植物科学

This is a protocol for quantitative determination of storage and total carbohydrates in algae and cyanobacteria. The protocol is simple, fast and sensitive and it requires only few standard chemicals. Great advantage of this protocol is that both storage and total saccharides can be determined in the cellular pellets that were already used for chlorophyll and carotenoids quantification. Since it is recommended to perform the pigments measurement in triplicates, each pigment analysis can generate samples for both total saccharide and glycogen/starch content quantification.

The protocol was applied for quantification of both storage and total carbohydrates in cyanobacteria Synechocystis sp. PCC 6803, Cyanothece sp. ATCC 51142 and Cyanobacterium sp. IPPAS B-1200. It was also applied for estimation of storage polysaccharides in Galdieria (IPPAS P-500, IPPAS P-507, IPPAS P-508, IPPAS P-513), Cyanidium caldarium IPPAS P-510, in green algae Chlorella sp. IPPAS C-1 and C-1210, Parachlorella kessleri IPPAS C-9, Nannochloris sp. C-1509, Coelastrella sp. IPPAS H-626, Haematococcus sp. IPPAS H-629 and H-239, and in Eustigmatos sp. IPPAS H-242 and IPPAS C-70.

We analyzed the reactive oxygen species (ROS) accumulation in the colony-forming green microalga Botryococcus braunii in response to several stress inducers such as NaCl, NaHCO₃, salicylic acid (SA), methyl jasmonate, and acetic acid. A staining assay using the fluorescent dye CellROX Green was used. CellROX Green is a fluorogenic probe used for measuring oxidative stress in live cells. The dye is weakly fluorescent inside cells in a reduced state but exhibits bright green photostable fluorescence upon oxidation by ROS and subsequent binding to DNA. The large amount of liquid hydrocarbons produced and excreted by B. braunii, creates a highly hydrophobic extracellular environment that makes difficult to study short times defense responses on this microalga. The procedure developed here allowed us to detect ROS in this microalga even within a short period of time (in minutes) after treatment of cells with different stress inducers.

Plant including green algal cells are surrounded by a cell wall, which is a diverse composite of complex polysaccharides and crucial for their function and survival. Here we describe two simple protocols to visualize callose (1→3-β-D-glucose) and cellulose (1→4-β-D-glucose) and related polysaccharides in the cell walls of streptophyte green algae. Untreated or algal cells heated in NaOH are incubated in Calcofluor white (binding to β-glucans including cellulose) or Aniline blue (binding to callose), respectively. Both dyes can be visualized by epifluorescence microscopy.

Cryo-electron tomography (CET) is a well-established technique for imaging cellular and molecular structures at sub-nanometer resolution. As the method is limited to samples that are thinner than 500 nm, suitable sample preparation is required to attain CET data from larger cell volumes. Recently, cryo-focused ion beam (cryo-FIB) milling of plunge-frozen biological material has been shown to reproducibly yield large, homogeneously thin, distortion-free vitreous cross-sections for state-of-the-art CET. All eukaryotic and prokaryotic cells that can be plunge-frozen can be thinned with the cryo-FIB technique. Together with advances in low-dose microscopy, this has shifted the frontiers of in situ structural biology. In this protocol we describe the typical steps of the cryo-FIB technique, starting with fully grown cell cultures. Three recently investigated biological samples are given as examples.

Eukaryotic cilia/flagella are ideal organelles for the analysis of membrane trafficking, membrane assembly, and the functions of a variety of signal transduction molecules. Cilia are peninsular organelles and the membrane lipids, membrane proteins, and microtubular-associated components are selectively transported into cilia through the region formed by the basal body/transition region and tightly associated ciliary membrane. Cilia can be isolated from many organisms without disrupting cells and many will rapidly regenerate cilia (with the ciliary membrane lipids and proteins) to replace those that are released. Despite their ease of isolation, we have relatively little understanding of the mechanisms that regulate lipid and protein transport into ciliary membranes (Pazour and Bloodgood, 2008; Bloodgood, 2009; Bloodgood, 2012).

Chlamydomonas flagella shed membrane vesicles, also called ectosomes (Wood et al., 2013) from flagellar tips and these vesicles can be purified from the culture medium without damaging or deflagellating cells (McLean et al., 1974; Bergman et al., 1975; Snell, 1976; Kalshoven et al., 1990). Based on a comparison of biotinylated proteins on the shed vesicles with biotinylated proteins isolated from purified flagella and cell bodies, the ectosomes contain most, but not all, flagellar surface proteins and none of the major cell body proteins (Dentler, 2013). Although ectosomes have only been purified from Chlamydomonas cells, preliminary evidence indicates that similar vesicles are released from Tetrahymena cilia (Dentler, unpublished).

Flagellar (and ciliary) membranes or membrane proteins also can be released from purified flagella/cilia. Most membrane proteins can be solubilized by extracting purified cilia with nonionic detergent [Triton X-100 or X-114 or Nonidet P-40 (NP-40)] and pelleting the microtubules (axonemes). However, not all membranes are released by detergent (Dentler, 1980) and the supernatant also contains all of the flagellar proteins that are not attached to the microtubules.

Intact membrane vesicles can be released from flagella by agitation of flagella, often with low concentrations of nonionic detergents or freeze-thawing (Witman et al., 1972; Snell, 1976; Dentler, 1980; Dentler, 1995; Bloodgood and May, 1982; Pasquale and Goodenough, 1987; Iomini et al., 2006; Huang et al., 2007). Once released, they can be purified from axonemes by differential centrifugation.

Each of these methods may enrich for different populations of axonemal and membrane proteins and lipids. The different solubility of membranes may reveal local differences in lipid or protein composition (Bloodgood, 2009). The ectosomes contain most but not all surface proteins found on purified Chlamydomonas flagella (Dentler, 2013). The ectosomes vesicles may be enriched in different soluble flagellar proteins than those trapped as vesicles are released from purified flagella. The detergent-solubilized “membrane+matrix” will contain all soluble membrane proteins as well as all of the soluble proteins in the flagellar compartment.

In this paper, a method to purify ectosomes vesicles released from the tips of living Chlamydomonas cells is presented as are two methods to release flagellar membrane vesicles and proteins from purified flagella.

Eukaryotic cilia/flagella are one of the only cellular structures that can be removed without injuring cells, can be highly purified for biochemical analysis, and, in many cells, can be completely reassembled within 90 minutes. Following amputation, the expression of many flagellar genes is up-regulated, and many are packaged and associated with intraflagellar transport (IFT) particles for transport to flagellar bases and into growing flagella. Studies of deciliation and ciliary growth provide insight to mechanisms that regulate microtubule assembly and length, mechanisms that regulate the transport of soluble cytoplasmic proteins into the ciliary compartment and their assembly into microtubules, and mechanisms that regulate trafficking of membrane proteins and lipids to the plasma membrane or to ciliary bases and their movement into and out of the cilium. These are important for motility and for signal transduction.

Deciliation methods for many cells have been developed and most require extracellular calcium ions and activation of signaling pathways that regulate microtubule severing (Quarmby, 2009). Deciliation occurs at the distal end of the basal bodies and, as soon as axonemes are severed, the membrane reseals and basal bodies begin to regenerate cilia.

Chlamydomonas is an ideal organism with which to study ciliary regeneration. Cells are easily and inexpensively cultured, flagellar amputation and regeneration is uniform in all cells in a population and growth can be assayed by observing fixed or living cells with a phase contrast microscope equipped and a 40x objective lens. Flagellar regeneration on individual living cells can be observed using paralyzed mutants immobilized in agarose. Because deflagellation leaves cells intact, the released flagella can be purified without contamination with cellular debris. The most reliable deciliation and regeneration method is the pH shock method developed by Reference 5 (also see References 4 and 11). Other methods are reviewed by Quarmby, (2009). The pH shock method is primarily used for Chlamydomonas but can be used for deciliation and regeneration of Tetrahymena cilia (Gaertig et al., 2013).

The protocol is very reliable and simple for inducing heat shock in unicellular green algae cells. The main purpose was to compare cellular response of three Chlorella species, isolated from different habitats: Chlorella vulgaris 8/1- thermophilic, Chlorella kesslery- mesophilic and C. vulgaris- extremophilic. Species were isolated from different habitats and differ in their temperature preferences and tolerance. Temperature induced stress response was measured as cell survival, induction of chloroplast HSP70B and DSBs induction and rejoining.

The method of immunoelectron microscopy is intended for localization of proteins inside the cells of Chlamydomonas reinhardtii or other microalgae and cyanobacteria. This protocol was used to study localization of carbonic anhydrase Cah3 with antibodies raised in rabbit, though it can be used to localize any other abundant protein. Primary rabbit antibodies are recommended because they react quickly and specifically with proteins of C. reinhardtii. If primary antibodies other than rabbit are used, the blocking procedure and time of incubation with primary and secondary antibodies should be adjusted.