Plant Science

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.

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.).

Here we describe two experimental protocols to measure the biomechanical properties of primary (growing) plant cell walls, with a focus on analyzing cell wall epidermal strips of onion scales. The first protocol measures cell wall creep (time-dependent irreversible extension) under constant force. Such creep is often mediated by the wall-loosening action of expansin or selective endoglucanases. The second protocol is based on two consecutive stretches of the wall and measures the wall’s elastic and plastic compliances, which depend on cell wall structure. These two assays provide complementary information that may be linked to cell wall structure and expansive growth of cells.

The shapes of chloroplasts and the architectures of internal thylakoid membranes are altered by growth and environmental changes (Lichtenthaler et al., 1981; Kutik, 1985; Terashima and Hikosaka, 1995). These morphological alterations proceed via transitional intermediates, during which dynamic and heterogeneous thylakoid membranes are observed in cells (Nozue et al., 2017). Light microscopy is useful for the detection of morphological differences in chloroplasts. The thylakoid architecture of such morphologically variable chloroplasts is confirmed by transmission electron microscopy (TEM). The method of monitoring structural variation by light microscopy in combination with electron microscopy is described.

Leaf epidermal cell size and number are positively correlated to leaf area. Stomata are specialized epidermal cells vital for gas exchange and water transpiration. So, observation and statistical analysis of the leaf epidermal cells are valuable for the study of leaf development and response to environmental stimulus. The classical method is using the scanning electron microscope (SEM), which is an expensive and time-consuming method, thus makes the large-scale screening of epidermis impractical. Here we provide simple but effective methods (agarose-based epidermal imprinting and tape-based epidermis tearing) for solving this problem without using the SEM.

Microtubules (MTs) support an astonishing set of versatile cellular functions ranging from cell division, vesicle transport, and cell and tissue morphogenesis in various organisms. This versatility is in large mediated by MT-associated proteins (MAPs). The neuronal MAP Tau, for example, is stabilizing MTs in axons of the vertebrate nervous system and thus provides the basis for enduring axonal transport and the long life span of neurons (Mandelkow et al., 1994). Tau has been shown to bind to MTs directly in vitro and also to promote their nucleation from α-/β-tubulin subunits (Goode et al., 1994). Recently, we identified a plant-specific protein family called “companion of cellulose synthase” (CC), which was shown to bind MTs and enhance dynamics of the cortical MT array in plant cells under salt stress (Endler et al., 2015). The CCs were therefore hypothesized to help plant cells cope with stress conditions and thereby maintain biomass production under adverse growth conditions. Here, we provide detailed experimental information on in vitro MT binding assays, which allow assessing whether a protein of interest is binding to MTs. The assay utilizes the high molecular weight of MTs in a spin down approach and enables the determination of the dissociation constant K_d, a measure for the protein’s binding strength to MTs.

Light microscopy is the standard tool for studying sub-cellular structures however, owing to the diffractive properties of light, resolution is limited to 200 nm. Super-resolution microscopy methods circumvent this limit, offering greater resolution, particularly when studying fluorescently labeled sub-cellular structures. Super-resolution methods such as 3D-SIM (Structured Illumination Microscopy) fill a useful niche between confocal and electron microscopy. We have previously had success using fixed plant tissue samples with 3D-SIM (Bell and Oparka, 2014). However, sensitive structures can be altered by fixation and embedding procedures, so we developed a method for imaging live cells. In this protocol we used 3D-SIM to image the ER and Hechtian Strands in live, plasmolysed BY2 cells.

Chloroplasts accumulate to weak light and escape from strong light. These light-induced responses have been known from the 19th century (Böhm, 1856). Up to now, many scientists have developed different methods to investigate these dynamic phenomena in a variety of plant species including the model plant Arabidopsis thaliana, a terrestrial dicot (Wada, 2013). Especially, a serial recording to trace the position of individual chloroplast for the analysis of its mode of movement is critical to understand the underlying mechanism. An aquatic monocot Vallisneria (Alismatales: Hydrocharitaceae, Figure 1A) has contributed over a century to such investigation (Senn, 1908; Zurzycki, 1955; Seitz, 1967), because Vallisneria leaves have rectangular parallelepiped-shaped epidermal cells aligned orderly in a monolayer (Figure 1B), providing an excellent experimental system for microscopic studies. Here we describe a protocol for the up-to-date time-lapse imaging procedures to analyze Vallisneria chloroplast movement. Using this and prototype procedures, the relevant photoreceptor systems (Izutani et al., 1990; Dong et al., 1995; Sakai et al., 2015), association with actin cytoskeleton (Dong et al., 1996; Dong et al., 1998; Sakai and Takagi 2005; Sakurai et al., 2005), and regulatory roles of Ca²⁺ (Sakai et al., 2015) have been strenuously investigated.


Figure 1. Vallisneria plant. A. Whole plant body; B. A bright-field image of adaxial epidermal cells containing a large number of chloroplasts; C. Culture facilities.

Hydroxyproline-rich glycoproteins (HRGPs) are major protein components in dicot primary cell walls and generally account for more than 10% of the wall dry weight. As essential members of the HRGP superfamily, extensins (EXTs) presumably function in the cell wall by assembling into positively charged protein scaffolds (Cannon et al., 2008) that direct the proper deposition of other wall polysaccharides, especially pectins, to ensure correct cell wall assembly (Hall and Cannon, 2002; Lamport et al., 2011a). Extensins are recalcitrant to purification as they are rapidly cross-linked into a covalent network after entering the cell wall but there exists a short time window in which newly synthesized extensin monomers can be extracted (Smith et al., 1984; Smith et al., 1986) by salt elution. A detailed protocol for extraction of extensin and other wall structural proteins has been described earlier (Lamport et al., 2011b). The protocol elaborated here provides an approach to studying the self-assembly of extensins and potentially of other cell wall components in vitro using AFM.

Plants maintain capacity to form new organs such as leaves, flowers, lateral shoots and roots throughout their postembryonic lifetime. Lateral roots (LRs) originate from a few pericycle cells that acquire attributes of founder cells (FCs), undergo series of anticlinal divisions, and give rise to a few short initial cells. After initiation, coordinated cell division and differentiation occur, giving rise to lateral root primordia (LRP). Primordia continue to grow, emerge through the cortex and epidermal layers of the primary root, and finally a new apical meristem is established taking over the responsibility for growth of mature lateral roots [for detailed description of the individual stages of lateral root organogenesis see Malamy and Benfey (1997)]. To examine this highly dynamic developmental process and to investigate a role of various hormonal, genetic and environmental factors in the regulation of lateral root organogenesis, the real time imaging based analyses represent extremely powerful tools (Laskowski et al., 2008; De Smet et al., 2012; Marhavý et al., 2013; Marhavý et al., 2014). Herein, we describe a protocol for real time lateral root primordia (LRP) analysis, which enables the monitoring of an onset of the specific gene expression and subcellular protein localization during primordia organogenesis, as well as the evaluation of the impact of genetic and environmental perturbations on LRP organogenesis.