Plant Science

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.

Citrus fruits encompass a diverse family, including oranges, mandarins, grapefruits, limes, kumquats, lemons, and others. In citrus, Agrobacterium tumefaciens–mediated genetic transformation of Hongkong kumquat (Fortunella hindsii Swingle) has been widely employed for gene function analysis. However, the perennial nature of woody plants results in the generation of transgenic fruits taking several years. Here, we show the procedures of Agrobacterium-mediated transient transformation and live-cell imaging in kumquat (F. crassifolia Swingle) fruit, using the actin filament marker GFP-Lifeact as an example. Fluorescence detection, western blot analysis, and live-cell imaging with confocal microscopy demonstrate the high transformation efficiency and an extended expression window of this system. Overall, Agrobacterium-mediated transient transformation of kumquat fruits provides a rapid and effective method for studying gene function in fruit, enabling the effective observation of diverse cellular processes in fruit biology.

Studies on chromosomal status are a fundamental aspect of plant cytogenetics and breeding because changes in number, size, and shape of chromosomes determine plant physiology/performance. Despite its significance, the classical cytogenetic study is now frequently avoided because of its tedious job. In general, root meristems are used to study the mitotic chromosome number, even though the use of root tips was restricted because of sample availability, processing, and lack of standard protocols. Moreover, to date, a protocol using shoot tips to estimate chromosome number has not yet been achieved for tree species’ germplasm with a large number of accessions, like mulberry (Morus spp.). Here, we provide a step-by-step, economically feasible protocol for the pretreatment, fixation, enzymatic treatment, staining, and squashing of meristematic shoot tips. The protocol is validated with worldwide collections of 200 core set accessions with a higher level of ploidy variation, namely diploid (2n = 2x = 28), triploid (2n = 3x = 42), tetraploid (2n = 4x = 56), hexaploid (2n = 6x = 84), and decosaploid (2n = 22x = 308) belonging to nine species of Morus spp. Furthermore, accession from each ploidy group was subjected to flow cytometry (FCM) analysis for confirmation. The present protocol will help to optimize metaphase plate preparation and estimation of chromosome number using meristematic shoot tips of tree species regardless of their sex, location, and/or resources.

Expansion microscopy is an innovative method that enables super-resolution imaging of biological materials using a simple confocal microscope. The principle of this method relies on the physical isotropic expansion of a biological specimen cross-linked to a swellable polymer, stained with antibodies, and imaged. Since its first development, several improved versions of expansion microscopy and adaptations for different types of samples have been produced. Here, we show the application of ultrastructure expansion microscopy (U-ExM) to investigate the 3D organization of the green algae Chlamydomonas reinhardtii cellular ultrastructure, with a particular emphasis on the different types of sample fixation that can be used, as well as compatible staining procedures including membranes.

Graphical overview

Since the genetic transformation of Chinese cabbage (Brassica rapa) has not been well developed, in situ RT-PCR is a valuable option for detecting guard cell–specific genes. We reported an optimized protocol of in situ RT-PCR by using a FAMA homologous gene Bra001929 in Brassica rapa. FAMA in Arabidopsis has been verified to be especially expressed in guard cells. We designed specific RT-PCR primers and optimized the protocol in terms of the (a) reverse transcription time, (b) blocking time, (c) antigen-antibody incubation time, and (d) washing temperature. Our approach provides a sensitive and effective in situ RT-PCR method that can detect low-abundance transcripts in cells by elevating their levels by RT-PCR in the guard cells in Brassica rapa.

Genome sizes of Zygnema spp. vary greatly, being unknown whether polyploidization occurred. The exact number of chromosomes in this genus is unknown since counting methods established for higher plants cannot be applied to green algae. The massive presence of pectins and arabinogalactan proteins in the cell wall interferes with the uptake of staining solutions; moreover, cell divisions in green algae are not restricted to meristems as in higher plants, which is another limiting factor. Cell divisions occur randomly in the thallus, due to the intercalary growth of algal filaments. Therefore, we increased the number of cell divisions via synchronization by changing the light cycle (10:14 h light/dark). The number of observed mitotic stages peaked at the beginning of the dark cycle. This protocol describes two methods for the visualization of chromosomes in the filamentous green alga Zygnema. Existing protocols were modified, leading to improved acetocarmine and haematoxylin staining methods as investigated by light microscopy. A freeze-shattering approach with liquid nitrogen was applied to increase the accessibility of the haematoxylin dye. These modified protocols allowed reliable chromosome counting in the genus Zygnema.

Key features

• Improved method for chromosome staining in filamentous green algae.

• Optimized for the Zygnema strains SAG 698-1a (Z. cylindricum), SAG 698-1b (Z. circumcarinatum), and SAG 2419 (Zygnema ‘Saalach’).

• This protocol builds upon the methods of chromosomal staining in green algae developed by Wittmann (1965), Staker (1971), and Fujii and Guerra (1998).

• Cultivation and synchronization: 14 days; fixation and permeabilization: 24 h; staining: 1 h; image analysis and chromosome number quantification: up to 20 h.

In vivo microscopy of plants with high-frequency imaging allows observation and characterization of the dynamic responses of plants to stimuli. It provides access to responses that could not be observed by imaging at a given time point. Such methods are particularly suitable for the observation of fast cellular events such as membrane potential changes. Classical measurement of membrane potential by probe impaling gives quantitative and precise measurements. However, it is invasive, requires specialized equipment, and only allows measurement of one cell at a time. To circumvent some of these limitations, we developed a method to relatively quantify membrane potential variations in Arabidopsis thaliana roots using the fluorescence of the voltage reporter DISBAC₂(3). In this protocol, we describe how to prepare experiments for agar media and microfluidics, and we detail the image analysis. We take an example of the rapid plasma membrane depolarization induced by the phytohormone auxin to illustrate the method. Relative membrane potential measurements using DISBAC₂(3) fluorescence increase the spatio-temporal resolution of the measurements and are non-invasive and suitable for live imaging of growing roots. Studying membrane potential with a more flexible method allows to efficiently combine mature electrophysiology literature and new molecular knowledge to achieve a better understanding of plant behaviors.

Key features

• Non-invasive method to relatively quantify membrane potential in plant roots.

• Method suitable for imaging seedlings root in agar or liquid medium.

• Straightforward quantification.