植物科学

Identifying microscopic mycorrhizal fungal structures in roots, i.e., hyphae, vesicles and arbuscules, requires root staining procedures that are often time consuming and involves chemicals known to present health risks from exposure. By modifying established protocols, our root staining method stains roots using a safe ink- and vinegar-based staining solution, followed by a 2-16 h-long de-staining period. The entire procedure can be completed in less than 6 h (plus up to 16 h de-staining overnight) and roots are suitable for semi-permanent and permanent slide mounting for light microscopy. We tested our method on hundreds of wild-sourced roots from two different plant species: Lycopodiella inundata, a herbaceous clubmoss with tough water-resistant roots, and Sambucus nigra, a temperate woody shrub. Both plants associate with endomycorrhizae, L. inundata predominantly with Mucoromycotina fine root endophytes (MucFRE) and S. nigra with Glomeromycota arbuscular mycorrhizal fungi (AMF). Here we describe a simple, efficient, repeatable and safe method to detect the presence of fungal structures using light microscopy.

Although it is widely accepted that actin plays an important role in regulating pollen germination and pollen tube growth, how actin exactly performs functions remains incompletely understood. As the function of actin is dictated by its spatial organization, it is the key to reveal how exactly actin distributes in space in pollen cells. Here we describe the protocol of revealing and quantifying the spatial organization of actin using fluorescent phalloidin-staining in fixed Arabidopsis pollen grains and pollen tubes. We also introduce the method of assessing the stability and/or turnover rate of actin filaments in pollen cells using the treatment of latrunculin B.

Natural hosts for the fungal endophyte Epichloë festucae include Festuca rubra (fine fescue) and Festuca trachyphylla (hard fescue). Some strains also form stable associations with Lolium perenne (perennial ryegrass). L. perenne is a suitable host to study fungal endophyte–grass interactions, such as endophytic fungal growth within the plant and epiphyllous growth on the plant surface. Here we provide a detailed protocol based on work by, for artificial inoculation of E. festucae into L. perenne, and newly developed staining and visualization techniques for observing endophytic and epiphyllous hyphae and the expressorium, an appressorium-like structure used by the fungus to exit the plant. The staining method uses a combination of glucan binding aniline blue diammonium salt (AB) and chitin binding wheat germ agglutinin-conjugated Alexa Fluor^®488 -(WGA-AF488). This protocol will be a useful tool to study Epichloë-grass interactions, particularly the comparison of different Epichloë-grass associations, various endophyte-host developmental stages, as well as the analysis of mutant Epichloë strains.

To investigate the chromosome dynamics during mitosis, it is convenient to mark the discrete chromosome foci and then analyze their spatial rearrangements during prophase condensation and telophase decondensation. To label the chromosome regions in plant chromosomes, we incorporated the synthetic nucleotide, 5-ethynyl-2’-deoxyuridine (EdU), which can be detected by click-chemistry, into chromatin during replication. Here, we described a protocol of a method based on the application of semi-thin sections of Nigella damascena L. roots embedded in LR White acrylic resin. The thickness of semi-thin (100-250 nm) sections is significantly lower than that of optical sections even if a confocal microscope was used. This approach may also be suitable for work with any tissue fragments or large cells (oocytes, cells with polytene chromosomes, etc.).

The opening of stomata in plants in response to blue light is driven by the plasma membrane H⁺-ATPase in guard cells. To evaluate the activation of the H⁺-ATPase in vivo, we can use H⁺-pumping by guard cells in response to blue light and fusicoccin. To do this, it is required to prepare a large amount of guard cell protoplasts and measure H⁺-pumping in the protoplasts. It is also necessary to determine the protein amount of H⁺-ATPase. In this protocol, we describe the procedures required for these preparations and measurements.

Membrane damage is a hallmark of both biotic and abiotic stress responses. The membrane determines the ability of a cell to sustain altered environmental conditions and hence can be used as a biomarker to assess stress-induced cell damage or death. We present an easy, quick, cost-effective, staining and spectrophotometric method to assess membrane stability of plant cells. In this method, Evan’s blue, an azo dye, is used to assay for cell viability. More specifically, Evan’s blue dye can penetrate through ruptured or destabilized membranes and stain cells. Thus, when plant cells are subjected to stress that compromises membrane integrity, the number of cells that are permeated by Evan’s blue dye will be increased compared to control cells that are not stressed. In contrast, live, healthy cells that are capable of maintaining membrane integrity do not take up Evan’s blue dye. Cells that have taken up Evan’s blue dye will have an accumulation of a blue protoplasmic stain and these stained cells can be qualitatively documented under bright field microscopy with or without the use of a camera. Furthermore, the dye can be extracted from cells that are stained by Evan’s blue dye and can be quantified spectrophotometrically. Using this analysis, the accumulation of dye in positively-stained cells correlates with the extent of cell membrane damage and thus the amount of cells that are stained with Evan’s blue dye under various conditions can be used as an indicator of cellular stress.

Confocal laser scanning microscopy in combination with fluorescent proteins is a powerful tool for the study of sexual reproduction and other developmental processes in plants. In order to understand the origin and localization of fluorescent signals in a complex tissue, staining of cell outlines is often mandatory. Cell wall staining with SCRI Renaissance 2200 (SR2200) has recently been described as a method of choice to study plant reproductive processes (Musielak et al., 2015). In this protocol, we present detailed instructions on the use of SR2200 to stain cell walls in different Arabidopsis tissues.

Callose is an amorphous homopolymer, composed of β-1, 3-glucan, which is widespread in higher plants. Callose is involved in multiple aspects of plant growth and development. It is synthetized in plants at the cell plate during cytokinesis, in several stages during pollen development and is deposited at plasmodesmata to regulate the cell-to-cell movement of molecules. Moreover, it is produced in response to multiple biotic and abiotic stresses (Chen and Kim, 2009). Callose is considered to act as a physical barrier by strengthening the plant cell well to slow pathogen infection and to contribute to the plant’s innate immunity. Thus the callose staining method is useful to quantify activity of plant immunity. In addition, this staining can be used to visualize structures in plant tissue, where the callose may be implied whether during the development of plants or response against pathogen infection. This method is based on the use of methyl blue which reacts with (1→3)-β-glucans to give a brilliant yellow fluorescence in UV light. Moreover, calcofluor stains chitin present in fungal cell membranes and also binds to cellulose at locations where the cuticle is damaged.

Silicon (Si) is a biologically important element for plants in the order Poales (Yamamoto et al., 2011; Kido et al., 2015). In rice, Si is mainly deposited in the motor cells and the cell walls of the leaf epidermis. However, the molecular basis of this overall process has not been elucidated. Thus, we propose a protocol for the histochemical staining of the silica body based on specific hydrogen bonding between silanol group and the carboxylate group of crystal violet lactone (Ichimura et al., 2008), as described by Isa et al. (2010), but with minor modifications. This modified protocol can be used for observing Si accumulation during rice development.

Export of transcribed mRNAs from nucleoplasm to cytosol is an essential process for the translation of genes into proteins. This process is tightly regulated by nuclear pores, composed of about 30 nucleoporin proteins (Nups). Whether or not the mRNAs are able to be appropriately exported to cytoplasm is of an importance for understanding the role of Nups. Here, we describe a practical protocol to detect the intracellular localization of mRNAs in mesophyll cells of Nicotiana benthamiana (N. benthamiana). This protocol is based on poly (A) in situ hybridization method using an oligo d(T) probe conjugated with Alexa Fluor-488.