Plant Science

Cryo-electron tomography (cryo-ET) is a formidable technique to observe the inner workings of vitrified cells at a nanometric resolution in near-native conditions and in three-dimensions. One consequent drawback of this technique is the sample thickness, for two reasons: i) achieving proper vitrification of the sample gets increasingly difficult with sample thickness, and ii) cryo-ET relies on transmission electron microscopy (TEM), requiring thin samples for proper electron transmittance (<500 nm). For samples exceeding this thickness limit, thinning methods can be used to render the sample amenable for cryo-ET. Cryo-focused ion beam (cryo-FIB) milling is one of them and despite having hugely benefitted the fields of animal cell biology, virology, microbiology, and even crystallography, plant cells are still virtually unexplored by cryo-ET, in particular because they are generally orders of magnitude bigger than bacteria, viruses, or animal cells (at least 10 μm thick) and difficult to process by cryo-FIB milling. Here, we detail a preparation method where abaxial epidermal onion cell wall peels are separated from the epidermal cells and subsequently plunge frozen, cryo-FIB milled, and screened by cryo-ET in order to acquire high resolution tomographic data for analyzing the organization of the cell wall.

Arabidopsis seed coat epidermal cells deposit a significant quantity of mucilage, composed of the cell wall components pectin, hemicellulose, and cellulose, into the apoplast during development. When mature seeds are hydrated, mucilage extrudes to form a gelatinous capsule around the seed. Determining the monosaccharide composition of both extruded mucilage and whole seeds is an essential technique for characterizing seed coat developmental processes and mutants with altered mucilage composition. This protocol covers growth of plants to produce seeds suitable for analysis, extraction of extruded mucilage using water and sodium carbonate (used for mutants with impaired mucilage release), and extraction of alcohol insoluble residue (AIR) from whole seeds. The prepared polysaccharides are then hydrolyzed using sulfuric acid, which hydrolyses all polysaccharides including cellulose. Sensitive and reproducible quantification of the resulting monosaccharides is achieved using high-performance anion exchange chromatography coupled with pulsed amperometric detection (HPAEC-PAD).

All bacteria, fungi and plant cells are surrounded by a cell wall. This complex network of polysaccharides and glycoproteins provides mechanical support, defines cell shape, controls cell growth and influences the exchange of substances between the cell and its surroundings. Despite its name, the cell wall is a flexible, dynamic structure. However, due to the lack of non-invasive methods to probe the structure, relatively little is known about the synthesis and dynamic remodeling of cell walls. Here, we describe a non-invasive method that quantifies a key physiological parameter of cell walls, the porosity, i.e., the size of spaces between cell wall components. This method measures the porosity-dependent decrease of the plasma membrane-localized fluorescent dye FM4-64 in the presence of the extracellular quencher Trypan blue. This method is applied to bacteria, fungi and plant cell walls to detect dynamic changes of porosity in response to environmental cues.

Plant cell walls consist of different polysaccharides and structural proteins, which form a rigid layer located outside of the plasma membrane. The wall is also a very dynamic cell composite, which is characterized by complex polysaccharide interactions and various modifications during cell development. The visualization of cell wall components in situ is very challenging due to the small size of cell wall composites (nanometer scale), large diversity of the wall polysaccharides and their complex interactions. This protocol describes immunogold labeling of different cell wall epitopes for high-resolution transmission electron microscopy (TEM). It provides a detailed procedure for collection and preparation of plant material, ultra-thin sectioning, specimen labeling and contrasting. An immunolabeling procedure workflow was optimized to obtain high efficiency of carbohydrates labeling for high-resolution TEM. This method was applied to study plant cell wall characteristics in various plant tissues but could also be applied for other cell components in plant and animal tissues.