas reinhardtii serves as a model organism for investigating metal ion metabolism. Considering that LROs contain polyphosphate and various metal elements, the purification strategy is based on their higher density by fractionating cell lysate through OptiPrep density gradient ultracentrifugation. Here, we optimized a method for purifying LROs from C. reinhardtii cells that have reached stationary phase (sta-LROs) or are overloaded with iron (Fe-LROs). Our protocol provides technical support for further investigations on the biogenesis and function of LROs in C. reinhardtii.

Plants use CO₂, water, and light energy to generate carbohydrates through photosynthesis. During daytime, these carbohydrates are polymerized, leading to the accumulation of starch granules in chloroplasts. The catabolites produced by the degradation of these chloroplast starch granules are used for physiological responses and plant growth. Various staining methods, such as iodine staining, have previously been used to visualize the accumulation of chloroplast starch granules; however, these staining methods cannot be used to image live cells and/or provide confocal images with non-specific signals. In this study, we developed a new imaging method for the fluorescent observation of chloroplast starch granules in living plant cells by staining with fluorescein, a widely available fluorescent dye. This simple staining method, which involves soaking a leaf disk in staining solution, shows high specificity in confocal images. Fluorescent images of the stained tissue allow the cellular starch content of living cells to be quantified with the same level of accuracy as a conventional biochemical method (amyloglucosidase/α-amylase method). Fluorescein staining thus not only enables the easy and clear observation of chloroplast starch granules but also allows for precise quantification in living cells.

Extracellular vesicles are membrane-bound organelles that play crucial roles in intercellular communication and elicit responses in the recipient cell, such as defense responses against pathogens. In this study, we have optimized a protocol for isolating extracellular vesicles (EVs) from Sorghum bicolor apoplastic wash. We characterized the EVs using fluorescence microscopy and correlative light and electron microscopy.

Stomata are pores surrounded by a pair of specialized cells, called guard cells, that play a central role in plant physiology through the regulation of gas exchange between plants and the environment. Guard cells have features like cell-autonomous responses and easily measurable readouts that have turned them into a model system to study signal transduction mechanisms in plants. Here, we provide a detailed protocol to analyze different physiological responses specifically in guard cells. We describe, in detail, the steps and conditions to isolate epidermal peels with tweezers and to analyze i) stomatal aperture in response to different stimuli, ii) cytosolic parameters such as hydrogen peroxide (H₂O₂), glutathione redox potential (E_GSH), and MgATP^-2 in vivo dynamics using fluorescent biosensors, and iii) gene expression in guard cell–enriched samples. The importance of this protocol lies in the fact that most living cells on epidermal peels are guard cells, enabling the preparation of guard cell–enriched samples.

The roots of herbaceous and woody plants growing in soil are complex structures that are affected by both natural and artificial fungal colonization to various extents. To obtain comprehensive information about the overall distribution of fungi or oomycetes inside a plant root system, rapid, effective, and reliable screening methods are required. To observe both fine roots, i.e., a common site for penetration of fungi and oomycetes, and mature roots, different techniques are required to overcome visual barriers, such as root browning or tissue thickening. In our protocol, we propose using fast, cost-effective, and non-harmful methods to localize fungal or oomycete structures inside plant roots. Root staining with a fluorescent dye provides a quick initial indication of the presence of fungal structures on the root surfaces. The protocol is followed by clearing and staining steps, resulting in a deeper insight into the root tissue positioning, abundance, and characteristic morphological/reproductive features of fungal or oomycete organisms. If required, the stained samples can be prepared by using freeze-drying for further observations, including advanced microscopic techniques.